FE R R E T S, R A B B I T S, and R O D E N T S C l i nical Medicine and Surgery T H I R D
E D I T I O N
Katherine E. Quesenberry DVM, MPH, Diplomate ABVP (Avian) Service Head Avian and Exotic Pet Service The Animal Medical Center New York, New York
James W. Carpenter MS, DVM, Diplomate ACZM Professor of Zoological Medicine Department of Clinical Sciences College of Veterinary Medicine Kansas State University Manhattan, Kansas
3251 Riverport Lane St. Louis, Missouri 63043
FERRETS, RABBITS, AND RODENTS: CLINICAL MEDICINE AND SURGERY Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
ISBN: 978-1-4160-6621-7
Some material was previously published. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Images in Chapter 34 © Stephen J. Divers. Notice Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods, they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. With respect to any drug or pharmaceutical products identified, readers are advised to check the most current information provided (i) on procedures featured or (ii) by the manufacturer of each product to be administered, to verify the recommended dose or formula, the method and duration of administration, and contraindications. It is the responsibility of practitioners, relying on their own experience and knowledge of their patients, to make diagnoses, to determine dosages and the best treatment for each individual patient, and to take all appropriate safety precautions. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Previous editions copyrighted 2004 and 1997. Library of Congress Cataloging-in-Publication Data Ferrets, rabbits, and rodents : clinical medicine and surgery / [edited by] Katherine E. Quesenberry, James W. Carpenter. — 3rd ed. p. ; cm. Includes bibliographical references and index. ISBN 978-1-4160-6621-7 (pbk. : alk. paper) I. Quesenberry, Katherine E. II. Carpenter, James W. (James Wyman), 1945[DNLM: 1. Ferrets—surgery. 2. Rabbits—surgery. 3. Rodentia—surgery. 4. Surgical Procedures, Operative—veterinary. SF 997.3] 636.932’2—dc23 2011039100 Vice President and Publisher: Linda Duncan Publisher, Veterinary Medicine: Penny Rudolph Associate Developmental Editor: Brandi Graham Publishing Services Manager: Catherine Jackson Senior Project Manager: Carol O’Connell Design Direction: Paula Catalano Printed in the United States of America Last digit is the print number: 9 8 7 6 5 4 3 2 1
Contributors Sean Aiken, DVM, MS, Diplomate ACVS Veterinary Specialty Hospital San Diego, California Natalie Antinoff, DVM, Diplomate ABVP (Avian) Gulf Coast Veterinary Specialists Avian and Exotics Houston, Texas Heather W. Barron, DVM, Diplomate ABVP (Avian) Professor and Chair Department of Veterinary Clinical Sciences School of Veterinary Medicine St. Matthew’s University Grand Cayman, Cayman Islands, British West Indies Louise Bauck, DVM, MVSc Professor of Biology Department of Math and Science Brenau University Gainesville, Georgia Teresa Bradley Bays, DVM, CVA Director Belton Animal Clinic and Exotic Care Center Animal Urgent Care of Cass County Belton, Missouri Judith A. Bell, DVM, PhD Department of Population Medicine Ontario Veterinary College University of Guelph Guelph, Ontario, Canada R. Avery Bennett, DVM, MS, Diplomate ACVS Chief of Surgery The Animal Medical Center New York, New York Cynthia R. Bishop, DVM Assistant Professor Department of Veterinary Clinical Sciences Seattle Pacific University Seattle, Washington Cynthia Brown, DVM, Diplomate ABVP (Avian) Avian and Exotic Medicine New England Veterinary Medical Center Mystic, Connecticut Susan A. Brown, DVM Rosehaven Exotic Animal Veterinary Service North Aurora, Illinois Michelle L. Campbell-Ward, BSc, BVSc (Hons I), DZooMed, MRCVS Taronga Western Plains Zoo Dubbo, NSW, Australia
Vittorio Capello, DVM, Diplomate ABVP (Exotic Companion Mammal), Diplomate ECZM (Small Mammal) Exotic Companion Mammal Medicine and Surgery Clinica Veterinaria S.Siro Clinica Veterinaria Gran Sasso Milano, Italy Stephen J. Divers, BSc (Hons), BVetMed, Diplomate ACZM, Diplomate ECZM (Herpetology), Diplomate ZooMed, FRCVS Professor of Zoological Medicine Department of Small Animal Medicine & Surgery College of Veterinary Medicine University of Georgia Athens, Georgia Thomas M. Donnelly, BVSc, Diplomate ACLAM Warren Institute Ossining, New York Peter G. Fisher, DVM Director Pet Care Veterinary Hospital Virginia Beach, Virginia Anthony J. Fischetti, DVM, MS, Diplomate ACVR Department Head of Diagnostic Imaging The Animal Medical Center New York, New York James G. Fox, DVM, MS, Diplomate ACLAM Professor and Director Division of Comparative Medicine Massachusetts Institute of Technology Cambridge, Massachusetts Carley J. Giovanella, DVM, Diplomate ACVIM (Neurology) Gulf Coast Veterinary Neurology and Neurosurgery Houston, Texas Jennifer Graham, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal), Diplomate ACZM Avian and Exotic Medicine Angell Animal Medical Center Boston, Massachusetts Michelle G. Hawkins, VMD, Diplomate ABVP (Avian) Associate Professor, Companion Avian and Exotic Pet Medicine Department of Medicine and Epidemiology School of Veterinary Medicine University of California, Davis Davis, California Laurie Hess, DVM, Diplomate ABVP (Avian) Veterinary Center for Birds and Exotics Bedford Hills, New York
iii
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CONTRIBUTORS
Heidi L. Hoefer, DVM, Diplomate ABVP (Avian) Island Exotic Veterinary Care Huntington Station, New York; Consultant and Adjunct Clinician Exotics Emergency and Critical Care Long Island Veterinary Specialists Plainview, New York Sharon M. Huston, DVM, Diplomate ACVIM (Cardiology) San Diego Veterinary Cardiology San Diego, California Evelyn Ivey, DVM, Diplomate ABVP (Avian) Four Corners Veterinary Hospital Concord, California Jeffrey R. Jenkins, DVM, Diplomate ABVP (Avian) Avian and Exotic Animal Hospital San Diego, California
Marla Lichtenberger, DVM, Diplomate ACVECC Department of Emergency and Critical Care Milwaukee Emergency Center for Animals Greenfield, Wisconsin Teresa Lightfoot, DVM, Diplomate ABVP (Avian) Avian and Exotic Service BluePearl Veterinary Partners Tampa, Florida Andrew S. Loar, DVM, Diplomate ACVIM ALX Laboratories The Animal Medical Center New York, New York Lori Ludwig, VMD, MS, Diplomate ACVS Veterinary Surgical Care, LLC Mt. Pleasant, South Carolina
Cathy A. Johnson-Delaney, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal) Avian and Exotic Animal Medicine Center Kirkland, Washington
Douglas R. Mader, MS, DVM, Diplomate ABVP (Canine and Feline) Marathon Veterinary Hospital Marathon Sea Turtle Hospital Conch Republic
Amy S. Kapatkin, DVM, MS, Diplomate ACVS Associate Professor of Orthopedic Surgery Department of Surgical and Radiological Sciences College of Veterinary Medicine University of California–Davis Davis, California
Christoph Mans, MedVet Special Species Health Service Department of Surgical Sciences School of Veterinary Medicine University of Wisconsin Madison, Wisconsin
Eric Klaphake, DVM, Diplomate ACZM, Diplomate ABVP (Avian) Animal Medical Center Bozeman, Montana
Mark A. Mitchell, DVM, MS, PhD, Diplomate ECZM (Herpetology) Professor, Zoological Medicine College of Veterinary Medicine University of Illinois Urbana, Illinois
Marc S. Kraus, DVM, Diplomate ACVIM (Cardiology, Internal Medicine) Senior Lecturer Department of Clinical Sciences College of Veterinary Medicine Cornell University Ithaca, New York Pamela Ming-Show Lee, DVM, MS Cardiology Service The Animal Medical Center New York, New York Angela M. Lennox, DVM, Diplomate ABVP (Avian) Avian and Exotic Animal Clinic Indianapolis, Indiana; Adjunct Assistant Professor Department of Veterinary Clinical Sciences School of Veterinary Medicine Purdue University West Lafayette, Indiana
James K. Morrisey, DVM, Diplomate ABVP (Avian) Senior Lecturer Exotic and Wildlife Medicine Department of Clinical Sciences College of Veterinary Medicine Cornell University Ithaca, New York Robert D. Ness, DVM Ness Exotic Wellness Center Lisle, Illinois Barbara L. Oglesbee, DVM, Diplomate ABVP (Avian) Clinical Associate Professor Department of Veterinary Clinical Science College of Veterinary Medicine The Ohio State University Capital Veterinary Referral and Emergency Center Columbus, Ohio
CONTRIBUTORS Connie Orcutt, DVM, Diplomate ABVP (Avian, Exotic Companion Mammals) Avian and Exotic Animal Medicine Putnam Veterinary Clinic Topsfield, Massachusetts Peter J. Pascoe, BVSc, Diplomate ACVA, DVA, Diplomate ECVAA Professor Department of Surgical and Radiological Sciences School of Veterinary Medicine University of California, Davis Davis, California Joanne Paul-Murphy, DVM, Diplomate ACZM Professor Department of Companion Avian and Exotic Pets School of Veterinary Medicine University of California, Davis Davis, California Anthony A. Pilny, DVM, Diplomate ABVP (Avian) Avian and Exotic Pet Medicine Veterinary Internal Medicine and Allergy Specialists New York, New York Christal G. Pollock, DVM, Diplomate ABVP (Avian) Veterinary Consultant Lafeber Company Cleveland, Ohio Lauren V. Powers, DVM, Diplomate ABVP (Avian) Avian and Exotic Pet Service Carolina Veterinary Specialists Huntersville, North Carolina Karen L. Rosenthal, DVM, MS Associate Professor of Special Species Medicine School of Veterinary Medicine University of Pennsylvania Philadelphia, Pennsylvania Jonathan Rubinstein, DVM Avian and Exotic Service BluePearl Veterinary Partners Tampa, Florida Andrea Siegel, DVM ALX Laboratories The Animal Medical Center New York, New York Kathy Tater, DVM, Diplomate ACVD Master’s of Public Health Candidate in Quantitative Methods Harvard School of Public Health; Clinical Assistant Professor Cummings School of Veterinary Medicine Tufts University North Grafton, Massachusetts
v
Thomas N. Tully, Jr., DVM, MS, Diplomate ABVP (Avian), Diplomate ECZM (Avian) Professor of Zoological Medicine Department of Veterinary Clinical Sciences School of Veterinary Medicine Louisiana State University Baton Rouge, Louisiana Alexandra van der Woerdt, DVM, MS, Diplomate ACVO, Diplomate ECVO Staff Ophthalmologist The Animal Medical Center New York, New York David Vella, BSc, BVSc (Hons), Diplomate ABVP (Exotic Companion Mammal) North Shore Veterinary Specialist Centre; Animal Referral Hospital Sydney, New South Wales, Australia James Walberg, DVM, Diplomate ACVP Consultant Department of Pathology The Animal Medical Center New York, New York Bruce H. Williams, DVM, Diplomate ACVP Senior Pathologist Veterinary Pathology Service Joint Pathology Center Washington, D.C. Nicole R. Wyre, DVM, Diplomate ABVP (Avian) Chief, Special Species Section Matthew J. Ryan Veterinary Hospital School of Veterinary Medicine University of Pennsylvania Philadelphia, Pennsylvania Ashley Zehnder, DVM, Diplomate ABVP (Avian) Post Doctoral Scholar Department of Comparative Medicine Stanford University Stanford, California
I dedicate this book to all of my friends and very close colleagues who have supported me through some very difficult times in these last 5 years. Without their encouragement and support, pulling this book together in the face of both personal and professional challenges would not have been possible. In particular, I thank Connie Orcutt, Laurie Hess, Tom Donnelly, Heidi Hoefer, Susan Orosz, and my co-editor, Jim Carpenter, all of whom I have worked with and learned from for many years in this profession. I thank my sister, Marcia Quesenberry, for her unwavering support, encouragement, and love. I also give special thanks and love to my children, Zachary and Chelsea Messinger, who are always at the center of my life and being. Katherine E. Quesenberry I wish to acknowledge all those who have contributed to our knowledge and understanding of small mammal medicine; Dr. Kathy Quesenberry for graciously cajoling me into collaborating on yet another edition of the Pink Book; and the many colleagues, interns and residents, and students who have inspired my professional life. I also wish to thank veterinary students Caitlin Burrell, Richard Brooksby, and Amy Guersey for their “office assistance,” and especially Dr. Chris Marion for his editorial assistance in the preparation of this edition of Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery. I would like to dedicate this book to my family (wife, Terry; son, Michael; and daughter, Erin, and her family–husband, Steve, and kids, Kylie, Hayden, and Asher) who have supported me as I pursue my passion for zoological medicine. James W. Carpenter
Preface In the 15 years since the first edition of Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, the specialty of exotic pet mammal medicine and surgery has become more mainstream in veterinary practice, and, as a matter of fact, has become an integral component of most small animal clinical practices. The knowledge base has expanded tremendously as interest in small mammals has prompted research in these animals not as just as laboratory species, but as companion animals. As information about these species has become much more accessible via Internet websites and chat groups, the public has increasingly recognized that these species are valued as pets and deserving of high quality veterinary care. Many pet owners look beyond the financial value of these animals and expect state of the art veterinary care at the same level as that given to dogs and cats. The numbers of small mammals, particularly pet rabbits, chinchillas, and guinea pigs, presented to veterinarians for health care has grown steadily, as a consequence of more accessible information about their care to the public as well as the increase in spending on companion animal care in general. As veterinarians, we therefore must be able to provide a high level of medical and surgical care, based on a solid knowledge base, for these pets. The number of books, publications, and websites that are now devoted to the husbandry and veterinary care of small pet mammals is enormous. Whereas previously only a few veterinary texts were published about these species, now there are many books, serial publications, and journal articles available on various topics ranging from medicine, surgery, imaging, and clinical techniques to behavior. Some of this work is original, and some works only present the same material reworded into different formats.
With this third edition of Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, we have tried to stay true to our original mission of publishing up-to-date information in a reader-friendly, comprehensive yet concise format. All of the chapters have been updated, with new authors on many of the topics and new chapters added on “Emergency and Critical Care” and “Behavior.” As the information about these species has exploded, we have tried to focus on the most pertinent and reliable information to present to our readers. Our authors are among the most respected in veterinary medicine and encompass a broad range of specialists in exotic pet mammals, internal medicine, surgery, critical care, and laboratory animal medicine. We are proud to present this third edition of Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery to our colleagues and readers, and we are especially pleased that this edition is in color. We are very grateful to the chapter authors and to our team at Elsevier, especially Penny Rudolph, Brandi Graham, and Carol O’Connell, for their patience and very hard work in bringing this publication together. We are confident that the format, presentation, information, and reliability of the “Pink Book” will continue to set it apart as the standard in this subspecialty of veterinary medicine.
Katherine E. Quesenberry, DVM, MPH, Diplomate ABVP (Avian)
James W. Carpenter, MS, DVM, Diplomate ACZM
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SECTION ONE
Ferrets
CHAPTER
1
Basic Anatomy, Physiology, and Husbandry
Lauren V. Powers, DVM, Diplomate ABVP (Avian), and Susan A. Brown, DVM
Domestication History Uses Anatomy and Physiology Integument Gastrointestinal System Urogenital System Cardiovascular and Lymphatic Systems Respiratory System Endocrine System Musculoskeletal System Neurologic System and Special Senses Physiology and Reproduction Physiology Body Size and Seasonal Weight Variation Reproduction Husbandry Housing Environmental Enrichment Nutrition
DOMESTICATION HISTORY Ferrets belong to the family Mustelidae and are related to weasels, mink, otters, badgers, stoats, and martens. There are currently three living species of ferrets (also known as polecats in Europe and Asia): the European polecat (Mustela putorius), the Steppe or Siberian polecat (Mustela eversmanni), and the blackfooted ferret (Mustela nigripes). All three species live primarily solitary social lives and are very efficient hunters supporting Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
their obligate carnivore lifestyle. The European polecat is found in various areas from the Atlantic to the Ural mountains and dwells along the edges of woodlands and wetlands.12 The Siberian polecat is found in Eurasia from the thirtieth to the sixtieth degree of latitude, may be larger than the European polecat, and lives primarily in open areas such as steppes, slopes of ravines, and semi-deserts.12 The black-footed ferret is native to the prairies of North America. It almost became extinct in the wild because of habitat destruction and the decimation of its main food source, the prairie dog, from poisoning and hunting.12 Currently, captive breeding and reintroduction programs are under way in an attempt to reestablish the black-footed ferret into its native range. It is illegal to own this endangered species. The origin of the domestic ferret, which is traditionally referred to as Mustela putorius furo, is shrouded in mystery. The Latin name translates loosely as “mouse-eating (mustela) smelly (putorius) thief (furo).” Currently there is a move toward using nomenclature that differentiates the wild progenitors of a domesticated species from the domesticate, and some mammologists are moving toward referring to the domesticated ferret as Mustela furo.5,11 The domesticated ferret may have originated from either the Siberian or the European polecat, or possibly both.5,8,33 It is difficult to find archaeological evidence of domestication, possibly because of the ferret’s small skeleton, which may have deteriorated rapidly or was indistinguishable from wild ferrets living in the environment, or the lack of paraphernalia associated with the ferret, making them archaeologically unimportant.5 Ferrets have been domesticated for approximately 2,000 to 3,000 years.5 The first clear reference to domestication is in the writings of the Spaniard Isidore of Seville in 622 ad.5 There is a high probability that ferrets were brought by Romans or Normans during their invasions, but there are currently no references that irrefutably link Romans and Greeks with domestication of ferrets.5 It is likely that ferrets 1
2
SECTION I Ferrets
were first domesticated over a wide area somewhere in the south to southeastern portion of Europe near the Mediterranean.5 Over the centuries, numerous references have been made to the use of ferrets in Europe for rodent control in homes, farms, and ships and for hunting rabbits both for damage control and for human food, as well as for their pelts. The domestic ferret was introduced into Australia from Europe in the 1800s to control the populations of European rabbits that had been previously released.19 Fortunately, enough other predators, such as foxes, dingoes, and hawks, preyed on the ferret so that feral populations never developed.19 However, when they were introduced into New Zealand for the same reason in the late 1800s along with stoats and weasels, there were no predators to control their numbers.19 Feral populations of domestic ferrets therefore developed and are still present today.8,19 The impact of feral ferrets on native wildlife has been controversial. The domestic ferret was probably introduced into the United States from Europe by the shipping industry in the 1700s. They may have come as pets or as hunting companions.8,19 Ferrets were also used for their pelts and the town of New London, Ohio, became known as Ferretville because of its huge breeding population of ferrets around 1915.19 At the turn of the twentieth century, hunting with ferrets was banned in some states to protect against destruction of the native rabbit population.19 The pet ferret has changed from its wild progenitor in the process of domestication in the areas of physiology, reproduction, and behavior. Reproductively, ferrets mate two or more times a year compared to polecats at generally once a year.5 The ferret’s litter size average of eight is larger than the polecat’s at six.5 Ferret coat color has changed, as is the case with most animals that are domesticated. Albino ferrets have been bred for centuries and are often preferred for hunting because of their high visibility. Albino ferrets may also have alterations in their vision and hearing.5 Other changes include a 15% to 20% smaller cranial capacity, a wider postorbital constriction, and dental crowding.5 Behaviorally, ferrets appear to be more gregarious than their wild counterparts, but this may be due to juvenilization, which is a common side effect of domestication that allows ferrets to live in “litter groups” rather than their more naturally solitary lifestyle.5 The most noticeable behavioral change is a loss of the innate fear of humans, as well as a lack of fear toward unfamiliar objects in their environment.5
USES Historically, humans have not domesticated animals primarily for the purpose of companionship. Animals needed to serve an economic purpose and the same holds true for the ferret. Early references to ferrets record their use for rodent or rabbit control.8,19 Ferrets are efficient little predators that can bring down prey quite a bit larger than themselves and can maneuver in small spaces more effectively than cats. Ferrets were used on ships in colonial days to control the rat populations.8 In the early 1900s, the U.S. Department of Agriculture encouraged the use of ferrets as a means of controlling rabbits, raccoons, gophers, mice, and rats around granaries and farms.8 One needed only to call the local “ferret master” to bring out his ferrets, which were set loose to do their work and then recaptured to work another day. Large facilities kept their own ferrets on site. Ferrets are still used for rodent and rabbit control in some areas of Europe and Australia today. However, hunting with ferrets is prohibited in the United States.
Ferrets have long been used to hunt rabbits—not only for control, but as a food source for human beings. “Ferreting” was a common sport in the United Kingdom and many other areas of Europe. It is still practiced today but to a much lesser degree. Ferrets are released in a rabbit warren area, where they investigate burrows and flush out rabbits. The rabbits are then caught in nets or by dogs or shot by the waiting hunter as they exit their burrows. Domestic ferrets have been bred for their pelts. A coat made of ferret fur is referred to as fitch. Ferret hair has also been used in other products such as artist’s brushes.19 Ferret fur never really took hold in the United States, but it still exists in a few areas of Northern Europe. An entertainment peculiar to English pubs and still found in a few isolated areas of the United Kingdom is called ferret- legging. This is a sport in which a man securely ties his trouser legs closed at the ankles and then places two ferrets, each with a full set of teeth, into his trousers. He then securely ties the trousers closed at the waist. The contest is to see how long he can stand having the ferrets in his trousers. If a ferret bites, it can only be dislodged from the outside of the trousers. The record of 5 hours and 26 minutes was set by a 72-year-old Yorkshire man.8 Domestic ferrets have also been used to transport cables through long stretches of conduit. They have been used to string cable for oilmen of the North Sea, for camera crews, in jets, and for telephone companies.8 Ferrets have been used in biomedical research since the early 1900s, when they were used to study human influenza and other viral diseases.8 Today ferrets are used in the fields of virology, reproductive physiology, anatomy, endocrinology, and toxicology.8 Although the use of ferrets in research is very distasteful to some, much of the information gained has directly benefited the pet ferret as well. The main use for ferrets today, however, particularly in the United States, is as a companion animal. Their popularity has increased dramatically over the past few decades. There has been a proliferation of ferret organizations dedicated to the wellbeing of this pet. It is difficult to say when the first ferret was kept strictly as a pet, but it is hard to imagine people in the distant past not feeling some attraction to the engaging personality of this animal. Ferrets make suitable pets for many people. They are small, clean, and very interactive with human beings and each other. However, as with all companion animals, the prospective owner should be educated on their husbandry requirements and behavior. For instance, ferrets (as with most pets) are not suitable for children younger than 6 years. Another consideration is that the majority of ferrets in the United States will likely be afflicted by one or more neoplastic diseases as they age. In addition, certain legal restrictions relate to the ownership of ferrets. Ferrets are still not considered domestic animals in most areas of the United States despite their long history of domestication. In some areas, owning a ferret as a pet is illegal, and in other areas permits must be obtained for ownership. With the advent of an approved rabies vaccine for the domestic ferret, restrictions on their use as pets have been lifted in many parts of the United States. However, in some localities, even if the ferret is appropriately vaccinated, it can be seized and destroyed if it bites a human being. Veterinarians should therefore be familiar with legislation not only in their state, but in their specific county or city regarding the keeping of ferrets before they engage in ferret veterinary care.
CHAPTER 1 Basic Anatomy, Physiology, and Husbandry
ANATOMY AND PHYSIOLOGY The basic body plan of the domestic ferret is similar to that of other carnivores. The following is a brief review of the clinically relevant anatomic and physiologic features of the ferret. Skeletal anatomy is depicted in Figure 1-1, visceral anatomy in Figure 1-2, and normal radiographic anatomy in Figures 1-3 and 1-4. For radiographic views of select pathologic conditions, anatomy, see Chapter 35. Selected physiologic values are detailed in Table 1-1. The reader is directed to publications containing more extensive reviews of ferret anatomy and physiology.7,9,10,17,31
INTEGUMENT Coat The domestic ferret possesses a fine undercoat and coarse, long guard hairs that provide excellent insulation.7 There are no specific breeds of ferrets, but color and pattern standards exist (Fig. 1-5). The American Ferret Association recognizes the following color standards for the purposes of show and breeding—albino, black, black sable, champagne, chocolate, dark-eyed white, and sable. Recognized pattern standards for these colors include solid, standard, color point (Siamese), blaze, panda, roan, and dark-eyed white. Mask configuration can change from season to season and from year to year, making photography an unreliable method of individual identification. Ferrets living outdoors tend to be darker in color.20 Ferrets undergo a heavy shed in the spring and the fall as seasonal weight changes occur. The coat may be shorter in summer months and longer in the fall, and lighter in color in the winter and darker in the fall. Sexually altered ferrets of either gender
3
have a less dramatic molt and color change than intact animals. Clients should be warned that fur shaved for a procedure may not be replaced for weeks to months, and that the fur may initially discolor the skin a bluish hue before erupting and may have a different color or texture than surrounding fur.
Skin and Associated Glands The thick skin and muscle found on the neck and shoulders of a ferret protect it from trauma during fighting and mating. Ferrets have very active sebaceous glands, which account for their strong musky odor.14 During the breeding season, intact animals have increased sebaceous secretions; this increase results in a noticeable increase in odor, yellow to orange discoloration of the undercoat, and oily fur.14 Ferrets lack sweat glands, and in part for this reason they are very susceptible to heat prostration.14,15,25
Anal Glands Ferrets possess a pair of well-developed anal glands, as do all mustelids. These glands produce a serous yellow liquid with a powerful odor. Ferrets that are frightened or threatened can express their anal glands but, unlike skunks, are unable to project the fluid over long distances.14,15 The anal gland ducts are located at about 4 o’clock and 8 o’clock and open into the anal canal. The glands are typically about 10 mm × 5 mm in size.20 Striated external anal sphincter muscle encloses the duct of each anal sac.7,15,20 Ferrets raised at large commercial breeding facilities in the United States are routinely descented between 5 and 6 weeks of age. This is despite the fact that the majority of the odor from a ferret arises from the sebaceous glands.14
Fig. 1-1 Skeletal anatomy of a ferret. 1, Calvaria; 2, hyoid apparatus; 3, larynx; 4, seven cervical vertebrae; 5, clavicle; 6, scapula; 7, 15 thoracic vertebrae; 8, five lumbar vertebrae; 9, three sacral vertebrae; 10, 18 caudal vertebrae; 11, first rib; 12, manubrium; 13, sternum; 14, xiphoid process; 15, humerus; 16, radius; 17, ulna; 18, carpal bones; 19, accessory carpal bone; 20, metacarpal bones; 21, ilium; 22, ischium; 23, pubis; 24, femur; 25, patella; 26, fabella; 27, tibia; 28, fibula; 29, tarsal bones; 30, calcaneus; 31, metatarsal bones; 32, talus; 33, os penis. (Adapted from An NQ, Evans HE. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:14.)
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SECTION I Ferrets
Fig. 1-2, see legend on opposite page.
CHAPTER 1 Basic Anatomy, Physiology, and Husbandry
5
Fig. 1-2 A, Ventral aspect of the viscera of a ferret in situ. B, Anatomy of the viscera and most important blood vessels as seen after removal of the lungs, liver, and gastrointestinal tract. 1, Larynx; 2, trachea; 3, right cranial lobe of lung; 4, left cranial lobe of lung; 5, right middle lobe of lung; 6, right caudal lobe of lung; 7, left caudal lobe of lung; 8, heart; 9, diaphragm; 10, quadrate lobe of liver; 11, right medial lobe of liver; 12, left medial lobe of liver; 13, left lateral lobe of liver; 14, right lateral lobe of liver; 15, stomach; 16, right kidney; 17, spleen; 18, pancreas; 19, duodenum; 20, transverse colon; 21, jejunoileum; 22, descending colon; 23, uterus; 24, ureter; 25, urinary bladder; 26, right common carotid artery; 27, left common carotid artery; 28, vertebral artery; 29, costocervical artery; 30, superficial cervical artery; 31, axillary artery; 32, right subclavian artery; 33, right internal thoracic artery; 34, left internal thoracic artery; 35, branch to thymus; 36, left subclavian artery; 37, brachiocephalic (innominate) artery; 38, cranial vena cava; 39, aortic arch; 40, right atrium; 41, pulmonary trunk; 42, left atrium; 43, right ventricle; 44, left ventricle; 45, caudal vena cava; 46, aorta; 47, esophagus; 48, hepatic veins; 49, celiac artery; 50, cranial mesenteric artery; 51, left adrenolumbar vein; 52, left adrenal gland; 53, right adrenal gland; 54, left renal artery and vein; 55, left kidney; 56, suspensory ligament of ovary; 57, left ovarian artery and vein; 58, left ovary; 59, left deep circumflex iliac artery and vein; 60, caudal mesenteric artery; 61, broad ligament of uterus; 62, left external iliac artery; 63, right common iliac vein; 64, left internal iliac artery; 65, rectum. (Adapted from An NQ, Evans HE. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:14.)
1 2 3 4
11
5 2
2 6
7 12
8
13 9 8
10
L
A
R
R
B Fig. 1-3 A, Ventrodorsal radiograph of a 1-year-old, spayed female ferret. Note normal positioning of thoracic and abdominal viscera. B, Same radiograph as (A): 1, trachea (endotracheal tube within lumen); 2, lung; 3, cranial mediastinum; 4, left primary bronchus; 5, heart; 6, liver; 7, stomach; 8, spleen; 9, left kidney; 10, urinary bladder; 11, right primary bronchus; 12, small intestine; 13, right kidney. (Silverman S, Tell LA. Radiology of rodents, rabbits, and ferrets: an atlas of normal anatomy and positioning. St. Louis: Elsevier Saunders; 2005:233.)
L
SECTION I Ferrets
6
A 1
2
3
4
5
2
10
11 2 10
2
6
7
8
9
B 12
13
8
14
Fig. 1-4 A, Lateral (right lateral recumbency) radiograph of a 1-year-old, spayed female ferret. Note normal positioning of thoracic and abdominal viscera. B, Same radiograph as (A): 1, trachea (endotracheal tube within lumen); 2, lung; 3, pulmonary vasculature; 4, bronchus; 5, pulmonary vein; 6, stomach; 7, kidney; 8, spleen; 9, colon; 10, intrathoracic adipose tissue; 11, heart; 12, liver; 13, small intestine; 14, urinary bladder. (Silverman S, Tell LA. Radiology of rodents, rabbits, and ferrets: an atlas of normal anatomy and positioning. St. Louis: Elsevier Saunders; 2005:232.)
GASTROINTESTINAL SYSTEM Teeth and Salivary Glands The ferret dentition is typical of carnivores, consisting of long curved canine teeth and resilient molars and premolars. The permanent teeth erupt between 50 and 74 days of age. There are 34 adult teeth, and the dental formula of the adult ferret is 2(I33 C11 P33 M12).14,15,18,20 The incisors and canine teeth possess a single root. The premolars have two roots each except for the upper third premolar (carnassial tooth), which has three roots. The upper molar and first lower molar have three roots, and the tiny second lower molar has only one.7 This formula differs slightly from other carnivores in that ferrets have three rather than four premolars.14,18 Supernumerary incisors are common.3,14,20 Ferrets possess five major pairs of salivary glands—the parotid, mandibular, sublingual, molar, and zygomatic glands.15,20
Esophagus, Stomach, and Intestines The muscle of the ferret esophagus is striated along the entire length cranial to the diaphragm.15 There is not a true gastroesophageal sphincter, and ferrets are able to vomit.15,18,20 However, in contrast to other carnivores, ferrets with gastrointestinal obstruction usually are not presented with a history of vomiting.20 The stomach is relatively simple, being roughly J-shaped and consisting of a cardia, body, pyloric antrum, and fundus.14,20 The stomach is separated from the liver by the lesser omentum.7 The stomach is capable of enormous distention18,20 and can easily hold 50 mL/kg or more.20 The small intestine is comparatively short, with reported lengths of 182 to 198 cm in the adult, and with a ratio of intestinal length to body length of about 5:1.7,15,18,28 This short intestinal length contributes to the comparatively short gastrointestinal transit time of 3 to 4 hours in the adult ferret.4
CHAPTER 1 Basic Anatomy, Physiology, and Husbandry
7
Table 1-1 Selected Biologic Values for the Domestic Ferret9,14,15,20 Parameter
Gender
Value
Body weight
Intact male Intact female Neutered, both genders
1-2 kg 0.6-1.0 kg 0.8-1.2 kg
Life span Rectal temperature Heart rate Blood volume Systolic blood pressure, awake Respiratory rate Tidal volume Stomach capacity Gastrointestinal transit time Urinary output Bladder capacity Urine pH Puberty
Reproductive life span
Male Female Male Female
Male Female Male Female
Gestation Litter size Birth weight Eyes and ears open Weaning age Maintenance fluid needs Maintenance caloric needs
6-12 yr 101.8°F (range 100-104°F [37.8-40°C]) 200-400 beats/min 60 mL 40 mL 161 mm Hg 133 mm Hg 33-36 breaths/min 10-11 mL/kg 50 mL/kg when distended 2.5 to 3.6 hr (meat); liquids may reach rectum within 1 hr 1 mL/hr (range 0.33-5.8 mL/hr) Approximately 5 mL/kg (higher under increased pressure) 6.5-7.5 9 months (as early as 23 wk with photoperiod manipulation) 8-12 months (as early as 16 wk with photoperiod manipulation) Throughout life 2-5 yr 41 days (range 39-42 days) 8 kits (range 1-18 kits) 6-12 g 28-34 days 6-8 wk Unknown, estimated at 60 mL/kg/day 200-300 kcal/kg/day
indistinguishable and are coiled together as the jejunoileum.15 The ferret lacks a cecum and appendix, and therefore the ileocolic junction is grossly indistinguishable but is typically defined as the region at which the ileocolic and jejunal arteries join.15,18 The large intestine is about 10 cm in length, consisting of the colon, rectum, and anus. There is an ascending, transverse, and descending colon.7
Liver, Gallbladder, and Pancreas
Fig. 1-5 Sable coloring of a domestic ferret. (Photography subjects provided by J. Ball.) The duodenum is about 10 cm in length and consists of three portions—the shorter (2 cm) cranial portion, the descending portion (5 cm), and the ascending portion (3 cm). The mesoduodenum encloses the right limb of the pancreas and a portion of the lesser omentum.7 The jejunum and ileum are
The liver is relatively large in the ferret and consists of six lobes—left lateral, left medial, quadrate, right medial, right lateral, and caudate.7,15,18,20 The gallbladder sits in a fossa between the quadrate and right medial liver lobes and averages 2 cm in length and 1 cm in width.7,15,20 The cystic duct joins the left, right, and central hepatic ducts to form the common bile duct, although variations of this formula exist.7 The pancreas is V-shaped and divided into right and left lobes connected by a body that lies close to the pyloris and is contained within the mesoduodenum.7 The left lobe extends along the dorsal caudal stomach and medial to the spleen. The right lobe closely follows the descending duodenum. Ducts from the left and right lobes connect to form the common pancreatic duct that joins the bile duct, which opens into the duodenal lumen about 3 cm caudal to the cranial duodenal flexure.7
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SECTION I Ferrets
UROGENITAL SYSTEM Kidneys, Ureters, and Bladder The kidneys are bean-shaped in the ferret and average 2.4 to 3.0 cm in length, 1.20 to 1.35 cm in width, and 1.10 to 1.35 in thickness.7 Both kidneys are retroperitoneal and covered by a thin peritoneal membrane. The cranial margin of the right kidney sits in a fossa of the caudate lobe of the liver.7 The ureters pass from the renal pelvis and extend caudally along the ventral aspect of the psoas muscle, entering the dorsolateral bladder just caudal to the its neck.7 The urinary bladder sits ventrally in the abdomen just cranial to the pelvic inlet. Although the bladder is small, it can easily hold 10 mL of urine at low pressure.15,31
Male Reproductive Tract The reproductive tract of the intact male (hob) resembles that of the dog, with a palpable os penis.14,15 Unlike the dog, however, the tip to the os penis is J-shaped, making urethral catherization difficult.15 The prepuce is a fold of skin reflected over the penis, containing fur externally but bare within.7 The preputial opening lies just caudal to the umbilical area on the ventral abdomen. The scrotum lies just caudal to the caudal margin of the os penis.7 The prostate gland is the single accessory reproductive gland in the male ferret. It is a fusiform structure that surrounds the proximal urethra and measures approximately 1.5 cm long and 0.6 cm across.16 Each ductus deferens opens into the urethra at the level of the prostate.7
Female Reproductive Tract The female reproductive tract of the intact female ferret (jill) closely resembles that of other carnivores. Two long, tapering uterine horns are present, as well as a short uterine body and a single cervix.7,15,29 The ovaries are paired and located just caudal to the kidneys. The ovary is attached to the wall of the abdominal cavity by the suspensory ligament cranially and to the uterine horn caudally by the proper ligament. The uterus is suspended by the broad and round ligaments. The urethra opens into the vaginal floor at the urethral orifice.7 The vulva consists of the vestibule, clitoris, and labia, and is located in the perineum ventral to the anus. In the nonestrous jill, the urogenital opening appears as a slit. During estrus, the vulva can swell considerably and resemble a doughnut. There are three to five pairs of nipples present in both jills and hobs.14,15
CARDIOVASCULAR AND LYMPHATIC SYSTEMS Heart and Blood Vessels The heart lies in a comparatively caudal location within the thoracic cavity, between the sixth and eighth ribs, with the apex to the left.15,18,20 The cardiodiaphragmatic ligament can contain varying amounts of fat.15A single brachiocephalic artery exits the aorta just proximal to the left subclavian artery. At the level of the thoracic inlet, this single artery divides into the right and left common carotid arteries and the right subclavian artery.7,18 This variation of the brachiocephalic artery is speculated to assist in maintaining cerebral blood flow during extreme rotation of the head and neck.32
Lymphatic Structures The thymus is located in the cranial mediastinum and can vary in size with age.7 Mediastinal lymph nodes are also present.15 The mediastinum is believed to be complete in ferrets.15 The
ferret possesses a robust system of lymph nodes and organs. The palatine tonsil is a flattened and ovoid structure that lies in the tonsillar fossa lateral to the ventral sulcus of the soft palate.7,15 The mandibular lymph node lies just rostral to the mandibular salivary gland and can easily be confused with this structure. The abdominal cavity has several prominent lymph nodes, including a prominent, palpable node at the root of the mesentery.27 The spleen sits in the left hypogastric region and parallels the greater curvature of the stomach.7 The reported normal size of the ferret spleen is 5.1 cm in length and 1.8 cm in width.7 However, benign extramedullary hematopoiesis is common in adult ferrets and results in a moderately to severely enlarged spleen.15 When enlarged, the spleen can extend diagonally from the upper left to the lower right abdomen, crossing the midline.
RESPIRATORY SYSTEM The lungs, contained within the slender and elongated thoracic cavity, are comparatively long in ferrets. The right lung is divided into cranial, middle, caudal, and accessory lobes. The left lung consists of a cranial and caudal lobe.7,14,15,18,20 Ferret lungs have a remarkably large filling capacity at about three times the predicted value for body size.14,18,30 Pinpoint yellow foci on the surface of the lungs observed at necropsy are foci of alveolar histiocytosis, the significance of which is unknown.14 The trachea is wide and very long, which results in comparatively lower airway resistance.14 The trachea bifurcates at the fifth intercostal space.15
ENDOCRINE SYSTEM Adrenal Glands The adrenal glands are located near each kidney, embedded in fat and covered by peritoneum.7 Each gland lies ventral to its ipsilateral adrenolumbar artery. The right gland lies in close apposition to the caudal vena cava and is draped by the caudate lobe of the liver. The right adrenal gland is slightly larger and longer than the left, at approximately 8 to 11 mm in length in one study.13 In another study, the adrenal gland length in females ranged from 5.0 to 10.0 mm for the left and 5.0 to 10.0 mm for the right; in males, it was 7.0 to 10.5 mm for the left and 7.5 to 13.5 for the right.26 Blood supply to either gland arises from the ipsilateral renal artery, with branches arising directly from the aorta, as well as the right adrenolumbar artery for the right adrenal gland.13 Variation to the adrenal blood supply exists. Accessory adrenal tissue can occasionally be found.7,15
Thyroid and Parathyroid Glands The thyroid gland is located ventrally along the neck between the third and eleventh tracheal rings, with each lobe positioned just lateral to the trachea.7,15 The parathyroid glands are small, pinkish structures that lie along the medial surface of the cranial portion of the thyroid, contacting the trachea at the fourth to fifth tracheal ring.7,15 The glands may be paired, occasionally single, on either side.7
MUSCULOSKELETAL SYSTEM The domestic ferret possesses a slender, elongated body, allowing it to fit through narrow spaces in pursuit of prey.7,29 The skeleton is lightweight but very flexible and strong.7,18 The skull is long and lacks sutures in the adult and the nasal openings are small compared to other mammals.7,20 Like all mustelids, ferrets
CHAPTER 1 Basic Anatomy, Physiology, and Husbandry possess strong muscles of mastication and can inflict a powerful and unrelenting bite.20 The vertebral formula of the ferret is C7, T15, L6 (5 or 7), S3, Cd18.7,15,18 The thorax is comparatively large with a narrow thoracic inlet and small first ribs.7,18,20 There are 15 pairs of ribs (occasionally 14), the first 10 of which are attached to the sternum and the last 5 comprising the costal arch.7,18 The spine is remarkably flexible, allowing the ferret to easily turn 180 degrees in a narrow passage. Despite possessing short legs, ferrets can climb remarkably well. Each foot possesses five digits with non-retractable claws. There are three phalangeal bones for each digit except the first digit, which has two.
NEUROLOGIC SYSTEM AND SPECIAL SENSES Brain and Spinal Cord The ferret brain has a typical mammalian design, and has been extensively reviewed elsewhere.17 The spinal cord and peripheral nerves of the ferret appear to be similar to those of the dog.15 The cauda equina begins at about the level of the last lumbar vertebra in the ferret.15
Special Senses Sight. The ferret eye has a prominent third eyelid, large cornea, horizontal elliptical pupillary opening, and a spherical lens.18,24 The eyes are open at 4 to 5 weeks after birth.18 Ocular movements are less pronounced in the ferret than in dogs and cats, and ferrets appear to track moving objects with movements of the head.15 Dorsal and ventral nasolacrimal puncta are present, although the dorsal punctum is smaller.15 The retina is similar to that of the dog and has a well-defined tapetum lucidum. The ferret is adapted to nocturnal living, and as such eyesight in the ferret is relatively poor compared to its olfactory and auditory senses.18 However, the ferret is a skilled hunter and can respond to movements of 25 to 45 cm/sec. Ferrets are believed to have limited color discrimination, at best.15,18,24 Hearing. The ferret pinna is set close to the head and points forward.7,18 The structure of the middle and inner ear is similar to that in the dog, although the ferret lacks a distinct tubular ear canal.18 Auditory function in the ferret is similar to the cat, although the auditory response may be more primitive.18 Although hearing range of ferrets is between 4 and 15 kHz (0.5 to 5 kHz in humans), lactating females can hear distress calls as high as 100 kHz from kits.15,18 Kits can hear by 32 days of age.18 Taste and Olfaction. Ferrets rely extensively on sense of smell.18 Ferrets have an elaborate nasal turbinate system like other carnivores, and appear to develop their olfactory preferences for food items during the first few months of life. These preferences may explain why diet changes in pet adult ferrets can be quite challenging.1 Taste sensation arises from the fungiform, vallate, and perhaps the foliate tongue papillae.15
PHYSIOLOGY AND REPRODUCTION PHYSIOLOGY Physiologic values of the domestic ferret are presented in Table 1-1.
BODY SIZE AND SEASONAL WEIGHT VARIATION Ferrets typically reach adult size by 6 months of age.7 Intact hobs weigh between 1 and 2 kg, in comparison to a weight between 0.6 and 1 kg for intact jills.18,23,29 If ferrets are neutered
9
before weaning, jills tend to become comparatively larger and hobs comparatively smaller, with a range of 0.8 to 1.2 kg for the two sexes. Neutered hobs lack the pronounced, muscular neck and shoulders characteristic of intact hobs. Ferrets tend to gain weight as winter approaches and lose weight in the spring.14 This seasonal weight fluctuation can be dramatic, with body weight variability approaching 40% in some individuals.10,15
REPRODUCTION Jills normally reach sexual maturity at 8 to 12 months of age, usually in the first spring after birth.10,14,22,29 Hobs typically reach puberty at about 9 months of age.29 The domestic ferret is seasonally polyestrus.14 Ferrets require alternating periods of long days and short days to have a normally functioning annual cycle.22 In both sexes, fertility increases as the days get longer. Spermatogenic activity in the hob occurs from December to July, and the testicles enlarge during this time. If not bred, intact jills will remain in persistent estrus from late March into early August.22,29 To an inexperienced observer, copulation appears violent, with the hob biting and dragging the jill by the neck.14,29 A receptive jill will remain limp and not fight back. For successful mating, it is suggested to wait until the jill has been in estrus for 10 days before placing with the hob. The jill and the hob can be left together for up to 48 hours or be bred for shorter periods on 2 consecutive days.22,29 Jills are induced ovulators, requiring neck restraint and intromission, and ovulation generally occurs 30 to 36 hours after copulation.14,22,29 Gestation length in the domestic ferret is 41 days on average, with a range of 39 to 42 days. If fertilization does not occur after ovulation is induced, a pseudopregnancy lasting 40 to 42 days may occur.14,15,22 If ovulation is not induced mechanically or chemically, the jill will remain in estrus until a changing photoperiod occurs. Prolonged estrus introduces the risk of severe anemia due to bone marrow suppression caused by persistent hyperestrogenism.18 Jills deliver an average of 8 kits, with a range of 1 to 18. Kits weigh 6 to 12 grams at birth and are born blind and deaf with a thin coat of white fur.10,14,22 Jills raise the kits alone. Kits begin eating soft food by 21 days of age, often before their eyes open. Kits are generally weaned by 6 to 8 weeks of age.14,18
HUSBANDRY The following discussion of husbandry is an overview of the keeping of ferrets as pets. A wealth of information is now available on all these topics, providing more details. The literature also contains ample information about maintaining ferrets as laboratory animals; thus this topic is not addressed here. Behavior of the domestic ferret is discussed in Chapter 39.
HOUSING Ferrets can be housed either indoors or outdoors depending on the climatic conditions of the area. Ferrets are intelligent, curious animals that should not be continuously confined in a small cage. Pets need a safe play area where they can investigate a variety of objects, such as boxes, bags, and plastic pipes. Ferrets should be allowed a minimum of 2 hours a day of exercise. Lewington21 has an extensive description of an entire “ferretarium” and other outside enclosures for ferrets that are rich in environmental stimuli. A play or living area for ferrets must first be “ferret proofed”— that is, all holes to the outside or to areas from which the ferrets
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SECTION I Ferrets
cannot be retrieved must be blocked. In addition, ferrets like to burrow into the soft foam rubber of furniture and mattresses. Owners should be advised to cover the bottom of all couches, chairs, and mattresses with a piece of thin wood, or a sturdy wire mesh. The burrowing is not only destructive but also potentially life-threatening because ferrets may swallow the foam rubber and develop gastrointestinal obstructive disease. Reclining chairs have been implicated in the crushing deaths of many ferrets and should be removed from the environment. In addition, all access to foam or latex rubber items, such as dog and cat toys, athletic shoes, rubber bands, stereo speakers, headphones, and pipe insulation, should be eliminated. Ferrets will often chew these materials and ingest them. Ingestion of rubber foreign bodies is the most frequent cause of gastrointestinal obstruction, particularly in ferrets younger than 1 year. Up to two ferrets can use a wire cage of 24 × 24 × 18 inches as a home base when it is necessary to confine them. The floor can be either solid or wire. Glass tanks are not suitable for caging ferrets because they provide poor ventilation. Custom-built wooden cages can also be used, but care must be taken to protect corners, the lower third of walls, and the floor from contamination with urine and feces. A moisture-proof material such as linoleum or plastic or vinyl molding can be used for this purpose. If ferrets are kept outdoors, a portion of the cage should be shaded for protection from extremes of heat and cold, and a well-insulated nest box should be provided. Ferrets do not tolerate temperatures above 90°F (32°C), especially in the presence of high humidity, and may need to be brought indoors. In climates where the temperature drops below freezing, particularly if the temperature falls below 20°F (−7°C), a heated shelter is necessary. When caring for ferrets in a clinical setting, ensure that cages are escape proof. Ferrets have been known to squeeze between the bars of a standard dog or cat hospital cage. Using a piece of Plexiglas over cage bars that are too wide can keep the patient secure. Ferrets need a dark, enclosed sleeping area. This is also essential in the clinical setting, because the patient may become more anxious and stressed if denied access to such a “safe” area. Towels, old shirts, and cloth hats can be used, in addition to specific products designed for ferrets to sleep in, such as cloth tubes and tents. For the occasional ferret that insists on eating its cloth sleeping material, use a small cardboard, plastic, or wooden box with an access hole cut into it. Some owners use slings, hammocks, or shelves that are built into the cage to provide additional sleep and play areas. In a multiple-ferret household, at least one sleep area should be provided per ferret to allow the choice to be alone and to reduce fighting. Ferrets can be trained to use a litter box relatively easily. Because ferrets like to back up in corners to defecate or urinate, the litter box sides should be high enough to contain the excreted material. Pelleted litter material is recommended instead of clay or clumping litter. Because of the ferret’s short digestive transit time, the pet may not always reach the cage to use the litter box if it is not close by. Therefore owners should be advised to have several litter boxes available in various rooms of the house for use by the pet when it is uncaged.
ENVIRONMENTAL ENRICHMENT Environmental enrichment is of vital importance to the health and mental well-being of any animal kept in captivity. A sterile environment is an inhumane environment. The goal of enrichment should be to encourage healthy, natural behaviors and to
minimize injurious unnatural behaviors or even lack of activity. Consider these five areas when planning enrichment strategies: dietary, occupational activity, physical environment, sensory, and social. Specifics of ferret nutrition are discussed in the next section, but method of delivery of the food and the variety of food can be an enrichment in itself. Feeding the same diet, day in and day out, always available in a clearly visible container does not allow natural hunting behaviors. It is beneficial to alter feeding times, put food in different places, hide the food, give food choices, use food as part of positive reinforcement training, and rotate through a variety of food textures (hard to soft) and tastes to simulate more natural feeding patterns. Occupational therapy would include activities that stimulate physical exercise and mental stimulation. There are countless possibilities for creativity. For instance, ferrets like to be in underground burrows, so providing a network of tunnels and tubes made out of PVC pipe or other sturdy material stimulates movement. Ferrets also enjoy climbing up ramps and cloth- or carpetcovered objects. Digging boxes are greatly enjoyed and allow burrowing. Boxes filled with clean dirt can be given on a limitedtime basis in an area that is easy to clean up such as a bathtub or a secure outdoor enclosure. Some ferrets like to play in water, so an occasional trip to the bathtub with an inch of water in the bottom can be a special treat. Ferrets love the movement of a “prey” animal such as cat toys on a string or mechanical toys that can be introduced for a period of time into the environment. Enrichment of the physical environment includes the type and size of the cage or play area, the cage furniture, and the toys. Secure, private sleeping areas are very important to ferrets. Provide several choices with different textures and sizes. Have at least one hiding area per ferret that has an opening of a size the ferret can “defend” from other ferrets. Other furniture may include the climbing and hiding areas described under occupational activity. Toys for ferrets should not include any latex rubber toys intended for dogs or cats because these can end up as a gastrointestinal obstruction when ferrets chew on them. Instead, paper bags, cloth toys for cats or babies, or cat or human infant toys made of hard plastic or metal can be used. Toys that move or make noise are especially appreciated. Sensory enrichment is a part of many of the strategies already discussed. Stimulate olfactory, visual, tactile, auditory, and taste experiences. Food, toys, and the cage environment should be rich in sensory experiences. Using different scents, such as food scents, or safe essential oil scents on toys can turn them into a new item. If the ferret is an indoor pet, if at all possible arrange regular forays outside in a safe environment to promote further mental stimulation and exposure to novel experiences. Social enrichment can be fulfilled with contact with other ferrets and with other species, including humans. Play both contact and noncontact games with the ferret. Consider engaging in positive reinforcement training, which not only strengthens the bond between pet and caregiver, but also empowers the ferret to make choices, to continue to learn, and to be mentally stimulated.
NUTRITION Ferrets are obligate carnivores designed to eat whole, small prey animals including small to medium-sized mammals, birds, eggs, frogs, crustaceans, fish, worms, and insects. The ferret’s polecat ancestors would bring their kill home and store the excess in the den and eat small frequent meals rather than gorging.2
CHAPTER 1 Basic Anatomy, Physiology, and Husbandry Ferrets have a very short gastrointestinal tract with minimal gut flora and few brush border enzymes, so they cannot use carbohydrates efficiently or digest fiber.2 Ferrets in nature would only encounter carbohydrates as found in the partially digested stomach contents of their prey. Like many other obligate carnivores, young ferrets imprint on food by smell at a very young age and develop strong food preferences by the time they are a few months old. Therefore, regardless of the diet strategy that is chosen, ferrets should be exposed to a variety of food tastes, textures, smells, and different protein sources as juveniles so their diet will have more flexibility as an adult. This can be extremely helpful when ferrets experience medical conditions that may require restricted or altered diets at an older age. Ferrets should be fed a diet high in animal fat for energy, high in good-quality meat (not plant) protein, with minimal carbohydrate and fiber. A whole-prey diet or a balanced fresh or freeze-dried carnivore diet is the most appropriate for a ferret, and such diets are currently fed in many areas of the world with great success. Clean sources of prey food such as chicks, mice, and rats are now available in many areas of the country thanks to the reptile market, which uses these foods for carnivorous pets. If an owner does not want to feed a 100% whole-prey or raw diet, consider the occasional “treat” of a whole mouse or chick as valuable environmental enrichment. The stools of a ferret on a whole-prey diet are very firm and of low volume and odor. The most common diet fed to pet ferrets in the United States is dry kibble. Although there have been advancements in dry ferret food formulation, these diets still contain grain, which is necessary to hold the food in its solid shape. Very high levels of plant proteins in the diet can lead to urolithiasis.2 The hardness of the kibble may promote excessive dental wear and disease.6 Furthermore, excess dietary carbohydrates may affect the pancreas and may contribute to disease of the beta cells. Unfortunately, ferrets seem to enjoy sweet foods, and some commercial pet food companies have capitalized on this preference by producing ferret treats that are little more than sugar-coated grains. These treat foods are particularly dangerous to the health of the pet ferret. The stools of a ferret eating a dry kibble diet are formed but are soft, voluminous, and may contain visible undigested grain. If a dry diet is fed to the ferret, the owner should read the diet ingredients carefully. The crude protein should be 30% to 35% and composed primarily of high-quality meat sources, not grains; the fat content should be 15% to 20%.32 Dry food ingredients are listed on the label in descending order of their amount in the product. The first three ingredients of a ferret diet should be meat products. Because the diet is dry, it can be left out at all times. However, this is not very mentally stimulating and the ferret may establish stashes of food around the house, mimicking the storage of extra prey in its ancestral den. It is preferable not to leave the dry food out all the time, but rather to offer it two or three times a day and increase mental stimulation by varying the times and locations of feeding. Growing kits need 35% protein and 20% fat, and lactating females require 20% fat and twice the calories of the nonpregnant ferret.2 Acceptable supplemental foods to a dry diet include fresh human food grade raw organ or muscle meat and raw egg. Cooking the meat or eggs may not be absolutely necessary if they are fresh and are suitable for human consumption; however, raw or undercooked meat and eggs introduce the risk of certain enteric pathogens such as Salmonella species, Campylobacter jejuni, and E. coli. Adding a small amount of high-quality canned cat food
11
can add variety in texture, taste, and protein sources. Omega-3 oils, fish oils, or meat fat can be added to increase the fat content of the diet provided these additions are not allowed to become rancid, and are not fed in excess of about 20% of the diet. Dairy products have also been used as a fat and protein supplement, but some ferrets develop soft stools when fed these products. Even though ferrets enjoy eating fruits, they should only be given occasionally in very small amounts. Owners often overfeed these items, leading to a reduction in the consumption of a healthier diet and the overfeeding of sugars and fiber. Perhaps one of the best strategies for feeding a ferret is to offer a variety of food items throughout the ferret’s life, including a minimum of weekly whole-prey foods, daily high-quality ferret kibble, and small amounts of high-quality canned cat food or other meat-based treats fed two to three times a week. This diet would cover many nutritional bases, increase the flexibility of the ferret’s diet preferences, and is mentally enriching. Because of the short gastrointestinal transit time, fasting a ferret for longer than 3 hours is not necessary to check the fasting blood glucose level or to empty the GI tract for a surgical procedure. Ferrets older than 2 years in the United States are prone to develop insulinoma, and a longer fast could result in a serious hypoglycemic condition. Water should always be available in either a sipper bottle or a heavy crock-type bowl. Ferrets love to play in the water, so the bowl should not be easy to overturn. Supplements should not be added to the ferrets’ water supply.
References 1. Apfelbach R. Olfactory sign stimulus for prey selection in polecats. Zeitschrift fur Tierpsychol. 1973;33:270-273. 2. Bell JA. Ferret nutrition. Vet Clin North Am Exot Anim Pract. 1999;2:169-192. 3. Berkovitz BKB. Supernumerary deciduous incisors and the order of eruption of the incisor teeth in the albino ferret. J Zool Lond. 1968;155:445-449. 4. Bleavins MR, Aulerich RJ. Feed consumption and food passage time in mink (Mustela vison) and European ferrets (Mustela putorius furo). Lab Anim Sci. 1981;31:268-269. 5. Church B. Ferret-polecat domestication: genetic, taxonomic and phylogenetic relationships. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. Philadelphia: WB Saunders; 2008:122-150. 6. Church RR. The impact of diet on the dentition of the domesticated ferret. Exot DVM. 2007;9:30-39. 7. Evans HE, An NQ. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:19-69. 8. Fox JG. Taxonomy, history, and use. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:3-18. 9. Fox JG. Normal clinical and biologic parameters. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:183-210. 10. Fox JG, Bell JA. Growth, reproduction, and breeding. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:211-227. 11. Gentry A, Clutton-Brock J, Groves CP. The naming of wild animal species and their domestic derivatives. J Archaeol Sci. 2004;31:645-651. 12. Grzimek B. Grzimek’s encyclopedia of mammals. Vol 3. New York: McGraw-Hill; 1990;388–449. 13. Holmes RL. The adrenal glands of the ferret, Mustela putorius. J Anat. 1961;95:325-336.
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14. Hrapkiewicz K, Medina L. Clinical laboratory animal medicine: an introduction. 3rd ed. Ames, Iowa: Blackwell Publishing; 2007. 15. Ivey E, Morrisey J. Ferrets: examination and preventive medicine. Vet Clin North Am Exot Anim Pract. 1999;2:471-494. 16. Jacob S, Poddar S. Morphology and histochemistry of the ferret prostate. Acta Anat. 1986;125:268-273. 17. Lawes INC, Andrews PLR. Neuroanatomy of the ferret brain. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:71-102. 18. Lewington JH. Ferrets. In: O’Malley B, ed. Clinical anatomy and physiology of exotic species: structure and function of mammals, birds, reptiles and amphibians. Philadelphia: WB Saunders; 2005:237-261. 19. Lewington JH. Classification, history and current status of ferrets. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. Philadelphia: WB Saunders; 2008:3-14. 20. Lewington JH. External features and anatomy profile. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. Philadelphia: WB Saunders; 2008:15-33. 21. Lewington JH. Accommodation. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. Philadelphia: WB Saunders; 2008:34-56. 22. Lindeberg H. Reproduction of the female ferret (Mustela putorius furo). Reprod Domest Anim. 2008;43(suppl 2):150-156. 23. MacDonald D. The velvet claw: a natural history of the carnivores. London: BBC Books; 1992.
24. Miller PE. Ferret ophthalmology. Semin Avian Exot Pet Med. 1997;6:146-151. 25. Moody KD, Bowman TA, Lang CM. Laboratory management of the ferret for biomedical research. Lab Anim Sci. 1985;35:272-279. 26. Neuwirth L, Collins B, Calderwood-Mays M, et al. Adrenal ultrasonography correlated with histopathology in ferrets. Vet Radiol Ultrasound. 1997;38:69-74. 27. Paul-Murphy J, O’Brien RT, Spaeth A, et al. Ultrasonography and fine needle aspirate cytology of the mesenteric lymph node in normal domestic ferrets (Mustela putorius furo). Vet Radiol Ultrasound. 1999;38:69-74. 28. Poddar S, Murgatroyd L. Morphological and histological study of the gastro-intestinal tract of the ferret. Acta Anat. 1976;96: 321-334. 29. Purcell K, Brown SA. Essentials of ferrets: a guide for practitioners. 2nd ed. Lakewood, Colorado: AAHA Press; 1999. 30. Vinegar A, Sinnett EE, Kosch PC, et al. Pulmonary physiology of the ferret and its potential as a model for inhalation toxicology. Lab Anim Sci. 1985;35:246-250. 31. Whary MT, Andrews PLR. Physiology of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:103-148. 32. Willis LS, Barrow MV. The ferret (Mustela putorius furo) as a laboratory animal. Lab Anim Sci. 1971;21:712-716. 33. Zeuner FE. A history of domesticated animals. New York: Harper & Row; 1963.
CHAPTER
2
Basic Approach to Veterinary Care
Katherine E. Quesenberry, DVM, MPH, Diplomate ABVP (Avian), and Connie Orcutt, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal)
Restraint and Physical Examination Restraint Physical Examination Preventive Medicine Vaccinations Parasites Hospitalization Clinical and Treatment Techniques Venipuncture Intravenous Catheters Fluid Therapy Antibiotic and Drug Therapy Pain Management Nutritional Support Urine Collection and Urinalysis Urinary Catheterization Splenic Aspiration Bone Marrow Collection Tracheal Wash Blood Transfusion Blood Pressure Monitoring Diagnostic Peritoneal Lavage
Ferrets are commonly seen in many small animal veterinary practices. Special equipment needs are minimal, and the approach to handling ferrets is similar in many ways to that for dogs and cats. Ferret owners regularly seek veterinary care for a variety of reasons: ferrets need preventive vaccinations for canine distemper and rabies; ferrets have a relatively short life span compared with that of cats and dogs; ferrets in the United States and in some European countries have a high incidence of endocrine, gastrointestinal, and neoplastic diseases; and many of the diseases common to ferrets are not easily ignored by the pet owner (e.g., alopecia resulting from adrenal disease and hypoglycemic episodes caused by insulinoma). Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
RESTRAINT AND PHYSICAL EXAMINATION RESTRAINT Most ferrets are docile and can be easily examined without assistance. However, an assistant is usually needed when taking the rectal temperature, when administering injections or oral medications, or if an animal has a tendency to bite. Young ferrets often nip, and nursing females and ferrets that are handled infrequently may bite. Unlike dogs and cats, which growl, ferrets will bite without warning. Therefore always ask the owner if the ferret will bite before handling it and take precautions accordingly. Make sure to obtain the rabies vaccination history before physical examination, as reporting and rabies protocols for animal bites from vaccinated and unvaccinated ferrets differ (see below). Ferrets that are prone to bite and are not currently vaccinated for rabies may need tranquilization for procedures that require restraint. Depending on the ferret’s disposition, several basic manual restraint methods can be used for physical examination. For tractable animals, lightly restrain the ferret on the examination table. Examine the mucous membranes, oral cavity, head, and integument. Then pick the ferret up and use one hand for support under its body while using the second hand to auscultate the thorax and palpate the abdomen. The ferret can be scruffed at any time for vaccination, ear cleaning, or other procedures that may elicit an attempt to escape or bite. For a very active animal or one that bites, scruff the ferret at the back of its neck and suspend it with all four legs off the table (Fig. 2-1). Most ferrets become very relaxed with this hold, and the veterinarian is able to examine the oral cavity, head, and body; auscultate the chest; and palpate the abdomen easily. However, this method may not work for very fractious animals. To manually restrain a ferret for procedures such as venipuncture or ultrasound, hold it firmly by the scruff of its neck and around the hips without pulling the legs back. Most ferrets struggle if their legs are extended by pulling on the feet. Some animals can be distracted during a procedure by feeding a meat-based canned food (a/d Prescription Diet, Hill’s Pet Nutrition, Topeka, KS; Eukanuba Maximum-Calorie, The Iams 13
14
SECTION I Ferrets
Fig. 2-1 Restrain an active ferret by scruffing the loose skin on the back of the neck. The ferret will relax and allow you to palpate the abdomen or administer a vaccine.
Company, Dayton, OH) or a small amount of a supplement such as FerreTone (8-in-1 Pet Products, Islandia, NY) by syringe. Avoid products containing sugar, which can affect blood glucose values, particularly in ferrets with insulinoma. For very fractious or anxious animals or for procedures requiring lengthy restraint, light tranquilization or sedation may be indicated (see Chapter 31).
PHYSICAL EXAMINATION Most ferrets strenuously object to having their temperature taken with a rectal thermometer. If a ferret struggles during the examination, the temperature taken at the end of the examination may be artificially high. Therefore measure the rectal temperature early in the physical examination with a flexible digital thermometer that is well lubricated. The normal range of rectal temperature of a ferret is 100.5°F to 102.5°F (38.0°C to 39.2°C); a mean of 102°F (38.8°C), with a wider range of 100°F to 104°F (37.8°C to 40.0°C), is also reported.21 Interestingly, in normal ferrets housed outdoors at a fur farm in very cold ambient temperatures (21°F [−6.1°C]), the mean 24-hour mean core body temperature measured by sterile thermosensitive data loggers implanted in their abdomens was 99.3°F (37.4°C), with a range of 97.3°F to 101.1°F (36.3°C to 38.4°C).41 The physical examination of a ferret is basically the same as that of any small mammal and can be performed quickly and efficiently if a few simple guidelines are followed. Observe the attitude and alertness of the animal. Ferrets may sleep in the carrier in the veterinary office; however, once awakened for the examination, a ferret should be alert and responsive. Assess hydration by observing the skin turgor of the eyelids, tenting of the skin at the back of the neck, and moistness of the oral mucous membranes. However, skin turgor can be difficult to evaluate in a cachectic animal. Estimate the capillary refill time by digitally pressing on the gingiva.
Examine the eyes, nose, ears, and facial symmetry. Cataracts can develop in both juvenile and adult animals. Retinal degeneration is another ophthalmic disorder seen in ferrets and may be indicated by abnormal pupil dilation. Inspect for nasal discharge and ask the owner about any history of sneezing or coughing. The ears may have a brown waxy discharge, but the presence of excessive brown exudate may indicate infestation with ear mites (Otodectes cynotis). Bruxism often indicates gastrointestinal discomfort. The teeth of ferrets should be clean and the gingiva pink. Dental tartar is commonly present in pet ferrets. The amount of plaque may be exacerbated by the feeding of soft foods or sugary treats, such as raisins, and is possibly related to a dry kibble diet.14 Tartar most commonly accumulates on the first and second maxillary premolars. Excessive dental tartar should be removed by dental techniques used in dogs and cats, and measures to prevent tartar buildup should be implemented. As a preventive, a pet dentifrice or tartar control toothpaste25,32 can be applied to the teeth to decrease formation of calculus. Gingivitis, which manifests as erythematous gingival tissue that sometimes bleeds, is a common sequela of excessive dental tartar. Ferrets often break off the tip of one or both canine teeth; however, they rarely exhibit clinical signs of sensitivity or pain associated with a fractured canine. If the tooth turns dark or the ferret exhibits sensitivity when eating, recommend a root canal or extraction, depending on the degree of damage to the tooth (see Chapter 32). Rarely, an infected root of a fractured canine can cause swelling of the ipsilateral submandibular lymph node. If swelling is present, dental radiographs, canine tooth extraction, and possibly lymph node biopsy are indicated. Observe the symmetry of the face. Although uncommon, salivary mucoceles occur in ferrets and are noticeable as a unilateral swelling on the side of the face, usually in the cheek or temporal area (see Chapter 3). Palpate the regional lymph nodes of the neck and axillary, popliteal, and inguinal areas. Nodes should be soft and may sometimes feel enlarged in large or overweight animals because of surrounding fat. Any degree of firmness or asymmetry in one or more nodes is suspicious and warrants a fine-needle aspirate or a biopsy. If two or more nodes are enlarged and firm, a full diagnostic workup is indicated. Auscultate the heart and lungs in a quiet room. Ferrets have a rapid heart rate (180 to 250 beats/min) and often a pronounced sinus arrhythmia. If a ferret is excited and has a very rapid heart rate, subtle murmurs may be missed. Valvular disease, cardiomyopathy, and congestive heart failure are seen in ferrets, and any murmur or abnormal heart rhythm should be investigated further (see Chapter 5). Palpate the abdomen while holding the ferret off the table, either by scruffing the neck or supporting the ferret with one hand. This allows the abdominal organs to displace downward, facilitating palpation. If the history is consistent with an intestinal foreign body or urinary blockage, palpate gently to avoid causing iatrogenic injury, such as a ruptured stomach or bladder. Palpate the cranial abdomen, paying particular attention to the presence of gas or any irregularly shaped mass in the stomach area, especially in ferrets with a history of vomiting, melena, or chronic weight loss. The spleen is commonly enlarged in ferrets; this may or may not be significant, depending on other clinical findings (see Chapter 5). Palpate a large spleen gently to avoid iatrogenic damage. A very enlarged spleen may indicate systemic disease or, very rarely, idiopathic hypersplenism, and
CHAPTER 2 Basic Approach to Veterinary Care further diagnostic workup is warranted. Always note any degree of splenic enlargement in the medical record so that this finding can be rechecked at future examinations. Examine the genital area, observing the size of the vulva in females. Vulvar enlargement in a spayed female is consistent with either adrenal disease or an ovarian remnant; the latter is rare. If the vulva is of normal size, show this to the owner so that any vulvar enlargement in the future will be noticed. Examine the preputial area and size of the testicles of male ferrets; preputial and testicular tumors are sometimes seen. Check the fur coat for evidence of alopecia. Alopecia of the tail tip is common in ferrets and may be incidental and transient or an early sign of adrenal disease. Symmetric, bilateral alopecia or thinning of the hair coat that begins at the tail base and progresses cranially is a common clinical finding in ferrets with adrenal disease. Examine the skin on the back and neck for evidence of scratching or alopecia. Pruritus may be present with adrenal disease (common) or with ectoparasites (fleas, Sarcoptes scabiei). Check closely visually and by searching through the hair coat with your fingers for evidence of skin masses. Mast cell tumors are common and can range in diameter from a few millimeters to over a centimeter. Often, the fur around a mast cell tumor is parted and matted with dark blood from the animal’s scratching. Other types of skin tumors, such as sebaceous adenomas and basal cell tumors, are also common (see Chapter 9). Perform an excisional biopsy of any lump found on the skin.
PREVENTIVE MEDICINE Young, recently purchased ferrets need serial distemper vaccinations until they are 13 to 14 weeks of age.2 Rabies vaccines should be given annually beginning at 3 months of age.15 Ferrets should be examined annually until they are 4 or 5 years of age; middle-aged and older animals should be examined twice yearly because of the high incidence of metabolic disease and neoplasia. Annual blood tests (consisting of a complete blood count and plasma or serum biochemical analysis) are recommended for older animals. Measure the blood glucose concentration twice yearly in healthy middle-aged and older ferrets; more frequent monitoring is needed in ferrets with insulinoma. An endocrine panel is indicated in ferrets with hair loss on the tail or other clinical signs suggestive of early adrenal disease (see Chapter 7). Testing for infectious diseases may be warranted, especially in new ferrets that will be introduced into a multi-ferret household or those that are taken to ferret shows. Currently, ferrets can be tested for Aleutian disease virus and ferret enteric coronavirus by polymerase chain reaction (PCR) testing (Michigan State University, Diagnostic Center for Population and Animal Health, www.animalhealth.msu.edu; Veterinary Molecular Diagnostics, www.vmdlabs.com). Serologic tests for Aleutian disease by enzyme-linked immunosorbent assay (ELISA) and counterimmunoelectrophoresis (CIEP) are also available (see Chapter 5).
VACCINATIONS Canine Distemper Ferrets must be vaccinated against canine distemper virus. Currently, one vaccine is approved by the U.S. Department of Agriculture for use in ferrets: PureVax (Merial, Athens, GA). Because PureVax is a canarypox-vectored recombinant vaccine
15
it does not contain adjuvants or the complete distemper virus; thus many of the postvaccination risks have been reduced. This product has a wide safety margin and has proved effective in protecting ferrets against canine distemper infection.58 Another distemper vaccine that was widely used previously (Fervac-D, United Vaccines, Inc., Madison, WI) is no longer available. Fervac-D was a modified live virus vaccine propagated in avian cell lines. Another modified live canine distemper vaccine (Galaxy D, Merck/Schering-Plough Animal Health, Whitehouse Station, NJ) has been studied for safety and efficacy in ferrets. This product, derived from the Onderstepoort distemper strain and attenuated in a primate cell line, proved effective in preventing canine distemper in young ferrets challenged after serial vaccination.64 However, duration of immunity with this product is not known, and its use in clinical animals is extralabel, requiring informed owner consent. Although no vaccine reactions were reported in the study, the incidence of vaccine reactions with Galaxy D is unknown because experience with repeated longterm use in ferrets has been limited.64 Because of the possibility of vaccine-induced disease, especially in immunosuppressed or sick ferrets, do not use combination canine vaccines or vaccines of ferret cell or low-passage canine cell origin. In young ferrets, the half-life of maternal antibody to canine distemper virus is 9.43 days.2 Vaccinate young ferrets for distemper at 8 weeks of age, then give two additional boosters at 3-week intervals for a total of three vaccinations. Give booster vaccines annually.
Rabies All ferrets should be vaccinated against rabies.15 A killed rabies vaccine is approved for use in ferrets (Imrab-3 or Imrab-3 TF, Merial, Duluth, GA) and is effective in producing immunity for at least 1 year.55 Current recommendations are to vaccinate healthy ferrets at 3 months of age at a dose of 1 mL administered subcutaneously. Give booster vaccinations annually. Titers develop within 30 days of rabies vaccination.55 In ferrets that were experimentally inoculated intramuscularly with skunk-origin rabies virus, the mean incubation period was 33 days and the mean morbidity period was 4 to 5 days.42 Clinical signs were ascending paralysis, ataxia, cachexia, bladder atony, fever, hyperactivity, tremors, and paresthesia. Virus antigen was present in the brain tissue of all ferrets with clinical signs of rabies, and virus was isolated from the salivary gland of one ferret. In a similar study of ferrets inoculated with a raccoon rabies isolate, the mean incubation period was 28 days. Virus was isolated from the salivary glands of 63% of rabid ferrets, and 47% shed virus in saliva. Virus excretion began from 2 days before until 6 days after the onset of illness.43 In an earlier study of ferrets with experimentally induced rabies, only mild clinical signs were observed before death.7 Infected ferrets exhibited restlessness and apathy, and some showed paresis. Sick animals did not attempt to bite when threatened, and virus was not excreted in the submaxillary salivary glands of animals that died. In this study, the authors concluded that ferrets are 50,000 times less susceptible to rabies than fox and 300 times less susceptible than hares. In another study, ferrets that were fed up to 25 carcasses of mice infected with rabies did not develop the disease; in contrast, skunks become fatally infected after the consumption of only one carcass.4 Ferrets are considered currently immunized 28 days after the initial rabies vaccination and immediately after a booster vaccination.15 If a healthy pet ferret bites a person, current
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SECTION I Ferrets
recommendations of the Compendium of Animal Rabies Prevention and Control are to confine and observe the animal for 10 days; the ferret should not be vaccinated during this period.15 If signs of illness develop, this should be reported to the local health department and a veterinarian should evaluate the animal. If signs suggest rabies, the ferret must be euthanized and protocols for rabies evaluation should be followed. If a stray ferret bites a person, the ferret must be euthanized and submitted immediately for rabies testing. For a vaccinated ferret exposed to a possible rabid animal, recommendations are to revaccinate the ferret immediately and quarantine for 45 days. An unvaccinated animal that is exposed to a rabid animal should be euthanized immediately and submitted for rabies testing. See the website of the Centers for Disease Control and Prevention (www.cdc. gov/mmwr/preview/mmwrhtml/rr5702a1.htm) or the National Association of Public Health Veterinarians (www.nasphv.org/) for specific guidelines.
Vaccine-Associated Adverse Events In ferrets, adverse events associated with vaccination are primarily type I hypersensitivity reactions or anaphylaxis.37 Type I hypersensitivity reactions involve lymphoid tissue associated with mucosal surfaces (skin, intestines, and lungs) and result from the interaction of antigen and immunoglobulin E in mast cells or basophils. Ferrets with mild reactions may exhibit pruritus and skin erythema. More severe reactions are typified by vomiting, diarrhea, piloerection, hyperthermia, cardiovascular collapse, or death. Vaccine reactions are most common after distemper vaccination but may also occur after rabies vaccination. In a study of vaccine reactions in 3,857 ferrets, the incidence of adverse events associated with rabies vaccine alone, distemper vaccine alone, and rabies and distemper vaccines together were 0.51%, 1.0%, and 0.85%, respectively. The incidence of adverse events did not differ significantly among these three groups; however, the cumulative number of distemper vaccinations received was significantly associated with the occurrence of an adverse event. The distemper vaccines used in this population of ferrets were PureVax and Fervac D; however, the two distemper vaccines were grouped collectively in the analysis, and the incidence of adverse events associated with the individual distemper vaccines was not reported. Sex, age, and body weight were not associated with occurrence of an adverse event. All reactions occurred immediately after vaccination and most commonly consisted of vomiting and diarrhea. In another study of 143 ferrets, the incidence of adverse events after administering distemper (5.9%) (Fervac D), rabies (5.6%) (Imrab-3), or both vaccines (5.6%) was not significantly different. In a 2001 report of vaccine reactions in ferrets reported to the United States Pharmacopeia Veterinary Practitioners’ Reporting Program, 65% (54 of 83) of reports involved administration of FerVac D; 24% (20 of 83) involved concomitant administration of FerVac D and Imrab; and 11% (9 of 83) involved administration of Imrab alone (PureVax was not approved for use at the time these data were collected).37 According to Merial’s product information, the incidence of vaccine reactions with PureVax is 0.3%. No data are available for products not licensed for use in ferrets. Veterinarians are not required to report vaccine-associated adverse events, and surveillance of these events is passive, relying on voluntary reporting by practitioners.37 Vaccine-associated adverse events can be reported to the Center for Biologics, U.S. Department of Agriculture (1-800-752-6255; www.aphis. usda.gov/animal_health/vet_biologics/vb_adverse_event.shtml).
Always follow the manufacturer’s instructions for vaccine administration and inform the owner of the possibility of a reaction before vaccinating. Have the owner monitor the ferret in the waiting area for 30 minutes or more after vaccination with any product. As stated, most reactions occur almost immediately after vaccination. If a ferret has an adverse reaction, administer an antihistamine (e.g., diphenhydramine hydrochloride [Benadryl, ParkeDavis, Morris Plains, NJ], 0.5 to 2.0 mg/kg intravenously [IV] or intramuscularly [IM]), epinephrine (20 μg/kg IV, IM, subcutaneously [SC], or intratracheally), or a short-acting corticosteroid (e.g., dexamethasone sodium phosphate, 1 to 2 mg/kg IV or IM), and give supportive care. For any biologic product, veterinarians must assess risk versus benefit of vaccination. The treatment options for ferrets that have had a vaccine reaction include not vaccinating if the risk of exposure is minimal; administering diphenhydramine (2 mg/kg orally [PO] or SC) at least 15 minutes before vaccination; or, for distemper, administering a different product. Vaccine injection-site sarcomas have been described in ferrets.39,40 In one report, 7 of 10 fibrosarcomas in ferrets were from locations used for vaccination.39 Fibrosarcomas from injection sites had a high degree of cellular pleomorphism and similar histologic, immunohistochemical, and ultrastructural features as those reported for feline vaccine-associated sarcomas. In the reported cases in ferrets, no definitive association could be made between the fibrosarcoma and the type of vaccine. In cats, adjuvanted vaccines are most likely to be involved in tumor development. However, although injection-site sarcomas may occur in ferrets, ferrets appear less prone than cats to tumor development. In a study of vaccine reactions in ferrets, mink, and cats, cats had more lymphocytes at the injection site than either ferrets or mink after vaccination with three different rabies vaccines.11 Results of this study suggest a lower species susceptibility to vaccine-associated sarcomas in ferrets than in cats.
PARASITES Endoparasites Gastrointestinal parasitism is uncommon in domestic ferrets. Rarely, pet ferrets may become infected with parasites from other natural hosts through intermediate hosts or vectors. Protozoan parasites are occasionally seen. Therefore perform routine fecal flotations and direct fecal smears for all young ferrets at the initial examination. Coccidiosis (Isospora species) is seen infrequently, usually in young ferrets, which shed oocysts between 6 and 16 weeks of age.3 The infection is usually subclinical; occasionally, however, ferrets may have loose stool or bloody diarrhea. Treatment of ferrets with coccidiosis is similar to that of other small animals and should be continued for at least 2 weeks. Coccidiostats, such as sulfadimethoxine and amprolium, are effective and safe. The Isospora species that infect ferrets may cross-infect dogs and cats; therefore other pets in the household should be checked for coccidia and treated as needed. Giardiasis is occasionally seen in ferrets. Results of a recent study on molecular characterization of Giardia duodenalis isolates from pet ferrets show that genetic sequences from isolates in ferrets differ from isolates of humans and other animals, suggesting that Giardia isolates from ferrets may be host specific.1 Giardia species can be detected by identifying cysts or
CHAPTER 2 Basic Approach to Veterinary Care trophozoites in a fresh fecal smear or zinc sulfate flotation, or by fecal ELISA. Treat ferrets with giardiasis with metronidazole (20 mg/kg PO q12h) for 5 to 10 days. Fenbendazole (50 mg/kg PO q24h for 3 to 5 days) is used in dogs and cats, but safety and efficacy in ferrets are unknown. Cryptosporidiosis can occur in a high percentage of young ferrets.53 Infection is usually subclinical in both immunocompetent and immunosuppressed animals. Although most immunocompetent animals recover from infection within 2 to 3 weeks, infection can persist for months in immunosuppressed animals. Oocysts of Cryptosporidium are small (3 to 5 μm) and difficult to detect but can be found in samples of fresh feces examined immediately after acid-fast staining.3,53 No treatments exist for Cryptosporidium infection. Because of the zoonotic potential, ferrets may be a source of infection for human beings, especially immunocompromised individuals with acquired immunodeficiency syndrome (AIDS).53 Heartworms (Dirofilaria immitis) can cause disease in ferrets. Ferrets that are housed outdoors in heartworm-endemic areas are most susceptible to infection; however, all ferrets in endemic areas should be given preventive medicine. Oral administration of ivermectin is currently the most practical preventive measure because it is administered once per month (see Chapter 5 and Appendix).
Ectoparasites Ear mites (Otodectes cynotis) are common in ferrets, but affected animals rarely exhibit pruritus or irritation. This mite species also infects dogs and cats, and animals in households with multiple pets can transmit mites to other animals. A red-brown, thick, waxy discharge in the ear canal and pinna characterizes infection. A direct smear of the exudate reveals adult mites or eggs. Because ferrets normally have brown ear wax, the color or appearance of debris in the ear canal is not pathognomonic for mites. At the initial examination, check all ferrets for ear mites and do follow-up checks at the annual examination in ferrets kept in multiple-pet households. Several products, including selamectin, are effective in treatment (see Chapter 9). Flea infestation (Ctenocephalides species) is most common in ferrets kept in households with dogs or cats. Ferrets with chronic infestation can become severely anemic. Check all ferrets during the physical examination for signs of fleas or flea dirt. Treat infested animals with products safe for use in cats and institute flea control measures (see Chapter 9). Ticks are rarely seen in domestic ferrets, and Lyme disease in ferrets has not been reported.
HOSPITALIZATION Ferrets can be hospitalized in standard stainless steel hospital cages with some adaptations. Ferrets are agile escape artists and can squeeze through even very small openings. In many standard cages designed for veterinary hospitals, the bar spacing is too wide, allowing an easy avenue of escape. For housing ferrets, use only cages with small spacing between vertical bars or use cages with small crossbars. Alternatively, adapt standard cages for use by attaching a Plexiglas plate to the front of the cage at least half the height of the cage door or higher. The plate will prevent escape through the bars yet can be easily detached and cleaned. Commercial hospital cages with Plexiglas fronts and access ports can be used for ferrets. There is no avenue of escape, and
17
ferrets are visible at all times. Acrylic or laminate animal intensive care cages or incubators also can be used to house ferrets and are especially useful for animals that need supplemental heat or oxygen. The cage should be large enough to accommodate a sleeping area or box and an area for defecation and urination. Ferrets are very careful about not soiling their sleeping area, even when very sick. All ferrets like to burrow and should be given opportunity to do so while hospitalized. Clean towels make excellent burrowing material. Alternatively, a mound of shredded paper provides much satisfaction to hospitalized animals. If not provided with burrowing material, many ferrets will burrow underneath the cage paper. Extra-small padded pet beds and fleece pet “pockets” work well as sleeping areas. An oxygen cage should be available for use with dyspneic animals. Monitor the temperature in commercial oxygen cages closely, because ferrets can become hypothermic quickly at cool cage temperatures that are used for dogs and cats. Conversely, ferrets can overheat at temperatures used for avian patients. Provide water for hospitalized ferrets in either water bottles or small weighted bowls. Ask the owner which type of watering system the ferret is accustomed to before hospitalization. Ferrets can be finicky eaters and should be fed their regular diet while hospitalized, if possible. Otherwise, feed a very palatable ferret food or a premium-quality, high-protein cat or kitten chow. If dietary changes are needed in the regular diet, recommend that changes be made gradually after the ferret has been released from the hospital. For animals that are anorexic, force-feed a high-calorie semisolid food or supplement until the animal is eating on its own (see later discussion).
CLINICAL AND TREATMENT TECHNIQUES VENIPUNCTURE Obtaining a blood sample from a ferret is relatively easy and usually does not require anesthesia. Several venipuncture sites are readily accessible; the technique and site chosen depend on how much blood is needed and the availability of assistants for restraint. Anesthesia or tranquilization can be used if assistants are unavailable, but anesthesia may affect hematologic values (see later discussion).35 Ferrets often can be distracted during restraint for venipuncture by offering semi-solid food or a product such as FerreTone (8-in-1 Pet Products) by syringe. Avoid using supplements with corn syrup or other sugars, as this will affect blood glucose levels, and collect blood for glucose determination or other fasting samples before offering food. Most veterinary laboratories offer small mammal hematologic and biochemical panels that can be done with 1.0 mL or less of blood. In-clinic point-of-care analyzers require very small sample sizes (usually 100 μL). The blood volume of healthy ferrets is approximately 40 mL in average-sized females weighing 750 g and 60 mL in males weighing 1 kg.21 Up to 10% of the blood volume can be safely withdrawn at one time in a normal ferret, but collect only the minimum amount needed for analysis. Repeated blood drawing can contribute to anemia in sick animals hospitalized for long periods. Two sites are commonly accessed to obtain large blood volumes in ferrets. The jugular vein can be approached in the neck by using the conventional technique used in cats, with the forelegs extended over the edge of a table and the neck extended up (Fig. 2-2). Use a 25-gauge needle with a 1- to 3-mL syringe for
18
SECTION I Ferrets
Fig. 2-2 Restraint for jugular venipuncture in a ferret. Restrain the ferret similar to a cat, with the legs pulled down and the head back. Shave the neck to improve visibility of the jugular vein in the lateral neck. After the vein is punctured, the head can be “pumped” up and down slowly to facilitate blood flow.
venipuncture in most ferrets; a 22-gauge needle can be used in large males. Shave the neck at the venipuncture site to enhance visibility of the jugular vein. The vein is relatively superficial and is located more lateral in the neck than it is in dogs or cats, and it is sometimes difficult to locate in heavy males. Once the needle is inserted, the blood should flow easily into the syringe; if the neck is overextended and the head is arched back, the blood may not flow readily from the vein. Relax the hold on the head or gently “pump” the vein by moving the head slowly up and down to enhance blood flow into the syringe. With ferrets that resist limb extension, a towel-wrap technique can be used.9 Scruff the ferret with its front legs extended caudally against the ventral thorax, and wrap the animal’s body firmly with a towel from the base of the neck down. An assistant is needed to restrain the toweled ferret in dorsal recumbency while scruffing the cranial neck. Apply pressure lateral to the thoracic inlet to visualize or palpate the jugular vein lying between the thoracic inlet and the base of the ear. However, with very fractious animals, even this technique may be difficult without tranquilization. The second venipuncture site to obtain large blood samples is the cranial vena cava. The actual site of venipuncture has been called the thoracic portion of the jugular vein12,60; however, anatomically it is more likely the right or left brachiocephalic trunk or the anterior vena cava itself, depending on the point of entry and the depth of needle penetration (Fig. 2-3, A). This technique is safe in ferrets because of the long anterior vena cava and the caudal location of the heart in the thoracic cavity, which is approximately 3 cm from the thoracic inlet. However, rare instances of hemorrhage into the anterior thoracic cavity can occur. Restrain the ferret on its back with the forelegs pulled caudally and the head and neck extended (Fig. 2-3, B). In an unanesthetized ferret, two assistants are usually needed, one for restraint of the forelegs and head and the other for restraint
of the rear just cranial to the pelvis. Insert a 25-gauge needle with an attached 1-mL or 3-mL syringe into the thoracic cavity between the first rib and the manubrium at an angle 30 to 45 degrees to the body. Direct the needle toward the opposite rear leg or most caudal rib and insert it almost to the hub. Pull back on the plunger as the needle is slowly withdrawn until blood begins to fill the syringe. If the ferret struggles, quickly withdraw the needle and wait until the ferret is quiet before making a second attempt. In very fractious or active ferrets, jugular venipuncture or use of tranquilization are safer choices to avoid lacerating the vessels. The lateral saphenous or cephalic vein can be used if only a small amount of blood is needed to measure a packed cell volume or blood glucose level. To prevent collapse of the vein during venipuncture, use an insulin syringe with an attached 27- or 28-gauge needle. The saphenous vein lies just proximal to the hock joint on the lateral surface of the leg; the cephalic vein is in the same anatomic location as in a dog. Before venipuncture, shave the fur from the area to enhance visibility of the vein. Although rarely used in pet ferrets, venipuncture of the tail artery is described to obtain blood samples.8 Venipuncture at this site can be painful; anesthetize the ferret for this technique. The artery is located 2 to 3 mm deep to the skin. Insert a syringe with a 21-gauge needle into the ventral midline of the tail directed toward the body. Once the artery is entered, slowly withdraw the plunger until blood fills the syringe. Apply pressure to the venipuncture site for 2 to 3 minutes after the needle has been withdrawn.
Reference Ranges Published reference intervals for hematologic, biochemical, and plasma electrophoresis values in ferrets are listed in Tables 2-1, 2-2, and 2-3. Other published sources of reference intervals for ferrets are available.21,22 Additionally, most clinical veterinary laboratories routinely provide reference intervals for ferret hematologic and biochemical values. Published reference intervals for white blood cell (WBC) counts in ferrets vary from 2.5 to 19.1 × 103 cells/μL21,27,61; however, WBC counts generally tend to be low in ferrets. In one study, mean WBC values were 5.7 and 5.6 × 103 cells/μL in male and female ferrets, respectively.22 High WBC counts are not seen as commonly in ferrets as in dogs and cats, perhaps in part because infectious bacterial diseases are comparatively uncommon in ferrets. Isoflurane anesthesia can cause decreases in all hematologic values beginning at induction of anesthesia and reaching maximal effects at 15 minutes after induction.35 Therefore the complete blood count values of blood samples collected while a ferret is anesthetized must be carefully interpreted. Reference intervals for blood coagulation times in ferrets have been published. In a recent study, blood samples were collected into sodium citrate in a ratio of 9:1 from 18 clinically healthy ferrets (12 males, 6 females, all neutered).6 Results showed some variation in values obtained by the method used for measurement. Mean prothombin time (PT) was 12.3 seconds (range, 11.6 to 12.7 seconds) measured by fibrometer and 10.9 seconds (range, 10.6 to 11.6 seconds) measured by an automated coagulation analyzer. Mean activated partial thromboplastin time (aPTT) was 18.7 seconds (range, 17.5 to 21.1 seconds) by fibrometer and 18.1 seconds (range, 16.5 to 20.5 seconds) by automated coagulation analyzer. Mean fibrinogen concentration was 107.4 mg/dL (range,
CHAPTER 2 Basic Approach to Veterinary Care
19
JV LBT RBT
AVC
A
B Fig. 2-3 A, Dissection of the thoracic cavity of a ferret illustrating the site for blood collection using the anterior vena cava technique. The sternum and ventral ribs are removed. The site of venipuncture is either the right brachiocephalic trunk (RBT) or left brachiocephalic trunk (LBT) or the anterior vena cava (AVC), depending on the point of entry and depth of penetration (see marker). The jugular vein (JV) is usually lateral and cranial to the venipuncture site. The base of the first two ribs are shown by arrows. B, A ferret is restrained for venipuncture of the anterior vena cava. Both forelegs are pulled back, hindlegs are restrained, and the neck is extended.
Table 2-1 Reference Intervals for Hematologic Values in Ferrets ALBINO37
FITCH16
Value
Combined Sexa
Maleb
Female
Malec
Female
Hematocrit (%) Hemoglobin (g/dL) Red blood cells (×106/μL) Reticulocytes (%) White blood cells (×103/μL) Neutrophils (%) (cells/μL) Lymphocytes (%) (cells/μL) Monocytes (%) (cells/μL) Eosinophils (%) (cells/μL) Basophils (%) (cells/μL) Bands (cells/μL) Platelets (×103/μL) Mean corpuscular volume (fL) Mean corpuscular hemoglobin (g/dL) Mean corpuscular hemoglobin concentration (g/dL)
36-48 12.2-16.5 7.01-9.65 4.3-10.7 18-47 41-73 0-4 0-4 0-2 200-459 50-54 15-18 32-35
44-61 16.3-18.2 7.30-12.18 1-12 4.4-19.1 11-82 12-54 0-9 0-7 0-2 297-730 -
42-55 14.8-17.4 6.77-9.76 2-14 4.0-18.2 43-84 12-50 2-8 0-5 0-1 310-910 -
46-57 15.2-17.7 5.6-10.8 616-7020 1728-4704 0-432 112-768 0-112 0-972 -
47-51 15.2-17.4 2.5-8.6 725-2409 1475-5590 100-372 50-516 0-172 0-248 -
aCombined
male and female pet ferrets (n = 60). From Cray C, Avian and Wildlife Laboratory, Miller School of Medicine, University of Miami, Miami, FL. bIntact males. cCastrated males.
20
SECTION I Ferrets Table 2-2 Reference Intervals for Biochemical Values in Ferrets SERUM Analyte
Plasmaa
Albinob
Fitchc
Alanine aminotransferase (U/L) Albumin (g/dL) Alkaline phosphatase (U/L) Amylase (U/L) Aspartate aminotransferase (U/L) Bilirubin, total (mg/dL) Blood urea nitrogen (mg/dL) Calcium (mg/dL) Carbon dioxide (mmol/L) Cholesterol (mg/dL) Chloride (mmol/L) Creatinine (mg/dL) Creatine phosphokinase (U/L) Glucose (mg/dL) Gamma glutamine transferase (U/L) Lactate dehydrogenase (U/L) Phosphorus (mg/dL) Potassium (mmol/L) Sodium (mmol/L) Total protein (g/dL) Triglycerides (mg/dL) Uric acid (mg/dL)
65-128 2.5-4.0 25-60 26-36 70-100 0.2-0.5 18-32 8.1-9.5 22-29 119-163 0.2-0.5 55-93 80-117 8-34 200-1400 5.1-6.5 4.5-6.1 142-148 4.5-6.2 30-140 1.3-1.9
2.6-3.8 9-84 28-120 <1.0 10-45 9.0-11.8 16-25 64-296 106-125 0.4-0.9 94-207 4.0-9.1 4.5-7.7 137-162 5.1-7.4 -
82-289 3.3-4.1 25-60 12-43 8.6-10.5 102-121 0.2-0.6 62-134 5.6-8.7 4.3-5.3 146-160 5.3-7.2 -
aCombined
male and female pet ferrets (n = 60). From Cray C, Avian and Wildlife Laboratory, Miller School of Medicine, University of Miami, Miami, FL. bCombined values of male (n = 40) and female (n = 24) ferrets.37 cCombined values of intact male, female, and castrated male ferrets (n = 13), age 4-8 months.16
Table 2-3 Reference Intervals for Plasma Protein Electrophoresis in Ferrets Analyte Albumin (g/dL) Alpha-1 globulin (g/dL) Alpha-2 globulin (g/dL) Beta globulin (g/dL) Gamma globulin (g/dL) Albumin/globulin ratio
Combined Sexa 2.50-3.31 0.33-0.56 0.36-0.60 0.83-1.20 0.31-0.81 1.05-1.33
aCombined
male and female ferrets, n = 60. From Cray C, Avian and Wildlife Laboratory, Miller School of Medicine, University of Miami, Miami, FL.
90.0 to 163.5 mg/dL), and mean antithrombin activity (AT) was 96% (range, 69.3% to 115.3%).6 In another study, the mean PT was 15.7 seconds (range, 14.4 to 16.5 seconds) in male ferrets.61 In a study of 6 ferrets, values of clotting time of whole blood were 2.0 ± 0.5 minutes in glass tubes and 3.0 ± 0.9 minutes in siliconized tubes; mean PT was 10.3 ± 0.1 seconds; aPPT was 18.4 ± 1.4 seconds; and thrombin time was 28.8 ± 8.7 seconds.31 In the same study, mean values for individual coagulation factors were also determined31 and have been reported elsewhere.17 In a study of 30 intact ferrets (15 male, 15 female), mean bleeding time was less than 2 minutes; PT was 8 to 11 seconds for females and 9 to 10.6 seconds for
males; and aPPT was 16 to 21 seconds for females and 17 to 25 seconds for males (E. Ivey, DVM, unpublished data, 2000). In this study, PT and aPPT were measured with the ACT II (Medtronic, Parker, CO).
INTRAVENOUS CATHETERS Indwelling intravenous catheters are routinely used in ferrets. Catheters can be placed in the lateral saphenous or cephalic vein (Fig. 2-4). Jugular vein catheters are difficult to place and are rarely used. Except in very depressed animals, catheters are placed with the ferret tranquilized or anesthetized. Applying a topical anesthetic containing lidocaine 2.5% and prilocaine 2.5% (EMLA cream, AstraZeneca LP, Wilmington, DE) on the venipuncture site approximately 30 to 60 minutes before the procedure may facilitate catheterization in the unanesthetized ferret. Shave and sterilely prepare the skin over the vein. Puncture the skin over the vein with a 20- or 22-gauge needle, taking care to avoid the vein; then introduce a short 22-, 24-, or 26-gauge over-the-needle catheter into the vein. Secure the catheter hub or a tape butterfly wrapped around the hub directly to the skin with suture, tape, or tissue adhesive. Attach a T-connector and wrap the leg with a soft padded bandage. Closely monitor ferrets with indwelling catheters to prevent the fluid line from entangling, and frequently check the leg distal and proximal to the catheter for soft tissue swelling. Most ferrets do not chew a catheter once it is placed and do not require an Elizabethan collar.
CHAPTER 2 Basic Approach to Veterinary Care
Fig. 2-4 Placing an indwelling catheter in the cephalic vein of a ferret. The hair over the vein is shaved and the vein held off. A 24-gauge, three-quarter–inch intravenous catheter is introduced into the cephalic vein and taped in place.
In ferrets that are collapsed with poor blood pressure or in young or very small ferrets, attempts to place an intravenous catheter may be unsuccessful. An intraosseous catheter can be placed in these animals. Contraindications to intraosseous catheterization include local infection or burns, fracture of the ipsilateral extremity, osteopenia, or osteopetrosis.59 The proximal tibia is the most common site for intraosseous catheter placement in small mammals,28 but the proximal femur can also be used; the latter site allows the patient more range of movement. Unless the ferret is very depressed, anesthesia is required to place the catheter. Sterilely prepare the insertion site, and in the conscious patient, infiltrate the area with 1% to 2% lidocaine (maximum dose 1 mg/kg).28 Insert a 20- or 22-gauge, 1.5-inch spinal needle into the marrow cavity. Alternatively, use a 20- or 22-gauge hypodermic needle with a surgical steel wire inserted into the lumen to prevent the needle from occluding during insertion.46 The intraosseous catheter should occupy approximately 33% to 67% of the marrow cavity at the narrowest portion.28 In humans, medications and blood products intended for intravenous administration are considered safe to administer via an intraosseous catheter.59 Therapeutic agents should be administered in small volumes while applying minimal pressure to prevent leakage from the insertion site.59 If possible, change to an intravenous catheter as soon as the animal is rehydrated or blood pressure improves. Intraosseous catheters should not be left in place for more than 72 hours.28 Vascular access ports, consisting of an indwelling intravenous catheter attached to an injection port placed in subcutaneous tissue, have been used in ferrets for repeated administration of chemotherapeutic medications. These ports can be used when repeated vascular access is required for any reason.52 The technique used to place the catheter and port has been described and illustrated.47
FLUID THERAPY Hospitalized ferrets usually require fluid therapy to maintain hydration and correct dehydration. Daily fluid requirements of ferrets have not been determined; however, calculating fluid
21
requirements based on rates used in cats (60 to 70 mL/kg per day) appears to be adequate for maintenance. One source estimates daily water consumption of adult ferrets as 75 to 100 mL/day.38 Provide additional fluids to compensate for ongoing fluid loss and to correct dehydration calculated as a percentage of the body weight. Give fluids subcutaneously or intravenously; intravenous fluids are preferred in ill animals. Administer subcutaneous fluids in the loose skin along the back and dorsal cervical area, dividing the calculated daily fluid volume into doses given two or three times daily. Ferrets often react painfully to subcutaneous fluid administration, and good restraint is needed to prevent a ferret from biting its handler. If possible, administer intravenous fluids by continuous rate infusion. Alternatively, administer fluids by dividing the calculated daily fluid volume into two or three doses administered by a Buretrol (Baxter Healthcare, Glendale, CA) or syringe pump. Depending on the clinical condition of the ferret, supplements and drugs can be added to fluids by using the same criteria and calculations as for dogs and cats. Colloids are effective in improving intravascular fluid volume and oncotic pressure in ferrets that are hypoproteinemic or in shock. Dosage and administration are similar to those in small animals (see Chapter 38). Most commonly, hydroxyethyl starch (hetastarch) is given at a dosage of 10 to 20 mL/kg per day. When hetastarch is coadministered with crystalloids, reduce the crystalloid fluid volume by 33% to 50%. In ferrets in shock, hetastarch can be given as a bolus at 5 mL/kg over a 15- to 30-minute period; this can be repeated to a total dose not exceeding 20 mL/kg per day.
ANTIBIOTIC AND DRUG THERAPY Ferrets are given antibiotics and other drugs at dosages similar to those used in cats (see Appendix). Intravenous antibiotics are preferred in sick animals if an indwelling catheter is in place, but the catheterized extremity should be monitored carefully for phlebitis and cellulitis, particularly when caustic medications are administered. Intramuscular antibiotics can be given, but subcutaneous administration is preferred because of the limited muscle mass in cachectic animals if therapy continues over several days. Because pills are very difficult to administer, oral medications are most easily given in a liquid form. Most compounding pharmacists can prepare suspensions of drugs that are not commercially available as liquids. Ferrets generally accept chicken and beef flavors; avoid fish flavors in compounded formulas, as ferrets do not generally like this taste.
PAIN MANAGEMENT Pain management is important in the postoperative period, with diseases that create significant discomfort (such as gastrointestinal ulceration), and for traumatic injuries (see Chapter 31). Ferrets in pain may exhibit tachypnea, a stiff gait, a strained facial expression, teeth grinding, shivering, half-closed eyelids, aggression, focal muscle fasciculations, hiding, general malaise, bristling of tail fur, and being “tucked” in the abdomen.36,62 Many analgesic agents are used effectively in ferrets, including opioids, alpha-2 agonists, nonsteroidal anti-inflammatory drugs (NSAIDs), and local anesthetics.26 Clinically, buprenorphine, butorphanol, and meloxicam are most commonly used for hospitalized animals and outpatients (see Chapter 31 and Appendix).
22
SECTION I Ferrets
For severe pain, combination therapy, such as with an opioid and an NSAID, may be most effective.26 Epidural administration of analgesics preoperatively appears to be effective in helping to control postoperative pain in ferrets (see Chapter 31). Epidural anesthesia and analgesia is most effective in procedures involving the abdomen, spine, pelvis, hind legs, tail, and perineum.18 In a study of ferrets undergoing ovariohysterectomy and bilateral anal sacculectomy, both physiologic and behavioral manifestations of pain were attenuated in ferrets that received epidural morphine preoperatively compared with control ferrets, and beneficial effects were seen for at least 24 hours.57 The technique has been well described and illustrated.18,23 Place the anesthetized ferret in sternal recumbency with the legs flexed to open the lumbosacral space. Clip a large rectangle of hair over the injection site (so that anatomic landmarks can easily be identified) and aseptically prepare the area. The injection site is at the intersection of a line drawn between the cranial aspect of the wings of the ilea and a line drawn between the dorsal spinous processes of the last lumber and first sacral (S1) vertebrae; the dorsal spinous process of S1 is easily palpable. Either a 22-gauge, 1.5-inch spinal needle or a 25-gauge hypodermic needle can be used, although a spinal needle will prevent creation of a skin plug that can obstruct the flow of cerebrospinal fluid (CSF) or set up a nidus for infection and inflammation.18 Insert the needle on the midline perpendicular to the skin and with the bevel facing cranially. A correctly placed needle will drop through the lumbosacral space until it encounters bone on the ventral floor of the spinal canal. If using a spinal needle, remove the stylet and observe the needle hub for blood or CSF. Confirm correct epidural placement by attaching a 1-mL syringe containing 0.2 mL of sterile saline and 0.2 mL of air and applying gentle suction; no CSF or blood should be seen in the needle hub. No resistance should be encountered when the fluid is injected, and the air space above the fluid should not be compressed. Once correct epidural placement is confirmed, inject analgesic or anesthetic fluids slowly to avoid an increase in intracranial pressures or extended cranial infiltration of the anesthetic block. Morphine is often used for epidural analgesia at a dose of 0.1 mg/kg.23 Epidural injection is contraindicated in cases of coagulopathy, sepsis, hypovolemia, skin infection, and local fractures.18 Be careful in the immediate postoperative period when using drugs such as butorphanol, which can cause a pronounced sedative effect. Ferrets can remain very lethargic and immobile for long periods and can overheat quickly if a heating lamp or other strong heat source is used. Therefore closely monitor the body temperature of any immobile or lethargic ferret given pain medication to prevent overheating when using heat lamps or forced air warming devices. The frequent temperature spike described in ferrets 30 minutes or longer after surgery or sedation may be related to administration of various analgesics, similar to a syndrome seen in cats; however, responses in ferrets are inconsistent.29 Like cats, ferrets are sensitive to acetaminophen toxicity.16 The activity of UDP-glucuronosyltransferase in their livers is similar to that of cats. Therefore acetaminophen glucuronidation is slower in ferrets than in other non-felid species. Unlike cats, however, no genetic mutations are associated with this slow metabolism, and the exact cause is not known. When dosed inappropriately, ibuprofen can also be toxic in ferrets.13 Therefore use any NSAIDs with caution. Do not use NSAIDs in ferrets already being treated with a corticosteroid for insulinoma or other disease.
NUTRITIONAL SUPPORT Many sick ferrets are either cachectic or have minimal body fat and require nutritional support. Force-feeding is also important to prevent hypoglycemia in anoretic ferrets with insulinomas. Ferrets can be syringe fed meat-based soft foods marketed for hospitalized dogs and cats such as Maximum-Calorie (The Iams Company) or Canine a/d (Hill’s Pet Nutrition). Alternatively, products formulated as recovery diets for carnivores are available and readily accepted by most ferrets (Carnivore Care, Oxbow Animal Health, Murdock, NE; Emeraid Carnivore, LafeberVet.com, Cornell, IL). Although readily accepted, commercial supplement gels based on corn syrup should be offered cautiously or not at all because of the high sugar content. Force-feed anorectic ferrets as much as they will accept comfortably, usually 8 to 12 mL fed three or four times daily. Use a syringe to administer food. Once a ferret develops a taste for the food, it may eat it directly from a bowl. Although rarely used clinically, esophagostomy feeding tubes can be placed in ferrets to manage debilitated animals over the long term. The technique is similar to that used in cats.20 Gastric feeding tubes have been placed in ferrets both experimentally and clinically.5,10 In a study of 14 ferrets, gastrostomy tubes were placed percutaneously by a nonendoscopic technique. A gastrostomy tube was placed in a ferret after surgical repair of an esophageal perforation caused by an esophageal foreign body.10 The tube was maintained successfully throughout the postoperative healing period. Total nutrient admixtures have been formulated to provide partial parenteral nutrition to ferrets.46,54 Parenteral nutrition should be considered if an esophageal, gastric, or intestinal disorder precludes the use of enteral formulations, for example, in cases of malabsorptive diarrhea. The total nutrient mixture is formulated from a mixture of lipid and dextrose supplemented with amino acids, electrolytes, water-soluble vitamins, minerals, and enough fluids to meet daily fluid volume requirements. Depending on the osmolarity of the solution, parenteral nutrition solutions can be delivered via a central, peripheral, intraosseous, or intraperitoneal catheter; however, the relatively high protein requirements of ferrets usually result in a solution of 600 to 800 mOsm/L, which should be administered into a large (central) vein.54 Ferret owners often prepare homemade diets of “duck soup” or “chicken gravy” to nurse their pets at home. Many different recipe variations are available online. These recipes are usually based on canned dog food, kibble, or whole chicken with additives ranging from beef fat, nutritional supplement gels, or brewer’s yeast to Echinacea capsules. The “duck soup” variations all provide a soft, porridge-consistency food that is usually readily eaten by sick and convalescing ferrets. Although many of these recipes appear to be acceptable, some are very high in fat and carbohydrates. Unless the homemade diets are based on or used with a commercial ferret diet, they should not be used long term because of possible nutritional imbalances and deficiencies. Discuss any particular recipe that a ferret owner is using before endorsing it for long-term use.
URINE COLLECTION AND URINALYSIS Urine samples can be collected by cystocentesis or by free catch after natural voiding or gentle manual expression of the bladder. The techniques for manually expressing the bladder and
CHAPTER 2 Basic Approach to Veterinary Care Table 2-4 Reference Values for Urinalysis in Ferrets Value
Mean ± SD
Volume (mL/24 hr) Males Females pH Urine protein (mg/dL) Males Females Exogenous creatinine clearance (mL/min/kg) Insulin clearance (mL/min/kg) Endogenous creatinine clearance (mL/min/kg)
24.9 ± 14.319 2637 2837
3.3 ± 2.219
Range 8-4837 8-14037 6.5-7.537 7-3337 0-3237
3.0 ± 1.819 2.5 ± 0.919
cystocentesis are the same as those used in dogs and cats. Anesthetize fractious ferrets to avoid trauma to the thin bladder wall. Use a 25-gauge needle for cystocentesis. Reference values for urinalysis are listed in Table 2-4. In one study, the reference range for urine pH in ferrets was reported as 6.5 to 7.5; however, urine pH can vary according to the diet, and the normal urine pH in ferrets fed a high-quality, meat-based diet is approximately 6.0.
URINARY CATHETERIZATION Urinary catheterization is commonly indicated in male ferrets, but the procedure can be challenging. Although techniques have been described for both sexes,34 clinical indications to place a urinary catheter in females are rare. For females, tranquilize or anesthetize the ferret, then position it in ventral recumbency with the rear quarters elevated with a rolled towel. With a vaginal speculum, locate the urethral opening in the floor of the urethral vestibule, approximately 1 cm cranial to the clitoral fossa. Introduce a 3.5-French, red rubber urethral catheter fitted with a wire stylet into the urethral orifice. In male ferrets, urethral blockage is a common sequela of adrenal disease. Androgens produced by the adrenal gland stimulate the prostate gland to enlarge, which subsequently constricts the urethra. Placing a urinary catheter is difficult because the urethral opening is very small and located on the ventral surface of the penis, below the J-hook in the end of the os penis. Also, in ferrets with urethral blockage, the tip of the penis and the preputial area are often very swollen, and introducing a catheter can be challenging. If needed, a small incision can be made in the prepuce to facilitate exteriorizing the penis. Surgical magnifying loupes may be helpful in locating the urethral orifice. To place a catheter, use a 3.0-French polytetrafluroethylene urinary catheter (Slippery Sam Tomcat Urethral Catheters, Surgivet, Smiths Medical, Norwell, MA; Ferret Urinary Catheter, Mila International, Erlanger, KY) or a 3.5-French rubber feeding catheter (see also Chapter 4). If using a long rubber catheter, estimate the length of the catheter that must be inserted to reach the bladder before placing it. Use a stylet or flexible KE wire to stiffen the catheter while passing. Another option is to use a 20or 22-gauge, 8-inch jugular catheter with the stylet removed.46 If needed, the stylet can be left in but retracted slightly to provide stiffness, but be very careful when rounding the pelvic flexure to
23
avoid perforating the urethra. Use sterile technique to prepare the urethral and preputial area and pass the catheter. If the urethral opening is difficult to see, dilate the opening by passing a 24-gauge intravenous catheter just inside the tip of the urethra and flushing gently with saline. Then slip the tip of the lubricated urinary catheter gently into the dilated opening alongside the intravenous catheter and, while gently flushing with saline solution, pass the catheter into the bladder. Often resistance is met at the pelvic flexure; if this occurs, try repeated gentle flushing and relubricating the catheter until it passes. Once in place, secure the catheter by suturing the hub to the prepuce if using a Slippery Sam catheter. If using a rubber catheter, place butterfly tape strips around the catheter just as it enters the urethra and at another point 3 to 5 cm distal and suture these to the skin. Attach a sterile urinary collection device, and tape the tubing to the tail to further prevent tension. If needed, bandage the ferret’s abdomen to minimize rotation of the catheter and to restrict the ferret from traumatizing it. Soft Elizabethan collars are needed in some ferrets to prevent chewing at the catheter. Maintain sterility of the collection system, and drain the bag by needle and syringe rather than opening the system (see also Chapter 4). Temporary tube cystostomy has been used successfully to manage male ferrets with urinary obstruction caused by adrenal disease. In four ferrets treated surgically, a 5- or 8-French Foley catheter was placed in the bladder at the time of adrenalectomy and left in place for 5 to 14 days.44 In these ferrets, immediate treatment of urinary blockage was by cystocentesis. A cystostomy catheter also can be placed by interventional radiographic techniques. This is an important option in obstructed ferrets in which a urethral catheter cannot be passed and surgery is considered high risk because of the poor condition of the patient. A cystostomy catheter allows azotemic ferrets to be managed with diuresis before surgery; alternatively, in cases in which surgery is not an option, the catheter can remain in place until response to medical management with leuprolide acetate occurs.
SPLENIC ASPIRATION Splenic aspiration is a common diagnostic technique that is used in ferrets with enlarged spleens (see Chapters 5 and 36). The technique is simple and usually can be done in unanesthetized ferrets. However, if a ferret is fractious, use an injectable sedative or inhalant anesthesia administered by face mask. Restrain the ferret on its back or in lateral recumbency, and shave and prepare the abdominal skin in the area over the spleen. Palpate and immobilize the spleen directly under the prepped area with one hand while inserting a 25-gauge needle into the spleen several times with the other hand. Then attach an airfilled syringe to express the contents of the needle onto slides. This technique will minimize blood contamination. Alternatively, attach a 25-gauge needle to a 3-mL syringe and aspirate quickly after directing the needle into the spleen. A positive aspirate appears bloody. Detach the needle, fill the syringe with air, and express the samples onto slides. Obtain samples from two sites and prepare several slides for cytologic staining. If an abnormal mass is found on ultrasound examination, perform an ultrasound-guided aspirate to improve chances of a positive result. The two most common findings on cytologic examination of a splenic aspirate are extramedullary hematopoiesis and lymphoma.
SECTION I Ferrets
24
BONE MARROW COLLECTION Evaluating a bone marrow sample is a valuable diagnostic tool for many disease conditions, including anemia, thrombocytopenia, pancytopenia, proliferative abnormalities, and suspected hematopoietic malignancies. Anesthesia is necessary to aspirate the bone marrow or perform a core biopsy. Although the proximal femur is usually the most readily accessible site, the iliac crest, tibial crest, and humerus can also be used to collect bone marrow samples (Fig. 2-5). After the ferret is anesthetized, place it in lateral recumbency and shave and aseptically prepare the area around the collection site. For the proximal femur,50 make a small incision through the skin over the greater trochanter with a no. 15 scalpel blade. Hold and stabilize the femur with one hand while inserting a 20-gauge, 1.5-inch spinal needle into the bone medial to the greater trochanter. Use steady pressure and an alternating rotating motion to advance the needle into the marrow cavity. Withdraw the stylet, and attach a 6- to 12-mL syringe to the needle. Aspirate the marrow sample into the syringe, stopping suction as soon as the sample is visible (to prevent blood contamination). To collect a core biopsy sample, use the same technique, but use a 1.5-inch, 18-gauge needle in place of the spinal needle.63 Collect samples from alternate sites by using the same basic technique. Try to prepare at least four to eight slides for cytologic evaluation. To do this, forcibly expel the bone marrow sample from the syringe onto glass slides. The slide can be held vertically to allow contaminating blood to drain, leaving only bony spicules. Place a clean slide on top of the slide with the sample and allow the marrow to spread between the slides, then draw the two slides apart in a horizontal plane.63
TRACHEAL WASH Ferrets will occasionally have clinical and radiographic evidence of respiratory disease. In these animals, a tracheal wash may be indicated to obtain samples for cytologic examination and bacterial culture and sensitivity testing. The procedure is similar to that used in a cat. Anesthetize the ferret and intubate with a sterile endotracheal tube. An 8-French pediatric suction catheter (Safe-T-Vac Suction, Kendall Healthcare Products, Mansfield, MA) connected to a specimen container (Argyle
A
Lukens Specimen Container, Sherwood Medical, St. Louis, MO) and attached to a wall suction outlet will maximize the volume of sample collected. Pass the tip of the suction catheter through the endotracheal tube, preferably to the level of tracheal bifurcation. Inject up to 2 mL of warm, sterile saline solution, then aspirate the fluid into the specimen container.
BLOOD TRANSFUSION Blood transfusions may be needed in ferrets that are anemic from chronic disease, blood loss, or estrogen toxicosis or in ferrets that are thrombocytopenic. As in other species, evaluate the need for a transfusion based on the packed cell volume or platelet count and clinical status of the ferret. Consider a transfusion if the packed cell volume is 25% or less in a ferret that exhibits clinical signs of anemia or requires surgery or if a ferret is thrombocytopenic and exhibits ecchymosis, petechiation, or bleeding. Ferrets lack detectable blood groups and there is little risk of transfusion reaction, even without cross-matching.33 Because they have a larger blood volume, large male ferrets are preferred over females as blood donors. Depending on the size of the donor ferret, 6 to 12 mL of blood can be safely collected for transfusion. Collect blood into an anticoagulant such as acidcitrate-dextrose at a ratio of 1 mL of anticoagulant to 6 mL of donor blood.24 Always use a filter when transfusing whole blood, and use at least a 22-gauge catheter (to prevent cell lysis). Intraosseous blood transfusions can be given to ferrets if an intravenous catheter cannot be placed. Use of a hemoglobin-based oxygen-carrying solution obviates the need for a donor ferret and a filter for administration, and the solution can be administered through a catheter of any size. A hemoglobin-based oxygen-carrying solution (Oxyglobin, Biopure Corp., Cambridge, MA) has been used in anemic ferrets at a dose of 11 to 15 mg/kg infused over a 4-hour period and administered once or twice during a 24-hour period.48,49 However, this product is currently unavailable.
BLOOD PRESSURE MONITORING Indirect blood pressure monitoring techniques in ferrets, using both Doppler ultrasound and oscillometric methods, have been described.30,45,56 However, both methods have shown poor
B Fig. 2-5 A, Preparing to collect a bone marrow sample from the humerus of a ferret. The site over the proximal humerus is shaved and aseptically prepared. B, Collection of a bone marrow sample from the femur of a ferret with an enlarged vulva. The femur is stabilized while inserting a 20-gauge, 1.5-inch spinal needle medial to the greater trochanter.
CHAPTER 2 Basic Approach to Veterinary Care correspondence with direct arterial blood pressure measurements. In one study using 10 healthy adult female ferrets to compare direct blood pressure measurement from the carotid artery to indirect blood pressure measurement from the tail and limbs using both Doppler and oscillometric methods, the indirect systolic blood pressure measurements were approximately 28 to 30 mm Hg less than the direct systolic blood pressures. Accurate indirect blood pressure readings were deemed very difficult to obtain because the neonatal blood pressure cuff did not fit securely on the short limbs of the ferrets (Doppler method) and the tail artery did not produce sufficiently high pressure changes to be detected by the oscillometric system. The authors concluded that indirect blood pressure assessment, using a sphygmomanometer and Doppler probe, could still be considered useful for evaluating changes in blood pressure from a known baseline or for surgical monitoring of general trends.45 In another study using 14 male ferrets, indirect blood pressure measurements using an oscillometric sphygmomanometer and a veterinary high definition oscillometry monitor on the tail, forelimb, and hindlimb were compared with direct arterial blood pressure measurements. Measurements using the tail were considered most reliable; however, the oscillometric sphygmomanometer consistently overestimated the systolic arterial pressure. In addition, the indirect measurements of systolic, mean, and diastolic arterial pressures obtained with the highdefinition oscillometry monitor were consistently higher than the corresponding direct blood pressure measurements during hypotensive states but were substantially lower than the corresponding direct blood pressure measurements in hypertensive states.56 Therefore, be aware of the above described inconsistencies of indirect methods when compared with results of direct blood pressure measurement, and interpret clinical results to measure trends more than for accurate measurements. In clinical practice, the Doppler method of indirect blood pressure measurement is most commonly used in ferrets and other small mammals. Shave the hair on the ventral carpus or tarsus (overlying the radial artery on the front leg or the digital branch of the tibial artery on the rear leg), and place the ferret in lateral or sternal recumbency. Place a pneumatic cuff (infant or number 1) above the carpus or tarsus or on the distal humerus, and attach it to a sphygmomanometer. Place the Doppler transducer probe crystal on the shaved skin in a bed of ultrasonic gel and tape or hold in place. Inflate the cuff bladder to a pressure exceeding the systolic blood pressure, which will diminish the Doppler signal of blood flow. Deflate the blood pressure cuff gradually; the first sound heard denotes the systolic pressure.30 Normal systolic blood pressure in ferrets is similar to that of other small mammals (80 to 120 mm Hg).
DIAGNOSTIC PERITONEAL LAVAGE Ferrets are commonly presented for disease involving the abdominal cavity. Diagnostic peritoneal lavage has been widely used in human and veterinary medicine for assessment of abdominal fluid accumulation secondary to organ rupture, hemorrhage, and neoplasia. In humans, diagnostic peritoneal lavage is a more sensitive diagnostic test than abdominocentesis, and less fluid is required for diagnosis. In dogs and cats, accuracy of diagnosis is higher with diagnostic peritoneal lavage than with abdominocentesis.51 The technique described in ferrets is similar to that used in dogs and cats.51 Anesthetize or sedate the ferret, depending
25
on the stability of the patient. Place the ferret in dorsal recumbency to facilitate identification and retraction of the spleen, and, if possible, empty the urinary bladder. Shave and aseptically prepare the skin caudal to the umbilicus, retracting the spleen as necessary. Infuse the skin and body wall at the entry site with 2% lidocaine, and make a small stab incision through the skin. Elevate the body wall with sterile forceps, and inset an 18- to 20-gauge over-the-needle catheter through the body wall, being cautious to avoid the spleen. The catheter can be sterilely fenestrated before insertion to optimize fluid recovery; however, take care to remove all cut pieces. Advance the catheter caudodorsally, and remove the stylette. Aspirate the catheter; if no fluid appears, instill 20 to 22 mL/kg of warm sterile isotonic saline. Rock the ferret gently or massage the abdomen for 1 to 2 minutes. Aspirate the fluid and place into sterile containers for evaluation. If necessary, repeat fluid instillation at up to half the initial volume used. Remove the catheter, and suture or glue the incision. Treat with a systemic analgesic subsequent to the procedure.
References 1. Abe N, Tanoue T, Noguchi E, et al. Molecular characterization of Giardia duodenalis isolates from domestic ferrets. Parasitol Res. 2010;106:733-736. 2. Appel MJ, Harris WV. Antibody titers in domestic ferret jills and their kits to canine distemper virus vaccine. J Am Vet Med Assoc. 1988;193:332-333. 3. Bell JA. Parasites of domesticated pet ferrets. Compend Cont Educ Pract Vet. 1994;16:617-620. 4. Bell JF, Moore GJ. Susceptibility of carnivore to rabies virus administered orally. Am J Epidemiol. 1971;93:176-182. 5. Benson KG, Paul-Murphy J, Carr A. Percutaneous placement of a gastric feeding tube in the ferret. Lab Anim. 2000; 29:44-46. 6. Benson KG, Paul-Murphy J, Hart AP, et al. Coagulation values in normal ferrets (Mustela putorius furo) using selected methods and reagents. Vet Clin Pathol. 2008;37:286-288. 7. Blancou J, Aubert MFA, Artois M. Experimental rabies in the ferret (Mustela [putorius] furo): susceptibility—symptoms— excretion of the virus. Rev Med Vet. 1982;133:553-557. 8. Bleakley SP. Simple technique for bleeding ferrets (Mustela putorius furo). Lab Anim. 1980;14:59-60. 9. Brown SA. Clinical techniques in domestic ferrets. Sem Avian Exot Pet Med. 1997;6:75-85. 10. Caligiuri R, Bellah JR, Collins BR, et al. Medical and surgical management of esophageal foreign body in a ferret. J Am Vet Med Assoc. 1989;195:969-971. 11. Carroll EE, Dubielzig RR, Schultz RD. Cats differ from mink and ferrets in their response to commercial vaccines: a histologic comparison of early vaccine reactions. Vet Pathol. 2002;39:216-227. 12. Castanheira de Matos RE, Morrisey JK. Common procedures in the pet ferret. Vet Clin North Am Exot Anim Pract. 2006;9:347-365. 13. Cathers TE, Isaza R, Oehme F. Acute ibuprofen toxicosis in a ferret. J Am Vet Med Assoc. 2000;216:1426-1428:1412. 14. Church RR. Impact of diet on the dentition of the domesticated ferret. Exot DVM. 2007;9:30-39. 15. Compendium of Animal Rabies Prevention and Control, 2008. National Association of State Public Health Veterinarians. Available at www.nasphv.org/documentsCompendia.html. Accessed January 31, 2011. 16. Court MH. Acetaminophen UDP-glucuronosyltransferase in ferrets: species and gender differences, and sequence analysis of ferret UGT1A6. J Vet Pharmacol Ther. 2001;24:415-422.
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17. Dodds WJ. Rabbit and ferret hemostasis. In: Fudge AM, ed. Laboratory medicine: avian and exotic pets. Philadelphia: WB Saunders; 2000:285-290. 18. Eshar D, Wilson J. Epidural anesthesia and analgesia in ferrets. Lab Anim. 2010;39:339-340. 19. Esteves MI, Marini RP, Ryden EB, et al. Estimation of glomerular filtration rate and evaluation of renal function in ferrets (Mustela putorius furo). Am J Vet Res. 1994;55:166-172. 20. Fisher PG. Esophagotomy feeding tube placement in the ferret. Exot DVM. 2001;2:23-25. 21. Fox JG. Normal clinical and biologic parameters. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:183-210. 22. Fudge AM. Ferret hematology. In: Fudge AM, ed. Laboratory medicine: avian and exotic pets. Philadelphia: WB Saunders; 2000:269-272. 23. Harms CA, Sladky KK, Horne WA, Stoskopf MK. Epidural analgesia in ferrets. Exot DVM. 2002;4.3:40-42. 24. Hoefer HL. Transfusions in exotic species. In: Hohenhaus AE, ed. Transfusion medicine. Philadelphia: JB Lippincott; 1992: 625-635. 25. Johnson-Delaney CA. Diagnosis and treatment of dental disease in ferrets. J Exot Pet Med. 2008;17:132-137. 26. Ko J, Marini RP. Anesthesia and analgesia in ferrets. In: Fish RE, Danneman PJ, Brown M, et al. eds. Anesthesia and analgesia in laboratory animals. 2nd ed. San Diego: Academic Press, an imprint of Elsevier; 2008:443-456. 27. Lee EJ, Moore WE, Fryer HC, et al. Haematological and serum chemistry profiles of ferrets (Mustela putorius furo). Lab Anim. 1982;16:133-137. 28. Lennox AM. Intraosseous catheterization of exotic animals. J Exot Pet Med. 2008;17:300-306. 29. Lennox AM. Post-surgical hyperthermia in ferrets. Proc Assoc Exot Mam Vet. 2009:9-10. Available at www.aemv.org/documents/ AEMV2009ScientificSession.pdf. Accessed March 5, 2011. 30. Lichtenberger M, Ko J. Critical care monitoring. Vet Clin North Am Exot Anim Pract. 2007;10:317-344. 31. Lewis JH. Comparative hemostasis in vertebrates. New York: Plenum Press; 1996. 32. Mann PH, Harper DS, Regnier S. Reduction of calculus accumulation in domestic ferrets with two dentifrices containing pyrophosphate. J Dent Res. 1990;69:451-453. 33. Manning DD, Bell JA. Lack of detectable blood groups in domestic ferrets: implications for transfusion. J Am Vet Med Assoc. 1990;197:84-86. 34. Marini RP, Esteves MI, Fox JG. A technique for catheterization of the urinary bladder in the ferret. Lab Anim. 1994;28:155-157. 35. Marini RP, Jackson LR, Esteves MI, et al. Effect of isoflurane on hematologic variables in ferrets. Am J Vet Res. 1994;55:1479-1483. 36. Mayer J. Use of behavior analysis to recognize pain in small mammals. Lab Anim. 2007;36:43-48. 37. Meyer EK. Vaccine-associated adverse events. Vet Clin North Am Small Anim Pract. 2001;31:493-514. 38. Moody KD, Bowman TA, Lang CM. Laboratory management of the ferret for biomedical research. Lab Anim Sci. 1985;35:272-279. 39. Munday JS, Stedman NL, Richey LJ. Histology and immunochemistry of seven ferret vaccination-site fibrosarcomas. Vet Pathol. 2003;40:288-293. 40. Murray J. Vaccine injection-site sarcoma in a ferret [letter]. J Am Vet Med Assoc. 1998;213:955. 41. Mustonen AM, Puukka M, Rouvinene-Watt K, et al. Response to fasting in an unnaturally obese carnivore, the captive European polecat Mustela putorius. Exp Biol Med (Maywood). 2009;234:1287-1295.
42. Niezgoda M, Briggs DJ, Shaddock J, et al. Pathogenesis of experimentally induced rabies in domestic ferrets. Am J Vet Res. 1997;58:1327-1331. 43. Niezgoda M, Briggs DJ, Shaddock J, et al. Viral excretion in domestic ferrets (Mustela putorius furo) inoculated with a raccoon rabies isolate. Am J Vet Res. 1998;59:1629-1632. 44. Nolte DM, Carberry CA, Gannon KM, et al. Temporary tube cystostomy as a treatment for urinary obstruction secondary to adrenal disease in four ferrets. J Am Anim Hosp Assoc. 2002;38:527-532. 45. Olin JM, Smith TJ, Talcott MR. Evaluation of noninvasive monitoring techniques in domestic ferrets (Mustela putorius furo). Am J Vet Res. 1997;58:1065-1069. 46. Orcutt C. Emergency and critical care of ferrets. Vet Clin North Am Exot Anim Pract. 1998;1:99-126. 47. Orcutt C. Use of vascular access ports in exotic animals. Exot DVM. 2000;2.3:34-38. 48. Orcutt C. Oxyglobin administration for the treatment of anemia in ferrets. Exot DVM. 2000;2.3:44-46. 49. Orcutt C. Update on oxyglobin use in ferrets. Exot DVM. 2001;3.3:29-30. 50. Palley LS, Marini RP, Rosenblad WD, et al. A technique for femoral bone marrow collection in the ferret. Lab Anim Sci. 1990;40:654-655. 51. Powers L. Use of the diagnostic peritoneal lavage in the domestic ferret. Proc Assoc Exot Mam Vet. 2010:53-59. Available at www.aemv.org/members_only/SmallBook2010FINAL.pdf. Accessed March 13, 2011. 52. Rassnick KM, Gould WJ, Flanders JA. Use of a vascular access system for administration of chemotherapeutic agents to a ferret with lymphoma. J Am Vet Med Assoc. 1995;206:500-504. 53. Rehg JE, Gigliotti F, Stokes DC. Cryptosporidiosis in ferrets. Lab Anim Sci. 1988;38:155-158. 54. Remillard RL. Parenteral nutrition support in rabbits and ferrets. J Exot Pet Med. 2006;15:248-254. 55. Rupprecht CE, Gilbert J, Pitts R, et al. Evaluation of an inactivated rabies vaccine in domestic ferrets. J Am Vet Med Assoc. 1990;196:1614-1616. 56. Shoemaker NJ, Bosman IH. Intra-arterial blood pressure in ferrets compared to peripheral blood pressure. Proc Assoc Exot Mam Vets. 2009:3-4. Available at www.aemv.org/ documents/AEMV2009ScientificSession.pdf. Accessed March 5, 2011. 57. Sladky KK, Horne WA, Goodrowe KL, et al. Evaluation of epidural morphine for postoperative analgesia in ferrets (Mustela putorius furo). Contemp Top Lab Anim Sci. 2000;39:33-38. 58. Tanner PA, Tseggai T, Rice Conlon JA, et al. Minimum protective dose (MPD) and efficacy determination of a recombinant canine distemper virus vaccine for ferrets. Proc 81st Ann Meet Conf Research Workers Animal Dis. 2000:Abstract 156. 59. Tay ET, Hafeez W. Intraosseous access. eMedicine. Updated April 12, 2009. Available at http://emedicine.medscape.com/article/ 80431-overview. Accessed March 2, 2011. 60. Taylor B. Alternate technique for venipuncture in ferrets. Exot DVM. 2001;2.6:37-38. 61. Thornton PC, Wright PA, Sacra PJ, et al. The ferret, Mustela putorius furo, as a new species in toxicology. Lab Anim. 1979:119-124. 62. van Oostrom H, Schoemaker NJ, Uilenreef JJ. Pain management in ferrets. Vet Clin North Am Exot Anim Pract. 2011;14:105-116. 63. Williams BH. Disorders of rabbit and ferret bone marrow. In: Fudge AM, ed. Laboratory medicine: avian and exotic pets. Philadelphia: WB Saunders; 2000:276-284. 64. Wimsatt J, Jay MT, Innes KE, et al. Serologic evaluation, efficacy, and safety of a commercial modified-live canine distemper vaccine in domestic ferrets. Am J Vet Res. 2001;62:736-740.
CHAPTER
3
Gastrointestinal Diseases
Heidi L. Hoefer, DVM, Diplomate ABVP (Avian), James G. Fox, DVM, MS, Diplomate ACLAM, and Judith A. Bell, DVM, PhD
General Gastrointestinal Disorders Dental Disease Salivary Mucocele Oral Neoplasia Esophageal Disease Gastritis and Ulceration Helicobacter Mustelae Gastritis Gastrointestinal Polyps Gastric Distention (Bloat) Gastrointestinal Foreign Bodies Liver Disease Gastrointestinal Parasitism Enteritis and Diarrhea Salmonellosis Mycobacteriosis Campylobacteriosis Viral Diarrhea Inflammatory Bowel Disease and Eosinophilic Gastroenteritis Proliferative Bowel Disease Clinical Signs and Diagnosis of Proliferative Bowel Disease Treatment of Proliferative Bowel Disease Rectal Disease Neoplasia General Approach to Vomiting General Approach to Diarrhea Differentiation of Emaciation (“Wasting Disease”) with Diarrhea Steps in Diagnosis of Wasting Disease Treatment of Ferrets with Wasting and Diarrhea
Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
GENERAL GASTROINTESTINAL DISORDERS Disease of the gastrointestinal (GI) tract is common in ferrets. Clinicians should be familiar with the more common GI disorders in ferrets and be able to recognize clinical signs and differentiate among potential diagnoses.
DENTAL DISEASE Ferrets are obligate carnivores with specialized tooth form and function designed to consume animal tissue. Compared with New Zealand feral ferrets, North American pet ferrets have a much greater amount of dental pathology. This is speculated to be a result of dental trauma from inappropriate chewing behavior and kibble-mediated disease.14 Dry kibble, the mainstay of most pet ferret diets, may be responsible for structural changes to the teeth. These low-moisture, hard crunchy diets appear to be quite abrasive to ferret teeth and result in significant wear to the cheek teeth and molars.14 Although moist or semimoist diets have been associated with the formation of dental calculi and periodontal disease in experimental cases,40 most ferrets on a dry kibble diet develop tartar and gingivitis that progresses with age (Fig. 3-1). Periodontal disease is considered pervasive in pet ferrets (see Chapter 32). Chewing inappropriate objects (cage bars and toys) can lead to damage. Biting and gnawing habits often result in discoloration, wearing, and breaking of the tips of the canine teeth. Broken canine teeth do not usually result in obvious discomfort or pain unless the dental pulp is exposed. Root canal restoration or surgical removal of the affected teeth may be necessary in some ferrets.44 Tooth root abscesses are uncommon in ferrets. Although dysphagia and drooling are sometimes seen, dental disease is often an incidental finding during physical examination. Dental extractions and scaling can be performed with the animal under anesthesia. Follow the basic principles for dental disease management that apply in the care of the dog or cat. Offering a natural prey diet or moistening the dry kibble may decrease dental abrasions. 27
28
SECTION I Ferrets
Fig. 3-1 Broken canine teeth and dental tartar are common in ferrets.
Fig. 3-3 Oral squamous cell carcinoma in situ, multiple sites, in the mouth of a ferret.
of the affected salivary gland is ideal for avoiding recurrence (see Chapter 11). It may be possible to inject contrast medium into the mucocele in an effort to trace the origin of the saliva. Before attempting surgical excision of a salivary gland, review the superficial anatomy of the head and neck region of the ferret.66 Recurrence is possible.
ORAL NEOPLASIA
Fig. 3-2 Surgical correction of a salivary mucocele. The medial aspect of the mucocele is marsupialized into the mouth.
SALIVARY MUCOCELE Ferrets have five major pairs of salivary glands: the parotid, submandibular, sublingual, molar, and zygomatic.66 Trauma to a gland can result in extravasation of saliva and salivary mucocele formation. Although this lesion is uncommon in ferrets, mucocele diagnosis and treatment have been described.5,57 Diagnosis of a mucocele is relatively straightforward. Facial swellings are often seen in the commissures of the mouth or in the orbital area in the case of a zygomatic mucocele. Other locations also are possible. Aspirate the mass to obtain samples for cytologic analysis. The fluid is viscous or mucinous and clear or blood-tinged. Cytologic examination reveals amorphous debris and occasional red blood cells. Treatment for salivary mucoceles is usually surgery. In one reported case, scalpel blade lancing of the medial wall of the mucocele resulted in drainage and no recurrence.5 Marsupialization into the mouth with the use of a wide circular incision in the medial wall of the mucocele may be effective for mucoceles that bulge into the oral cavity (Fig. 3-2). Surgical excision
The oral cavity is an uncommon site of neoplasia in ferrets. Squamous cell carcinoma is the most commonly reported oral tumor in ferrets and typically manifests as a firm swelling of the upper or lower mandible.36,38,47 These masses are usually solitary but can appear in situ, in multiple sites (Fig. 3-3). Treat squamous cell carcinoma with wide surgical excision, including maxillectomy or mandibulectomy as required. In one report, surgical resection of the mass was followed with radiation therapy.36
ESOPHAGEAL DISEASE Diseases of the esophagus are rare in ferrets. Acquired megaesophagus has been reported in ferrets8,39 and is occasionally seen in practice. Megaesophagus describes an esophagus that is enlarged (dilated) on radiographic examination and that lacks normal motility. Recognizing this disease is important because the prognosis in ferrets with megaesophagus is poor. Clinical signs include lethargy, inappetence or anorexia, dysphagia, and weight loss. Regurgitation is common. Coughing or choking motions are sometimes described, and some ferrets have labored breathing. Differential diagnoses includes the presence of an esophageal or GI foreign body, gastritis, influenza, and respiratory diseases. Diagnosis of megaesophagus is based on clinical signs and radiographic evidence. On radiographs, the esophagus is often dilated in both the cervical and thoracic segments (Fig. 3-4). Food may be visualized in the esophagus. Aspiration pneumonia and gastric gas are sometimes evident in addition to esophageal dilation. In suspect cases, always take radiographs of the abdomen to exclude lower GI disease. Administer barium
CHAPTER 3 Gastrointestinal Diseases
A
29
B Fig. 3-4 A, Lateral thoracic radiograph of a ferret with megaesophagus. Note the subtle dilation of the thoracic esophagus (arrows). B, Ventrodorsal radiograph of the same ferret in A. The cranial thoracic esophagus is dilated (arrow) and is much easier to visualize in this view than in the lateral view.
(10 mL/kg by mouth [PO]) to delineate the esophagus and to evaluate mural lesions, strictures, or obstructions (Fig. 3-5). An endoscope can also be used to evaluate the esophagus. Use fluoroscopy, if available, to determine the motility of the esophagus after a barium swallow. The cause of megaesophagus in ferrets is unknown. Consider possibilities in the differential diagnosis as for dogs, and tailor the diagnostic workup accordingly. To test for myasthenia gravis, serum acetylcholinesterase antibody testing can be performed (Comparative Neuromuscular Laboratory, University of California San Diego, La Jolla, CA; http://vetneuromuscular. ucsd.edu/) and edrophonium chloride (Tensilon) testing is possible, albeit difficult to administer and interpret. Myasthenia gravis has been documented in a young ferret, but an association between megaesophagus and myasthenia was not reported.43 The management of ferrets with megaesophagus is similar to that of canine patients but is less successful. Supportive care and antibiotics are palliative at best. Administration of a GI motility enhancer such as metoclopramide (0.2 to 1 mg/kg PO or subcutaneously [SC] q6-8h) may be helpful. Cisapride, which until recently was marketed for gastroesophageal reflux and gastroparesis in humans, reduces the frequency of regurgitation in dogs with megaesophagus when given at 0.5 mg/kg PO q8-24h.80 This drug has been removed from the market for human use in the United States because of adverse cardiac effects in some people but is available through veterinary compounding pharmacies. Its use in ferrets has not been evaluated. If esophagitis is suspected, add an H2-receptor blocker, such as cimetidine, ranitidine, or famotidine. The prognosis for ferrets with megaesophagus is poor; generally, they die or are euthanized within days of diagnosis. Affected ferrets are debilitated and may suffer from malnutrition, hepatic lipidosis, and aspiration pneumonia.
Fig. 3-5 Lateral radiograph of a ferret with megaesophagus. Orally administered barium sulfate delineates the esophagus.
Other causes of esophageal disease in ferrets are rare. Esophageal foreign body has been reported in a ferret and was successfully managed surgically.12 One of the authors (HH) has seen a ferret with a sponge foreign body lodged in its distal esophagus; the sponge was broken into smaller pieces by using a 2.7-mm rigid endoscope. The foreign material then passed through the gastrointestinal tract without incident.
GASTRITIS AND ULCERATION Gastric and duodenal ulcers have been reported in laboratory ferrets and are relatively common in pet ferrets (Fig. 3-6). Causes of GI ulceration are foreign body or toxin ingestion, Helicobacter mustelae infection, neoplasia of the intestinal tract, treatment
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SECTION I Ferrets
Fig. 3-6 Ruptured duodenal ulcer with marked inflammation and hemorrhage in a ferret.
with nonsteroidal anti-inflammatory drugs (NSAIDs), and azotemia caused by renal disease. The laboratory ferret is used as an animal model for the study of H. pylori infection in humans. Helicobacter mustelae isolated from the gastric mucosa of ferrets shares many molecular and biochemical features of H. pylori. Infection with H. mustelae in ferrets is associated with varying degrees of gastritis, with or without duodenitis, and can result in ulcer formation (see discussion later in this chapter).26 Ulcerogenic drugs such as nonsteroidal and steroidal antiinflammatory agents can be associated with ulcer formation in many species. It is rare, however, for ferrets to have GI bleeding with corticosteroids, even at dosages as high as 2 to 3 mg/kg/day. Be careful with NSAIDs in ferrets; overdose of anti-inflammatory agents such as ibuprofen (see Chapter 5) can cause ulceration with prolonged or inappropriate use. Severe uremia and associated melena can develop in ferrets with primary renal disease, but this is uncommon. Gastritis in ferrets may be acute or chronic, or subclinical in some cases. Affected ferrets may hypersalivate, paw at the roof of the mouth, or display teeth-grinding, all of which indicate nausea and abdominal pain. Vomiting is not always reported, but owners sometimes describe “coughing” or gagging episodes that may actually represent gastric reflux. Some ferrets may not display obvious clinical signs, or signs may be missed if the ferret is part of a multi-ferret cage setup. Significant weight loss may be the only indication of a problem. Gastric or duodenal ulceration results in melena, anorexia, lethargy, and weight loss. Basic diagnostic testing includes whole-body radiography and screening blood tests. Fast the ferret for a short time (4 to 6 hours) to facilitate visualization of a gastric foreign body or hairball. The stomach should be empty, and any residual foodlike material may represent ingested hair or other material. The diagnosis of H. mustelae gastritis may be a diagnosis of exclusion of other common disorders, such as the presence of a GI foreign body. Treatment for H. mustelae gastritis is often based on a presumptive diagnosis. Treat gastritis and gastric ulceration with both specific therapy (according to the diagnosis) and supportive care. Hospitalize sick and anorexic ferrets for fluid therapy and parenteral treatment. Antibiotics are indicated for sick ferrets; consider choosing a combination that will target Helicobacter (e.g., amoxicillin and metronidazole in combination with bismuth or proton-pump inhibitor—see Helicobacter
treatment later in this chapter). For ferrets that are not vomiting, offer multiple small feedings of a bland, moist diet such as a/d Canine/Feline (Hill’s Pet Nutrition, Inc., Topeka, KS) or a recovery diet formulated for carnivores such as Carnivore Care (Oxbow Pet Products, Murdock, NE) or Emeraid Carnivore (LafeberVet.com, Cornell, IL). Avoid dry, high-fiber foods. For vomiting animals, withhold food for 6 to 8 hours while closely monitoring for any signs of hypoglycemia (older ferrets often have subclinical insulinomas); then, if vomiting has resolved, introduce small, frequent feedings. Bismuth compounds have action against pepsin, a proteolytic enzyme believed to be an important factor in the development of peptic ulcers. Administer bismuth subsalicylate at a dose of 1 mL/kg PO q8h. Sucralfate is a cytoprotective agent that binds to the erosion site and helps to form a protective barrier. It is a safe and useful adjunct to ulcer treatment and can be given orally in suspension form (100 mg/kg) every 6 hours. Systemic H2-receptor antagonists, such as ranitidine or famotidine, are often used to treat gastric ulcers because they block the histamine receptor on the gastric parietal cell and reduce gastric acid secretion. Famotidine is convenient to use because it is available for parenteral administration and has long dosing intervals (0.5 mg/kg PO, intravenously [IV], or SC q24h). The proton pump inhibitors, such as omeprazole, are occasionally used in ferrets. One quarter of the contents of a 10-mg capsule can be mixed with soft food and given orally.
HELICOBACTER MUSTELAE GASTRITIS Helicobacter mustelae is a gram-negative rod morphologically similar to Campylobacter species that requires a microaerobic environment for growth on artificial media. It is antigenically related and biochemically similar to H. pylori, a human pathogen associated with gastritis and ulcers.26 Virtually all North American ferrets are likely to be exposed to H. mustelae as kits, becoming persistently infected at weaning and developing some degree of gastritis.22 Colonization of the antral area of the stomach and pyloric area of the duodenum with H. mustelae is common in domestic ferrets, unless they are specifically treated or hand reared in isolation.17,22 Colonization is accompanied by a specific immune response, but infection persists despite high serum antibody titers.22 Although infection is common, clinical gastritis and ulcers occur relatively infrequently. Severe gastritis may be evident in gastric biopsy samples from ferrets showing no signs of clinical disease.22 The histopathologic lesions of H. mustelae–associated gastritis in ferrets, like H. pylori gastritis in humans, consist of mucus depletion, loss of glands, epithelial hyperplasia, foci of dysplasia, and leukocyte infiltration. The organism can be observed in silver-stained histologic sections of gastric mucosa.26 Severely affected ferrets that die usually have a single large pyloric ulcer or many small ones, and the stomach and intestinal tract contain digested blood and mucus, causing the ingesta to appear very dark. Cultures of H. mustelae from fecal samples are usually difficult to obtain, even when the organism is readily identified histologically or by culture in gastric biopsy samples. In humans, chronic infection with H. pylori leads to varying clinical and pathologic outcomes, including chronic gastritis, peptic ulcer disease, and gastric adenocarcinoma or mucosaassociated lymphoid tissue (MALT) lymphoma.61 The severity and distribution of the H. pylori–induced inflammation are
CHAPTER 3 Gastrointestinal Diseases key determinants of these outcomes.60 Gastritis involving the antrum is associated with excessive acid secretion and a high risk of duodenal ulcer. Gastritis involving the acid-secreting corpus region of the stomach is associated with hypochlorhydria, gastric atrophy, gastric ulcers, and increased risk of gastric cancer.32 As in humans infected with H. pylori, transient hypochlorhydria develops in ferrets approximately 4 weeks after experimental infection.53 This condition probably facilitates fecal-oral transmission as well as recovery of H. mustelae from feces.55 Infection is associated with urease produced in abundance by H. mustelae, which is detectable in gastric biopsy samples and can be used for presumptive diagnosis. Urease production correlates with the degree of colonization and the occurrence of gastritis in biopsy results.22 A urease breath test is available for humans to aid in diagnosis, and a similar test has been used in ferrets under research conditions but is not practical for clinical use.55 Gastric atrophy and hypochlorhydria are associated with the ability of H. mustelae to persistently colonize the stomach; hypochlorhydria due to loss of parietal cells allows non–ureaseproducing bacteria to colonize the stomach as well.22 Helicobacter species inhibit secretion of acid by parietal cells in vitro, and this mechanism may also contribute to hypochlorhydria.26 In humans, chronic infection with H. pylori is associated with release of cytokines that impair function of enterochromaffin cells, which are neuroendocrine cells in the gastric mucosa that control acid secretion by releasing histamine. The impaired secretory function of these cells may predispose to hypochlorhydria and gastric carcinogenesis. Loss of parietal cells due to chronic inflammation is, however, the primary cause of achlorhydria in humans and ferrets colonized with their respective Helicobacter species.67 Gastrin is a hormone that stimulates gastric acid secretion and is secreted by the G-cells of the gastric antrum. In humans and probably in ferrets, high levels of gastrin may initiate GI mucosal damage and ulceration. Hypergastrinemia is probably a response to the presence of H. mustelae in the antrum or to its associated inflammation. Hypergastrinemia is abolished after antibiotic therapy eradicates the Helicobacter infection.22 In humans, Helicobacter pylori–associated gastritis is a risk factor for gastric adenocarcinoma and gastric lymphoma, and H. pylori has been classified as a Class I carcinogen by the World Health Organization (WHO).22 There is some evidence that this is also the case in ferrets. As H. pylori does in humans,41 infection with H. mustelae in ferrets stimulates cell proliferation in the gastric mucosa.27 Under research conditions, gastric adenocarcinoma developed in aged ferrets that were naturally infected with H. mustelae and treated with a known gastric carcinogen.22 Spontaneously occurring gastric adenocarcinoma has been reported in pet ferrets. Although in some cases H. mustelae was neither cultured from the lesions nor identified histologically,69,76,78 in other cases, silver-stained organisms that were morphologically compatible with H. mustelae were present in the neoplastic tissues.27 Lymphoid follicles are observed in the gastric mucosa of humans colonized with H. pylori91 and in that of ferrets colonized with H. mustelae26 but not in uninfected individuals. In humans this condition may progress to MALT lymphoma. Eradicating H. pylori usually causes early tumors to regress, implicating the infection as the cause of neoplasia.90,91 Gastric MALT lymphoma associated with H. mustelae infection has also been reported in adult ferrets17 (Fig. 3-7). None of the affected ferrets
31
Fig. 3-7 MALT lymphoma in the stomach of a ferret infected with Helicobacter mustelae.
Fig. 3-8 Melena in a ferret with upper gastrointestinal bleeding from gastric ulcers.
were treated with antibiotics to eradicate H. mustelae either before or during the illness associated with neoplasia.
Clinical Signs and Diagnosis of H. Mustelae Gastritis with Ulcers Illness may develop in ferrets 12 to 20 weeks of age under conditions of stress caused by a combination of factors, such as rapid growth, dietary changes or inadequacy, and concurrent diseases. Infection is lifelong in untreated ferrets, and the severity of chronic gastritis increases with age.29 In mature ferrets, the disease may become clinically apparent in animals that are stressed by concurrent disease or by surgery for other conditions such as adrenal disease or insulinoma. Ferrets with severe H. mustelae gastritis and ulcers are lethargic and anorexic, and they rapidly become emaciated. Chronic vomiting may occur. Excessive salivation and pawing at the mouth, which are signs of nausea in ferrets, may be evident. Affected ferrets are often moderately to severely dehydrated and may have mild anemia and melena (Fig. 3-8). Black, tarry feces often stains the fur of the tail and perineal region. Definitive diagnosis of Helicobacter infection is confirmed by histopathologic examination of a gastric mucosal sample obtained by endoscopic or surgical biopsy. Gastric mucosa or fecal samples can be submitted for polymerase chain reaction (PCR)-based analysis (Taconic Rockville, Rockville, MD, www.taconic.com; Veterinary
32
SECTION I Ferrets Table 3-1 Summary of Suggested Treatment Regimens for Helicobacter mustelae Gastritis, Inflammatory Bowel Disease, Proliferative Bowel Disease, and Eosinophilic Gastroenteritis Disease Helicobacter mustelae gastritis Original triple therapya
Alternative therapya
H2 receptor blocker
Drug
Dosage
Amoxicillin Metronidazole Bismuth subsalicylate
10 mg/kg PO q12h 20 mg/kg PO q12h 17 mg/kg (1 mL/kg) PO q12h
Clarithromycin Ranitidine bismuth citrate or Clarithromycin Omeprazole Metronidazole Famotidine
12.5 mg/kg PO q8h36 24 mg/kg PO q8h36 50 mg/kg PO q24h1 4 mg/kg PO q24h1 75 mg/kg PO q24h1 0.5-1 mg/kg PO, SC q24h
Inflammatory bowel disease
Azathioprine Prednisone Sucralfate Hypoallergenic diet
0.9 mg/kg PO q24-72h 1 mg/kg PO q24h 100 mg/kg PO q6h
Proliferative bowel disease
Chloramphenicol
50 mg/kg PO, IM, SC q12h
Eosinophilic gastroenteritis
Prednisone Ivermectin
1.25-2.5 mg/kg PO q24h 0.4 mg/kg SC, PO once; repeat in 14 days
IM, Intramuscular; PO, per os; SC, subcutaneous. Chloramphenicol, ranitidine bismuth citrate, azathioprine, and metronidazole can be prepared as suspensions by compounding pharmacists. aTreat for a minimum of 14 days.
Molecular Diagnostics, Milford, OH, www.vmdlabs.com). Specialized techniques are necessary for culturing the organism, which is not shed consistently in feces of infected ferrets.31
Treatment of H. Mustelae–Associated Gastritis with Ulcers Treatment for Helicobacter infection in humans usually consists of “triple therapy” consisting of two antibiotics from different classes, such as amoxicillin or metronidazole and clarithromycin, and a proton pump inhibitor. Differing combinations of antibiotics and proton pump inhibitors are used, with varying success. In humans with resistant Helicobacter infections, “quadruple therapy” with an added bismuth compound is often used. Bismuth interferes with the colonization of H. pylori in humans and suppresses colonization of H. mustelae in ferrets.77 Bismuth also has direct antimicrobial actions against Helicobacter.64 In ferrets, the initial treatment for Helicobacter species in ferrets is commonly a triple-therapy combination of amoxicillin, metronidazole, and bismuth subsalicylate, administered q12h for at least 2 weeks (Table 3-1). Oral veterinary or pediatric amoxicillin suspensions are palatable and well accepted by most ferrets; metronidazole can be compounded into a suspension for oral administration. Although H. mustelae is sensitive to either amoxicillin or metronidazole, treatment with one of these antibiotics alone or single treatment with other antibiotics is ineffective for eradication therapy. Other drug combinations have been used in ferrets to eradicate H. mustelae, with advantages of improved palatability and convenience of dosing. Clarithromycin (Biaxin, Abbott Laboratories, North Chicago, IL) (12.5 mg/kg PO q8h) in combination with ranitidine
bismuth (24 mg/kg q8h) has been shown to eradicate H. mustelae in ferrets.53 Ranitidine bismuth citrate tablets (not available in the United States) may be crushed and mixed with a palatable liquid or compounded, and clarithromycin is available as a pediatric suspension. Both drugs are administered for 14 to 21 days (dosages are given in Table 3-1). A combination of clarithromycin, metronidazole, and omeprazole or clarithromycin and omeprazole proved more effective than triple therapy with amoxicillin, metronidazole, and omeprazole in eradicating H. mustelae in research ferrets.2 Resistance to clarithromycin has not yet been reported in ferrets but does occur in humans.53 To prevent development of macrolide-resistant strains, clarithromycin should be combined with a second antibiotic not in the macrolide class.2 Chloramphenicol has no effect on H. mustelae.61 Colloidal bismuth subcitrate (8 mg/kg PO q8h) may be substituted for bismuth subsalicylate. Antacid therapy may not be helpful in the early treatment of Helicobacter infection because affected ferrets usually develop transient hypochlorhydria.31 However, once affected with gastritis and inappetence, antacids can decrease discomfort, improve appetite, and reduce effects of acid reflux on esophageal mucosa. Famotidine (0.50-1.0 mg/kg PO q24h) or other H2-receptor blockers, sucralfate suspension (25 to 100 mg/kg PO q8h), or a proton-pump inhibitor such as omeprazole (4 mg/kg PO q24h) (if not used in triple therapy) may be helpful in very sick animals that are bleeding from extensive gastric ulcers. Although eradication of H. mustelae is accompanied by decreasing antibody titers, lesions may take longer to resolve.53 Successfully treated ferrets can be reinfected with H. mustelae through contact with infected ferrets.4 For treated ferrets to
CHAPTER 3 Gastrointestinal Diseases
A
33
B Fig. 3-9 Lateral (A) and ventrodorsal (B) radiographic views of a ferret with a pyloric outflow obstruction from a foreign body. The stomach is greatly distended with fluid and gas.
remain free of H. mustelae, they should not be exposed to ferrets of unknown Helicobacter infection status until the newly introduced ferrets have also been treated.
GASTROINTESTINAL POLYPS Two ferrets with GI polyps have been seen at the Animal Medical Center (New York, NY). Both ferrets showed lethargy, inappetence, melena, and weakness from anemia. Abdominal radiographs suggested GI abnormalities. On surgical abdominal exploration, one ferret had a gastric polyp and the other had a small intestinal polyp. Both ferrets did well after surgical resection of the polyps, which were histologically benign.
GASTRIC DISTENTION (BLOAT) Pet ferrets occasionally are seen with an acute gastric or small intestinal foreign body blockage that results in a distended, fluidfilled stomach. These ferrets are acutely very weak, reluctant to ambulate, and anorexic. To confirm the diagnosis, take full-body radiographs (Fig. 3-9). These animals are in shock and need immediate aggressive therapy that includes intravenous fluids and decompression. Relieve gastric pressure by placing an orogastric tube (8- or 10-French red rubber feeding tube), treat the hypovolemic shock, and prepare these cases for surgery when stable. Pyloric stenosis and gastric outflow obstruction can manifest as an acute bloat in the ferret. Pyloric adenocarcinoma in ferrets and in humans has been associated with infection with H. mustelae.69,76 Pyloric stenosis caused by muscular hypertrophy of the pylorus has been seen clinically by one author (HH), with one case occurring in a 4-month-old ferret. Pyloromyotomy and dilation of the pyloric outflow is the recommended treatment, especially where Helicobacter and cancer are not present. Other than associated with foreign body disease, gastric bloat is rarely seen in pet ferrets, but it has been reported on
domestic ferret farms and in black-footed ferrets (Mustela nigripes).21,73 Clinical signs are usually observed in weanling ferrets and include acute gastric distention, dyspnea, and cyanosis. Sudden death can occur. The cause is unknown but is thought to be related to an overgrowth of Clostridium perfringens (previously called C. welchii). Certain conditions may predispose to clostridial overgrowth, including increased concentration of carbohydrates in the GI tract from overeating, dietary changes, and intestinal hypomotility. Clostridium perfringens multiplies rapidly, producing enterotoxins that attack the villous epithelial cells of the gut. Gas production by the bacteria results in abdominal distention.
GASTROINTESTINAL FOREIGN BODIES Gastrointestinal foreign bodies are common in ferrets.58,84 Ferrets are naturally inquisitive and like to chew on miscellaneous environmental objects, particularly rubber or sponge products. Rubber or foam foreign bodies are most commonly ingested by young ferrets (younger than 2 years of age); in contrast, trichobezoars (hair balls) are more common in older ferrets (Fig. 3-10). Linear foreign bodies, commonly ingested by cats, are very rare in ferrets. The most common clinical signs of a GI foreign body in ferrets are lethargy, inappetence or anorexia, and diarrhea. Vomiting is sometimes not reported by the owner. However, if vomiting is observed, consider a GI foreign body (Table 3-2). Some ferrets display signs of nausea, including bruxism, ptyalism, and face rubbing. Weakness can be profound in acutely obstructed animals; some of these ferrets are recumbent and reluctant to ambulate. Trichobezoars can manifest as chronic, intermittent gastric problems: bruxism, weight loss, inappetence, and melena may be reported. Some of these cases can become acute outflow obstructions, whereas in other cases, a more subtle, chronic history can be seen, depending on where the hairball is located in the stomach.
SECTION I Ferrets
34
If a GI foreign body is suspected, palpate the abdomen carefully. Foreign bodies in the small intestine often are associated with localized discomfort or pain and can usually be palpated, especially under sedation. Gastric foreign bodies are more difficult to palpate because the stomach is under the ribcage. Hold the sedated ferret vertically from the axillae to allow the spleen with the attached stomach to drop down for easier palpation. Hairballs tend to be firm, tubular structures (see Fig. 3-10) and can often be palpated in the stomach through the gastric wall.
Fig. 3-10 Trichobezoars surgically removed from a ferret stomach.
If a foreign body is suspected, take whole-body survey radiographs. Make sure to include the thorax to evaluate the esophagus. Abnormal abdominal radiographic findings are segmental ileus, gaseous distention of the stomach, and, occasionally, a visible foreign object or trichobezoar (Fig. 3-11). Contrast (barium or negative gastrogram with air) studies can be done if needed and can be helpful to determine the presence of hairballs84 (Fig. 3-12). Base the diagnosis on history, clinical signs, palpation, and the radiographic results. At a minimum, submit a blood sample for a complete blood count (CBC) and plasma biochemical analysis in a ferret that is sick for longer than 24 hours. Ferrets rarely pass GI foreign bodies unassisted. Occasionally, a small, partially obstructing object may pass after treatment with intestinal lubricants (hairball laxatives) q8h and replacement fluids. However, most GI foreign bodies must be removed surgically (see Chapter 11). Stabilize debilitated ferrets before surgery. Parenteral fluids are essential because these ferrets are usually dehydrated. If the ferret is stable, alert, and ambulatory, fluids can be given subcutaneously until prepping for surgery; then, place an IV catheter and administer fluids intravenously. Perform an exploratory laparotomy as soon as possible. Collect biopsy specimens as needed from the stomach or intestines if ulcerated or abnormal in appearance. Some ferrets also may have H. mustelae–associated gastritis or GI lymphoma. Check the adrenal glands and the pancreas in older ferrets; discovery of concurrent abdominal disease is not unusual during surgery. In
Table 3-2 Differentiation of Common Gastrointestinal Diseases That Cause Weight Loss and Diarrhea in Ferrets by Typical History, Clinical Findings, and Laboratory and Radiographic Resultsa Typical Diarrhea
Vomiting/ Bruxism
Prolapsed Rectum/ Tenesmus
Eosinophilic gastroenteritis
Mucoid, green
Possible
No
± Thickened intestinal loops
Ferret enteric coronavirus/ (Epizootic catarrhal enteritis) Foreign body
Acute: profuse, Possible green Chronic: grainy (“bird seed”)
No
Thickened or fluidfilled intestines
Black, tarry or mucoid, green
Yes
No
Palpable gastric or intestinal gas Painful abdominal palpation
Helicobacter mustelae gastritis
Black, tarry or mucoid, green
Yes
No
Inflammatory bowel disease
Mucoid, green
Possible
No
Proliferative bowel disease
Mucoid, green
Rare
Yes
Disease
aAlthough
Physical Findings
Laboratory/ Radiographic Results
Comments
Eosinophilia Rare; multiple tissue ± Reactive hepatitis involvement (visceral lymph nodes, spleen) ± Reactive hepatitis Acute onset, can become chronic; exposure to new or young ferrets
± Reactive hepatitis Acute or chronic Anemia (chronic) Young ferrets—rubber Gas in stomach or objects, toys intestinal loops Older ferrets— hairballs more common Enlarged mesenteric ± Reactive hepatitis Recent stress (i.e., lymph nodes ± Anemia surgery); can Gas in stomach increase in severity with age ± Enlarged Reactive hepatitis May develop mesenteric lymph ± High globulins secondary to other nodes ± High lipase G1 disease Palpably thickened ± Globulins Primarily affects large bowel; young ferrets proliferative rectal mucosa
typical findings are listed, clinical signs and physical findings are variable in any of the described diseases.
CHAPTER 3 Gastrointestinal Diseases most cases, recovery is rapid after GI foreign body removal, and ferrets are able to eat soft foods within 12 hours after surgery. Most ferrets can be discharged within 48 hours after surgery. Prevention of foreign body obstructions includes recommending the regular use of a hairball laxative preparation during active shedding seasons and “ferret-proofing” the household. Ferrets should not be left uncaged or unsupervised. Advise owners to avoid giving small rubber “squeak” toys to pet ferrets
and remove soft rubber objects (e.g., rubber-soled shoes and earphones) from the ferret’s accessible environment.
LIVER DISEASE Lymphoma is the most common hepatic neoplasm seen in ferrets. Other reported hepatic neoplasms include hemangiosarcoma, adenocarcinoma, hepatoma, biliary cystadenomas, and
B
A
Fig. 3-11 Lateral (A) and ventrodorsal (B) radiographic views of a ferret with a gastrointestinal foreign body. The proximal small intestine is markedly dilated with fluid and gas, compatible with segmental ileus. The foreign body is not visualized.
A
B
35
C
Fig. 3-12 A, Ventrodorsal radiograph of a ferret with a gastric trichobezoar. The trichobezoar appears as a tubular soft tissue density poorly delineated in the stomach. Contrast studies were performed. B, lateral and C, ventrodorsal radiographic views of the same ferret 30 minutes after administration of barium sulfate. Note the delineation of at least two trichobezoars.
36
SECTION I Ferrets
hepatocellular adenoma.16,37,47 Pancreatic islet cell tumors can metastasize to the liver. Prognosis is guarded regardless of tumor type. Other than neoplastic diseases, primary hepatopathies are uncommon in ferrets. Vascular shunts have not been reported. Cholelithiasis and cholestasis have been described in ferrets and may be underdiagnosed in clinical practice.37a,40a Hepatic lipidosis can be found in association with long-term anorexia. Chronic GI diseases (e.g., trichobezoar formation) can lead to hepatic lipidosis. Foreign bodies in the proximal duodenum can obstruct the bile ducts as they enter the small intestine and cause increased liver enzyme and bilirubin levels. Steroid hepatopathy is rare in ferrets, even with long-term steroid administration or hyperadrenocorticism. Chronic-lymphocytic portal hepatitis has been found on histologic examination of hepatic biopsy samples. The cause is unknown but may be related to chronic visceral inflammation such as inflammatory bowel disease. Chronic cholangiohepatitis with biliary hyperplasia of variable intensity was reported in 8 of 34 cohabitating ferrets.34 Three ferrets had neoplastic lesions in the liver. Spiral-shaped bacteria were identified in the livers of three ferrets, and bacteria with 97% similarity to Helicobacter species were identified by PCR in the feces of one ferret. Because of the clustering of cases and the pathologic findings, a possible infectious cause was suggested. Copper toxicosis was diagnosed in two sibling ferrets on the basis of high hepatic copper concentrations and histologic changes in hepatic tissue.33 Clinical signs in these two ferrets were mostly nonspecific and included severe central nervous system depression with hypothermia and hyperthermia, respectively. One ferret was icteric. Both ferrets died within a few days of clinical evaluation despite supportive care. A genetic predisposition to copper toxicosis in these two ferrets was proposed because they were siblings with the same phenotypic coat color and because no environmental source of copper could be identified. In most ferrets with liver disease, a high concentration of alanine aminotransferase (ALT greater than 275 IU/L) is present on plasma biochemical analysis. Alkaline phosphatase concentration is sometimes increased. High total bilirubin levels are uncommon, and ferrets are rarely icteric. Be careful in diagnosing liver disease in ferrets with high enzyme concentrations; these laboratory findings are also common in ferrets with intestinal disease. Base the diagnosis of liver disease on the presence of persistently high liver enzyme concentrations, radiographic and ultrasound findings, and, for definitive diagnosis, analysis of liver biopsy samples. Ultrasound-guided needle biopsy of the liver is possible, but a full abdominal exploratory is often recommended because of the likelihood of concomitant disease in ferrets.
GASTROINTESTINAL PARASITISM Intestinal parasites are uncommon in ferrets. However, in any ferret with diarrhea, perform a complete fecal parasite check, including a direct fresh wet mount and fecal flotation. In juvenile ferrets, nematodiasis is rare, but coccidiosis and giardiasis are occasionally seen. Coccidiosis can be subclinical in ferrets or it may be associated with diarrhea, lethargy, and dehydration.9 Rectal prolapse is possible. Diagnose coccidiosis by results of fecal testing, either direct wet mount and microscopic examination or fecal flotation. Follow the same treatment protocols as for canine and feline patients with coccidiosis.
Cryptosporidiosis is described in ferrets but may not result in clinical disease.1,68 Young ferrets can have a subclinical infection with Crytosporidium parvum that can persist for several weeks. The oocysts can be shed in the feces of clinically normal ferrets. Histologically, the organism may be associated with an eosinophilic infiltrate in the lamina propria of the small intestine. The zoonotic potential of the ferret genotype of C. parvum to infect humans is unknown1; however, if oocysts are detected, discuss the potential of transmission to immunocompromised owners.
ENTERITIS AND DIARRHEA SALMONELLOSIS Salmonellosis is a contagious disease characterized by fever, bloody diarrhea, and lethargy. Conjunctivitis and anemia also may be present. Salmonella newport, S. typhimurium, and S. choleraesuis may be involved.52 The incidence of salmonellosis in pet ferrets is very low and the infection may be associated with feeding of raw or undercooked meat, poultry, and meat by-products. Isolation of Salmonella organisms usually requires collecting multiple fecal samples and the use of selective media. Treatment consists of aggressive supportive care, antimicrobials, and treatment for shock as needed.
MYCOBACTERIOSIS Ferrets can be naturally or experimentally infected by several species of mycobacteria.10,20,74 Mycobacterium bovis and M. avium infections have been recognized in research, farm, and feral ferrets in England, Europe, and New Zealand. These infections have been associated with the feeding of raw meat and poultry and unpasteurized dairy products, or, in feral ferrets, feeding on carrion infected with M. bovis.50 Infections with M. avium, M. genavense, M. abscesses, and M. celatum have been reported in domestic ferrets.48,49,65,71,74,82 Ferrets with mycobacterial infections can have a variety of clinical symptoms, which may include weight loss, lymph node enlargement, conjunctival lesions, splenomegaly, and pneumonia. At necropsy, granulomas with acid-fast organisms can be found in the liver, lungs, lymph nodes, intestines, stomach, and trachea. Mycobacteriosis is diagnosed by results of tissue biopsy, including histopathologic examination with acid-fast staining, polymerase chain reaction (PCR) testing, and culture. In several reported cases, treatment regimens usually involving rifampicin alone or in combination with enrofloxacin and azithromycin have been used with variable success. The zoonotic potential of infection is unknown but should be discussed with owners before beginning treatment in any ferret diagnosed with mycobacteriosis.
CAMPYLOBACTERIOSIS Campylobacter jejuni is a bacterial enteric pathogen that is associated with diarrhea and enterocolitis in humans and many animal species, including dogs, cats, calves, and sheep. Campylobacter jejuni can be isolated from the feces of normal ferrets, and during the 1980s, it was suspected to be the cause of proliferative colitis, or proliferative bowel disease (PBD), in ferrets.24,25 However, inoculation of C. jejuni into 54 conventionally reared
CHAPTER 3 Gastrointestinal Diseases
37
1 mg/kg q12h for 14 days) and changing the diet to an easily absorbed food may speed recovery. More recently, a ferret systemic coronavirus (FSCV) has been identified as the causative agent of a progressive systemic pyogranulomatous disease in ferrets that resembles the dry form of feline infectious peritonitis (FIP).35,54,63 Affected ferrets exhibit weight loss, palpable abdominal mass or masses, diarrhea, hypergammaglobulinemia, anemia, and sometimes central nervous system signs. The disease is progressive and carries a high mortality rate, with the duration of clinical illness averaging 67 days.35 Partial gene sequencing indicates that the ferret systemic and enteric coronaviruses are closely related but not identical, and that FSCV is more closely related to FECV than to other group 1 coronaviurses.35,87 There is no treatment for this form of coronavirus in ferrets. Similar to FIP in cats, this disease carries a poor prognosis.
Rotavirus Fig. 3-13 Grainy loose feces in a ferret with chronic diarrhea, consistent with infection ferret enteric coronavirus (epizootic catarrhal enteritis).
and 2 gnotobiotic ferrets caused diarrhea but not the full spectrum of clinical signs and histopathologic lesions seen in PBD (see later discussion).6 The Campylobacter-like organism that causes of PBD was subsequently thought to be a Desulfovibrio species,28 but is now known to be Lawsonia intracellularis, the agent that causes porcine proliferative enteropathy (see later discussion).20 The importance of C. jejuni as a primary pathogen in pet ferrets is not known.
VIRAL DIARRHEA Coronavirus Epizootic catarrhal enteritis (ECE) is a highly transmissible diarrheal disease of ferrets that first appeared in 1993 in several rescue and breeder operations in the eastern United States. The causative agent of ECE is now attributed to a coronavirus.86,88 In intestinal biopsy samples of ferrets infected with ferret enteric coronovirus (FECV), histologic findings include lymphocytic enteritis, villous atrophy, and blunting or degeneration of apical epithelium. Ferrets infected with FECV initially develop a profuse, green mucoid diarrhea that may progress to a loose, grainy stool resembling birdseed (Fig. 3-13). Adult ferrets are most susceptible to ECE, and the typical history includes recent exposure to a new, young ferret that acts as an asymptomatic carrier. The incubation period is 48 to 72 hours, and affected ferrets are anorexic and lethargic. Coronavirus may be detected by PCR testing of fecal or intestinal tissue (jejunum and ileum) samples of affected ferrets (Diagnostic Center for Population and Animal Health, Michigan State University, Lansing, MI, www.dcpah.msu.edu; Veterinary Molecular Diagnostics, Milford, OH, www.vmdlabs.com). Treat sick ferrets with signs of ECE with aggressive fluid therapy, antibiotics, and supportive care, and isolate these ferrets from asymptomatic or unexposed ferrets. Although the morbidity rate can be high, the mortality rate is low in ferrets that are treated appropriately. After recovering from ECE, some adult ferrets develop a persistent, intermittent malabsorption syndrome with diarrhea. The clinical course can be prolonged in these ferrets, lasting weeks to months. Treatment with a short course of steroids (prednisone
Infection with rotavirus causes diarrhea in very young ferrets. Farm outbreaks of diarrhea are associated with high morbidity and mortality rates in neonatal kits from 2 to 6 weeks of age.7,81 In a report of an outbreak of diarrhea in 1-week-old ferrets, a group C rotavirus was identified and appears to be highly prevalent.89 The morbidity is low in adult ferrets, but infection may result in a transient, green, mucoid diarrhea. The virus is shed in the stool, and transmission is by contact with infected animals or the environment. Diagnosis is by PCR testing of fecal or tissue samples (jejunum or ileum) (Diagnostic Center for Population and Animal Health, Michigan State University, Lansing, MI; www.animalhealth.msu.edu). Treatment is supportive with fluids and antibiotics.
Canine Distemper Virus Distemper is caused by a highly contagious paramyxovirus that causes fatal disease in unvaccinated ferrets. Clinical signs can vary but often include diarrhea in conjunction with nasal and ocular discharges and a generalized orange-tinged dermatitis (see Chapter 6). Diarrhea may be acute or intermittent. The widespread practice of vaccinating ferrets against canine distemper virus has greatly limited the occurrence of distemper and it is now a rare disease in ferrets. Cold-like symptoms and diarrhea in newly purchased, unvaccinated ferrets should arouse suspicion. Distemper virus is generally considered incurable and fatal in ferrets, although vitamin A has been shown to have promising antiviral properties when administered as a supplement (30 mg) to experimentally infected ferrets.70
Influenza Virus Ferrets infected with influenza (an orthomyxovirus) sometimes have transient diarrhea. The virus also causes upper respiratory disease associated with coughing, sneezing, inappetence, and lethargy. Affected ferrets are often febrile (see Chapter 6).
INFLAMMATORY BOWEL DISEASE AND EOSINOPHILIC GASTROENTERITIS Inflammatory bowel disease (IBD) is a relatively common cause of gastroenteritis in ferrets.11 The cause is unknown but may be related to dietary intolerance, hypersensitivity reaction, or another aberrant immune response. The inflammation typically is lymphoplasmacytic and should be distinguished from eosinophilic gastroenteritis, which often involves multiple tissues.
38
SECTION I Ferrets
This condition is easily overlooked in ferrets because it resembles viral diarrhea (ECE), dietary indiscretion, and Helicobacterassociated gastroenteritis. Affected ferrets can have loose grainy stools, intermittent nausea, occasional vomiting, and weight loss. Clinical signs can be subtle and chronic, or some ferrets can have acute vomiting and lethargy. Ferrets with IBD are usually young or middle-aged adults and in multiple-ferret households; typically only one ferret in the household is affected. Results of blood tests may reveal an increase in liver enzyme activities and plasma globulin concentrations, and lymphocytosis is occasionally present. In some ferrets, laboratory results are unremarkable. Diagnosis is based on clinical signs, a detailed clinical history that eliminates the possibility of exposure to coronavirus, and results of diagnostic tests such as radiographs and routine blood tests. Diseases such as Helicobacter gastroenteritis and intestinal lymphoma should be ruled out. Definitive diagnosis can only be made by histologic examination of full-thickness gastric and intestinal biopsy samples. Treating without a histologic diagnosis is common because of the risks and costs of an abdominal exploratory procedure to collect biopsy samples. Therapy is aimed at suppressing the immune response and dietary management. Corticosteroids such as prednisone (1 mg/kg PO q12–24h) are often used, but some ferrets with inflammatory bowel disease respond poorly to long-term treatment with steroids. Azathioprine (Imuran, Prometheus Laboratories, San Diego, CA) (0.9 mg/kg PO q24–72h) is another treatment option and seems to be well tolerated in ferrets.11 Hypoallergenic diets made for cats (z/d feline; Hill’s Pet Nutrition, Topeka, KS) can be tried, but ferrets are often reluctant to make dietary conversions. A chicken-free diet formulated for ferrets is available (Totally Ferret Turkey-VenisonLamb Meal Formula, Performance Foods, Broomfield, CO; www.totallyferret.com) but efficacy for IBD is unknown. A grain-free ferret diet is also available (ZuPreem, Shawnee, KS; www.zupreem.com). Eosinophilic gastroenteritis is a rare type of inflammatory bowel disease that occurs in ferrets and has also been reported in dogs, cats, horses, and humans.3,83,92 In all reported cases in ferrets, animals were older than 6 months of age; however, because of the small number of reports, the incidence of disease in young animals is not known. No specific causative agent has been found in ferrets,13,18,62 dogs,75 or humans,79 but food allergy is implicated in most humans and in some dogs. In specific cases in humans and in other species, clinical signs were relieved when appropriate treatment for food allergies or parasitism was instituted. Peripheral eosinophilia is a common but not a constant finding in affected dogs and humans,75 but it has been reported in most of the relatively few ferrets diagnosed with this disease.62 Eosinophilia in ferrets, however, is highly suggestive of the disease. No reports of food elimination tests in affected ferrets have been published. The lesion of eosinophilic gastritis in ferrets, as in other animals and humans, is a mild to extensive infiltration of the mucosa, submucosa, and muscularis of the stomach and small intestine with eosinophils. Focal eosinophilic granulomas may be found in the mesenteric lymph nodes or abdominal organs of affected ferrets.62 No pathogens have been observed in or isolated from the lesions of affected ferrets. In humans and other affected species, granulomas may cause partial bowel obstruction. Affected animals typically have chronic diarrhea, with or without mucus and blood, and severe weight loss. Granulomas
and a thickened lower bowel may be palpable. Vomiting, anorexia, and dehydration are variable signs. Signs may be clinically indistinguishable from those of gastritis, persistent ECE, or GI obstruction by a foreign body. Humans, dogs, and cats with eosinophilic gastroenteritis usually respond to steroid treatment. Because the disease in ferrets resembles that in other species, prednisone administration has been the treatment of choice.62 Remission has occurred in ferrets treated with prednisone (1.25 to 2.5 mg/kg PO q24h for 7 days and q48h thereafter) until the ferret is clinically normal. Immediate recovery also followed removal of an enlarged mesenteric lymph node in one ferret and treatment with ivermectin (0.4 mg/kg SC) in another.62 When eosinophilic gastroenteritis is a response to the presence of parasites, eliminating the parasites is preferable to prolonged treatment with corticosteroids to relieve clinical signs. In a recent report of a case series of dogs with gastrointestinal masses composed primarily of eosinophilic infiltrates, an underlying cause was not ascertained. Interestingly however, most dogs that were treated with corticosteroids and ivermectin improved clinically, with resolution of the eosinophilic infiltrates and prolonged survival. In contrast, all dogs treated surgically to remove the eosinophilic masses died of complications of their disease.51
PROLIFERATIVE BOWEL DISEASE Proliferative bowel disease, which has been recognized for decades in pigs and hamsters, was first diagnosed in ferrets in 1982. The cause in swine is a bacterium classified as Lawsonia intracellularis.56 This same agent causes PBD in hamsters and in ferrets23 and has more recently been implicated in proliferative enteropathies of other species, including rabbits,72 white-tailed deer, ratite birds, and domestic foals.15 Lawsonia intracellularis is an obligate intracellular organism that cannot be propagated on artificial media but can be grown in embryonated chicken eggs. Two tests that detect this organism have been developed and are used in ferret tissues under research conditions: a PCR test specific for the swine isolate, and an indirect fluorescent antibody test that identifies the omega antigen common to organisms found in PBD lesions of swine, hamsters, and ferrets.23 However, diagnosis of clinical cases usually depends on observing clinical signs and gross or histopathologic lesions. Areas of intestine affected by PBD can be palpated, appear grossly thickened, and are often discolored on the serosal surface. The colon and less commonly the small intestine may be involved. Ridges of proliferative tissue, distinct from adjacent normal tissue, are obvious on the mucosal surface (Fig. 3-14). Occasionally the affected bowel perforates and causes fatal peritonitis. On histologic examination, epithelial proliferation with hypertrophy of the muscularis and infiltration of the bowel wall with either monocytic or granulocytic inflammatory cells, or both, are present.30 In silver-stained sections, commashaped organisms can be found inside enterocytes lining crypts or glands. The normal architectural pattern of the mucosa is lost. Normally, straight tubular glands are covered evenly with enterocytes and numerous goblet cells. In PBD, the irregular, branching, proliferative glands lack goblet cells, and necrotic debris accumulates in the crypts. Severe glandular hyperplasia resembles neoplasia and may translocate to liver and regional lymph nodes.23
CHAPTER 3 Gastrointestinal Diseases
39
Fig. 3-14 Mucosal surface of a ferret with proliferative bowel disease. The mucosa appears thickened and hemorrhagic.
CLINICAL SIGNS AND DIAGNOSIS OF PROLIFERATIVE BOWEL DISEASE Proliferative bowel disease occurs most frequently in rapidly growing juveniles, 10 to 16 weeks of age. Environmental and nutritional stress factors appear to play a role in resistance of infected animals to clinical disease. Lawsonia intracellularis is probably transmitted by the oral-fecal route,23 and all ferrets that are housed in groups presumably will be equally exposed to the agent. However, clinical disease develops in only a small percentage (usually 1% to 3%) of group-housed juvenile ferrets. Improvements in the quality of care and nutrition of pet ferrets may be responsible for the apparently decreasing incidence of PBD in recent years. Affected ferrets have chronic diarrhea that may vary from dark, liquid feces streaked with bright red blood to scant, mucoid stool, often with bright green mucus. The fur of the tail and perineal area may be stained and wet with fecal material, and the preputial area of males is often wet with urine. Rectal tissue may continuously or intermittently prolapse (Fig. 3-15). Affected animals moan or cry while straining. Some continue to eat but lose weight at an alarming rate. If not appropriately treated, a ferret weighing 800 g may lose 400 g in less than 2 weeks. These animals are moderately to severely dehydrated and may be hypoalbuminemic. They are weak and sleep most of the time. Because of their general debility, ferrets with PBD are more susceptible to other infectious diseases. They may have upper respiratory tract infections that do not affect other healthy ferrets housed with them and often develop clinical gastritis or ulcers. Severely affected animals will die if not treated appropriately, and most of those that die despite treatment have proliferative ileitis alone or in combination with colitis. Diagnosis is based on clinical signs, history, and response to treatment. A PCR assay is available for fecal, rectal swab, or ileal biopsy tissue samples (Zoologix, Inc., Chatsworth, CA; www.zoologix.com).
TREATMENT OF PROLIFERATIVE BOWEL DISEASE Lawsonia intracellularis is sensitive to chloramphenicol. No other antibiotic consistently resolves PBD in ferrets. Chloramphenicol is administered at a dose of 50 mg/kg q12h IM or SC
Fig. 3-15 Prolapsed rectum in a young ferret. (chloramphenicol sodium succinate) or orally (chloramphenicol palmitate oral suspension) for at least 10 days.45 A ferret with colitis of recent onset improves quickly with this treatment and gains 50 to 100 g/day within a few days of the first dose. Although the organism is also sensitive to tylosin, tetracyclines, tiamulin, and several other antimicrobials that are used to treat PBD in pigs,56 and to erythromycin, which is commonly used in affected foals,46 treatment of ferrets with any of these drugs is disappointing. Treatment of infected but clinically normal 6to 9-week-old ferrets with oral tylosin (5 mg/kg mixed in soft food once daily) appears to reduce the incidence of clinical PBD in a colony, but only chloramphenicol produces a dramatic improvement in sick ferrets. Repair of rectal prolapse with a purse-string suture is rarely necessary because as the colon heals, the prolapse usually disappears spontaneously. It may appear intermittently for weeks but causes no apparent distress. If a purse-string suture is used, the owner must closely monitor the ferret to make sure that it can defecate, especially when the stool regains its normal consistency. Sutures should be removed in 2 to 3 days.
RECTAL DISEASE Rectal prolapse can occur in ferrets. It is most often associated with diarrhea and is usually a disease of young ferrets. Possible causes include coccidiosis, PBD, colitis, and neoplasia. Some young ferrets protrude the rectal mucosa from poor surgical analsacculectomy technique. Straining from adrenal-associated prostatic disease, urinary outflow obstruction, or an enlarged sublumbar lymph node (lymphosarcoma) may result in protrusion of the rectum. Anal gland impactions or abscesses are rare in ferrets that have been surgically descented. Neoplasia is rare in the rectal area, with one recent report of anal sac apocrine adenocarcinoma.59 One author (HH) has seen a descented ferret with leiomyosarcoma that surrounded the rectal opening. The ferret presented for a rectal prolapse, and a tumor was found on palpation. Treatment of rectal neoplasia involves surgical debulking, possible rectoplasty, and possible localized radiation therapy. Prognosis is poor. Include a careful rectal examination (visualization and palpation) in all physical examinations in ferrets. Undescented ferrets may develop anal gland disease, including impactions and
40
SECTION I Ferrets
abscessation. Palpation of the anal area may reveal either unilateral or 360-degree perianal swelling. Manage anal gland disease as in dogs. Be forewarned: anal gland odor is quite noxious. Anal gland removal is described in Chapter 11. Diagnostic tests should include radiographs in the ferrets that present for straining, and a fecal wet mount and flotation to check for parasites. Other than coccidiosis, GI parasitism is uncommon. Rectal prolapse often resolves with treatment of the causative condition. Treat with antibiotics and antiparasitic agents, as indicated. Although rarely needed, rectal purse-string sutures can be placed if the prolapse is extensive; these sutures can be left in place for 2 to 3 days.
NEOPLASIA The GI tract is not an uncommon site of primary neoplasia in ferrets. Squamous cell carcinoma of the oral cavity manifests as locally aggressive tumors usually involving the jaw bone.36,85 Wide surgical resection is the only treatment, and radiation therapy has been applied. Pyloric adenocarcinoma has been reported,69,76 and may be related to chronic gastritis induced by Helicobacter infection. Intestinal adenocarcinoma has been seen clinically in several ferrets by one author (HH). Clinically, these cases presented for nonresponsive diarrhea and signs of intestinal obstruction or rupture. In all cases, adenocarcinoma was rapidly progressive and fatal. Lymphoma frequently affects the GI tract of ferrets. Visceral and mesenteric lymph nodes, liver, and spleen are common sites for lymphoma; intestinal lymphoma is less common. Intestinal lymphoma results in chronic weight loss and diarrhea and is often overlooked because it resembles other more common causes of chronic diarrhea such as ECE and inflammatory bowel disease. Intestinal rupture from an affected segment of bowel can be seen in some cases of intestinal lymphoma. These ferrets usually have an acute abdomen and septic peritonitis. Diagnosis can only be made by exploratory laparotomy and intestinal biopsy, although if other organs such as the liver or spleen are involved, needle aspirate or biopsy of these organs can be performed. Treatment for intestinal lymphoma carries a poor prognosis. Chemotherapy for lymphoma is described in Chapter 8.
GENERAL APPROACH TO VOMITING Owners may describe “vomiting” in their ferrets, but some of these animals may actually be regurgitating. In light of this, the differential diagnoses for emesis in ferrets include both esophageal diseases and gastroenteric disorders. In the clinical history, vomiting is not as frequently described in ferrets as it is in dogs or cats. For example, ferrets rarely vomit hairballs, and often vomiting is not part of the history associated with foreign body ingestion. The reason for this is unclear. No anatomic feature prevents emesis in ferrets; in fact, ferrets have long been laboratory animal models for human emesis studies because vomiting can be induced readily in ferrets in a laboratory setting.19 The major differential diagnoses for vomiting or regurgitation in ferrets include the presence of a GI foreign body, H. mustelae gastritis, gastroenteritis, and, rarely, megaesophagus. It is uncommon for ferrets with metabolic problems such as azotemia or hepatic disease to vomit. Although definitive diagnosis is not always possible, recognizing whether medical or surgical treatment is required is important. For example, most
obstructions caused by a foreign body require surgery, whereas gastroenteritis is a medical disease. However, differentiating these two diagnoses is often quite challenging (see Table 3-2). Diagnosis begins with the history. Pointedly question the owner regarding the chewing habits of the ferret: Does the ferret have a squeak toy? Is it unsupervised in the household or usually caged? Has vomiting been observed? The description of any vomiting behavior is significant. Also question the owner regarding the animal’s appetite and obtain a description of the feces. Ferrets that live in groups will need to be separated for observation. On physical examination, some foreign bodies in the small intestine can be distinctly palpated. However, enlarged mesenteric lymph nodes can feel like foreign objects. Also remember that foreign bodies in the stomach are difficult to detect on palpation. Proliferative bowel disease may result in palpably thickened intestines in the ferret; however, vomiting is not usually a feature of this disease. Radiography is the most important diagnostic test in the workup of a vomiting ferret. Always include the whole body in a survey radiograph. Radiographic signs of megaesophagus can be subtle. The heart may appear small because of hypovolemia from dehydration. Varying amounts of gas can be seen with a foreign body–related obstruction, and sometimes the incriminating object is visible. Segmental ileus or a dilated and gas- or fluid-filled stomach is a typical radiographic sign of obstruction (see Fig. 3-11). Not all cases of GI foreign body are obvious on radiographs. If evidence of foreign body obstruction is not well defined, consider medical therapy and repeat radiographs in 24 hours. Alternatively, give barium sulfate (8 to 10 mL/kg PO) for a series of contrast-enhanced films. Ferrets will readily take barium force-fed from a syringe. If there is a strong indication of the presence of a foreign body, perform abdominal exploratory surgery as soon as possible, preferably after parenteral fluid therapy has been started (see Chapter 11). Obtain tissue biopsy samples as needed (e.g., the liver or gastric mucosa), and save any foreign object to show to the owner. Always check the entire intestinal tract for lesions, and examine the pancreas and adrenal glands, especially in older ferrets. If a foreign body is not found, collect gastric and duodenal mucosal biopsy samples to submit for special staining or PCR testing for H. mustelae. Infection is associated with gastritis, especially in the antral region and the proximal duodenum (see earlier discussion). Although results of exploratory surgery may be negative for a foreign object, histologic examination of biopsy samples may or may not reveal a diagnosis. The possibility of negative findings should be discussed with the owner before surgery. If surgery is not an option or is not recommended, consider treatment for H. mustelae–associated gastritis (see earlier discussion) and administer fluid therapy as needed. If obstruction is still a possibility, administer a petrolatum hairball preparation at 1 mL q8–12h. Carefully examine all feces passed in the hospital; foreign objects or matter may sometimes be found in the stool.
GENERAL APPROACH TO DIARRHEA Normal ferrets nibble on food all day. Their GI transit time is short (3 hours), so defecation is frequent in the healthy state. The normal stool is slightly soft and formed. Diarrhea can range from mucoid and green to hemorrhagic. Anorexic ferrets may
CHAPTER 3 Gastrointestinal Diseases produce a very dark green (bile) stool that can resemble melena. Some owners describe a “birdseed” type of diarrhea that may be caused by malabsorption and is often associated with ECE or inflammatory bowel disease (see Fig. 3-13). Unlike canine patients, diarrhea in ferrets is difficult to classify as originating in the small intestine or the large intestine. More important are the onset, duration, and severity of the diarrhea, as well as concurrent clinical signs. Causes of diarrhea can be separated into diseases of young or older ferrets, as well as infectious or noninfectious causes. The most common noninfectious causes of diarrhea include stress, dietary indiscretion, foreign body ingestion, lymphosarcoma, and inflammatory bowel disease. Occasionally, severe metabolic disease can result in a green (bile-tinged), mucoid diarrhea. Eosinophilic gastroenteritis typically affects mature ferrets but is uncommon (see Table 3-2 and earlier discussion). Infectious agents are rare causes of diarrhea in closed groups or isolated ferrets, such as those kept as individual household pets. Ferrets do not usually have GI parasites, but coccidia can be present in young, newly purchased ferrets. Rotavirus can cause outbreaks of severe diarrhea, but most reports of this are in very young, unweaned ferrets. Ferrets that have been exposed to unfamiliar ferrets, such as show ferrets, may be susceptible to ferret enteric coronavirus. Newly acquired young ferrets can also act as asymptomatic carriers of coronavirus and expose naïve, older ferrets in a household group. Proliferative bowel disease is usually seen in young ferrets. Helicobacter-associated gastritis may also be present. Canine distemper virus in the epitheliotropic form can cause diarrhea in conjunction with respiratory and integumentary disease in unvaccinated ferrets. The clinical approach to the diagnosis of diarrhea depends on the severity and duration of clinical signs. Obtain a vaccination and dietary history and perform a direct fecal wet mount and centrifugation to check for GI parasites. Sick ferrets need a more comprehensive workup that includes radiographs to check for obstructive lesions and a CBC and a plasma biochemical analysis to assess metabolic conditions. If simple diagnostic tests do not reveal a cause and therapy is unsuccessful, consider exploratory surgery to evaluate the GI tract and obtain biopsy samples. Endoscopy can be difficult in ferrets because of their small size but may be an alternative to surgery. Consider culture of the feces for Salmonella species, especially if the ferret is febrile or the feces are hemorrhagic. Treat ferrets with mild diarrhea, without anorexia or vomiting, on an outpatient basis with an antibiotic such as amoxicillin or chloramphenicol. Metronidazole is a good enteric antibiotic, especially when paired with amoxicillin for Helicobacter therapy. Ferrets find metronidazole strongly distasteful, even when it is compounded into a suspension with fruit or chicken flavor. Hospitalize sick or dehydrated ferrets for supportive care and a diagnostic workup. Give fluids subcutaneously if a ferret is stable or intravenously if it is weak and dehydrated. Administer antibiotics parenterally if possible. A short course of a kaolin/pectin suspension (1 to 2 mL/kg PO q2–6h as required [prn]) or bismuth subsalicylate can be administered as a GI protectant until a more definitive diagnosis is established. Drugs that affect the motility of the GI tract should not be administered without an initial diagnosis, although in ferrets with severe diarrhea, loperamide can be administered (0.2 mg/kg q12h). Ferrets with chronic diarrhea
41
may have diminished levels of cobalamin from intestinal malabsorption.42 Cobalamin administration can be given following the feline protocol: 250 μg SC every 7 days for 6 weeks, then 250 μg SC every 14 days for 6 weeks, then monthly.42
DIFFERENTIATION OF EMACIATION (“WASTING DISEASE”) WITH DIARRHEA Helicobacter mustelae–associated gastritis and PBD may occur independently, sequentially, or concurrently in the same animal. Proliferative bowel disease is a sufficient stressor to induce clinical gastritis in a ferret colonized with H. mustelae. Although these two diseases are most common in ferrets 12 to 16 weeks of age, sufficiently stressed mature ferrets may also be affected. However, clinical disease in adult animals is more often associated with Helicobacter-associated gastritis than with PBD. Although eosinophilic gastroenteritis has been confirmed only in adults, it also may occur in young animals. Any of the wasting diseases can be diagnosed by gastric and intestinal biopsy. However, a presumptive diagnosis may be based on the clinical examination, an accurate history, and results of routine diagnostic tests, which may include radiographs, CBC, plasma biochemical analysis, and fecal examination. Diagnosis may be “confirmed” by the response to appropriate treatment. Characteristics of GI diseases that cause diarrhea and weight loss are summarized in Table 3-2. Other important differential diagnoses for diarrhea and weight loss in domestic ferrets are chronic GI foreign bodies, lymphoma, coronavirus, Aleutian disease, and rarely, mycobacteriosis.
STEPS IN DIAGNOSIS OF WASTING DISEASE History Question the owner of a lethargic, anorexic ferret with diarrhea and sudden weight loss about changes in the ferret’s diet, feeding schedule, and access to water. The stress factor most commonly associated with wasting diseases is restriction of food for any reason, including the following. Self-Denial of Food. Ferrets resist changing to a food that differs in flavor and texture from the one to which they are accustomed and may fast for several days rather than eat the new food. Fasting depletes fat stores, which should not be confused with the loss of muscle mass associated with wasting diseases. Restricted Access to Water. Ferrets consume about three times as much water as dry food pellets and cannot meet their nutritional requirements if water is restricted. Restricted Access to Food. Food hoppers used with some types of ferret cages may be easily blocked by large food pellets or pellets with unusual shapes, and the owner may not realize that the ferret is unable to get its food. Children caring for ferrets are less likely than adults to understand the significance of an unchanging level of food in the hopper for several days. In addition, some ferrets habitually dig their food out of the container and refuse to eat food that becomes wet or contaminated on the cage floor. Inappropriate or Nutritionally Deficient Diet. Occasionally, new owners provide ferrets with inappropriate foods, such as dog food or poor-quality cat food, or offer them excessive amounts of sweet treats, especially raisins, which are palatable
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SECTION I Ferrets
but contain almost 100% sugar and no protein. Rapidly growing young animals with nutritional deficiencies are much more susceptible to infectious diseases. Environmental Stress. Exposure to extremes of temperature, particularly heat, is very stressful to ferrets. Animals may be stressed during inclement weather if they are housed outdoors without adequate protection from wind and rain, especially if their food is of poor quality or subject to wetting, caking, and molding. Ask the owner if the affected pet is his or her first ferret. You may identify stressors that the owner would not have taken into consideration.
Physical Examination Palpate the abdomen of an emaciated ferret. Grossly thickened areas of bowel in ferrets with PBD and eosinophilic gastroenteritis are sometimes palpable. A focal area of pain in the abdomen is more typical of the presence of a GI foreign body. Splenomegaly is common in ferrets in association with many diseases, and mesenteric lymph nodes are likely to be enlarged in ferrets with any of the wasting diseases. Projectile vomiting has been reported in one ferret with eosinophilic gastroenteritis.18 Although rectal prolapse is not pathognomonic for PBD, this diagnosis is highly probable in a ferret with prolapse associated with diarrhea and weight loss. In young ferrets (younger than 16 weeks of age and usually younger than 10 weeks), coccidiosis may be associated with diarrhea and rectal prolapse but is rarely associated with significant weight loss. Both Aleutian disease and lymphosarcoma are insidious; thin ferrets will have lost condition over a period of weeks or months and may not have diarrhea. Radiography is the most useful tool for detecting a GI foreign body (see earlier discussion). Contrast radiographs are sometimes helpful in identifying obstruction with a radiolucent foreign body, but radiographs may also suggest areas of gastric ulceration or intestinal mucosal proliferation in ferrets with either PBD or eosinophilic gastroenteritis. To help rule out the possibility of lymphosarcoma or eosinophilic gastroenteritis, obtain a blood sample for a CBC. Ferrets with eosinophilic gastroenteritis usually have dramatic eosinophilia (10% to 35% eosinophils compared with 3% to 5% in normal ferrets). Ferrets with lymphosarcoma may not be leukemic, and further tests, such as peripheral lymph node biopsy or a splenic aspirate, are necessary for diagnosis. Inflammation associated with PBD often causes leukocytosis with neutrophilia and a left shift. Ferrets with bleeding ulcers are usually anemic (normal hematocrit is 40% to 55% in spayed or neutered pets, lower in jills in estrus). Dehydration may mask mild anemia and hypoproteinemia in emaciated animals; therefore repeat the CBC after rehydration. Aleutian disease may cause diarrhea, anemia, leukocytosis, and wasting. Serologic tests for Aleutian disease virus are available (see Chapter 5), but many ferrets with positive test results are asymptomatic for Aleutian disease. Clinicians should not assume that illness in ferrets that are seropositive for Aleutian disease virus is caused by the virus.
TREATMENT OF FERRETS WITH WASTING AND DIARRHEA Debilitated ferrets with diarrhea need rehydration with either intravenously (preferred) or subcutaneously administered balanced electrolyte solutions. These ferrets are often very weak and
usually allow a saphenous or cephalic catheter to be inserted with little resistance. However, if a catheter cannot be placed, fluids administered subcutaneously usually are well and rapidly absorbed. Alternatively, an intraosseous catheter can be used until hydration is improved and an intravenous catheter can be placed. Hospitalize an emaciated, dehydrated ferret until fluid and electrolyte balances are reestablished. Most cachectic ferrets with diarrhea that do not have a GI foreign body or eosinophilia presumably have both ileitis/ colitis and H. mustelae–associated gastritis with ulcers. While waiting for results of diagnostic tests, if the ferret is treated for only one disease, the time required for recognizing treatment failure may be the factor that ultimately decides if the ferret will survive. Unfortunately, the effective drugs used for treating the diseases associated with wasting are different, and therapy necessitates multiple daily doses of several drugs for at least 2 weeks. Because the gut flora of ferrets is very simple and plays no vital role in digestion, long-term administration of broad-spectrum antibiotics does not cause dysbiosis and diarrhea in ferrets as it does in many species. Emaciated ferrets that die despite proper treatment usually have either very extensive gastric ulcers or severe ileitis, each of which drastically reduces the absorption of essential nutrients. The age of the animal may sometimes help in the differential diagnosis; young ferrets are more likely to develop PBD, whereas older ferrets develop inflammatory bowel disease or severe, chronic gastritis associated with H. mustelae infection.29 Also, H. mustelae–associated disease is more often associated with stress factors common in mature pet ferrets, such as concurrent disease or surgery (see Table 3-2). Emaciated animals have no energy reserve and should receive intensive care. Offering a variety of premium ferret foods and high-calorie, easily absorbed foods is important. Some animals refuse to eat their regular diet of dry pellets but do accept the same food mixed with water and heated in a microwave until it develops a porridge-like consistency. Supplemental foods formulated specifically for carnivores such as Carnivore Care (Oxbow Animal Health) or Emeraid Carnivore (Lafebervet.com) are readily accepted by most ferrets when fed by syringe. Alternatively, offer nutritional recovery foods formulated for dogs and cats such as Maximum-Calorie (The Iams Company, Dayton, OH) or Canine a/d (Hill’s Pet Nutrition); most sick ferrets accept these foods readily. Nutritional recovery diets such as these can be used as the sole source of nutrition for several weeks if necessary. When using these diets, calculate a minimum daily intake of 400 kcal per kilogram of body weight. Sick ferrets may not make the effort to get up and drink from a water bottle but do usually drink from a dish. Offer fresh food and water by hand several times daily during hospitalization and home care; ferrets often take a few mouthfuls of every new offering but never go back for more. The first 2 days are critical for an animal that has lost 40% to 50% of its body weight, and intensive supportive care is essential. When ferrets regain their appetites, often within 48 hours of the first doses of medication, and diarrhea has subsided, owners may be tempted to stop treatment. However, ferrets treated for less than 2 weeks often relapse, and some ferrets need antibiotic therapy and supportive care for an additional 2 to 3 weeks if they are to recover completely. Many ferrets with wasting diseases will survive with aggressive and continued treatment.
CHAPTER 3 Gastrointestinal Diseases
References 1. Abe N, Iseki M. Identification of genotypes of Cryptosporidium parvum isolates from ferrets in Japan. Parasitol Res. 2003;89:422-424. 2. Alder JD, Ewing PJ, Mitten MJ, et al. Relevance of the ferret model of Helicobacter-induced gastritis to evaluation of antibacterial therapies. Am J Gastroenterol. 1996;91:2347-2354. 3. Archer DC, Barrie Edwards G, Kelly DF, et al. Obstruction of equine small intestine associated with focal idiopathic eosinophilic enteritis: an emerging disease?. Vet J. 2006;171:504-512. 4. Batchelder M, Fox JG, Hayward A, et al. Natural and experimental Helicobacter mustelae reinfection following successful antimicrobial eradication ferrets. Helicobacter. 1996;1:34-42. 5. Bauck LS. Salivary mucocele in 2 ferrets. Mod Vet Pract. 1985;66:337-339. 6. Bell JA, Manning DD. Evaluation of Campylobacter jejuni colonization of the domestic ferret intestine as a model of proliferative colitis. Am J Vet Res. 1991;52:826-832. 7. Bernard S, Gorham JR, Ryland LM. Biology and diseases of ferrets. In: Fox JG, Cohen BJ, Loew FM, eds. Laboratory animal medicine. New York: Academic Press; 1984:385-397. 8. Blanco MC, Fox JG, Rosenthal K, et al. Megaesophagus in nine ferrets. J Am Vet Med Assoc. 1994;205:444-447. 9. Blankenship-Paris TL, Chang J, Bagnell CR. Enteric coccidiosis in a ferret. Lab Anim Sci. 1993;43:361-363. 10. Bryant JL, Hanner TL, Fultz DG, et al. A chronic granulomatous intestinal disease in ferrets caused by an acid-fast organism morphologically similar to Mycobacterium paratuberculosis. Lab Anim Sci. 1988;38:498-499. 11. Burgess M, Garner M. Clinical aspects of inflammatory bowel disease in ferrets. Exot DVM. 2002;4.2:29-34. 12. Caliguiri R, Bellah JR, Collins BR, et al. Medical and surgical management of esophageal foreign body in a ferret. J Am Med Vet Assoc. 1989;195:969-971. 13. Carmel B. Eosinophilic gastroenteritis in three ferrets. Vet Clin North Am Exot Anim Pract. 2006:707-712. 14. Church RR. Impact of diet on the dentition of the domesticated ferret. Exot DVM. 2007;9:30-39. 15. Cooper DM, Swanson DL, Gebhart CJ. Diagnosis of proliferative enteritis in frozen and formalin-fixed, paraffin-embedded tissues from a hamster, horse, deer and ostrich using a Lawsonia intracellularis–specific multiplex PCR assay. Vet Microbiol. 1997;54:47-62. 16. Darby C, Ntavlourou V. Hepatic hemangiosarcoma in two ferrets (Mustela putorius furo). Vet Clin North Am Exot Anim Pract. 2006;9(3):689-694. 17. Erdman SE, Correa P, Coleman LA, et al. Helicobacter mustelae–associated gastric MALT lymphoma in ferrets. Am J Pathol. 1997;151:273-280. 18. Fazakas S. Eosinophilic gastroenteritis in a domestic ferret. Can Vet J. 2000;41:707-709. 19. Florcyzk AP, Schurig JE, Bradner WT. Cisplatin-induced emesis in the ferret: a new animal model. Cancer Treat Rep. 1982;66:187-189. 20. Fox JG. Bacterial and mycoplasmal diseases. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:210-211. 21. Fox JG. Systemic diseases. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:258-259. 22. Fox JG. Bacterial and mycoplasmal diseases: Helicobacter mustelae. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:327-333. 23. Fox JG. Bacterial and mycoplasmal diseases: proliferative bowel disease— Desulfovibrio spp. (Lawsonia intracellularis). In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:335-339.
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24. Fox JG, Ackerman JI, Newcomer CE. Ferret as a potential reservoir for human campylobacteriosis. Am J Vet Res. 1983;44:1049-1052. 25. Fox JG, Ackerman JI, Taylor NS, et al. Campylobacter jejuni infection in the ferret: an animal model of human campylobacteriosis. Am J Vet Res. 1987;48:85-90. 26. Fox JG, Correa P, Taylor NS, et al. Helicobacter mustelae– associated gastritis in ferrets. An animal model of Helicobacter pylori gastritis in humans. Gastroenterology. 1990;99:352-361. 27. Fox JG, Dangler CA, Sager W, et al. Helicobacter mustelae– associated gastric adenocarcinoma in ferrets (Mustela putorius furo). Vet Pathol. 1997;34:225-229. 28. Fox JG, Dewhirst FE, Fraser GJ, et al. Intracellular Campylobacter-like organism from ferrets and hamsters with proliferative bowel disease is a Desulfovibrio sp. J Clin Microbiol. 1994;32:1229-1237. 29. Fox JG, Marini RP. Helicobacter mustelae infection in ferrets: pathogenesis, epizootiology, diagnosis, and treatment. Semin Avian Exot Pet Med. 2001;10:36-44. 30. Fox JG, Murphy JC, Ackerman JI, et al. Proliferative colitis in ferrets. Am J Vet Res. 1982;43:858-864. 31. Fox JG, Paster BJ, Dewhirst FE, et al. Helicobacter mustelae isolation from feces of ferrets: evidence to support fecaloral transmission of a gastric Helicobacter. Infect Immun. 1992;60:606-611. (Published erratum appears in Infect Immun. 1992;60:4443.) 32. Fox JG, Wang TC. Inflammation, atrophy, and gastric cancer. J Clin Invest. 2007;117(1):60-69. 33. Fox JG, Zeman DH, Mortimer JD. Copper toxicosis in sibling ferrets. J Am Vet Med Assoc. 1994;205:1154-1156. 34. Garcia A, Erdman SE, Xu S, et al. Hepatobiliary inflammation, neoplasia, and argyrophilic bacteria in a ferret colony. Vet Pathol. 2002;39:173-179. 35. Garner MM, Ramsell K, Morera N, et al. Clinicopathologic features of a systemic coronavirus-associated disease resembling feline infectious peritonitis in the domestic ferret (Mustela putorius). Vet Pathol. 2008;45(2):236-246. 36. Graham J, Fidel J, Mison M. Rostral maxillectomy and radiation therapy to manage squamous cell carcinoma in a ferret. Vet Clin North Am Exot Anim Pract. 2006;9:701-706. 37. Goad ME, Fox JG. Neoplasia in ferrets. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:281-282. 37a. Hall BA, Ketz-Riley CJ. Cholestasis and cholelithiasis in a domestic ferret (Mustela putorius furo). J Vet Diagn Invest. 2011;23:836-839. 38. Hamilton TA, Morrison WB. Bleomycin chemotherapy for metastatic squamous cell carcinoma in a ferret. J Am Vet Med Assoc. 1991;1998:107-108. 39. Harms CA, Andrews GA. Megaesophagus in a domestic ferret. Lab An Sci. 1993;43:506-508. 40. Harper DS, Mann PH, Regner S. Measurement of dietary and dentifrice effects upon calculus accumulation rates in the domestic ferret. J Dent Res. 1990;69:447-450. 40a. Hauptman K, Jekl V, Knotek Z. Extra-hepatic biliary obstruction in 2 ferrets (Mustela putorius furo). J Small Anim Prac. 2011;52:371-375. 41. Havard TJ, Sarsfield P, Wotherspoon AC, et al. Increased gastric epithelial cell proliferation in Helicobacter pylori associated follicular gastritis. J Clin Pathol. 1996;49:68-71. 42. Hoppes S. The senior ferret (Mustela putorius furo). Vet Clin North Am Exot Anim Pract. 2010;13(1):107-122. 43. Huynh M. Myasthenia gravis in a ferret. Proc Assoc Exot Mam Vet. 2009;93. 44. Johnson-Delaney CA, Nelson WB. A rapid procedure for filling fractured canine teeth of ferrets. J Sm Exot Anim Med. 1992;1:100-102.
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SECTION I Ferrets
45. Krueger KL, Murphy JC, Fox JG. Treatment of proliferative colitis in ferrets. J Am Vet Med Assoc. 1989;194:1435-1436. 46. Lavoie JP, Drolet R, Parsons D, et al. Equine proliferative enteropathy: a cause of weight loss, colic, diarrhea and hypoproteinaemia in foals on three breeding farms in Canada. Eq Vet J. 2000;32:418-425. 47. Li X, Fox JG, Padrid PA. Neoplastic diseases in ferrets: 574 cases (1968-1997). J Am Vet Med Assoc. 1998;212(9):1402-1406. 48. Lucas J, Lucas A, Furber H, et al. Mycobacterium genavense infection in two aged ferrets with conjunctival lesions. Aust Vet J. 2000;78:685-689. 49. Lunn JA, Martin P, Zaki S, et al. Pneumonia due to Mycobacerium abscessus in two domestic ferrets (Mustelo putorius furo). Aust Vet J. 2005;83:542-546. 50. Lugton IW, Wobeser G, Morris RS, et al. Epidemiology of Mycobacterium bovis infection in feral ferrets (Mustela furo) in New Zealand: II. Routes of infection and excretion. N Z Vet J. 1997;45:151-157. 51. Lyles SE, Panciera DL, Saunders GK, et al. Idiopathic eosinophilic masses of the gastrointestinal tract in dogs. J Vet Int Med. 2009;23:818-823. 52. Marini RP, Adkins JA, Fox JG. Proven or potential zoonotic diseases of ferrets. J Am Vet Med Assoc. 1989;195:990-993. 53. Marini RP, Fox JG, Taylor NS, et al. Ranitidine bismuth citrate and clarithromycin, alone or in combination, for eradication of Helicobacter mustelae in ferrets. Am J Vet Res. 1999;60:1280-1286. 54. Martinez J, Renacher M, Perpinan D, et al. Identification of group 1 coronavirus antigen in multisystemic granulomatous lesions in ferrets (Mustela putorius furo). J Comp Pathol. 2008;138(1):54-58. 55. McColm AA, Bagshaw JA, O’Malley CF. Development of a 14C-urea breath test in ferrets colonised with Helicobacter mustelae: effects of treatment with bismuth, antibiotics, and urease inhibitors. Gut. 1993;34:181-186. 56. McOrist S, Mackie RA, Lawson GHK. Antimicrobial susceptibility of ileal symbiont intracellularis isolated from pigs with proliferative enteropathy. J Clin Microbiol. 1995;33:1314-1317. 57. Miller PE, Pickett JP. Zygomatic salivary gland mucocele in a ferret. J Am Vet Med Assoc. 1989;194:1437-1438. 58. Mullen HS, Scavelli TD, Quesenberry KE, et al. Gastrointestinal foreign body in ferrets: 25 cases (1986-1990). J Am Anim Hosp Assoc. 1989;28:13-19. 59. Nakata M, Miwa Y, Nakayama H, et al. Localized radiotherapy for a ferret with possible anal sac aprocrine adenocarcinoma. J Small Anim Pract. 2008;49:476-478. 60. Naito Y, Yoshikawa T. Molecular and cellular mechanisms involved in Helicobacter pylori–induced inflammation and oxidative stress. Free Radic Biol Med. 2002;33:323-336. 61. Otto G, Fox JG, Wu P-Y, et al. Eradication of Helicobacter mustelae from the ferret stomach: an animal model of Helicobacter (Campylobacter) pylori chemotherapy. Antimicrob Agents Chemother. 1990;34:1232-1236. 62. Palley LS, Fox JG. Eosinophilic gastroenteritis in the ferret. In: Kirk RW, Bonagura JD, eds. Kirk’s current veterinary therapy XI: small animal practice. Philadelphia: WB Saunders; 1992:1182-1184. 63. Perpinan D, Lopez C. Clinical aspects of systemic granulomatous inflammatory syndrome in ferrets (Mustela putorius furo). Vet Rec. 2008;162:180-183. 64. Phillips RH, Whitehead MW, Lacey S, et al. Solubility, absorption, and anti-Helicobacter pylori activity of bismuth subnitrate and colloidal bismuth subcitrate: in vitro data do not predict in vivo efficacy. Helicobacter. 2000;5:176-182. 65. Piseddu E, Trotta M, Tortoli E, et al. Detection and molecular characterization of Mycobacterium celatum as a cause of splenitis in a domestic ferret (Mustela putorius furo). J Comp Pathol. 2011;144(2-3):214-218.
66. Poddar S, Jacob S. Gross and microscopic anatomy of the major salivary glands of the ferret. Acta Anat (Basel). 1977;98:434-443. 67. Prinz C, Zanner R, Gratzl M. Physiology of gastric enterochromaffin-like cells. Annu Rev Physiol. 2003;65:371-382. 68. Rehg JE, Gigliotti F, Stokes DC. Cryptosporidiosis in ferrets. Lab Anim Sci. 1988;38:155-158. 69. Rice LE, Stahl SJ, McLeod Jr C. Pyloric adenocarcinoma in a ferret. J Am Vet Med Assoc. 1992;200:1117-1118. 70. Rodeheffer C, von Messling V, Milot S, et al. Disease manifestations of canine distemper virus infection in ferrets are modulated by vitamin A status. J Nutr. 2007;137:1916-1922. 71. Saunders GK, Thomsen BV. Lymphoma and Mycobacterium avium infection in a ferret (Mustela putorius furo). J Vet Dian Invest. 2006;18:513-515. 72. Shoeb TR, Fox JG. Enterocecocolitis associated with intraepithelial Campylobacter-like bacteria in rabbits (Oryctolagus cuniculus). Vet Pathol. 1990;27:73-80. 73. Schulman FY, Montali RJ, Hauer PJ. Gastroenteritis associated with Clostridium perfringens type A in black-footed ferrets (Mustela nigripes). Vet Pathol. 1993;30:308-310. 74. Schultheiss PC, Dolginow SZ. Granulomatous enteritis caused by Mycobacterium avium in a ferret. J Am Vet Med Assoc. 1994;204:1217-1218. 75. Sherding RG, Johnston SE. Diseases of the intestines. In: Sherding RG, ed. Saunders manual of small animal practice. 2nd ed. Philadelphia: WB Saunders; 2000:787-815. 76. Sleeman JM, Clyde VL, Jones MP, et al. Two cases of pyloric adenocarcinoma in the ferret (Mustela putorius furo). Vet Rec. 1995;17:272. 77. Stables R, Campbell C, Clayton N, et al. Gastric anti-secretory, mucosal protective, anti-pepsin and anti-Helicobacter properties of ranitidine bismuth citrate. Aliment Pharmacol Ther. 1993;7:237-246. 78. Stauber E, Kraft S, Roninette J, et al. Multiple tumors in a ferret. J Sm Exot Anim Med. 1991;1:87-88. 79. Talley NJ, Shorter RG, Phillips SF, et al. Eosinophilic gastroenteritis: a clinicopathological study of patients with disease of the mucosa, muscle layer, and subserosal tissues. Gut. 1990;31:54-58. 80. Tams TR. Cisapride: clinical experience with the newest GI prokinetic drug. Proc 12th Annu Meet Am Col Vet Int Med Forum. 1994:100-101. 81. Torres-Medina A. Isolation of an atypical rotavirus causing diarrhea in neonatal ferrets. Lab Anim Sci. 1987;37:167-171. 82. Valheim M, Djonne B, Heiene R, et al. Disseminated Mycobacterium celatum (type 3) infection in a domestic ferret (Mustela putorius furo). Vet Pathol. 2001;38:460-463. 83. Van de Gaag I, van der Linde-Sipman JS. Eosinophilic granulomatous colitis with ulceration in a dog. J Comp Pathol. 1987;97:179-185. 84. Wagner R, Finkler MR. Diagnosing gastric hairballs in ferrets. Exot DVM. 2008;10:9-23. 85. Williams B. Pathology of the domestic ferret squamous cell carcinoma, 1999: Retrieved March 2010 from www.afip.org/ consultation/vetpath/ferrets/SCC/SCC.html. 86. Williams BH, Kiupel M, West KH, et al. Coronavirusassociated epizootic catarrhal enteritis in ferrets. J Am Vet Med Assoc. 2000;217:526-530. 87. Wise AG, Kiupel M, Garner MM, et al. Comparative sequence analysis of the distal one-third of the genomes of a systemic and an enteric ferret coronavirus. Virus Res. 2010;149:42-50. 88. Wise AG, Kiupel M, Maes RK. Molecular charcterization of a novel coronavirus associated with epizootic catarrhal enteritis (ECE) in ferrets. Virology. 2006;349:164-174. 89. Wise AG, Smedley RC, Kiupel M, et al. Detection of group C rotavirus in juvenile ferrets (Mustela putorius furo) with diarrhea by reverse transcription polymerase chain reaction: sequencing and analysis of the complete coding redion of the VP6 Gene. Vet Pathol. 2009;46:985-991.
CHAPTER 3 Gastrointestinal Diseases 90. Wotherspoon AC. A critical review of the effect of Helicobacter pylori eradication on gastric MALT lymphoma. Curr Gastroenterol Rep. 2000;2:494-498. 91. Wotherspoon AC. Gastric lymphoma of mucosa-associated lymphoid tissue and Helicobacter pylori. Ann Rev Med. 1998;49:289-299.
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92. Yun MY, Cho YU, Park IS, et al. Eosinophilic gastroenteritis presenting as small bowel obstruction: a case report and review of the literature. World J Gastroenterol. 2007;13:1758-1760.
CHAPTER
4
Disorders of the Urinary and Reproductive Systems
Christal G. Pollock, DVM, Diplomate ABVP (Avian)
Disorders of the Urinary System Renal Cysts Polycystic Kidney Disease Hydronephrosis Renal Disease and Renal Failure Aleutian Disease Nephrocalcinosis Pyelonephritis Renal Neoplasia Ureteral Rupture Urolithiasis Cystitis Bladder Neoplasia Urinary Incontinence Urethral Obstruction Prostatic Cysts Prostatitis and Prostatic Abscess Prostatic Tumors Paraurethral Cysts or Paraurethral Disease Disorders of the Reproductive System The Male Ferret The Female Ferret Periparturient Disease Normal Breeding Management of Breeding Ferrets Diseases of the Jill Diseases of the Kit
DISORDERS OF THE URINARY SYSTEM The ferret has a classic bean-shaped kidney that sits in the retroperitoneal space; its dorsal surface lies in direct contact with sublumbar musculature.21 Renal pathology is a common necropsy finding in ferrets,25,31 whereas renal disease, such as renal failure, cystitis, and urolithiasis, is uncommon in the pet ferret. The only 46
exception is prostatic disease, which is relatively common in the United States, because of the high prevalence of adrenal disease.
RENAL CYSTS Renal cysts are a common incidental finding in ferrets. At one institution, cysts were found in 10% to 15% of ferrets submitted for necropsy.25 Renal cysts were detected sonographically at percentages ranging from 31.6% to 62.8% of ferrets.27,31,59,60 Renal cysts may range from 1 to 25 mm in diameter.19,38,69 They are usually present singly or in small numbers and can be present in one or both kidneys.20 Cysts are commonly found in the cortices, in close relationship to the pelvic recesses.59,60 The cause of renal cysts in ferrets is uncertain. There is no known hereditary basis for cyst formation, but they are thought to be congenital.60 Renal cysts may be detected during physical examination (as one or more smooth masses on the renal surface), during sonographic examination (as smooth-walled, hypoechoic areas), or grossly (as translucent swellings visible through the renal capsule). Cysts may pit the kidney surface creating a wedge of cortical pallor and renal congestion.31 In ferrets with renal cysts, submit samples for a complete blood count (CBC), serum biochemical analysis, and urinalysis. Use ultrasonography to evaluate renal architecture. Pyelography with intravenous contrast media or nuclear scintigraphy may be used to evaluate renal function in clinically affected ferrets. There is no specific treatment for renal cysts. Monitor affected ferrets with periodic palpation and, if clinically indicated, ultrasound examination, serum biochemical analysis, and urinalysis. In humans, simple cysts are usually clinically silent, although they occasionally hemorrhage and cause acute pain.10 If a cyst becomes very large or painful, consider unilateral nephrectomy; however, be sure the contralateral kidney is functioning adequately before surgery.
POLYCYSTIC KIDNEY DISEASE The quantity, location, and size of the typical ferret renal cyst are very different from polycystic kidney disease,60 which is rare in ferrets. Unlike the smooth masses palpable with renal cysts, Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 4 Disorders of the Urinary and Reproductive Systems
47
Fig. 4-1 Polycystic kidney in a 3-year-old female ferret in acute renal failure. Capsular aspect. polycystic kidneys are palpable as slightly irregular, enlarged, firm, oval masses that may grossly appear pale to tan or gray in color.21,31 Cysts may also be diffusely distributed throughout other parenchyma, particularly the liver. If sufficient renal architecture is disrupted, cysts will lead to renal failure (Fig. 4-1). Ferrets may have anorexia, weight loss, lethargy, and signs of gastrointestinal dysfunction.21 Use ultrasonography or intravenous pyelography to confirm disease. Unilateral nephrectomy carries a good prognosis if function of the remaining kidney is normal. Assess the contralateral kidney using CBC, serum biochemistry analysis, and urinalysis. Several renal cysts were found at necropsy in an adult ferret with seizure activity. Neurologic signs were presumed to be caused by uremic encephalopathy, although the brain was unavailable for examination.15 In a second case report, polycystic kidneys and bilateral perinephric pseudocysts were reported in an adult ferret. Pseudocyst formation led to marked abdominal distention and tachypnea. Ultrasound-guided paracentesis was used initially as a palliative treatment, but fluid reaccumulated rapidly. Although plans were made to resect the renal capsule, the ferret declined rapidly and was euthanatized instead.57
HYDRONEPHROSIS Hydronephrosis and hydroureter are uncommon findings in ferrets usually attributed to ligation of the ureter during ovariohysterectomy.46 There have also been rare reports of hydronephrosis associated with obstruction caused by ureteral calculi, carcinoma involving the renal pelvis, and cystitis and herniation of the bladder.9,37,53 Affected ferrets typically have progressive abdominal distention. The hydronephrotic kidney is palpable as renomegaly or an abdominal mass. If hydronephrosis is suspected, submit samples for CBC, serum biochemistry analysis, and urinalysis. Use imaging to further characterize the swelling. On survey radiographs, the hydronephrotic kidney appears as a radiopaque, mid-abdominal mass with a large fluid density. Confirm your diagnosis via ultrasound or intravenous contrast pyelography. Fine-needle aspiration of the fluid-filled mass should reveal a transudate unless secondary bacterial infection is present. Treatment of hydronephrosis relies on unilateral nephrectomy, and carries a good prognosis if function of the remaining kidney is normal.
RENAL DISEASE AND RENAL FAILURE Chronic Renal Failure Variable degrees of chronic interstitial nephritis are commonly found at necropsy in ferrets older than 4 years of age.20,31,35 Advanced interstitial nephrititis, as well as pyelonephritis, glomerulonephritis, bilirubinuric nephrosis (C. G. Pollock, personal observation, 2003), and immune complex–mediated glomerulonephropathy caused by Aleutian disease (see discussion later in this chapter), can cause renal failure.54 Use of nephrotoxic drugs may also result in renal disease, and renal failure has also been reported as a result of an epinephrine overdose given to a ferret with a vaccine reaction.16 Although uncommon, renal tumors are another potential cause of renal failure in ferrets.9
Acute Renal Failure The most common cause of acute renal failure (ARF) in the ferret is urethral obstruction secondary to prostatic disease or, less commonly, urolithiasis.21,40 Another important, but uncommon, cause of ARF is toxic exposure. The minimum lethal dose of ibuprofen is 220 mg/kg, which means that ingestion of one tablet could be fatal in a small ferret. Clinical signs appear within 48 hours of ibuprofen ingestion and as rapidly as in 4 hours in 40% of ferrets. The most common clinical signs are neurologic, and more than half of ferrets also have gastrointestinal signs. Renal signs may include polyuria and polydipsia. In rare instances, acetaminophen ingestion may also cause ARF.17
Diagnosis and Treatment Clinical signs may include polyuria, polydipsia, oral ulcers, nonspecific signs of illness (such as anorexia, weight loss, and lethargy), signs of gastrointestinal upset (including melena and, in rare instances, vomiting), pelvic limb paresis, and even ataxia. Physical examination findings can include dehydration, pallor, and irregularities or asymmetry in the size and shape of the kidneys. Abnormal serum biochemistry results may include hyperphosphatemia, hyperkalemia, reduced total carbon dioxide levels, and azotemia. Interestingly, increases in serum creatinine levels are relatively moderate (generally less than 2 mg/dL). Kawasaki35 described two ferrets with severe renal disease confirmed by histologic examination; both ferrets had a creatinine level of 1.1 mg/dL in conjunction with blood urea nitrogen values of 140 and 320 mg/dL. The reported normal mean
48
SECTION I Ferrets
creatinine level in ferrets is lower (0.4 to 0.6 mg/dL) and the range is narrower (0.2 to 0.9 mg/dL) than in cats and dogs.19,23 Reference values have been established for endogenous and exogenous creatinine and insulin clearance tests used to evaluate glomerular filtration in ferrets.18,19 Although reference values for urine protein/creatinine ratios (UP:C) have not been established for ferrets, UP:C accurately reflects protein excretion in cats and dogs over a 24-hour period regardless of sex, urine collection method, time of day, or whether the animal has been fasted or fed. Canine reference values are generally less than 1.0 and cats are less than 0.6.68 If renal disease is diagnosed, attempt to identify the underlying cause with ultrasound examination and, if necessary, ultrasound-guided renal biopsy (as in cats). Sonographically, healthy ferret kidneys show a median resistive index (RI) of 0.54 ± 0.04 and a median pulsatility index (PI) of 0.83 ± 0.10. Even with severe renal parenchymal changes no significant alterations in blood flow parameters are seen; however, these indices do increase with age. Interestingly, male ferret kidneys have significantly higher blood flow velocities than females.32 At necropsy, kidneys with significant disease are often grossly pitted with large, focal depressions in the outer cortex secondary to scarring.21 Although treatment should be aimed at the underlying cause, nonspecific therapy of renal failure includes supportive care such as vitamin and iron supplementation, nutritional support, omega-3 fatty acid supplementation, erythropoietin for anemia, and fluid therapy. Normal daily fluid intake is estimated to be 75 to 100 mL/kg/day. Beware of overhydration in ferrets. Monitor patients for evidence of serous nasal discharge, tachypnea, or dyspnea and monitor body weight at least twice daily. Begin antibiotics if clinically indicated and use culture and sensitivity results when available. Discontinue any potentially nephrotoxic drugs,21 and administer phosphate binders and diuretics as needed. The use of small animal renal diets is difficult to impossible in ferrets since most individuals do not find these products palatable. Furthermore, the effects of longterm protein restriction require study in the ferret. Lewington37 has proposed feeding ferrets with renal disease discrete meals instead of free choice. The prognosis for ferrets with renal failure depends on laboratory findings and response to therapy.
ALEUTIAN DISEASE Many ferrets infected with the parvovirus that causes Aleutian disease are asymptomatic carriers. Clinical signs are usually consistent with chronic wasting and ataxia. Additional clinical signs may be attributed to immune complex deposition. Deposits in the kidney may cause membranous glomerulonephritis and tubular interstitial nephritis, and possibly renal failure. Aleutian disease virus (ADV) may also cause liver failure, intestinal disease including melena, and central nervous system disease.54 See Chapter 5 for diagnosis and treatment of ADV.
NEPHROCALCINOSIS In three separate sonographic surveys, hyperechogenicity of the renal medulla was seen in 40% to 58% of European pet ferrets.27,59,60 This sonographic finding was not identified in laboratory ferrets. Histologic evaluation showed calcium deposits in the renal tubules consistent with renal calcification. Crystalline
structures within renal parenchyma that could not be found sonographically were believed to be urates. Dietary analyses suggested that the presumptive nephrocalcinosis was related to excess dietary calcium and phosphorus.60
PYELONEPHRITIS Pyelonephritis is an uncommon finding in the ferret usually associated with an ascending bacterial urinary tract infection or sepsis. Hemolytic Escherichia coli and Staphylococcus aureus are the most common causative agents. Clinical signs include anorexia, lethargy, fever, and pain on palpation of the kidneys.20 Severe suppurative pyelonephritis progressing to endstage chronic renal failure was reported in a ferret treated for lymphoma. Immunosuppression from long-term corticosteroid administration may have led to cystitis and secondary pyelonephritis.61 It can be difficult to differentiate pyelonephritis from lower urinary tract disease using history and physical examination alone. Urinalysis will reveal hematuria, pyuria, renal tubular cells, and cellular casts. Abdominal ultrasound or intravenous pyelography may help confirm the diagnosis. Provide supportive care and administer antibiotics for 3 to 6 weeks based on culture and sensitivity results.
RENAL NEOPLASIA Renal tumors are very rare in the ferret.9,34,39 The most commonly reported primary tumor is renal pelvic transitional cell carcinoma.20 Other primary renal tumors reported include renal adenocarcinoma, renal adenoma, and papillary tubular cystadenoma.9 The kidney may also be the site of metastatic disease such as malignant lymphoma.39 Stage IV lymphoma invaded the kidneys in 8 of 18 ferrets in one survey.2 Primary renal tumors can be incidental necropsy findings since they tend to be slow growing with little metastatic potential.69 Clinical signs reported in a 7-year-old ovariectomized pet ferret with pleomorphic renal adenocarcinoma included hematuria, dysuria, and incontinence, as well as nonspecific signs of illness such as anorexia, lethargy, and weight loss. Abdominal effusion and peripheral lymphadenopathy were detected on physical examination, and metastases to the lungs and pleura were found at necropsy. The neoplasm consisted of a large, solid mass and a cyst measuring 3 cm in diameter.34 Diagnosis of renal tumors relies on abdominal palpation, urinalysis, and imaging. Renal tumors usually appear as cystic areas on ultrasound, and may initially be mistaken for renal cysts.69 Definitive diagnosis is achieved via ultrasound-guided needle biopsy or at necropsy. Primary renal tumors may be treated with nephrectomy and surgical excision. There is little information on the use of chemotherapy in domestic animals, and no information is available for ferrets.
URETERAL RUPTURE Traumatic avulsion of the ureter was reported in a ferret with blunt trauma severe enough to also create a diaphragmatic hernia. No specific urinary tract signs or abnormal clinical pathologic findings were observed. Excretory urography was used to detect ureteral leakage, and treatment included ureteronephrectomy.69
CHAPTER 4 Disorders of the Urinary and Reproductive Systems
UROLITHIASIS Urolithiasis is characterized by solitary or multiple calculi found anywhere throughout the urinary tract or by the presence of sandy material within the bladder and urethra. Urinary calculi used to be a common cause of stranguria in ferrets; however, improvements in diet have made urolithiasis rare in ferrets on ferret food or high-quality cat food. Urolithiasis is also rare in working ferrets and pet ferrets on fresh meat diets in New Zealand, Australia, and Europe.37 The most common urinary calculus reported is magnesium ammonium phosphate or struvite. Dietary factors are believed to play an important role in struvite crystal formation. Urine pH is greatly influenced by diet, specifically by the source of dietary protein. Metabolism of animal protein tends to produce acidic urine, whereas plant-based protein diets, such as dog food or inexpensive cat foods, produce alkaline urine. Struvite crystals commonly form at urine pH exceeding 6.6. Significant crystalluria leads to the development of calculi or sandy material in the bladder and urethra. In one report, 6 of 43 ferrets (14%) fed dog food had renal or cystic calculi at necropsy.48 Urolithiasis is seen most commonly in adult males. Calculi may also be associated with ascending cystitis in pregnant jills, usually caused by urease-positive bacteria such as Staphylococcus or Proteus species.8,37 Mixed uroliths have been reported in ferrets including 60% struvite and 40% calcium oxalate,14 and there are also rare reports of cystine stones.18,21 The cause of cystine urolithiasis in ferrets is unknown, but has been speculated to be dietary or hereditary.
Diagnosis Obtain a complete history, including dietary history, from the owner. Clinical signs of urolithiasis include stranguria, dysuria, pollakiuria, urine dribbling, frequent licking of the prepuce, and hematuria. Ferrets with urethral obstruction may strain violently or cry when attempting to urinate, and owners may misinterpret the straining as “constipation.” Tenesmus may even lead to diarrhea in some cases. Occasionally, a ferret with blockage will be treated for lethargy, weakness, inappetance, and even collapse without obvious signs of dysuria. If not corrected, urinary obstruction can result in severe metabolic disturbances, coma, and death. Affected jills may be asymptomatic or show intermittent straining for days or weeks. Eventually the jill will show real distress when cystic calculi reach a large size. By this time, there may be evidence of urine dribbling and possibly vulvar scald. Although urethral obstruction is most common in male ferrets, females can also become obstructed, potentially straining hard enough to cause rectal or vaginal prolapse and potentially fatal hemorrhage.8,37,52 Cystic calculi or sand is often palpable in ferrets without obstruction, and a distended bladder is readily palpable in obstructed ferrets. Abdominal radiographs serve as a valuable diagnostic tool. Evaluate the entire urinary tract for radiodense uroliths and other abnormalities. Calculi lodged at the os penis can be difficult to detect. Use abdominal ultrasound to evaluate the urinary tract, prostate, and adrenal glands. In affected ferrets, submit samples for CBC, serum biochemistry analysis, urinalysis, and ideally urine bacterial culture and sensitivity. The reported range for normal ferret urine pH is
49
6.5 to 7.5.21 Urine should be more acidic in ferrets fed a highquality, meat-based diet.
Therapy If the ferret is not obstructed, provide supportive care, including parenteral fluids, and then schedule cystotomy to remove cystic calculi and flush the bladder. Submit calculi for mineral analysis, and send crushed calculi and bladder mucosa for bacterial culture and sensitivity. Renal calculi may be managed by antibiotics and dietary modification (see later discussion), unless clinical signs warrant surgical removal. Begin antibiotics after surgical removal of calculi or preoperatively if you suspect infection. Select a broad-spectrum antibiotic that reaches high levels in the urinary tract until culture and sensitivity results are available. Administer antibiotics for a minimum of 10 to 14 days, but use urinalysis and urine culture results to guide the duration of therapy. Continue antimicrobials several days past resolution of clinical signs. Treatment of urinary obstruction in male ferrets is a challenge. Place a urinary catheter (see “Urethral Obstruction” later in this chapter), then flush the urolith into the urinary bladder for future removal via cystotomy.20 Convert the ferret to an animal protein–based diet. Because a ferret on a high-quality diet has a urinary pH of approximately 6.0, urinary acidifiers are usually unnecessary. Attempts to feed feline magnesium-restricted acidifying diets (e.g., feline s/d [Hill’s Pet Nutrition, Topeka, KS] or feline Urinary SO [Royal Canin, St. Charles, MO]) are generally unsuccessful. These diets probably also contain insufficient protein for long-term use in ferrets. Use of a protein-restrictive diet for advanced renal disease (Hill’s Prescription diet u/d) has been described for dietary management of cystine urolithiasis. The ferret was also fed a protein supplement and hemoglobin and albumin levels were monitored.18 Two cases of cystine urolithiasis in which owners did not modify diet postoperatively have also been reported. Calculi did not recur postoperatively.20 The prognosis is good for urethral or cystic calculi with aggressive treatment. The long-term prognosis for bilateral renal calculi is guarded.21
CYSTITIS Spontaneous bacterial cystitis is rare in ferrets, although it is more common in jills than hobs. Bacteria commonly associated with cystitis include S. aureus, Proteus species, and E. coli. If cystitis is identified, screen the ferret for underlying disease such as adrenal-associated prostatomegaly (see later discussion) or urolithiasis (see earlier discussion). In rare instances, cystitis has also been associated with neoplasia such as transitional cell carcinoma.20 Infection may initially be asymptomatic, but by the time of treatment most ferrets demonstrate pollakiuria, dysuria, hematuria, and urine staining of the perineum. Ferrets with advanced cystitis may have anorexia, lethargy, weakness, or even collapse. The abdomen may be painful and the bladder wall may feel thickened. Obtain a sample for urinalysis and bacterial culture and sensitivity, ideally via cystocentesis. Urinalysis results may include hematuria, pyuria, bacteruria, and tubular casts. Normal ferret urine pH is approximately 6.0. CBC and serum biochemistry results are often unremarkable although an inflammatory leukogram may be seen with an ascending bacterial infection.
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SECTION I Ferrets
Ferret 3-0 urinary catheter Tip of os penis
Fig. 4-2 Urethral catheterization of a male ferret using a 3-French urinary catheter. Note the J-shape of the penis tip. (From Lennox AM, Lichtenberger M. A new type of urinary catheter for catheterization of the male ferret. Exot DVM. 2008;10:5-6.)
Because primary cystitis is rare, consider survey abdominal radiographs, abdominal ultrasound, and adrenal hormone analysis to identify the underlying cause of infection. Administer a broad-spectrum antibiotic until culture results are available, then adjust antibiotics accordingly. Continue treatment for a minimum of 10 days and for at least 4 weeks in complicated cases. Provide fluids and supportive care as needed and ensure that the ferret is fed a high-quality diet with a meatbased protein source (as for urolithiasis).
BLADDER NEOPLASIA There are rare reports of transitional cell carcinoma of the urinary bladder in ferrets. Initial signs are often subtle but may include hematuria, dysuria, polyuria, or incontinence. The urinary bladder may palpate abnormally, and excessive numbers of transitional cells, some abnormal in appearance, are seen on urinalysis. Use ultrasonography or contrast radiography to visualize bladder lesions. Definitive diagnosis is based on biopsy results. The prognosis is poor.58,69
URINARY INCONTINENCE True incontinence is rare in ferrets. More commonly, ferrets are treated for dribbling due to overfill of the urinary bladder as a result of urolithiasis or prostatomegaly. True incontinence may be associated with bladder neoplasia, severe cystitis, or Aleutian disease virus. Additionally, although rabies virus is extremely rare in domestic ferrets, bladder atony and incontinence were reported in 65% of experimentally infected ferrets.37,47
URETHRAL OBSTRUCTION Although the overall incidence of urethral obstruction is probably uncommon, it is an important reason for emergency treatment of the male ferret. In the United States, urethral obstruction is most frequently caused by adrenal-associated prostatic disease (see later discussion) or less commonly urolithiasis (see earlier discussion). On physical examination, the urinary bladder is distended and painful. Laboratory results may include azotemia, hyperkalemia, hyperphosphatemia, and metabolic acidosis. Urethral catheter placement is challenging in the male ferret due to its small size and J-shaped os penis (Fig. 4-2). Use general
Fig. 4-3 The Slippery Sam (top) and the Tomcat urethral catheter with stylet in place (bottom). Both catheters are flexible with a fine flexible stylet, and the latter has a rounded tip to facilitate introduction into the distal urethra. (From Lennox AM, Lichtenberger M. A new type of urinary catheter for catheterization of the male ferret. Exot DVM. 2008;10:5-6.) anesthesia to achieve adequate skeletal muscle relaxation. Mask induce ferrets with isoflurane or sevoflurane or administer etomidate (1 to 2 mg/kg intravenously [IV]) with diazepam (0.1 mg/kg IV) or propofol (4 to 6 mg/kg IV) with diazepam (0.25 to 0.5 mg/kg IV). After induction, intubate ferrets if jaw tone allows and maintain on inhalant anesthesia.40 Avoid ketamine because it is excreted by the kidneys. Provide supplemental heat and appropriate preemptive analgesia such as buprenorphine (0.01 to 0.03 mg/kg subcutaneously [SC], intramuscularly [IM], IV q8–12h), butorphanol (0.1 to 0.5 mg/kg SC, IM q4–6h or 0.025 to 0.1 mg/kg/hr IV), or fentanyl (2.5 to 5.0 μg/kg/hr IV).26,40 Whenever possible use a catheter designed for use in ferrets such as the 3.0-French 11-cm open-ended silicone catheter (Slippery Sam, Smiths Medical PM, Waukesha, WI) (Fig. 4-3).36 The Slippery Sam catheter is flexible with a rounded tip and fine flexible stylet to facilitate introduction into the distal urethra, which is difficult to visualize in this species (A. Lennox, personal communication, 2009). A 24-gauge catheter may also be used to identify and dilate the small, slit-like urethral opening. In addition to the Slippery Sam catheter, a 20- or 22-gauge 8-inch jugular catheter may serve as a makeshift urinary catheter, and a 3.5-French red rubber catheter may be placed in a large male (Fig. 4-4).43 Some clinicians routinely fit ferrets with an Elizabethan collar at the time of catheter placement. Secure the collar by criss-crossing gauze under the forelimbs in a figure-eight pattern.36 Patients alert enough to attempt to disrupt the catheter may also be maintained on low-dose opioids by injection or constant rate infusion (CRI) (M. Lichtenberger and A. Lennox, personal observation, 2009). Carefully monitor the electrocardiogram for evidence of hyperkalemia such as loss of the P wave, widening of the QRS complex, and peaked T waves. Relief of obstruction and forced diuresis are usually sufficient in the management of hyperkalemia. Medical treatment is indicated if an arrhythmia is present in addition to poor perfusion or altered mentation. Give calcium gluconate (50 to 100 mg/kg slow bolus IV) or insulin (0.2 U/kg IV) with 50% dextrose (1 to 2 g per unit insulin IV).40 To provide additional relief for urethral obstruction, some clinicians use smooth muscle relaxants such as diazepam
CHAPTER 4 Disorders of the Urinary and Reproductive Systems
Fig. 4-4 Urethral catheterization using a 3.5-French red rubber catheter. The catheter is sutured in place using butterfly tape. (Courtesy Dr. Marla Lichtenberger.)
(0.5 mg/kg PO, IM, IV q6–8h), midazolam (0.5 to 1.0 mg/kg SC, IM), or, in rare instances, alpha-adrenergic antagonists such as phenoxybenzamine (Dibenzyline, SmithKline Beecham, Philadelphia, PA; 3.75 to 7.5 mg PO q24–72h) (D. Mader, personal communication, 2010). Use alpha-adrenergic antagonists with caution because of the potential for adverse gastrointestinal and cardiovascular effects. Monitor urine production in catheterized ferrets using a closed line attached to a small (150 to 250 mL) IV bag. Urinary output must be at least 1 to 2 mL/kg/hr and with diuresis, urine production may be up to 140 mL/day.40 Maintain the urinary catheter for 1 to 3 days50 postoperatively while providing aggressive supportive care: correct metabolic and acid-base disturbances, and administer fluids to correct perfusion abnormalities and rehydrate the patient. Maintenance fluid requirements in ferrets are estimated at 75 to 100 mL/kg/day. Monitor patients carefully for signs of overhydration. If attempts at urinary catheterization are unsuccessful, consider cystocentesis to help stabilize the patient preoperatively,7 although there is some risk of urinary bladder rupture and uroabdomen.40 Perform an exploratory laparotomy and emergency cystotomy if needed. If normograde passage of a urinary catheter is unsuccessful, an emergency perineal urethrostomy is required. Tube cystostomy has also been described in ferrets with urinary obstruction (Fig. 4-5) (see Chapter 11).49
PROSTATIC CYSTS The prostate is the only accessory reproductive gland of the ferret. Whereas the canine prostate is a compact, relatively prominent enlargement, the ferret prostate is a small, fusiform mass measuring approximately 15 mm long by 6 mm wide. The ferret prostate surrounds the neck of the bladder and the proximal urethra, and is surrounded by a fibromuscular capsule from which numerous septae extend.25,30,38,50 Prostatic disease is the leading cause of urinary tract disease and urethral obstruction in middle-aged to older male castrated ferrets.52 Neutering of ferrets is routinely performed at 6 weeks of age in the United States,39 and adrenocortical disease in the ferret has been correlated with early gonadectomy. Adrenal
51
Fig. 4-5 Diagram illustrating percutaneous cystostomy tube. A Foley catheter is placed within the bladder and the bladder wall is sutured to the abdominal wall. (Courtesy Dr. Marla Lichtenberger.)
Fig. 4-6 The prostate is expanded by cysts of various sizes filled with keratin and proteinaceous debris. (From Coleman GD, Chavez MA, Williams BH. Cystic prostatic disease associated with adrenocortical lesions in the ferret. Vet Pathol 1998;35:547-549.)
tumors in gonadectomized ferrets are theorized to be due to chronic stimulation by luteinizing hormone.66 Outside of the United States, most ferrets are gonadectomized at several months of age, and adrenal disease is relatively uncommon. Prostatic disease is also uncommon in the intact hob.20 There are rare reports of prostatic disease associated with transitional cell carcinoma of the bladder.21 Adrenocortical disease in the male ferret is associated with the development of sterile prostatic cysts. Although the exact pathogenesis is unclear, elevated circulating sex steroid hormone levels appear to stimulate proliferation of prostatic tissue (see Chapter 7).12 Low to moderate numbers of neutrophils and fewer lymphocytes, macrophages, and plasma cells may infiltrate glandular tissue.12 Squamous metaplasia of prostatic ductular epithelium causes the prostate to expand with multiple, fluid-filled cysts of various sizes filled with keratin, neutrophils, and proteinaceous debris (Fig. 4-6). Prostatic cysts can measure
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SECTION I Ferrets
Fig. 4-7 The urinary bladder with two associated fat pads, one on either side, are displaced to the left. The prostatic cyst is on the right. (Courtesy Dr. Lauren Powers.)
up to 1 cm in diameter and they can compress the urethra, leading to urine stasis.12 Extensive prostatomegaly leads to urethral obstruction.
Diagnosis Although prostatic disease may be seen at any age, disease is most common in middle-aged (3 to 4 years) to older, neutered male ferrets. Signs of lower urinary tract disease secondary to prostatomegaly include pollakiuria, hematuria, stranguria, dysuria, or even anuria. Ferrets may lick excessively, creating a red prepuce. Owners may misinterpret signs of straining and present their pets for constipation. Tenesmus may even lead to diarrhea or, in rare instances, rectal prolapse. Signs of adrenal disease, such as alopecia, pruritus, or behavioral changes, may be seen but are frequently absent.3,49,52 The enlarged prostate may be palpable as a variably sized fluctuant mass dorsal to the urinary bladder and near the base of the bladder. The prostate may even be larger than the bladder (Fig. 4-7), although not all ferrets with prostatic disease demonstrate prostatomegaly.52 Ferrets with urethral obstruction may have a history of anorexia, lethargy, weakness, or even collapse. Do a full medical workup in affected ferrets. On survey radiographs, prostatomegaly may appear as a mass lesion caudodorsal to the urinary bladder, displacing the bladder cranioventrally. Use ultrasound to evaluate the adrenal glands, kidneys, urinary bladder, and prostate. Prostatic cysts contain hypoechoic to anechoic fluid. Collect cyst fluid via fine-needle aspirate for cytology and aerobic and anaerobic culture if cysts are overly large. Ideally cyst aspiration should be ultrasound guided in a sedated patient. If ultrasound is unavailable, contrast cystography may illustrate an irregular blockage around the bladder neck (Fig. 4-8). Use the ferret adrenal panel available through the University of Tennessee College of Veterinary Medicine Endocrinology Lab (www.vet.utk.edu/diagnostic/endocrinology/index. php) to confirm the presence of adrenocortical disease, particularly if ultrasound results are equivocal.
Treatment Treatment involves management of urethral obstruction if present (see earlier discussion) and management of adrenocortical disease (see Chapter 7). Surgical therapy involves exploratory
Fig. 4-8 Contrast cystography illustrating irregular mass lesions near the bladder neck due to prostatitis. Note the caudal extension of the inflammed prostate gland.
laparotomy and adrenalectomy. Surgical debulkment is sometimes necessary for large prostatic cysts. Cystic prostatic hypertrophy begins to resolve within 2 to 3 days postoperatively.3 Drug therapy may serve as an adjunct or an alternative to surgery in patients that are a poor surgical risk or when there are financial constraints. Medical inhibition of gonadotropin production can reduce sex steroid hormone production and improve clinical signs, but medical management does not alter adrenal tumor growth. Therapies are extrapolated from drug use in other species, and there is considerable individual variation in clinical response.66 The depot formulation of the synthetic gonadotropin releasing hormone (GnRH) agonist, leuprolide acetate, is most commonly used (100 to 250 μg/kg/month IM).33,62,66 In many individuals, high-dose leuprolide acetate administration causes prostatic tissue to shrink within 12 to 48 hours so that the ferret may even begin to urinate around the urethral catheter. Other GnRH agonists, such as goserelin acetate or a slowrelease deslorelin acetate implant (3 mg SC), have also been used, particularly in countries where leuprolide is not readily available.62 Use ferret adrenal hormone panel results to guide selection of other drug therapies. Some veterinarians have advocated the use of the antiandrogen agent flutamide (10 mg/kg PO q12–24h). Effects may be seen within days in some ferrets, but use caution in patients with liver disease. Other antiandrogen agents used include bicalutamide (5 mg/kg PO q24h) in conjunction with leuprolide or finasteride (5 mg/kg PO q24h). The antiestrogen agent anastrazole (0.1 mg/kg PO q24h) may be useful as an adjunct to leuprolide when excess estradiol levels are present.33 If left untreated, prostatic disease can lead to urethral blockage, acute renal failure, and death. Fortunately, the prognosis for prostatic cysts is generally good in ferrets that undergo aggressive therapy for underlying adrenocortical disease. Ensure that prostatic cysts resolve completely by following their sonographic appearance. Long-term persistence of cysts may promote the development of bacterial prostatitis and abscesses. Also monitor patient progress by following sex steroid hormone levels.
CHAPTER 4 Disorders of the Urinary and Reproductive Systems
Fig. 4-9 Gross appearance of a prostatic abscess. The bladder is the smaller structure to the left. (Courtesy Dr. Sandra Mitchell.)
PROSTATITIS AND PROSTATIC ABSCESS Pathogenesis of bacterial prostatitis and prostatic abscess is poorly understood in the ferret. Urine stasis secondary to adrenal-associated prostatomegaly may promote bacteruria and regional migration of bacteria into prostatic tissue (see “Prostatic Cysts” earlier in this chapter). Bacteria may also enter the prostate hematogenously. Prostatitis has been described in a ferret with Sertoli cell tumor associated with a retained testicle and there are rare reports of prostatic abscesses associated with transitional cell tumors of the bladder.56 Prostatitis has been associated with heavy growth of Staphylococcus species, nonhemolytic Streptococcus species, E. coli, Proteus species, and Pseudomonas species. There have also been reports of clostridial infections in canine paraprostatic cysts.56
Diagnosis Prostatic disease may be seen at any age, but disease is most common in middle-aged to older, neutered male ferrets. A thick, white to yellow, opaque penile discharge may be seen, and purulent discharge may also be associated with urination. Round structures may be palpable near the urinary bladder (Fig. 4-9) and the abdomen may be swollen, tense, and/or painful (see “Prostatic Cysts” earlier in this chapter for additional clinical signs of prostatic disease).52,56 An inflammatory leukogram may be observed, although serum biochemistry is frequently unremarkable.56 Urinalysis results are variable. Many ferrets have concurrent bacterial cystitis, but ferrets may have prostate infection without evidence of bacteria or inflammation in the urine.50 Abdominal ultrasound of the prostate shows hypoechoic to anechoic fluid-filled cysts with hyperechoic sediment. Perform fine-needle aspiration of prostatic cysts, ideally using ultrasound in a sedated patient. Aspirated fluid may be turbid or flocculent and range from yellow to green in color.56 Cytology will reveal suppurative exudate. Submit the sample for aerobic and anaerobic bacterial culture and sensitivity.
Treatment Select a lipid-soluble antibiotic known to penetrate the prostatic capsule such as potentiated sulfas or fluoroquinolones. Use culture and sensitivity results to adjust therapy and administer
53
antibiotics for at least 4 to 6 weeks.3,4,7 Although antimicrobials are an important ancillary treatment, antibiotics generally fail to achieve effective tissue levels throughout the prostate, and surgical management of prostatitis or prostatic abscess is recommended. Perform exploratory laparotomy and adrenalectomy as indicated. Both marsupialization and omentalization have been advocated in the surgical management of prostatic abscess (see Chapter 11).3,4,7 In marsupialization, the infected prostate is flushed and marsupialized to the abdominal wall. Marsupialization allows for repeated lavage. Because drainage is external to the abdominal cavity, the risk of peritonitis may be reduced.3,52 Omentalization is currently the surgical technique of choice for management of prostatic abscess in dogs,3,4 and this procedure has been described in the ferret.3,7,52,56 The prostatic abscess is opened and flushed, and as much of the cranial capsule is excised as possible. The caudal aspect of the abscess may extend deep into the pelvic canal.56 The remaining abscess cavity is irrigated, then a portion of greater omentum is inserted into the abscess and sutured into place. Omentum promotes tissue adhesion, angiogenesis, hemostasis, peritoneal lymphatic drainage, fibrinolysis, and immune system function.29 Regardless of the surgical technique selected, potential complications include recurrent cystitis and peritonitis.4 Prostatic cysts may extend into bladder or urethral tissue, increasing the risk of urinary incontinence, uroabdomen, or urethrocutaneous fistula formation.4 No complications were reported with marsupialization in five ferrets, although permanent cystostomy was reported in one ferret in another case report.4,52 Prognosis is fair to poor for prostatic abscess when disease is managed aggressively. The prognosis worsens as the size of the lesions increases.50 Culture prostatic fluid and urine 2 to 4 weeks after completion of antibiotic therapy, and use serial abdominal ultrasound to follow the progress of the prostatic abscess. Follow underlying adrenal disease with sex steroid hormone levels.
PROSTATIC TUMORS Prostatic tumors such as prostatic seminoma and carcinoma have been reported. The prognosis is poor.58
PARAURETHRAL CYSTS OR PARAURETHRAL DISEASE Paraurethral cysts are a rare lesion reported in six adult ferrets (four males, two females).38 There is also a separate case report in a 3-year-old female spayed ferret. Single or multiple, semispherical to bilobulated, fluid-filled, thin-walled cysts of variable size were reported on the dorsal surface of the urinary bladder trigone region and the proximal urethra. Variable intraluminal communication existed between the cysts and the bladder and/ or urethra. Paraurethral or bladder cysts arise from embryonic remnants of the urogenital ducts, müllerian structures, or mesonephric duct. In the single female ferret, the cystic paraurethral mass was described as reproductive in origin and was presumed to be a remnant of the cervix.52 Clinical signs included dysuria and hematuria in three of six ferrets, as well as alopecia and vulvar swelling. On physical examination, cysts were palpable as a fluctuant mass caudodorsal to the urinary bladder. Adrenal hyperplasia or neoplasia was detected in five of six ferrets; the adrenal gland was not evaluated
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SECTION I Ferrets
in the sixth ferret. Although treatment was not attempted in the six ferrets, initial therapy would involve relief of urethral obstruction, hydration, and treatment of secondary bacterial infection. Definitive therapy for paraurethral disease is surgical. In the single female ferret, a compartmentalized mass containing purulent material was identified and marsupialized at exploratory surgery. Bacterial culture of the material was negative.38,52
DISORDERS OF THE REPRODUCTIVE SYSTEM Primary reproductive tract disease is uncommon in the United States where ferrets are routinely desexed at 6 weeks of age.20 Outside of the United States, most ferrets are gonadectomized at 5 to 6 months of age, if at all, and therefore disease of reproductive organs and prolonged estrus seasons are much more common.37 Ferrets normally reach puberty between 4 and 12 months of age. The onset of puberty is earlier when ferrets are exposed to long days of more than 12 hours of light.8,41 Gonadal activity is seasonal and the breeding season occurs from September to December in the Southern Hemisphere and March to August in the Northern Hemisphere with peak breeding activity in April.1,37,62 Ferrets are “long-day” breeders and greater than 12 hours of light promotes reproductive activity.1
THE MALE FERRET Plasma testosterone levels increase at the end of January in the Northern Hemisphere and a peak plateau is maintained from the end of February until late July. Increases in testicular size begin in late January and peak in April.20 The stages and the cycle duration of ferret spermatogenesis are similar to those reported in other carnivores.45
Cryptorchidism Ferret testes usually descend into the scrotum during fetal development, but complete descent may take several months after birth.20 Cryptorchidism is uncommon and was reported in less than 1% of 1,597 male ferrets.11
Tumors of the Male Reproductive Tract Testicular tumors have been reported in middle-aged to older ferrets,25 most commonly in cryptorchid testes.69 Reported neoplasms include Leydig or interstitial cell tumor, seminoma, Sertoli cell tumor, and carcinoma of the rete testis.25,28 Metastasis to the liver was reported in one ferret with a Sertoli cell tumor.69 Tumors may be associated with unilateral or bilateral testicular enlargement. Swelling may be fluctuant, soft, or firm.25,39 Increases in sex steroid hormone levels may also increase sexual behavior and aggression, and increased dermal sebaceous gland activity creates a greasy coat and a distinctive musky odor. Total body alopecia and severe pruritus were described in a ferret with Sertoli cell tumor, presumably due to hyperestrogenism.69 One report describes Sertoli cell tumor in a ferret with elevated levels of estradiol and 17-OH progesterone. Androstenedione levels were normal.56 The treatment of choice for testicular tumors is castration. Preputial gland mass lesions reported in intact and neutered male ferrets include preputial gland adenoma, adenocarcinoma (Fig. 4-10), and apocrine cysts.39,69 These tumors are located in the subcutis and appear white, pink, or darkly pigmented. The treatment of choice is careful surgical resection.39
Fig. 4-10 Preputial gland adenocarcinoma (arrows) in a 4-yearold castrated male ferret.
Prostatic Lesions See “Disorders of the Urinary System” earlier in this chapter.
Penile Lesions Penile lesions are occasionally seen in male ferrets kept on hay bedding when grass awns catch in the prepuce. Breeding males may also incur injury as they move along the ground pressing their groin down to mark their territory. Remove foreign bodies, ideally using general anesthesia, and apply a topical antibioticcorticosteroid ointment.37
THE FEMALE FERRET Female ferrets are seasonally polyestrous. Proestrus usually occurs in January or February,20 and is signaled by increasing vulvar tumescence.41 The vulva can increase up to 50 times its normal size. Proestrus is characterized by a rising number of superficial epithelial cells on vaginal cytology. Estrus is typically seen between late March and early August. The jill may eat and sleep less and she may become irritable. In addition to an enlarged, edematous vulva,20 there may be some blood associated with estrus.37 Superficial epithelial cells make up 90% or more of epithelial cells in vaginal cytology, and after several days, cells are fully keratinized. Neutrophils are common during all stages of the estrous cycle. Until mated, jills may exhibit constant estrus for up to 5 months and hyperestrogenism (see next section) may occur.20,41
CHAPTER 4 Disorders of the Urinary and Reproductive Systems Ferrets are induced ovulators and coital stimulus is required to trigger ovulation.1 Once ovulation is induced, either pregnancy or pseudopregnancy occurs. By several days following induced ovulation, superficial vaginal cells decline in numbers to anestrus levels and the vulva decreases in size and turgidity.41
Hyperestrogenism Hyperestrogenism, also known as estrogen toxicity or postestrus anemia,37 has become uncommon in the United States where breeding farms routinely spay ferrets at 6 weeks of age. As induced ovulators, approximately half of estrous female ferrets remain in estrus until bred or artificially stimulated to ovulate.41 Estrus lasting more than 1 month is associated with hyperestrogenism, which leads to bone marrow depression and hypoplasia of all cell lines.28 In one survey of 20 females left in heat, the first death related to aplastic anemia occurred 1 month after estrus began. Twelve ferrets became anestrous and eight ferrets died, with the last death occurring 6 months later.64 In addition to persistent estrus, other potential causes of estrogen toxicity include cystic ovary and an estrogen-secreting ovarian remnant,37,59 which are also sometimes cystic. Adrenocortical disease rarely leads to hyperestrogenism and only after long-standing disease has remained untreated. Diagnosis and Treatment. Hyperestrogenism is usually seen in jills over 1 year of age. Clinical signs may include anorexia, lethargy, vulvar swelling, vulvar discharge, pallor, and systolic murmur. Extended hyperestrogenism may cause melena, petechial or ecchymotic hemorrhages, and dorsal bilaterally symmetric alopecia. Concurrent metritis, pyometra, or vaginitis may also be seen. Pelvic limb paresis, ataxia, and paralysis have been reported secondary to subdural hematoma.25 Early CBC results may show thrombocytosis and neutrophilic leukocytosis; however, by the time of presentation the CBC will generally reveal nonregenerative anemia (hematocrit less than 20% to 25%), nucleated red blood cells, neutropenia, and thrombocytopenia (platelet count less than 50,000/μL).28,58,64 Hemorrhage occurs when platelet counts fall below 20,000/μL.64 Confirm nonregenerative anemia with a reticulocyte count. When results of the CBC or reticulocyte count are equivocal, aspirate bone marrow for cytologic evaluation. Ferrets usually die from hemorrhage secondary to thrombocytopenia. Necropsy findings may include pale mucous membranes and bone marrow, hematomyelia, hydrometra, and petechial, ecchymotic, or subcutaneous hemorrhages.25,58 Neutropenia may predispose jills to bacterial infections such as pyometra, vaginitis, and bronchopneumonia. Histologically, centrilobular hepatic degeneration, panmyelophthisis, and cystic endometrial hyperplasia may be observed.25 The primary goal of therapy is to reduce estrogen levels. Definitive treatment calls for ovariohysterectomy or surgical removal of an ovarian remnant. Ovarian remnants may be found near the ovarian pedicle or within mesenteric fat. Because these ferrets are often poor surgical and anesthetic risks, aggressive supportive care is often required preoperatively. Give blood transfusions as needed. Ferrets do not possess blood groups, and multiple transfusions from multiple donors are possible.58 The hemoglobin-based oxygen carrier Oxyglobin (Biopure Corporation, Cambridge, MA; 11 to 15 mL/kg IV over a 4-hour period) can be used in anemic ferrets.51 Less commonly, bone marrow transfusions have also been described. Additional supportive care measures may include anabolic steroids, corticosteroids, and iron dextran. Erythropoietin has been tried but
55
is reportedly ineffective, possibly because of inhibition by estrogen.51,58 Medical manipulation of hyperestrogenism is most likely to be successful when estrogen has been elevated for less than 4 weeks. Administer GnRH (20 μg/ferret SC, IM) or human chorionic gonadotropin (hCG) (20 to 100 IU/ferret IM).25,37,58 Give hCG at least 10 days after the onset of estrus, then repeat in 7 days if vulvar swelling has not resolved.53,58 The use of second-generation progestogens such as proligestone (Covinan, Intervet Australia Schering-Plough Animal Health, Victoria, Australia) (50 mg SC) has been proposed when the jill has been in estrus for 10 days. Proligestone has also been recommended in the United Kingdom to prevent estrus just before breeding season.37 A jill may also be converted to anestrus by breeding her with a vasectomized male or by mechanically inducing ovulation with a vaginal probe such as a cotton-tipped applicator. The results of these procedures are unpredictable and often unsuccessful in moribund ferrets. The prognosis for estrogen toxicity has classically relied on hematocrit (Hct) value. Hct greater than 25% is associated with a fair to good prognosis, Hct between 15% and 25% is associated with a guarded prognosis, and Hct less than 15% is associated with a poor to grave prognosis. Estrogen toxicity caused by persistent estrus is prevented by routine spaying of all female ferrets not intended for breeding by 6 months of age. At surgery, carefully identify all ovarian tissue that may be located within large fat deposits. Prevent breeding jills from remaining in estrus for more than 2 to 4 weeks.37
Tumors of the Female Reproductive Tract Tumors of the female reproductive tract are relatively common in intact female ferrets. In a retrospective study of 4,774 ferrets, 2.3% of 639 tumors involved the reproductive system,39 with the most common being ovarian leiomyoma.38 Other tumors reported include uterine leiomyoma, leiomyosarcoma, fibroleiomyoma, papillary adenocarcinoma, ovarian and uterine teratoma, luteoma, techoma, uterine adenoma, fibromyoma, fibrosarcoma, granulosa cell tumor, arrhenoblastoma, dysgerminoma, sex cord stromal tumors, and undifferentiated carcinoma of the uterine stump.69 Tumors attributed to the ovarian pedicle or ovarian remnants include granulosa cell tumor, leiomyoma, and fibrosarcoma. Ovarian tumors are commonly identified as incidental findings during ovariohysterectomy since affected ferrets may be asymptomatic.69 If clinical signs are seen, they may include lethargy, anorexia, and persistent estrus (vulvar swelling, anemia, endocrine alopecia). Older jills may breed but fail to conceive.37 Teratomas may become large enough to palpate during physical examination or identify on survey radiographs.69 Case reports involving tumors of the ovarian pedicle have documented elevated levels of androstenedione and 17-hydroxyprogesterone in one account and elevated estradiol and progesterone in another.32,55 Metastasis is generally not seen with reproductive tract tumors and surgical excision is generally curative.69 Ovariohysterectomy is the treatment of choice.58 Pyometra and Mucometra. Pyometra is uncommon in clinical practice in the United States where ferrets are spayed at 6 weeks of age, but this condition is often documented in intact females, particularly jills with estrogen toxicity or in older, pseudopregnant jills.37 Pyometra may occur secondary to ascending
56
SECTION I Ferrets
infection of the vagina.20 Various bacteria have been cultured including E. coli, Staphylococcus, Streptococcus, and Corynebacterium species.25 Some authors have documented pyometra and mucometra only during the breeding season, and it has been proposed that pyometra in the ferret may be under the influence of estrogens and not progestins.62 Pyometra may be open or closed and clinical signs may include lethargy, depression, and sometimes fever. An enlarged uterus may be palpable and purulent vaginal discharge may be present.25 Stump pyometra is occasionally seen due to elevated sex steroid hormone levels secondary to adrenocortical disease. Stump pyometra was reported in a 4-year-old spayed female ferret. Clinical signs included endocrine alopecia, pollakiuria, and mild vulvar swelling. A turgid mass was palpable on physical examination. Peritonitis developed as a result of fine-needle aspiration of the mass.22 Determine whether the ferret with pyometra is suffering from estrogen-induced bone marrow depression (see “Hyperestrogenism” earlier in this chapter).25 Definitive treatment of pyometra or mucometra involves ovariohysterectomy or surgical excision of the infected stump and adrenalectomy. Provide systemic antibiotics and supportive care such as fluids, analgesia, and anti-inflammatories. Medical management of pyometra has also been described. Give prostaglandin F2-alpha (Lutalyse, Pharmacia and Upjohn, Kalamazoo, MI; 0.1 to 0.5 mg IM), followed in 1 hour by oxytocin (5-10 IU IM) to stimulate myometrial contraction and expulsion of pus.26,37 Hydrometra. Hydrometra is the accumulation of aseptic fluid within the uterus in the presence of persistent corpora lutea. Hydrometra in association with ovarian tumor is one of the most commonly diagnosed uterine diseases in ferrets (Figs. 4-11 and 4-12).55 Segmental atresia of the uterus associated with hydrometra has also been reported in a 2-year-old ferret.6 Definitive treatment relies on ovariohysterectomy. Vaginitis. Vaginitis is sporadically seen in female ferrets.25 This condition may be associated with anything that causes chronic and/or severe vulvar swelling such as hyperestrogenism caused by persistent estrus, ovarian remnant disease, or adrenal disease. Less commonly, vulvar swelling develops with cystitis, crystalluria, or even aggressive hob mating behavior.20 Poor husbandry and inadequate sanitation may also promote vaginitis in breeding jills kept on particulate bedding when hay, straw, or shavings adhere to the swollen vulva during estrus. Vaginitis is associated with overgrowth of bacteria such as E. coli, Staphylococcus, Streptococcus, Proteus, or Klebsiella.25 Vaginitis may be differentiated from the swelling associated with estrus by history, vaginal cytology,20 and the presence of mucopurulent vaginal discharge. Concurrent metritis and fever are occasionally present.23 Ascending vaginitis can also lead to a secondary cystitis. Give systemic antibiotics and resolve the underlying problem(s). Reduce estrogen levels surgically by adrenalectomy or removal of the ovarian remnant, or medically with the use of hCG or leuprolide acetate.
thrusts last a variable length of time up to 3 minutes. Between pelvic thrusts there are periods of rest in which the hob simply lies over the jill still gripping her neck. Total mating time lasts on average 1 hour (range: 15 minutes to 3 hours).41 Ferrets are induced ovulators. Ovulation is induced by pressure on the cervix, which leads to endogenous release of luteinizing hormone (LH). This LH surge stimulates preovulatory follicles to mature with an average of 12 oocytes (5 to 13) per female ovulated 30 to 40 hours after copulation. Because the ovary is encapsulated within a fatty bursa that completely encloses the ovary, ovulated eggs cannot be shed into the abdomen. Ferret oocytes are most capable of being fertilized up to 12 hours after ovulation with a maximum of 30 to 36 hours. Embryos enter the uterus over several days starting on day 5. Implantation is central, with rapid invasion of the uterine epithelium by the trophoblast over a broad area that eventually becomes a zonary band of endotheliochorial placenta.41 Approximately 1.5 weeks before whelping, the jill loses her hair coat and hairless rings develop around her teats. Gestation lasts 41 days (range: 39 to 42 days).1,8,41 Kits usually die in utero around day 43.37 The gestation period is slightly shorter for primiparous jills. Parturition normally lasts approximately
Fig. 4-11 Gross image of hydrometra in association with an ovarian tumor (far right). (Courtesy Dr. Vladimir Jekl.)
PERIPARTURIENT DISEASE NORMAL BREEDING Mating begins with genital and neck sniffing. When ready to breed, the receptive jill becomes flaccid and submissive, allowing the hob to grasp her nape with his teeth and grip her body by wrapping his forelegs around her ribcage. Repeated pelvic
Fig. 4-12 Survey radiograph of hydrometra in an adult female ferret. (Courtesy Dr. Vladimir Jekl.)
CHAPTER 4 Disorders of the Urinary and Reproductive Systems 2 to 3 hours.8 Approximately 5 kits are born per hour, although some jills take longer. Progress should be steady, and no signs of distress should be seen. The jill usually will not begin to nurse her litter until all kits are born; she will then lie down in a semicircular position on her side.8,37 The domestic ferret gives birth to an average of 8 or 9 kits (range: 7 to 15).1,41 Newborn kits are altricial and weigh 6 to 12 g at birth.8 Females will return to estrus within 2 weeks after weaning if exposed to the appropriate photoperiod. If kits are removed at birth, the jill will return to estrus 8 weeks after mating as do pseudopregnant females and females with resorbed fetuses. If a jill gives birth to a small number of kits (five or fewer), she may return to estrus while nursing.24
MANAGEMENT OF BREEDING FERRETS Successful breeding requires constant supervision during gestation, parturition, and lactation. Breeders should monitor jills closely for any change in appetite or body condition, particularly late in gestation.5,8 Place the pregnant jill’s cage in a quiet area well before parturition. Minimizing stress is particularly important in young, primiparous jills. Avoid environmental stressors such as excessive heat. Room temperature should not exceed 70°F (21°C). Place a heat lamp over only part of the nesting box so that the jill and kits can select warmth as necessary. The risk of poor mothering also increases with crowding or unusual noise and activity nearby. Jills become excited and may bury the kits in bedding, or place them in a pile in a corner or in food or water containers. Some jills will cannibalize the first few or all of their kits as they are born. Handling kits does not appear to cause rejection of their litter.8 The jill must also be able to easily enter and exit the nesting box without traumatizing her mammary glands. Shredded aspen shavings, recycled paper bedding, or small cloth towels are practical choices for whelping nest substrate. Do not use large towels because kits can get lost in them.8 Feed ferrets proper nutrition containing 36% to 40% animalbased protein and 18% to 20% fat.24 Make food and fresh water available at all times.37
DISEASES OF THE JILL Pregnancy Toxemia Pregnancy toxemia is a life-threatening disease seen sporadically in jills during late gestation, particularly young, primiparous jills.5,20 Negative energy balance promotes abnormal energy metabolism and subsequent hyperlipidemia, hypoglycemia, ketosis, and hepatic lipidosis during the 10 days before parturition. This energy deficiency is caused by either inadequate dietary intake or excess demand for nutrients due to an exceptionally large litter (15 or even 20 kits).5,8,37 Pregnancy toxemia is observed in late gestation. Common clinical signs include acute onset of severe lethargy, anorexia, dehydration, weight loss, and excess shedding.5,13,25,37 There may also be diarrhea and possibly melena. Pregnancy toxemia is also a possible cause of sudden death in jills.5,8,25 Hematologic and biochemical abnormalities may include anemia, hypoproteinemia, azotemia, hypocalcemia, hyperbilirubinemia, elevated liver enzymes, and hypoglycemia.5,25 Urinalysis may reveal ketonuria. In a survey of 10 jills with pregnancy toxemia, the three surviving jills were not anemic and
57
they had less pronounced azotemia, hypoproteinemia, and liver enzyme activity increases. Hepatic lipidosis was observed grossly in all jills that died.5 Provide aggressive treatment in the form of cesarean section and intense supportive care. Correct fluid and electrolyte imbalances and provide nutritional support. Consider esophagostomy tube placement for frequent force feedings. Gastroprotectants are commonly indicated because of the high incidence of concurrent gastrointestinal ulcers. Postoperatively, surviving jills frequently produce no milk for at least several days and cross fostering of kits is often necessary (see “Agalactia” later in this chapter).8 Because of the guarded prognosis, prevention through client education is much more effective than treatment. Advise breeders to provide consistent, proper nutrition, avoid stressors, and be vigilant for any changes in jill appetite or body condition.5 Make palatable food available at all times, and owners should provide several small food dishes so that accidental spillage or contamination does not restrict the jill’s intake. During the last week of gestation, even one overnight fast can induce toxemia in a jill with a large litter. Provide plenty of fresh water since jills will also stop eating without adequate water. Many breeders also give nutritional supplements, especially to jills predicted or known to have large litters (Nutrical, EVSCO Pharmaceuticals, Buena, NJ; Ensure Plus, Abbott Laboratories, Columbus, OH).8,37
Dystocia Dystocia is defined as labor exceeding 12 to 24 hours or whenever there are signs of difficulty. Dystocia occurs at a rate of about 1% in a large group of ferrets. Potential causes of dystocia include pregnancy in an older jill or an elevated environmental temperature (above 70°F [21°C]). Kits of very large size (more likely with small litter size), posterior or sideways presentation, and deformed or anasarcous fetuses may also promote dystocia.8 Depending on the underlying cause of dystocia, jills may be managed medically with oxytocin (5-10 IU IM) or surgically via cesarean section.37
Pseudopregnancy False pregnancy is caused by failure of implantation, which is associated with reduced light intensity 1 month before the start of breeding, or failure to conceive.25,37 For instance, jills may be bred with infertile hobs, particularly males less than 6 months of age. Pseudopregnancy has also been associated with termination of estrus through hCG use or mating with vasectomized hobs.20 In pseudopregnancy, the physical and behavioral changes normally associated with pregnancy may be seen such as weight gain, mammary gland enlargement, and nesting behavior. Jills may also mother inanimate objects. One important difference is that pseudopregnant jills develop a full, beautiful hair coat approximately 1.5 weeks before “whelping” whereas pregnant jills lose their coat and develop hairless rings around their teats. After “whelping,” pseudopregnant jills return to estrus if it is early in the breeding season, or they become quiescent if it is late in the breeding season.8,20,37 Minimize the risk of pseudopregnancy by ensuring jills are exposed to maximum light intensity during the spring, and artificially extend light hours in late summer. Also use mature, sperm-tested hobs for mating (older males will have higher sperm counts).37
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SECTION I Ferrets
Agalactia Agalactia is defined as failure of the jill to lactate enough milk for a normal litter of eight or nine kits. Agalactia may be seen when the jill is stressed because of environmental factors such as overcrowding, noise, visits from strangers, or overheating (temperature exceeding 70°F [21°C]). Agalactia is also more common in litters with less than five kits and in jills on a poor diet or suffering from systemic disease or chronic mastitis.8,20 Since the jill will not begin to nurse her litter until all kits are born, any delay in nursing such as dystocia or prolonged delivery will also predispose the jill to lactation failure.37 Agalactia leads to thin kits that cry and move around restlessly. Determine the underlying cause while offering the best-quality diet available to lactating jills.8
Mastitis Mastitis is occasionally seen in pet ferrets and frequently in laboratory ferrets.25 Acute mastitis develops soon after whelping or after the third week of nursing when milk production peaks and kits become more aggressive feeders. Because kits demand greater quantities of milk, this may stress the jill or their teeth can damage the nipples. Potential etiologic agents may include Staphylococcus species and coliforms such as hemolytic E. coli.8,25 The inguinal mammary glands are most commonly affected with firm, painful swelling in one or more glands and red or purple discoloration of the overlying skin.37 Milk may be discolored and clotted. If not recognized and treated early, infection will spread rapidly to nearby glands.25 Abscesses may form in severe cases. Acute mastitis often becomes gangrenous within a few hours. The skin turns black and the jill becomes toxic with signs of systemic illness such as depression, fever, anorexia, and dehydration.25 Chronic mastitis may develop secondary to acute mastitis, or more commonly this condition develops insidiously 3 weeks postpartum. Microorganisms such as Staphylococcus intermedius may enter secondary to trauma. Clinical signs may be quite subtle. Mammary glands appear to be full of milk, but most glandular tissue has been replaced by scar tissue leading to glands that palpate firm, but are not painful or discolored. The most obvious sign of chronic mastitis is poor kit condition. Kits will continue to grow in size but not weight.8,44 Begin systemic antibiotics such as clavulanic acid–amoxicillin or chloramphenicol for both acute and chronic mastitis.8,29 For acute mastitis, culture milk before starting antibiotics and apply warm compresses to affected glands. More aggressive treatment is indicated for gangrenous mastitis: perform wide surgical resection of the involved gland and adjacent tissue in combination with analgesia, fluid therapy, nutritional support, and systemic antibiotics.8,29,47 The kits will also require nutritional support. One option is to remove and hand-feed kits, although neonates are very difficult to hand-raise (see “Caring for Ill Kits” later in this chapter). If kits must be left with the dam, provide supplemental feedings at least three or four times daily. These kits will occasionally develop diarrhea, and should then be placed on the same antibiotic selected for the jill. Kits are also at risk for neonatal conjunctivitis (see later discussion). Kits may also spread bacteria to unaffected glands.8,20,37 Finally, kits may also be fostered, but there is significant risk of infecting the foster dam. Both acute and chronic mastitis are extremely infectious. Suggestions for minimizing risk include bathing kits and waiting a few hours before placing kits with the foster mother so their gastrointestinal tracts
have emptied of infected milk. Some clinicians also routinely treat kits and foster mother with oral antibiotics.8 Isolate affected jills from other breeders. Feed and treat mastitis jills and kits last. Human handlers should also wash their hands after handling kits so that they do not inadvertently transmit infection.8 The prognosis for mastitis is guarded. Jills that survive acute mastitis may lose a gland or they may be predisposed to recurrent mastitis. In chronic mastitis, microorganisms may be sensitive to multiple antimicrobials in vitro but are rarely effective in vivo.8
Mammary Gland Neoplasia Mammary gland tumors are a rare differential diagnosis for severely swollen, firm glands in domestic ferrets.25,69 Mammary gland tumors are similar to those seen in cats and dogs, and can vary from adenomas to carcinomas. A mixed mammary tumor was reported in a 7-year-old female ferret.25
Postparturient Hypocalcemia Postparturient hypocalcemia, or “milk fever,” was reported on a commercial ferret farm in New Zealand. The condition was seen in primiparous jills 3 to 4 weeks postpartum. Signs included posterior paresis, sensitivity to stimuli, and seizure activity. A rapid clinical response was achieved with intraperitoneal calcium borogluconate injections. Prevent hypocalcemia by providing dietary calcium supplementation.25
Metritis Metritis is defined as inflammation of the uterus that develops in the immediate postpartum period and occasionally after abortion or breeding. Metritis may also be associated with retained fetuses or placentas. On physical examination, the uterus is distended and a red vaginal discharge may be seen. Definitive treatment involves ovariohysterectomy. Medical management of metritis relies on the use of analgesia and prostaglandin F2-alpha (Lutalyse, 0.5 mg IM) to evacuate the uterus, and systemic antibiotics. Select an antimicrobial that reaches high urinary levels to reduce the risk of ascending cystitis and secondary urolithiasis.8,37
DISEASES OF THE KIT The Normal Kit Ferret kits are altricial with little ability to maintain a normal body temperature for the first 2 weeks of life. A healthy litter normally lies quietly close to the jill, nursing and sleeping except when the jill leaves the nest. When kits reach 3 weeks of age they begin to explore and nibble on soft food, even though their eyes are still closed. Kits generally weigh 8 to 10 g at birth. Body weight is 30 g by 1 week, 60 to 70 g at 2 weeks, and 100 g at 3 weeks. Eyes usually open at day 30 to 35, although eyes may open as early as day 25. Kits are weaned at 6 to 8 weeks.8
Caring for Ill Kits First provide supplemental heat, offering kits a temperature gradient of 85°F to 104°F (30°C to 40°C).42 Hypothermic kits do not nurse, so it is prudent to also administer a few drops of 50% dextrose solution by mouth or to rub Karo syrup on the gums.8 Then give warmed subcutaneous fluids (50 to 100 mL/kg). Feed the kit only once it is normothermic and hydrated. Neonatal ferrets have voracious appetites and are difficult to hand-rear. Thirty-six attempts to hand-rear neonates resulted
CHAPTER 4 Disorders of the Urinary and Reproductive Systems in only a single survivor.42 Kits require ferret milk for the first 7 to 10 days of life.8,42 The composition of ferret milk varies over the course of lactation (Table 4-1).63 Some success has been described sip-feeding neonates with pipettes every 1 to 1.5 hours. By day 10 to 21, kits may be offered puppy or kitten milk replacer four to six times daily.24 Some authors recommend enriching milk replacer with cream until fat content is 20%, and additional hand-feeding recipes are available.8,37,42 At week 4 to 5, offer a slurry of milk replacer and kibble mixed with solid food. Kits are weaned at 6 to 8 weeks.8 Because hand-rearing is so challenging, consider supplemental hand-feedings when the jill’s milk production is reduced because of illness. The stimulus of nursing may promote lactation as the jill improves. Another important alternative is cross fostering. In fact, it is best to breed jills in pairs so one may serve as a foster mother if problems arise. Most jills accept kits of any size or age at any stage of lactation. Merely remove kits from both litters for a short time, mix the two litters together, and then replace all kits with the foster mother.8,20
Neonatal Mortality and Deformities The neonatal mortality rate is greatest during the first 3 to 4 days of life, then drops dramatically after day 5. Common causes of death include cannibalism, stillbirth, and severe congenital defects such as agenesis of limbs, anencephaly, hemivertebrae, scoliosis, gastroschisis, and cranioschisis. Other congenital abnormalities that have been reported include cleft lip, cleft palate, corneal dermoids, kinked tail, and short tail.24,44
Entangled Umbilical Cords Kits in large litters are occasionally born so rapidly the dam is unable to chew the placenta off each individual, creating a mass of kits bound together by their umbilical cords. The entanglement may be exacerbated if kits are born on coarse, sharp-edged shavings. The mass of entangled kits cannot nurse so they quickly become hypoglycemic and the jill cannot curl around them so they also become hypothermic. If tangling of cords and placenta occurs, carefully dissect placentas from each kit’s umbilicus with blunt scissors as far as possible from the kit’s abdomen.24 If the placenta has become dry, soften tissue with warm water. Minimize the risk of entangled cords by closely supervising whelping. If need be, pick up kits as they are born and separate the placenta.8,24,44
Diarrhea Diarrhea in kits may be caused by ferret rotavirus alone, concurrent rotaviral and bacterial infections (i.e., Campylobacter jejuni, E. coli, Proteus species, S. aureus, Enterobacter cloacae), or bacterial infection alone. Ferret rotavirus is carried by adults, and
Table 4-1 Composition of Ferret Milk63 Postpartum (Days)
Mean % Fat
Mean % Protein
Mean % Lactose
5 11 19 25 33 39
7.8-8.5 9.3-10.5 8.9-10.8 8.8-9.5 9.2-10.3 9.0-13.0
7.2-8.8 6.3-7.9 6.0-8.3 5.0-7.9 8.6-9.8 8.4-10.6
2.7-4.2 2.8-4.4 3.8-4.2 3.3-4.2 3.0-4.1 1.5-3.2
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may cause diarrhea in stressed kits or even unstressed kits if they possess no passive immunity.8,65 The jill typically grooms away all evidence of diarrhea, but the kits will appear wet and dehydrated. Disease may be mild and self-limiting in older kits but is potentially life-threatening during the first week of life. Fortunately, the prognosis is good if kits receive aggressive supportive care. Provide fluids and oral, broad-spectrum antibiotics for 5 to 7 days. Monitor the jill closely since anorexia in the kits may promote mastitis and/or agalactia.8,24
Neonatal Conjunctivitis In neonatal conjunctivitis, or ophthalmia neonatorum, purulent discharge collects in the conjunctival sac behind the unopened eyelids of the kit. A variety of pathogens have been cultured. The route of infection is unknown, but has been theorized to develop secondary to minuscule eyelid punctures acquired as the kit is dragged around the nest box. Unilateral or bilateral conjunctivitis is typically seen in kits between a few days to 3 weeks of age. Kits typically stop nursing because of the pain associated with pressing their eyes against the dam. Open the eyelids by cutting along the natural suture line with a small scalpel blade or 25-gauge needle bevel. Flush debris and apply a broad-spectrum ophthalmic ointment. Since littermates are often affected, carefully examine the entire litter regularly. The prognosis for neonatal conjunctivitis is good in 3-week-old kits since the eyelids will stay open. In younger kits the eyelids may reseal and infection may recur.8,20,37
Splay-Legged Kits Splay-legged kits, or “swimmers,” are tetraparetic. Affected kits must lie on their sternum leading to rib compression and death secondary to anoxia by 8 weeks of age. Although the cause is unknown, “splay leg” has been theorized to be either hereditary or husbandry related when a rapidly growing kit housed on smooth flooring places excessive weight on its immature limbs.37
ACKNOWLEDGMENTS I thank Dr. Elizabeth V. Hillyer for authoring the first edition of this chapter on urogenital diseases and Dr. Judith Bell for authoring the second edition of this chapter on periparturient and neonatal diseases. Their work formed the basis for this chapter.
References 1. Amstislavsky S, Ternovskaya Y. Reproduction in mustelids. Anim Repro Sci. 2000;60-61:571-581. 2. Antinoff N. Lymphoma in ferrets: review and preliminary findings. Proc Assoc Exotic Mammal Vet Sci Progr. 2007:99-100. 3. Bartlett LW. Ferret soft tissue surgery. Semin Avian Exot Pet Med. 2002;11:221-230. 4. Basinger RR, Robinette CL, Spaulding KA. Prostate. In: Slatter D, ed. Textbook of small animal surgery. Philadelphia: Elsevier Science; 2002:1542-1556. 5. Batchelder MA, Bell JA, Erdman, et al. Pregnancy toxemia in the European ferret (Mustela putorius furo). Lab Anim Sci. 1999;49:372-379. 6. Batista-Arteaga M, Alamo D, Herráez P, et al. Segmental atresia of the uterus associated with hydrometra in a ferret. Vet Rec. 2007;161:759-760. 7. Beeber NL. Abdominal surgery in ferrets. Vet Clin North Am Exot Anim Pract. 2000;3:647-662.
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SECTION I Ferrets
8. Bell JA. Ferrets: periparturient and neonatal diseases. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits and rodents: clinical medicine and surgery. 2nd ed. Philadelphia: WB Saunders; 2003: 50-57. 9. Bell RC, Moeller RB. Transitional cell carcinoma of the renal pelvis in a ferret. Lab Anim. 1990;40:537-538. 10. Bisceglia M, Galliani CA, Senger C, et al. Renal cystic diseases: a review. Adv Anat Pathol. 2006;13:26-56. 11. Bodri MS. Theriogenology question of the month. J Am Vet Med Assoc. 2000;217:1465-1466. 12. Coleman GD, Chavez MA, Williams BH. Cystic prostatic disease associated with adrenocortical lesions in the ferret (Mustela putoris furo). Vet Pathol. 1998;35:547-549. 13. Dalrymple EF. Pregnancy toxemia in a ferret. Can Vet J. 2004;45:150-152. 14. del Angel-Caraza J, Chávez-Moreno O, García-Navarro S, et al. Mixed urolith (struvite and calcium oxalate) in a ferret (Mustela putorius furo). J Vet Diagn Invest. 2008;20:682-683. 15. Dillberger HE. Polycystic kidneys in a ferret. J Am Vet Med Assoc. 1985;186:74-75. 16. Donnelly TM, Orcutt CJ. Acute ataxia in a young ferret following canine distemper vaccination. Renal failure after epinephrine overdose. Lab Anim. 2001;30:25-27. 17. Dunayer E. Toxicology of ferrets. Vet Clin North Am Exot Anim Pract. 2008;11:301-314. 18. Dutton MA. Treatment of cystine bladder urolith in a ferret. Exot Pet Pract. 1996;1:7. 19. Esteves MI, Marini RP, Ryden EB, et al. Estimation of glomerular filtration rate and evaluation of renal function in ferrets (Mustela putorius furo). Am J Vet Res. 1994;55:166-172. 20. Fisher PG. Ferrets: urogenital and reproductive system disorders. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Quedgeley: BSAVA; 2009:1-12. 21. Fisher PG. Exotic mammal renal disease: diagnosis and treatment. Vet Clin North Am Exot Anim Pract. 2006;9:69-96. 22. Fisher PG. Stump pyometra in a female ferret. Exot Pet Pract. 1998;1:7. 23. Fox JG. Normal clinical and biologic parameters. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:183-210. 24. Fox JG, Bell JA. Growth, reproduction and breeding. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:211-227. 25. Fox JG, Pearson RC, Bell JA. Diseases of the genitourinary system. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:247-272. 26. Gamble C, Morrisey JK. Ferrets. In: Carpenter JW, ed. Exotic animal formulary. 3rd ed. St. Louis: WB Saunders; 2005:455. 27. Gorgas D. Sonography of the kidneys in ferrets. [German] Sonographische Untersuchungen der Nieren beim Frettchen (Mustela putorius f. furo L. 1758). Institut fur Tieranatomie. Munich, Ludwig Maximillians University: Faculty of Veterinary Medicine. Dr Med Vet. 2004:72. 28. Hart JE. Endocrine pathology of estrogens: species differences. Pharmacol Ther. 1990;47:203-218. 29. Hosgood G. The omentum—the forgotten organ: physiology and potential surgical applications in dogs and cats. Compend Contin Educ Pract Vet. 1990;12:45-51. 30. Jacob S, Poddar S. Morphology and histochemistry of the ferret prostate. Acta Anat. 1986;125:268-273:(Basel). 31. Jackson CN, Rogers AB, Maurer KJ, et al. Cystic renal disease in the domestic ferret. Comp Med. 2008;58:161-167. 32. Jekl V, Hauptman K, Jeklová E, et al. Hydrometra in a ferret-case report. Vet Clin North Am Exot Anim Pract. 2006;9:695-700. 33. Johnson-Delaney CA. Medical therapies for ferret adrenal disease. Semin Avian Exot Pet Med. 2004;13:3-7. 34. Kawaguchi H, Miyoshi N, Souda M, et al. Renal adenocarcinoma in a ferret. Vet Pathol. 2006;43:353-356.
35. Kawasaki TA. Normal parameters and laboratory interpretation of disease states in the domestic ferret. Semin Avian Exot Pet Med. 1994;3:40-47. 36. Lennox AM, Lichtenberger M. A new type of urinary catheter for catheterization of the male ferret. Exot DVM. 2008;10:5-6. 37. Lewington JH. Ferret husbandry, medicine and surgery. Edinburgh: Elsevier; 2002. 38. Li X, Fox JG, Erdman SE, et al. Cystic urogenital anomalies in ferrets (Mustela putorius furo). Vet Pathol. 1996;33:150-158. 39. Li X, Fox JG, Padrid PA. Neoplastic disease in ferrets: 574 cases (1968-1997). J Am Vet Med Assoc. 1998;212:1402-1406. 40. Lichtenberger M. Treatment of urinary obstruction in the male ferret. Exot DVM. 2006;8:26-30. 41. Lindeberg H. Reproduction of the female ferret (Mustela putorius furo). Reprod Domest Anim. 2008;2(suppl 43):150-156. 42. Manning DD, Bell JA. Derivation of gnotobiotic ferrets: perinatal diet and hand-rearing requirements. Lab Anim Sci. 1990;40:51-55. 43. Marini RP, Esteves MI, Fox JG. A technique for catheterization of the urinary bladder in the ferret. Lab Anim. 1994;28:155-157. 44. McLain DE, Harper SM, Roe DA, et al. Congenital malformations and variations in reproductive performance in the ferret: effects of maternal age, color, and parity. Lab Anim Sci. 1985;35:251-255. 45. Nakal M, Van Cleef JK, Bahr JM. Stages and duration of spermatogenesis in the domestic ferret (Mustela putorius furo). Tissue Cell. 2004;36:439-446. 46. Nelson WB. Hydronephrosis in a ferret. Vet Med. 1984;79:516-521. 47. Niezgoda M, Briggs DJ, Shaddock J, et al. Pathogenesis of experimentally induced rabies in domestic ferrets. Am J Vet Res. 1997;58:1327-1331. 48. Nguyen HT, Moreland AF, Shields RP. Urolithiasis in ferrets (Mustela putorius). Lab Anim Sci. 1979;29:243-245. 49. Nolte DM, Carberry CA, Gannon KM, et al. Temporary tube cystostomy as a treatment for urinary obstruction secondary to adrenal disease in four ferrets. J Am Anim Hosp Assoc. 2002;38:527-532. 50. Oglesbee BL. Prostatitis and prostatic abscesses. In: Oglesbee BL, ed. The 5-minute veterinary consult: ferret and rabbit. Ames: Blackwell Publishing; 2006:134-137. 51. Orcutt C. Oxyglobin administration for the treatment of anemia in ferrets. Exotic DVM. 2000;2:44-46. 52. Orcutt CJ. Treatment of urogenital disease in ferrets. Exot DVM. 2001;3:31-37. 53. Orcutt CJ. Ferret urogenital diseases. Vet Clin North Am Exot Anim Pract. 2003;6:113-138. 54. Palley LS, Corning BF, Fox JG, et al. Parvovirus-associated syndrome (Aleutian disease) in two ferrets. J Am Vet Med Assoc. 1992;201:100-106. 55. Patterson MM, Rogers AB, Schrenzel MD, et al. Alopecia attributed to neoplastic ovarian tissue in two ferrets. Comp Med. 2003;53:213-217. 56. Powers LV, Winkler K, Garner MM, et al. Omentalization of prostatic abscesses and large cysts in ferrets (Mustela putorius furo). J Exot Pet Med. 2007;16:186-194. 57. Puerto DA, Walker LM, Saunders HM. Bilateral perinephric pseudocysts and polycystic kidneys in a ferret. Vet Radiol Ultrasound. 1998;39:309-312. 58. Purcell K. Essentials of ferrets: a guide for practitioners. 2nd ed. Lakewood: AAHA Press; 1999;62-63. 59. Reese S, Frings B. Abdominal sonography in ferrets (Mustela putorius f. furo L. 1758). Tierärztl Prax. 2004;32:182-189. 60. Riechert M. Selected kidney diseases in the ferret. [German] Untersuchungen ausgewählter Nierenkrankheiten beim Frettchen (Mustela putorius f. furo L. 1758). Institut für Tieranatomie der Tierärztlichen Fakultät. München, Ludwig- Maximilians-Universität. Dr Med Vet. 2005:109.
CHAPTER 4 Disorders of the Urinary and Reproductive Systems 61. Rosenbaum MR, Affolter VK, Usborne AL, Beeber NL. Cutaneous epitheliotropic lymphoma in a ferret. J Am Vet Med Assoc. 1996;209:1441-1444. 62. Schoemaker NJ, van Deijk R, Muijlaert B, et al. Use of a gonadotropin releasing hormone agonist implant as an alternative for surgical castration in male ferrets (Mustela putorius furo). Theriogenology. 2008;70:161-167. 63. Schoknecht PA, Cranford JA, Akers RM. Variability in milk composition of the domestic ferret (Mustela putorius). Comp Biochem Physiol A Comp Physiol. 1985;81:589-591. 64. Sherrill A, Gorham J. Bone marrow hypoplasia associated with estrus in ferrets. Lab Anim Sci. 1985;35:280-286. 65. Torres-Medina A. Isolation of an atypical rotavirus causing diarrhea in neonatal ferrets. Lab Anim Sci. 1987;37:167-171.
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66. Wagner RA, Bailey EM, Schneider JF, et al. Leuprolide acetate treatment of adrenocortical disease in ferrets. J Am Vet Med Assoc. 2001;218:1272-1274. 67. Weisse C, Aronson LR, Drobatz K. Traumatic rupture of the ureter: 10 cases. J Am Anim Hosp Assoc. 2002;38:188-192. 68. Welles EG. Urine protein:creatinine ratio. In: Côté E, ed. Clinical veterinary advisor: dogs and cats. St. Louis: Mosby Elsevier; 2007:1502. 69. Williams BH, Weiss CA. Ferrets: neoplasia. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. Philadelphia: WB Saunders; 2003:91-106.
CHAPTER
5
Cardiovascular and Other Diseases
James K. Morrisey, DVM, Diplomate ABVP (Avian), and Marc S. Kraus, DVM, Diplomate ACVIM (Cardiology, Internal Medicine)
PART I CARDIAC DISEASE General Principles History and Clinical Signs Physical Examination Diagnosis Radiography Electrocardiography Echocardiography Treatment Dilated Cardiomyopathy Hypertrophic Cardiomyopathy Valvular Heart Disease Myocarditis Neoplasia Heartworm Disease PART II OTHER DISEASES Aleutian Disease Clinical Signs Diagnosis Treatment and Prevention Splenomegaly Anemia Ibuprofen Toxicosis
PART I CARDIAC DISEASE Cardiac disease is relatively common in pet ferrets and is suspected based on the presence of radiographic cardiomegaly, a heart murmur on auscultation, or clinical signs consistent with congestive heart failure (CHF). In addition, a thorough cardiac evaluation should be considered for any ferret undergoing anesthesia. For these reasons, practitioners working with ferrets should be familiar with the clinical signs and diagnosis of heart 62
disease as well as the treatment options available. Reference data for common testing modalities, including echocardiography, electrocardiography (ECG), and radiography, have been published for ferrets. This section provides readers with this information as well as our own accumulated clinical experience to facilitate the diagnosis and treatment of cardiovascular diseases in ferrets. To date, acquired heart disease is the only form of heart disease reported in ferrets. Congenital cardiac disease is not commonly recognized, and no case reports have been published.
GENERAL PRINCIPLES HISTORY AND CLINICAL SIGNS Most ferrets presented because of cardiac disease are middle-aged or older (i.e., older than 3 years of age). Typical presenting clinical signs may be confused with those of other diseases, so a thorough history is important. The owner should be questioned about changes in the animal’s appetite, activity level, sleep patterns, exposure to other ferrets, and exposure to the human influenza virus. Ferrets with cardiac disease show a variety of clinical signs similar to those observed in other species, including lethargy, exercise intolerance, weight loss, anorexia, ascites, coughing, and dyspnea.14,40,61 In addition, ferrets with cardiac disease may present with hind limb weakness. It is unclear why the hind limbs are preferentially affected compared with the front, but thromboembolic disease does not appear to be a cause. The diagnosis of cardiac disease may be an incidental finding in some ferrets; either these ferrets have compensated for mild cardiac disease or the owner has not noticed the often insidious signs.
PHYSICAL EXAMINATION A thorough and systematic physical examination should be performed in ferrets with suspected cardiac disease. Examine the oral mucous membranes for color and refill time (less than 2 seconds is normal). Pale or cyanotic mucous membranes with a prolonged capillary refill time may be related to congestive heart failure (CHF) or reduced cardiac output. Jugular pulses may be visible in ferrets with right-sided CHF. Femoral pulses are Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 5 Cardiovascular and Other Diseases palpated for quality, character, and the presence of any deficits. Pulses may be weak, irregular, or normal with cardiac disease. Bounding pulses are uncommon but could potentially be seen with a severe aortic insufficiency or a patent ductus arteriosus (PDA). Abdominal palpation of ferrets with heart disease may reveal ascites, hepatomegaly, and splenomegaly. Other findings on physical examination may include hypothermia, lethargy, generalized weakness, and dehydration. The ferret heart is auscultated between the sixth and eight ribs, much more caudally than the dog or cat. The normal heart rate is between 180 and 250 beats per minute. Ferrets may have a pronounced sinus arrhythmia that causes a dramatic decrease in heart rate or sinus pauses during auscultation. Relevant findings on auscultation are bradycardia, tachycardia, murmurs, gallop rhythms, and muffled heart sounds. Murmurs are usually associated with valvular insufficiency and increased outflow tract velocities. An S3 gallop occurs secondary to dilation of the left ventricle, and an S4 gallop occurs secondary to an accentuated atrial contraction filling a hypertrophied and noncompliant ventricle. Ferrets with cardiac disease may show signs of dyspnea and tachypnea. Auscultation of the lungs may reveal crackles, muffled lung sounds, or increased bronchovesicular sounds.
63
A
DIAGNOSIS A suspected diagnosis of heart disease can be made based on the history and physical examination findings previously discussed. A definitive diagnosis of cardiac disease requires an ECG, thoracic radiographs, and echocardiogram. These testing modalities are discussed in detail later. For all patients with documented or suspected heart disease, complete a minimum database— including a complete blood count (CBC), biochemical profile, and urinalysis—before instituting therapy. Heartworm testing is indicated in ferrets with clinical signs suggestive of infection and for those from heartworm-endemic areas. If thoracocentesis or abdominocentesis is done, cytologic and fluid analysis of the aspirated sample may provide some insight into the cause of the effusion.
RADIOGRAPHY Standard two-view thoracic radiographs can yield important information for the diagnosis of cardiac disease and CHF. Because problems in the anterior mediastinum are common, the entire thorax should be included in the radiographs. Physical restraint is usually adequate to obtain radiographs, although a short dose of inhalant anesthesia or midazolam (0.3-0.5 mg/kg) may be used safely. The typical landmarks used for dogs and cats are inaccurate in ferrets because of the shape of the ferret’s thorax, which is elongated and flattened dorsoventrally.71 The normal ferret heart is more globoid in appearance than a dog or cat heart and is located approximately between the sixth and eighth intercostal spaces. The right ventricle is slightly in contact with the sternum (Fig. 5-1). If heart disease is present, the cardiac silhouette may appear enlarged, resulting in tracheal elevation, rounding of the cardiac silhouette, and increased sternal contact on the lateral view. On the ventrodorsal view, the heart may be elongated and widened, filling up a larger portion of the thorax. The size of the cardiac silhouette can be quantified by the use of a modified vertebral heart score (VHS).71 The technique
B Fig. 5-1 Normal lateral (A) and ventrodorsal (B) thoracic radiographs of a 3-year-old, clinically normal ferret. involves measuring the length and width of the heart on the right lateral (RL) radiograph and expressing this measurement in units of vertebral length (Fig. 5-2). This method corrects for differences in size between ferrets. Because the presence of pericardial fat on the ventrodorsal (VD) view makes the edges of the cardiac silhouette more difficult to discern, the lateral view is more accurate. On the lateral view, the long axis of the heart is the length from the ventral border of the bifurcation of the trachea to the apex of the heart, and the short axis is the maximum length from the cranial to the caudal edges. On the ventrodorsal view, the long axis is the length of the heart along midline from the cranial border of the heart to the apex. The short axis is the maximum width of the heart measured perpendicular to the long axis. These measurements are compared with the length of the thoracic vertebrae (v), beginning with the cranial edge of the fifth thoracic vertebra (T5) to end of eighth thoracic vertebra (T8) and estimated to the nearest 0.25 vertebra. The heart length and width measurements in verterbral lengths from a single projection are then added to obtain a total score. The median VHS on the right lateral projection is 5.33 v, with a range of 5.23 to 5.47 v. The heart length plus width is 5.35 cm, with a range of 5.02 to 5.48 cm. The median VHS on
64
SECTION I Ferrets
Fig. 5-2 Drawings of lateral (right) and ventrodorsal (left) views of the thorax indicating measurements of the cardiac silhouette in both long axis (LA) and short axis (SA). The sum of the LA and SA measurements is expressed in terms of vertebral length, beginning at the cranial edge of the fifth thoracic vertebra (T5) and estimated to the nearest 0.25 vertebra. The vertebral length and width measurements are then added to obtain a vertebral heart score. (Adapted from Stepien RL, Benson KG, Forrest LJ. Radiographic measurement of cardiac size in normal ferrets. Vet Radiol Ultrasound. 1999;40:606-610.) the ventrodorsal view is 6.0 v, with a range of 5.73 to 6.15 v. Basically, the thoracic vertebrae in ferrets are 1 cm long between T5 and T8. Therefore you can measure length plus width, and the sum of these approximates the VHS. Radiographically, CHF may be seen as pleural effusion, pulmonary edema, or pulmonary venous congestion. Pulmonary edema shows most typically as a patchy interstitial and alveolar pattern. Abdominal radiographs may reveal hepatomegaly, splenomegaly, and ascites.
ELECTROCARDIOGRAPHY In ferrets, the electrocardiogram (ECG) is used primarily to determine the presence of abnormal rhythms and conduction disturbances. Reference values for ECGs in ferrets have been published and are summarized in Table 5-1.6,16,17,68 A variety of common arrhythmias have been seen in ferrets (Fig. 5-3). Sinus rhythm and sinus tachycardia are the most common rhythms seen on presentation with cardiac disease. Atrial and ventricular premature contractions may be recorded. Atrial fibrillation can occur in the presence of significant atrial enlargement. Sinus bradycardia can be associated with hypoglycemia, commonly seen in ferrets with insulinomas. Second-degree atrioventricular (AV) block can occur as a normal finding in healthy ferrets. Highgrade second-degree AV block and complete (third-degree) AV
block are rare. Complete AV block can be successfully treated by implanting an epicardial transdiaphragmatic pacemaker (Fig. 5-4). Other reported ECG changes include tall R waves, prolonged QRS complexes, and ST-segment depression.69 The ECG is ideally done without sedation with the animal in right lateral recumbency (Fig. 5-5). The ECG clips can be flattened to produce a smooth surface that does not result in pinching. Because most ferrets object to the use of alcohol, ECG coupling gel (or ultrasound gel) is recommended. The ferret can be distracted by offering semisoft food such as a/d (Hill’s Pet Nutrition, Inc, Topeka, KS) or a nonsugary treat such as FerreTone (8 in 1, Islandia, NY) from a syringe while the ECG is being recorded.
ECHOCARDIOGRAPHY The echocardiogram remains the test of choice for diagnosing structural and functional cardiac abnormalities.46 In addition, the echocardiogram can be useful in identifying mediastinal masses and as an aid in detecting heartworm disease.65 It can often be obtained without the use of sedation. If sedation is required, either a short-acting injectable anesthetic or an inhalant anesthetic may be used. The echocardiogram is done with the animal in both right and left lateral recumbency. Imaging planes similar to those obtained in other species are recorded.77 Two-dimensional echocardiography provides an assessment of
CHAPTER 5 Cardiovascular and Other Diseases
65
Table 5-1 Electrocardiographic Values for 52 Clinically Normal Ferretsa Parameter Age (mo) Male/female ratio Body weight (kg) Rhythm Normal sinus Sinus arrhythmia Heart rate (beats/min) Mean electrical axis, frontal plane (degrees) Lead II measurements P amplitude (mV) P duration (sec) PR interval (sec) QRS duration (sec) R amplitude (mV) QT interval (sec) aAll
Mean, or Mean ± SD (RANGE) (N = 25)6
Value (N = 27)17
10–20 All male 1.4 ± 0.2
5.2 1.25 Not available
Not available Not available 196 ± 26 (140–240) +86.1 ± 2.5 (79.6–90.0)
67% 33% 233 ± 22 +77.2 ± 12.0
Not available Not available 0.056 ± 0.0086 (0.04–0.08) 0.044 ± 0.008 (0.035–0.06) 2.21 ± 0.42 (1.4–3.0) 0.11 ± 0.02 (0.08–0.14)
0.122 ± 0.007 0.024 ± 0.004 0.047 ± 0.003 0.043 ± 0.003 1.46 ± 0.84 0.12 ± 0.04
ferrets were sedated with ketamine-xylazine.
A
B Fig. 5-4 Radiograph of a ferret with an implanted cardiac pacemaker. The ferret exhibited signs of collapse and weakness with profound bradycardia due to complete AV block.
C
D Fig. 5-3 Electrocardiograms recorded from four ferrets, showing normal rhythm (A), second-degree atrioventricular (AV) block in an asymptomatic ferret (B), paroxysmal ventricular tachycardia (C), and complete AV block (D).
cardiac size and function. Standard M-mode measurements are obtained, including chamber dimensions, wall thickness, and indices of systolic function. Spectral Doppler imaging is used to quantitate the velocity of normal and abnormal blood flow. Color-flow Doppler echocardiography provides a visual inspection of blood flow direction and detection of turbulent flow, including valvular regurgitation. Echocardiographic reference
Fig. 5-5 Ferret restrained in right lateral recumbency for recording an ECG.
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SECTION I Ferrets Table 5-2 M-Mode and Doppler Echocardiographic Values for Normal Ferretsa Value (Mean ± SD)72
Parameter Interventricular septum end-diastole (IVSd) (cm) Interventricular septum end-systole (IVSs) (cm) Left ventricular free wall end-diastole (LVWd) (cm) Left ventricular free wall end-systole (LVWs) (cm) Left ventricular internal dimension end-diastole (LVIDd) (cm) Left ventricular internal dimension end-systole (LVIDs) (cm) Fractional shortening (%) Ejection fraction (%) Left atrial diameter (LA) (cm) Aortic diameter (AO) (cm) LA/AO (ratio) Pulmonary artery diameter (cm) Right ventricular free wall end-diastole (cm) Right ventricular internal dimension end-diastole (cm) Aorta maximum velocity (m/s) Pulmonary artery maximum velocity (m/s) Mitral valve E velocity (m/s) Mitral valve A velocity (m/s) Mitral valve velocity E/A ratio aAll
0.36 ± 0.07 0.48 ± 0.11 0.42 ± 0.11 0.58 ± 0.99 0.88 ± 0.15 0.59 ± 0.15 33 ± 14 69 ± 19 0.71 ± 0.18 0.53 ± 0.10 1.33 ± 0.27 0.48 ± 0.90 0.12 ± 0.03 0.38 ± 0.10 0.89 ± 0.20 1.10 ± 0.14 0.70 ± 0.10 0.52 ± 0.11 1.38 ± 0.32
ferrets were sedated with ketamine and midazolam (0.2 mg/kg).
Table 5-3 Acute Dosages and Treatment Goals of Drugs Commonly Used to Treat Congestive Heart Failure in Ferrets Drug
Indication
Furosemidea
Overt CHF*
Dobutamine
Inotropic support, acute CHF Nitroglycerin 2% Overt CHF, preload ointment reduction aWhen
Dose (mg/kg)
Frequency
2.0-4.0 IV preferable q8-12h or IV bolus, then CRI at 0.7-1 mg/kg per hour 5-10 μg/kg per minute IV Constant rate infusion Topical 1/16-1/8 in area inside the ear pinna or inguinal region
q12-24h
to initiate ACE inhibitor therapy for impending CHF is controversial.
values in the sedated ferret have been published and are summarized in Table 5-2.72 The echocardiogram is used in conjunction with good-quality thoracic radiographs to properly identify the severity of underlying cardiac disease and the presence of CHF.
TREATMENT Basic therapeutic principles of CHF and cardiac disease are applied to ferrets. If a published ferret dose is not available, feline doses are often used as a starting point. Therapy for acute CHF (Table 5-3) focuses on improving oxygenation and reducing preload and afterload. Oxygen is provided by placing the animal in a closed cage or incubator with supplemental oxygen. Diuretics reduce preload by reducing blood volume.50 Furosemide is the most commonly used diuretic; it is most effective when given either intravenously or intramuscularly in the acute setting. Ferrets tolerate doses of 1 to 4 mg/kg every 8 to
12 hours. If respiratory rate does not decrease with pulse therapy with intravenous furosemide, consider treating with a constant rate infusion (CRI) of furosemide at a dose of 0.7 to 1 mg/kg per hour. Nitroglycerin 2% ointment is a venous dilator that can be applied to the skin in the axilla or inguinal area on a hairless body surface. Ferrets are sensitive to the hypotensive effects of nitroglycerin. Angiotensin-converting enzyme (ACE) inhibitors are given to reduce afterload by causing arterial vasodilation and to reduce preload by decreasing salt and water retention.37 The dose is usually started at 0.5 mg/kg PO given every 48 hours and then titrated up to every 24 hours if tolerated. In the first 24 to 48 hours of therapy, treatment with ACE inhibitors may result in hypotension or lethargy. The use of ACE inhibitors and concurrent diuretic therapy in ferrets with CHF can result in severe azotemia. Renal perfusion is reduced in CHF secondary to use of diuretics, decreased cardiac output, dehydration, and hypotension.28 If significant pleural effusion is present, resulting in an increased inspiratory effort, perform
CHAPTER 5 Cardiovascular and Other Diseases
67
Table 5-4 Maintenance Dosages of Drugs Commonly Used to Treat Congestive Heart Failure (CHF) in Ferrets Drug
Indication
Furosemide ACE inhibitor Enalaprilb Benazeprilb Spironolactonec
Overt or impending CHF Overt or impending CHFa
Digoxin (pediatric elixir) Diltiazem Amlodipinee Pimobendan
Aldosterone escape Electrolyte control Atrial fibrillation Atrial fibrillation or SVTd (rare) Afterload reduction Systolic dysfunction
Maintenance Dose (mg/kg)
Frequency
1.0-2.0
q12hr
0.25-0.5 0.25-0.5 1-2
q12-24hr q24hr q12hr
0.005-0.01 1.5-7.5 2-6 0.2-0.4 0.5
q12hr q12hr q12hr q12hr q12hr
aWhen
to initiate ACE inhibitor therapy for impending CHF is controversial. at 0.25 mg/kg and increase to 0.5 mg/kg after ensuring that renal function is normal. cPreserves magnesium and potassium while antagonizing aldosterone action. dSupraventricular tachycardia. eInitiate at 0.1 mg/kg and titrate upward weekly while monitoring blood pressure. bInitiate
pleurocentesis and submit the fluid for fluid analysis and cytologic examination. In acute disease, monitor ferrets closely for respiratory rate and effort, mucous membrane color, heart rate and rhythm, body weight, and hydration as well as electrolyte, blood urea nitrogen (BUN), and creatinine values. Obtain serial radiographs to assess the response to therapy. Adjust medication dosages based on the listed monitoring parameters. Chronic therapy typically includes furosemide, ACE inhibitors, digoxin, and a new positive inotrope, pimobendan (Vetmedin, Boehringer Ingelheim Vetmedica, St Joseph, MO) (Table 5-4). Furosemide is given orally at a dosage of 1 to 4 mg/kg q8-12h; ACE inhibitors are given at 0.5 mg/kg q24-48h as tolerated. Digoxin therapy is used as a positive inotrope and to depress AV nodal conduction with supraventricular arrhythmias.37 Digoxin elixir (0.05 mg/mL) is recommended because of the small doses required in ferrets. The recommended starting dosage is 0.01 mg/kg PO q12-24h.61 Calculated dosages for digoxin are based on lean body weight, usually 70% of body weight. Begin therapy once daily and then titrate up to twice daily depending on the clinical response and serum digoxin levels. Side effects of digoxin include inappetence, lethargy, vomiting, diarrhea, and arrhythmias.69 To date, no pharmacokinetic studies of digoxin in ferrets have been published. To measure digoxin levels, serum samples are taken approximately 6 to 8 hours after drug administration.69 Reference values for dogs and cats are extrapolated to interpret values in ferrets (0.8-2.0 ng/mL). Contraindications for the use of digoxin include significant azotemia, hypokalemia, bradyarrhythmias including AV block, and severe ventricular arrhythmias.37 In our clinic, digoxin is not used as the first-line positive inotrope because of the plethora of side effects and narrow therapeutic range. Pimobendan is used at a dosage of 0.625 to 1.25 mg/kg q12h. The positive inotropic effect is caused by the increased sensitivity of troponin C to calcium and also by inhibition of phosphodiesterase III. The latter mechanism results in vasodilation. In addition, this drug has neurohormonal effects, which include decreased levels of inflammatory cytokines (tumor necrosis factor [TNF] alpha, interleukin-1) and plasma epinephrine. Clinical studies in dogs
reveal improved quality of life and survival with pimobendan therapy; whether or not this beneficial effect exists in ferrets is unknown. However, some experimental studies with pimobendan in ferrets have been performed.23,38,39,41 Long-term management includes periodic monitoring for recurrence of CHF and changes in body weight, renal values, and heart rate and rhythm. If CHF recurs with standard therapy protocols, additional diuretics may be used, including thiazide diuretics or potassium-sparing diuretics such as spironolactone. Published feline dosages are used, and the dosage is titrated based on clinical response. Close monitoring of electrolytes after beginning more aggressive and mixed diuretic therapy is recommended. In ferrets with CHF receiving steroid therapy for lymphoma, spironolactone should be considered early in the disease process because of its effects on the aldosterone receptor in the distal renal tubule.50 The use of antiarrhythmic drugs is not well documented in ferrets. Atenolol titrated to effect can be given for many supraventricular and ventricular arrhythmias. Diltiazem can be used for many supraventricular arrhythmias to either reduce spontaneous occurrence or slow AV nodal conduction.36 Intravenous lidocaine can be used in ferrets with ventricular tachycardia; however, the dose should be reduced from that used in cats and then titrated to effect. With this class of antiarrhythmic drugs, close observation for the development of neurologic and gastrointestinal signs is essential.36
DILATED CARDIOMYOPATHY Dilated cardiomyopathy has been described in ferrets and has clinical characteristics similar to those observed in dogs and cats.14,20,40 The disease results in a dilated left ventricle, right ventricle, or both, with global systolic dysfunction. The cause of the disease in ferrets is not known; in one ferret, it was reported in association with a cryptococcal infection.20 Lethargy, dyspnea, anorexia, and weight loss are common complaints in the history. Physical examination findings can include hypothermia, heart murmur, tachycardia, pallor, weakness, or ascites.
SECTION I Ferrets
68
V
1
LV
LA
After the ferret’s condition is initially stabilized, therapy for chronic disease begins with diuretics, ACE inhibitors, and pimobendan and/or digoxin (for severe systolic dysfunction, the combination may be beneficial). The dose of diuretics should be reduced to the lowest dose that prevents reaccumulation of pleural effusion and pulmonary congestion. A low-salt diet and exercise restriction are theoretically beneficial in the management of CHF but can be difficult practices to institute. To date, no cases of taurine-responsive dilated cardiomyopathy have been documented in ferrets. Long-term therapy includes periodic monitoring of thoracic radiographs, ECG findings, and plasma BUN, creatinine, and electrolyte concentrations. If digoxin is administered, monitor serum digoxin levels 6 to 8 hours after dosing. Large amounts of data are not available to estimate the prognosis in ferrets with dilated cardiomyopathy. The clinical response to therapy is often good, and ferrets tend to respond better to treatment than do dogs or cats with similar echocardiographic findings.
HYPERTROPHIC CARDIOMYOPATHY Fig. 5-6 Two-dimensional echocardiogram of a ferret with dilated cardiomyopathy. The left ventricle (LV) is dilated, with rounding of the left ventricular apex, and the left atrium (LA) is enlarged.
Pleural effusion is seen as an increased inspiratory effort with muffled heart sounds. Pulmonary edema is heard as moist rales and increased respiratory sounds. In many cases, both pulmonary edema and pleural effusion are present simultaneously, resulting in a combination of the clinical signs mentioned. Echocardiography is used to make a definitive diagnosis of dilated cardiomyopathy. Typical echocardiographic changes are similar to those observed in other species.7,45,79 The left ventricle appears dilated, with increased end-systolic dimensions, and the left atrium is typically dilated (Fig. 5-6). Fractional shortening, a commonly used index of systolic function, is reduced. If the right side of the heart is involved, the right ventricle is dilated, right ventricular systolic motion is reduced, and the right atrium is enlarged. In advanced disease, mitral and tricuspid regurgitation secondary to dilation of the valve annulus is seen. Left ventricular outflow tract velocities can be normal or reduced. Radiographically, the cardiac silhouette appears enlarged. Congestive heart failure is seen as pleural effusion or pulmonary edema. If the abdomen is included in the radiograph, hepatomegaly, ascites, or splenomegaly may be present. A variety of ECG abnormalities may be present, including ventricular premature contractions, atrial premature contractions, atrial tachycardia, ventricular tachycardia, and atrial fibrillation.45,79 Therapy for acute clinical CHF was described earlier and includes oxygen, diuretics, nitroglycerin, and pleurocentesis if necessary. Place the ferret in an oxygen-rich environment and give furosemide at 1 to 4 mg/kg IV or IM. Take a thoracic radiograph after the animal is assessed as being stable and the degree of pleural effusion is determined. The venodilator nitroglycerin reduces pulmonary edema in the acute management of CHF.37 The ointment can be placed on a hairless area of skin every 12 to 24 hours. Monitor the response to initial management of heart failure by closely observing the respiratory rate and effort and ausculting lung sounds. An initial database should be obtained, including values for electrolytes, BUN, and creatinine.
Hypertrophic cardiomyopathy (HCM) occurs in ferrets, but the clinical characteristics have not been well described and no reports of HCM have been published. Physiologically, the presence of left ventricular hypertrophy results in impaired filling of the left ventricle (diastolic function). Left ventricular diastolic pressure progressively increases, and the subsequent increase in left atrial pressure results in the development of left-sided CHF.2 Left ventricular hypertrophy has not been documented secondary to hypertension or hyperthyroidism in ferrets. The course of disease is likely to be similar to that in cats, remaining silent until the onset of CHF, thromboembolic events, or sudden death.2,62 Hypertrophic cardiomyopathy should be a differential diagnosis in ferrets with suspected cardiac disease based on auscultation, radiographic findings, or clinical signs of CHF. Echocardiography is used to definitively diagnosis this disease. The entire left ventricle or isolated segments of the left ventricle can be hypertrophied. The left ventricular diastolic and systolic dimensions are reduced, and the fractional shortening is normal or increased. Left atrial enlargement may be present. Systolic anterior mitral valve motion may be seen and is often associated with interventricular septal hypertrophy. Doppler echocardiographic abnormalities can include turbulence in the left ventricular outflow tract secondary to dynamic obstruction and mitral regurgitation. Treatment is aimed at improving the diastolic efficiency of the left ventricle and relieving signs of CHF.19 The most commonly used therapeutic agents for the treatment of HCM include beta-adrenergic blocking drugs (e.g., atenolol at 3.1256.25 mg/kg PO q12-24h) or calcium channel blockers (e.g., diltiazem at 3.75-7.5 mg/kg PO q12h).70 These drugs reduce heart rate and contractility, resulting in better filling of the left ventricle.19 Therapy is titrated to achieve an effective reduction in heart rate and clinical improvement. With both medications, side effects include lethargy, inappetence, bradycardia, and hypotension. If clinical signs of CHF are present, treatment with diuretics is recommended (see earlier discussion). Therapeutic monitoring includes periodic echocardiograms to determine the extent of progressive cardiac hypertrophy and atrial enlargement. In those ferrets with CHF, results of thoracic radiographs, ECGs, and plasma biochemical profiles
CHAPTER 5 Cardiovascular and Other Diseases should be monitored. Thromboembolic disease secondary to cardiac disease in ferrets has not been reported, but one case of atrial thrombosis has been seen by one of the authors (JKM, unpublished, 1997).
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V
1
VALVULAR HEART DISEASE Valvular heart disease most commonly occurs in middle-aged to older ferrets and is being recognized with increasing frequency. Clinical signs vary depending on the severity of the underlying disease. Mitral regurgitation is auscultated as a systolic murmur over the left apical region, and tricuspid regurgitation is auscultated in the right parasternal location. A diastolic murmur of aortic insufficiency is rarely heard on physical examination. Femoral pulses can be hyperdynamic with significant aortic regurgitation, but they most often remain normal or reduced with CHF. Labored breathing and moist rales (crackles) are present in ferrets with CHF. Thoracic radiographs provide a general impression of cardiac size and are required to diagnose CHF definitively. Pulmonary edema is seen as a mixed patchy alveolar and interstitial pattern more prominent in the caudodorsal lung regions. The ECG signs vary but may indicate atrial arrhythmias associated with atrial enlargement. Echocardiography shows thickening of the affected valves and atrial enlargement (Fig. 5-7). The left ventricular diastolic dimension is increased, with a normal systolic dimension and preserved fractional shortening. Left ventricular wall thickness is normal. Regurgitation can be identified and quantified with the use of Doppler echocardiography. Aortic regurgitation is a common incidental finding on echocardiograms in ferrets and is rarely clinically significant. Therapy is recommended if CHF is present or if cardiac enlargement is significant. Treatment with ACE inhibitors blunts the neurohormonal activation that occurs with advanced cardiac disease and CHF.37 If radiographic evidence of CHF is present, administer furosemide, and titrate the dose to effect while monitoring renal values at periodic intervals. Pimobendan (with digoxin as discussed above) can be given after the ferret is stabilized on the therapy described. Indications for its use include stabilization of heart rate, positive inotropic effects, and control of supraventricular arrhythmias. The management of CHF is described above. Several factors influence the prognosis of ferrets with chronic valvular disease, including renal function, coexisting disease, and cardiac rhythm. Insufficient data are available to predict the prognosis in ferrets with CHF secondary to chronic degenerative valve disease.
MYOCARDITIS Myocarditis typically manifests as infiltration of the myocardium with inflammatory cells that results in reduced myocardial function, arrhythmias, and fibrous tissue replacement of normal myocardium.21 Causes of myocardial inflammation are systemic vasculitis and parasitic, autoimmune, bacterial, and viral disorders. A Toxoplasma-like organism has been described as causing multifocal myocardial necrosis.78 Aleutian disease virus can cause fibrinoid necrosis and mononuclear cell infiltration in arterioles of the heart.12 Because of the presence of cytokines and other inflammatory factors, systemic inflammatory diseases, including sepsis, can result in reduced myocardial function.
AV 2
LA
A V
1
2
MV
B Fig. 5-7 Two-dimensional short-axis echocardiograms from a ferret with mitral and aortic regurgitation. A, The aortic valve (AV) is thickened and the left atrium (LA) is severely dilated. B, The mitral valve (MV) is thickened.
The gold standard for the diagnosis of myocarditis requires histologic evaluation of the myocardium, making antemortem verification difficult. However, because cardiac biopsies are clinically difficult to obtain, the use of the cardiac biomarker troponin is the best alternative. Cardiac troponin I (cTnI) is a highly sensitive and specific biomarker of myocardial injury in human medicine that is used to diagnose and provide prognostic information in patients with myocardial infarction, sepsis, and traumatic myocardial injury.29,64,83 Blood cTnI levels have been reported in clinically normal cats and dogs as well as in diseased states (gastric dilation and volvulus, CHF, chemotherapy,
70
SECTION I Ferrets
heat stroke, cardiomyopathy, pericardial disease).3,25,42,54,66,67 Recently, a point-of-care analyzer (i-STAT 1, Abaxis Union City, CA) has become available and validated for measurement of cTnI in humans.83 The company has published reference values for cats, dogs, and horses. However, the accuracy of this instrument in detecting ferret troponin is unknown. The amino acid sequence of cTnI is highly conserved and greater than 90% identical to those from human, canine, and feline cTnI proteins, and the epitopes detected by commercial analyzers are highly conserved among the species.83 These findings substantiate the measurement of cTnI in ferrets and the use of these commercial analyzers. To date, we are unaware of published data on cTnI in ferrets, but based on other species, the “normal” value should be less than 0.1 ng/mL. Myocarditis should be suspected if arrhythmias and acute myocardial dysfunction are found in the presence of a multisystemic illness in conjunction with an increased cTnI. The primary therapeutic goal is directed at the underlying cause of the systemic disorder. Cardiovascular support, including diuretics or antiarrhythmic agents and corticosteroids, may be necessary.
NEOPLASIA
Necropsy reveals heartworms present in the right atrium, right ventricle, cranial vena cava, and pulmonary arteries (see Chapter 6). Right ventricular and right atrial enlargement develop secondary to pulmonary hypertension and mechanical obstruction. Thickening of the tricuspid valve may result from chronic damage caused by the presence of worms. Pleural effusion, ascites, and congestion of abdominal organs may be present. Microfilaria may be seen in alveolar capillaries and large vessels. On histopathologic examination, calcified worms, thrombosis, pulmonary endarteritis, and congestion are typically seen. Chronic passive congestion of the liver and renal hemosiderosis are reported. Aberrant larval migration has not been reported in ferrets. The presence of black feces in the colon has been reported but is not a consistent finding.44,56,61 Clinical signs in affected ferrets include coughing, lethargy, weakness, and dyspnea. Signs of right-sided heart failure, including pleural effusion and ascites, may be present. Hypothermia is a common finding on physical examination. Sudden cardiac death occurs, most likely from pulmonary artery obstruction.44,56 Thoracic radiographs should be obtained and may show pleural effusion and cardiomegaly (Fig. 5-8). Enlargement of the right atrium, caudal vena cava, and right ventricle
Neoplasia involving the myocardium or pericardium has not been reported in ferrets. Lymphoma, a common tumor found in ferrets, can involve the cranial mediastinum. Ferrets with a cranial mediastinal mass are often presented because of dyspnea associated with the presence of pleural effusion.15 Ultrasonography is useful to rule out the presence of cardiac disease and to visualize any mass that may be present. Fine-needle aspiration of the tumor, or in some cases cytologic examination of the fluid, can be used to make a diagnosis.
HEARTWORM DISEASE Natural and experimental infections with the canine heartworm Dirofilaria immitis have been reported in ferrets.4,8,44,47,56 Ferrets living in or originating from an endemic area are most susceptible to infection. The susceptibility and life cycle of this parasite have been studied in ferrets and are similar to those of heartworm in dogs; however, because of the small size of ferrets, the clinical presentation more closely resembles that of infected cats.76 The life cycle begins when a ferret is bitten by a mosquito containing infective D. immitis larvae, which are deposited in the subcutaneous tissues. Larvae migrate to the vascular system and can be found within the small pulmonary arteries. Microfilaria have been reported in ferrets in both natural and experimental infections. Microfilaria are present in 50% to 60% of infected animals.76 Ferrets can be severely affected by the presence of only a single worm. With natural infections, worm burdens ranging from 1 to 21 worms have been reported.44,47,56 The worms can be found within the pulmonary arteries, right side of the heart, and vena cava. Because of the ferret’s small size, infection with a small number of worms can cause mechanical obstruction to blood flow, resulting in clinical signs of right-sided heart failure. In dogs, the host response to living worms within the pulmonary vasculature can include pneumonitis, granuloma formation, pulmonary endarteritis, thromboembolism, and pulmonary hypertension. However, these classic histopathologic changes have not been identified in ferrets, possibly because of the low worm numbers and their primary location within the right side of the heart.
A
B Fig. 5-8 Lateral (A) and ventrodorsal (B) radiographs of a ferret with pleural effusion secondary to heartworm disease.
CHAPTER 5 Cardiovascular and Other Diseases are commonly seen. Unlike in dogs, radiographic peripheral pulmonary artery changes are not severe in ferrets, probably because the worms reside primarily in the right side of the heart and in the main pulmonary artery.44,56,75 Angiographic findings in ferrets experimentally inoculated with D. immitis have been published.75 Common findings are right-sided heart enlargement and filling defects in the right side of the heart, pulmonary artery, and vena cava. Although angiography can be a useful test in the diagnosis of heartworm disease, its clinical use is limited by technical inexperience and the requirement for specialized equipment. Echocardiography is a superior test because it is noninvasive and widely available. Echocardiographic examination can identify the presence of intracardiac parasites. The worms may be visualized in the pulmonary artery, right ventricle, and right atrium. Right ventricular and right atrial dilation may be seen.65 The presence of pulmonary hypertension should be also suspected and can be diagnosed with the use of Doppler echocardiography.30 Diagnosis of heartworm disease is based on clinical signs, radiographic and echocardiographic findings, and results of heartworm blood testing. Microfilaria are seen in only about 50% of infected ferrets. The antigen test used most commonly to diagnose heartworm infection is an enzyme-linked immunosorbent assay (ELISA).70 Antigen is shed by adult female heartworms into the circulation. Because antigen testing detects only female worms, there is less likelihood of positively identifying ferrets with low worm burdens. Studies are needed to further evaluate the sensitivity and specificity of heartworm testing in ferrets. Successful therapy for heartworm disease depends on early, accurate detection. The current recommended treatment protocol for ferrets is ivermectin (50 μg/kg SC q30d) given until clinical signs resolve and microfilaremia is absent. Previous treatment protocols using adulticide therapy with melarsomine (Immiticide, Rhone Merieux, Athens, GA) have fallen out of favor because of adverse reactions (N. Antinoff, personal communication, 2010). Management of pleural effusion with diuretics or pleurocentesis may be necessary. Prednisone (0.5 mg/kg PO q12-24h) is recommended throughout this period and until clinical signs resolve completely. Whether pretreatment with doxycycline for Wolbachia (a bacteria present within Dirofilia immitis that contributes to renal and pulmonary pathology) will enhance the efficacy of heartworm therapy in ferrets is unknown (www.heartwormsociety.org). A follow-up ELISA for heartworm antigen should be performed approximately 3 months after starting therapy and testing repeated at monthly intervals until the results are negative. If antigen test results remain positive, further diagnostic tests (radiographs, echocardiography) may be necessary to determine whether heartworm infection persists. Most ferrets become seronegative within 4 months after successful adulticide therapy. Preventive therapy is recommended for those ferrets previously infected with heartworm disease and for all ferrets in heartworm-endemic areas. Ivermectin (0.05 mg/kg) can be given orally or subcutaneously once a month for 1 month before and after the heartworm season. One quarter of the smallestsize canine or feline ivermectin oral tablet (yield, approximately 14-17 μg) (Heartgard-30 and Heartgard-Feline, Merck & Co. Inc., Whitehouse Station, NJ) is adequate for ferrets. Because the drug deteriorates once the pill is broken, the remainder of the pill must be discarded. Selemectin topically and milbemycin oxime orally can also be used effectively in ferrets (see Appendix for recommended dosages).
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PART II OTHER DISEASES
ALEUTIAN DISEASE Aleutian disease is caused by a parvovirus and was first reported as a disease of mink in the 1940s.18 Mink homozygous for the Aleutian (blue) gene are most severely affected, thus giving the disease its name. The disease is an immune complex-mediated disorder that causes hypergammaglobulinemia. Deposits of immune complexes in various organs are responsible for the clinical signs, which include glomerulonephritis, bile duct proliferation, arteritis, and progressive wasting. In persistently infected mink, other clinical signs are infertility, abortion, and neonatal interstitial pneumonitis.26 Affected animals are immunosuppressed and are more susceptible to other infections, such as influenza, viral enteritis, and distemper. The disease was first seen in ferrets in the late 1960s.32,34 It has been seen in ferrets in the United States, United Kingdom, Sweden, New Zealand, Japan, and Canada.43 There are several strains of the Aleutian disease virus (ADV) in mink, the most common of which is ADV-Utah. These strains vary in the immune response elicited and therefore in virulence. Although the mink virus can infect ferrets, at least three separate viral strains that are distinct from the mink ADV have been documented in ferrets.48,58 The most common strain is called ADVF. Because the hypervariable capsid region of the ferret ADV strains is similar to that of the mink virus, these ferret strains are thought to be mutant strains of the mink ADV.63 Mink can be infected with ferret ADV, but the virulence is lower.12,49,58 The disease is transmitted by aerosol or direct contact with infected saliva, urine, blood, or feces. Fomites, including humans, may be a source of infection, as parvoviruses are typically stable in the environment. Vertical transmission has not been reported in ferrets but is known to occur in mink. Asymptomatic animals have been shown to have persistent viral shedding for more than a year without clinical signs.57
CLINICAL SIGNS In ferrets experimentally infected with mink or ferret strains of ADV, infection persisted for up to 180 days, but the clinical signs that are seen in naturally infected ferrets did not develop.33 The clinical signs in naturally infected ferrets relate to the immune complex deposition and can vary significantly. Some ADVinfected ferrets die in good body condition without any clinical signs.12 Most ferrets show signs of a chronic wasting disease, with varying degrees of progressive weight loss, weakness, ataxia, and posterior paresis. Central nervous system (CNS) signs can include tremors, convulsions, and paralysis.53,73,82,84 Hepatomegaly, splenomegaly, pallor, and melena are also consistent findings. In a recent report, respiratory signs including acute dyspnea were described.80 Ferrets infected at older ages usually show fewer clinical signs; however, ferrets can be infected for years before clinical signs become apparent. Most reported cases were in ferrets between 2 and 4 years of age. Clinical pathologic findings in infected ferrets can also vary. The most consistent findings are hypoalbuminemia and hypergammaglobulinemia. Protein electrophoresis of plasma samples from these animals usually demonstrates gamma globulins amounting to 20% to 60% of the total protein concentration
SECTION I Ferrets
72
TP 10.6 g/dL -globulin 5.75 g%
Alb
2
1
1
2
TP 6.5 g/dL -globulin 1.89 g%
Alb
1
2
1
2
Fig. 5-9 Serum protein electrophoretograms from two ferrets with syndromes associated with Aleutian disease. Top, Ferret 1: Notice the pronounced hypergammaglobulinemia. The gamma globulin fraction (γ) equals 54% of the total protein concentration. Bottom, Ferret 2: The gamma globulin fraction equals 29% of the total protein concentration. Hypergammaglobulinemia is the hallmark of Aleutian disease virus infection. Alb, Albumin; TP, total protein. (From Palley LS, Corning BF, Fox JG, et al. Parvovirusassociated syndrome [Aleutian disease] in two ferrets. J Am Vet Med Assoc. 1992;201:100-106.)
(Fig. 5-9). Rare cases with normal protein fractions have been recorded.80 Because the immune complexes damage internal organs, other biochemical abnormalities, such as azotemia and high liver enzyme concentrations, may be present. Anemia of chronic inflammation has also been reported.55
DIAGNOSIS A presumptive diagnosis is based on the history and clinical signs and a high gamma globulin concentration. Antemortem diagnosis can be made on the basis of clinical signs and a positive serum titer coupled with hypergammaglobulinemia, positive polymerase chain reaction (PCR) results from a fecal swab sample, histologic evidence of associated lymphoplasmacytic inflammation in tissue biopsy samples, or positive PCR results of intestinal or tissue biopsy samples. Counterimmunoelectrophoresis (referred to as CIEP or CEP) historically has been the standard for antibody testing in mink and in ferrets and is still used a screening tool on most mink farms. At present, ferret ADV CIEP testing is available through Taconic (Animal Health Diagnostics, Taconic, Rockville,
MD; www.taconic.com) and Blue Cross Animal Hospital (Burley, Idaho; FerretADV.com). Other available diagnostic tests for ADV testing include ELISA (Avecon Diagnostics Inc., Bath, PA; www.avecon.com), and PCR testing of fecal swab or intestine or splenic tissue samples (Diagnostic Center for Population and Animal Health, Michigan State University, Lansing, MI, www.an imalhealth.msu.edu; Veterinary Molecular Diagnositics, Milford, OH, www.vmdlabs.com; Infectious Disease Laboratory, University of Georgia, Athens, GA; www.vet.uga.edu/SAMS/IDL/), and is the standard screening test in mink for all ADV strains. It is used routinely in ferrets and has been shown to be an effective method of serologically identifying ferrets with ADV antibodies.53,73,82,84 Results are reported as positive, negative, or “no test,” which means that an interference with test results has occurred in the sample. Results from the retesting of “no test” samples usually are clearly positive or negative. The ELISA test is a sensitive and specific test that detects antibodies to the nonvirion NS1 protein of ADV-F only. This is a protein produced by replicating virus, thus confirming active infection; however, a false-negative result is possible if the virus is not actively replicating. This test uses blood (serum/plasma) or saliva samples. A Quick-Check pointof-care test (Avecon) is also available for in-house testing of saliva or blood samples or client testing of saliva samples. A second ELISA developed by the University of Georgia that detects two major capsid proteins of the ADV that are present during replication and latency. False negatives with this test can occur if the host only produces antibodies to the nonstructural viral proteins. For this reason, both ELISAs may be used concurrently to detect structural and nonstructural ADV antigens. The PCR method can be added to detect viremia in an active ADV infection and may be especially useful in detecting asymptomatic animals. The presence of ADV antibody in a ferret is not necessarily diagnostic of disease. In serologic surveys of large pet ferret populations, 8.5% and 10% of ferrets surveyed were antibodypositive without clinical signs of disease.82 Also, immunocompetent adult ferrets experimentally infected with ferret ADV can develop persistent infection (with viral shedding) without clinical disease. As in mink, ferrets probably develop persistent, nonprogressive infection as well as nonpersistent, nonprogressive infection with ADV.5,55 Ferrets infected with these forms of ADV could be seropositive without clinical disease. The judicious use of the ELISA tests, PCR, and DNA in situ hybridization may be best to characterize infection status in ferrets. At necropsy, gross changes in ferrets with Aleutian disease include hepatomegaly, splenomegaly, lymphadenopathy, and thymic enlargement. A positive diagnosis is made by demonstrating lesions consistent with ADV infection on histopathologic examination, such as periportal hepatitis with aggregates of lymphocytes, plasmacytes, and neutrophils. Bile duct hyperplasia and periportal fibrosis are common findings, and mild membranous glomerulonephrosis may also be seen. In the CNS form of the disease, perivascular cuffing in the brain and spinal cord and lymphoplasmacytic meningitis may be present. The DNA of ADV can be detected by in situ hybridization, confirming infection and identifying infected cells.22 This test can be also be used on formalin-fixed, paraffinembedded biopsy samples as an antemortem screening method to aid in the diagnosis and prognosis of the disease.
TREATMENT AND PREVENTION No definitive treatment exists for ADV infection in ferrets. In mink, immunosuppressive therapy with cyclophosphamide has been used to control infection for up to 16 weeks, but viral
CHAPTER 5 Cardiovascular and Other Diseases titers did not change in treated animals.10 Treatment of infected mink kits with gamma globulin-containing ADV antibody has decreased mortality rates.1 In ferrets with clinical disease, supportive care, use of anti-inflammatory agents, and immunosuppressive therapy with prednisone or cyclophosphamide should be considered. No vaccine exists for Aleutian disease, and vaccination is probably contraindicated given the immune-mediated nature of the disease. In mink farms, testing by CIEP and removal of animals that test positive has been effective in eradicating the disease11; testing and removal or isolation should be considered as a control method in ferret colonies and shelters as well. Excellent hygiene is imperative in light of the ability of humans to act as vectors; in addition, strict quarantine protocols should be followed, since asymptomatically infected animals can shed virus in bodily secretions.
SPLENOMEGALY Splenomegaly is a very common clinical finding in ferrets older than 1 year of age. A variety of conditions can cause splenomegaly, including extramedullary hematopoiesis (EMH), neoplasia, hypersplenism, and heart disease. Splenic torsion, abscess, and rupture are rare in ferrets. Splenomegaly is usually a normal finding, unrelated to any disease state. Often, splenomegaly is present concurrently and as an incidental finding with diseases such as adrenal disease and insulinoma. In some ferrets, the spleen is so enlarged and pendulous that the animal has difficulty lifting its abdomen off the ground. In most cases, the enlarged spleen is of little or no clinical significance. The normal ferret spleen is quite large for the size of the animal, measuring approximately 5 cm in length, 2 cm in width, and 1 cm in thickness. The spleen may enlarge slightly with age in ferrets, for reasons that are poorly understood. Chronic immune stimulation or compensation for erythroid bone marrow insufficiency may play a role. More research is needed to determine if splenomegaly is indicative of other disease problems. The most common cause of splenomegaly in ferrets is extramedullary hematopoiesis (EMH). This appears grossly as a smooth dark-red spleen. The cause of this syndrome is unknown, although compensation for myeloid insufficiency has been suggested.16 Most ferrets with EMH do not show evidence of anemia or other hematologic deficiencies. Splenic lymphoproliferation and resultant splenomegaly in response to chronic immune stimulation has also been suggested as a cause. In mice, evidence exists that chronic infection with Helicobacter felis induces this change13; a similar situation may exist in ferrets infected with Helicobacter mustelae. Although hemangioma, hemangiosarcoma, and other tumors can occur, lymphoma is the most common neoplasia of the ferret spleen. On gross examination, a spleen with lymphoma often has irregular borders and texture as well as white to tan nodules on the surface and within the parenchyma. Splenic neoplasia often involves other organs, such as the liver and intestinal lymph nodes. Hypersplenism is rare in ferrets. This disease involves destruction of one or more blood cell lines by the reticuloendothelial system within the spleen. Affected ferrets have blood dyscrasias such as anemia, leukopenia, thrombocytopenia, or pancytopenia. The bone marrow may be normal or hypercellular. Clinical signs are associated with the predominantly decreased blood cell count; they include weakness, pallor, petechiae, and
73
secondary infections. Diagnosis is made on the basis of abnormal CBC findings coupled with cytologic results indicating a normal to hypercellular bone marrow and no other sign of blood loss, infection, or neoplasia. Results of radiographs and plasma biochemical analyses are usually normal but help to rule out other possible causes of disease. The relationship of EMH to hypersplenism is unclear. In one report, ferrets with anemia of undetermined cause responded to splenectomy, but the histologic appearance of the spleen was found to be that of EMH.16 Two of these ferrets developed systemic disease (lymphoma and multifocal granulomatous disease) within 4 months after splenectomy. In the diagnostic approach to an enlarged spleen in a ferret, perform a thorough examination with particular attention to palpation of the spleen. With EMH, the spleen is usually of normal shape and consistency with regular borders; neoplasia of the spleen may cause irregular borders and palpable lumps within the spleen. Palpation is best done with care, because some diseases cause the spleen to be more friable and splenic rupture is possible. Radiographs may be helpful to define the borders of the spleen and to evaluate for other abnormalities that may be associated with splenomegaly, such as hepatomegaly, cardiomegaly, and other systemic problems. Aspiration or biopsy can best determine the cause of splenomegaly. Splenic aspiration with a 22- or 25-gauge needle and a 3-mL syringe can usually be performed with manual restraint. On cytologic examination of an aspirate, EMH typically shows a mixed population of mature and immature red cells, whereas lymphoma or other lymphoproliferative diseases show a predominant or homogenous population of lymphocytes (see Chapter 36). Biopsy of the spleen is best performed surgically or endoscopically; however, surgery for the sole purpose of obtaining a splenic biopsy is rarely indicated unless other tests are nondiagnostic and primary splenic disease is suspected. Obtaining biopsy samples is the best method to maintain cellular architecture for an accurate diagnosis. Percutaneous biopsy equipment should not be used because it is too large and too long for use in ferrets. Ultrasonography of the spleen should be used to evaluate for areas of altered echogenicity. Typically the spleen is of uniform echogenicity with EMH, whereas with neoplasia, the splenic parenchyma appears mottled. If the splenic parenchyma is irregular, a diagnostic sample should be obtained by ultrasound-guided aspiration. Treatment of splenomegaly depends on the cause. No treatment is known or required for EMH; however, if the spleen has become so large that movement is hampered, splenectomy may be warranted. Do this cautiously, because anemia can result. Splenectomy is recommended for hypersplenism, rupture, torsion, infection, or neoplasia of the spleen. Ferrets with peripheral lymphomas do not necessarily benefit from splenectomy.
ANEMIA As in other species, anemia in ferrets is caused by decreased production of erythrocytes, destruction of existing erythrocytes, or blood loss. The normal hematocrit of ferrets (46%-61%) is higher than that of other species; therefore mild anemia may go unrecognized unless clinicians keep this fact in mind. Erythrocyte numbers are also higher; cell numbers can be as high as 17.4 × 106/μL.59 As in other species, hemoglobin concentration and mean cell volume, respectively, can be used to characterize
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SECTION I Ferrets
the anemia as normochromic or hypochromic and as normocytic, macrocytic, or microcytic. Normal reticulocyte counts in ferrets can be as high as 10%. Counts higher than 12% indicate a regenerative response. Examination of the bone marrow is indicated in any ferret with a nonregenerative anemia that does not respond to treatment within 3 to 6 days. Decreased production of erythrocytes can be caused by chronic disease or inflammation, bone marrow suppression, or neoplasia. Anemia of chronic disease, also called anemia of chronic inflammation, can develop with any long-term illness. It results from decreased iron availability, decreased erythrocyte survival, or decreased response to the anemia.81 Cytokines secreted during inflammation mediate the response and cause a nonregenerative normocytic normochromic anemia. In animals with this type of anemia, results of bone marrow examination are usually normal. In pet ferrets, hyperestrogenemia resulting from an ovarian remnant or adrenal disease is the most common cause of bone marrow suppression. Chronic estrus as a cause of hyperestrogenemia is rare in the United States because pet ferrets are typically spayed before purchase. Neoplasia that affects the bone marrow (e.g., leukemia, myeloma) can suppress erythroid series production because the marrow is replaced by tumor or fibrosis.35 This typically results in a normocytic normochromic anemia with a low reticulocyte count. Systemic neoplasia can also cause an anemia of chronic disease. Erythrocytes can be destroyed by immune-mediated disease, toxins, parasites, or septicemia. Idiopathic immune-mediated hemolytic anemia has not been documented in ferrets, and diagnosis would be difficult because ferret antibody-specific reagents for the Coombs test are not available. No viral diseases or blood parasites that elicit immune-mediated hemolysis of red blood cells are known to exist in ferrets. Heavy-metal toxicosis and certain drugs are possible causes of hemolytic anemia. Zinc toxicosis as a cause of hemolytic anemia in a ferret has been documented.74 This type of anemia is uncommon in ferrets, but it may be underdiagnosed because of the inherent difficulties in identifying the toxic agents. Anemia secondary to blood loss can be caused by trauma, bleeding lesions, parasites, or hemostatic disorders. Trauma is common in ferrets because of their inquisitive nature, and hemorrhage can be internal or external. Bleeding ulcers develop commonly with gastritis caused by Helicobacter mustelae infection, gastrointestinal foreign bodies or trichobezoars, or other causes (e.g., chronic use of nonsteroidal anti-inflammatory drugs [NSAIDs]). Parasitism is a less common cause of anemia in ferrets, although coccidiosis in young ferrets or flea infestation can be severe enough to cause anemia. Hemostatic disorders such as thrombocytopenia can develop from estrogen toxicity. Other coagulopathies, such as rodenticide poisoning and disseminated intravascular coagulation, can also occur. An acute hemorrhagic syndrome of ferrets was reported in young ferrets between 8 and 24 weeks of age.31 These ferrets were all recently shipped to distributors or pet stores. Clinical signs started with epistaxis and oral ulceration, with hemorrhage that progressed to petechiae, ecchymoses, internal hemorrhage, and death. Treatment with vitamin K and supportive care was successful in early cases. The cause of this syndrome was unknown, but exposure to toxins (including NSAIDs) or viral agents was considered. To diagnose anemia in ferrets, obtain a complete history, perform a thorough physical examination, and submit samples
for a CBC, reticulocyte count, and, if indicated, bone marrow examination. A plasma biochemical analysis, radiographs, and an abdominal ultrasound examination may be helpful depending on the differential diagnosis. In analyzing test results, pay particular attention to the red blood cell values and the reticulocyte count. The treatment of the anemia is tailored to the specific cause. Specific supportive care can include one or more transfusions of fresh whole blood, iron dextran therapy, hemoglobin solutions, and erythropoietin. Ferrets lack blood types and transfusion reactions are difficult to induce, even with repeated transfusions between the same animals. Indications for transfusion in a ferret are a low packed cell volume (PCV), worsening clinical signs, and the possibility of continued blood loss. The decision to transfuse a ferret is based on its clinical status and the PCV. Transfusion in a ferret with acute blood loss may be considered if the PCV falls to less than 25%. If the anemia has developed gradually, the ferret may tolerate a lower PCV, although its clinical status may benefit from a transfusion if the PCV is less than 25%. In determining the need for transfusion, take into account the volume of blood that has been extracted for diagnostic tests. The blood volume required for transfusion can be estimated by the following formula: BVdonor = BVrecipient × [(PCVpost − PCVpre ) / PCVdonor ]
where BV is blood volume in milliliters (calculated as 8% of the body weight in kilograms) and PCVpost is the desired PCV after transfusion. The value of PCVpost is ideally within the reference range but it is more often 5% to 10% higher than the pretransfusion PCV (PCVpre). A simpler method is to administer 10% to 20% of the recipient’s blood volume in a single transfusion given over at least 4 hours. A filter is advised to prevent clots from being administered. Donor blood is collected in sodium citrate or acid-citrate-dextran (ACD). If citrate is used, 0.1 mL is used for each 0.9 mL of blood collected; for ACD, 1 mL per 6 mL of blood is recommended.27 Long-term storage of ferret blood is possible with the use of typical mammalian storage media, but it is not currently practiced. Because they are inexpensive and generally safe, oral iron supplements are preferred for treatment in small animals. Ferrous salts are absorbed better than ferric salts. A total daily dosage of 15 mg/kg, divided two or three times is recommended in small animals.24 Continue treatment for several weeks after the anemia resolves to replenish whole-body iron stores. Hemoglobin substitutes, such as hemoglobin glutamer-200 (bovine) solution (Oxyglobin, Biopure Corporation, Cambridge, MA), are safe and effective in ferrets; however, at the time of this writing, Oxyglobin is not available. This drug has colloidal properties and should be administered slowly in normovolemic animals and in animals with kidney disease, heart disease, or risk of pulmonary edema. Side effects include discoloration of serum or plasma, skin, mucous membranes, sclerae, and urine, which is nonpathogenic and transient. Plasma or serum biochemical values may be affected by this discoloration, so a blood sample for these tests should be submitted before this solution is administered. The dose is 11 to 15 mL/kg administered intravenously or intraosseously over 4 hours.51 Although erythropoietin is rarely indicated, it can be used in ferrets with chronic renal failure at 100 U/kg three times weekly until the PCV is stabilized and then twice weekly.
CHAPTER 5 Cardiovascular and Other Diseases
IBUPROFEN TOXICOSIS Ibuprofen toxicosis has been documented in many ferrets9,60 and is seen clinically, often as an emergency presentation. Prostaglandin inhibition by ibuprofen and other NSAIDs can cause altered renal blood flow, ulceration of the gastrointestinal tract, and platelet dysfunction. Clinical signs include vomiting, CNS depression, anorexia, diarrhea, and melena. Severe overdose can cause renal failure, resulting in azotemia and oliguria or anuria. Gastrointestinal signs develop as early as 2 to 6 hours after ingestion; renal signs may develop within 12 hours after ingestion or as long as 5 days later.52 Seizures can occur with massive overdosage. Treatment depends on the clinical signs, time elapsed since ingestion, electrolyte imbalances, and azotemia. Treatment is directed at preventing absorption (if possible), managing bleeding ulcers, and maintaining renal perfusion and electrolyte balance. Induction of emesis is effective only within the first 30 to 60 minutes after ingestion, although activated charcoal may be beneficial if given within the first few hours. Intravenous fluids (0.45% NaCl and 2.5% dextrose) should be administered even if no signs of renal compromise are apparent. To control hemorrhage and hypotension, electrolyte solutions, colloids, and transfusions may be needed. Gastrointestinal protectants, such as ranitidine (24 mg/kg q8h), cimetidine (5-10 mg/kg q8-12h), omeprazole (0.7 mg/kg q24h), or sucralfate (25 mg/kg q8h), are helpful in controlling ulceration.
References 1. Aasted B, Alexandersen S, Hansen M. Treatment of neonatally Aleutian disease virus (ADV) infected mink kits with gammaglobulin containing antibodies to ADV reduces the death rate of mink kits. Acta Vet Scand. 1988;29:323-330. 2. Atkins CE, Gallo AM, Kurzman ID, et al. Risk factors, clinical signs, and survival in cats with a clinical diagnosis of idiopathic hypertrophic cardiomyopathy: 74 cases (1985-1989). J Am Vet Med Assoc. 1992;201:613-618. 3. Baumwart RD, Orvalho J, Meurs KM. Evaluation of serum cardiac troponin I concentration in boxers with arrhythmogenic right ventricular cardiomyopathy. Am J Vet Res. 2007;68:524-528. 4. Blair LS, Campbell WC. Suppression of maturation of Dirofilaria immitis in Mustela putorius furo by single dose of ivermectin. J Parasitol. 1980;66:691-692. 5. Bloom ME, Race RE, Wolfinbarger JB. Identification of a nonvirion protein of Aleutian disease virus: mink with Aleutian disease have antibody to both virion and nonvirion proteins. J Virol. 1982;43:608-616. 6. Bone L, Battles AH, Goldfarb RD, et al. Electrocardiographic values from clinically normal, anesthetized ferrets (Mustela putorius furo). Am J Vet Res. 1988;49:1884-1887. 7. Calvert C, Brown J. Use of M-mode echocardiography in the diagnosis of congestive cardiomyopathy in Doberman pinschers. J Am Vet Med A-ssoc. 1986;189:293-297. 8. Campbell WC, Blair LS. Dirofilaria immitis: experimental infections in the ferret (Mustela putorius furo). J Parasitol. 1978;64:119-122. 9. Cathers TE, Isaza R, Oehme F. Acute ibuprofen toxicosis in a ferret. J Am Vet Med Assoc. 2000;216:1426-1428. 10. Cheema A, Henson JB, Gorham JR. Aleutian disease of mink: prevention of lesions by immunosuppression. Am J Pathol. 1972;55:543-546. 11. Cho JG, Greenfield J. Eradication of Aleutian disease of mink by eliminating positive counter immunoelectrophoresis reactors. J Clin Microbiol. 1978;7:18-21.
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12. Daoust PY, Hunter DB. Spontaneous Aleutian disease in ferrets. Can Vet J. 1978;19:133-135. 13. Enno A, O’Rourke JL, Howlett CR, et al. MALToma-like lesions in the murine gastric mucosa after long-term infection with Helicobacter pylori-induced gastric lymphoma. Am J Pathol. 1995;147:217. 14. Ensley PK, Van Winkle T. Treatment of congestive heart failure in a ferret (Mustela putorius furo). J Zoo Anim Med. 1982;13:23-25. 15. Erdman SE, Brown SA, Kawasaki TA. Clinical and pathologic findings in ferrets with lymphoma: 60 cases (1982-1994). J Am Vet Med Assoc. 1996;208:1285-1290. 16. Erdman SE, Xiantang L, Fox JG. Hematopoietic diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Philadelphia: Lippincott Williams & Wilkins; 1998:231-246. 17. Fox JG. Normal clinical and biologic parameters. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:159-173. 18. Fox JG, Pearson RC, Gorham JR. Viral diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Philadelphia: Lippincott Williams & Wilkins; 1998:355-374. 19. Fox PR. Feline cardiomyopathies. In: Fox PR, Sisson D, Moise NS, eds. Textbook of canine and feline cardiology: Principles and clinical practice. 2nd ed. Philadelphia: WB Saunders; 1999:896-923. 20. Greenlee PG, Stephens E. Meningeal cryptococcosis and congestive cardiomyopathy in a ferret. J Am Vet Med Assoc. 1984;184:840-841. 21. Haas GJ. Etiology, evaluation, and management of acute myocarditis. Cardiol Rev. 2001;9:88-95. 22. Haas L, Lochelt M, Kaaden OR. Detection of Aleutian disease virus DNA in tissues of naturally infected mink. J Gen Virol. 1988;69:705-710. 23. Haggstrom J, Boswood A, O’Grady M, et al. Effect of pimobendan or benazepril hydrochloride on survival times in dogs with congestive heart failure caused by naturally occurring myxomatous mitral valve disease: the QUEST study. J Vet Intern Med. 2008;22:1124-1135. 24. Harvey JW, French TW, Meyer DJ. Chronic iron deficiency anemia in dogs. J Am Anim Hosp Assoc. 1982;18:946-960. 25. Herndon WE, Kittleson MD, Sanderson K, et al. Cardiac troponin I in feline hypertrophic cardiomyopathy. J Vet Intern Med. 2002;16:558-564. 26. Hillyer EV. Cardiovascular diseases: Part II. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders; 1997:71-76. 27. Hoefer HL. Transfusions in exotic species. Prob Vet Med. 1992;4:625-635. 28. Hollenberg NK. Control of renal perfusion and function in congestive heart failure. Am J Cardiol. 1988;62:72E-76E. 29. John J, Awab A, Norman D, et al. Activated prtein C improves survival in severe sepsispatients with elevated troponin. Intens Care Med. 2007;33:2122-2128. 30. Johnson L, Boon J, Orton EC. Clinical characteristics of 53 dogs with Doppler-derived evidence of pulmonary hypertension: 1992-1996. J Vet Intern Med. 1999;13:440-447. 31. Johnson-Delaney KA, Reavill DR. Ferret acute hemorrhagic syndrome. Proceedings. 29th Annual Assoc Avian Vet Conf. 2008;49-50. 32. Kenyon AJ, Howard E, Buko L. Hypergammaglobulinemia in ferrets with lymphoproliferative lesions (Aleutian disease). Am J Vet Res. 1967;28:1167-1172. 33. Kenyon AJ, Kenyon BJ, Hanhn ED. Protides of the Mustelidae: immunoresponse of mustelids to Aleutian mink disease virus. Am J Vet Res. 1978;39:1011-1015. 34. Kenyon AJ, Magnano T, Helmboldt CF, et al. Aleutian disease in the ferret. J Am Vet Med Assoc. 1966;149:920-924. 35. Kisseberth WC, MacEwen EG. Complications of cancer and its treatment. In: Withrow SJ, MacEwen EG, eds. Small animal clinical oncology. 2nd ed. Philadelphia: WB Saunders; 2001:198-218.
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36. Kittleson MD. Diagnosis and treatment of arrhythmias (dysrhythmias). In: Kittleson MD, Kienle RD, eds. Small animal cardiovascular medicine. St Louis: Mosby; 1998:449-494. 37. Kittleson MD. Management of heart failure. In: Kittleson MD, Kienle RD, eds. Small animal cardiovascular medicine. St Louis: Mosby; 1998:149-194. 38. Komukai K, Kurihara S. Mechanisms of the inotropic effects of UD-CG 212 Cl, an active metabolite of pimobendan, on ferret papillary muscles. J Cardiovasc Pharmacol. 1996; 27:673-679. 39. Lee JA, Allen DG. EMD 53998 sensitizes the contractile proteins to calcium in intact ferret ventricular muscle. Circ Res. 1991;69:927-936. 40. Lipman NS, Murphy JC, Fox JG. Clinical, functional and pathologic changes associated with a case of dilatative cardiomyopathy in a ferret. Lab Anim Sci. 1987;37:210-212. 41. Lombard CW, Jons O, Bussadori CM. Clinical efficacy of pimobendan versus benazepril for the treatment of acquired atrioventricular valvular disease in dogs. J Am Anim Hosp Assoc. 2006;42:249-261. 42. Mellor PJ, Mellanby RJ, Baines EA, et al. High serum troponin I concentration as a marker of severe myocardial damage in a case of suspected exertional heatstroke in a dog. J Vet Cardio. 2006;8:55-62. 43. Meredith A. Ferrets: systemic viral diseases. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester, UK: BSAVA; 2009:330-335. 44. Miller WR, Merton DA. Dirofilariasis in a ferret. J Am Vet Med Assoc. 1982;180:1103-1104. 45. Moise NS, Dietze AE, Mezza LE, et al. Echocardiography, electrocardiography, and radiography of cats with dilated cardiomyopathy, hypertrophic cardiomyopathy, and hyperthyroidism. Am J Vet Res. 1986;7:1477-1485. 46. Moise NS, Fox PR. Echocardiography and Doppler imaging. In: Fox PR, Sisson D, Moise NS, eds. Textbook of canine and feline cardiology: Principles and clinical practice. 2nd ed. Philadelphia: WB Saunders; 1999:130-172. 47. Moreland AF, Battles AH, Nease JH. Dirofilariasis in a ferret. J Am Vet Med Assoc. 1986;188:864. 48. Murakami M, Matsuba C, Une Y, et al. Nucleotide sequence and polymerase chain reaction/restriction fragment length polymorphism analyses of Aleutian disease virus in ferrets in Japan. J Vet Diagn Invest. 2001;13:337-340. 49. Ohshima K, Shen DT, Henson JB, et al. Comparison of the lesions of Aleutian disease in mink and hypergammaglobulinemia in ferrets. Am J Vet Res. 1978;39:653-657. 50. Opie LH. Diuretics. In: Opie LH, Hersh BJ, eds. Drugs for the heart. 5th ed. Philadelphia: WB Saunders; 2000:83-104. 51. Orcutt C. Oxyglobin administration for the treatment of anemia in ferrets. Exot DVM. 2001;2.3:44-46. 52. Owens-Clark J, Dorman CD. Toxicities from newer over-thecounter drugs. In: Bonagura JD, ed. Kirk’s current veterinary therapy XIII: small animal practice. Philadelphia: WB Saunders; 2000:227-230. 53. Oxenham M. Aleutian disease in the ferret. Vet Rec. 1990;126:585. 54. Oyama MA, Sisson DD. Cardiac troponin-I concentration in dogs with cardiac disease. J Vet Intern Med. 2004;18:831-839. 55. Palley LS, Corning BF, Fox JG, et al. Parvovirus-associated syndrome (Aleutian disease) in two ferrets. J Am Vet Med Assoc. 1992;201:100-106. 56. Parrott TY, Greiner EC, Parrott JD. Dirofilaria immitis infection in three ferrets. J Am Vet Med Assoc. 1984;184:582-583. 57. Pennick KE, Stevenson MM, Latimer DS, et al. Persistent viral shedding during asymptomatic Aleutian mink disease parvoviral infection in a ferret. J Vet Diagn Invest. 2005;17:594-597.
58. Porte HG, Porter DD, Larsen AE. Aleutian disease in ferrets. Infect Immun. 1982;36:379-386. 59. Quesenberry KE. Basic approach to veterinary care. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders; 1997:14-25. 60. Richardson JA, Balabuszko RA. Ibuprofen ingestion in ferrets: 43 cases. J Vet Emerg Crit Care. 2001;11:53-59. 61. Rosenthal K. Ferrets. Vet Clin North Am Small Anim Pract. 1994;24:1-21. 62. Rush JE, Freeman LM, Fenollosa NK, et al. Population and survival characteristics of cats with hypertrophic cardiomyopathy: 260 cases (1990-1999). J Am Vet Med Assoc. 2002;220:202-207. 63. Saifuddin M, Fox JG. Identification of a DNA segment in ferret Aleutian disease virus similar to a hypervariable capsid region in mink Aleutian disease parvovirus. Arch Virol. 1996;141:1329-1339. 64. Salim A, Velmahos GC, Jindal A, et al. Clinically significant blunt cardiac trauma: role of serum troponin levels combined with electrocardiographic findings. J Trauma. 2001;50:237-243. 65. Sasai H, Kato T, Sasaki S, et al. Echocardiographic diagnosis of dirofilariasis in a ferret. J Small Anim Pract. 2000;41:172-174. 66. Schober KE, Cornand C, Kirbach B, et al. Serum cardiac troponin I and cardiac troponin T concentrations in dogs with gastric dilatation-volvulus. J Am Vet Med Assoc. 2002;221:381-388. 67. Selting KA, Lana SE, Ogilvie GK, et al. Cardiac troponin I in canine patients with lymphoma and osteosarcoma receiving doxorubicin: comparison with clinical heart disease in a retrospective analysis. Vet Comp Oncol. 2004;2:142-156. 68. Smith SH, Bishop SP. The electrocardiogram of normal ferrets and ferrets with right ventricular hypertrophy. Lab Anim Sci. 1985;35:268-271. 69. Snyder PS, Atkins CE. Current uses and hazards of the digitalis glycosides. In: Kirk RW, Bonagura JD, eds. Kirk’s current veterinary therapy XI: small animal practice. Philadelphia: WB Saunders; 1992:689-693. 70. Stamoulis ME, Miller MS, Hillyer EV. Cardiovascular diseases. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders; 1997:63-76. 71. Stepien RL, Benson KG, Forrest LJ. Radiographic measurement of cardiac size in normal ferrets. Vet Radiol Ultrasound. 1999;40:606-610. 72. Stepien RL, Benson KG, Wenholz LJ. M-mode and Doppler echocardiographic findings in normal ferrets sedated with ketamine hydrochloride and midazolam. Vet Radiol Ultrasound. 2000;41:452-456. 73. Stewart JD, Rozengurt N. Aleutian disease in the ferret. Vet Rec. 1993;133:172. 74. Straube EF, Schuster NH, Sinclair AJ. Zinc toxicity in the ferret. J Comp Pathol. 1980;90:355-361. 75. Supakorndej P, Lewis RE, McCall JW. Radiographic and angiographic evaluations of ferrets experimentally infected with Dirofilaria immitis. Vet Radiol Ultrasound. 1995;36:23-29. 76. Supakorndej P, McCall JW, Jun JJ. Early migration and development of Dirofilaria immitis in the ferret, Mustela putorius furo. J Parasitol. 1994;80:237-244. 77. Thomas WP, Gaber CE, Jacobs GJ, et al. Recommendations for standards in transthoracic two-dimensional echocardiography in the dog and cat. Echocardiography Committee of the Specialty of Cardiology, American College of Veterinary Internal Medicine. J Vet Intern Med. 1993;7:247-252. 78. Thornton RN, Cook TG. A congenital toxoplasma-like disease in ferrets (Mustela putorius furo). N Z Vet J. 1986;34:31-33. 79. Tidholm A, Jonsson L. A retrospective study of canine dilated cardiomyopathy (189 cases). J Am Anim Hosp Assoc. 1997;33:544-550.
CHAPTER 5 Cardiovascular and Other Diseases 80. Une Y, Wakimoto Y, Nakano Y, et al. Spontaneous Aleutian disease in a ferret. J Vet Med Sci. 2000;62:553-555. 81. Waner T, Harrus S. Anemia of inflammatory disease. In: Feldman BF, Zinkl JG, Jain NC, eds. Schalm’s veterinary hematology. 5th ed. Philadelphia: Lippincott Williams & Wilkins; 2000:205-209. 82. Welchman D. de B, Oxenham M, Done SH: Aleutian disease in domestic ferrets: diagnostic findings and survey results. Vet Rec. 1993;132:479-484.
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83. Wells SM, Sleeper M. Cardiac troponins. J Vet Emerg Crit Care. 2008;18:235-245. 84. Wolfensohn HHL. Aleutian disease in laboratory ferrets. Vet Rec. 1994;134:100.
CHAPTER
6
Respiratory Diseases
Heather W. Barron, DVM, Diplomate ABVP (Avian), and Karen L. Rosenthal, DVM, MS
Canine Distemper Virus History and Physical Examination Diagnosis Treatment Prevention Influenza History and Physical Examination Diagnosis Treatment Prevention Pneumonia History and Physical Examination Diagnosis Treatment Prevention Pulmonary Mycoses History and Physical Examination Cryptococcosis Blastomycosis Coccidioidomycosis Other Causes of Respiratory Signs
Although there are few causes of primary respiratory disease in captive ferrets, several diseases may present with respiratory symptoms. Canine distemper virus (CDV) and influenza virus are the most common causes of primary respiratory disease. While distemper is invariably fatal in ferrets, influenza often resolves over the course of a couple of weeks unless it is complicated by bacterial pneumonia. Dyspnea, tachypnea, coughing, and other respiratory signs may manifest with a variety of conditions, including Aleutian disease, heartworm disease, congestive heart failure, lymphoma, trauma, anemia, heatstroke, anaphylactic reactions, and metabolic disturbances. Normal respiratory rate in ferrets is 33 to 36 breaths per minute; documenting this rate should be part of every physical examination along with careful auscultation. 78
CANINE DISTEMPER VIRUS Canine distemper virus belongs to the Morbillivirus genus, which also includes the measles virus (MV). Although different strains of the virus vary in virulence, canine distemper is typically a fatal disease in ferrets. It is one of the most prevalent viral diseases of dogs, and because it is ubiquitous, ferrets risk being exposed to this virus through canine companions. Reservoirs of CDV include members of the families Canidae, Mustelidae, and Procyonidae. The virus is most commonly transmitted by aerosol exposure.2 Direct contact with conjunctival and nasal exudates, urine, feces, and skin can also cause infection.21 Ferrets shed virus in all body excretions, and shedding begins about 7 days after exposure.2 Fomites are also implicated in transmission; on gloves, the virus is viable for up to 20 minutes.21 Once in a ferret’s body, the virus appears to spread by viremia.43 The incubation period in ferrets is typically 7 to 10 days, although incubation periods of up to 56 days have been reported in natural infections.21,44
HISTORY AND PHYSICAL EXAMINATION Infection with CDV should be suspected in any unvaccinated, exposed ferret showing compatible clinical signs. Unvaccinated ferrets of any age are equally susceptible to this disease. In dogs, pyrexia develops 3 to 6 days after infection with CDV and is soon followed by anorexia and a serous nasal discharge.2 A serous ocular discharge then appears; this discharge quickly becomes mucopurulent. In ferrets, the first sign of disease is usually a papular dermatitis on the chin, followed by a cheilitis characterized by swelling and crusting. These changes may be accompanied by dermatitis on the anus and inguinal area,21 which is orange-tinged in some ferrets. Other clinical signs are anorexia, depression, dyspnea, pyrexia, photophobia, pruritus, blepharospasm, and abundant mucopurulent oculonasal discharge. Hyperkeratosis of the planum nasale and footpads (Fig. 6-1) often occurs. Vomiting and diarrhea, which are seen in dogs with CDV, are uncommon in ferrets. The respiratory system is the preferred site for the virus to replicate.43 Secondary bacterial infections, which are responsible Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 6 Respiratory Diseases
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may be present. In the central nervous system, inflammatory cell invasion with demyelination is observed.2 History of exposure and clinical signs can be highly suggestive of infection. Nonspecific test results can include leukopenia, alpha- and beta-hyperglobulinemia, and radiographic evidence of lung congestion or consolidation.21,44,57 Nonregenerative anemia and increased serum levels of alpha and beta globulins are the most common routine laboratory changes.44 Clinical signs of infection with CDV can initially resemble those of influenza. However, within 1 to 2 days, the nasal and ocular discharge turns from serous to mucopurulent, and dermatitis develops around the chin and lips. When it is present with other clinical signs typical of CDV, this dermatitis is pathognomonic for CDV infection. Also, ferrets infected with CDV tend to be much sicker than those with influenza.
TREATMENT Fig. 6-1 Hyperkeratotic footpads in a ferret diagnosed with canine distemper virus (CDV). Photograph courtesy of Dr. David Perpinan.
for many of the severe respiratory symptoms and death, are caused by the immunosuppressive effects of the virus.57 Seizures and blindness are common in dogs with CDV infection,57 and neurologic signs may manifest without previous systemic signs.2 In ferrets with advanced CDV infection, incoordination, torticollis, and nystagmus can be present.2
DIAGNOSIS In the past, most laboratory tests available for the diagnosis of distemper were hampered by low sensitivity, low specificity, or both. A plasma sample can be submitted to measure an antibody titer against CDV. However, because both infected and vaccinated ferrets can have a positive titer, a positive result is not diagnostic of disease. In practical terms, if a ferret that has not been vaccinated has a positive titer, this test can confirm CDV infection. A fluorescent antibody test can be done on conjunctival smears, mucous membrane scrapings, or blood smears to identify CDV antigen in cells.21 However, this test is useful only in the first few days of disease, and false-negative results are possible. Modified live viral strains used for vaccination do not interfere with this test.57 Polymerase chain reaction (PCR) assays have been developed and may be used for ante- or postmortem diagnosis. Recent research suggests that nested PCR assays are more sensitive for antemortem diagnosis than reverse transcriptase PCR (RT-PCR).30,51 Samples of blood, urine, feces, or tissues or deep pharyngeal swabs should be submitted to commercial laboratories for CDV PCR testing. False-positive results may be seen in the first few weeks after vaccination with modified live vaccines. Killed and vector-recombinant vaccines do not interfere with PCR testing. A positive postmortem diagnosis can be made by fluorescent antibody staining of imprints from lymph nodes, bladder epithelium, and cerebellum.2 Histopathologic examination of affected cells can also confirm the disease. Inclusion bodies of CDV are usually intracytoplasmic but can be intranuclear. Inclusions are generally found in the epithelial cells of the trachea, urinary bladder, skin, gastrointestinal tract, lymph nodes, spleen, and salivary glands.21 Diffuse interstitial pneumonia
No specific treatment exists for CDV infection in ferrets, and the mortality rate may be up to 100%. Death generally occurs 12 to 16 days after exposure to ferret-adapted CDV strains and 21 to 35 days after exposure to canine wild virus strains. Euthanasia of affected ferrets is usually the most humane option. Palliative treatment consists of supportive care and antibiotics for secondary bacterial infections. Administration of anti-canine distemper hyperimmune serum may be useful if given early in the course of the disease.44
PREVENTION Vaccination is the best way to prevent CDV infection in ferrets. However, CDV vaccines are insufficient to induce protection in very young ferrets because of interference from maternal antibodies acquired via colostrum in the first few days of life. Because CDV is closely related to MV, vaccination with MV vaccine may offer cross protection at 5 to 6 weeks of age without interference from maternal antibodies.64 Otherwise, CDV vaccinations may be started at 6 or 8 weeks of age for kits from nonimmune or immune dams, respectively, and then continued every 3 to 4 weeks until the kits are at least 12 to 14 weeks old. At present, the recommendation is to revaccinate yearly. However, results of currently ongoing research demonstrate that antibody titers may remain high for several years, suggesting that boosters could be given less frequently. In other species, titers of 1:32 are considered protective, and in a clinical trial involving 66 CDV-vaccinated ferrets, average antibody titers were found to be over 1:1000 a year postvaccination (HL Heller, 2009, personal communication). Currently, only one CDV vaccine approved by the U.S. government is available for ferrets: PureVax Ferret Distemper Vaccine (Merial, Athens, GA). Production of Fervac-D (United Vaccines, Madison, WI), the only other approved ferret distemper vaccine, was permanently discontinued by the manufacturer. Avoid use of multivalent canine vaccines, which can be associated with adverse effects. Vaccine strains of CDV that have been propagated in cell lines of canine origin may induce distemper disease in ferrets. Signs of vaccine-induced distemper may include mild purulent upper respiratory tract infection with pyrexia that resolves in a week or progresses to fulminating distemper during the same time frame (see Chapter 2). Anaphylactic reactions in ferrets have been reported after vaccination.24,40 Most of these reactions occur after vaccination
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with canine distemper vaccines. Most reactions usually happen within 30 minutes after vaccination, with clinical signs of vomiting, diarrhea, pale mucous membranes, weak pulses, tachycardia, and lethargy. If a reaction occurs, treat the ferret for anaphylactic shock with epinephrine, parenteral fluids, steroids, antihistamines, and oxygen therapy as needed. It is prudent to suggest that a ferret owner remain in the hospital for up to 30 minutes after CDV vaccination in case a reaction should occur. PureVax ferret distemper vaccine is a recombinant canarypox vector vaccine that appears to be less likely to cause anaphylaxis. A possible link between myofasciitis and distemper vaccination has been suggested.22 If an outbreak of CDV occurs in a group of susceptible ferrets, all affected animals should be removed and the healthy ferrets vaccinated immediately. However, vaccinating nonimmunized ferrets may not stop infection and subsequent death in the face of an outbreak.21 CDV is relatively labile and its infectivity is destroyed by heat, drying, detergents, and disinfectants.57 Routine cleaning and disinfection procedures effectively destroy CDV on hard surfaces.
INFLUENZA Ferrets are susceptible to infection from both influenza type A and B viruses of the class Orthomyxoviridae. Natural outbreaks or clinical cases of influenza in ferrets have occurred with common human influenza type A viruses, the human strain of pandemic H1N1 virus, and swine-origin H1N1 influenza virus.10,42,55,58 The pathogenicity of type B influenza virus in ferrets appears to be low. In a recent report documenting natural cases of pandemic H1N1 influenza in ferrets, transmission was most likely from infected humans in the household.58 The influenza virus H3N8 that emerged in dogs in 2004 is most closely related to the equine influenza virus, whereas influenza A viruses affecting ferrets appear to have a pattern of viral attachment more similar to avian and human influenza subtypes.15,62 Although there is a theoretical potential of the virus being transmitted from ferrets to humans,36 there is only one report, from the 1930s, that documents a probable transmission of virus to humans. In that report, an animal-passage influenza strain was inoculated into a laboratory ferret, and a laboratory investigator was infected after close contact with the animal.54 Ferrets have long been important animal models of transmission, pathogenicity, and treatment studies of influenza virus in humans. Influenza virus is transmitted primarily by aerosol droplets from ferret to ferret or from human to ferret. The virus can be transmitted beginning at the height of pyrexia and continuing for the next 3 to 4 days.56 In ferrets as in people, influenza primarily causes upper respiratory disease. The different subtypes of influenza A viruses vary in virulence and likelihood of developing secondary bacterial infections, which accounts for the difference in severity of clinical signs.4,30,46
HISTORY AND PHYSICAL EXAMINATION Ferrets contract influenza after being exposed to infected people or other infected ferrets. All ferrets are susceptible to influenza, although the disease is typically more severe in neonates than in older ferrets. After a short incubation period, the body temperature increases and then decreases approximately 48 hours later.14,21,36,49 The fever may be biphasic throughout the course
of the disease. Bouts of sneezing, epiphora, and a mucoid or mucopurulent nasal discharge are common. Clinical signs can appear within 48 hours of exposure.14,36 Affected ferrets may become lethargic and anorexic,56 and photophobia and conjunctivitis may be present.5 Neonates develop a much more severe upper respiratory tract infection than adults, and death may ensue from lower airway obstruction.9,53 Clinical signs involving the lower respiratory tract are less common than those of the upper respiratory tract. Infection of the lower respiratory tract is usually confined to the bronchial epithelium51 and results from secondary bacterial infection. Death can ensue from secondary pulmonary infection with Lancefield group C hemolytic streptococci.36 Neonates are more likely than older ferrets to develop bronchiolitis and pneumonia14 and to die from lower respiratory tract infection.51 Influenza virus can infect the cells of the intestinal mucosa and cause a limited enteritis.23 The potential for hepatic dysfunction has been described in ferrets infected experimentally with influenza.31 Hearing loss has also been associated with influenza infection in ferrets.48
DIAGNOSIS The availability of anti-influenza drugs, which must be given early in infection in order to be effective, has emphasized the need for early diagnosis. Traditionally, a diagnosis of influenza was based on the presence of clinical signs typical of infection, a history of exposure to infected individuals, isolation of the virus from nasal secretions, and a high antibody titer.36 However, serologic testing and virus isolation are primarily retrospective tools. Experimentally, an enzyme-liked immunosorbent assay can detect antibodies against influenza A and can be used to rapidly establish a serologic diagnosis.10 Antibodies against influenza virus have been detected within 3 days after infection.38,41 More recently, polymerase chain reaction (PCR) assays and in-clinic antigen detection assays like Directigen Flu (Becton Dickinson, Franklin Lakes, NJ), have become available and are able to differentiate between human influenza types A and B with a simple nasopharyngeal swab taken within the first 48 to 72 hours after clinical signs develop. An important differential is CDV; Table 6-1 highlights important distinctions between influenza and distemper. Diagnostic tests available for influenza include rapid immunoassay, immunofluorescence assay, PCR assay, serology, and viral culture. A transient leukopenia can be seen with this disease. Increases in concentrations of blood urea nitrogen, creatinine, alanine aminotransferase, potassium, and albumin have been reported in infected ferrets, but plasma biochemical values are usually within reference intervals.31
TREATMENT Influenza has a 7- to 14-day course in adult ferrets and is associated with a low mortality rate. Most ferrets can be treated at home. Instruct owners to offer favorite foods, chicken or beef broth, or specialized diets (e.g., Carnivore Care, Oxbow Animal Health, Murdock, NE; Eukanuba Maximum-Calorie, The Iams Company, Dayton, OH). Force-feed and offer water by syringe as needed. If pneumonia is not a complicating factor, use a pediatric cough suppressant without alcohol (at the pediatric dosage on a per weight basis). Also a bronchodilator, such as aminophylline (4 mg/kg PO, IM q12h), may be used for symptomatic
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Table 6-1 Clinical Distinctions between Canine Distemper Virus and Influenza Virus Infections Clinical Findings
Canine Distemper Virus
Influenza
Nasal and ocular discharge Sneezing Coughing Pyrexia Dermatitis (chin, lips, inguinal) Footpad hyperkeratosis Central signs Outcome
+++ (Mucopurulent) + + +++ (> 40°C) +++ ++ +a Almost 100% fatal
++ (Mucoserous) +++ +++ ++b — — — Self-limitingc
Frequency of clinical signs: +, may be present; ++, common; +++, usual presentation; —, absent. aCentral nervous system signs seen in advanced stages of disease (rarely the only signs). bPyrexia occurs early in the course of disease and may be resolved by the time of presentation. cInfluenza virus infection can be fatal in neonates.
therapy. As with any flu patient, parenteral fluids to maintain hydration and antibiotics to treat secondary bacterial infections may be indicated. To relieve nasal congestion, intranasal delivery of phenylephrine can be effective.7 The antiviral medication amantadine (6 mg/kg PO q12h) (Symmetrel; ENDO Pharmaceuticals, Chadds Ford, PA) has been experimentally effective in treating ferrets with influenza, although resistance in humans is widely reported.15 Other antiviral medications include neuraminidase inhibitors like zanamivir (12.5 mg/kg as a one-time intranasal dose) (Relenza; GlaxoSmith Kline, Research Triangle Park, NC), and oseltamivir (5 mg/kg PO q12h × 10 days) (Tamiflu; Roche, Nutley, NJ). These have been shown to prevent and treat influenza infection and either agent may be used to greater effect in combination with amantadine.4,19,28 However, resistance to oseltamivir appears to be emerging among some influenza strains.52 Because ferrets are a good model with which to study influenza infection in people, they are frequently used in experimental studies to develop new anti-influenza drugs.63 Anti-influenza drugs used in humans may therefore be used to treat pet ferrets. Antibiotics can be used to control secondary bacterial infections of the respiratory tract. In neonates, death typically results from secondary bacterial infections; antibiotic therapy may thus reduce neonate mortality.25 The use of antipyretic drugs to control fever is of questionable merit because fever is an important host defense mechanism. In one study, ferrets given aspirin had lowered body temperature, but they shed more virus and their viral levels decreased less rapidly than those of ferrets not treated with an antipyretic.26 In a recent meta-analysis of the use of antipyretics in animal models of influenza virus, risk of mortality increased with the use of antipyretics (aspirin, paracetamol, and diclofenac).17 This suggests that fever is instrumental in restricting the severity of infection.26,53
PREVENTION Controlling influenza rests mainly on preventing exposure of susceptible ferrets to infected individuals. Newborn ferrets are protected from disease by milk-derived antibodies in immunized dams.27 Experimentally, ferrets remain resistant to infection from the same influenza strain 5 weeks after primary infection.21
Vaccinating ferrets against influenza virus is not generally r ecommended for several reasons. Influenza is a relatively benign disease in ferrets, and the wide antigenic variation of the virus makes vaccination difficult. Also, vaccination seems to confer only short-term immunity.21 However, if a vaccine is being given, the use of a live or recombinant rather than an inactivated vaccine should be considered, because they may induce a greater protective effect.18,33
PNEUMONIA Pneumonia is not a common diagnosis in ferrets. Viral causes of pneumonia include CDV and influenza virus. Aleutian disease virus, a parvovirus, is associated with interstitial pneumonia in mink kits1 and should be considered as a possible cause of pneumonia and dyspnea in ferrets, especially the young.60 Respiratory syncytial virus has been shown to cause rhinitis and infection in the lungs of ferrets, but clinical signs of pneumonia have not been seen.47 Pyogranulomatous pneumonia has recently been reported in association with a systemic coronavirus infection in ferrets; it appears to produce a disease syndrome similar to the dry form of feline infectious peritonitis (see Chapter 3).23,37,45 Bacterial pneumonia (Fig. 6-2) is characterized by a suppurative inflammatory process that affects the bronchial tree, the lung lobes, or both. Reported primary bacterial pathogens that cause pneumonia in ferrets are Streptococcus zooepidemicus, other streptococcal species, and numerous mycobacterial species.12,32,50,61 Gram-negative bacteria such as Escherichia coli, Klebsiella pneumoniae, and Pseudomonas aeruginosa have been isolated from ferrets.20 Other bacteria that have been isolated from the lungs of ferrets include Bordetella bronchiseptica and Listeria monocytogenes. An acute hemorrhagic syndrome has recently been described in young ferrets (8-24 weeks of age) and may result in interstitial pneumonia. Affected ferrets have a prolonged prothrombin time (PT) and activated partial thromboplastin time (APTT) when compared with unaffected ferrets.29 Pneumocystis carinii is known to infect the lungs of ferrets. Latent infections can become active with immune suppression.3,11 Diagnosis is based on identifying the organism in a tracheal or lung wash. Treatment recommendations for P. carinii
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SECTION I Ferrets
Fig. 6-2 Bacterial pneumonia in a ferret. The lungs are diffusely congested with dark areas of consolidation.
pneumonia, based on those for dogs, include pentamidine isethionate or trimethoprim-sulfamethoxazole. Two cases of mild endogenous lipid pneumonia have been documented on histologic examination of ferrets at necropsy at the Animal Medical Center (New York, NY) (K. Quesenberry, personal communication, 2003), and one case of mortality due to endogenous lipid pneumonia was confirmed at necropsy (D. Perpinan, personal communication, 2009). In suspected cases, lung aspiration or bronchiolar lavage may help to provide antemortem diagnosis, although biopsy is needed for definitive diagnosis. Documented cases in rodents and other mustelids appear to be idiopathic or secondary to other disease processes.6 Successful treatment of lipid pneumonia has been achieved in people with prednisolone and thus may be a therapeutic option in ferrets.8 Although exogenous lipid pneumonia has not been documented in ferrets, caution should be exercised in treating animals with a mineral oil-based preparation for gastrointestinal disease (e.g., trichobezoar). Chronic aspiration of mineral oil products has been associated with lipid pneumonia in cats and people.13,39
HISTORY AND PHYSICAL EXAMINATION Ferrets with pneumonia exhibit typical clinical signs such as labored breathing, dyspnea, cyanotic mucous membranes, increased lung sounds, nasal discharge, fever, lethargy, and anorexia. Fulminant pneumonia leading to sepsis and death has been reported.20
DIAGNOSIS The diagnosis of pneumonia should be based on the clinical signs, radiographic findings, and results of supportive diagnostic tests. Results of the CBC may reveal leukocytosis caused by a neutrophilia with a left shift. In young ferrets with evidence of interstitial pneumonia, positive results of serologic tests and high concentrations of gamma globulins may support a diagnosis of Aleutian mink disease. Early in the disease, radiographs may show an interstitial pattern that changes to an alveolar pattern (Fig. 6-3) as the pneumonia progresses. If aspiration pneumonia is present,
Fig. 6-3 Ventrodorsal radiograph demonstrating an alveolar pattern and air bronchograms in a ferret with pneumonia. Photograph courtesy of Dr. Nico Schoemaker.
dependent lung lobes are primarily involved. Marked bronchial patterns suggest primary airway disease. Microbial cultures of tracheal or lung wash samples are invaluable in establishing a diagnosis and in treating ferrets with pneumonia. Submit samples for culture (aerobic or anaerobic bacterial, fungal, mycobacterial, or other) based on cytologic analysis of the collected fluid and debris. Cytologic assessment of tracheal wash samples from a ferret with pneumonia typically reveals septic inflammation and degenerating neutrophils. Results may also suggest the severity, cause, and chronicity of disease.
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TREATMENT Treat ferrets with pneumonia with good supportive care, including fluid therapy, force-feeding, and oxygen therapy as needed as well as with antimicrobials tailored according to test results. First-line antibiotics to consider before the results of culture and sensitivity testing are known are the quinolones, trimethoprimsulfamethoxazole, chloramphenicol, or the cephalosporins. Anecdotally, azithromycin at a dose of 5 mg/kg PO q24h also appears to be effective. In a report of two ferrets with mycobacterial pneumonia, both responded successfully to clarithromycin.32 Combination antibiotic therapy may be indicated. The prognosis depends on the cause of pneumonia and response to treatment. Most ferrets with bacterial pneumonia respond to antibiotic therapy and supportive care.
PREVENTION Bordetellosis is rare in ferrets. Nonetheless there is pervasive information in the lay literature about disease prevention. The best way to prevent B. bronchiseptica infection is to avoid hospitalizing ferrets where dogs, rabbits, or other common carriers are present. Anectdotally, a killed, injectable Bordetella bacterin may be effective in preventing bordetellosis in ferrets when used in accordance with manufacturer recommendations for dogs; however, no published reports supports this claim. The canine modified live intranasal Bordetella bacterin may cause disease in ferrets and is not recommended.
PULMONARY MYCOSES Pulmonary mycoses are uncommon in pet ferrets. Because ferrets in the United States are usually indoor pets, exposure to mycotic spores, which are mainly found in the soil, is unlikely.
HISTORY AND PHYSICAL EXAMINATION Not all animals with mycoses exhibit signs consistent with pulmonary disease. If lesions develop in the lungs, animals usually cough. Other signs consistent with a mycotic infection are wasting, lethargy, anorexia, lymph node enlargement, lameness, ocular and nasal discharge, and draining tracts unresponsive to antibiotic therapy.16,59 The prognosis for ferrets with pulmonary mycoses is poor.
CRYPTOCOCCOSIS Cryptococcosis, caused by Cryptococcus bacillisporus (formerly C. neoformans var gattii) and C. neoformans var grubii, has been diagnosed in a small number of ferrets.34,35 Infection can cause rhinitis, pneumonia, and pleuritis. Additionally, regional lymph node involvement is common and may also be expected to cause dyspnea when the retropharyngeal or mediastinal lymph nodes are involved.34 Invasive cryptococcal rhinitis has been successfully treated with itraconazole and surgical debulking.35
BLASTOMYCOSIS Blastomycosis, caused by Blastomyces dermatitidis, is endemic in the southeastern United States, the Mississippi River Valley, and the Ohio River Valley.59 Experimentally, the incubation period is 5 to 12 weeks. The mycelial phase is found in the soil, and
Fig. 6-4 The heart of a ferret that presented for moderate dyspnea and coughing. Necropsy demonstrated 3 female and 7 male Dirofilaria immitis worms in the heart.
the yeast form is found in the tissues. Diagnosis is made on the basis of a history of travel to an endemic region, clinical signs consistent with disease, results of cytologic assessment, positive periodic acid-Schiff reaction, or culture of B. dermatitidis. Amphotericin B and ketoconazole or itraconazole are recommended for treatment.59 Dosages should be based on those used for cats.
COCCIDIOIDOMYCOSIS Coccidioides immitis, the causative agent of coccidioidomycosis, is endemic in the southwestern United States and parts of Latin America. Primary infection develops after a susceptible host inhales the mycelia. Once in the host, spherules form and then produce endospores.16,59 Pulmonary signs develop 1 to 3 weeks after infection. Diagnosis of this disease is based on identifying the spherules on cytologic examination; they appear as refractile double-walled bodies.54 Recommended treatment, which is based on that for cats with coccidioidomycosis, includes the use of amphotericin B and ketoconazole or itraconazole.16,59
OTHER CAUSES OF RESPIRATORY SIGNS Differential diagnoses for tachypnea, dyspnea, and respiratory distress are similar to those for other small animals. After the history and physical examination, chest and abdominal radiography is the most important tool to differentiate the causes of lower respiratory tract symptoms. Ferrets that have severe traumatic injuries, such as from a fall from a great height, can develop pneumothorax or diaphragmatic hernia. These animals should be managed as one would a dog or cat with the same injuries. Ferrets with heartworm disease often present with coughing and tachypnea as the only clinical signs, even with moderate worm burdens (see Chapter 5) (Fig. 6-4).
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SECTION I Ferrets
References 1. Alexandersen S, Larsen S, Aasted B, et al. Acute interstitial pneumonia in mink kits inoculated with defined isolates of Aleutian mink disease parvovirus. Vet Pathol. 1994;21:216-228. 2. Appel M. Canine distemper virus. In: Appel M, ed. Virus infections of carnivores. New York: Elsevier Science; 1987:133-139. 3. Bauer NL, Paulsrud JR, Bartlett MS, et al. Pneumocystis carinii organisms obtained from rats, ferrets, and mice are antigenically different. Infect Immun. 1993;61:1315-1319. 4. Boltz DA, Rehq DE, McClaren J, et al. Oseltamivir prophylactic regimens prevent H5N1 influenza morbidity and mortality in a ferret model. J Infect Dis. 2008;197(9):1315-1323. 5. Buchman CA, Swarts JD, Seroky JT, et al. Otologic and systemic manifestations of experimental influenza A virus infection in the ferret. Otolaryngol Head Neck Surg. 1995;112:572-578. 6. Caswell JL, Williams KJ. Respiratory system. In: Grant Maxie M, ed. Jubb, Kennedy, and Palmer’s pathology of domestic animals. 2nd ed. Edinburgh, UK: Saunders Elsevier; 2007:523-653. 7. Chen KS, Bharaj SS, King EC. Induction and relief of nasal congestion in ferrets infected with influenza virus. Int J Exp Pathol. 1995;76:55-64. 8. Chin NK, Hui KP, Sinniah R, et al. Idiopathic lipoid pneumonia in an adult treated with prednisolone. Chest. 1994;105:956-957. 9. Collie MH, Rushton DI, Sweet C, et al. Studies of influenza virus infection in newborn ferrets. J Med Microbiol. 1980;13:561-571. 10. de Boer GF, Back W, Osterhaus AD. An ELISA for detection of antibodies against influenza A nucleoprotein in humans and various animal species. Arch Virol. 1990;115:47-61. 11. Dei-Cas E, Brun-Pascaud M, Bille-Hansen H, et al. Animal models of pneumocystosis. FEMS Immunol Med Microbiol. 1998;22:163-168. 12. de Lisle GW, Kawakami RP, Yates GF, et al. Isolation of Mycobacterium bovis and other mycobacterial species from ferrets and stoats. Vet Microbiol. 2008;132(3-4):402-407. 13. De Souza HJM, dos Santos AE, Ferreira AMR, et al. Chronic lipidic pneumonia in a cat. Feline Pract. 1998;26:16-19. 14. Doggart L. Viral disease of pet ferrets. Part II. Aleutian disease, influenza, and rabies. Vet Tech. 1988;9:384-389. 15. Dubovi EJ, Bradley LN. Canine influenza. Vet Clin North Am Small Anim Pract. 2008;38(4):827-835. 16. DuVal-Hudelson KA. Coccidioidomycosis in three European ferrets. J Zoo Wildl Med. 1990;21:353-357. 17. Eyers S, Weatherall M, Shirtcliffe P, et al. The effect on mortality of antipyretics in the treatment of influenza infection: systematic review and meta-analysis. J R Soc Med. 2010;103(10): 403-411. 18. Fenton RJ, Clark A, Potter CW. Immunity to influenza in ferrets. XIV: comparative immunity following infection or immunization with live or inactivated vaccine. Br J Exp Pathol. 1981; 62:297-307. 19. Fenton RJ, Morley PJ, Owens IJ, et al. Chemoprophylaxis of influenza A virus infections, with single doses of zanamivir, demonstrates that zanamivir is cleared slowly from the respiratory tract. Antimicrob Agents Chemother. 1999;43:2642-2647. 20. Fox JG. Bacterial and mycoplasmal diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Philadelphia: Lippincott Williams & Wilkins; 1998:321-354. 21. Fox JG, Pearson RC, Gorham JR. Viral diseases. In: Fox JG, ed. Biology and diseases of ferrets. 2nd ed. Philadelphia: Lippincott Williams & Wilkins; 1998:355-374. 22. Garner MM, Ramsell K, Shoemaker NJ, et al. Myofasciitis in the domestic ferret. Vet Pathol. 2007;44:25-38. 23. Garner MM, Ramsell K, Morera N, et al. Clinicopathologic features of a systemic coronavirus-associated disease resembling feline infectious peritonitis in the domestic ferret. Vet Pathol. 2008;45(2):236-246.
24. Greenacre CB. Incidence of adverse events in ferrets vaccinated with distemper or rabies vaccine: 143 cases (1995-2001). J Am Vet Med Assoc. 2003;223(5):663-665. 25. Husseini RH, Collie MH, Rushton DI, et al. The role of naturally-acquired bacterial infection in influenza-related death in neonatal ferrets. Br J Exp Pathol. 1983;64:559-569. 26. Husseini RH, Sweet C, Collie MH, et al. Elevation of nasal viral levels by suppression of fever in ferrets infected with influenza viruses of differing virulence. J Infect Dis. 1982;145:520-524. 27. Husseini RH, Sweet C, Overton H, et al. Role of maternal immunity in the protection of newborn ferrets against infection with a virulent influenza virus. Immunology. 1984;52:389-394. 28. Ilyushina NA, Hoffman E, Saloman R, et al. Amantadineoseltamivir combination therapy for H5N1 influenza virus infection in mice. Antivir Ther. 2007;12(3):363-370. 29. Johnson-Delaney CA, Reavill DR. Ferret acute hemorrhagic syndrome, In Proceedings. Assoc Avian Vets and Assoc of Exot Mam Vets, 2008; 49–50. 30. Jóżwik A, Frymus T. Comparison of the immunofluorescence assay with RT-PCR and nested PCR in the diagnosis of canine distemper. Vet Res Commun. 2005;29(4):347-359. 31. Kang ES, Lee HJ, Boulet J, et al. Potential for hepatic and renal dysfunction during influenza B infection, convalescence, and after induction of secondary viremia. J Exp Pathol. 1992;6:133-144. 32. Lunn JA, Martin P, Zaki S, Malik R. Pneumonia due to Mycobacterium abscesses in two domestic ferrets (Mustela putorius furo). Aust Vet J. 2005;83(9):542-546. 33. Mahmood K, Bright RA, Mytle N, et al. H5N1 VLP vaccine induced protection in ferrets against lethal challenge with highly pathogenic H5N1 influenza viruses. Vaccine. 2008;26(42):5393-5399. 34. Malik R, Alderton B, Finlaison D, et al. Cryptococcosis in ferrets: a diverse spectrum of clinical disease. Aust Vet J. 2002;80(12):749-755. 35. Malik R, Martin P, McGill J, et al. Successful treatment of invasive nasal cryptococcosis in a ferret. Aust Vet J. 2000;78(3): 158-159. 36. Marini RP, Adkins JA, Fox JG. Proven or potential zoonotic diseases of ferrets. J Am Vet Med Assoc. 1989;195:990-994. 37. Martinez J, Ramis AJ, Reinacher M, et al. Detection of feline infectious peritonitis virus-like antigen in ferrets. Vet Rec. 2006;158:523. 38. McLaren C, Butchko GM. Regional T- and B-cell responses in influenza-infected ferrets. Infect Immunol. 1978;22:189-194. 39. Midulla F, Strappini PM, Ascoli V, et al. Bronchoalveolar lavage cell analysis in a child with chronic lipid pneumonia. Eur Respir J. 1998;11:239-242. 40. Moore GE, Glickman NW, Ward MP, et al. Incidence of and risk factors for adverse events associated with distemper and rabies vaccine administration in ferrets. J Am Vet Med Assoc. 2005;226(6):909-912. 41. Ochi A, Danesh A, Seneviratne C, et al. Cloning, expression and immunoassay detection of ferret IFN-gamma. Dev Comp Immunol. 2008;32(8):890-897. 42. Patterson AR, Cooper VL, Yoon KJ, et al. Naturally occurring influenza infection in a ferret (Mustela putorius furo) colony. J Vet Diag Invest. 2009;21(4):527-530. 43. Pearson RC, Gorham JR. Viral disease models. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Philadelphia: Lippincott Williams & Wilkins; 1998:487-498. 44. Perpinan D, Ramis A, Tomas A, et al. Outbreak of canine distemper in domestic ferrets (Mustela putorius furo). Vet Rec. 2008;163(8):246-250. 45. Perpinan D, Lopez C. Clinical aspects of systemic granulomatous inflammatory syndrome in ferrets (Mustela putorius furo). Vet Rec. 2008;162:180-184.
CHAPTER 6 Respiratory Diseases 46. Peltola VT, Boyd KL, McAuley JL, et al. Bacterial sinusitis and otitis media following influenza virus infection in ferrets. Infect Immunology. 2006;74(5):2562-2567. 47. Prince GA, Porter DD. The pathogenesis of respiratory syncytial virus infection in infant ferrets. Am J Pathol. 1976;82:339-352. 48. Rarey KE, DeLacure MA, Sandridge SA, et al. Effect of upper respiratory infection on hearing in the ferret model. Am J Otolaryngol. 1987;8:161-170. 49. Ryland LM, Gorham JR. The ferret and its diseases. J Am Vet Med Assoc. 1978;173:1154-1158. 50. Saunders GK, Thomsen BV. Lymphoma and Mycobacterium avium infection in a ferret (Mustela putoris furo). J Vet Diagn Invest. 2006;18(5):513-515. 51. Shin YJ, Cho KO, Cho HS, et al. Comparison of one-step RTPCR and a nested PCR for the detection of canine distemper virus in clinical samples. Aust Vet J. 2004;82(1-2):83-86. 52. Sy CL, Lee SS-J, Liu M-T, et al. Rapid emergence of oseltamivir resistance [letter]. Emerg Infect Dis. Apr 2010;16(4). http:// www.cdc.gov/EID/content/16/4/723.htm. Accessed 1/12/2011. 53. Smith H, Sweet C. Lessons for human influenza from pathogenicity studies with ferrets. Rev Infect Dis. 1988;10:56-75. 54. Smith W, Stuart-Harris CH. Influenza infection of man from the ferret. Lancet. 1936;228:121-123. 55. Spickler AR. Influenza. Technical Fact Sheet. The Center for Food Security and Public Health, Iowa State University. Available at http://www.cfsph.iastate.edu/DiseaseInfo/disease. php?name=influenza&lang=en. Accessed 1/12/2011. 56. Squires S, Belyavin G. Free contact infection in ferret groups. J Antimicrob Chemother. 1975;1:35-42.
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57. Swango LJ. Canine viral diseases. In: Ettinger SJ, ed. Textbook of veterinary internal medicine. Philadelphia: WB Saunders; 1989:298-311. 58. Swenson SL, Koster LG, Jenkins-Moore M, et al. Natural cases of 2009 pandemic H1N1 influenza A virus in pet ferrets. J Vet Diagn Invest. 2010;22(5):784-788. 59. Taboada J. Systemic mycoses. In: Ettinger SJ, Feldman EC, eds. Textbook of veterinary internal medicine. 5th ed. Philadelphia: WB Saunders; 2000:453-476. 60. Une Y, Wakimoto Y, Nakano Y, et al. Spontaneous Aleutian disease in a ferret. J Vet Med Sci. 2000;62(5):553-555. 61. Valheim M, Djonne B, Heiene R, Caugant DA. Disseminated Mycobacterium celatum (Type 3) infection in a domestic ferret (Mustela putorius furo). Vet Pathol. 2001;38:460-463. 62. Van Riel D, Munster VJ, McAuley AL, et al. Human and avian influenza viruses target different cells in the lower respiratory tract of humans and other mammals. Am J Pathol. 2007;171(4):1215-1223. 63. Yoshimoto J, Yagi S, Ono J, et al. Development of anti-influenza drugs: II. Improvement of oral and intranasal absorption and the anti-influenza activity of Stachyflin derivatives. J Pharm Pharmacol. 2000;52:1247-1255. 64. Welter J, Taylor J, Tartaglia J, et al. Vaccination against Canine Distemper Virus Infection in Infant Ferrets with and without Maternal Antibody Protection, Using Recombinant Attenuated Poxvirus Vaccines. J Virol. 2000;74(14):6358-6367.
CHAPTER
7
Endocrine Diseases
Karen L. Rosenthal, DVM, MS, and Nicole R. Wyre, DVM, Diplomate ABVP (Avian)
Adrenal Gland Disease History and Physical Examination Clinical Pathology and Diagnostic Testing Possible Concurrent Abnormalities Management Adrenal Histopathology Prognosis Pheochromocytomas Thyroid Disease Diabetes Mellitus History and Physical Examination Clinical Pathologic and Diagnostic Testing Treatment Prognosis Pancreatic Islet Cell Tumors Etiology Pathophysiology Clinical Features Clinical Pathologic Abnormalities Differential Diagnoses for Fasting Hypoglycemia Diagnostic Approach Diagnostic Imaging Management of Islet Cell Tumors Histopathology Prognosis
ADRENAL GLAND DISEASE Adrenocortical disease has been recognized for almost 25 years as a common malady affecting pet ferrets both in the United States and in many countries around the world.11,20,25,35,39,53, 54,58,68,72,91,93,109 It is typically seen in middle-aged to older ferrets and is most commonly characterized by hair loss in both sexes and by vulvar enlargement in females.35,58,91 Clinical signs of adrenocortical disease in ferrets differ from those of classic Cushing’s disease in dogs; moreover, plasma cortisol 86
concentrations are rarely increased in ferrets. Instead, the concentrations of estradiol, 17-hydroxyprogesterone, or one or more of the plasma androgens may be increased as a result of adrenocortical hyperplasia, adenoma, or adenocarcinoma.93 There has been much speculation regarding the underlying cause of the pathologic changes in the adrenal glands of these ferrets. Suggested causes include early neutering, genetic predisposition, light-dark cycle disruptions, and diet.12,13,42,46,85,97,98 The role of early neutering has garnered the most attention and study. Historically, gonadectomy at an early age in some strains of mice was observed to lead to adrenocortical nodular hyperplasia or neoplasia of one or both adrenal glands. These glands hypersecrete estrogens or androgens.30,70,99 Like mice that undergo gonadectomy early in life, most commercially raised ferrets in the United States undergo ovariohysterectomy or castration before they are 6 weeks of age. Another possible explanation is not necessarily the age of neutering but the time period between neutering and the onset of disease.97 To this end, gonadotrophic hormones appear to play a role in the pathogenesis of hyperadrenocorticism in ferrets.98 Specifically, this condition has been defined as a disease resulting from the expression of luteinizing hormone (LH) receptors on adrenocortical cells that produce sex steroids. The speculative pathogenesis is that, after neutering, LH and follicle stimulating hormone (FSH) persistently stimulate the adrenal cortices as a result of the loss of negative gonadal feedback on hypothalamic gonadotropin releasing hormone (GnRH), resulting in adrenocortical hyperplasia. To further support this hypothesis, LH receptors that could trigger abnormal adrenal gland growth have been found in the adrenal glands of normal ferrets.97 In a small study of adult ferrets, there was a high incidence of adrenal gland disease during a 1- to 7-year period after surgery.42 These results may be further evidence that adrenal gland disease is influenced more by lack of negative feedback than the practice of early neutering.12 As the ability to study disease at the molecular level becomes more robust, the underlying pathogenicity of adrenal gland disease in ferrets will become more evident.77 Other possible causes of adrenal gland disease in ferrets may be related to husbandry. One speculated risk factor for adrenal Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 7 Endocrine Diseases gland disease is the artificial light:dark cycle to which indoor ferrets are exposed.46,69,85 The long light cycle, as would be present with ferrets living indoors, stimulates the release of GnRH and LH while simultaneously inhibiting the production of melatonin. The increased LH concentrations combined with decreased circulating melatonin (a known antigonadotropic hormone) stimulate receptors in adrenal gland tissue, leading to adrenal gland disease.85 Therefore housing ferrets indoors may be a contributing risk factor to the pathogenesis of adrenal gland disease.69,85
HISTORY AND PHYSICAL EXAMINATION Progressive alopecia is the most common historic finding. Hair loss typically begins in the late winter or early spring and may continue until the ferret is partially or completely bald. Occasionally the hair coat may regrow fully during the fall. During the next winter or spring, alopecia commonly begins again. This sequence can recur over a period of 2 to 3 years until the hair does not regrow. Spayed female ferrets with adrenocortical disease frequently have a history of vulvar enlargement with or without a mucoid discharge. Male ferrets may have a history of dysuria, urinary blockage, or increased aggressive behavior. In at least one study, most ferrets reported with adrenocortical disease were female.91 Many owners are aware that an enlarged vulva in a female ferret is a cause for concern. The vulva enlarges normally during estrus, and many owners know that prolonged estrus can result in estrogen-induced bone marrow toxicosis. Thus although the disease is reported more frequently in females than in males, this may be caused by a presentation bias rather than an actual sex predilection and may not reflect the true incidence of disease.11 In one study, the average age at which signs of adrenal disease were first observed by ferret owners was approximately 3.5 years.91 A smaller study has reported an older age distribution for this disease.72 Alopecia is the most common clinical manifestation of adrenocortical disease and develops in both male and female ferrets (Fig. 7-1). More than 90% of ferrets with adrenal gland disease have some hair loss. The hair epilates easily. Alopecia is usually symmetrical, beginning on the rump, the tail, or the flanks and
Fig. 7-1 A 4-year-old female spayed ferret with extensive alopecia as a result of adrenal gland disease. Hair was first noticed to be thinning one year earlier.
87
progressing to the lateral trunk, dorsum, and ventrum. These patterns of alopecia should be documented in the medical record for comparison as the disease progresses. In more than one-third of ferrets with adrenal gland disease, owners report the ferret as pruritic. Although pruritus usually accompanies hair loss, in some ferrets pruritus is the only clinical sign. Pruritus is most frequently observed on the dorsum between the shoulder blades. The skin is often erythematous in these areas. More than 70% of female ferrets with adrenal gland disease have an enlarged vulva (Fig. 7-2).91 The vulva can be slightly to grossly enlarged and become turgid and edematous, resembling the vulva of a jill in estrus. A seromucoid discharge may be present, and results of cytologic examination may show localized vaginitis. The perivulvar skin may appear dark and bruised. Partial or complete urinary blockage in male ferrets is occasionally associated with adrenocortical disease.20,88 Affected ferrets have dysuria or stranguria. Periurethral cysts develop in the region of the prostate (possibly originating from hormoneresponsive cells) and cause urethral narrowing. Because of the urethral narrowing, passing a urinary catheter into the bladder of these ferrets can be difficult (see Chapter 2). A ferret with a urethral blockage may have a life-threatening metabolic derangement; therefore these cases usually constitute emergencies. In some ferrets, removing the diseased adrenal tissue and draining the cysts resolves the urinary blockage within 1 or 2 days. In others, the prostatic tissue is infected and aggressive surgical treatment along with antibiotic and hormonal therapy is necessary (see Chapter 11). On physical examination, enlarged adrenal glands are sometimes palpable (Fig. 7-3). The left adrenal gland is more easily identified than the right. The left gland is usually engulfed in a large fat pad cranial to the left kidney. It may feel like a small, firm, round mass. Because the right gland has a more cranial location and is under a lobe of the liver, it is more difficult to palpate. Enlarged mesenteric lymph nodes may be palpable. The spleen may be palpably enlarged, but it usually has smooth borders and is not painful. In some instances, the texture of the spleen is irregular and knobby. In most older ferrets, an enlarged spleen is an incidental finding on physical examination.
Fig. 7-2 An enlarged vulva in a female ferret with adrenal gland disease.
88
SECTION I Ferrets Abdominal ultrasound is also useful for detecting concurrent diseases, such as renal or hepatic disease, metastasis from a pancreatic insulinoma, or enlarged lymph nodes. The presence of prostatic enlargement or uterine disease associated with adrenal gland disease can also be found by ultrasonography.11 Although not done routinely, advanced imaging studies such as computed tomography (CT) or magnetic resonance imaging (MRI) of a ferret’s abdomen can demonstrate adrenal gland abnormalities. In ferrets with right adrenal gland tumors, CT or MRI may be useful as a presurgical screening tool in determining involvement or invasion of the vena cava.9
Ancillary Diagnostic Tests
Fig. 7-3 A female spayed ferret with confirmed adrenal gland disease. The left adrenal gland was palpable on physical examination and was prominent when the ferret was in dorsal recumbency.
CLINICAL PATHOLOGY AND DIAGNOSTIC TESTING The presumptive diagnosis of adrenal gland disease is based on history, clinical signs, and results of imaging techniques and steroid hormonal assays. Diagnosis is confirmed by histologic examination of adrenal tissue obtained during surgical biopsy or adrenalectomy. Results of the complete blood count (CBC) are usually unremarkable. Rarely adrenocortical disease is associated with nonregenerative anemia. If disease is severe or prolonged, pancytopenia may rarely be present. These changes mimic those in ferrets with estrogen-induced bone marrow toxicosis (see Chapter 4). In ferrets with anemia and pancytopenia, a packed cell volume less than 15% carries a grave prognosis. Results of the biochemical profile are also usually within reference intervals. The alanine aminotransferase concentration is occasionally high, but the association of this finding with adrenal gland disease is unknown. Because insulinomas are also common in older ferrets, hypoglycemia from a pancreatic beta cell tumor (insulinoma) may be present. A urinalysis is not helpful in diagnosing adrenal gland disease. Radiographs are generally not helpful in diagnosing this disease. An enlarged adrenal gland rarely displaces other organs or calcifies; therefore organ displacement and mineralized glands are not visible radiographically. Lung metastasis of adrenal gland tumors is very rare. However, radiographs are useful as a screening tool for other conditions, such as heart disease or splenomegaly. Abdominal ultrasound is useful for detecting abnormal adrenal glands. The size, side of enlargement, and architecture can often be determined.11,53,54,71,74 In one study in normal ferrets, the mean dimensions (length and width) of the right adrenal gland were 7.6 + 1.8 by 2.6 + 0.4 mm, and those of the left adrenal gland were 7.2 + 1.8 by 2.8 + 0.5 mm.74 In another study, the mean dimensions of 28 abnormal left adrenal glands (length and thickness) were 9.2 ± 3.2 by 6.3 ± 3.0 mm, and dimensions of 19 abnormal right adrenal glands were 8.5 ± 2.5 by 5.2 ± 3.3 mm.53 In the same study, abnormal adrenal glands appeared on ultrasound as rounded with an enlarged cranial or caudal pole or both, a heterogeneous structure, and increased echogenicity; they were either with or without mineralization.53
Several diagnostic tests used in dogs with adrenocortical disease are not useful in ferrets. The adrenocorticotropic hormone (ACTH) stimulation test and the dexamethasone suppression test cannot be used to diagnose adrenal disease in ferrets. Both normal ferrets and those with adrenal disease respond equally well to an ACTH stimulation test, probably because most ferrets with this disease do not produce abnormally high concentrations of cortisol.90 Plasma concentrations of ACTH and alpha-melanocyte-stimulating hormone in ferrets with adrenal disease are similar to those of normal ferrets, suggesting that adrenal disease in ferrets is independent of ACTH and alphamelanocyte-stimulating hormone.96 Urinary cortisol/creatinine ratios were higher in 12 ferrets with adrenocortical tumors than in 51 clinically normal ferrets.39 In dogs, the urinary cortisol/ creatinine ratio is a sensitive but not a specific indicator of hyperadrenocorticism. Further studies are needed to evaluate urinary cortisol/creatinine ratios in ferrets with diseases other than hyperadrenocorticism. The measurement of serum or plasma concentrations of steroid hormones is a reliable means of diagnosing adrenal disease in ferrets. Hormone panels that measure estradiol, androstenedione, and 17-hydroxyprogesterone in blood samples are commercially available (Clinical Endocrinology Laboratory of the Department of Comparative Medicine at the University of Tennessee [http://www.vet.utk.edu/diagnostic/endocrinology/ index]). In a normal neutered ferret, these steroids are found in minute quantities, whereas in ferrets with adrenal disease, the serum concentrations of one or more of these compounds may be high.89 In Table 7-1, reference intervals are given for androstenedione, dehydroepiandrosterone sulfate, estradiol, and 17-hydroxyprogesterone in intact female and neutered ferrets.
Differential Diagnosis A ferret with an ovarian remnant or an intact female ferret may present with an enlarged vulva and alopecia, resembling a ferret with adrenal disease. However, ferrets with ovarian remnants are uncommon and tend to exhibit an enlarged vulva at an earlier age (first onset of estrus at one year of age) compared with the usual age of onset of adrenal disease (3-4 years of age). Several diagnostic methods can be used to differentiate the two conditions. In one method, human chorionic gonadotropin (HCG) 100 IU IM is administered and then repeated in 7 to 10 days. If the ferret is intact or if an ovarian remnant is present, the vulva usually decreases in size. Alternatively, imaging methods such as abdominal ultrasound or measuring steroid hormone concentrations may help differentiate the two conditions. If the concentrations of androgens—such as androstenedione, dehydroepiandrosterone sulfate, or 17-hydroxyprogesterone— are high, an adrenal tumor is likely. If only the estradiol
CHAPTER 7 Endocrine Diseases
89
Table 7-1 Serum Concentrations of Steroid Hormones in Intact and Neutered Normal Ferrets and Ferrets with Adrenal Disease NEUTERED FERRETSa
INTACT FEMALE FITCH FERRETS (N = 11)c
NORMAL FERRETS (N = 26) (13 MALE, 13 FEMALE) Steroid Hormone Androstenedione (nmol/L) Dehydroepiandrosterone sulfate (mmol/L) Estradiol (pmol/L) 17-Hydroxyprogesterone (nmol/L)
Adrenal Disease Mean 67 (n = 25) 0.03 (n = 27) 167 (n = 28) 3.2 (n = 20)
Mean 6.6 0.01 106 0.4
Reference Intervalb
Mean
<0.1–15
58.3
30–108 <0.1–0.8
165.5 7.7
Reference Interval 20–96 122–210 2.3–13.1
aData
from Rosenthal K and Peterson M.89 ranges currently in use by the Clinical Endocrinology Laboratory, College of Veterinary Medicine, University of Tennessee, Knoxville, TN. Courtesy Jack W. Oliver, DVM, PhD. cRamer J and Oliver JW, unpublished data. bReference
concentration is elevated, then either condition may be present. An ultrasound examination of the abdomen of a female ferret may differentiate between an adrenal tumor and an intact genital tract. Surgery is the definitive method to confirm the presence of ovarian tissue or an abnormal adrenal gland. Some ferrets, particularly males, exhibit seasonal alopecia of the tail. After several weeks, the hair typically regrows. This condition does not appear to be related to an adrenal tumor.
POSSIBLE CONCURRENT ABNORMALITIES Pancreatic beta-cell tumors and adrenal gland disease are both seen in older ferrets and often occur concurrently. However, it is not known whether a correlation exists between the two diseases. Medications (prednisone and diazoxide) used to control the clinical signs of insulinoma do not appear to interfere with therapy of adrenal disease. If adrenalectomy is elected, pancreatic beta-cell tumors can be debulked simultaneously. Many ferrets with adrenal disease have splenic enlargement. Histopathologic examination of the spleen usually shows extramedullary hematopoiesis. Infrequently, neoplasia, including lymphoma and hemangiosarcoma, causes splenic enlargement. Nodular hyperplasia is also seen. In any ferret with an enlarged spleen that undergoes adrenalectomy, obtain a biopsy specimen of the spleen at the time of surgery (see Chapter 5). Middle-aged and geriatric ferrets frequently have heart disease, which can be clinical or subclinical (see Chapter 5). Before surgery, all older ferrets should be evaluated for subclinical heart disease to prevent the decompensation that can accompany the effects of anesthesia or fluid administration. Lymphoma is also common in ferrets. Abnormal nodes can be found incidentally during a diagnostic workup or abdominal exploratory surgery. At surgery, obtain biopsy samples of any abnormal subcutaneous or mesenteric lymph nodes. There is one case report of concurrent primary hyperaldosteronism in a ferret with classic signs of adrenal gland disease. Persistent hypokalemia, high blood pressure measurements, and a high concentration of circulating aldosterone led to the tentative diagnosis of hyperaldosteronism, with a definitive diagnosis at necropsy.24
MANAGEMENT The two treatment modalities for adrenocortical tumors are surgically removing all or part of an affected adrenal gland(s) or medical management. Surgical removal is the preferred treatment for most ferrets, as it is for dogs with adrenocortical tumors.52,101 With medical management, the growth of the adrenal tumor may not stop but clinical signs may regress either temporarily or permanently. However, medical management may be preferred for economic reasons or in ferrets that are geriatric or have concurrent disease.
Surgical Therapy Preoperative testing of a ferret with an adrenal tumor can include a CBC, biochemical profile, abdominal and thoracic radiographs, and abdominal and cardiac ultrasound examination. Withhold food for 4 to 6 hours before surgery. If a ferret is suspected of having an insulinoma, place an intravenous catheter during most or all of this fasting period to provide intravenous fluid support with added dextrose. Otherwise, place an intravenous catheter at induction. Adrenalectomy techniques are described in Chapter 11. At surgery, fully explore the abdomen before addressing the diseased adrenal gland(s). Other structures to be examined are the liver, the lymph nodes, the pancreas, the kidneys, and the spleen. Because bilateral adrenal gland disease can be present, both adrenal glands must be examined, palpated, and compared. Perform a unilateral adrenalectomy if only one adrenal gland is diseased.101,110,112 If both glands are diseased, a subtotal adrenalectomy, with total removal of one gland and partial removal of the other, is indicated (see Chapter 11). Total bilateral adrenalectomy carries a high risk that drug therapy may be necessary for an unspecified time to replace corticosteroids and mineralocorticoids. In a retrospective study, slightly less than 10% of the ferrets undergoing surgery had bilateral adrenalectomy, but survival rates for that particular group were not reported.101 After surgery, administer maintenance and replacement fluids as needed. Unless concurrent gastrointestinal surgery was performed or complications develop, feed the ferret soon after surgery. Ferrets rarely need postoperative corticosteroid
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SECTION I Ferrets
replacement therapy after simple unilateral adrenalectomy. If the ferret has a concurrent insulinoma or has had a subtotal adrenalectomy and appears very lethargic after surgery, giving prednisone at a starting dose of 0.1 mg/kg q12h for 2 to 3 days after surgery for glucocorticoid replacement may be considered. Monitor the blood glucose levels to determine the effectiveness of or need for medication. If a total or subtotal bilateral adrenalectomy has been performed, monitor electrolyte concentrations closely after surgery. Not all ferrets undergoing these procedures will need supplementation; therefore monitoring electrolyte levels, following trends in blood glucose levels, and performing serial physical examinations is required after surgery. If necessary, mineralocorticoid supplementation may be given as needed. Parenteral administration of mineralocorticoids has been reported with desoxycorticosterone pivalate (Percortin, Novartis, Greensboro, NC) at 2 mg/kg IM q21d, or oral supplementation can be used with fludrocortisone acetate (Florinef, Bristol-Myers Squibb, Princeton, NJ) at a dosage of 0.05 to 0.1 mg/kg PO q24h or divided q12h.48
Medical Management At present, medical management of adrenal gland disease is aimed at moderating the clinical signs rather than curing the disease. Medical management may be considered if the owner cannot afford surgery, if the ferret is a poor surgical candidate or is geriatric, if it has bilateral adrenal tumors that cannot be totally resected, or if it has recurrent disease in the remaining gland after previous unilateral adrenalectomy. Several medical strategies are used to decrease the clinical signs of adrenal disease. Drug therapy includes mitotane, androgen receptor blockers, aromatase inhibitors, melatonin, and GnRH analogs. However, medical therapy is not always successful, and it is impossible to predict which ferret will respond to a particular medication. Numerous attempts with different classes of medications may be needed to find a drug that is effective. Mitotane. Mitotane, or o,p-DDD (Lysodren, Bristol-Myers Squibb Oncology, Princeton, NJ), is effective in dogs for treating pituitary-dependent hyperadrenocorticism. However, this form of hyperadrenocorticism has not been recognized in ferrets, which may explain why treatment with mitotane has rarely been successful in ferrets. Additionally, the use of this drug can result in severe, life-threatening hypoglycemia in ferrets. With the development of newer chemotherapeutic strategies to treat adrenal gland disease in ferrets, mitotane treatment is not recommended. Ketoconazole. Because of its ability to inhibit the steroid biosynthetic pathway at several steps, ketoconazole is used in the treatment of adrenocortical disease in other species.95 However, it is not effective in ferrets. Androgen Receptor Blockers. Androgen receptor blockers reverse the clinical signs of adrenal gland disease but do not inhibit the growth of an abnormal adrenal gland. Theoretically these compounds act at the receptor site to block the actions of androgens. In human medicine, these drugs are used to treat men with benign prostatic hyperplasia or prostatic carcinoma.1,7,22,64 Like all medical therapies used in ferrets with adrenal disease, androgen receptor blockers are effective in some but not all ferrets. Flutamide (Eulexin, Schering Corporation, Kenilworth, NJ) and bicalutamide (Casodex, AstraZeneca Pharmaceuticals LP, Wilmington, DE) have been used in ferrets with adrenal disease.48 Flutamide has been used in ferrets for behavioral research.108
Aromatase Inhibitors. Another class of drugs that can depress the signs of adrenal disease in ferrets comprises the aromatase inhibitors. One drug that has been used is anastrozole (Arimidex, AstraZeneca Pharmaceuticals LP, Wilmington, DE). This drug specifically inhibits aromatase, the enzyme that catalyzes the final step in estrogen production. In people treated with continued dosing of anastrozole, plasma concentrations of estradiol decrease approximately 80% from baseline.38,80 Anastrozole has little or no effect on central nervous system, autonomic, or neuromuscular function. Melatonin. In the mink industry, melatonin implants are used to force early molt to produce optimal winter pelt development for the fur industry; they have also been used in studies of seasonal pelage cycles.49,85 In one study, oral administration of melatonin in ferrets did not stop the progression of adrenal gland disease and/or inhibit the continued growth of the affected adrenal glands. However, melatonin treatment improved hair growth and resulted in reduced prostatic or vulvar size in ferrets with adrenal gland disease for at least 8 months.85 Oral administration of melatonin may not be practical in pet ferrets, but a long term implantable form of melatonin (Ferretonin, Melatek, Middleton, WI) is available. According to the manufacturer, Ferretonin at either a 5.4 mg or 2.7 mg depot of melatonin gives four months of relief to signs of adrenal gland disease in ferrets. Gonadotropin-Releasing Hormone Analogs. GnRH analogs are a class of compounds that are widely used in human medicine to treat prostatic cancer, endometriosis, and breast cancer.49 In ferrets, these compounds have been used to control the signs of adrenal gland disease. There are two general types of analogs: GnRH agonists and GnRH antagonists. During short-term or intermittent therapy, GnRH agonists have the same stimulatory action as GnRH, but long-term therapy suppresses the production and/or release of LH and FSH by downregulating the receptors at the pituitary.105 This downregulation of receptors is the mode of action of the agonists. Two currently used injectable GnRH agonists used in ferrets with adrenal gland disease are leuprolide acetate (Lupron Depot, TAP Pharmaceuticals Inc., Lake Forest, IL) and deslorelin acetate (Suprelorin, Peptech Animal Health Pty Limited [Virbac], NSW, Australia).49,105 Leuprolide acetate (Lupron) was the first GnRH agonist to be widely used in the treatment of ferret adrenal gland disease.49,106 Currently, two types of depot formulations of this agent are commercially available: a 1- or 4-month formulation.48,49 The 1-month formulation appears to be more consistently effective.48 The reported dose range is 100 to 250 mcg/kg or 100 to 200 mcg per ferret IM. The response to this injection may last from 1 to 4 months.48,106 Deslorelin is similar in action to leuprolide but appears to provide a longer disease-free interval than leuprolide.105,107 In one report, the response to a single 4.7-mg implant of deslorelin acetate lasted between 8 and 30 months.105 It is likely that once deslorelin is approved for use and distribution in the United States, it will become the drug of choice for treating ferret adrenal gland disease. In humans, GnRH antagonists are being studied for treatment of many of the same diseases in which agonists are currently used. Theoretically antagonists should be more effective than agonists but at a much lower dose. In human medicine, an antagonist called abarelix is being evaluated to determine its ability to inhibit the action of GnRH. This or similar drugs may eventually replace GnRH agonists in the treatment of ferret adrenal disease.
CHAPTER 7 Endocrine Diseases
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ADRENAL HISTOPATHOLOGY On histopathologic examination, adrenal gland disease is described as adrenocortical hyperplasia, adenoma, or carcinoma.91 One study describes spindle cell-variant adrenal tumors in adrenal gland disease.73 The spindle cell component of these variant adrenal tumors is smooth muscle in origin and may be associated with a more malignant grade of this tumor.73 Another study describes a variant of adrenocortical carcinoma with prominent mucin production, interpreted as myxoid differentiation. In ferrets, adrenocortical carcinoma with myxoid differentiation appears to be more malignant than carcinoma, its well-differentiated counterpart.79 Although adenomas appear to be most common, the clinical behavior of these tumors and hyperplasia is usually identical. Metastasis is uncommon; however, some tumors locally invade the vena cava or liver. Very rarely, carcinomas can metastasize to the lungs. Pheochromocytomas occur uncommonly in ferrets (see below).
Fig. 7-4 Pheochromocytoma in the adrenal gland of a ferret. Neoplastic cells have finely granular, amphophilic cytoplasm with a large nuclei and are arranged in tight packets. Hematoxylin and eosin stain, × 40.
PROGNOSIS The prognosis with surgical treatment is good in the hands of an experienced and skilled surgeon. In one study of 130 ferrets that were treated with surgery for adrenal gland disease, the 1- and 2-year survival rates were 98% and 88% respectively.101 In most ferrets, the associated clinical signs resolve after the diseased adrenal gland has been removed. Owners should be instructed to watch for recurrence of adrenal gland disease, especially if not all of the adrenal gland tissue is removed. Clinical signs of recurrence are alopecia, vulvar enlargement, and pruritus. Later complications of surgical treatment include recurrence of adrenal tumor because of metastasis (rare) or the development of an adrenocortical tumor in the remaining adrenal gland.101 In one study, recurrence of signs after bilateral adrenalectomy developed in 15% of cases, with a mean long-term follow-up period of 30 months.110 Because the results of medical treatment are equivocal, the prognosis with medical treatment is unpredictable. If the effects of the adrenal disease are only cosmetic (alopecia), the prognosis is good. The prognosis worsens if prostatic disease, bone marrow suppression, or tumor-related mechanical interference with the vena cava develops or if the tumor metastasizes.
PHEOCHROMOCYTOMAS Pheochromocytomas occur rarely in ferrets.34,101 Pheochromocytomas arise from the adrenal medulla and produce excessive amounts of catecholamines (Fig. 7-4). Clinical signs are primarily associated with the effects of catecholamines on the cardiovascular system. In clinical cases seen at the Animal Medical Center in New York City, several ferrets exhibited clinical signs consistent with those seen in dogs with pheochromocytomas: tachycardia, dyspnea, and cardiovascular collapse (K. Quesenberry, personal communication, 2002). High blood pressure is a common finding in other species; however, blood pressure was not measured in these ferrets. Because clinical signs of pheochromocytomas are not obvious, retrospective studies are used to determine the incidence of pheochromocytomas in relation to adrenocortical disease. In one study, 5 pheochromocytomas were associated with 131 adrenal gland masses (3.8%).101 In another study, the percentage of pheochromocytomas in adrenal tumors was similar (3%).78 However, the
incidence of pheochromocytomas in ferrets without clinical signs attributed to adrenocortical disease is unclear. Histologic diagnosis of a pheochromocytoma is confirmed by immunohistochemical staining. Animals with pheochromocytomas respond poorly to chemotherapy, and surgical excision is the treatment of choice. Prognosis in ferrets with pheochromocytomas is poor.
THYROID DISEASE Clinical hyperthyroidism and hypothyroidism have not been reported in ferrets. In one report of medullary thyroid carcinoma diagnosed at necropsy, functional hyperthyroidism was not confirmed antemortem.34 Resting values for thyroxin and tri-iodothyronine and results of thyroid-stimulating hormone testing reported in one study of normal intact male ferrets are presented in Table 7-2.43 Suspected pseudohypoparathyroidism was reported in a 1.5-year-old neutered male ferret.114 Pseudohypoparathyroidism, a hereditary condition in humans, results from a lack of response to circulating parathyroid hormone rather than hormone deficiency. The ferret was seen because of intermittent seizures, and results of diagnostic tests revealed low serum calcium, high serum phosphorus, and high serum parathyroid hormone concentrations. The ferret responded to long-term treatment with dihydrotachysterol, a vitamin D analog, and calcium carbonate.
DIABETES MELLITUS HISTORY AND PHYSICAL EXAMINATION Spontaneous diabetes mellitus is very uncommon in ferrets.10,14 Most ferrets develop iatrogenic diabetes from aggressive pancreatectomy to debulk beta-cell tumor nodules. The clinical signs of diabetic ferrets are similar to those of other species. The severity of such signs depends on the severity and chronicity of the disease. Affected ferrets are polyuric and polydipsic and may lose weight despite a good appetite. They often appear lethargic, especially if a metabolic derangement such as ketoacidosis is present.
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SECTION I Ferrets
Table 7-2 Reference Values for Endocrine Tests in Ferrets Hormone Cortisol (nmol/L)a Thyroxine (μg/dL)b Tri-iodothyronine (ng/mL)b Mean thyroxine (μg/dL)c
Sex M F M F M
Values 25.9-235 (mean, 73.8) 1.01-8.29 (mean, 3.24) 0.71-3.43 (mean, 1.87) 0.45-0.78 (mean, 0.58) 0.29-0.73 (mean, 0.53) 2.69 at 0 hours; 3.37 at 2 hours; 3.97 at 4 hours; 3.45 at 6 hours
aAdministration of cosyntropin, 1 μg/kg IM, generally caused a threeto fourfold increase in plasma cortisol concentration. Data from Rosenthal KL, Peterson ME, Quesenberry KE, et al.90 bData from Garibaldi BA, Pequet Goad ME, and Fox JG.37 cMean thyroxine at baseline (0 hours) and 2, 4, and 6 hours after intravenous administration of 1 IU of thyroid-stimulating hormone (n = 8 intact males). Data from Heard DJ, Collins B, Chen DL, et al.43
The findings on physical examination are often unremarkable. Ferrets may be thin and have distended urinary bladders.
CLINICAL PATHOLOGIC AND DIAGNOSTIC TESTING The diagnosis of diabetes mellitus in ferrets is based on some or all of the following: compatible clinical signs (polyuria/polydipsia), a history of recent insulinoma surgery, repeated high blood glucose concentrations, low blood insulin concentration, and normal to high blood glucagon concentration. In diabetic ferrets, a profound hyperglycemia is usually present. Although a blood glucose concentration of 400 mg/dL or higher is suspicious for diabetes, repeated high blood glucose measurements are needed to confirm consistent hyperglycemia. In uncomplicated diabetes, results of the biochemical analysis are usually unremarkable. However, the same metabolic derangements that are present in other mammals with complicated diabetes can occur in ferrets. Results of the CBC are usually unremarkable. However, if a concurrent bladder infection is present, the WBC may be high. A consistent glucosuria is present and, in severe cases, ketones are detected. As with other animals with diabetes, ferrets with diabetes can have an active urine sediment. Although radiography and ultrasound are not useful for diagnosing diabetes mellitus, they are useful to screen for other conditions such as splenomegaly, hepatic enlargement, and cardiac disease.
Ancillary Diagnostic Tests Diabetes mellitus in ferrets can be caused by a lack of insulin, insulin resistance, or a glucagonoma. Insulin concentration is routinely measured in ferrets with insulinomas. This test is preferably performed by a diagnostic laboratory that has validated the test for ferrets. Theoretically, a low insulin concentration concurrent with hyperglycemia can confirm the diagnosis of diabetes mellitus. A normal or high insulin concentration could represent either insulin resistance or the presence of a glucagonoma. Therefore, measuring glucagon concentration can be an important indicator in ferrets with hyperglycemia.
Unfortunately routine measurement of glucagon concentration is difficult because it must be performed in a laboratory that has validated the assay for ferrets. Currently no validated assay is available.
TREATMENT Treatment of diabetes mellitus depends on the severity of hyperglycemia and other metabolic disturbances. Insulin therapy should be instituted in ferrets if the blood glucose concentration is higher than 300 mg/dL on repeated measurements. The treatment principles used for dogs and cats can be followed. Unfortunately success is limited in tightly regulating the blood glucose concentrations of diabetic ferrets. In hospitalized animals, measure serial blood glucose concentrations while administering insulin twice daily. Neutral protamine Hagedorn insulin can be used, starting it at an empiric dose of 0.5 to 1 U of insulin per ferret twice daily. In an effort to give injections only once per day, some ferrets receive Ultralente insulin, which may have a longer duration of action than neutral protamine Hagedorn insulin. Insulin glargine (0.5 U twice daily) has been used successfully in one ferret.43a Increase or decrease the insulin dose as dictated by the blood glucose concentration while monitoring the urine for the presence of glucose and ketones. After the blood glucose concentration is stabilized between 125 and 200 mg/dL, discharge the ferret to its owner with a prescribed insulin regimen. Instruct owners to check for the presence of ketones and glucose in the urine with urine dipsticks; if no glucose is detected in the urine, the the next dose of insulin should not be given. If trace amounts of glucose are found, the insulin dose is not changed. If the amount of glucose in the urine is large, then the insulin dose is increased slightly. Most diabetic ferrets are difficult to regulate. Realistically, the goal is to have negative ketones and a small amount of glucose in the urine.
PROGNOSIS Many ferrets that develop iatrogenic hyperglycemia immediately after insulinoma surgery may be transient diabetics. Their prognosis as it relates to diabetes is good because the hyperglycemia usually normalizes without treatment during the first 1 to 2 weeks after surgery. Occasionally, the diabetes resolves spontaneously after 4 to 6 weeks of treatment. The prognosis is worse or, at best, unpredictable for ferrets with diabetes mellitus that occurs spontaneously or is detected weeks to months after insulinoma surgery. Blood glucose concentration is usually difficult to regulate in these animals.
PANCREATIC ISLET CELL TUMORS Since first reported in 1984,50 pancreatic islet cell tumors have become the most commonly reported neoplasm in ferrets in the United States.57 Affecting mostly middle-aged to older ferrets, the clinical signs associated with this disease are secondary to hypoglycemia resulting from the excess secretion of insulin by these tumors.
ETIOLOGY The islets of Langerhans are the pancreatic endocrine centers.62 These groups of cells are found throughout the pancreas but comprise only about 2% of the total pancreatic
CHAPTER 7 Endocrine Diseases tissue.62 Normal pancreatic islets contain four cell types that each secrete a different peptide: alpha cells secrete glucagon, beta cells secrete insulin, delta cells secrete somatostatin, and P (F) cells secrete pancreatic polypeptide.41 Neuroendocrine tumors of the pancreas are neoplasms arising from the islets of Langerhans and are called islet cell tumors.15 These tumors are further classified by the type of peptide they secrete. Tumors secreting biologically active peptides that result in clinical symptoms are called functional islet cell tumors.15 The most frequently reported functional tumor in small animals and specifically in ferrets is the insulin-secreting beta-cell tumor, or insulinoma.18,27,31,62,84 In one study, 94% of beta-cell tumors in ferrets were reported to be functional.57 In fact, islet cell tumors are reported to be the most common endocrine neoplasm seen in ferrets, with an incidence of 22%57 of all reported neoplasms in a 1998 retrospective study and an incidence of 25% of all reported neoplasms in a 2003 retrospective study.57,113
PATHOPHYSIOLOGY In healthy animals and in humans, the ratio between the blood insulin and glucose concentrations remains constant because of control by the beta cells of the pancreatic islets.31 In normal animals, the insulin secretion rate increases when the blood glucose concentration exceeds 110 mg/dL and is inhibited when the blood glucose concentration decreases below 60 mg/dL.31 When neoplastic beta cells are present, they synthesize and release insulin autonomously despite hypoglycemia.31 This excessive insulin secretion leads to hypoglycemia by suppressing endogenous glucose production by the liver and stimulating glucose utilization in muscle, liver, and adipose tissues. Clinical signs associated with the resultant hypoglycemia are manifested by both neuroglycopenia and stimulation of the sympathoadrenergic system. Clinical signs are related to the rate of decrease of serum glucose, the concentration of serum glucose, and the duration of hypoglycemia.55 Neuroglycopenic signs result from a decreased glucose supply to the brain, which leads to central neurologic signs such as lethargy, ataxia, weakness, bizarre behavior (“staring off into space”), disorientation, collapse, hind limb weakness, seizures, and coma.27,31 Sympathoadrenergic signs result from decreased blood glucose to the hypothalamus, with resultant stimulation of the sympathetic nervous system and release of catecholamines. These are signs associated with the “fight or flight” response such as muscle tremors, nervousness, restlessness, hunger, vocalization, tachycardia and mydriasis. In states of chronic hypoglycemia, as is commonly seen in ferrets with insulinomas, the blood glucose concentration drops slowly and does not activate the sympathetic nervous system. These animals can tolerate very low blood glucose levels (30 to 40 mg/dL) for prolonged periods without exhibiting clinical signs. They become symptomatic when their blood glucose levels decrease slightly, as with exercise or fasting.31 Counterregulatory hormones such as epinephrine, cortisol, glucagon, and growth hormone help to antagonize the effects of insulin during hypoglycemic episodes by increasing the release and formation of glucose while decreasing its use.27 This allows animals to remain normal between hypoglycemic episodes and can prevent owners from bringing their animal to the veterinarian when initial signs are observed.
93
Table 7-3 Incidence of Clinical Signs Associated with Pancreatic Islet Cell Tumors in 161 Ferrets Clinical Sign Lethargy Weakness Ptyalism Pawing at mouth Weight loss Posterior weakness Collapse Seizures Ataxia Vomiting Twitching Abnormal behavior Trembling Disorientation
Ferrets (n)
Percent
125 107 62 40 40 31 30 22 22 11 10 2 2 1
78 66 39 25 25 19 19 14 14 7 6 1 1 <1
Based on cases from references 16, 17, 25, 26, 33, 47, 50, 51, 59, 60, 61, 65, and 111.
CLINICAL FEATURES Signalment Pancreatic islet tumors typically occur in middle-aged and older ferrets. The reported mean age at the time of diagnosis is 5 years, with an age range of 2 to 7 years.17,26,111 This is consistent with the average age of incidence of all neoplastic disease in ferrets, which is 4 to 7 years.57 While these tumors are rarely reported in younger ferrets, there is one report of a functional islet cell tumor in a 2-week-old ferret.57 In three studies, male ferrets were diagnosed slightly more frequently than female ferrets, although this may reflect a sex bias of ferrets presenting to these hospitals rather than a true sex predilection.17,26,111
Clinical Signs Clinical signs of pancreatic islet cell tumors may be observed by the owner as an acute or chronic onset over the course of weeks to many months.84 In one retrospective study of insulinomas in ferrets, the average duration of clinical signs before presentation to a veterinarian was 90 days.26 The clinical signs are caused by neuroglycopenia and an increase in circulating catecholamine concentrations.31 The most common clinical signs in ferrets are lethargy and weakness, followed by ptyalism, pawing at the mouth, and weight loss (Table 7-3). Of interest is that ptyalism, which is reported in 39% of ferrets, and pawing at the mouth, which is reported in 25% of ferrets, is not a reported clinical sign in cats or dogs with insulinomas.27,31 Ptyalism has also been observed in humans treated with overdoses of insulin for shock therapy of mental disorders; it is thought to be related to the stimulation of the autonomic nervous system.4,44 Ferrets with insulinoma may be presented for profound pawing at the mouth and can appear to have something stuck in their mouths. This can be quite dramatic and may result in excoriations or bleeding from the mouth, lips, or perioral region. Numbness and tingling of the lips and tongue is described by humans with insulinomas and is thought to be associated with a peripheral neuropathy associated with hypoglycemia.6,23 These clinical signs have also been suggested to be associated with nausea.3
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SECTION I Ferrets
Weight loss, which is seen in 25% of ferrets, is not seen in dogs with insulinomas. Conversely, dogs with insulinomas are reported to have weight gain.27,31 As reported in canine, feline, and humans patients, clinical signs in ferrets may last several minutes to several hours and usually end with spontaneous recovery after the patient eats or is given a glucose solution.15,27,31,84 As with dogs, most ferrets exhibit multiple clinical signs that are often episodic but tend to become more frequent and severe if left untreated.27,60 One of the most severe clinical signs of hypoglycemia is seizures. While seizures are the most common clinical sign in dogs with insulinomas, affecting 56% to 77% of dogs, they have been reported in only 14% of ferrets (see Table 7-3).27,31 This may be related to differences in the tolerance to hypoglycemia, the rate at which hypoglycemia develops, and typical husbandry of these species. For example, ferrets are usually fed ad libitum as opposed to feedings at time-restricted intervals for dogs, and compared with dogs, ferrets have a decreased activity level due to cage restriction.65 However, generalized seizures secondary to hypoglycemia constitute an emergency and must be treated as such to prevent permanent damage to the cerebral cortex.31
Physical Examination
Fig. 7-5 Hypoglycemic ferret with ptyalism, a common sign seen in ferrets with hypoglycemia secondary to pancreatic islet cell tumors. This clinical sign is not observed in dogs with insulinomas. (Courtesy Dr. Margaret Fordham.)
Physical examination findings in ferrets with pancreatic islet cell tumors vary and are often unremarkable. The most common abnormalities are nonspecific and may include lethargy, generalized weakness, posterior weakness, and weight loss (see Table 7-3). Because most ferrets that present for insulinomas are middle aged or older, signs associated with aging or concurrent disease such as alopecia, skin masses, vulvar enlargement, prostatomegaly, and cardiac arrhythmias or murmurs may be present. A ferret presenting as an emergency may appear disoriented with “glazed” eyes, have ptyalism (Fig. 7-5), and have oral ulcerations from pawing at the mouth. In more severe cases, the ferret may be having a seizure or be postictal, obtunded (Fig. 7-6), or comatose.
CLINICAL PATHOLOGIC ABNORMALITIES The minimum diagnostic evaluation for ferrets suspected of having an insulinoma includes a CBC and serum or plasma biochemical profile. Aside from hypoglycemia, defined as a blood glucose concentration lower than 60 mg/dL, results of the CBC and biochemical profile are usually unremarkable.27 High concentrations of alanine aminotransferase and aspartate aminotransferase, leukocytosis, neutrophilia, and monocytosis have been reported, but these findings are nonspecific and usually not helpful in achieving a definitive diagnosis.17,61
DIFFERENTIAL DIAGNOSES FOR FASTING HYPOGLYCEMIA Although hypoglycemia (blood glucose <60 mg/dL) in a ferret with neuroglycopenic signs suggests the presence of an insulinoma, other causes must also be considered. In animals and humans, hypoglycemia can also result from excessive glucose utilization by neoplastic cells, impaired hepatic gluconeogenesis/ glycogenolysis as seen with severe hepatic disease, a deficiency in glucose counterregulatory hormones as seen with hypoadrenocorticism (potentially with bilateral adrenalectomy), increased tissue utilization as seen with sepsis, increased glucose utilization secondary to polycythemia and artifactual hypoglycemia.31
Fig. 7-6 Obtundation secondary to severe or prolonged hypoglycemia in a ferret. Such patients must be immediately stabilized with intravenous dextrose and fluid therapy and monitored as being in critical condition. (Courtesy Dr. Margaret Fordham.)
Many of these diseases can be ruled in or out with a thorough anamnesis and physical examination coupled with routine blood work or abdominal ultrasound. Artifactual hypoglycemia is commonly encountered if whole blood is stored before being analyzed.21 Decreases in the glucose concentration as high as 10% per hour have been reported.21 Previously, sodium fluoride-treated tubes have been recommended for collecting blood samples to assess glucose concentration.94 More recent information indicates that hemolysis is common when blood is collected in these tubes, resulting in falsely decreased blood glucose values.31 Handheld portable blood glucose monitoring devices measure blood glucose values lower than actual glucose values determined by benchtop devices (i.e., glucose oxidase and hexokinase methods).31 Failure to consider this “error” can result in
CHAPTER 7 Endocrine Diseases
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Table 7-4 Published Reference Ranges for Insulin Values in Ferrets Insulin Value Insulin Value Fasted/Time (μU/mL) (pmol/L) Number (sex) of Fast
Method
6.4-17.6
46-126
4.6-43.3 5.0-35.0 aAnimal
Yes/4 hr
Radioimmunoassay65
33-311
15 (8 male, 7 female) 30
Yes/overnight
36-251
—
—
Immunoreactive insulin radioimmunoassay63,a No longer commercially available84,b
Health Diagnostic Laboratory, Michigan State University, Lansing, MI.
bClinical Endocrinology Laboratory, College of Veterinary Medicine, University of Tennessee, Knoxville, TN.
an incorrect diagnosis of hypoglycemia.31 Additionally, to date, there is no handheld portable glucose monitoring device that has been validated for use with ferret blood. This is particularly important because in humans, handheld devices have been shown to be extremely inaccurate with higher hematocrit levels.102 Because ferrets normally have a higher hematocrit than humans (mean 52% in ferrets vs. mean 44% in humans), the results must be interpreted with caution.45,83 However, blood glucose values obtained by a handheld device can be used in screening for suspect insulinoma cases. If blood glucose values obtained are low, the results must be confirmed by submitting a blood sample for measurement by conventional laboratory methods.
DIAGNOSTIC APPROACH Beyond Whipple’s Triad Whipple’s triad, which is the demonstration of hypoglycemia with relevant signs and response to treatment with glucose, has been used by practitioners in the past in diagnosing insulinsecreting tumors.31 However, this triad is only suggestive of an insulinoma and should not be used as a definitive diagnosis of the disease. There are noninvasive tests to further support a clinical diagnosis of insulinoma, but only histologic diagnosis is definitive.
Determination of Baseline Insulin and Glucose Concentrations The goal of testing blood insulin and glucose concentrations is to establish that the blood insulin concentration is high when hypoglycemia is present. Insulin secretion should be inhibited when the blood glucose concentration drops below 60 mg/dL.31 Therefore the relative excess of insulin is easiest to recognize when the blood glucose concentration is low.31 If the blood glucose level is low and the insulin level is above the normal range, the relative or absolute excess of insulin can be explained by the presence of an insulin-secreting tumor that is insensitive to hypoglycemia. If a ferret with a suspected insulinoma has a resting blood glucose of less than 60 mg/dL, a blood sample can be collected at that time to measure the insulin and blood glucose levels simultaneously.31 If the ferret’s resting blood glucose is above 60 mg/dL, a closely monitored short-term fast can be performed in a hospital setting. During the fast, check the ferret’s blood glucose concentration every 1 to 2 hours until it is less than 60 mg/ dL. At that time, collect a blood sample for measurement of simultaneous blood glucose and insulin levels. It is important
to take care with this technique and, if at any point the ferret develops clinical signs, feed the ferret immediately and monitor it closely. Because insulin values measured by different methods have been shown to be unequal, plasma or serum insulin concentrations should be measured at a laboratory that has validated the assay for ferrets.63,65 Hemolysis of the blood sample renders the measurement of insulin unreliable.40 Currently, two veterinary laboratories that offer insulin assays for ferrets are the Animal Health Diagnostic Center, College of Veterinary Medicine, Cornell University, Ithaca, New York (http://ahdc.vet.cornell.edu/) and the Diagnostic Center for Population and Animal Health, Michigan State University, Lansing, Michigan (www.dcpah.msu.edu/). Reference values are listed in Table 7-4. Normal ferret insulin concentrations are reported to range from about 5 to 40 μU/mL (36-288 pmol/L), depending on the laboratory. In reports of ferrets diagnosed with insulinomas, insulin concentrations have ranged from normal to a high of 2,670 μU/mL (19,157 pmol/L).17,33,47,51,61,65 In a retrospective study, 40 out of 48 ferrets with histologically confirmed insulinomas had high insulin levels, with a range of 7.9 to 372 μU/mL (57-2,670 pmol/L).17 In the same study, 57 of 57 ferrets with confirmed insulinomas had low serum glucose levels with a range of 2 to 60 mg/dL.17 In a study of 6 ferrets with confirmed insulinomas, the mean serum insulin concentration after a 4-hour fast was 58 μU/mL (416 pmol/L), and the mean serum glucose concentration was 44 mg/dL.65 Because insulin is released in pulsatile manner, if a single insulin value is lower than expected in a suspect insulinoma case, the measurement should be repeated after a controlled, observed fast as described above.94 If the blood glucose concentration is less than 60 mg/dL, insulin levels should be low. Therefore even a “normal” insulin concentration with low blood glucose is suggestive of an insulinoma.94
Insulin:Glucose Ratios In the past, various ratios such as the insulin:glucose, glucose:insulin, and amended insulin:glucose ratios were used to diagnose the presence of an insulinoma. These ratios are less specific and give more false-positive results in animals without insulin-secreting tumors and are therefore no longer recommended.32,55
Provocative Testing Several tests have been described that use agents to stimulate insulin secretion by normal and neoplastic beta cells. These include the glucagon tolerance test, the oral and intravenous
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SECTION I Ferrets
glucose tolerance test, the tolbutamide tolerance test, and the epinephrine stimulation test. These tests have never been validated in ferrets and are not recommended for use in dogs suspected of having an insulinoma because of the potential risk of prolonged and severe hypoglycemia.31,32,84
Additional Testing Evaluation of glycosylated protein, such as fructosamine and glycosylated hemoglobin, has been assessed in the diagnosis of insulinomas in dogs.28,82,103 In these proteins the degree of glycosylation depends on the serum glucose concentration, with hypoglycemia causing a decreased glycosylation. Thus levels would be expected to be low in patients with insulinomas. This has been observed in some cases,82 but results in the normal range have also been observed.28 To date there are no reports of measurement of these proteins in ferrets.
DIAGNOSTIC IMAGING Radiography In both dogs and ferrets, abdominal radiographs are not helpful in establishing the diagnosis of insulin-secreting tumors.17,31 In a retrospective study of 28 ferrets, pancreatic masses or metastases were not observed in lateral and ventrodorsal radiographic views in any of the ferrets, although masses were found during surgery or at necropsy.17 However, both thoracic and abdominal radiographs are warranted if a concurrent disease is suspected or as a part of a thorough presurgical workup.
Ultrasonography In general, transabdominal ultrasound has a low sensitivity in detecting islet cell tumors. In dogs and humans, bowel gas, obesity, body conformation, and intestinal content are common causes of poor pancreatic image quality.86 In ferrets, low sensitivity may be because of the small, sometimes microscopic size of the tumors.18,84 When insulinomas are detected by ultrasound, they are seen as solitary nodules, multiple nodules, or an ill-defined area of abnormal echogenicity (Fig. 7-7).75 Benign pancreatic nodular hyperplasia, which appears as welldefined hypoechoic to isoechoic nodules, can be confused with insulinomas.75
PANCREAS
1
2
1 L 1.64 mm 2 L 2.28 mm
Fig. 7-7 Pancreatic islet cell tumor in a ferret. Two hypoechoic nodules are present at the distal portion of the left limb of the pancreas, measuring 1.64 (lesion 1) and 2.3 (lesion 2) mm respectively. Both lesions were surgically removed and identified as well-encapsulated islet cell tumors.
In one report of insulinoma detection by ultrasound in 14 dogs, only 5 (36%) dogs had masses that were correctly identified.86 The median size of the pancreatic masses that were correctly identified was 20 mm, whereas the mean size of unidentified lesions was 15 mm.86 Another report in 44 dogs found that tumors in the left lobe of the pancreas were identified by ultrasound more often than those in the body or the right lobe.31 In a retrospective study of 23 ferrets, pancreatic masses were seen in 5 (22%) on abdominal ultrasound examination. Interestingly, hepatic metastases were detected in 3 of the ferrets.17 Therefore abdominal ultrasound is still warranted from a prognostic standpoint and should be included as part of a thorough presurgical workup to look for concurrent disease.
Advanced Imaging Abdominal CT and MRI are used for insulinoma localization in humans, but they are sensitive only for cases with hepatic or lymph node metastasis or pancreatic tumors larger than 1 cm in diameter.100 In one prospective study of 14 dogs with insulinomas that underwent CT, primary lesions were correctly identified in 10 (71%) dogs, but specificity was low and accuracy in identifying metastatic lesions was poor.86 Endoscopic ultrasound is a common preoperative localizing technique in human patients with insulinomas, with reported sensitivities up to 94%.29,104 Nuclear scintigraphy using radiolabeled octreotide or indium 111 (111In) pentetreotide octreotide has been used in human and veterinary medicine.31,62 Octreotide, a somatostatin analog, is useful for detecting occult insulinomas possessing somatostatin receptors. Somatostatin receptor scintigraphy was effective in identifying the primary insulin-secreting tumor in 5 of 7 dogs, metastases to the regional lymph nodes in 3 of 3 dogs,31 and metastases to the liver in 2 of 3 dogs.56 However, the expense and the need for specialized facilities makes nuclear scintigraphy a limited option for veterinary patients.31
MANAGEMENT OF ISLET CELL TUMORS Surgical Therapy In patients stable enough to undergo anesthesia, surgical therapy remains the treatment of choice for ferrets, dogs, and humans with insulinomas.5,15,31,40,84 Surgical therapy in ferrets is usually not curative but may stop or slow the progression of the insulinoma.84 Among dogs with insulinomas, those treated surgically followed by as-needed medical management are more likely to become euglycemic for longer periods and have longer survival times compared with dogs managed only medically.5 This longer survival time has similarly been shown in ferrets treated with surgical plus medical therapy versus medical therapy alone (Tables 7-5 and 7-6).
Table 7-5 Survival Time of Ferrets Diagnosed with Pancreatic Islet Cell Tumors and Treated with Medical Therapy Alone Survival Time (days) 180-26717 9-50426 36-273111
Ferrets (n) 3 3 10
CHAPTER 7 Endocrine Diseases Before fasting the ferret for surgery, place an indwelling intravenous catheter and administer maintenance fluids with added 5% dextrose to prevent signs of hypoglycemia during the fast. Because of their rapid gastrointestinal transit time (3-4 hours8), ferrets can be fasted for a shorter period of time (3-6 hours) than cats and dogs.84 If possible, schedule surgery early in the day so that fasting time is minimal and the patient can be closely monitored in the immediate postoperative period. Monitor blood glucose concentrations before, during, and immediately after surgery. One or two pancreatic nodules are usually found at surgery, but as many as seven to nine nodules have been reported.17,111 Nodulectomy, partial pancreatectomy, or both are performed depending on the number and location of the pancreatic nodules (see Chapter 11). In ferrets, metastasis at the time of surgery is very uncommon; however, metastases to mesenteric and pancreatic lymph nodes, liver, and spleen are reported.17,33,65,111 Therefore it is important to biopsy or resect any suspicious lesions during surgery and perform a full abdominal exploration because concurrent, unrelated disease has been reported in many ferrets undergoing surgery for insulinoma removal.17,25,26,57,111 Ferrets usually recover quickly from surgery and, unlike the case in dogs, complications related to pancreatic surgery appear to be rare. Postoperative pancreatitis has been reported in only two ferrets.17,111 In routine cases, ferrets may be offered small amounts of water when they have recovered from anesthesia and are able to swallow. If the ferret is able to drink water without problems, offer small amounts of food in the form of kibble or highly palatable canned food (such as Hill’s Prescription Diet A/D, Hill’s Pet Nutrition, Topeka, KS). Administer intravenous fluid maintenance therapy until the ferret is eating and drinking well on its own. The addition of dextrose at 2.5% to 5% to maintenance fluids may be necessary if the ferret remains hypoglycemic after surgery. Hand feeding the ferret every 4 to 6 hours can be helpful after surgery to encourage eating and prevent signs of hypoglycemia. Routine postoperative pain management is necessary as well. Many ferrets that undergo stress from surgery manifest clinical signs of Helicobacter mustelae infection.84 These signs may include anorexia, melena, and bruxism.36 Therefore prophylactic treatment for Helicobacter with triple therapy may be considered.84 While the patient has an IV catheter in place, this can be accomplished with intravenous ampicillin and metronidazole, switching to oral amoxicillin, metronidazole and an H2 blocker when the ferret is discharged from the hospital (see Chapter 3).
While the ferret is hospitalized after surgery, monitor the blood glucose concentrations at least twice daily. In the immediate postoperative period, 48% to 94% of ferrets become euglycemic.17,26,65,111 Postoperative hyperglycemia has been reported only occasionally47,65 and is usually transient, resolving within 2 to 3 weeks.84 In these rare cases, the blood glucose level should be measured periodically until hyperglycemia resolves.84 Postoperative diabetes mellitus is rare but can occur.17,111 Most ferrets that have been treated surgically require medical management for recurrent hypoglycemia. In one study of 50 ferrets that were surgically treated for insulinoma, 42 (84%) eventually became hypoglycemic.17 In another study of 17 ferrets that were treated surgically for insulinoma, 10 (59%) eventually became hypoglycemic or redeveloped clinical signs.26 Therefore in any ferret that has had surgery for insulinoma, the blood glucose concentration should be checked 7 to 14 days after surgery and at 3-month intervals thereafter.84
Medical Therapy for an Acute Hypoglycemic Crisis Therapy for the acute onset of clinical signs associated with hypoglycemia depends on the severity of clinical signs and the location of the patient—either in the home or hospital setting. Advise owners about the clinical signs to watch for, such as lethargy, ptyalism, a fixed stare, pawing at the mouth, collapse, or seizures. If a ferret has mild clinical signs such as lethargy, hind end weakness, or ptyalism and is still alert enough to eat and swallow, it should be fed a small high-protein meal. If the ferret has a more severe hypoglycemic episode at home such as collapse, seizure, or coma, instruct the owner to put a sugar solution (Karo syrup, honey) on his or her fingers and rub it onto the ferret’s buccal mucosa. One should not place fingers into the ferret’s mouth because they might be bitten, and the sugar solution should not be poured directly into the ferret’s mouth because the liquid might be aspirated.31 If the ferret responds to the buccal glucose administration, the owner should feed a small high-protein meal once the ferret is more alert and able to swallow. The ferret should be kept quiet and transported to the veterinary hospital. The ferret should not be transported during an active seizure unless the administration of glucose to the buccal membranes has not controlled the seizure.31 If a ferret is presented comatose or seizing, place an IV catheter and administer a slow IV bolus of 50% dextrose (0.25-2 mL) to effect.84 The goal is to administer enough dextrose to alleviate the clinical signs without causing overstimulation of the tumor.
Table 7-6 Published Survival Times and Disease Free Intervals (Mean or Range) of Ferrets Diagnosed with Pancreatic Islet Cell Tumors and Treated with Surgical and Medical Therapy Treatment Surgery and medical therapy Surgery only Pancreatic nodulectomy Partial pancreatectomy + nodulectomy
97
Survival Time (days)
Disease-Free Interval (days)
Ferrets (n)
46-50365 483 (0-1100)26 — 510 (14-1207)17 456 (93-826)111 668 (219-1002)111
— — 240 (0-545)26 — 234 (0-546) 365 (0-690)
6 20 17 20 27 29
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SECTION I Ferrets
Rapid infusion of IV dextrose can result in a massive release of insulin into the circulation and subsequent rebound, severe hypoglycemia.31 Once the clinical signs have been controlled with judicious IV administration of dextrose and the patient is able to swallow, offer frequent feedings of a small amount of a high-protein food.31 Continuous IV infusion of 2.5% to 5% dextrose may be necessary.84 Ferrets that have seizures or are comatose must be monitored as critical patients and maintained on a continuous IV dextrose infusion. Rarely, anticonvulsant therapy such as diazepam is needed to control seizures until supportive care becomes effective.84 In such cases, follow anticonvulsant protocols used for dogs and cats in status epilepticus, making sure that hypoglycemia is properly addressed.84 The prognosis is generally poor for ferrets that remain comatose or continue to have seizures after 24 hours despite therapy with dextrose infusion and anticonvulsants.
Medical Management of Chronic Hypoglycemia Medical management for chronic hypoglycemia is begun when surgery cannot be performed or when clinical signs of hypoglycemia return after surgery.31 The goals of medical therapy are to reduce the frequency and severity of clinical signs and avoid an acute hypoglycemic crisis, not to establish euglycemia.31 Medical therapy of insulinoma is progressive: as signs of hypoglycemia worsen, increased frequency of feedings, increased doses of implemented drugs, and additional drugs are usually necessary.27 1. Food/diet: The first step in medical management is dietary management.27 Animals with insulinomas should be fed a diet that is high in protein and fat.66 Ferrets should be fed free choice and owners must ensure that the ferret is actually eating throughout the day. If the ferret is not eating on its own, the owner must hand-feed a high-fat, high-protein meal every 4 to 6 hours. Commercially available high-protein, high-fat, low-carbohydrate feline diabetic diets may be useful.3 Additionally, instruct owners to discontinue all treats that are high in simple sugars such as raisins or supplements containing corn syrup, sugar, molasses, or fructose.18 The rapid increase in blood glucose from the ingestion of these simple sugars can induce a rebound release of insulin, thus triggering a hypoglycemic episode.92 If a ferret is being fed an inappropriate diet at the time of insulinoma diagnosis, the owner must slowly transition the ferret to a more appropriate low-sugar, high-fat, high-protein diet. Rapid transition to a new diet may result in refusal to eat the new food or gastrointestinal upset, which can lead to a hypoglycemic episode.18 2. Glucocorticoids: Glucocorticoid therapy is begun when dietary manipulations are no longer effective in preventing the signs of hypoglycemia.31 a. Mechanism of action: Glucocorticoids increase glucose in the blood by stimulating hepatic glycogenolysis and providing the necessary substrates for hepatic gluconeogenesis.19 Prednisone and prednisolone are the most common glucocorticoids administered to ferrets with insulinomas. Liquid suspensions should be used, as they offer more accurate dosing.3 Such suspensions are commercially available or can be made by a compounding pharmacy in meat flavors that appeal to ferrets. Because ferrets have a noxious reaction to liquids containing alcohol, make sure that the suspension is alcohol-free. Although ferrets enjoy the taste of sweet flavors, owners and compounding
pharmacies must not add flavors or additives that contain sugar to the medications. b. Adverse effects: In general, ferrets are resistant to the typical side effects seen in dogs receiving glucocorticoid therapy, such as polydipsia, polyuria, and polyphagia. One report describes abdominal weight gain and impaired hair growth at shaved sites in ferrets on long-term steroid therapy.18 c. Dosage: Although there is a wide dose range for prednisone in ferrets, the minimal dose required to alleviate significant hypoglycemia should be given. Generally, it is recommended to begin at 0.25 to 0.5 mg/kg PO q12h. If this controls the signs of hypoglycemia, the medication is continued without dose adjustment. If signs persist or recur, the dose of prednisone should be gradually increased until signs of hypoglycemia abate.31 The immunosuppressive dosage of steroids is reported to be 2 to 4 mg/kg per day in dogs and 2 to 8 mg/kg per day in cats.19 The immunosuppressive dosage of steroids in ferrets is not reported, and it is not uncommon to use dosages of up to 2 mg/kg PO q12h.3,84 d. Monitoring: After initiating steroid therapy, the blood glucose concentration should be checked after 5 to 7 days of therapy. Because the goal of this therapy is to alleviate signs of hypoglycemia and not to attain euglycemia, the owner must monitor for signs of hypoglycemia at home, as this is more important in determining when the dose of prednisone must be increased or diazoxide must be added. After the patient’s clinical signs are initially stabilized, monthly rechecks are recommended. 3. Diazoxide: Diazoxide is usually administered in conjunction with glucocorticoids if diet and glucorticoid therapy alone are not sufficient in controlling signs of hypoglycemia. a. Mechanism of action: Diazoxide (Proglycem; Baker Norton Pharmaceuticals, Miami, FL) is a benzothiadiazide diuretic that increases glucose concentration in the bloodstream by inhibiting insulin release from the beta cells, stimulating hepatic gluconeogenesis and glycogenolysis, and inhibiting cellular uptake of glucose.31 b. Adverse effects: In dogs, the most commonly reported adverse reactions to diazoxide administration are anorexia and vomiting.31 Other potential complications are diarrhea, tachycardia, bone marrow suppression, aplastic anemia, thrombocytopenia, pancreatitis, diabetes mellitus, cataract formation, and sodium and fluid retention.31 These adverse effects have not been reported in ferrets. Because diazoxide is metabolized in the liver, lower doses may be necessary in patients with concurrent hepatic dysfunction.31 The major disadvantage of diazoxide is the expense of the commercial suspension. Although compounding pharmacists can make suspensions of diazoxide from tablets, the concentration of the resultant suspension cannot be guaranteed.84 c. Dosage: Initially, diazoxide should be administered at 5 mg/kg PO q12h. If clinical signs of hypoglycemia do not improve or recur, the dose of diazoxide can be slowly increased but should not exceed 60 mg/kg/day.31 d. Monitoring: The animal should be monitored as is recommended for glucocorticoid therapy. 4. Somatostatin: Somatostatin is used rarely for treatment. a. Mechanism of action: Octreotide (Sandostatin, Novartis Pharmaceuticals, East Hanover, NJ) is an analog of
CHAPTER 7 Endocrine Diseases somatostatin that inhibits the synthesis and secretion of insulin by normal and neoplastic beta cells. The responsiveness of insulin-secreting tumors to the suppressive effects of octreotide depends on the presence and affinity of somatostatin binding receptors on the tumor cells.31 In dogs, only one type of somatostatin receptor has been identified on insulin-secreting tumors versus five receptors that have been identified in humans.15,87 Studies to determine somatostatin receptor binding in insulin-secreting tumors in ferrets have not been done. Depending on the reference, success rate in alleviating clinical signs in dogs with insulinomas is 40% to 75%, although most eventually become refractory to the somatostatin.31,62,66 In four reported cases of its use in ferrets, only one ferret had an improvement in clinical signs while a second ferret had “equivocal” results.17,66,84 b. Adverse effects: There are no reported adverse effects in dogs or ferrets with the use of somatostatin, although reports in the human literature site the development of cholelithiasis15 and hypoglycemia due to suppression of glucagon.76 c. Dose: In one report of somatostatin use in a ferret, the recommended dosage is 1-2 mcg/kg every 8 to 12 hours.66 d. Monitoring: Because of reports of hypoglycemia in some human patients treated with somatostatin, close monitoring of blood glucose concentration is recommended in the initial stages of treatment. 5. Other medications: Phenytoin, an anticonvulsant that inhibits the release of insulin, and propanolol, a nonselective beta-adrenergic blocker that may block insulin secretion, have been reportedly used in humans with beta-cell tumors. These drugs have not been critically evaluated in dogs or ferrets and their use is not recommended.31 6. Chemotherapy: Treatment with streptozotocin, alloxan, and doxorubicin have been used in humans with pancreatic beta cell tumors.15,31,66 Streptozotocin and alloxan have been used in studies in dogs with equivocal results, and adverse effects associated with their use are potentially severe.31,67,81 The use of these drugs in ferrets has not been reported.
HISTOPATHOLOGY Neuroendocrine tumors are characterized histologically by the presence of neurosecretory granules. Under light microscopy, these tumors exhibit a bland, monotonous appearance with relatively uniform, well-differentiated cells with homogenous small nuclei, few if any nucleoli, abundant cytoplasm, and a low mitotic rate.15 More specifically, insulinomas contain cords and nests of polyhedral cells with moderate eosinophilic cytoplasm along with a delicate fibrovascular stroma.33 Primary masses can be fully, partially, or nonencapsulated depending on the histologic grade47,50,60,61 while metastatic masses have be described as nonencapsulated.33 More aggressive tumors can infiltrate locally into the surrounding tissue. Definitive diagnosis of insulinoma requires a histopathologic diagnosis acquired through surgical biopsy or necropsy. Tumors in ferrets have been described as beta-cell hyperplasia, adenoma, and carcinoma. The specific tumor may be a combination of any of these types and a single ferret may have several nodules of different histologic classification. In one study of 57 ferrets with insulinomas, 34 (60%) were reported to have carcinoma, 1 (2%) had adenoma, and 22 (38%) had a combination
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of carcinoma with either hyperplasia or adenoma.17 In a similar study of 20 ferrets, 11 (55%) cases were reported as adenoma and 9 (45%) as carcinoma.26 Immunohistochemical evaluation has been used to further characterize pancreatic islet cell tumors in ferrets. In two studies, weak immunoreactivity was reported for somatostatin, pancreatic polypeptide, and glucagon while strong immunoreactivity was reported for insulin.2,33 Immunohistochemical staining of one metastatic lesion was positive for glucagon, which is not seen in canine metastatic lesions.33 Immunohistochemical staining was positive for additional neuroendocrine markers such as chromogranin A and neuron-specific enolase in 22 ferret pancreatic islet cell tumors, including tumors that stained insulin-negative.2 These markers may be diagnostically useful in characterizing poorly differentiated tumors or those of indeterminant origin.2
PROGNOSIS Unlike dogs, which have a 50% metastasis rate,5 insulinomas in ferrets have a very low rate of metastasis. In 161 ferrets with reported insulinomas,* metastasis has been reported in only 7 ferrets (4%).17,33,65,111 The reported metastatic sites included the spleen,20 mesenteric lymph nodes,65 pancreatic lymph nodes,17,111 and liver.17,111 Although the metastasis rate is low, multiple pancreatic nodules are identified in up to 75% of affected ferrets at initial exploration, versus 15% in dogs.5 The prognosis for disease-free interval or survival time is similar for ferrets diagnosed with carcinomas and those with adenomas.26 Survival time based on medical therapy alone is difficult to determine as it has been reported in a total of only 16 ferrets, but it is reported to range from 9 to 504 days (see Table 7-5). Medical management may be effective in controlling clinical signs for periods of 6 months to 1.5 years.84 The paucity of reported cases treated with medical management alone is probably because definitive diagnosis of insulinoma requires a tissue sample, usually acquired via surgical biopsy. Survival time based on medical plus surgical therapy is reportedly longer than with medical therapy alone, with a range of 0 to 1204 days, depending on the reference (see Table 7-6). Where documented, the disease-free interval ranged from 0 to 690 days. In one report, pancreatic nodulectomy combined with partial pancreatectomy trended toward a longer survival time and disease-free interval versus pancreatic nodulectomy alone, although statistical significance was not shown.111 In retrospective studies of ferrets with insulinomas, the duration of clinical signs appears to be a negative prognostic indicator.26 In 20 ferrets with insulinomas, those that had a longer duration of clinical signs before diagnosis had a shorter diseasefree interval and shorter survival time.26
References 1. Ayub M, Levell MJ. Suppression of plasma androgens by the antiandrogen flutamide in prostatic cancer patients treated with Zoladex, a GnRH analogue. Clin Endocrinol. 1990;32:329-339. 2. Andrews GA, Myers NC. Immunohistochemistry of pancreatic islet cell tumors in the ferret (Mustela putorius furo). Vet Pathol. 1997;34:387-393. *References 16,17,25,26,33,47,50,51,59,60,61,65,111
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3. Antinoff N, Hahn K. Ferret oncology: diseases, diagnostics, and therapeutics. Vet Clin North Am Exot Anim Pract. 2004;7:579-625. 4. Bachrach WH. Action of insulin hypoglycemia on motor and secretory functions of the digestive tract. Physiol Rev. 1953;33:566-592. 5. Bailey DB, Page RL. Tumors of the endocrine system. In: Withrow SJ, Vail DM, eds. Withrow and MacEwen’s small animal clinical oncology. 4th ed. St. Louis: Saunders Elsevier; 2007:583-609. 6. Bazil CW, Pack A. Insulinoma presenting as seizure disorder. Neurol. 2001;56:817-818. 7. Bélanger A, Labrie F, Dupont A, et al. Endocrine effects of combined treatment with an LHRH agonist in association with flutamide in metastatic prostatic carcinoma. Clin Invest Med. 1988;11:321-326. 8. Bell J. Ferret nutrition and diseases associated with inadequate nutrition, in Proceedings. North Am Vet Conf. 1993:719-720. 9. Bennett R, Larraio L, Weisse C, Rosenthal K, et al. Collateral circulation during caval occlusion in ferrets, in Proceedings. Ann Conf Assoc Exot Mam Vet. 2008;105. 10. Benoit-Biancamano M, Morin M, Langlois I. Histopathologic lesions of diabetes mellitus in a domestic ferret. Can Vet J. 2005;46:895-897. 11. Besso JG, Tidwell AS, Gliatto JM. Retrospective review of the ultrasonographic features of adrenal lesions in 21 ferrets. Vet Rad Ultrasound. 2000;41:345-352. 12. Bielinska M, Kiiveri S, Parviainen H, et al. Gonadectomyinduced adrenocortical neoplasia in the domestic ferret (Mustela putorius furo) and laboratory mouse. Vet Pathol. 2006;43:97-117. 13. Bielinska M, Parviainen H, Kiiveri S, et al. Review paper: origin and molecular pathology of adrenocortical neoplasms. Vet Pathol. 2009;46:194-210. 14. Boari A, Papa V, Di Silverio F, et al. Type 1 diabetes mellitus and hyperadrenocorticism in a ferret. Vet Res Com. 2010;34(Suppl. 1):S107-110. 15. Brentjens R, Saltz L. Islet cell tumors of the pancreas: the medical oncologist’s perspective. Surg Clin North Am. 2001;81:527-542. 16. Buchanan KC, Belote DA. Pancreatic islet cell tumor in a domestic ferret. Contemp Top Lab Anim Sci. 2003;42:46-48. 17. Caplan ER, Peterson ME, Mullen HS, et al. Diagnosis and treatment of insulin-secreting pancreatic islet cell tumors in ferrets: 57 cases (1986-1994). J Am Vet Med Assoc. 1996;209:1741-1745. 18. Chen S. Pancreatic endocrinopathies in ferrets. Vet Clin North Am Exot Anim Pract. 2008;11:107-123. 19. Cohn LA. Glucocorticoid therapy. In: Ettinger S, ed. Textbook of veterinary internal medicine. 6th ed. St. Louis: Elsevier Saunders; 2005:503-508. 20. Coleman GD, Chavez MA, Williams BH. Cystic prostatic disease associated with adrenocortical lesions in the ferret (Mustela putorius furo). Vet Pathol. 1998;35:547-549. 21. Coles EH. Carbohydrate metabolism. In: Coles EH, ed. Veterinary clinical pathology. 4th ed. Philadelphia: WB Saunders Co; 1986:152-156. 22. Couzinet B, Pholsena M, Young J, et al. The impact of a pure anti-androgen (flutamide) on LH, FSH, androgens and clinical status in idiopathic hirsutism. Clin Endocrinol. 1993;39:157-162. 23. Danta G. Hypoglycemic peripheral neuropathy. Arch Neurol. 1969;21:121-132. 24. Desmarchelier M, Lair S, Dunn M, et al. Primary hyperaldosteronism in a domestic ferret with an adrenocortical adenoma. J Am Vet Med Assoc. 2008;233:1297-1301. 25. Eatwell K. Two unusual tumours in a ferret (Mustela putorius furo). J Small Anim Pract. 2004;44:454-459.
26. Ehrhart N, Withrow SJ, Ehrhart EJ, et al. Pancreatic beta cell tumor in ferrets: 20 cases (1986-1994). Am Vet Med Assoc. 1996;209:1737-1740. 27. Elie MS, Zerbe CA. Insulinoma in dogs, cats and ferrets. Compend Cont Ed Pract Vet. 1995;17:51-59. 28. Elliot DA, Nelson RW, Feldman ED, et al. Glycosylated hemoglobin concentrations in the blood of healthy dogs and dogs with naturally developing diabetes mellitus, pancreatic betacell neoplasia, hyperadrenocorticism, and anemia. J Am Vet Med Assoc. 1997;211:723-727. 29. Fein J, Gerdes H. Localization of islet cell tumors by endoscopic ultrasonography. Gastroenterology. 1992;103:711-712. 30. Fekete E, Woolley G, Little CC. Histological changes following ovariectomy in mice: I. dba high tumor strain. J Exp Med. 1941;74:1-8. 31. Feldman EC, Nelson RW. Beta-cell neoplasia: insulinoma. In: Feldman EC, Nelson RW, eds. Canine and feline endocrinology and reproduction. 3rd ed. St. Louis: WB Saunders; 2004:616-644. 32. Feldman EC, Schall WD, Kruth SA, et al. Amended insulin:glucose ratio. J Am Vet Med Assoc. 1986;188:1227-1230. 33. Fix AS, Harms CA. Immunocytochemistry of pancreatic endocrine tumors in three domestic ferrets (Mustela putorius furo). Vet Pathol. 1990;27:199-201. 34. Fox JG, Dangler CA, Snyder SB, et al. C-cell carcinoma (medullary thyroid carcinoma) associated with multiple endocrine neoplasms in a ferret (Mustela putorius). Vet Pathol. 2000;37:278-282. 35. Fox JG, Goad ME, Garibaldi BA, et al. Hyperadrenocorticism in a ferret. J Am Vet Med Assoc. 1987;191:343-344. 36. Fox JG. Bacterial and mycoplasmal diseases. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:258-259. 37. Garibaldi BA, Pequet Goad ME, Fox JG. Serum thyroxine (T4) and tri-iodothyronine (T3) radioimmunoassay values in the normal ferret. Lab Anim Sci. 1987;37:544-547. 38. Goss PE, Gwyn KM. Current perspectives on aromatase inhibitors in breast cancer. J Clin Oncol. 1994;12:2460-2470. 39. Gould WJ, Reimers TJ, Bell JA, et al. Evaluation of urinary cortisol:creatinine ratios for the diagnosis of hyperadrenocorticism associated with adrenal gland tumors in ferrets. J Am Vet Med Assoc. 1995;206:42-46. 40. Grant CS. Insulinoma. Best Pract Res Clin Gastroenterology. 2005;19:783-798. 41. Guyton AC, Hall JE. Insulin, glucagon, and diabetes mellitus. In: Guyton AC, Hall JE, eds. Textbook of medical physiology. 10th ed. Philadelphia: WB Saunders; 2000:884-894. 42. Harms CA, Stoskopf MK. Outcomes of adoption of adult laboratory ferrets after gonadectomy during a veterinary student teaching exercise. J Am Assoc Lab Anim Sci. 2007;46:50-54. 43. Heard DJ, Collins B, Chen DL, et al. Thyroid and adrenal function tests in adult male ferrets. Am J Vet Res. 1990;51:32-35. 43a. Hess L. Treatment of diabetes mellitus in a ferret with insulin glargine. J Am Vet Med Assoc. In press. 44. Himwich HE, Frostig JP, Fazekas JF, et al. The mechanism of the symptoms of insulin hypoglycemia. Am J Psychiatry. 1939;96:373-385. 45. Isselbacher KJ. Laboratory values of clinical importance. In: Isselbacher KJ, Martin JB, Braunwald E, et al, eds. Harrison’s principles of internal medicine. 13th ed. New York: McGrawHill, Inc; 1994:2489-2496. 46. Jallageas M, Mas N, Boissin J, et al. Seasonal variations of pulsatile luteinizing hormone release in the mink (Mustela vison). Comp Biochem Physiol C Pharmacol Toxicol Endocrinol. 1994;109:9-20. 47. Jergens AE, Shaw DP. Hyperinsulinism and hypoglycemia associated with pancreatic islet cell tumor in a ferret. J Am Vet Med Assoc. 1989;194:269-271.
CHAPTER 7 Endocrine Diseases 48. Johnson D. Current therapies for ferret adrenal disease, in Proceedings. Atlantic Coast Vet Conf. 2006: 1-7. 49. Johnson-Delaney CA. Medical therapies for ferret adrenal disease. Sem Avian Exot Pet Med. 2004;13:3-7. 50. Kaufman J, Schwarz P, Mero K. Pancreatic beta cell tumor in a ferret. J Am Vet Med Assoc. 1984;185:998-1000. 51. Kemmerer DW. Pancreatic beta-cell tumors in two domestic ferrets (Mustela putorius furo). Compan Anim Pract. 1988;2:29-30. 52. Kintzer PP, Peterson ME. Mitotane treatment of 32 dogs with cortisol-secreting adrenocortical neoplasms. J Am Vet Med Assoc. 1994;205:54-61. 53. Kuijten AM, Schoemaker NJ, Voorhout G. Ultrasonographic visualization of the adrenal glands of healthy ferrets and ferrets with hyperadrenocorticism. J Am Anim Hosp Assoc. 2007;43:78-84. 54. Kupersmith DS, Bauck L. Hyperadrenocorticism in a ferret: diagnosis (using ultrasound) and treatment. J Small Exot Anim Med. 1991;1:66-68. 55. Leifer CE, Peterson ME, Matus RE. Insulin-secreting tumor: diagnosis and medical and surgical management in 55 dogs. J Am Vet Med Assoc. 1986;188:60-64. 56. Lester NV, Newell SM, Hill RC, et al. Scintigraphic diagnosis of insulinoma in a dog. Vet Radiol Ultrasound. 1999;40:174-178. 57. Li X, Fox JG, Padrid PA. Neoplastic diseases in ferrets: 574 cases (1968-1997). J Am Vet Med Assoc. 1998;212:1402-1406. 58. Lipman NS, Marini RP, Murphy JC, et al. Estradiol-17 betasecreting adrenocortical tumor in a ferret. J Am Vet Med Assoc. 1993;203:1552-1555. 59. Lloyd CG, Lewis WG. Two cases of pancreatic neoplasia in British ferrets (Mustela putorius furo). J Small Anim Pract. 2004;45:558-562. 60. Lumeij JT, van der Hage MH, Dorrestein GM, et al. Hypoglycemia due to a functional pancreatic islet cell tumor (insulinoma) in a ferret (Mustela putorius furo). Vet Rec. 1987;120:129-130. 61. Luttgen PJ, Storts RW, Rogers KS, et al. Insulinoma in a ferret. J Am Vet Med Assoc. 1986;189:920-921. 62. Luyre JC, Behrend EN. Endocrine tumors. Vet Clin North Am Small Anim Pract. 2001;31:1083-1111. 63. Mann FA, Stockham SL, Freeman MB, et al. Reference intervals for insulin concentrations and insulin:glucose ratios in the serum of ferrets. J Small Exot Anim Med. 1993;2:79-83. 64. Marcondes JA, Minnani SL, Luthold WW, et al. Treatment of hirsutism in women with flutamide. Fertil Steril. 1992;57:543-547. 65. Marini RP, Ryden EB, Rosenblad WD, et al. Functional islet cell tumor in six ferrets. J Am Vet Med Assoc. 1993;202:430-433. 66. Meleo KA, Caplan ER. Treatment of insulinomas in the dog, cat, and ferret. In: Bonagura JR, ed. Current veterinary therapy XIII. Philadelphia: WB Saunders; 1999:357-361. 67. Moore AS, Nelson RW, Henry CJ, et al. Streptozotocin for treatment of pancreatic islet cell tumors in dogs: 17 cases (1989-1999). J Am Vet Med Assoc. 2002;221:811-818. 68. Mor N, Qualls Jr CW, Hoover JP. Concurrent mammary gland hyperplasia and adrenocortical carcinoma in a domestic ferret. J Am Vet Med Assoc. 1992;201:1911-1912. 69. Murray J. Melatonin implants: an option for use in the treatment of adrenal disease in ferrets. J Exot Mam Med Surg. 2005;3:1-6. 70. Murthy AS, Brezak MA, Baez AG. Postcastrational adrenal tumors in two strains of mice: morphologic, histochemical, and chromatographic studies. J Natl Cancer Inst. 1970;45:1211-1222. 71. Neuwirth L, Collins B, Calderwood-Mays M, et al. Adrenal ultrasonography correlated with histopathology in ferrets. Vet Radiol Ultrasound. 1997;38:69-74.
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72. Neuwirth L, Isaza R, Bellah J, et al. Adrenal neoplasia in seven ferrets. Vet Radiol Ultrasound. 1993;34:340-346. 73. Newman SJ, Bergman PJ, Williams B, et al. Characterization of spindle cell component of ferret (Mustela putorius furo) adrenal cortical neoplasms–correlation to clinical parameters and prognosis. Vet Comp Onc. 2004;2(3):13-124. 74. O’Brien RT, Paul-Murphy J, Dubielzig RR. Ultrasonography of adrenal glands in normal ferrets. Vet Radiol Ultrasound. 1996;37:445-448. 75. Penninck D. Ultrasonographic features of pancreatic disorders. In: Penninck D, d’Anjou MA, eds. Atlas of small animal ultrasonography. Ames: Blackwell Publishing; 2008:322-337. 76. Perry RR, Vinik AI. Diagnosis and management of functioning islet cell tumors. J Clin Endocrinol Metab. 1995;8:2273-2278. 77. Peterson 2nd RA, Kiupel M, Bielinska M, et al. Transcription factor GATA-4 is a marker of anaplasia in adrenocortical neoplasms of the domestic ferret (Mustela putorius furo). Vet Pathol. 2004;41:446-449. 78. Peterson 2nd RA, Kiupel M, Capen CC. Adrenal cortical carcinomas with myxoid differentiation in the domestic ferret (Mustela putorius furo). Vet Pathol. 2003;40:136-142. 79. Peterson RA, Kiupel M, Capen CC. Adrenal cortical carcinomas with myxoid differentiation in the domestic ferret (Mustela putorius furo). Vet Pathol. 2003;40:136-142. 80. Plourde PV, Dryoff M, Dukes M. Arimidex: a potent and selective fourth-generation aromatase inhibitor. Breast Cancer Res Treat. 1994;30:103-111. 81. Plumb DC. Plumb’s veterinary drug handbook. 5th ed. Stockholm: Blackwell; 2005. 82. Polton GA, White RN, Brearley MJ, et al. Improved survival in a retrospective cohort of 28 dogs with insulinoma. J Small Anim Pract. 2007;48:151-156. 83. Quesenberry KE, Orcutt C. Basic approach to veterinary care. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders an imprint of Elsevier; 2003:13-24. 84. Quesenberry KE, Rosenthal KR. Endocrine diseases. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders an imprint of Elsevier; 2003:79-90. 85. Ramer JC, Benson KG, Morrisey JK, et al. Effects of melatonin administration on the clinical course of adrenocortical disease in domestic ferrets. J Am Vet Med Assoc. 2006;229:1743-1748. 86. Robben JH, Pollak YW, Kirpensteijn J, et al. Comparision of ultrasonography, computed tomography, and single-photon emission computed tomography for the detection and localization of canine insulinoma. J Vet Intern Med. 2005;19:15-22. 87. Robben JH, Visser-Wisselaar HA, Rutteman GR, et al. In vitro and in vivo detection of functional somatostatin receptors in canine insulinomas. J Nucl Med. 1997;38:1036-1042. 88. Rosenthal K, Peterson M. Clinical case conference: stranguria in a castrated male ferret. J Am Vet Med Assoc. 1996;209:462-464. 89. Rosenthal K, Peterson M. Plasma androgen concentrations in ferrets with adrenal gland disease. J Am Vet Med Assoc. 1996;209:1097-1102. 90. Rosenthal KL, Peterson ME, Quesenberry KE, et al. Evaluation of plasma cortisol and corticosterone responses to synthetic adrenocorticotropic hormone administration in ferrets. Am J Vet Res. 1993;54:29-31. 91. Rosenthal KL, Peterson ME, Quesenberry KE, et al. Hyperadrenocorticism associated with adrenocortical tumor or nodular hyperplasia in ferrets: 50 cases (1987-1991). J Am Vet Med Assoc. 1993;203:271-275. 92. Rosenthal KL. Feeding the hypoglycemic ferret, in Proceedings. North Am Vet Conf. 2009:1766. 93. Rosenthal KL. Adrenal gland disease in ferrets. Vet Clin North Am Small Anim Pract. 1997;27:401-418.
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94. Rosenthal KL. Ferret and rabbit endocrine disease diagnosis. In: Fudge AM, ed. Laboratory medicine: avian and exotic pets. Philadelphia: WB Saunders Co; 2000:319-324. 95. Saadi HF, Bravo EL, Aron DC. Feminizing adrenocortical tumor: steroid hormone response to ketoconazole. J Clin Endocrinol Metab. 1990;70:540-543. 96. Schoemaker NJ, Mol JA, Lumeij JT, et al. Plasma concentrations of adrenocorticotrophic hormone and alpha-melanocytestimulating hormone in ferrets (Mustela putorius furo) with hyperadrenocorticism. Am J Vet Res. 2002;63:1395-1399. 97. Schoemaker NJ, Schuurmans M, Moorman H, et al. Correlation between age at neutering and age at onset of hyperadrenocorticism in ferrets. J Am Vet Med Assoc. 2000;216:195-197. 98. Schoemaker NJ, Teerds KJ, Mol JA, et al. The role of luteinizing hormone in the pathogenesis of hyperadrenocorticism in neutered ferrets. Mol Cell Endocrinol. 2002;197:117-125. 99. Sharawy MM, Liebelt AG, Dirksen TR, et al. Fine structural study of postcastrational adrenocortical carcinomas in female CE-mice. Anat Rec. 1980;198:125-133. 100. Stabile BE. Islet cell tumors. Gastroenterologist. 1997;5:213-232. 101. Swiderski JK, Seim 3d HB, MacPhail CM, et al. Long-term outcome of domestic ferrets treated surgically for hyperadrenocorticism: 130 cases (1995-2004). J Am Vet Med Assoc. 2008;232:1338-1343. 102. Tang Z, Lee JH, Louie RF, et al. Effects of different hematocrit levels on glucose measurements with handheld meters for point-of-care testing. Arch Pathol Lab Med. 2000; 124:1135-1140. 103. Thoreson SI, Aleksandersen M, Lonaas L, et al. Pancreatic insulin-secreting carcinoma in a dog: fructosamine for determining persistent hypoglycaemia. J Small Anim Pract. 1995;36:282-286. 104. Tucker ON, Cotty PL, Conlon KC. The management of insulinomas. Brit J Surg. 2006;93:264-275.
105. Wagner R, Finkler M, Fecteau K, et al. The treatment of adrenal cortical disease in ferrets with 4.7-mg deslorelin acetate implants. J Exot Pet Med. 2009;18:146-152. 106. Wagner RA, Bailey EM, Schneider JF, et al. Leuprolide acetate treatment of adrenocortical disease in ferrets. J Am Vet Med Assoc. 2001;218:1272-1274. 107. Wagner RA, Piché CA, Jöchle W, et al. Clinical and endocrine responses to treatment with deslorelin acetate implants in ferrets with adrenocortical disease. Am J Vet Res. 2005;66:910-914. 108. Weaver C, Baum M. Differential regulation of brain aromatase by androgen in adult and fetal ferrets. Endocrinology. 1991;128:1247-1254. 109. Weiss CA, Scott MV. Clinical aspects and surgical treatment of hyperadrenocorticism in the domestic ferret: 94 cases (19941996). J Am Anim Hosp Assoc. 1997;33:487-493. 110. Weiss CA, Williams BH, Scott JB, et al. Surgical treatment and long-term outcome of ferrets with bilateral adrenal tumors or adrenal hyperplasia: 56 cases. J Am Vet Med Assoc. 1999;215:820-823. 111. Weiss CA, Williams BH, Scott MV. Insulinoma in the ferret: clinical findings and treatment comparison of 66 cases. J Am Anim Hosp Assoc. 1998;34:471-475. 112. Wheeler J, Bennett RA. Ferret abdominal surgical procedures. Part I. Adrenal gland and pancreatic beta-cell tumors. Comp Contin Ed Pract Vet. 1999;21:815-822. 113. Williams BH, Weiss CA. Ferret neoplasia. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2003:91-106. 114. Wilson GH, Greene CE, Greenacre CB. Suspected pseudohypoparathyroidism in a domestic ferret. J Am Vet Med Assoc. 2003;222:1093-1096.
CHAPTER
8
Neoplasia
Natalie Antinoff, DVM, Diplomate ABVP (Avian), and Bruce H. Williams, DVM, Diplomate ACVP
Etiology Incidence and Behavior Tumors of the Endocrine System Insulinoma Adrenocortical Neoplasms Thyroid Neoplasms Tumors of the Hemolymphatic System Classification of Lymphoma Signalment and Clinical Signs Laboratory Evaluation Diagnostic Imaging Cytologic/Histologic Description Treatment Tumors of the Skin and Subcutis Tumors of the Gastrointestinal Tract Tumors of the Reproductive Tract Tumors of the Musculoskeletal System Tumors of the Nervous System Tumors of the Urinary System Tumors of the Respiratory System Vascular Neoplasms Miscellaneous Neoplasms
With the exception of routine vaccinations, neoplasms today may represent the most common reason for presenting a ferret for veterinary care. The probability is good that most ferrets will develop a neoplasm of the endocrine system during the “golden age” for tumors (4 to 6 years) and excellent that some type of neoplasm will become evident over the course of a lifetime. In ferrets derived from American bloodlines, the incidence of three neoplasms—adrenocortical neoplasia, insulinoma, and malignant lymphoma—likely exceeds the incidence of all other neoplasms combined. The increasing popularity of ferrets as both pets and laboratory animals has facilitated the compilation of impressive Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
collections of neoplasms that provides a fairly accurate look at the distribution of neoplasia in this species.4,16,36,37,44,60,62,68 In the previous edition of this book, the authors provided a review of common neoplasia as well as frequency data based on the extensive collection of the Armed Forces Institute of Pathology.68 With the current expansion of literature regarding neoplastic disease in this species as well as our desire to expand the discussion of available and emerging clinical treatments, frequency data of common neoplasms are presented based on the available literature for this species. Clinical discussion is derived from the authors’ clinical experiences as well as a comprehensive review of available literature. Space does not permit us to cover every type of neoplasm that has been reported in the ferret (as rarer and rarer tumors find their way into single case reports); thus we present only the major types of neoplasms, including their diagnosis, treatment, and prognosis. One tenet should be considered by all veterinarians treating ferrets and their neoplasms: a ferret is not a cat or a dog. The clinical behavior, prognosis, and paraneoplastic syndromes in ferrets are often far different from what is seen with similar neoplasms in dogs or cats. For example, insulinoma in the ferret is a neoplasm that rarely metastasizes and behaves in a benign fashion, as opposed to the same neoplasm in dogs and cats, which metastasizes widely and rapidly and results in short survival times. Adrenocortical carcinoma, a neoplasm prone to metastasize widely in the dog, metastasizes only late in the course of disease in ferrets; with early removal, it warrants a good prognosis. Mast cell tumors, often malignant (and fatal) in the dog, are invariably benign and associated with a good prognosis in ferrets. Thus, practitioners who extrapolate diagnostic and therapeutic options from comparable syndromes in more traditional pet species may find themselves in difficult and unexpected situations.
ETIOLOGY While we now have tremendous information on the frequency and distribution of neoplasia, there is still little definitive information on the causes of many common neoplasms in the ferret (as is often the case in human neoplasms). A large number of theories abound, but only rarely with supportive evidence. 103
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Three main schools of thought exist for ferrets, and it is likely that many neoplasms may be the result of several coexisting factors at once: 1. Husbandry issues. The “domestication” of the ferret as a pet species in many countries of the world involves varying degrees of environmental and surgical manipulation of the animal itself. Proof surrounding the effects of early neutering of ferrets and the subsequent development of adrenocortical neoplasia is now published and this mechanism is widely accepted.7,53,54 Dietary manipulation, especially the common practice of feeding high-carbohydrate diets and treats, has been suggested to be a primary cause for the increased incidence of insulinoma (when compared with the incidence in animals fed raw whole prey [rats, mice, etc.] as the dietary staple).53 Finally, the modern ferret owner’s predilection for indoor housing and artificial lighting may also play a role in the development of certain types of neoplasms. In Europe, where most ferrets are housed outdoors and exposed to natural lighting cycles, the incidence of neoplasia, especially adrenocortical, is greatly decreased.53 However, other factors, including delayed neutering, may also affect the development of neoplasia in European ferrets. 2. Genetic (familial) predisposition. While genetic or chromosomal aberrations are yet to be studied in domestic ferrets, the tremendous incidence of neoplasia in American bloodlines of ferrets as compared with their European counterparts certainly lends credence to this widely held belief. A recent case report by Fox22 documents a syndrome of multiple neoplasms in an adult ferret that closely resembles (multiple endocrine neoplasia (MEN) type 2 in humans, a condition known to be the result of a genetic point mutation. 3. Infectious agents. Suspicious cluster outbreaks of malignant lymphoma in laboratory colonies and rescue operations3,18 have sparked the investigation of a possible viral etiology for this neoplasm in ferrets. Transforming retroviruses are known to be responsible for the development of lymphoma in a number of other species, including humans, cats, and rabbits among others. Erdman20 demonstrated the transmissibility of this neoplasm between ferrets using cell-free inocula, thus furthering this theory, although a prolonged incubation time was required. Helicobacter mustelae, a ubiquitous inhabitant of the stomach of ferrets, has been circumstantially incriminated in the development of gastric adenocarcinoma33 (which is enhanced when coupled with the ingestion of chemical carcinogens as promoters)21,23 as well as in the development of gastric B-cell (MALT) lymphomas.17 In addition, a progression from chronic inflammatory intestinal disease to neoplasia has been proposed in cats.70
INCIDENCE AND BEHAVIOR Most studies agree that the endocrine system appears to be the most common site of neoplasia in ferrets.8,36,37,68 Pancreatic islet cell tumors (also known as insulinomas) are the most common neoplasms overall, with adrenocortical neoplasms being the second most common.18,36,37,68 In all studies, lymphoma was both the most common hematopoietic neoplasm and the most common malignancy.8,36,37,68 Between 12% and 20% of cases in each study had multiple tumor types, with insulinoma and adrenocortical carcinoma being seen concurrently most often.8,36,37,68 However, the presence of multiple tumor types in an individual should not be interpreted as a neoplastic
syndrome arising from a common tumorigenic mechanism. In a study of 66 cases in which ferrets had multiple concurrent neoplasms, Li36 found no evidence of an association between tumor type and multiplicity. Because endocrine neoplasia is extremely common in American ferrets today (and is increasing in global frequency), it comes as no surprise that middle-aged and geriatric ferrets have multiple tumors developing over time.
TUMORS OF THE ENDOCRINE SYSTEM By far most neoplasms in domestic ferrets in North America arise in the endocrine system; chiefly, tumors of the pancreatic islets and the adrenal cortex (the two overall most common neoplasms in the ferret, as previously discussed).
INSULINOMA While multiple studies have pointed to insulinomas (tumors of the beta cells of the pancreatic islets) as the most common neoplasm of the ferret, they may be slightly overrepresented because of their relatively obvious symptomatology, response to surgical excision, and tendency to recur over time. This neoplasm is most commonly seen in middle-aged ferrets, with no gender predilection. Insulinoma in the ferret exhibits a far different behavior than in the dog or cat.9,62 In the dog and cat, these are highly malignant neoplasms with marked metastatic potential, leading to a short survival time. In ferrets, these same neoplasms have low metastatic potential and tend to respond well to medical management for long periods of time; their removal may result in a symptom-free or medication-free interval.62 In truth, some reports have clouded the issue of metastatic potential of insulinoma in the ferret. True metastasis involves the translocation, either via blood or lymph, to another organ; this is seen in the dog and cat, where the metastasis of islet cell tumors to local lymph nodes, liver, or other visceral organs is common. Some papers have incorrectly referred to the additional development of insulinoma within the pancreas over time as “metastasis,” whereas recurrence would be a more appropriate term. Other papers have labeled islet cell tumors as malignant (“islet cell carcinomas”) based solely on microscopic features of the tumor cells without evidence of intraorgan translocation or recurrent disease. The diagnosis of insulinoma is not exceedingly difficult in the ferret and is generally based on a combination of the characteristic clinical signs and a low fasting blood glucose level in the absence of a nonendocrine etiology (see Chapter 7). The hypoglycemia resulting from the inappropriate secretion of insulin by these tumors generally results in a constellation of neurologic signs ranging from mild (ataxia or disassociation from the surroundings) to severe (seizures, coma). Blood glucose levels of less than 60 mg/dL in the ferret are generally diagnostic for insulinoma even in the absence of clinical signs. Some individuals may present with a history of neurologic disease and a normal fasting blood glucose; in the early stages of this condition, insulin release may be sporadic and clinical signs may be intermittent. The determination of insulin levels is rarely indicated prior to institution of therapy and is of no value in cases where blood glucose is above 60 mg/dL. Therapeutic approaches for the treatment of insulinoma in ferrets are widely reported. In our experience, surgical excision is the preferred course of treatment for symptomatic animals with documented hypoglycemia.
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In a clinical study,62 partial pancreatectomy yielded the longest disease-free intervals and survival times (365 and 668 days, respectively), followed by simple nodulectomy (234 and 456 days, respectively), although surgery did not eliminate the need for concurrent medical management in all cases. Medical treatment alone resulted in a mean disease free interval of 22 days and a mean survival time of 186 days. While the potential for affected animals to develop multiple islet cell tumors over time is well known (up to 40% develop additional tumors within 10 months), the tumorigenic process remains unclear. Owners should be well aware of the potential for recurrence before surgical removal.
ADRENOCORTICAL NEOPLASMS The second most common neoplasm in the domestic ferret is also of endocrine origin and originates in the adrenal cortex. In the intact ferret, seasonal stimulus of the hypothalamus results in liberation of a range of hormones, including luteinizing hormone, which stimulates sex steroid production from the ovaries or testes. In neutered animals, the absence of gonads results in a lack of negative feedback for the hypothalamus and, under constantly elevated levels of luteinizing hormone, pluripotent cells of the zona reticularis differentiate into cells capable of producing estrogen and other intermediate sex steroid metabolites, including androstenedione and hydroxyprogesterone.7,52 Multiple studies have reported the average age of ferrets with adrenal disease at approximately 4.5 years,8,61,68 but the disease has also been reported in ferrets under 12 months of age,36 with no gender predilection. The relatively obvious clinical signs exhibited by most ferrets with adrenal disease contribute significantly to the frequency of their presentation for treatment. Affected ferrets exhibit a constellation of cutaneous, behavioral, and reproductive signs that make them easily identifiable. Follicular atrophy resulting from excessive levels of estrogen results in a characteristic bilateral truncal alopecia in about half of affected animals, although irregular, patchy hair loss may be a presenting sign in a minority; in rare cases, no alopecia is appreciated. Vulvar swelling, similar to that seen in females in estrus may be seen in up to 90% of affected neutered jills, although absence of vulvar swelling does not rule out this disease.51,57 The effects of estrogen on the prostatic glandular epithelium in male ferrets may result in dysuria due to prostatic cysts or abscesses; if not treated promptly, azotemia, obstruction, and ultimately uremia are probable sequelae. Finally, the presence of elevated levels of testosterone in the male or estrogen in the female may result in a return to intact sexual behavior such as mounting, urine marking, and aggression. To confirm enlargement, determine affected side, and screen for concurrent disease. For best detail, use a 14.5-MHz probe. Look for the right adrenal gland medial to the cranial pole of the right kidney, cranial to the origin of the cranial mesenteric artery, and adjacent or adherent to the caudal vena cava. The left adrenal gland is cranial and medial to the cranial pole of the left kidney, lateral to the aorta, and cranial to the left renal artery. Adenomas and adenocarcinomas can occur and cannot be differentiated without histopathology.6 Laboratory evaluation of circulating sex steroids is occasionally performed in cases where clinical signs are subtle or may be masked by concurrent disease. Estrogen, androstenedione, and 17-hydroxyprogesterone have been identified as the most
Fig. 8-1 Carcinoma of the right adrenal gland in a ferret (open arrow), demonstrating the proximity between these neoplasms and the caudal vena cava (closed arrow). Note that degree of malignancy cannot be determined by size or degree of invasiveness as visualized at surgery.
sensitive hormones for the detection of adrenocortical lesions in the ferret and are over 95% predictive in the diagnosis of adrenocortical disease in this species.51 Practitioners are cautioned that hyperadrenocorticism in ferrets is not a form of Cushing’s disease, and cortisol testing is not useful in routine diagnosis. Elevations in cortisol52 and even aldosterone13 levels have been documented in ferrets with adrenal neoplasia, but these are rare and inconsistent findings and are not considered diagnostic. From a surgical standpoint, the incidence of the lesion appears equivalent between the left and right adrenal glands, and approximately 20% are bilateral.68 A wide range of surgical approaches exist for removing affected adrenal glands, and surgery is considered the treatment of choice for this condition.64 Because of its proximity to the vena cava (Fig. 8-1), surgical excision of the right adrenal gland often proves to be a challenge for most practitioners; a range of successful surgical options—including cryotherapy, laser dissection, and microvascular techniques—has been described for right adrenalectomy in the ferret.68 In cases where the neoplasm occludes the vena cava by 50% or more, the neoplasm and the affected section of vena cava may be excised en bloc, but this is not recommended by the authors. Excision of the vena cava carries a high risk of complications including renal failure and death, and these risks are greater than the risk of death from adrenal disease. While presurgical ultrasound examination may be used to identify the side (or sides) at which an affected neoplasm is located, the absence of adrenomegaly or an identifiable nodule via ultrasound does not obviate the need for surgery in affected individuals, since functional lesions may be present in normal-sized adrenal glands.6 Several options for medical management have emerged in recent years (see Chapters 7 and 11 for more in-depth discussions of surgical and medical treatments). From a histologic (and prognostic) standpoint, proliferative lesions in the adrenal cortex of affected individuals fall into a spectrum ranging from hyperplastic lesions to benign or malignant neoplasms. A good prognosis appears warranted in the case of all surgically removed lesions, regardless of location (right vs. left), histologic grade, or completeness of excision.57,68 However, in all cases caution owners that lesions in
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the contralateral adrenal gland occasionally develop, resulting in recurrent disease at a later date, and metastatic disease may be seen in a low percentage of highly anaplastic carcinomas.46 Medical treatment, largely directed at decreasing circulating levels of sex steroids in affected animals, is generally reserved for nonsurgical candidates. These drugs are temporarily effective and are useful in ameliorating clinical signs; however, their effectiveness in halting the growth of established lesions or diminishing the risk of metastatic disease or hemoperitoneum associated with large tumors is still in question.60 Other neoplasms may be seen arising from adrenal glands in ferrets. First described in 1995,24 leiomyosarcomas of the adrenal capsule are often encountered in the ferret and may result in confusion on the part of the practitioner as well as the pathologist. These very firm neoplasms may lead practitioners to perform adrenalectomy on normally functioning adrenal glands. Unfortunately these cannot be differentiated without histopathology, so the practitioner is forced to make an intraoperative decision. The presence of the tumor may also mask the presence of proliferative adrenocortical lesions unless multiple sections at 1 mm or more are examined. These neoplasms have demonstrated estrogen receptors on the smooth muscle cells in several of these tumors, suggesting a possible etiology for their development as well as the common finding of smooth muscle proliferation in adrenocortical tumors.41
THYROID NEOPLASMS Thyroid neoplasms are extremely uncommon in this species. Nonfunctional thyroid follicular adenocarcinoma was reported in one ferret with infiltration into the surrounding tissue,69 and another was found with metastasis to cervical lymph nodes and liver.10 One case of a C-cell carcinoma22 (seen along with concurrent adrenocortical adenoma and insulinoma) has been reported. Clinical signs were not observed.
TUMORS OF THE HEMOLYMPHATIC SYSTEM Lymphoma (malignant lymphoma, lymphosarcoma) is the most common malignancy in the domestic ferret and the third most common neoplasm overall (after islet cell tumors and adrenocortical neoplasia). Lymphoma denotes solid-tissue tumors composed of neoplastic lymphocytes in visceral organs or lymph nodes throughout the body. These neoplasms most commonly arise spontaneously; however, horizontal transmission of malignant lymphoma in ferrets by using cell or cell-free inoculum has been documented.20 This finding, coupled with the occasional clustering of lymphomas in a single facility, has prompted speculation that some variants of lymphoma in the ferret may be the result of a retroviral infection.19 A viral agent has not as yet been isolated from cases of lymphosarcoma in the ferret, and associations with feline leukemia virus and Aleutian disease (parvovirus) have been disproved.18,20
CLASSIFICATION OF LYMPHOMA Today there is substantial variation in the classification of lymphoma, which leads to a lack of consistency in the evaluation of any form of cumulative data for comparison of disease or prognostic outcome. There is no universally accepted classification scheme for ferrets; even among dogs and cats, pathologists differ in the descriptive information they routinely include in
Table 8-1 Recommended Grading System for Ferret Tumors Based on Cell Morphology Nuclear size (relative to red blood cell [RBC] size) Small Medium Large
≤1 RBC >1 but <3 RBC ≥3 RBC
Mitotic index (per high-power field) Low Intermediate High
<3 3-8 >8
Additional descriptive grading information that might also be included Nuclear morphology Round Indented/asymmetric/irregular
Nucleoli Distinct Indistinct
histopathology reports. As an example of the importance of a uniform system, three papers have described ferret lymphomas as low-, intermediate-, and high-grade, but each paper uses different criteria to define “low” and “high,” creating inconsistency in our ability to interpret or compare.1,16,43 Clinicians should obtain as much descriptive information as possible for all cases, following the guidelines below as closely as possible, to ensure the most appropriate diagnosis and treatment for each individual. This may also improve prognostic ability in the future. All diagnostic workups should include both grading (histologic description in as much detail as possible) and staging (classification of disease) information. Ideally, phenotyping (immunohistochemistry to define cell origin) would also be included, although this is not yet routine in general clinical practice. Grading provides a histologic description based on cell morphology independent of phenotype (B-cell or T-cell). This provides indices like low, intermediate, and high grade based on cellular size and mitotic indices. The most commonly accepted grading system in companion animal medicine is the National Cancer Institute Working Formulation (NCI-WF), which differentiates cells based on morphology.39,43 There is still some discrepancy, so we recommend following a standard protocol (Table 8-1). Staging. This is a clinical description of the disease, providing information about the location of the neoplasia as well as its extent of dissemination throughout the body. The most commonly accepted staging system in veterinary medicine is the World Health Organization (WHO) staging system, which is also generally accepted by the American College of Veterinary Pathologists (ACVP). This system is based on descriptions of the clinical presentation, anatomic location, and disease progression (Table 8-2).39 In cats, a secondary staging system is used based solely on anatomic location. This would also be extremely useful in ferrets, and we recommend that this information also be included in staging (Table 8-3). Examples of appropriately staged and graded lymphoma in a ferret might be as follows: • Stage: I, alimentary; Grade: small-cell, low mitotic activity, round nuclei, indistinct nucleoli • Stage: IV, multicentric; Grade: large-cell, intermediate mitotic activity, round nuclei, distinct nucleoli
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Table 8-2 Staging for Lymphoma Based on Anatomic Location and Clinical Presentation Stage
Characteristics
I II
Affecting a single node or tissue in a single organ Multiple lymph nodes in one area of the body (same side of the diaphragm) Generalized lymph node involvement (both sides of the diaphragm) Any of the above + liver or spleen Any of above + blood or bone marrow
III IV V
Table 8-3 Anatomic Staging for Lymphoma Staging
Characteristics
Multicentric
Multiple lymph nodes, usually on both sides of diaphragm May also involve liver, spleen, bone marrow, or other extranodal sites Solitary mass within gastrointestinal tract or mesenteric node Multiple masses ± regional involvement of intra-abdominal node Diffusely infiltrating any part of bowel Mediastinal lymph nodes Not usually involving thymus Other locations: Renal CNS Ocular Cardiac Sometimes included in extranodal
Alimentary
Mediastinal Extranodal
Cutaneous
Lastly, if any neoplastic lymphocytes are present in the bone marrow or the peripheral blood, a diagnosis of lymphocytic leukemia is appropriate. Chronic lymphocytic leukemia indicates the presence of excessive numbers of mature (small) lymphocytes in the peripheral blood, with total leukocyte counts ranging from normal into the hundreds of thousands. Acute lymphoblastic leukemia indicates the presence of immature lymphocytes (lymphoblasts) in the bone marrow as well as in the peripheral blood, with leukocyte counts well in excess of normal. True lymphomas are far more commonly seen than leukemias, at a ratio of approximately 10:1.68 Phenotyping. Phenotyping defines tumor etiology as either B-cell or T-cell in origin. This can only be determined with the use of immunohistochemical stains or flow cytometry.27 CD3 is a T-cell marker, and CD79α is a B-cell marker. Although this information is useful, it is not routinely assessed in ferrets at this time. However, flow cytometric assays are becoming commonly used in companion animal medicine, and as phenotyping becomes a more standard part of a diagnostic workup, this may yield important prognostic information for ferrets. Two other types of lymphoma should be discussed here. Cutaneous (epitheliotropic) lymphoma is of T-cell origin and possesses a mature lymphocytic phenotype and a profound affinity to infiltrate epithelial structures, such as the epidermis and hair follicles (Fig. 8-2). It alone among the ferret lymphomas does
Fig. 8-2 Cutaneous lymphoma in a ferret. Surgical excision of this ulcerated neoplasm (arrow) was accomplished and, despite several recurrences, the ferret was still alive 3 years later.
not warrant a poor prognosis at onset, as prolonged survival times (possibly up to 3-4 years) are associated with it, especially in cases where cutaneous lesions are rapidly surgically excised. Unlike epitheliotropic lymphoma (mycosis fungoides) in dogs and humans, the clinical picture does not necessarily progress to systemic involvement (Sézary’s syndrome). Epitheliotropic lymphoma is commonly seen in the feet and extremities of ferrets, resulting in grossly swollen, hyperemic, alopecic feet. Untreated, these lesions grow in size and multiple lesions will develop. Complete surgical excision of cutaneous lesions may result in prolonged disease-free intervals; chemotherapeutic attempts, both topical and systemic, have generally proved to be unsatisfactory.34,50 Gastric lymphomas, or mucosal-associated lymphoid tissue (MALT) lymphomas, have been reported in four ferrets (see also Chapter 3).17 Considered akin to lymphomas associated with Helicobacter pylori infection in humans, these neoplasms arose in the stomach of ferrets infected with Helicobacter mustelae. An interesting feature of this proposed form of lymphoma is that although neoplastic cells varied in phenotype (two lymphocytic and two lymphoblastic forms), all four cases were composed of monoclonal B lymphocytes.17
SIGNALMENT AND CLINICAL SIGNS There is no universal signalment or clinical presentation for lymphoma in ferrets. It may occur at any age and has been reported in ferrets as young as 2 months of age. There is no color or sex predisposition. One paper historically reported that young ferrets (<2 years of age) develop a lymphoblastic form char acterized by disseminated disease often involving spleen, liver, thymus, or mediastinum and rapid progression, while adult ferrets develop a slower, more insidious form consisting of mature, well-differentiated small lymphocytes that are accompanied by peripheral lymphadenopathy and slower progression.16 This paper has been quoted repeatedly throughout veterinary literature. However, there have been several more recent publications that failed to show this correlation. Although it was not the primary purpose of either investigation, these retrospective studies found that the lymphoblastic form exists commonly in all age groups (Fig. 8-3).1,43 In one paper, all multicentric lymphomas
108
SECTION I Ferrets number of ferrets, correlating with small intestinal tumor. Hypercalcemia was present in 2 of 28 ferrets (both with T-cell lymphoma).1
DIAGNOSTIC IMAGING
Fig. 8-3 Multicentric lymphoma in a 1-year-old ferret. Note the thymic mass (open arrow), and marked hepatosplenomegaly (closed arrows) as a result of massive infiltration by this neoplasm.
that were identified were comprised of blast cells and occurred largely in adult ferrets.43 Another paper identified visceral involvement in almost all ferrets necropsied, and again several ferrets representing all age groups had larger blast-like variants. Peripheral lymphadenopathy was rare, and again a correlation of the blast form to young ferrets was absent.1 Therefore the age of ferrets cannot be reliably used to determine type, extent, or prognosis for lymphoma. The clinical presentation of ferrets with lymphoma will be nonspecific and most likely will vary with the organ system affected. Ferrets may present with varying degrees of lethargy, inappetance, weakness, diarrhea, dyspnea or respiratory signs, or they may be completely asymptomatic. In one author’s (NA’s) practice, lymphoma in 24% of the ferrets so diagnosed was an incidental finding during evaluation (surgery or ultrasound) for another disease process.
LABORATORY EVALUATION Anemia is the most consistent laboratory abnormality in ferrets with lymphoma.1 All the reported anemias were nonregenerative. Lymphocytosis and thrombocytopenia were extremely rare, and neutropenia was only occasionally identified.1 This indicates that results of CBCs and peripheral blood smears may yield valuable information in some cases but are rarely diagnostic for most cases of lymphoma. Persistently elevated lymphocyte counts cannot be used as evidence of lymphoma; as in other species, chronic smoldering infection is the most common cause of lymphocytosis in the ferret. The ubiquitous nature of Helicobacter and coronavirus infection in the U.S. ferret population has tremendous potential for inciting this nonspecific change in ferrets. Plasma biochemical data are also inconsistent in patients with lymphoma, with abnormalities usually relevant to the location of the disease or organ involvement. One study found hyperproteinemia and hyperglobulinemia rarely in ferrets (all with T-cell lymphoma) and hypoalbuminemia in a small
Radiography is necessary in ferrets suspected of having lymphoma, although it is not considered diagnostic. Evaluate radiographs for the presence of mediastinal masses or thoracic lymphadenopathy and pleural effusion as well as enlargement of liver, spleen, or kidneys. However, the absence of radiographic abnormalities does not rule out the possibility of lymphoma. Ultrasonography is perhaps the most valuable clinical tool available to most practitioners in evaluating ferrets for lymphoma. In addition to evaluating the abdominal and mesenteric lymph nodes, ultrasound also enables the clinician to assess the liver, spleen, kidneys, mediastinum, and sometimes even the gastrointestinal (GI) tract for infiltration. Figure 8-4 shows a ferret with infiltrated liver/spleen; Fig. 8-5 shows a ferret with mesenteric lymphadenopathy (Figs. 8-4 and 8-5). In one author’s (NA’s) practice, all ferrets with alimentary lymphoma had sonographic evidence of mesenteric lymphadenopathy (not all ferrets with mesenteric lymphadenopathy had lymphoma; this is an important difference!). It is important to recognize that mesenteric and intestinal lymph nodes in ferrets often appear sonographically more prominent than those in dogs and cats, but this does not specifically indicate lymphoma. Even severely enlarged lymph nodes may represent diseases other than lymphoma. One paper evaluated ultrasonographic characteristics of the normal mesenteric lymph node in ferrets, which is described as round to ovoid, measuring 12.6 ± 2.6 mm ´ 7.6 ± 2.0 mm and uniformly hyperechoic.45 Once again, though, the absence of abnormality on ultrasound does not eliminate lymphoma as a possible diagnosis.
CYTOLOGIC/HISTOLOGIC DESCRIPTION Histology or cytology is the only reliable tool with which to diagnose lymphoma. The definitive diagnosis of lymphoma is best accomplished by a pathologist experienced in the evaluation of ferret lymph nodes, as there is often great overlap between the histologic picture of lymphoma and other nonneoplastic causes of lymphadenomegaly. Biopsy (either needle or excisional) of gastric lymph nodes should also be avoided whenever possible, as chronic GI inflammation, a common problem in older ferrets, may yield reactive changes almost indistinguishable from lymphoma. Peripheral nodes, such as popliteal and scapular nodes, are less likely to be affected by local inflammation; excisional biopsy of these nodes is easily accomplished, and complications of this procedure are extremely rare (Fig. 8-6). In many cases, aspirates are performed as part of an initial examination, especially when clinical signs point strongly to lymphoma. Aspirates of enlarged nodes may result in a diagnosis in the hands of a trained pathologist or experienced practitioner, but false readings due to sample preparation, reactive changes, and well-differentiated neoplasms may occur. The possibility of false negatives is increased when aspirates of visceral organs are performed. Ultrasonographically obtained fine-needle aspirates of the mesenteric lymph nodes in normal ferrets yielded 50 to 60 small lymphocytes, 2 to 3 lymphoblasts and prolymphocytes, and 0 to 1 macrophages, plasma cells, and nondegenerate neutrophils per 200´ field.45 Eosinophils
CHAPTER 8 Neoplasia
A
B
SPLEEN
109
LIVER
Fig. 8-4 Ultrasound images from a ferret with lymphoma demonstrating infiltrative disease in the spleen (A) and liver (B). Note the mottled appearance and hypoechoic regions (arrows) in both tissues.
A
SUB LUMBAR LN
B
MESENTERIC LN
Fig. 8-5 Ultrasound images from a ferret with lymphoma with multiple enlarged lymph nodes in the abdomen. The sublumbar lymph node (closed arrow) measures 16.6 mm in diameter (A); the two mesenteric lymph nodes (arrowheads) measure 9.8 and 7.0 mm (B). The colon is evident above the sublumbar lymph node (open arrow).
(up to 10 per 200´ field) were also identified in some normal ferret mesenteric lymph node aspirates and did not correlate with peripheral blood eosinophilia.45 The cytologic hallmarks of lymphoma are a monotonous population of lymphocytes and the absence of peripheral blood elements. A range of cell sizes and types or the presence of other types of white blood cells in aspirated nodes is not consistent with a diagnosis of lymphoma. In cases of suspected leukemia, form a bone marrow aspiration via the proximal femur by using an 18- to 20-gauge bone marrow-collection needle. An alternate technique for obtaining a sample is a core bone biopsy, which may provide a better diagnostic yield (Fig. 8-7). In most cases of leukemia, the bone marrow is hypercellular and often
monomorphic. Microscopic examination reveals a significant decrease or total absence of normal marrow elements such as fat. Pathologists are commonly asked to evaluate splenic aspirates from animals with enlarged spleens. In our experience, >95% of these cases are the result of extramedullary hema topoiesis (EMH), a stereotypical response to chronic (GI) inflammatory diseases (see Chapter 36). Evidence of erythrocyte precursors, megakaryocytes, and abundant peripheral blood on splenic aspirates should lead the prudent practitioner to a diagnosis of EMH. Alternatively, cases of splenic lymphosarcoma are identified by the presence of a monomorphic population of cells with large nuclei, prominent nucleoli, and
110
SECTION I Ferrets In human medicine, there is a phase called consolidation therapy, in which the goal is to reduce the number of neoplastic cells within the body. This phase may last several months and often involves the introduction of new drugs to prevent resistance of neoplastic cells. It may be more aggressive than the induction phase and is often determined by the nature of the tumor. The third phase (often the second phase in animals) is a maintenance phase, where chemotherapy drugs are administered to destroy any residual cancer cells and prevent their multiplication. This is a less intensive protocol that may be continued for months or even years. Few or no side effects may be observed during maintenance chemotherapy. In patients that come out of remission, a rescue protocol, or rescue phase, may be initiated. This is a more aggressive combination of drugs, or a drug that is novel to that particular tumor type in that patient. It may be a single agent or a combination of drugs or treatment modalities. L-asparaginase, doxorubicin, and radiation therapy are some of the more commonly used rescue protocols in ferrets.
Adverse Effects and Precautions for Chemotherapy.
Fig. 8-6 Presentation of submandibular lymph node enlargement (arrow) in a ferret with lymphoma.
minimal cytoplasm as well as an absence of erythrocyte precursors and minimal peripheral blood elements. Additionally, mitotic figures should be present within the monomorphic cell population.
TREATMENT Many therapeutic options are available for the treatment of lymphoma in ferrets. Currently, there are no comparative studies of treatment protocols, so it is impossible to recommend one protocol over another based on survival, remission rate, side effects, or general efficacy. However, certain treatment options may be preferable for individual ferrets based on cost to owners, availability of drugs, or ease of administration. Although the goal of any protocol is to achieve complete remission of cancer, it may be more realistic in ferrets to obtain a “regression” of the tumor while maintaining a good quality of life for the pet. Even in cases where neoplasia is still evident at a cellular level, masses can shrink dramatically and hematologic values can return to normal for extended periods, enabling the ferret to return to normal activities and functions without compromising quality of life. If this goal is made clear to owners, survival times may be extended as expectations are for time and quality rather than complete cure.
Chemotherapy Chemotherapy refers to any drug or combination of drugs used to kill cancer cells. Even medications such as steroids may be considered chemotherapeutic agents when used for this purpose. In general, it is advisable to begin a regimen with multiple drugs at once so as to achieve maximum killing potential initially. This is called the induction phase, aimed at immediate rapid killing of tumor cells, and lasts approximately 4 weeks. The goal of the induction phase is to achieve remission. The most severe side effects usually accompany the induction phase, as there may be killing of other rapidly dividing normal cells (in the GI tract, skin, hair) in addition to tumor cells.
Although many people are familiar with the severe side effects associated with chemotherapy in humans, in general, side effects seem subjectively less severe in animals. Loss of fur and whiskers is common in ferrets but insignificant to the animal. Gastrointestinal effects do occur, so providing antinausea drugs such as metoclopramide, ondansetron, or maropitant citrate (Cerenia, Pfizer Animal Health, Pfizer Inc., New York, NY) is recommended. These drugs are preferred as they target the chemoreceptor trigger zone and peripheral receptors, which are the primary mediators of chemotherapy-induced nausea. GI protectants may also be beneficial. More serious adverse effects are cytopenias, particularly neutropenia, which predisposes patients to secondary infection and potential sepsis. Since the low end of normal WBCs in ferrets is lower than that in dogs and cats, it is necessary to redefine parameters for neutropenia that are more specific to ferrets. Any patient receiving chemotherapy who develops neutropenia (WBC < 1000/μL) should receive prophylactic antibiotics. If fever develops, hospitalize the patient immediately for intravenous antibiotic and fluid administration; this is a true oncologic emergency. Patients who are hospitalized under these circumstances generally respond well within 24 to 36 hours, while those that do not receive intravenous antibiotic therapy may suffer from more serious effects and prolonged recovery or even death. If either of these adverse effects occurs, separate the interval for administration (if multiple drugs were administered concurrently) to determine which drug is responsible for the adverse effect and decrease the dosage by 20% for future administrations. Use standard precautions when chemotherapeutic agents are being administered. Chemotherapy gloves are thicker than regular gloves and therefore provide extra protection against contact. If these are not available, double-gloving is recommended. Protective eyewear should be worn. In addition to syringes and needles, all materials that come in contact with chemotherapeutic agents (gloves, catheters) must be disposed of in appropriate biohazard containers. Certain medications (cyclophosphamide, chlorambucil) can be compounded into oral suspensions for more accurate dosing. These drugs should be stored in double plastic bags and appropriately labeled. Do not send compounded chemotherapeutic liquids home with owners, as spillage can occur and lead to potential exposure of owners, children, and other pets.
CHAPTER 8 Neoplasia
A
C
111
B
D Fig. 8-7 Standard core bone biopsies in larger ferrets are obtained
E
by a Michele trephine or Jamshidi bone biopsy instrument; these have beveled tips with which to remove the core once the instrument penetrates marrow (closed arrow). However, these instruments are too large for use in ferrets. Instead, samples can be obtained from the humerus or femur with a standard bone marrow biopsy instrument, but the technique must be modified because these lack beveled tips (open arrow) (A). After a stab incision is made in the skin, the bone marrow needle is introduced without the stylet. Both cortices of bone are penetrated and the needle is advanced through the skin on the opposite side of the limb (B). Without removing the needle, a collection cassette is placed at the tip of the needle; the stylet is then introduced (open arrowhead) to expel the core bone biopsy (closed arrowhead) (C, D). Performing this technique at an oblique angle minimizes the risk for fracturing the long bone (E). Save samples in 10% formalin.
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SECTION I Ferrets
Table 8-4 CP Protocol for Ferret Lymphoma Used at Gulf Coast Veterinary Specialistsa Case #: ___________ Stage: _____________ Grade: __________________ The following drugs are given sequentially for the first year as described in the flow sheet below: Elspar (L-asparaginase), 400 IU/kg IM (premedicate with diphenhydramine, 1 mg/kg IM) Prednisone (pred), 1 mg/kg PO Vincristine, 0.12 mg/kg IV Cytoxan, 10 mg/kg PO or SC Protocol flow sheet (provide dates for each therapy administered; week numbers may vary if neutropenic):
Week 0 1 2 3 4 7 10 13 16 19 22 25 28 31 34 37 40 43 46 49 52
Date
CBC
WBC
Neut
Lymph
PCV
(-3 day)
Elspar
Vincristine
Cytoxan
X X X X X X X X X X X X X X X X X X X Restage
X
Pred
Notes
X X X X X X X X X X X X X X X X X X X X X
X X X X X X X X X X X X X X X X
q12h q12h q24h q24h q24h q24h q48hb q48h q48h q48h q48h q48h q48h q48h q48h q48h q48h q48h q48h
CBC, complete blood cell count; WBC, white blood cell count; Neut, neutrophils; Lymph, lymphocytes; PCV, packed cell volume. aAdminister chemotherapy if neutrophil count ≥ 1000/μL; do not administer drugs if < 1000 neutrophils/μL (if so, delay treatment by 5 to 7 days, then repeat CBC). bBeginning at week 13, prednisolone can be decreased to q48h or continued at q24h, depending on individual patient’s clinical response.
When intravenous administration of chemotherapy drugs is necessary, a clean stick is imperative to prevent extravasation of the drug. Because a CBC is often required before administering chemotherapeutic drugs, collect a blood sample for the CBC from the jugular vein to preserve the peripheral veins for catheters. Sedate the ferret for placement of an intravenous catheter to deliver the chemotherapeutic agent; the catheter can be removed immediately after administration. Alternatively, a vascular access system can be surgically implanted and subsequently used for all intravenous administration without sedation. The use of these ports in ferrets has been well described.2,47 Protocols. Several protocols have been described for the treatment of lymphoma in ferrets. There are no studies comparing treatment modalities to date, and no evidence supports the use of one protocol over another. However, in humans as well as dogs and cats, combinations of cyclophosphamide, vincristine (Oncovin), and prednisone (COP) and/or COP with hydroxydaunorubicin (CHOP) represent the standard of care for lymphoma. In dogs, protocols containing doxorubicin result in the best clinical response and longest length of response.30,48 Table 8-4 provides a COP protocol that is used in one author’s (NA’s) practice. The “Wisconsin protocol,” which includes doxorubicin
and is used in dogs and cats, provides a more aggressive approach and can also be used. Tufts University has created a “no IV” protocol that enables the administration of multiple drugs without the need for vascular access (Table 8-5). Other chemotherapeutic protocols have also been used (Table 8-6). Table 8-7 provides a list of chemotherapeutic drugs reported in publications that have been used in ferrets; it is probable that still more drugs have been used; their dosages may be available on various oncology list-serves and discussion groups. Although many drug doses are calculated out on a “square meter” basis, there is a nominal difference between that measure and body weight in patients weighing less than 1 kg. A conversion table is provided in Table 8-8 for ease of dosing and calculation.
Palliative Therapy Palliative therapy refers to the partial treatment of disease or disease symptoms without aiming at cure. In cases where owners elect against chemotherapy, treatment with steroids is often beneficial in improving quality of life and shrinking tumor cells. Additional drugs such as cyclophosphamide or chlorambucil may be added, as there are minimal side effects at low doses or long intervals. Chlorambucil will only be helpful only for
CHAPTER 8 Neoplasia
113
Table 8-5 Tufts University “No-IV” Chemotherapy Protocol Week
Treatmenta,b
Weight (kg)
Week 1
L-Asp _____ Ctx _____ Pred _____
____
Week 2
L-Asp _____ CBCd _____
Week 3
L-Asp _____ Cytosar_____
Week
Treatmenta,b
Weight (kg)
Week 13
Ctx _____
____
Week 15
Pcb _____
____
Week 16
CBCd _____
____
Week 17
CBC _____
____
Week 18 Week 20
Ctx _____ Cytosar _____ Leuk _____ both drugs x 2 days
____ ____
Week 23
Ctx _____
____
Week 26
Pcb _____
____
Week 27
CBC _____ Chem _____
____
____ ____
Cytosar x 2 days
Week 4
CBCc _____
____
Week 5
Ctx _____
____
Week 7
Mtx _____ CBC _____
____
Week 8 Week 9
CBCc _____ Ctx _____
____ ____
Week 11
Cytosar _____ Leuk _____ both drugs x 2 days CBCc _____
____
If not in remission, continue weeks 20-26 for three cycles
Week 12 aPred,
prednisone (non); 2 mg/kg PO daily x 1 week then q48h. L-Asp, L-asparaginase (non); 10,000 IU/m2 SC. Ctx, cytoxan (mod); 250 mg/m2 PO; give with 50 mL/kg of LRS once. Cytosar, cytosar (mod); 300 mg/m2 SC x 2 days (dilute 100 mg with 1 mL sterile water). Mtx, methotrexate (mild); 0.8 mg/kg IM. Leuk, leukeran (mild); 1 tab PO (or ½ tablet daily for 2 days). Pcb, procarbazine (mild); 50 mg/m2 PO daily for 14 days. bOther abbreviations: CBC, complete blood cell count; chem, serum biochemical analysis. cDose reductions: if CBC indicates severe myelosuppression, reduce dosage by 25% for next treatment. Mod, moderately myelosuppressive; mild, mildly myelosuppressive; non, nonmyelosuppressive. Courtesy of Dr. Joerg Mayer, University of Georgia. Tufts staging protocol for ferret lymphoma ■ CBC and platelet count
■ Thoracic radiographs
■ Chemistry profile
■ Abdominal ultrasound
■ Freeze serum
■ UA (culture if indicated)
■ Bone marrow aspirate
■ Histopathology
slow-growing small-cell lymphomas. However, if owners choose this route, they must be advised that switching to a more aggressive therapy later on is less likely to provide a positive response because of the potential development of drug resistance.
Radiation Treatment Lymphoma is a tumor type that is highly responsive to radiation therapy. With the advent of more advanced equipment such as linear accelerators, radiation therapy offers a safe modality for either primary or adjunctive treatment. It is especially beneficial when there is one large tumor, either in the abdominal
or thoracic cavity, but can also be used on a single peripheral lymph node. While multiple treatments to a site are preferred, even a single dose of radiation will shrink most nodules and can greatly improve control of disease. This can be used as an initial therapy, for example, in a ferret with a mediastinal mass to alleviate immediate dyspnea or respiratory distress prior to or concurrent with the onset of chemotherapy. It can also be used as a rescue treatment when a solitary mass is present in the abdomen and not responding to chemotherapy. Treatment with radiation can shrink that mass and enable reestablishment of tumor control. Half- and total-body irradiation protocols are
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SECTION I Ferrets
Table 8-6 Chemotherapy Protocol for Lymphoma in Ferrets Week Drug
Dosage
1
Vincristine Asparaginase Prednisone
2 3 4-6
Cyclophosphamide Doxorubicin As weeks 1-3, but discontinue asparaginase Vincristine Cyclophosphamide Vincristine Methotrexate
0.07 mg/kg IV 400 IU/kg IP 1 mg/kg PO q24h and continued throughout therapy 10 mg/kg SC 1 mg/kg IV
8 10 12 14a
0.07 mg/kg IV 10 mg/kg SC 0.07 mg/kg IV 0.5 mg/kg IV
IV, intravenously; IP, intraperitoneally; PO, per os (orally); SC, subcutaneously. aProtocol is continued in sequence biweekly after week 14. From Rosenthal K. Ferrets. Vet Clin North Am Small Anim Pract. 1994;24:1-23.
used in other species and can be considered in ferrets, but there are no data available on such use in ferrets.
Ancillary Treatments Perhaps the most important ancillary treatment in any cancer patient is the provision of proper nutrition. Cancer cachexia is a well-documented paraneoplastic syndrome in all species. It consists of progressive involuntary weight loss despite adequate nutritional intake. Humans with this syndrome have a decreased response to treatment, diminished quality of life, and decreased survival time as compared with humans with the same disease who do not suffer from cancer cachexia.42 Maintaining protein and amino acid requirements improves immune response, GI function, and surgical healing and may also improve a cancer patient’s chances of remission. Alterations in lipid metabolism can lead to immunosuppression. Some tumor cells make poor use of lipid as a source of energy. This implies that a high-fat, low-carbohydrate diet may result in a greater chance of remission and a longer survival time. Omega-3 fatty acids have been shown to inhibit tumorigenesis in animal models.42 Most high-quality ferret diets are ideal because of their high protein and fat content.
Table 8-7 Drugs Published in the Literature and Used to Treat Tumors In Ferrets Drug
Dose
Notes/Precautions
Bleomycin
10 U/m2 SC26
Chlorambucil
1 mg/kg PO q7d31 20 mg/m2 PO47 200 mg/m2 PO, SC x 4 consecutive days weekly31 10 mg/kg PO2,8,65 80 mg/m2 PO q24h x 3 days q2wk47 20 mg/m2 IV31 2 mg/kg31 2.8 mg/kg IV q3wk x 3 doses47 30 mg/m2 IV47 2 mg/kg q24h PO50
Repeated dosages may cause pulmonary fibrosis
Cyclophosphamidea
Doxorubicinb
Isotretinoin L-asparaginaseb Methotrexate Prednisone/ prednisolone Vincristinec
aInjectable
400 IU/kg SC, IM31,68 5000 IU47 0.5 mg/kg IV8,47 2 mg/kg q24h28,65 1 mg/kg q48h19 20 mg/m2 q24h x 2 mon then q48h31 0.75 mg/m2 IV q7d31 2 mg/m2 IV31 0.12 mg/kg IV2,8 0.2 mg/kg IV68 0.5 mg/kg IV q7d19
High dose used for salvage; hemorrhagic cystitis developed
Cutaneous epitheliotropic lymphoma
Minimal myelosuppression Rescue protocol
cyclophosphamide can be administered orally at the same dose but may require dilution in propylene glycol for appropriate dosing. Alternatively, an oral formulation can be compounded by a professional compounding pharmacy. It should be administered in the hospital with proper precautions to avoid unnecessary human contact or risk with the use of a liquid chemotherapeutic. bPremedicate with diphenhydramine, 1 to 2 mg/kg, IV or IM, 30 minutes prior to administration to prevent anaphylactic response. cVincristine must be administered intravenously via a clean stick to avoid extravasation of the drug.
CHAPTER 8 Neoplasia Supplementation can also be provided with formulas available for ferrets (Oxbow Carnivore Care, Murdock, NE; Lafeber’s Emeraid Carnivore, Cornell, IL) or, in their absence, commercial gruel diets used for cats. Eukanuba Maximum-Calorie Formula (The Iams Company, Dayton, OH) is recommended
Table 8-8 Conversion Table (m2/kg) for Use in Ferrets Weight (kg)
BSA (m2)
0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8 1.9 2.0 2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8 2.9 3.0
0.034 0.045 0.054 0.063 0.071 0.079 0.086 0.093 0.100 0.107 0.113 0.119 0.125 0.131 0.137 0.142 0.148 0.153 0.159 0.164 0.169 0.174 0.179 0.184 0.189 0.194 0.199 0.203 0.208
m2, meters squared; BSA, body surface area. Courtesy of Dr. Joerg Mayer, University of Georgia.
A
115
over other canine/feline products because of its higher fat and protein content. Most ferrets can be encouraged to eat or are easily force-fed. Although clients often ask about holistic or herbal therapies, free radical scavengers, or immune stimulants, avoiding these products is recommended for patients receiving chemotherapy. The goal of chemotherapy is to suppress the immune system and often to induce cell death and free radicals; immune stimulants may interfere with this process. If necessary, one may consult with both a veterinary oncologist and a veterinarian trained in holistic/herbal medicine before initiating any alternative therapies. Lastly, any ferret that has received chemotherapy should not be vaccinated for the remainder of its life. Vaccines stimulate the immune system, which may cause the animal to come out of remission. If it is absolutely necessary to vaccinate, consider performing titers to determine whether vaccination is indicated. Other types of hematopoietic neoplasms, generally arising from cells of leukocytic lineage, are rarely seen. The spleen is the most common site of origin for these neoplasms. Myelolipoma, a benign neoplasm of immature leukocytes admixed with well-differentiated adipocytes, may occasionally present as a space-occupying mass in the spleen, but it is of no clinical significance.35 Thymoma, a neoplasm involving the epithelial and mature lymphocytic elements of the thymus, may present as a mass lesion of the anterior thorax and be easily confused for thymic lymphoma. An account of thymoma in two 5-yearold ferrets58 reported vomiting, lethargy, and dyspnea in both cases.
TUMORS OF THE SKIN AND SUBCUTIS The skin and subcutis are also common sites of neoplasia in ferrets, accounting for approximately 20% of cases in most reports.8,36,44,68 The vast majority of neoplasms of the skin are benign and most are primary neoplasms.68 Benign tumors of basal cell origin, including sebaceous adenoma and sebaceous epithelioma, are the most common skin neoplasm in the ferret (Fig. 8-8). These warty exophytic neoplasms, which may attain a large size and ominous appearance
B
Fig. 8-8 Multiple sebaceous adenomas in a ferret on the thorax (A) and face (B). The neoplasm on the face did not involve the orbit. Occasionally, the centers of these neoplasms may be cavitated (arrow). Although these lesions were impressive in appearance, surgical removal was curative.
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(largely as a result of self-trauma), are almost invariably benign. They occasionally cause irritation, and the resulting self-trauma may result in local inflammation and infection. In extremely rare cases, these tumors may give rise to squamous cell carcinoma.68 Surgical excision is curative and should be accomplished early. Mast cell tumors are the second most common skin tumors in the ferret. Unlike such tumors in the dog and cat, mast cell tumors are universally benign in the ferret and warrant a good prognosis. Upon gross inspection, these neoplasms are flat and discrete, with a crusty yellow appearance. It is common for them to bleed and then form scabs, which subsequently fall off to reveal the familiar yellow crusty or slightly raised appearance until the cycle repeats. Most mast cell tumors in ferrets show minimal infiltration into the dermis and are easily excised, which is curative. Malignancy and metastatic disease have not been reported in cutaneous mast cell tumors in ferrets. A number of ferrets, however, may show multicentric development of mast cell tumors over time and require additional surgeries, but this finding is not of any adverse prognostic significance. Neoplasms of apocrine scent glands are the third most common neoplasms seen in the skin and subcutis in the ferret. Unlike the previously described neoplasms, tumors of apocrine glands are largely restricted to the deeper layers of the skin and subcutis and have a distinct predilection for malignant behavior. Apocrine neoplasms are most often seen in areas where scent glands are concentrated (head, neck, prepuce, and vulva). Approximately 75% of preputial neoplasms of apocrine origin (as well as 100% of the less common perianal and perivulvar tumors) are malignant, exhibiting aggressive infiltration of local tissues, metastasis to local nodes, and occasionally pulmonary metastasis. Complete surgical excision of apocrine malignancies is difficult because of their rapid and aggressive growth as well as the distinct possibility of presurgical metastasis. Excise any suspected apocrine neoplasms with wide surgical margins to minimize the potential for metastasis. Use advancement flaps or y-plasty, if necessary, for closure or reconstruction of the prepuce. In cases of apocrine carcinoma of the prepuce, appropriate surgical treatment may entail amputation of the prepuce and urethrostomy. Submit all excised tissues for evaluation of surgical margins. Radiation therapy may be a valuable postoperative treatment to aid in eliminating any local residual malignant cells, and chemotherapy can be used as an adjunct to decrease the risk of metastasis. Vascular neoplasms of the skin and subcutis are occasionally seen in the ferret. Histologically, these neoplasms are fairly evenly divided between histologically benign and malignant,68 but all neoplasms were cured after complete surgical excision. Coat color and pigmentation does not appear to play a significant role in the development of this neoplasm (as it does in other domestic species), likely because most pet ferrets in North America are exclusively indoor animals. Low-grade subcutaneous sarcomas are also occasionally seen in the subcutaneous tissues of the ferret. Most of these neoplasms are of smooth muscle origin and arise along the dorsal midline. Many of these neoplasms arise from smooth muscle (erector pili) associated with hair follicles (piloleiomyosarcomas).49 Cutaneous fibrosarcomas may also be seen in the skin of ferrets68 and have been reported in association with vaccination in this species.40 (However, these neoplasms do not manifest the aggressive behavior associated with vaccine-related sarcomas in the cat.40) Regardless of their origin, subcutaneous
sarcomas are generally low-grade malignancies with a slow rate of growth and low metastatic potential; they respond well to wide surgical excision. Mammary gland neoplasms are rarely seen in domestic ferrets. A previous report of 6 cases (2 simple, 4 complex) comprised only benign mammary neoplasms.68 Three cases of simple mammary hyperplasia have also been reported,38 two of which were seen in conjunction with adrenal carcinoma. Other benign neoplasms seen in the skin of ferrets include lipomas, squamous papillomas, and tumors of sebaceous or eccrine sweat glands.68 Malignant neoplasms include epitheliotropic lymphoma (discussed under Tumors of the Hemolymphatic System, above), squamous cell carcinoma (which may occasionally arise from the lining of the anal sac), and rare neoplasms such as ceruminous gland adenocarcinoma of the ear. An interesting cutaneous tumor of uncertain etiology has been reported in the literature and seen several times by one author (BW). Subcutaneous neoplasms of the ventral abdomen of ferrets with marked morphologic and immunohistochemical similarities to adrenocortical tumors have been reported in ferrets.56 In each case, the animal did not display systemic signs associated with endocrinopathy, and a primary adrenocortical tumor could not be identified. General guidelines for treating cutaneous neoplasms in ferrets are similar to those prescribed for more traditional pet species. Early surgical intervention is the rule with cutaneous neoplasms; most neoplasms are benign, and most malignancies are of low grade and can be successfully treated with early surgical excision with wide margins. Submit all neoplasms for histopathologic evaluation to provide an accurate diagnosis as well as recommendations for additional treatment, if any. Surgically excise of all preputial or perivulvar/perianal neoplasms as early as possible, after careful palpation and radiography to minimize the opportunity for metastasis.
TUMORS OF THE GASTROINTESTINAL TRACT Neoplasms of the gastrointestinal tract are commonly seen in the ferret. The liver is a particularly common site for metastasis, most commonly for malignant lymphoma, adrenocortical carcinoma, and a number of poorly differentiated neoplasms in which the primary neoplasm cannot be identified.68 The liver is also a relatively common site for the development of primary neoplasms. The most common of these by far is biliary cystadenoma/cholangioma; however, cholangiocarcinoma, hepatocellular carcinoma, and hepatoma are also observed.68 In most cases animals with hepatic neoplasia are presented for nonspecific weight loss, anorexia, and lethargy, but some may be asymptomatic. A cranial abdominal mass is generally identified by palpation or radiography. Elevations in alanine transaminase (ALT) or alkaline phosphatase (ALP) concentrations may be present, but clinicopathologic abnormalities are usually mild and nonspecific. Infiltrative or nodular disease is readily apparent ultrasonographically. The diagnosis of biliary cystadenoma is especially important in this species because of its predilection to exhibit malignant behavior (replacing one or more lobes of the liver and ultimately resulting in hepatic failure) (Fig. 8-9). The differentiation of biliary cystadenoma from biliary cyst (a common incidental finding in this species) is made on the basis of one or more of the following factors: the presence of clinical
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Fig. 8-10 Mandibular squamous cell carcinoma in a ferret. There Fig. 8-9 Biliary cystadenoma in a ferret. Because of their aggressive nature, these histologically benign tumors (arrows) are best treated with lobectomy or, at a minimum, wide surgical excision.
symptoms, abnormalities in liver-specific clinical pathology, or expansive growth over time documented by abdominal ultrasound. Hepatic carcinoma and cholangiocarcinoma will result in elevated hepatic enzymes, eventual hepatic failure, and other signs, such as profound anemia, hemoperitoneum, anemia, and ascites. Biopsies or aspirates should be obtained from all hepatic neoplasms, especially those involving multiple lobes. Ultrasound-guided aspirates can reveal a diagnosis in many cases. Pretreatment with vitamin K, whenever possible, can help to minimize the risk of bleeding. If the neoplasm is confined to one lobe of the liver, lobectomy is recommended. Lobectomy of two lobes is possible but may lead to further compromise of the patient. Because of the aggressive nature of biliary cystadenoma in this species, any cystic lesion of the liver should be removed with wide surgical margins or lobectomy. Neoplasms involving multiple lobes warrant a poor long-term prognosis; however, survival times of several months or more may be seen with hepatocellular carcinoma in ferrets. Animals possessing malignancies of the biliary system generally succumb within a short time frame. Neoplasms of the exocrine pancreas are occasionally seen in the ferret. Most neoplasms exhibit aggressive growth into the surrounding pancreas with seeding of the abdominal cavity as well as metastasis to additional organs.29,63 Complete surgical excision of these tumors is unlikely before the onset of extension into other areas of the abdominal cavity. The most common neoplasm affecting the GI tract is malignant lymphoma. This is understandable considering that the GI tract contains over half of all lymphoid tissue in the body of ferrets as well as the prevalence of chronic inflammatory disease in the stomach and intestine. Of the cases in which the GI tract was considered the primary site of origin, the intestine was the most common site, followed by the stomach, liver, colon, and, last, the oral cavity. Lymphoma of the intestine is considered to carry an extremely poor prognosis because of the disruption of the intestinal barrier and absorption of toxins from the GI tract and may be refractory to treatment.8 However, the incidence of intestinal lymphoma may be even greater than reported, as
is marked invasion of alveolar bone with tooth loss (arrow).
many patients with variable GI signs do not undergo biopsy and are undiagnosed. In many cases, steroids are likely used to alleviate clinical signs without biopsy, which may lead to incorrect assumptions regarding potential survival times and responses to therapy. Primary neoplasms of the gastrointestinal tract itself tend to be malignant, with adenocarcinomas arising primarily in the stomach and intestine. These neoplasms are locally aggressive, often infiltrating multiple layers of the wall with metastasis to local lymph nodes. The tendency of intestinal adenocarcinoma to incite a prominent scirrhous response often results in obstruction (as opposed to intestinal lymphomas, which do not result in a scirrhous response) and thus, clinical symptoms. This same scirrhous response, however, tends to achieve a type of containment of the neoplasm, allowing visualization of the tumor’s margins and facilitating complete excision. The prognosis at this point is heavily influenced by the presence or absence of presurgical metastasis. These masses may be identified by palpation, ultrasound, and/or a barium GI series. A complete resection and anastomosis should be performed whenever solitary intestinal masses are present without gross evidence of metastatic disease. In ferrets as opposed to other domestic species, smooth muscle tumors of the gastrointestinal tract, gastrointestinal stromal tumors, and neuroendocrine tumors are very uncommon. Tumors of the oral cavity are occasionally seen in ferrets and are usually associated with a poor prognosis. Squamous cell carcinoma, the most common of these, is an aggressive neoplasm of the gums that invades underlying bone, resulting in tooth loss, disfigurement, and inappetence (Fig. 8-10). There is one report of treatment of a mandibular squamous cell carcinoma with bleomycin at a dose of 20 U/m2, which reduced tumor mass.26 Surgical excision, if attempted, should be performed early and with wide surgical margins; a recent report combined rostal maxillectomy with radiation therapy as a treatment option.25 Intralesional chemotherapy may also be attempted; follow current recommendations for companion animals. Early, aggressive treatment provides the best opportunity for resolution of squamous cell carcinoma. Various sarcomas, including fibrosarcoma, have been reported in the oral cavity and respond poorly to all forms of treatment.8
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TUMORS OF THE REPRODUCTIVE TRACT Because of the prevalence of neutering among American pet ferrets, tumors of the reproductive system are rarely seen in clinical practice. While earlier reports indicated a high prevalence of these neoplasms,4,14 the rarity of intact animals in the current pet and laboratory populations of ferrets has significantly reduced the numbers of the tumors seen today. Clinical signs of reproductive neoplasia in ferrets are variable and often nonspecific. However, with rare exceptions (as noted below), surgical excision of affected gonads is curative. In spayed females showing signs of adrenal disease at very young ages or that are minimally responsive to medical management, consider residual reproductive tissue or, less likely, reproductive neoplasia, as a differential diagnosis, since ovarian remnants and stump granulomas are still possible sources of androgen production. Ultrasound examination of these animals will serve to identify residual reproductive tissue. Most neoplasms of the ferret uterus are of smooth muscle origin. While approximately 75% are malignant based on histologic appearance,68 metastatic disease has not been reported and surgical excision is curative. Nonmuscular tumors of the uterine tube are extremely rare in the ferret. One uterine adenocarcinoma and one deciduoma have been seen by one author (BW). Regressing implantation sites in female ferrets may be mistaken for uterine carcinoma by histologic examination as a result of the profound atypia of maternal presymplasmal cells. Ovarian tumors often result in no overt signs; in a few cases, affected animals exhibit reproductive failure. Ovarian tumors (granulosa cell tumors, teratomas, Leydig cell tumors, and sex cord-stromal tumors) have been reported, often as incidental findings during routine spays. Teratomas may attain a size that is obvious on routine palpation or may be identified via survey radiography as a result of the presence of bone within the tumor mass. One reported Leydig cell tumor metastasized to a regional lymph node.68 Testicular neoplasms are most commonly seen in retained testes—a finding similar to observations reported in other domestic species. Affected males may show signs of hyperestrogenism, including intact sexual behavior, aggression, a prominent musky odor, and a poor greasy hair coat. Multiple neoplasms may be seen in retained testes; in one testis, four distinct neoplasms (interstitial cell, seminoma, Sertoli cell, and a carcinoma of the rete testis) were seen.68 One Sertoli cell tumor metastasized to the liver.68
TUMORS OF THE MUSCULOSKELETAL SYSTEM Neoplasms of the skeletal system are occasionally seen in ferrets and generally result in a clinical appearance that is obvious to both owner and practitioner. Tumors of the skeletal muscles, however, are extremely rare.68 Chordomas are the most common neoplasm of the musculoskeletal system in the ferret. They most commonly appear as irregularly round, whitish gray, firm, club-like swellings of the tail tip. This low-grade malignancy (arising from primitive notochord) is most commonly seen at the tip of the tail but may arise in a vertebra in any region of the spinal column.15 These neoplasms are locally aggressive, destroy the vertebral body in which they arise, but have minimal metastatic potential (with only one report of metastasis, following surgical intervention).66 Radiography of affected vertebrae reveal a focally extensive vertebral lesion that is both lytic and proliferative.
Chordomas of the tail tip may be easily cured by amputation, but they carry a poor prognosis when they affect other parts of the spinal column. Because of their aggressive nature, extirpation of a chordoma from affected vertebrae is usually not feasible and eventual loss of function and pathologic fracture will result. This neoplasm is occasionally misdiagnosed as chondrosarcoma by pathologists unfamiliar with ferret tissues. True tumors of bone (osteomas and osteosarcomas) are occasionally seen in the ferret. Osteomas most commonly arise on flat bones, including the skull and ribs, and progress slowly. Surgical removal may occasionally be accomplished; however, many osteomas regrow when excision is incomplete. Osteosarcomas are rarely reported in the ferret64 and may arise on either flat or long bones. Amputate the affected limb if possible, as these malignancies are locally destructive. Although there are no published follow-up data for this disease in ferrets, metastasis is not reported and has not been seen by either of the present authors; therefore the prognosis for osteosarcoma may be better than in dogs. Surgeons are cautioned that noncore biopsies of malignant bone tumors may result in an errant diagnosis due to the presence of pronounced periosteal reactions overlying the osteosarcomas itself. Tumors of skeletal muscle are extremely rare in the ferret. Rhabdomyosarcomas—malignant tumors of skeletal muscle— have been reported.36,68 These neoplasms are treatable by radical excision or amputation if present on the limbs.
TUMORS OF THE NERVOUS SYSTEM Neoplasms of the nervous system are rare in ferrets, accounting for fewer than 0.5% of reported tumors.68 These tumors are equally divided between those of the central nervous system (affecting the brain and spinal cord) and those of the peripheral nervous system (affecting the peripheral nerves and ganglia). Tumors of the central nervous system (CNS) generally lead to the development of neurologic signs including ataxia and seizures, while those of the peripheral nervous system result in body-surface masses that owners usually notice before any neurologic signs develop. When the cause of neurologic signs in ferrets is being considered, intracranial tumors are a very unlikely cause, ranking as only the third most common cause of neurologic signs. Hypoglycemia due to insulinoma is by far the most common cause of neurologic signs in the ferret, followed (at a distance) by bacterial infections of the CNS. Because of their rarity, intracranial tumors should be considered only when these two previously mentioned syndromes are conclusively ruled out. Clinical signs associated with CNS tumors are quite variable and often nonspecific. Lateralizing signs—such as turning toward the side of the lesion, ataxia, cranial nerve deficits, normocellular cerebrospinal fluid, and uncontrolled seizure activity in the presence of normal blood glucose—are suggestive of a CNS neoplasm though not specific for one. There are currently no reports of successful treatment of CNS tumors. The most common neoplasm of the CNS in the ferret is malignant lymphoma, which may, of course, appear in any organ of the body. (This is also the most common neoplasm of the eye.) Primary brain tumors are rare and usually result in severe neurologic deficits over time. Astrocytomas appear to be the most common primary brain tumors, followed by granular cell tumors,55 meningiomas,36,68 primitive neuroendocrine tumors, and a choroid plexus papilloma.59 Of all of the primary
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B
Fig. 8-11 Preoperative (A) and postoperative (B) images of a malignant peripheral nerve sheath tumor (arrow) in a ferret. Removal was achieved with cryosurgery; however, facial nerve paralysis was encountered after surgery. (Courtesy Dr. Darrell Kraft, Pet’s Choice Animal Hospital, Woodinville, WA.)
brain tumors, meningiomas show the most promise for surgical excision because they are discrete neoplasms arising from the meninges and, in this limited number of cases, did not infiltrate the neuropil. Antemortem diagnosis of meningioma is a significant challenge facing practitioners. Computed tomography or magnetic resonance imaging will detect the presence of the tumor; meningioma should be suspected when any tumor is located along the meninges. Neoplasms of the peripheral nervous system, however, carry a significantly improved prognosis over those in the CNS, as they tend to be restricted to the skin and subcutis. Prognosis is based on the degree of malignancy and infiltration of local tissue. In one collection, both benign and malignant peripheral nerve sheath neoplasms were identified.68 Malignant peripheral nerve sheath tumors as a general rule exhibit rapid growth and tend to infiltrate adjacent tissue to a higher degree than their benign counterparts, rendering complete excision more difficult (Fig. 8-11). In many cases, repeat surgeries are required to effect a cure. While these neoplasms may be seen at any site in the body, the tissues of the head (and, interestingly, the eyelid) appear to be a common site of origin. Tumors of nerve sheath origin may be misdiagnosed as fibrosarcoma or leiomyosarcoma when immunohistochemical procedures are not used, with the prognosis of leiomyosarcoma being significantly improved over either peripheral nerve sheath tumors or fibrosarcomas. Perform wide surgical excision of these tumors as soon as possible after diagnosis, as growth in areas with high skin tension may result in large defects that are difficult to close. Radiation therapy may help minimize or prevent recurrence at the surgical site or may be used to reduce tumor size prior to surgery. Ganglioneuromas are rare neoplasms of the peripheral nerve ganglia.36,68 These well-differentiated neoplasms with neurons and glia in a matrix of neural tissue are reported in close proximity to the right adrenal gland. Close examination is required to differentiate these nodules from normal ganglia on a histologic basis; however, these tumors tend to be much larger than ganglia, ranging up to 1.5 cm in diameter. These tumors have no apparent clinical signs and on gross inspection are often misjudged to be adrenal tumors.
TUMORS OF THE URINARY SYSTEM Neoplasms involving the urinary system are rare in ferrets. Transitional cell carcinoma of the kidney is the most common; these neoplasms have also been reported in the urinary bladder.8 In the kidney, transitional cell carcinomas arise in the renal pelvis,5
eventually causing outflow obstruction and hydronephrosis. Metastasis has not been reported from this site; unilateral nephrectomy may be curative if early diagnosis is achieved. In the bladder, transitional cell carcinoma is generally associated with a poor prognosis. Because the presenting signs are vague, diagnosis is generally achieved only after extensive local invasion has occurred.8 Dysuria and incontinence may be presenting signs initially ascribed to cystic prostatic disease or crystalluria. Urinalysis, including the examination of urinary sediment, and contrast radiographic techniques may be helpful in identifying this neoplasm; definitive diagnosis is made by surgical biopsy.8 It is likely that these tumors, once identified, would be a surgical challenge, especially in the area of the trigone. For unresectable tumors, chemotherapeutic agents that inhibit COX-2 enzymes have shown promise in dogs and may ameliorate clinical signs and prolong life in ferrets; other more traditional agents such as doxorubicin, cisplatin, and cyclophosphamide may also be useful. However, appropriate dosages of all of these agents for the treatment of this and other types of invasive carcinomas have not been defined. Renal carcinomas and adenomas have also been reported in ferrets.8,32,36 These unilateral neoplasms of the kidney are most often encountered at necropsy, as most tend to be slow-growing with low metastatic potential. On ultrasound examination, renal neoplasms generally present as cystic areas; however, the high incidence of renal cysts in domestic ferrets would likely preclude further diagnostic workup based on this finding. Occasionally renal carcinoma may result in hemoperitoneum and require emergency nephrectomy.
TUMORS OF THE RESPIRATORY SYSTEM Neoplasms involving the lung are generally of metastatic origin, although one undescribed primary neoplasm of the lung has been reported.36 The lung is a common site for the metastasis of lymphoma; clinical signs may include marked pulmonary edema or effusion, which may be significant enough to mask the radiologic signs associated with a disseminated tumor. Pulmonary metastasis of other neoplasms would likely go unnoticed in most cases. Chemotherapy may be of benefit in metastatic lymphoma.
VASCULAR NEOPLASMS Hemangiomas and hemangiosarcomas are occasionally seen in the domestic ferret. Most arise in the skin or subcutis,68 although endothelial neoplasms are also seen in the liver,
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spleen, pancreas, lymph nodes, and free-floating in the abdomen. While over half of cutaneous vascular neoplasms exhibit histologic evidence of malignancy, metastasis is not seen.68 Even though most cutaneous vascular neoplasms are malignant, they are low-grade malignancies with slow growth and no metastatic potential. Complete excision of these tumors is considered curative. Rarely, multiple cutaneous hemangiosarcomas may be seen; however, the prognosis for these cases is no different than for animals with single neoplasms. However, the prognosis for animals with visceral hemangiosarcoma is significantly worse. These tumors tend to grow more aggressively within abdominal organs and may rupture at any time, seeding the abdomen with metastatic tumors12 or resulting in fatal hemorrhage. A guarded prognosis for animals with visceral hemangiosarcomas should be offered in all cases and early surgical intervention should be the rule when the neoplasm is restricted to a single site.12 An incidence of 21.7% of hepatic hemangiosarcoma was reported in one colony;11 the cause of this high frequency is uncertain, and this phenomenon has not been reported since.
MISCELLANEOUS NEOPLASMS Mesotheliomas are uncommon malignancies of ferrets that bode extremely poorly for any affected animal.67 These tumors arise in the abdominal cavity and spread extensively before the appearance of clinical signs. The most common clinical sign in affected animals is profound ascites (“malignant” ascites).67 Perform abdominocentesis in such cases; identification of rafts of atypical mesothelial cells may accomplish diagnosis. As normal mesothelial cells may be seen in any abdominal tap, exercise care to avoid misdiagnosis. Anaplastic neoplasms are those in which the level of cellular differentiation is below that needed to identify a cell of origin by its microscopic resemblance to normal tissue. Sophisticated techniques may yield clues to a tumor cell’s origin even if it does not resemble the parent tissue. Immunohistochemical procedures to identify tissue specific intermediate filaments or ultrastructural analysis of cellular organelles by electron microscopy may identify characteristic organelles for a particular cell type. Today, the use of these advanced techniques at large referral laboratories lends insight into the origin of a particular neoplasm; in doing so, it provides important information concerning therapeutic approaches and prognosis. However, many smaller laboratories are not equipped to perform these tests routinely, and a broad diagnosis of “poorly differentiated” carcinoma, sarcoma, or round cell tumor is often the result. In the collection of over 1600 ferret neoplasms at the Armed Forces Institute of Pathology and following advanced testing, the tissue of origin could not be identified in only 2% of cases; however, in 80% of these cases, a broad category of epithelial versus mesenchymal origin was obtainable.68 Even this limited classification has therapeutic importance, as epithelial and round cell tumors tend to be significantly more responsive to chemotherapy than the sarcomas. Sarcomas of the skin were the largest single classification of poorly differentiated tumors but likely the most responsive to treatment (i.e., surgery). As sarcomas of the skin tend to have low metastatic potential, a definitive identification of cell of origin (smooth muscle, skeletal muscle, fibrocyte, etc.) is of little clinical importance. However, the remainder of the poorly differentiated neoplasms generally carry a poor prognosis, especially those present in abdominal organs.
References 1. Ammersbach M, DeLay J, Caswell JL, et al. Laboratory findings, histopathology, and immunophenotype of lymphoma in domestic ferrets. Vet Pathol. 2008;45:663-673. 2. Antinoff N, Hahn KA. Ferret oncology: diseases, diagnostics, and therapeutics. Vet Clin North Am Exot Anim Pract. 2004;7:579-625. 3. Batchelder MA, Erdman SE, Li X, et al. A cluster of cases of juvenile mediastinal lymphoma in a ferret colony. Lab Anim Sci. 1996;46:271-274. 4. Beach JE, Greenwood B. Spontaneous neoplasia in the ferret (Mustela putorius furo). J Comp Pathol. 1993;108:133-147. 5. Bell RC, Moeller RB. Transitional cell carcinoma of the renal pelvis in a ferret. Lab Anim Sci. 1990;40:537-538. 6. Besso J, Tidwell A, Gliatto J. Retrospective review of the ultrasonographic features of adrenal lesions in 21 ferrets. Vet Radiol Ultrasound. 2000;41:345-352. 7. Bielinska M, Parviainen H, Kiiveri S. Review paper: origin and molecular pathology of adrenocortical neoplasms. Vet Pathol. 2009;46:194-210. 8. Brown SA. Neoplasia. In: Quesenberry KE, Hillyer E, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders; 1997:98-114. 9. Caplan ER, Peterson ME, Mullen HS, et al. Diagnosis and treatment of insulin-secreting pancreatic islet cell tumors in ferrets: 57 cases (1986-1994). J Am Vet Med Assoc. 1996;209:1741-1745. 10. Carpenter JW, Finnegan MV. A ferret with a midventral cervical mass. Vet Med. 2003;98:380-383. 11. Cross BM. Hepatic vascular neoplasms in a colony of ferrets. Vet Pathol. 1987;24:94-96. 12. Darby C, Ntavlourou V. Hepatic hemangiosarcoma in two ferrets (Mustela putorius furo). Vet Clin North Am Exot Anim Pract. 2006;9:689-694. 13. Desmarchelier M, Lair S, Dunn M, et al. Primary hyperaldosteronism in a domestic ferret with an adrenocortical adenoma. J Am Vet Med Assoc. 2008;233:1297-1301. 14. Dillberger JE, Altman NH. Neoplasia in ferrets: eleven cases with a review. J Comp Pathol. 1989;100:161-176. 15. Dunn DG, Harris RK, Meis JM, et al. A histomorphologic and immunohistochemical study of chordoma in twenty ferrets (Mustela putorius furo). Vet Pathol. 1991;28:467-473. 16. Erdman SE, Brown SA, Kawasaki TA, et al. Clinical and pathologic findings in ferrets with lymphoma: 60 cases (1982-1994). J Am Vet Med Assoc. 1996;208:1285-1289. 17. Erdman SE, Correa P, Coleman LA, et al. Helicobacter mustelae-associated gastric MALT lymphoma in ferrets. Am J Pathol. 1997;151:273-280. 18. Erdman SE, Kanki PJ, Moore FM, et al. Clusters of lymphoma in ferrets. Cancer Invest. 1996;14:225-230. 19. Erdman SE, Moore FM, Rose R, et al. Malignant lymphoma in ferrets: clinical and pathological findings in 19 cases. J Comp Pathol. 1992;106:37-47. 20. Erdman SE, Reimann KA, Moore FM, et al. Transmission of a chronic lymphoproliferative syndrome in ferrets. Lab Invest. 1995;72:539-546. 21. Fox JG, Dangler CA, Sager W, et al. Helicobacter mustelae- associated gastric adenocarcinoma in ferrets (Mustela putorius furo). Vet Pathol. 1997;34:225-229. 22. Fox JG, Dangler CA, Snyder SB, et al. C-cell carcinoma (medullary thyroid carcinoma) associated with multiple endocrine neoplasms in a ferret (Mustela putorius). Vet Pathol. 2000;37:278-282. 23. Fox JG, Wishnok JS, Murphy JC, et al. MNNG-induced gastric carcinoma in ferrets infected with Helicobacter mustelae. Carcinogenesis. 1993;14:1957-1961.
CHAPTER 8 Neoplasia 24. Gliatto JM, Alroy J, Schelling SH, et al. A light microscopical, ultrastructural and immunohistochemical study of spindle-cell adrenocortical tumours of ferret. J Comp Pathol. 1995;113:175-183. 25. Graham J, Fidel J, Mison M. Rostral maxillectomy and radiation therapy to manage squamous cell carcinoma in a ferret. Vet Clin North Am Exot Anim Pract. 2006;9:701-706. 26. Hamilton TA, Morrison WB. Bleomycin chemotherapy for metastatic squamous cell carcinoma in a ferret. J Am Vet Med Assoc. 1991;198:107-108. 27. Hammer AS, Williams B, Dietz HH, et al. High-throughput immunophenotyping of 43 ferret lymphomas using tissue microarray technology. Vet Pathol. 2007;44:196-203. 28. Hanley CS, Wilson GH, Frank P, et al. T cell lymphoma in the lumbar spine of a domestic ferret (Mustela putorius furo). Vet Rec. 2004;155:329-332. 29. Hoefer HL, Patnaik AK, Lewis AD. Pancreatic adenocarcinoma with metastasis in two ferrets. J Am Vet Med Assoc. 1992;201:466-467. 30. Hosoya K, Kisseberth WC, Lord LK, et al. Comparison of COAP and UW-19 protocols for dogs with multicentric lymphoma. J Vet Intern Med. 2007;21:1355-1363. 31. Hutson CA, Kopit MJ, Walder EJ. Combination doxorubicin and orthovoltage radiation therapy, single-agent doxorubicin, and high-dose vincristine for salvage therapy of ferret lymphosarcoma. J Am Anim Hosp Assoc. 1992;28:365-368. 32. Kawaguchi H, Miyoshi N, Souda M, et al. Renal adenocarcinoma in a ferret. Vet Pathol. 2006;43:353-356. 33. Lee A. Helicobacter infections in laboratory animals: a model for gastric neoplasias? Ann Med. 1995;27:575-582. 34. Li X, Fox JG, Erdman SE, et al. Cutaneous lymphoma in a ferret (Mustela putorius furo). Vet Pathol. 1995;32:55-56. 35. Li X, Fox JG, Erdman SE. Multiple splenic myelolipomas in a ferret (Mustela putorius furo). Lab Anim Sci. 1996;46: 101-103. 36. Li X, Fox JG, Padrid PA. Neoplastic diseases in ferrets: 574 cases (1968-1997). J Am Vet Med Assoc. 1998;212:1402-1406. 37. Miwa Y, Kurosawa A, Ogawa H, et al. Neoplastic diseases in ferrets in Japan: a questionnaire study for 2000 to 2005. J Vet Med Sci. 2009;71:397-402. 38. Mor N, Qualls Jr CW, Hoover JP. Concurrent mammary gland hyperplasia and adrenocortical carcinoma in a domestic ferret. J Am Vet Med Assoc. 1992;201:1911-1912. 39. Morrison WB. Lymphoma in dogs and cats. Jackson, WY: Teton NewMedia; 2005. 40. Munday JS, Stedman NL, Richey LJ. Histology and immunohistochemistry of seven ferret vaccination-site fibrosarcomas. Vet Pathol. 2003;40:288-293. 41. Newman SJ, Bergman PJ, Williams B, et al. Characterization of spindle cell component of ferret (Mustela putorius furo) adrenal cortical neoplasms, correlation to clinical parameters and prognosis. Vet Comp Oncol. 2004;2:113-124. 42. Ogilvie GK, Moore AS. Managing the veterinary cancer patient: a practice manual. Trenton, NJ: Veterinary Learning Systems Co; 1995. 43. Onuma M, Kondo H, Ono S, et al. Cytomorphological and immunohistochemical features of lymphoma in ferrets. J Vet Med Sci. 2008;70:893-898. 44. Parker GA, Picut CA. Histopathologic features and post-surgical sequelae of 57 cutaneous neoplasms in ferrets (Mustela putorius furo L.). Vet Pathol. 1993;30:499-504. 45. Paul-Murphy J, O’Brien RT, Spaeth A, et al. Ultrasonography and fine needle aspirate cytology of the mesenteric lymph node in normal domestic ferrets (Mustela putorius furo). Vet Radiol Ultrasound. 1999;40:308-310. 46. Peterson 2nd RA, Kiupel M, Capen CC. Adrenal cortical carcinomas with myxoid differentiation in the domestic ferret (Mustela putorius furo). Vet Pathol. 2003;40:136-142.
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47. Rassnick KM, Gould 3rd WJ, Flanders JA. Use of a vascular access system for administration of chemotherapeutic agents to a ferret with lymphoma. J Am Vet Med Assoc. 1995;206:500-504. 48. Rassnick KM, McEntee MC, Erb HN, et al. Comparison of 3 protocols for treatment after induction of remission in dogs with lymphoma. J Vet Intern Med. 2007;21:1364-1373. 49. Rickman BH, Craig LE, Goldschmidt MH. Piloleiomyosarcoma in seven ferrets. Vet Pathol. 2001;38:710-711. 50. Rosenbaum MR, Affolter VK, Usborne AL, et al. Cutaneous epitheliotropic lymphoma in a ferret. J Am Vet Med Assoc. 1996;209:1441-1444. 51. Rosenthal KL, Peterson ME. Evaluation of plasma androgen and estrogen concentrations in ferrets with hyperadrenocorticism. J Am Vet Med Assoc. 1996;209:1097-1102. 52. Schoemaker NJ, Kuijten AM, Galac S. Luteinizing hormonedependent Cushing’s syndrome in a pet ferret (Mustela putorius furo). Domest Anim Endocrinol. 2008;34:278-283. 53. Shoemaker NJ, Schuurmans M, Moorman H, et al. Correlation between age at neutering and age at onset of hyperadrenocorticism in ferrets. J Am Vet Med Assoc. 2000;216:195-197. 54. Schoemaker NJ, Teerds KJ, Mol JA, et al. The role of luteinizing hormone in the pathogenesis of hyperadrenocorticism in neutered ferrets. Mol Cell Endocrinol. 2002;197:117-125. 55. Sleeman JM, Clyde VL, Brenneman KA. Granular cell tumour in the central nervous system of a ferret (Mustela putorius furo). Vet Rec. 1996;138:65-66. 56. Smith M, Schulman FY. Subcutaneous neoplasms of the ventral abdomen with features of adrenocortical tumors in two ferrets. Vet Pathol. 2007;44:951-955. 57. Swiderski JK, Seim 3rd HB, MacPhail CM, et al. Long-term outcome of domestic ferrets treated surgically for hyperadrenocorticism: 130 cases (1995-2004). J Am Vet Med Assoc. 2008;232:1338-1343. 58. Taylor TG, Carpenter JL. Thymoma in two ferrets. Lab Anim Sci. 1995;45:363-365. 59. van Zeeland Y, Schoemaker N, Passon-Vastenburg M, et al. Vestibular syndrome due to a choroid plexus papilloma in a ferret. J Am Anim Hosp Assoc. 2009;45:97-101. 60. Wagner RA, Bailey EM, Schneider JF, et al. Leuprolide acetate treatment of adrenocortical disease in ferrets. J Am Vet Med Assoc. 2001;218:1272-1274. 61. Weiss CA, Williams BH, Scott JB, et al. Surgical treatment and long-term outcome of ferrets with bilateral adrenal tumors or adrenal hyperplasia: 56 cases (1994-1997). J Am Vet Med Assoc. 1999;215:820-823. 62. Weiss CA, Williams BH, Scott MV. Insulinoma in the ferret: clinical findings and treatment comparison of 66 cases. J Am Anim Hosp Assoc. 1998;34:471-475. 63. Whittington JK, Emerson JA, Satkus TM, et al. Exocrine pancreatic carcinoma and carcinomatosis with abdominal effusion containing mast cells in a ferret (Mustela putorius furo). Vet Clin North Am Exot Anim Pract. 2006;9:643-650. 64. Wilber J, Williams BH. Osteosarcoma in two domestic ferrets (Mustela putorius furo). Vet Pathol. 1997;34:486. 65. Williams BH. Therapeutics in ferrets. Vet Clin North Am Exot Anim Pract. 2000;3:131-153. 66. Williams BH, Eighmy JJ, Berbert MH, et al. Cervical chordoma in two ferrets (Mustela putorius furo). Vet Pathol. 1993;30:204-206. 67. Williams BH, Garner MM, Kawasaki TA. Peritoneal mesotheliomas in two ferrets (Mustela putorius furo). J Zoo Wildl Med. 1994;25:590-594. 68. Williams BH, Weiss CA. Neoplasia. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. Philadelphia: WB Saunders; 2004:91-106. 69. Wills TB, Bohn AA, Finch NP, et al. Thyroid follicular adenocarcinoma in a ferret. Vet Clin Pathol. 2005;34:405-408. 70. Zoran DL. Is it IBD? Managing inflammatory disease in the feline gastrointestinal tract. Vet Med. 2000;95:128-139.
CHAPTER
9
Dermatologic Diseases
Connie Orcutt, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal), and Kathy Tater, DVM, Diplomate ACVD
Anatomy, Physiology, and Husbandry Considerations Diseases Ectoparasites Viral Disease Fungal Disease Bacterial Disease Neoplasia Endocrine Disease Other Dermatologic Diseases
The initial approach to examining ferrets with dermatologic disease is very similar to that used for other small animals. The signalment is important in forming the initial list of differential diagnoses. A thorough history should include information on the origin of the ferret, housing, diet, vaccinations, condition of cage mate or mates, exposure to other animals, prior or current medical problems, and any skin problems affecting people in the household. Because certain skin conditions are present in conjunction with other clinical signs (e.g., pruritus and alopecia often accompany vulvar swelling in female ferrets with adrenal disease), a complete physical examination is extremely important. Diagnostic tests performed depend on the refined list of differential diagnoses. Direct microscopic examination of skin scrapings and aural debris aids in diagnosing parasitic diseases. If dermatophytosis is suspected, fungal cultures of hair and skin samples are indicated. In treating wounds and abscesses, cytologic evaluation of the exudate is helpful pending results of bacterial culture and sensitivity testing. With cutaneous or subcutaneous masses, fine-needle aspiration and cytologic evaluation of an impression smear can be performed initially, but biopsy is the diagnostic test of choice. In working with older ferrets (> 4 years of age) or any ferret suspected of having systemic disease, 122
a complete blood count (CBC), plasma or serum biochemical analysis, and radiographs are important. Other diagnostic tests may include abdominal ultrasonography (e.g., in suspected cases of systemic disease such as adrenal disease or neoplasia) and/or evaluation of plasma androgen and estrogen levels.
ANATOMY, PHYSIOLOGY, AND HUSBANDRY CONSIDERATIONS The skin of ferrets contains numerous sebaceous glands. Secretions of these glands sometimes cause the hair coat to have a greasy feel and a characteristic musky odor. Intact and sexually mature males and females have larger sebaceous glands than neutered ferrets,25 and glandular production appears to be under androgenic control. Secretions may be so profuse that intact male albino ferrets can appear yellow and dirty. Frequent bathing can remove essentials oils from the skin and result in pruritus and keratinopathies. Ferrets are quite fastidious animals, and routine bathing is not necessary. Ferrets have epitrichial (apocrine) sweat glands on the body and atrichial (eccrine) sweat glands on the footpads.25 They are predisposed to hyperthermia when exposed to high ambient temperatures. Paired musk-producing glands lateral to the anus store secretions that are expelled when ferrets are agitated, excited, or in estrus. These glands rarely become impacted, but the treatment is the same as for cats. Because most of a ferret’s body odor emanates from the dermal sebaceous glands, removal of the anal glands does not remove all scent. However, neutering does decrease much of the skin odor by reducing androgenic stimulation. Anal sacculectomy is usually performed in conjunction with neutering at breeding farms. Normal ferret skin has a compound hair follicle arrangement composed of one primary hair and a collection of secondary hairs, the latter increasing in number with the ferret’s age until maturity.25 Ferrets have a thick cream-colored undercoat with coarse guard hairs that define the hair-coat color. The color of the undercoat remains the same, but the guard hair color can change at different times during the ferret’s life, as can the facial Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 9 Dermatologic Diseases mask.3 The hair coat may become lighter as the ferret ages. Molting, which usually occurs twice a year, appears to be controlled by hormones responsive to changes in the photoperiod; thus estrogens cause hair and weight loss in ferrets.15 During the breeding season (March through August in the northern hemisphere), intact males increase sebaceous secretions in response to elevated hormone levels; the result can be an increased musky odor, oily fur, and a yellowish undercoat (see Chapter 1). In both intact and neutered ferrets, the hair coat normally thins in response to an increase in the number of daylight hours and in the ambient temperature (simultaneously with the breeding season)—that is, in the late spring in the northern hemisphere.15 This molt may result in bilaterally symmetric alopecia of the tail, perineum, and inguinal area, or the ferret may lose most of the guard hairs and appear “fluffy.” The spring molt is often more marked in females than in males and is usually more pronounced in intact ferrets. In the late fall, a less pronounced molt occurs coincident with an increase in density of the light-colored undercoat. As a result, the hair coat appears lighter during the fall and winter seasons. As the hair thins, red-brown waxy deposits, often sebaceous secretions, may be visible on the skin. Hair that is shaved during seasonal hair loss may not regrow for several weeks or months. Hair regrowth, such as occurs after shaving for surgery or after treatment of adrenal gland disease, sometimes imparts a bluish appearance to the skin, which may be mistaken for bruising or cyanosis.3 This apparent discoloration resolves as soon as hairs erupt from the skin. Both sexes accumulate subcutaneous fat beginning with the fall molt and resolving in the spring. As a result, body weight can fluctuate by 30% to 40%.15 The ideal ferret diet is rich in fat and animal protein and low in carbohydrate and fiber. Ferrets should be fed high-quality ferret, cat, or kitten food (see Chapter 1). If dietary requirements are not met, the hair coat can become dry and dull. The provision of a suitable diet, with or without short-term administration of a fat supplement, usually corrects this problem. In addition to inadequate levels of protein or fat, biotin deficiency can cause abnormalities in the hair coat. Avidin, an enzyme found in raw egg white, binds biotin and has been reported to cause bilaterally symmetric alopecia in ferrets fed diets containing more than 20% raw eggs.21 Feeding raw eggs to ferrets is not recommended. Wild ferrets spend considerable time in underground dens with high humidity. Pet ferrets housed in dry environments (e.g., homes that are heated in the winter) may scratch or have flaky skin. The use of a cool-air humidifier or application of an emollient skin spray has been recommended in these cases.3 Exposure to clay kitty litter can also dry a ferret’s coat and should be avoided.3 Certain skin or hair problems in ferrets result from selfmutilation or trauma. Ferrets need privacy and should be provided with hide boxes or artificial burrows. A ferret with inadequate bedding, nesting, or hiding spots may rub its face on the floor in an attempt to hide and subsequently cause facial abrasions. Broken hair shafts can resemble those seen with dermatomycoses. Intact females may pull hair from themselves to use as bedding material.15 Ferrets can be quite rough during mating and playing and can inflict scratches and bite wounds. Many pet ferrets are from large breeding farms where animals are tattooed at an early age after neutering. The tattoo is usually
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inside the pinna and appears as one or two gray or blue dots. On occasion these dots have been mistaken for skin lesions.
DISEASES The most common dermatologic lesions in ferrets are those associated with adrenal disease, benign neoplasia, or ear mite infestation. Malignant skin neoplasms, fungal disease, viral disease, and sarcoptic mange are uncommon. Treatment protocols for dermatologic disease, including medication selection and dosages, are often extrapolated from those used for cats.
ECTOPARASITES Ferrets are susceptible to most of the ectoparasites that affect dogs and cats. However, other than fleas and ear mites, ectoparasites are rarely seen.
Fleas Flea infestation is occasionally seen in ferrets. As with dogs and cats, Ctenocephalides species are usually involved,14 but infestations with Pulex irritans (the human flea), Paracaras meli (the badger flea), Ceratophyllus sciurorum (the squirrel flea), and Ceratopyllus vison (the mink flea) have also been reported, depending on the environment.4,44 Fleas are transmitted by direct contact with another animal or a flea-infested environment. Although some ferrets are asymptomatic, clinical signs generally include mild to intense pruritus, development of erythemic papules, and alopecia most often noted on the dorsal cervical and interscapular areas.47 Ferrets can less commonly develop signs of flea-bite hypersensitivity, including pruritic papulocrustous dermatitis over the tail base, ventral abdomen, or caudomedial thighs.14 Identifying fleas or flea excrement on the animal confirms the diagnosis. Treat the ferret as well as other animals in the household, and treat the environment concurrently. None of the treatments discussed below have been approved by the U.S. Department of Agriculture for use in ferrets. Traditional treatments include flea shampoos, dips, or powders containing pyrethrins, lindane, or carbamates;14 however, toxic reactions occasionally occur with some of these chemicals. Pyrethrins are among the least toxic.4 Organophosphates (malathion, ronnel, fenthion, cythionate, dichlorovos, neguvon) can be highly toxic in ferrets and are not recommended for topical use in this species.4 In addition to the risk of toxicity, dichlorvosimpregnated flea collars can get caught on objects in the environment. Because of the ferret’s small body size, caution is advisable in applying any topical medication. When using sprays, first spray a cloth and then rub the cloth on the ferret. Ferrets can ingest sprays and powders while grooming. Extreme care is advised in the use of pesticide dips; some dips can be toxic with prolonged exposure, and ferrets need to be dried and kept moderately warm after dips. Although newer products developed for flea control in dogs and cats have not been approved for use in ferrets, anecdotal reports of toxicity are uncommon.4 We prefer the use of these products for flea control in ferrets in our practice. Imidacloprid is a flea adulticide that kills on contact and has larvacidal activity in the environment.4,14 Topical imidacloprid (Advantage 40 for Cats, Bayer Corp., Shawnee Mission, KS) applied at a dose of 0.4 mL. 10% on the skin at the base of the skull was well tolerated in ferrets and was effective in eliminating an established flea population as well as in dealing with a subsequent flea
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SECTION I Ferrets
challenge for 3 weeks.9 In conjunction with environmental control, one application of imidacloprid might be sufficient to control a flea problem. Similarly, application of 0.4 mL imidacloprid 10%/moxidectin 1% spot-on (Advocate/Advantage Multi, Bayer Corp., Shawnee Mission, KS) on the interscapular skin in ferrets with established flea infestations resulted in a therapeutic efficacy of 100% on day 1 after treatment and a preventative efficacy of >90% for up to 30 days after treatment.44 Lufenuron (Program, Novartis Animal Health, Greensboro, NC), an insect growth regulator, kills immature fleas by interfering with chitin production. Because there is a lag time of approximately 6 to 8 weeks from the beginning of treatment with lufenuron and reduced numbers of adult fleas on the ferret, an adulticide may have to be used concurrently until the flea population in the environment has been controlled.4 A dosage of 45 mg lufenuron (half of a cat dose) given orally once a month has been used in ferrets.4 Rare side effects may include vomiting, lethargy, diarrhea, pruritus, or inappetance. Fipronil (Frontline, Merial Ltd., Duluth, GA) is a flea adulticide that also kills ticks. Topical application of a half to a full cat dose has been reported anecdotally in ferrets.4 Although the toxic action of fipronil is very specific to invertebrates and as such is not expected to affect mammals, the incidence of adverse reactions in a large population of ferrets is unknown.4 Either imidacloprid or fipronil can be used in conjunction with lufenuron. Selamectin (Revolution or Stronghold, Pfizer, New York, NY) is a semisynthetic avermectin that is approved for use in dogs and cats to control fleas, heartworms, ear mites, and sarcoptic mange mites. In a clinical trial, doses of either 6 or 18 mg/kg used monthly in ferrets for 4 months were 100% effective in preventing flea infestations for 7 to 21 days posttreatment.8 Treatment of the environment is the same as that in households with dogs and cats. Residual chemicals (e.g., microencapsulated formulas of pyrethrins with synergists and diazinon) or methoprene (insect growth retardant preventing fourth instar larval metamorphosis) have been recommended.14 Potentially less toxic forms of treatment that will also destroy parasites are steam cleaning of carpeting and the application of boric acid salts to the floor or carpet. These procedures will desiccate and destroy flea adults and larvae.4 Pets and humans should be prevented from coming in contact with any chemicals applied to the environment.
Mites Ear Mites. The same ear mite that affects dogs and cats (Otodectes cynotis) affects ferrets.14 The mite is transmitted by direct contact with other infested animals. Ferrets may shake their heads or scratch their ears, but they are more commonly asymptomatic. Clinical lesions vary from inflammation of the external ear canal with accompanying mild pruritus to severe pruritus with excoriations and crusting. Dark brown waxy aural debris is often present; however, this can also be seen in some normal ferrets. The mite is identified by microscopic examination of the aural debris. Although ear mites can reportedly colonize other parts of the body, specifically the perineum,5 this rarely occurs. Secondary otitis media or interna with neurologic deficits, most commonly head tilt, is uncommonly seen secondary to ear mite infestation in ferrets in our practice. Acute onset of head tilt may also result from exuberant ear cleaning with solutions that irritate the vestibular nerve if the tympanic membrane is compromised. Clinical signs in these ferrets generally resolve after treatment for mite infestation and secondary bacterial infection.
To eliminate ear mites, all susceptible animals in the household should be treated. Gently clean the ears before any treatment, but avoid solutions that can damage the middle ear if the tympanic membrane is ruptured. Selamectin applied topically at a dose of 45 mg in the dorsal interscapular area approximately every 30 days was reported to be safe and efficacious for the treatment of ear mite infestation in ferrets.28 One of the authors (CO) commonly uses selamectin for the treatment of ear mites in ferrets. Another commonly used treatment protocol is ivermectin injected subcutaneously at a dose of 0.2 to 0.4 mg/kg and repeated every 2 weeks for three treatments. Although this protocol is often successful, some mite infestations are now refractory to treatment. Aural topical treatments have alternatively been reported to be safe and efficacious in ferrets. In a study that compared three protocols for treating ear mites in ferrets (parenteral ivermectin, aural topical administration of ivermectin, and aural topical administration of an otic preparation of thiabendazole, dexamethasone, and neomycin [Tresaderm, Merck Agvet, Rahway, NJ]), results showed that treatment with either the thiabendazole product (2 drops in each ear canal q24h for 7 days, untreated for the next 7 days, then treatment reinstituted for 7 days) or topical ivermectin (1% ivermectin diluted 1:10 in propylene glycol; 0.4 mg/kg divided between each ear canal, then repeated 2 weeks later) was more effective in eliminating mites than parenteral treatment with ivermectin (1% ivermectin diluted 1:4 in propylene glycol; 0.4 mg/kg SC).34 To avoid toxicity, it is recommended not to treat with topical and parenteral ivermectin concurrently. Extra care is advised in treating pregnant animals with ivermectin, because high doses administered to pregnant jills have resulted in an increased rate of congenital defects in kits.14 Thiabendazole is expected to have a wider margin of safety in pregnant animals.34 Topical otic treatments used alone can fail for several reasons: the ear canal may be too narrow for the medications to penetrate, the ferret may resist treatment, or mites may be present on other areas of the body left untreated.14 Some clinicians have recommended treating the entire body with flea powder.14 The entire environment, including bedding, should be thoroughly cleaned as part of any treatment protocol. Sarcoptic Mange. Infestation with Sarcoptes scabiei, which also affects dogs and cats, is uncommon in ferrets. This zoonotic parasite is transmitted by direct contact or fomites. The mite is usually identified by examining a skin scraping; however, falsenegative results are not uncommon. Two different clinical syndromes are seen in ferrets with sarcoptic mange. In the generalized form, clinical signs include focal to generalized alopecia with intense pruritus.14 The localized form of the disease, in which only the feet are affected, is seen occasionally. The paws become inflamed, swollen, and crusted and can be very pruritic. In severe cases, the nails may become deformed or even slough. The layman’s term for this form of the disease is foot rot.14 Traditionally recommended treatment has been ivermectin injected subcutaneously at a dosage of 0.2 to 0.4 mg/kg every 14 days for three treatments.26 Alternative treatment recommendations have included dipping the ferret once weekly in 2% lime sulfur until 2 weeks after the signs resolve,42 but this agent has a strong odor and will discolor the fur. Treatment with either the same imidacloprid/moxidectin formulation mentioned above (Advocate/Advantage Multi Bayer Corp., Shawnee Mission, KS) or selamectin was reported to be highly effective
CHAPTER 9 Dermatologic Diseases against Sarcoptes scabiei in dogs,11 and although its use is offlabel in ferrets, its use carries low risk. Topical or systemic antibiotics are used for secondary bacterial infections. Treat affected feet with warm-water soaks, gentle debridement of crusts, and trimming of diseased claws. All affected animals as well as those in contact with the ferrets must be treated and the cages, bedding, and any other contaminated materials thoroughly cleaned. Demodectic Mange. A histopathologic study on the skin of normal ferrets found Demodex species within the hair follicles and sebaceous glands in the perianal, vulvar, preputial, facial, and caudal abdominal regions in 9 of 25 ferrets aged 4 to 32 months.25 However, clinical demodicosis has rarely been reported in ferrets. In dogs and cats, adult-onset demodectic mange has been associated with immunosuppression. Demodicosis has been reported in ferrets with adrenal disease and systemic lymphoma1 as well as subsequent to prolonged treatment with corticosteroids.32 Clinical signs of demodicosis can include alopecia, skin thickening, and/or discoloration, erythema, and pruritus affecting the ears, face, ventral abdomen, inguinal area, or tail. A brown ceruminous exudate is commonly seen in the otic canals. One of the authors (CO) has seen a persistent Demodex infestation in the otic canals of three geriatric ferrets from the same household, all of which had received ivermectin for several years for heartworm prophylaxis (55 μg orally once monthly) (Heartgard Plus for Cats, Merial, Duluth, GA). All ferrets had histories of adrenal disease and prolonged treatment with prednisone for presumptive insulinoma or inflammatory bowel disease. One male neutered ferret had several small plaques in the skin surrounding the prepuce, histopathologic exam of which revealed multiple cystic apocrine glands. Many of the glands contained cross sections of mites with an elongate morphology consistent with Demodex species. Several mites free
A
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within the superficial dermis appeared to have incited a granulomatous reaction (Fig. 9-1). In such cases deep skin scrapings and aural exudates should be examined for adult mites or larvae. Various treatments have been used singly in ferrets, including imidacloprid-moxidectin spot-on (once monthly for several months) (Advantage Multi; Bayer Health Care, Toronto, Canada),1 ivermectin administered orally (up to 0.3 μg/kg q24h),1 and amitraz (whole-body dips in a 0.0125% solution at 7-day intervals for three treatments and local topical application of the same solution in the ears every other day).32 Other Mite Infestations. Infestation by the fur mite, Lynxacarus mustelae, was reported in 5 ferret kits with ulcerative dermal lesions on the face.40 The lesions resolved after topical application of a powder containing permethrin on each kit and cleaning the animals’ cage with a permethrin-containing shampoo.
Ticks Ticks can be found on ferrets housed outdoors and are reportedly common in ferrets used for hunting in Great Britain.14 Remove ticks from ferrets as from other domestic animals by extracting the entire head from the skin. Because ticks can carry zoonotic diseases, wear gloves and use forceps to remove the tick. No cases of Lyme disease have been reported in ferrets, but this may be influenced by the lack of available diagnostic tests for ferrets.
Cutaneous Myiasis Cuterebra species can cause subdermal cysts in mustelids and have been uncommonly seen in ferrets.14 Granulomatous masses in the cervical area caused by larval stages of Hypoderma bovis are also uncommon.14 The moving larvae can often be seen through the open pore of the swollen area.
B
Fig. 9-1 Demodicosis in a geriatric male neutered ferret. Several small plaques in the skin surrounded the prepuce (A) of a geriatric male castrated ferret on chronic treatment with prednisone for presumptive inflammatory bowel disease. Histopathologic examination revealed a granulomatous reaction and cystic apocrine glands containing numerous cross sections of mites that were found in longitudinal section to be morphologically consistent with Demodex species (B) (hematoxylin and eosin stain, ´10). (Both photographs courtesy Pamela Mouser, DVM, MS, DACVP, Boston, MA.)
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SECTION I Ferrets
Fly strike, or infestation by the flesh fly (Wohlfahrtia vigil), has been reported by commercial mink and ferret ranchers and by owners who keep ferrets outdoors.14 Mink kits that are several weeks old are most commonly attacked during the summer months. Fly eggs laid on the face, neck, or flanks of the kits bore into the skin and cause irritation, and larvae localized in subcutaneous tissues can produce abscess-like lesions. In any ferret with cutaneous myiasis, attempt to remove the larvae intact so as to avoid leaving a nidus of infection or precipitating a systemic response. Debride the wound and apply topical antibiotic preparations, with or without the use of systemic antibiotics, to prevent or treat secondary bacterial infections. The wound is then allowed to heal by second intention.
VIRAL DISEASE Viral disease is a rare cause of dermatopathy in ferrets. Canine distemper virus (CDV) is the important exception. The causative agent, a paramyxovirus, is transmitted by direct contact, aerosol exposure to infected body fluids, or contact with fomites (see Chapter 6). Viremia has been detected 2 days after infection, subsequent to which the virus begins replicating in several tissues.16 However, in some cases of natural infection, the incubation period can be >56 days.35 Generally by about 10 days after exposure, the ferret becomes anorexic and febrile, may appear to be photosensitive, often develops a serous nasal and ocular discharge, and may cough and sneeze. Within 10 to 15 days of exposure, a characteristic erythematous and pruritic skin rash develops under the chin and then spreads to the inguinal or perianal region. This generalized dermatitis may be orange-tinged and can predispose to secondary pyoderma. Less commonly, generalized desquamation is reported.35 The nasal and ocular discharges become mucopurulent, resulting in brown crusts around the face, and the chin, lips, and eyelids can become swollen. A characteristic clinical sign is swelling and hyperkeratosis of the footpads (“hard pad”). Secondary bacterial infections may be observed. Definitive diagnosis of CDV is difficult. A presumptive diagnosis is based on the presence of distinctive clinical signs as well as a questionable vaccination history. The test of choice to rapidly confirm CDV is a fluorescent antibody test performed on a peripheral blood smear, buffy coat, or conjunctival scraping.16 In some cases, results of the test can be positive even before the ferret develops pyrexia. A polymerase chain reaction (PCR) assay has been used to test ferret peripheral mononuclear cells and conjunctival swabs.43,48 On histopathologic examination, eosinophilic cytoplasmic and intranuclear viral inclusions are generally widespread in tissues; however, inclusions are variably reported as being less common16 or frequently seen35 in skin. The mortality rate of CDV in ferrets is generally 100%. Death can occur from 12 to 35 days after exposure.16 Because of the untreatable and highly infectious nature of this disease, any ferret with a poor vaccination history or suspicious clinical signs should be isolated from other animals. The prognosis is grave. The primary treatment is supportive care, including administration of fluids, antimicrobials, bronchodilators, mucolytics, and nutritional support.35 In a recent outbreak of CDV in 14 young ferrets, the only survivor was reportedly treated early in the course of the disease with anti-CDV hyperimmune serum.35 A case of atypical CDV in a previously vaccinated ferret was reported. The course of the disease was prolonged (53 days), and neither respiratory nor neurologic signs were observed.48
The ferret’s primary clinical signs were pruritus and thickened erythematous skin with crusts over most of the body. On postmortem examination, eosinophilic intranuclear and intracytoplasmic inclusions were seen in multiple organs, including epidermal and follicular epithelium, and abundant CDV antigen was identified with immunohistochemistry.
FUNGAL DISEASE Dermatophytosis Fungal disease involving the skin is uncommon in pet ferrets. Although superficial mycotic skin infections (ringworm) in ferrets have been frequently seen by some clinicians,5 we rarely see this clinical entity in ferrets in our practice. Perhaps this is because of the geographic variation in incidence, which depends on climate. Ferrets are susceptible to infection with both Microsporum canis and Trichophyton mentagrophytes,42 although the former is more common.26 Microsporum canis can be transmitted by direct contact or by fomites. Clinical disease is more common in kits and young ferrets and can occur as a seasonal, self-limiting infection.13 Dermatophytosis may also affect ferrets of any age secondary to immunosuppressive disease. As with dogs and cats, dermatophytosis in ferrets is a zoonotic disease.13,42 Skin and hair lesions in ferrets with dermatophytosis are similar to those reported in other species. Dermatologic lesions can begin with small papules that spread peripherally. These can lead to large circumscribed areas of alopecia and inflammation involving all parts of the body. The skin becomes thickened, erythematous, and hyperkeratotic with superficial crusts, and hair shafts may appear broken. Affected animals are usually nonpruritic. One of the authors (CO) has seen fungal pododermatitis in a 7-year-old ferret subsequent to chemotherapy for systemic lymphosarcoma. Cytologic examination of a fine-needle aspirate of erythematous plaques on the feet showed pyogranulomatous inflammation with intralesional fungal hyphae, and a fungal culture grew Microsporum nanum. Although clinical signs may raise suspicion (especially if other animals or people in the household have skin lesions), dermatophytosis is definitively diagnosed by the results of a mycotic culture of a skin scraping or hair sample. Fungal organisms can also sometimes be appreciated on histologic examination of skin biopsy samples, especially with a periodic acid-Schiff (PAS) stain. M. canis may fluoresce with a Wood’s lamp, but Trichophyton species do not.13 Although some clinicians describe microscopic visualization of fungal arthrospores in skin scrapings and hair plucks that have been mixed with 10% potassium hydroxide,13 others have not found this method useful.5 Many authors report spontaneous remission of clinical signs of dermatophytosis in ferrets.13 In those that are treated, gently clip the hair around the lesions. A topical antifungal (e.g., miconazole cream) can be used19 but will not treat any infective spores present throughout the coat. Instead, lime sulfur dips can be applied weekly to the entire body as a topical antifungal treatment for dermatophytosis. Itraconazole (15 mg/kg PO q24h)37 can be used as a systemic therapy for dermatophytosis in conjunction with lime sulfur. If itraconazole is used, monitor liver enzyme concentrations periodically while the ferret is receiving treatment. Likewise, griseofulvin can be used at dosages recommended for other small mammals.42 If griseofulvin is used, monitor the CBC results every 2 weeks while the ferret is receiving treatment. Any treatment for the dermatophytosis must continue until two negative fungal cultures are
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obtained because clinical signs may improve prior to full resolution of the infection. The entire environment should be disinfected to eliminate infectious spores. Methods of decontaminating the environment include applying dilute chlorine bleach (1:10), high temperature professional steam-cleaning of carpets, changing air conditioning filters, cleaning heating ducts, and vacuuming with prompt disposal of vacuum bags. All animals in the household must be treated.
Other Fungal Infections Systemic mycoses with skin manifestations have been reported infrequently in ferrets; however, these diseases should be included on a list of differential diagnoses for persistent draining tracts and skin eruptions that are unresponsive to antibiotic therapy, especially if other systemic signs are evident (e.g., pyrexia, pneumonia, weight loss, or neurologic signs).47 Blastomycosis, coccidioidomycosis, and histoplasmosis have each been reported in ferrets,42 and treatments have included oral ketoconazole and intravenous amphotericin B. Mucormycosis, caused by infection with Absidia corymbifera, a widespread environmental saprophyte, has previously been reported to be common in ferrets raised for fur in New Zealand.13 Clinical signs included pruritus and inflammation of the external auditory canal as well as occasional neurologic deficits.
BACTERIAL DISEASE Primary bacterial skin infections are infrequently seen in ferrets and are most commonly caused by Staphylococcus aureus or Streptococcus species.42 However, ferrets can incur bite wounds while playing, mating, or fighting and can sustain puncture wounds from chewing on objects. These wounds can become infected, resulting in superficial or deep pyoderma, abscess, or cellulitis. Abscesses can involve the anal glands or remnants of these glands (Fig. 9-2). Abscesses usually become walled off, causing few systemic signs, but in some instances the affected ferret may be febrile and demonstrate leukocytosis. A tentative diagnosis may be reached by examining a Gram’s stain of discharge aspirated from an abscess. Definitive diagnosis is determined by aerobic or anaerobic bacterial culture and sensitivity testing of the exudate or infected tissue. Using sterile technique, lance and flush or debride the abscess. Depending on the size and location of the abscess, place drains in the wound, leave the wound open for flushing, or use wet-to-dry bandages over the wound. While culture results are pending, begin treatment with a broad-spectrum systemic antibiotic. Actinomycosis, or “lumpy jaw,” has rarely been reported in ferrets.12 Infection occurs when bacteria enter wounds to the oral mucosa or are swallowed or inhaled. Clinical signs include cervical masses with sinus tracts containing thick, yellowish-green, purulent material. Masses occasionally become large enough to cause dyspnea. As with any abscessed area, the lesions must be surgically debrided and drained. The results of bacterial culture and sensitivity testing then determine the choice of antimicrobial therapy. Empirical treatments that have been suggested for actinomycosis include penicillin (40,000 IU/kg SC q24h) or tetracycline (25 mg/kg PO q12h).26 Superficial spreading pyoderma has been reported in a ferret.20 This syndrome results from epidermolytic toxins produced by Staphylococcus species. Clinical signs in this ferret included skin erosions, ulcerations, crusts, and erythematous macules.
Fig. 9-2 Anal gland abscess in a ferret. Anal sacculectomy was performed and the abscess contents were submitted for aerobic culture. No bacteria were isolated.
Results of skin biopsy showed subcorneal epidermal pustules with inflammatory cells and gram-positive cocci along with dermal necrosis. Treatment included systemic antibiotic therapy based on bacterial culture and sensitivity testing.
NEOPLASIA A retrospective study of neoplastic diseases in 574 ferrets reported a 14% incidence of neoplasia involving the integumentary system, making this the third most common system affected.24 In a literature search of 214 cases of primary neoplasms of the integumentary system in ferrets, benign lesions were predominant, with mast cell tumors, basal cell tumors, and sebaceous cell tumors being the most common.22 Several of the tumor types described below appear very similar clinically. Excisional biopsy should be performed whenever a skin mass is detected.
Mast Cell Tumors In ferrets, mast cell tumors generally involve the skin and are benign, being composed predominantly of well-differentiated mast cells. Visceral involvement and malignant behavior of mast cell tumors are rare. In two reviews of neoplasia in ferrets, cutaneous mast cell tumors represented 21% to 24% of all neoplasms involving the integumentary system.22,24 Although mast cell tumors can occur anywhere on the body, they are most common on the head, neck, shoulders, or trunk. Mast cell tumors usually appear as single or multiple raised, well-circumscribed, hairless nodules that can vary in size from 0.2 to 1 cm. They are often hyperemic and may be ulcerated or covered with a black, crusty exudate. Some affected ferrets may present because their owners notice this crusty material even though the tumor underneath is still very small and barely raised from the skin surface (Fig. 9-3). Biologically active substances, such as histamine, present in mast cell granules often induce pruritus, and the resultant selfexcoriation can cause ulcers. However, mast cell tumors may remain static in appearance for months.36
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Fig. 9-3 Superficial black crusty debris overlying a dermal plaque in a ferret. This is a common clinical manifestation of both mast cell and sebaceous cell tumors.
Fig. 9-4 Sebaceous adenoma on the tail of a ferret. While the Surgical biopsy provides the definitive diagnosis, but analysis of an aspirate of the mass may reveal mature mast cells. We have not seen any systemic effects from degranulation of mast cells in ferrets after manipulating the dermal mass. Surgical resection is generally curative. Spontaneous resolution and recurrence of lesions has been described.43 New mast cell tumors occasionally develop in the same ferret over time.
appearance is typical for this tumor type, these neoplasms in ferrets are most commonly seen on the head, neck, or trunk.
Basal Cell Tumors, Sebaceous Adenomas, Epitheliomas, Cystadenomas In a retrospective study of 57 cutaneous neoplasms diagnosed by biopsy in ferrets, the incidence of basal cell tumors was 58%.33 Another review reported a 20% incidence.22 These tumors, which arise from the pluripotential basal cells of the epidermis, have been variably named basal cell tumors, sebaceous cell tumors, or basosquamous sebaceous tumors. Tumors of basal cell origin can develop anywhere on the body and are usually benign (see Chapter 8).22,33 Most basal cell tumors are sharply defined, firm, white to pink lesions that are pedunculated or plaque-like and sometimes ulcerate. Recurrence of a basal cell tumor excised from the tail was reported in one ferret, but no other recurrences or metastases have been reported.33 Other benign neoplasms include sebaceous adenomas, epitheliomas, and cystadenomas, all of which contain neoplastic cells that appear primarily as sebaceous cells.22 Sebaceous cell tumors can develop anywhere on the ferret’s body but are most commonly seen on the head or neck.36 They can be verrucous, inflamed, or ulcerated and up to 3 cm in diameter (Fig. 9-4).22 In our clinical practice, the prevalence of sebaceous cell tumors in ferrets nearly equals that of mast cell tumors; clinically, the 2 tumor types often look very similar. In some cases, both tumor types may be present in the same lesion (Fig. 9-5).
Squamous Cell Carcinoma Squamous cell carcinoma (SCC) uncommonly involves the skin of ferrets. Any area of the skin can be affected, but the perivulvar area of females is reportedly most common.44 These tumors tend to be firm, gray to white, single or multiple nodules or plaques in the skin that occasionally ulcerate. Local invasion and metastasis to other sites, such as lymph nodes, is common.22,44 One author (CO) has seen a 5-year-old neutered female ferret with a large, thickened, erythematous, ulcerated perianal squamous cell carcinoma that was nonresectable. The ferret had a second
Fig. 9-5 Sebaceous epithelioma and mast cell tumor in a ferret. This raised, ulcerated dermal mass was composed of two distinct tumor types.
mass in the right inguinal area with an enlarged right inguinal lymph node. Whenever possible, any masses should be surgically excised as soon as possible to decrease the extent of local invasion. Chemotherapy has not been rewarding for this tumor type.17,44 In animals with nonresectable tumors, external beam radiation or strontium-90 plesiotherapy may be a treatment option, as in other species.6,18 The prognosis for ferrets with SCC is generally poor. Unusual presentations of SCC in ferrets have recently been reported. An adult male ferret of unknown age was presented for fecal incontinence; the anus was dilated and the rectal mucosa partially exposed. A symmetric ring of firm perianal subcutaneous tissue was palpated on exam. The histopathologic diagnosis on necropsy was SCC arising from the anal sac.45 A 6-year-old male castrated ferret was presented with multiple black and tan proliferative skin lesions. Histopathologic evaluation of skin biopsies revealed multicentric SCC in situ as well as intranuclear
CHAPTER 9 Dermatologic Diseases
Fig. 9-6 Sebaceous gland adenocarcinoma in a 5-year-old male castrated ferret. The mass was excised with complete margins.
inclusion bodies within keratinocytes. Transmission electron microscopy demonstrated viral particles compatible with papillomavirus in dysplastic keratinocytes; immunohistochemical staining for papillomavirus was positive. The lesions were similar to those seen in cats with Bowenoid in situ carcinoma.38
Adenocarcinoma The perianal, perivulvar, and peripreputial regions in the ferret contain a large concentration of apocrine scent glands from which a variety of benign and malignant neoplasms originate.45 Most adenocarcinomas of apocrine scent glands develop in the perineal area. In some cases, tumors have been excised with no report of recurrence or metastasis.34 In other cases, metastasis and rapid recurrence of tumor has been reported subsequent to multiple excisions and radiation therapy.33 In a recent report, perianal masses diagnosed as apocrine adenocarcinoma presumed to be of anal gland origin recurred after surgical incision, which was incomplete because of deep rectal invasion. Radiotherapy using an orthovoltage unit at 4 Gy twice weekly was delivered for a total of 6 doses, after which no visible tumor was detected. However, radiotherapy was discontinued after the ferret developed pleural effusion. Cytologic evaluation was suggestive of mesothelioma, and no obvious metastatic disease from the original tumor was seen on thoracic radiographs. The ferret died 71 days after the original presentation.31 One of the authors (CO) has seen two cases of adenocarcinoma in ferrets. In one case, a slow-growing, broad-based, elevated skin mass was present on the lateral aspect of the tail of a young intact male ferret. On excisional biopsy, the mass was identified as a sweat gland adenocarcinoma with local infiltration. Within 2 months, the tumor had metastasized to the deep tissues of the right thigh and perianal area and a sublumbar mass (possible lymphadenopathy) was seen on radiographs. Two months later, the ferret died suddenly of unknown cause. The second case was a 5-year-old male neutered ferret with a 2.2-cm, broad-based, sessile, ulcerated dermal mass on the right lateral thigh. The mass was excised with complete margins, and the histopathologic diagnosis was sebaceous gland adenocarcinoma (Fig. 9-6). Operable tumors must be excised with wide tissue margins. Removal of tumors involving the prepuce may necessitate penile amputation and urethrostomy (see Chapter 11).10
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Fig. 9-7 Severe ulcerative lesions in a ferret with cutaneous lymphoma. (Courtesy Stephen White, DVM. Picture from the case files of the Veterinary Medical Teaching Hospital, School of Veterinary Medicine, University of California. Used with permission.)
Cutaneous Lymphoma Cutaneous (epitheliotrophic) lymphoma occasionally occurs in ferrets19,23,39 and should be included in the differential diagnosis of ferrets with chronic pruritus, dermatitis, or alopecia. Ulcerative lesions can also be significant (Fig. 9-7). Cutaneous lymphoma of T-cell origin was diagnosed in an 8-year-old neutered female ferret with generalized progressive, pruritic dermatitis.39 Treatment with isotretinoin (2 mg/kg PO q24h) resulted in significant improvement within 60 days, but the ferret was subsequently euthanized because of renal failure resulting from pyelonephritis. In a spayed female ferret with severe swelling of all 4 paws, erythematous skin, and malformed and missing nails, epitheliotropic lymphoma was diagnosed by histopathologic examination of a biopsy sample.19 The ferret’s clinical signs improved dramatically within 4 weeks of treatment with oral prednisone and a topical antibacterial/corticosteroid ointment, but the animal presented 6 months later with signs of advanced systemic lymphoma and was euthanized. Lymphoma involving the prepuce was reported in a 6-year-old intact male ferret.33 The tumor presented as a 2-cm ulcerated mass on the prepuce, and the inguinal lymph node was also involved.
Other Neoplasms Involving the Skin or Subcutis Hemangiomas have been reported in two middle-aged female ferrets.33 Both lesions were well-circumscribed dermal masses with one tumor on the pinna and the other in the dorsal lumbar area. No recurrence or metastasis of the masses was reported after resection. In a review of 57 cutaneous neoplasms in ferrets, 6 fibromas and 2 fibrosarcomas were described.33 There was no site predilection, and the affected ferrets ranged in age from 10 months to 4.5 years. Both tumor types were described as well-circumscribed dermal or subcutaneous masses. In ferrets that were followed up after excision, neither recurrence nor metastasis of either type was reported. Vaccination-associated sarcomas, similar to those occurring in cats, have also been reported in ferrets.30 Sarcomas were subcutaneous and more pleomorphic in morphology on histopathology than sarcomas occurring in locations not used for
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vaccination. In ferrets with vaccination-associated sarcomas, tumor recurrence after excision has been reported. Other cutaneous or subcutaneous tumors in ferrets include extraskeletal osteosarcoma,2 hemangiosarcoma,33 chordoma,29 rhabdomyosarcoma,33 neurofibroma,22,33 neurofibrosarcoma,22 histiocytoma,22 myxosarcoma of the subcutis,22 myelosarcoma,22 and leiomyosarcoma.27
ENDOCRINE DISEASE Endocrine disease, primarily adrenal gland disease, is a very common cause of dermatopathy in ferrets. In fact, adrenal gland disease is the most common cause of progressive and sustained alopecia, often with pruritus, in neutered ferrets (see Chapter 7). Affected ferrets may develop a strong odor from androgenic stimulation of sebaceous glands in the skin. Serpentine and annular erythematous macules have also been reported in association with adrenal disease in ferrets.41,42,46 Adrenalectomy did not result in resolution of the clinical signs (cutaneous telangiectasia) in one ferret.46 In another ferret, the dermatitis resolved after 20 days of treatment with a commercial omega 3/omega 6 fatty acid supplement.42 Other causes of alopecia in both intact and neutered ferrets are estrus, retained ovarian remnants, and neoplastic lesions involving ovarian tissue. No confirmed cases of hypothyroidism have been documented in ferrets.
OTHER DERMATOLOGIC DISEASES Apocrine cysts are benign lesions but can become quite large. Lesions are usually firm and filled with pale yellow or colorless fluid (Fig. 9-8). A pemphigus foliaceus-like skin disease was reported in one ferret.7 Clinical signs included pustules and bilaterally symmetric crusts on the face, pads, and prepuce. The ferret was moderately pruritic and had a decreased appetite and lethargy. Skin biopsy showed a pustule with occasional acantholytic cells. The negative dermatophyte culture and lack of response to antibiotics made pustular dermatophytosis and pyoderma, respectively, less likely. The lesions responded to prednisolone therapy, and a relapse was noted when the prednisolone was discontinued. Contact dermatitis can occur in ferrets; specific etiologies referenced for ferrets include frequent use of shampoos or insecticide sprays. One of the authors (CO) has seen a case
Fig. 9-8 Apocrine cyst in a ferret. These fluid-filled lesions are benign but can become very large.
of generalized erythema in a ferret subsequent to the use of a povidone-iodine scrub for surgical preparation. Chlorhexidine scrubs are recommended for use in ferrets. Presumptive atopy has been described in ferrets presenting with pruritus involving the trunk, rump, and paws.42 In the ferrets described, hypoallergenic diets were ineffective, but response to treatment with glucocorticoids or chlorpheniramine was good. One of the ferrets with similar clinical signs appeared to be food-hypersensitive, and clinical signs resolved after it was fed a hypoallergenic diet for cats (Innovative Veterinary Diets, venison; Royal Canin, St. Charles, MO).42 Erythema multiforme has been anecdotally reported in ferrets. We have seen a 5-year-old male castrated ferret with conjunctivitis and multifocal, erythematous, crusted and ulcerated plaques involving the skin of the neck, inguinal region, and paws (Fig. 9-9). The histopathologic diagnosis from a skin biopsy revealed epidermal orthokeratotic hyperkeratosis with foci of parakeratosis, moderate intracellular edema, dyskeratotic keratinocytes within the stratus spinosum, and foci of necrotic keratinocytes with infiltrates of neutrophils and lymphocytes; these findings were consistent with erythema multiforme. The ferret had been vaccinated for rabies 2 months before presentation but had been exposed to no medications before the development of the lesions. Results of immunohistochemistry for canine distemper virus was negative and characteristic viral cytopathic changes were not seen. An adrenal panel revealed markedly high levels of estradiol, androstenedione, and 17-hydroxyprogesterone; and yeast and bacteria were found on a skin scraping. The ferret was treated with enrofloxacin, selamectin (for previously diagnosed ear mites), and a shampoo containing chlorhexidine and miconazole. Leuprolide acetate (111 μg IM; Lupron Depot 3.75 mg, Abbott Laboratories, N. Chicago, IL) was administered once monthly. The owner reported that the skin lesions had completely resolved after 2 months of treatment but would begin to recur if the leuprolide acetate was not administered promptly at 1-month intervals.
Fig. 9-9 Erythema multiforme in a ferret. This 5-year-old castrated male had markedly elevated estradiol and androgen levels; bacteria and yeast were found on a skin scraping.
CHAPTER 9 Dermatologic Diseases
References 1. Beaufrere H, Neta M, Smith DA, et al. Demodectic mange associated with lymphoma in a ferret. J Exot Pet Med. 2009;18:57. 2. Berrocal A. Dermal and subcutaneous extraskeletal osteosarcomas in eight dogs, two cats, and one ferret. Vet Derm. 2004;15:61. 3. Brown SA. Small Mammal Health Series: Ferret grooming. Veterinary Partner.com. 2001. Retrieved October 26, 2010, from http://www.veterinarypartner.com/Content.plx?P=A&A=675& S=1&SourceID=43. 4. Brown SA. Small Mammal Health Series: Flea control for ferrets. Veterinary Partner.com. 2001. Retrieved October 26, 2010, from http://www.veterinarypartner.com/Content.plx? P=A&A=492&S=1&EVetID=0. 5. Burke TJ. Skin disorders of rodents, rabbits, and ferrets. In: Kirk RW, Bonagura JD, eds. Kirk’s current veterinary therapy XI: small animal practice. Philadelphia: WB Saunders; 1992:1170. 6. Cunha SC, Carvalho LA, Canary PC, et al. Radiation therapy for feline cutaneous squamous cell carcinoma using a hypofractionated protocol. J Feline Med Surg. 2010;12(4):306-313. 7. Eckerman-Ross C. Pemphigus foliaceus-like skin disease in a ferret. Exot DVM. 2007;9:5. 8. Fisher M, Beck W, Hutchinson MJ. Efficacy and safety of selamectin (Stronghold Revolution) used off-label in exotic pets. Intern J Appl Res Vet Med. 2007;5:87. 9. Fisher MA, Jacobs DE, Hutchinson MJ, et al. Efficacy of imidacloprid on ferrets experimentally infested with the cat flea, Ctenocephalides felis. Compend Contin Educ Pract Vet Suppl. 2001;23:8. 10. Fisher PG. Urethrostomy and penile amputation to treat urethral obstruction and preputial masses in male ferrets. Exot DVM. 2002;3:21. 11. Fourie LJ, Heine J, Horak IG. The efficacy of an imidacloprid/ moxidectin combination against naturally acquired Sarcopes scabiei infestations on dogs. Aust Vet J. 2006;84:17. 12. Fox JG. Bacterial and mycoplasmal diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:321. 13. Fox JG. Mycotic diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:393. 14. Fox JG. Parasitic diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:375. 15. Fox JG, Bell JA. Growth, reproduction, and breeding. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:211. 16. Fox JG, Pearson RC, Gorham JR. Viral diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:355. 17. Hamilton TA, Morrison WB. Bleomycin chemotherapy for metastatic squamous cell carcinoma in a ferret. J Am Vet Med Assoc. 1991;198:107. 18. Hammond GM, Gordon IK, Theon AP, et al. Evaluation of strontium Sr 90 for the treatment of superficial squamous cell carcinoma of the nasal planum in cats: 49 cases (1990-2006). J Am Vet Med Assoc. 2007;231(5):736-741. 19. Kelleher SA. Skin diseases of ferrets. Vet Clin North Am Exot Anim Pract. 2001;2:565. 20. King WW, Lemarie SL, Veazey RS, et al. Superficial spreading pyoderma and ulcerative dermatitis in a ferret. Vet Derm. 1996;7:43. 21. Lewington JH. Ferret husbandry, medicine and surgery. Edinburgh: Butterworth-Heinemann; 2000. 22. Li X, Fox JG. Neoplastic diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:405. 23. Li X, Fox JG, Erdman SE, et al. Cutaneous lymphoma in a ferret (Mustela putorius furo). Vet Pathol. 1995;32:55.
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24. Li X, Fox JG, Padrid PA. Neoplastic diseases in ferrets: 574 cases (1968-1997). J Am Vet Med Assoc. 1998;212:1402. 25. Martin AL, Irizarry-Rovira AR, Bevier DE, et al. Histology of ferret skin: preweaning to adulthood. Vet Dermatol. 2007;18:401. 26. Meredith A. Ferret dermatoses. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester: British Small Animal Veterinary Association; 2009;269. 27. Mikaelian I, Garner MM. Solitary dermal leiomyosarcomas in 12 ferrets. J Vet Diagn Invest. 2002;14:262. 28. Miller DS, Eagle RP, Zabel S, et al. Efficacy and safety of selamectin in the treatment of Otodectes cynotis infestation in domestic ferrets. Vet Rec. 2006;159:748. 29. Munday JS, Brown CA, Richey LJ. Suspected metastic coccygeal chordoma in a ferret (Mustela putorius furo). J Vet Dian Invest. 2004;16:454. 30. Munday JS, Stedman NL, Richey LJ. Histology and immunohistochemistry of seven ferret vaccination-site fibrosarcomas. Vet Pathol. 2003;40:288. 31. Nakata M, Miwa Y, Nakayama H, et al. Localized radiotherapy for a ferret with possible anal sac apocrine adenocarcinoma. J Small Anim Pract. 2008;49:476. 32. Noli C, van der Horst HH, Willemse T. Demodicosis in ferrets (Mustela putorius furo). Vet Q. 1996;18:28. 33. Parker GA, Picut CA. Histopathologic features and post-surgical sequelae of 57 cutaneous neoplasms in ferrets (Mustela putorius furo L.). Vet Pathol. 1993;30:499. 34. Patterson MM, Kirchain SM. Comparison of three treatments for control of ear mites in ferrets. Lab Anim Sci. 1999;49:655. 35. Perpiñan D, Ramis A, Tomás A, et al. Outbreak of canine distempter in domestic ferrets (Mustela putorius furo). Vet Rec. 2008;163:246. 36. Rakich PM, Latimer KS. Cytologic diagnosis of diseases of ferrets. Vet Clin North Am Exot Anim Pract. 2007;10:61. 37. Ramsey I, ed. BSAVA small animal formulary. 7th ed. Gloucester: BSAVA; 2010. 38. Rodrigues A, Gates L, Payne HR, et al. Multicentric squamous cell carcinoma in situ associated with papillomavirus in a ferret. Vet Path. 2010;47:964. 39. Rosenbaum MR, Affolter VK, Usborne AL, et al. Cutaneous epitheliotropic lymphoma in a ferret. J Am Vet Med Assoc. 1996;209:1441. 40. Schoemaker NJ. Selected dermatologic conditions in exotic pets. Exot DVM. 1999;1:5. 41. Scott DW, Gould WJ, Cayatte SM, et al. Figurate erythema resembling erythema annulare centrifugum in a ferret with adrenocortical adenocarcinoma-associated alopecia. Vet Derm. 1994;5:11. 42. Scott DW, Miller WH, Griffin CE. Dermatoses of pet rodents, rabbits and ferrets. In: Scott DW, Miller WH, Griffin CE, eds. Small animal dermatology. 6th ed. Philadelphia: WB Saunders; 2001:1417. 43. Stephensen CB, Welter J, Thaker SR, et al. Canine distemper virus (CDV) infection of ferrets as a model for testing Morbillivirus vaccine strategies: NYVAC- and ALVAC-based CDV recombinants protect against symptomatic infection. J Virol. 1997;71:1506. 44. Wenzel U, Heine J, Mengel H, et al. Efficacy of imidacloprid 10%/moxidectin 1% (Advocate/Advantage Multi) against fleas (Ctenocephalides felis felis) on ferrets (Mustela putorius furo). Parasitol Res. 2008;103:231. 45. Williams BH. Squamous cell carcinoma arising from the anal sac in a ferret. Exot DVM. 2002;4:7. 46. Williams BH, Fisher PG, Johnson TL. Diffuse cutaneous telangiectasia in a ferret with adrenal-associated endocrinopathy. Exot DVM. 2002;4:9. 47. Wolf TM. Ferrets. In: Mitchell MA, Tully TN, eds. Manual of exotic pet practice. St. Louis: Saunders; 2009:345. 48. Zehnder AM, Hawkins MG, Koski MA, et al. An unusual presentation of canine distemper virus infection in a domestic ferret (Mustela putorius furo). Vet Derm. 2008;19:232.
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10
Musculoskeletal and Neurologic Diseases
Natalie Antinoff, DVM, Diplomate ABVP (Avian), and Carley J. Giovanella, DVM, Diplomate ACVIM (Neurology)
Posterior Paresis, Ataxia, and Seizures Posterior Paresis Ataxia Seizures Diagnosis of Posterior Paresis, Ataxia, and Seizures Treatment of Posterior Paresis, Ataxia, and Seizures Spinal Disorders Aleutian Disease Spinal Tumors Intervertebral Disk Disease Intracranial Disorders Canine Distemper Rabies Osteoma Primary Neoplasia of the Central Nervous System Neuronal Vacuolation Musculoskeletal Disorders Disseminated Idiopathic Myofasciitis Myasthenia Gravis Miscellaneous Diseases Other Metabolic Diseases
Ferrets with neurologic, musculoskeletal, or systemic disease may present with rear limb weakness, ataxia/incoordination, or both. Primary neurologic and musculoskeletal disorders are not common in pet ferrets. Clinical signs that appear to be caused by primary neurologic disease, particularly posterior paresis, are frequently manifestations of systemic illness. Therefore a thorough and accurate history and physical examination supported by diagnostic imaging and laboratory testing are essential to establishing a diagnosis, treatment plan, and prognosis. Fracture management is covered in Chapter 33. Neurologic signs in ferrets are often a result of generalized disease, such as heart failure or hypoglycemia. Thus a 132
general health evaluation is important for all neurologic patients. A common cause of posterior paresis is hypoglycemia secondary to a pancreatic beta-cell tumor, or insulinoma (see Chapter 8). Hypoglycemia can also result from food deprivation or anorexia, vomiting, sepsis, neoplasia, severe hepatic disease, or any metabolic disorder. Cardiac disease, hypoxia, anemia, and toxin ingestion can result in weakness, ataxia, or central nervous system (CNS) depression.5,42 Toxicosis from ibuprofen ingestion has been reported to cause neurologic signs including ataxia, depression, coma, and tremors.41 Clostridium botulinum type C endotoxin causes signs that include dysphagia, ataxia, salivation, and paresis 12 to 96 hours after ingestion of contaminated food; signs can progress to death if the ferret is untreated.11 Proliferative bowel disease has been associated with paresis and ataxia.14 This may also occur secondary to discomfort and physical obstruction of limb movement from diseases such as splenomegaly, a caudal abdominal mass, inguinal or sublumbar lymphadenopathy, cystic calculi, peritonitis, prostatic enlargement, or urinary obstruction. Urinary and fecal incontinence may accompany posterior paresis if the underlying problem affects the caudal lumbar innervation of these structures.
POSTERIOR PARESIS, ATAXIA, AND SEIZURES POSTERIOR PARESIS Posterior paresis is synonymous with rear leg weakness. Generalized weakness in ferrets is often more pronounced in the rear legs and may be mistakenly attributed to primary neurologic disease. A ferret that is weak loses the normal upward arch in its back, so that the long axis of its body becomes parallel to the ground when it is standing or walking.
ATAXIA Ataxia is incoordination; it can be characterized as either cerebellar, vestibular, or proprioceptive. Cerebellar ataxia is caused by a disruption in transmission of sensory impulses from the vestibular system, cerebral cortex, and spinal cord via the cerebellum. Because the cerebellum does not initiate motor activity Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
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Seizures are a common neurologic presentation in middle-aged and older ferrets. As with paresis and ataxia, the most common cause of seizures is hypoglycemia secondary to insulinoma. Prolonged seizure activity can also result in hypoglycemia. Thus it is important to monitor blood glucose concentration long-term in patients that present with low blood glucose during or shortly after seizure activity. Other potential causes of seizures include toxin ingestion; CNS infection, inflammation, trauma, or neoplasia; and metabolic disturbances such as hepatic or renal failure. Idiopathic epilepsy has not been reported in ferrets.
Mallinckrodt Inc., St. Louis, MO); for MRI, use 0.2 mL/kg of gadolinium-diethylenetriamine pentaacetic acid (Gd-DTPA)49 (Fig. 10-2). If signs are localized to the spinal column, take spinal radiographs to look for possible fractures or bone abnormalities, such as proliferative or lytic lesions. Myelography is useful for localizing lesions of the spinal cord and determining a site for surgical approach if needed (see Fig. 10-3). If possible, perform a spinal tap to obtain a sample of cerebrospinal fluid (CSF) for analysis. Sites for CSF tap and myelography are the atlanto-occipital and the lumbar (L5-L6) regions. Use a 25-gauge spinal needle, as in canine and feline myelography; a suggested contrast medium is iohexol at 0.25 to 0.5 mL/kg50 (Fig. 10-4). Reference values for CSF fluid analysis are total protein 28 to 68 μg/μL and nucleated cells 0 to 8/μL.37 Some spinal lesions may be amenable to surgical resection or stabilization; many, however, carry a poor prognosis. For clinical signs localized to peripheral nerves or muscular defects, consult with a veterinary neurologist to perform electromyelography (EMG) and nerve conduction velocity (NCV) studies. Although normal values for ferrets have not been published, comparison of values from the contralateral limb (in unilateral disease) or from another, unaffected ferret will aid in interpreting the results. Tensilon testing (edrophonium HCL) and physostigmine testing can be performed in ferrets at standard canine doses (0.1 mg/kg IV).6,24,38,55 Obtain muscle biopsy samples in the same manner as in dogs and cats when indicated.
DIAGNOSIS OF POSTERIOR PARESIS, ATAXIA, AND SEIZURES
TREATMENT OF POSTERIOR PARESIS, ATAXIA, AND SEIZURES
Record the medical history and perform a complete physical examination, auscultating carefully for cardiac murmurs or arrhythmias. Ferrets may normally have a respiratory sinus arrhythmia. Palpate for peripheral pulses to evaluate the strength of cardiac contraction and to determine the presence of dropped or extra beats. Check mucous membranes for evidence of cyanosis. Palpate the abdomen carefully to detect discomfort, an abdominal mass, urinary calculi, or a distended urinary bladder. Include evaluation of the spine and neck for pain, fractures, and muscle asymmetry. Evaluate results of a complete blood count (CBC) and plasma biochemical analysis in any ferret with neurologic signs to rule out a metabolic or infectious component. Check the blood glucose concentration immediately to minimize artifactual decreases caused by faulty sample shipment or failure to separate plasma or serum from red blood cells. Correct any underlying abnormalities and reassess the animal for changes in neurologic status. Perform whole-body radiography and, if you suspect cardiac disease, perform echocardiography. If you suspect primary neurologic disease, perform a complete neurologic and orthopedic examination35 (Fig. 10-1). Characterize the signs as diffuse or focal, acute or chronic, progressive or static; localize the lesion to areas of brain or spinal cord.27 Evaluate reflexes and palpate for spinal cord pain or hyperesthesia. Keep in mind during examination that the ocular menace response in ferrets is normally diminished or absent. For signs that can be localized to the CNS, perform computed tomography (CT) or magnetic resonance imaging (MRI) if available. Administer an intravenous contrast medium to enhance brain lesions. For CT scanning, use 2.2 mL/kg of 400 mg/mL iodinated contrast medium (iothalamate sodium, Conray 400;
Treatment of posterior paresis, ataxia, and seizures in ferrets is tailored to the diagnosis. Follow the standard treatment regimens used for dogs and cats in managing a similar condition in a ferret. In any seizing ferret, the initial treatment must be directed toward arresting seizure activity (Fig. 10-5). Check the blood glucose concentration immediately in any animal presenting with seizures, ataxia, or other neurologic signs. If the glucose level is lower than 60 mg/dL, give an intravenous bolus of 50% dextrose solution (diluted 1:1 in crystalloid fluid) at a dose of 2 to 5 mL/kg or titrate to effect.10 Begin a dextrose drip infusion adequate to maintain normoglycemia while further diagnostic testing is done. Some ferrets require as much as 10% dextrose added to intravenous fluids to achieve normoglycemia. Administer prednisone or prednisolone to hypoglycemic patients to enhance hepatic gluconeogenesis and inhibit glycogenolysis. If an insulinoma is suspected, initiate additional therapy as indicated (see Chapter 8). If seizures persist and the blood glucose concentration is normal (or has been restored to normal), begin aggressive seizure management. Administer diazepam (0.5-1.0 mg/kg) intravenously10,24,38; if venous access is not readily available, administer diazepam or midazolam intramuscularly, intranasally (for more rapid absorption), or rectally. Double the dose for rectal or nasal administration.24 Repeat up to three times to arrest seizure activity. If grand mal or focal seizures persist, begin a constant-rate infusion of diazepam (0.1-1.0 mg/kg per hour added to IV fluids) or initiate phenobarbital (2-10 mg/kg per hour) as a constant rate infusion.38 Phenobarbital can also be administered by bolus; use 3 mg/kg slow IV q30-60min up to a total dose of 18 to 24 mg/kg, then 3 to 5 mg/kg q12h.24 Phenobarbital and diazepam can
but rather coordinates it, affected patients will demonstrate abnormal rate, range, or force of movement with intact strength. Paresis is not present with cerebellar dysfunction. Vestibular ataxia occurs when damage or disease affects the vestibular system within the inner ear. Motor activity is not initiated but is refined and coordinated via the vestibular system by controlling muscles used to maintain head position, eye movement, and equilibrium. Dysfunction results in loss of balance; animals often list or fall to one side and may have head tilt. Proprioceptive ataxia is caused by spinal disease, which will result in proprioceptive deficits that can be localized to the affected region of the spinal column. Trauma, intervertebral disk disease, and tumors arising within or compressing the spinal cord or nerves should be considered.35
SEIZURES
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NEUROLOGICAL EXAMINATION CHECKLIST FOR FERRETS 1. HISTORY a. When did the owner first notice the pet was not feeling well? b. What was the first sign? The next signs? c. Over what time period did they progress? d. Were any treatments administered? What was the result? e. How long has the pet been in the current condition? 2. MENTATION Alert Dull — Quiet, but alert and responsive Obtunded — Aware but not interested in the environment; only responds when stimulated Stuporous — Only responds to noxious stimulation Comatose — Unresponsive even to noxious stimulus 3. POSTURE AND GAIT/DESCRIPTION (Ambulatory, ataxia, paresis, what limbs affected?) General: Forelimbs: Hindlimbs: 4. CRANIAL NERVE EXAMINATION/DESCRIPTION (Normal or abnormal; describe if abnormal) Facial symmetry Facial sensation Facial expression Palpebral reflex R L Ocular position R L Ocular nystagmus (physiological present) Y N Ocular nystagmus (pathological, direction of fast phase) Menace Response R L PLR ( or ) Direct R L Consensual R L Gag reflex 5. PROPRIOCEPTION — scale: 0 (absent); 1 (decreased); 2 (normal) Conscious proprioception (paw placement): RF RH Table top testing RF RH (Placement of limbs when brought to table edge) Hopping RF RH Wheelbarrow____________
LF LF
LH LH
LF
LH
6. SPINAL REFLEXES — scale: 0 (absent), 1 (decreased), 2 (normal), 3 (increased), 4 (clonic) Withdrawal RF RH LF LH Patellar RH LH 7. SENSORY EVALUATION Sensation to all feet intact Any pain on spinal palpation? Normal range of motion in the neck?
Y Y Y
N N N
Fig. 10-1 Neurologic examination checklist for ferrets.
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A
C
B Fig. 10-2 Magnetic resonance imaging of a ferret with a cervical chordoma of C2-3. The spinal cord is compressed ventrally by the mass (arrow), which also extends into the bones of the skull. A, T1 Saggital view. B, Coronal view. C, Transverse view.
L
Fig. 10-3 Normal lateral myelogram of a ferret. Contrast can be seen as two parallel lines along the spinal cord.
be administered concurrently.38 Propofol can also be added; administer in aliquots of 2 mg/kg to effect, then 6 mg/kg per hour CRI to effect.24 If cerebral edema is suspected, administer dexamethasone (0.2 mg/kg) or prednisone or prednisolone (1 mg/kg IV) and mannitol (0.5-1.0 g/kg IV over 20 minutes).36,38 Once no seizures have occurred for 12 to 24 hours, taper the diazepam infusion slowly over the next 12 to 24 hours. Once seizures are controlled, use oral phenobarbital (1-2 mg/kg PO q8-12h) if necessary for long-term seizure management.38 Potassium bromide can also be used for seizure control; administer orally at 70 to 80 mg/kg per day if used alone or at 22 to 30 mg/kg per day in combination with phenobarbital.38 Check blood levels of phenobarbital within 2 to 3 weeks after starting therapy; potassium bromide levels may not reach a steady state for 60 to 90 days. Adjust dosages based on blood levels and clinical signs. The use of gabapetin, zonisamide, or levetiracetam has not been described in ferrets to date. Investigate any underlying disease stimulating the seizure activity after the patient is stable.
Physical therapy is an important but often overlooked adjunct treatment. For ferrets with paresis or paralysis and those debilitated or recumbent from seizures or metabolic disorders, begin passive range-of-motion exercises three to four times daily for affected limbs. This is essential to prevent contracture. Gently massage muscles to enhance blood flow. Implement active exercise as early as possible to preserve muscle tone and stimulate neural return.
SPINAL DISORDERS ALEUTIAN DISEASE Ferrets can be infected with Aleutian disease virus (ADV), a parvovirus that infects mink and ferrets (see Chapter 5). The virus is shed in saliva, urine, and feces; infection occurs by inhalation or ingestion. Clinical signs vary from mild incoordination to posterior ataxia, ascending paresis, persistent tremors, and quadriplegia.43,46 In one outbreak, clinical signs developed as soon as 24 hours or as long as 90 days after exposure.46 The most significant biochemical abnormality is a hypergammaglobulinemia, with serum gamma globulin concentrations increased to greater than 20% of the total serum protein concentration. However, not all affected ferrets have high gamma globulin concentrations.43,48 Histopathologic changes involving the brain and spinal cord include perivascular cuffing with lymphocytes
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C A
D B Fig.10-4 A, Cerebrospinal tap from the cisterna magna in a ferret. With the ferret in lateral recumbency at the edge of the table, the head is flexed so that the point of the nose is at 90 degrees to the long axis of the body. The wings of the atlas and the point of the occipital condyle are used as landmarks. B, Lumbar cerebrospinal fluid collection at L5-6. Only a few drops can be collected; to maximize the sample, the fluid can be collected directly into a microtainer (C) or hematocrit tube (D). and sometimes plasma cells, nonsuppurative meningitis, astrocytosis, mononuclear cell infiltration, and focal malacia.43,48 If you suspect Aleutian disease in a ferret, submit samples for testing by polymerase chain reaction (PCR), enzyme-linked immunosorbent assay (ELISA), or counterimmunoelectrophoresis (see Chapter 5). Tentative diagnosis is based on a positive result of ADV testing, a high gamma globulin concentration, and the presence of compatible clinical signs. Disease is confirmed at necropsy in suspected ferrets by demonstration of the presence of lymphocytic and plasmacytic infiltrates on histologic examination of tissue samples. Some infected ferrets may be asymptomatic but remain persistently infected, or the disease may be self-limiting and nonpersistent.48 The overall incidence of Aleutian disease in ferrets appears to be low; in one serologic study of 446 ferrets, the incidence of seropositive animals was 8.5%.48 Treatment consists of supportive care and isolation of suspected animals from unaffected ferrets.
SPINAL TUMORS Chordomas and Chondrosarcomas Chordomas are tumors that arise from remnants of notochord.29 In ferrets, these tumors develop most commonly at the tip of the tail,1,7,8,23,29 but they have also been described
in the cervical region.51 Ferrets with cervical chordoma can present with posterior paresis and ataxia localized to the area of the lesion.51 In such cases, perform spinal radiography and myelography to identify a site for surgical approach. Depending on location, these tumors may be amenable to surgical resection; however, the one reported case of recurrence and metastasis of a chordoma in a ferret was that of a cervical chordoma that had been surgically excised.51 In the tail, chordomas appear as lobulated, firm, nonencapsulated, ulcerated masses at or near the last caudal vertebra. Microscopically, these tumors consist of lobules of physaliphorous cells with areas of well-differentiated bone or cartilage throughout.8,23,51 Chondrosarcoma of the tail has also been described in ferrets.22,29 Clinical and morphologic descriptions are almost identical to those of chordoma. Differentiation must be made on the basis of immunohistochemical staining, with positive uptake of low-molecular-weight cytokeratin occurring in chordoma but not in chondrosarcoma.8,23 In ferrets with any distal tail mass, amputate several vertebrae proximal to the lesion. In cases of chordoma and chondrosarcoma, this is considered curative. Recurrence has not been reported.
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Seizure
Blood glucose 60
Blood glucose 60
Administer 50% dextrose IV diluted 1:1 with LRS or saline*, 1-3 cc to effect
Seizures stopped
Check history; has patient received glucose prior to obtaining blood glucose?
No supplemental dextrose administered
Still seizuring
Repeat glucose bolus Place on 2.5–10% IV dextrose drip, administer prednisone or adjust previous dose and begin treating for insulinoma
Seizures stopped
Still seizuring
Repeat blood glucose
Glucose 60 and still seizuring
Glucose 60
Administer diazepam* 0.5-1 mg/kg, IV bolus; repeat up to three times
Seizures stopped
Seizures continue, begin diazepam CRI 1 mg/kg/hr
Seizures stopped; continue diazepam CRI for 12-24 hours then taper slowly over 12-24 hours
Seizures continue; bolus phenobarbital 2-5 mg/kg
Seizures continue; phenobarbital CRI 2-10 mg/kg/hr
* If IV access not available, consider intraosseous catheter placement. Glucose and diazepam may be given orally or rectally. ** If prolonged seizures, consider steroids and mannitol for CNS swelling/cerebral edema.
Fig. 10-5 Treatment of seizures in ferrets. IV, Intravenous; CRI, continuous rate infusion.
Lymphoma T-cell lymphoma localized to the spine was identified in a ferret presenting with an acute onset of hindlimb paresis, proprioceptive deficits, diminished withdrawal responses, and an atonic bladder. Leukocytosis and lymphocytosis were present, and lysis of the vertebral body was evident radiographically. A mass associated with the soft tissues of the spine was identified by CT scan and ultrasonography. Ultrasound-guided biopsy of the mass confirmed lymphoma. The ferret did not respond
to immunosuppressive doses of prednisolone and deteriorated before radiation therapy was initiated.21
Fibrosarcoma Periosteal fibrosarcoma arising from the perivertebral connective tissue was diagnosed in a ferret that presented with hindlimb paralysis.34 Myelography was used to localize the lesion, and a dorsal laminectomy resulted in temporary improvement as well as providing samples for histologic diagnosis. Recurrence of the
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tumor was evident 35 days after surgery, and metastasis to the lungs was identified.34
INTERVERTEBRAL DISK DISEASE Intervertebral disk disease has been described rarely in ferrets;30,31 presumably, their long conformation and extreme flexibility of the body and spine may make them more resistant to herniation than other species. In two reports, ferrets with prolapse or herniation of an intervertebral disk presented with paresis or paralysis, accompanied by proprioceptive deficits caudal to the lesion.30,31 Clinical signs are similar to those seen in other mammals with intervertebral disk disease and may range in severity from mild proprioceptive deficits to complete paralysis. In one case, vertebral subluxation was also present radiographically.31 Confirm the diagnosis with myelography; CT and MRI are valuable tests for accurate localization of the lesion and planning of a surgical approach. Surgical decompression is the treatment of choice, and hemilaminectomy was performed successfully in both reported cases.30,31 Use guidelines for prognosis in disk disease in companion animals based on the presence or absence of deep pain perception and the duration of the clinical signs.
INTRACRANIAL DISORDERS CANINE DISTEMPER Canine distemper virus in the late stage affects the CNS of ferrets, although initial signs are usually localized to the respiratory and gastrointestinal tracts and the skin (see Chapter 7).16 Ferrets are highly susceptible to canine distemper virus and may seem to recover from the acute phase, only to die later from the neurotropic form of the disease. Affected ferrets can exhibit salivation, muscle tremors, seizures, and coma.16 The disease is almost 100% fatal, and infected animals may be a source of infection to other ferrets.16 Protect ferrets from distemper by vaccination (see Chapter 2).54
RABIES Although reports of rabies in ferrets are rare, ferrets are susceptible to this disease. Clinical signs include hyperactivity, lethargy, paresthesia (exaggerated grooming of a focal area), ataxia, and posterior paresis, sometimes followed by ascending paralysis.16,32 In experimentally induced rabies in ferrets, the mean incubation period was 28 to 33 days; the mean morbidity was 4 to 5 days.32,33 In one study virus was detected in the salivary glands of 63% and in saliva of 47% of the rabid ferrets.33 Suspect rabies in any unvaccinated ferret with clinical signs of neurologic disease and a history of exposure to rabid animals. A killed vaccine is approved for annual vaccination of ferrets (Imrab 3, Rhone Merieux, Athens, GA), but rabies protocol must be followed if the vaccinated animal bites a human (see Chapter 2). Be aware of local or state laws that may affect you or your clients and educate ferret owners at the time of vaccination.
OSTEOMA In ferrets, osteomas of the skull can arise from the zygomatic arch, parietal bone, or occipital bone.25,26,29,44 There is also a report of a multilobular osteoma originating from the neck and
extending from the base of the skull to the fifth cervical vertebra, causing extradural compression of the spinal cord.20 An osteoma presents as a firm, dense, bony mass arising from one of the bones of the skull. Although these are benign neoplasms, clinical signs are related to physical displacement or compression of normal structures.25,26,29,44 Obtain radiographs of any bony swelling to evaluate the extent of the lesion and the bone of origin. Biopsy may be difficult without surgical removal of the mass because of the extreme density of the tumor. The Jamshidi needle biopsy technique has been described.39 Histopathologic evaluation usually reveals compact lamellar bone, bony trabeculae, and mild to moderate osteoblastic and hematologic activity.25,26,44 Surgical removal is the treatment of choice and is usually curative if excision is complete.26
PRIMARY NEOPLASIA OF THE CENTRAL NERVOUS SYSTEM Although primary tumors of the CNS are uncommon overall, several recent case reports have documented neoplasms of the brain. One primary CNS neoplasm reported in a ferret was a granular cell tumor (sometimes called myoblastoma) in the cerebrum. Presenting signs included progressive head tilt, ataxia, and circling, followed by refractory seizures.45 There is also a report of a choroid plexus papilloma originating from the fourth ventricle in a ferret.47 This ferret presented with central vestibular signs including head tilt, circling, ataxia, hemiparesis, and loss of proprioception on the ipsilateral side of the lesion. Diagnosis of a mass in this case was made by the use of contrast-enhanced CT scanning and was histologically confirmed at necropsy.47 Consider intracranial neoplasia in a ferret with lateralizing progressive CNS signs or seizures and confirm with contrast CT or MRI imaging if available.
NEURONAL VACUOLATION Neuronal vacuolation was reported in one ferret that presented with rapidly progressive convulsions and incoordination. Although this is a change associated with transmissible spongiform encephalopathies, the case was ultimately confirmed to be a primary neuronal vacuolation; transmissible spongiform encephalopathy was ruled out based on negative testing.19
MUSCULOSKELETAL DISORDERS DISSEMINATED IDIOPATHIC MYOFASCIITIS Disseminated idiopathic myositis (DIM), or myofasciitis, is a recently described inflammatory disease of muscle and fascia that usually occurs in young ferrets ranging from 5 to 24 months of age. Clinical signs include acute to subacute onset of lethargy, pyrexia, recumbency, ataxia, posterior paresis, pain associated with movement or touch, anorexia, bruxism, and/or difficulty swallowing or drinking.18,40 Ferrets with this disease are most often recumbent and in extreme pain; they may vocalize or attempt to bite when touched or examined. They are usually febrile, with temperatures ranging from 104° to 108°F . Hematologic abnormalities associated with myofasciitis can be dramatic. The most remarkable is a mature neutrophilic leukocytosis, ranging from mild to severe, with reported total white blood cell (WBC) counts ranging from 8900 to 79,500 or higher. One author (NA) has seen WBC counts over 100,000 with this
CHAPTER 10 Musculoskeletal and Neurologic Diseases disease. The WBC count may also be normal at initial presentation but may then increase dramatically during the course of the disease. Mild to moderate anemia is also commonly present and is usually nonregenerative. Biochemical abnormalities reported with this disease are mild to moderate increases in concentrations of alanine aminotransferase (ALT) and glucose and mild hypoalbuminemia. Interestingly, increases in aspartate aminotransferase (AST) and creatine kinase (CK) concentrations are not associated with this disease.18,40 Gross pathologic changes at necropsy may be absent or may include esophageal dilation, red and white mottling of the esophagus, and white streaks in the heart, diaphragm, and intracostal muscles. Atrophy of the limb musculature and also the tongue and diaphragm is reported.18,40 Histopathologic lesions are moderate to severe areas of suppurative pyogranulomatous inflammation, most commonly in the skeletal muscle, esophagus, and heart but also identified in smooth muscle tunics and the submucosa of the stomach, small intestine, and urinary bladder. This profound inflammation occurs in the absence of any infectious or inciting agent. Inflammation is most commonly seen in the muscular tunics but also involves the submucosal and serosal tunics in some cases and frequently extends into the surrounding fascia and adipose tissue. Within inflamed foci, some small arteries, veins, and capillaries also have endothelial cell hypertrophy, neutrophil margination, and perivascular edema or fibrin deposition.18 Suspect myofasciitis in a recumbent ferret with unresponsive or cyclic pyrexia, muscle pain or sensitivity to touch, neutrophilic leukocytosis, and anorexia. Confirm the diagnosis antemortem with skeletal muscle biopsy from a hind limb. For necropsy diagnosis, be sure to include esophageal samples among those to be examined histologically. Treatment is often unrewarding, although some authors have reported success with aggressive supportive care along with the use of immunosuppressive doses of prednisolone (1 mg/kg PO q12h for 3 months or longer) and cyclophosphamide (10 mg/kg on days 1 and 14, then every 4 weeks for 3 months or longer) along with chloramphenicol (50 mg/kg PO q12h for 6-8 weeks). Aggressive supportive care can be prolonged (several weeks) and must include fluid therapy and nutritional supplementation; it should also include GI protectants and fever control.40 The author (NA) has treated some ferrets successfully with this protocol. The cause of this disease is unidentified to date. Investigations into bacterial, fungal, viral, environmental, husbandry, and vaccination status have failed to identify an agent. Results of PCR assays for Neospora caninum and Toxoplasma gondii have been negative, as have immunohistochemical stains for these organisms along with Sarcocystic neurona and feline and ferret coronavirus antigen.18
MYASTHENIA GRAVIS Acquired autoimmune myasthenia gravis was reported in a 7-month-old ferret. The clinical presentation included episodic hind limb weakness, flaccid paraparesis that progressed to tetraparesis, and mild megaesophagus. Diagnosis was made by nerve conduction velocity, which was diminished in the hind limbs; positive serologic testing for antiacetylcholine receptor antibodies; and response to neostigmine methylsulfate. Response to edrophonium hydrochloride can also be used in diagnostic testing. Treatment with pyridostigmine bromine (1 mg/kg PO q12h) initially produced a return to normal, but clinical
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signs progressed within 1 month, leading to euthanasia.6 Prednisolone at immunosuppressive doses is commonly used concurrently with pyridostigmine bromine in the treatment of myasthenia gravis and should be considered in the treatment of this disease. The authors have also diagnosed and treated this disease in a ferret.
MISCELLANEOUS DISEASES Systemic mycoses, which may contribute to CNS depression and lethargy, have been reported in ferrets.9,12 Cryptococcus species have been identified as a cause of meningitis in ferrets, including one that died from congestive heart failure after being treated with steroids for intervertebral disk disease.12 Blastomycosis has also been identified in ferrets, with multifocal granulomatous meningoencephalitis described in one ferret with systemic blastomycosis.12,28 One author (NA) has diagnosed a case of disseminated histoplasmosis in a ferret with fungal organisms present in the brain at necropsy. Diagnosis of these diseases is based on clinical signs, radiographic changes, and isolation of the causative organism. Impression smears of draining tracts or CSF analysis may be useful in identifying the organism.
OTHER METABOLIC DISEASES Toxemia of pregnancy is most common in young, primiparous jills late in gestation. Although hypocalcemia, hypophosphatemia, and ketosis may develop, neurologic signs are rare unless hepatic lipidosis is so severe that encephalopathy is present.3 One case of suspected pseudohypoparathyroidism was reported in a ferret that presented for seizures and had low serum calcium, high serum phosphorus, and high serum parathyroid hormone concentrations. This patient responded to lifelong treatment with vitamin D (dihydrotachysterol) and supplemental calcium.53 Paresis or paralysis of the hind limbs has also been described in ferrets with heartworm disease.2 An eosinophilic granulomatous infiltrate in the choroid plexus was reported in a ferret with diffuse eosinophilic gastroenteritis and multisystem involvement.15 Toxoplasmosis has been identified in ferrets; most likely disease develops after exposure to cat feces or raw meat.4,5,13 Copper toxicosis has been reported and was believed to be congenital in two ferrets with CNS depression. Diagnosis was based on histopathologic changes in the liver as well as copper levels.17 Iniencephaly in a litter of ferrets has also been described.52 Other congenital anomalies of the brain and spinal column probably occur but have not been reported. Additional diseases of the brain and spinal cord, as well as muscular and metabolic disorders that can cause clinical signs as described in this chapter, are likely to be identified as our diagnostic capability improves with newer technology.
References 1. Allison N, Rakich P. Chordoma in two ferrets. J Comp Pathol. 1989;100:161-176. 2. Antinoff N. Clinical observations in ferrets with naturally occurring heartworm disease, and preliminary evaluation of treatment with ivermectin with or without melarsomine. Proceedings. Heartworm Symposium, Austin: American Heartworm Society. 2002:45-47.
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3. Batchelder MA, Bell JA, Erdman SE, et al. Pregnancy toxemia in the European ferret (Mustela putorius furo). Lab Anim Sci. 1999;49:372-379. 4. Bell JA. Parasites of domesticated pet ferrets. Compend Contin Educ Pract Vet. 1994;16:617-620. 5. Besch-Williford C. Biology and medicine of the ferret. Vet Clin North Am Small Anim Pract. 1987;17:1155-1183. 6. Couturier J, Huynh M, Boussarie D, et al. Autoimmune myasthenia gravis in a ferret. J Am Vet Med Assoc. 2009;235:1462-1466. 7. Dillberger JE, Altman NH. Neoplasia in ferrets: eleven cases with a review. J Comp Pathol. 1989;100:161-176. 8. Dunn DG, Harris RK, Meis JM, et al. A histomorphologic and immunohistochemical study of chordoma in 20 ferrets. Vet Pathol. 1991;28:467-493. 9. DuVal-Hudelson KA. Coccidioidomycosis in three European ferrets. J Zoo Wildl Med. 1990;21:353-357. 10. Ford RB, Mazzaferro EM. Kirk and Bistner’s handbook of veterinary procedures and emergency treatment. St. Louis, MO: Saunders Elsevier; 2006. 11. Fox JG. Bacterial and mycoplasmal diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott, Williams & Wilkins; 1998:321-354. 12. Fox JG. Mycotic diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:393-403. 13. Fox JG. Parasitic diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:375-391. 14. Fox JG, Murphy JC, Ackerman JI, et al. Proliferative colitis in ferrets. Am J Vet Res. 1982;43:858-864. 15. Fox JG, Palley LS, Rose R. Eosinophilic gastroenteritis with Splendore-Hoeppli material in the ferret. Vet Pathol. 1992;29:21-26. 16. Fox JG, Pearson RC, Gorham JR. Viral diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:335-374. 17. Fox JG, Zeman DH, Mortimer JD. Copper toxicosis in sibling ferrets. J Am Vet Med Assoc. 1994;205:1154-1156. 18. Garner MM, Ramsell K, Schoemaker NJ, et al. Myofasciitis in the domestic ferret. Vet Pathol. 2007;44:25-38. 19. Hamir AN, Miller JM, Yaeger MJ. Neuronal vacuolation in an adult ferret. Can Vet J. 2007;48:389-391. 20. Hanley CS, Geiger T, Frank P. What is your diagnosis? Multilobular osteoma (MLO). J Am Vet Med Assoc. 2004;225:1665-1666. 21. Hanley CS, Wilson GH, Frank P, et al. T cell lymphoma in the lumbar spine of a domestic ferret (Mustela putorius furo). Vet Rec. 2004;155:329-332. 22. Hendrick MJ, Goldschmidt MH. Chondrosarcoma of the tail of ferrets. Vet Pathol. 1987;24:272-273. 23. Herron AJ, Brunnert SR, Ching SV, et al. Immunohistochemical and morphologic features of chordomas in ferrets. Vet Pathol. 1990;27:284-286. 24. Jasani S. Saunders solutions in veterinary practice; small animal emergency medicine. St. Louis, MO: Saunders Elsevier; 2011. 25. Jensen WA, Myers RK, Liu CH. Osteoma in a ferret. J Am Vet Med Assoc. 1985;187:1375-1376. 26. Jensen WA, Myers RK, Merkley DF. Diagnostic exercise: a bony growth of the skull in a ferret. Lab Anim Sci. 1987;37:780-781. 27. Lawes INC, Andrews PLR. The neuroanatomy of the ferret brain. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:71-102. 28. Lenhard A. Blastomycosis in a ferret. J Am Vet Med Assoc. 1985;186:70-72. 29. Xiantang Li, Fox JG. Neoplastic diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Lippincott Williams & Wilkins; 1998:405-447.
30. Lu D, Lamb CR, Patterson-Kane JC, et al. Treatment of a prolapsed lumbar intervertebral disk in a ferret. J Small Anim Pract. 2004;45:501-503. 31. Morera N, Valls X, Mascort J. Intervertebral disk prolapse in a ferret. Vet Clin North Am Exot Anim Pract. 2006;9:667-671. 32. Niezgoda M, Briggs DJ, Shadduck J, et al. Pathogenesis of experimentally induced rabies in domestic ferrets. Am J Vet Res. 1997;58:1327-1331. 33. Niezgoda M, Briggs DJ, Shadduck J, et al. Viral excretion in domestic ferrets (Mustela putorius furo) inoculated with a raccoon rabies isolate. Am J Vet Res. 1998;59:1629-1632. 34. Ohta G, Kobayashi M, Yanai T, et al. A case of fibrosarcoma on the perivertebral surface of a ferret with hind limb paralysis. Exp Anim. 2008;57:397-400. 35. Oliver JE, Lorenz MD, Kornegay JN. Handbook of veterinary neurology. 3rd ed. Philadelphia: WB Saunders; 1997. 36. Papich MG. Saunders handbook of veterinary drugs. 3rd ed. St. Louis: Elsevier Saunders; 2011. 37. Platt SR, Dennis PM, McSherry IJ, et al. Composition of cerebrospinal fluid in clinically normal adult ferrets. Am J Vet Res. 2004;65:758-760. 38. Plumb DC. Veterinary drug handbook. 6th ed. Ames: Blackwell Publishing Professional; 2008. 39. Powers BE, LaRue SM, Withrow SJ, et al. Jamshidi needle biopsy for diagnosis of bone lesions in small animals. J Am Vet Med Assoc. 1988;193:205-210. 40. Ramsell KD, Garner MM. Disseminated idiopathic myofasciitis in ferrets. Vet Clin North Am Exot Anim Pract. 2010;13:561-575. 41. Richardson JA, Balabuszko RA. Ibuprofen ingestion in ferrets: 43 cases. J Vet Emerg Crit Care. 2001;11:53-59. 42. Rosenthal K. Ferrets. Vet Clin North Am Small Anim Pract. 1994;24:1-23. 43. Rozengurt N, Stewart S, Sanchez S. Diagnostic exercise: ataxia and incoordination in ferrets. Lab Anim Sci. 1995;45:432-434. 44. Ryland LM, Gogolewski R. What’s your diagnosis? J Am Vet Med Assoc. 1990;197:1065-1066. 45. Sleeman JM, Clyde VL, Brennerman KA. Granular cell tumour in the central nervous system of a ferret (Mustela putorius furo). Vet Rec. 1996;138:65-66. 46. Une Y, Wakimoto Y, Nakano Y, et al. Spontaneous Aleutian disease in a ferret. J Vet Med Sci. 2000;62:553-555. 47. van Zeeland Y, Schoemaker N, Passon-Vastenburg M, et al. Vestibular syndrome due to a choroid plexus papilloma in a ferret. J Am Anim Hosp Assoc. 2009;45:97-101. 48. Welchman D deB, Oxenham M, Done SH. Aleutian disease in domestic ferrets: diagnostic findings and survey results. Vet Rec. 1993;132:479-484. 49. Westbrook C, Kaut C. MRI in practice. 2nd ed. Oxford: Blackwell Science; 1998. 50. Widmer WR, Blevins WE. Veterinary myelography: a review of contrast media, adverse effects, and technique. J Am Anim Hosp Assoc. 1991;27:163-176. 51. Williams BH, Eighmy JJ, Berbert MH, et al. Cervical chordoma in two ferrets. Vet Pathol. 1993;30:204-206. 52. Williams BH, Popek EJ, Hart RA, et al. Iniencephaly and other neural tube defects in a litter of ferrets. Vet Pathol. 1994; 31:260-262. 53. Wilson GH, Greene CE, Greenacre CB. Suspected pseudohypoparathyroidism in a domestic ferret. J Am Vet Med Assoc. 2003;222:1093-1096. 54. Wimsatt J, Jay MT, Innes KE, et al. Serologic evaluation, efficacy, and safety of a commercial modified-live canine distemper vaccine in domestic ferrets. Am J Vet Res. 2001;62:736-740. 55. Wingfield WE, Raffe MR. The veterinary ICU book. Jackson: Teton NewMedia; 2002.
CHAPTER
11
Soft Tissue Surgery
Teresa Lightfoot, DVM, Diplomate ABVP (Avian), Jonathan Rubinstein, DVM, Sean Aiken, DVM, MS, Diplomate ACVS, and Lori Ludwig, VMD, MS, Diplomate ACVS
General Surgical Principles in Ferrets Surgery of Cutaneous Neoplasia Exploratory Laparotomy Gastrointestinal System Salivary Mucocele Resection Intestinal Surgery Liver Biopsy Gallbladder Surgery Endocrine System Surgery of the Adrenal Gland Pancreatic Surgery Splenectomy Urogenital System Nephrectomy Cystotomy Perineal Urethrostomy Paraurethral/Prostatic Cysts Ovariohysterectomy Ovarian Remnant Pyometra Castration Preputial Masses Miscellaneous Surgical Procedures Anal Sacculectomy Heartworm Disease—Caval Syndrome
GENERAL SURGICAL PRINCIPLES IN FERRETS Soft tissue surgical procedures in domestic ferrets are similar to those performed in other small animal species treated in a veterinary practice. This chapter describes soft tissue surgical procedures that are common in ferrets. Because ferrets may be afflicted with surgical diseases similar to those of other small mammal species, a basic small animal surgery text should be Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
used in conjunction with this chapter to provide a complete understanding of the various surgical procedures. As with dogs and cats, appropriate preoperative patient preparation includes placing an intravenous or intraosseous catheter. Provide intraoperative crystalloid fluid therapy at the rate standard for small animals (5-10 mL/kg per hour) as well as appropriate antibiotic coverage when indicated. Appropriate pain management is paramount and can be provided with opiods, nonsteroidal anti-inflammatory drugs, epidurals, and local blocks or a combination thereof (see Chapter 31). In performing surgical procedures in ferrets, the use of some specialized equipment (which should be kept on hand) is highly recommended. Endotracheal tubes in sizes from 2.5 to 4.0 mm should be available. Microsurgical instrumentation and magnifying loupes greatly simplify surgical procedures (Fig. 11-1). The surgical suite should be stocked with a variety of synthetic absorbable suture materials ranging in size from 4-0 to 6-0 with both cutting and taper needles. Various forms of hemostasis, including monopolar and bipolar radiosurgical units, absorbable hemostatic material such as Gelfoam (Pfizer Inc., New York, NY) and Surgicel (Ethicon, Inc., Johnson and Johnson Healthcare, Piscataway, NJ), or vascular sealing devices should be available. Equip the surgical preparation area, surgical suite, and recovery area with devices needed to maintain the ferret’s body temperature throughout the anesthetic procedure. Circulating hot-water blankets, warm-air circulators, fluid warmers for intravenous fluids, and warm water bottles may be used. Heat lamps can burn patients and should be used with caution. Self-adherent clear plastic drapes are useful to allow visibility of these small patients during surgery. These drapes also aid in keeping patients warm and dry and can eliminate the need for towel clamps. Monitoring should include pulse oximetry, electrocardiography (ECG), blood pressure, and body temperature.
SURGERY OF CUTANEOUS NEOPLASIA Ferrets commonly present with a variety of cutaneous neoplasms (see Chapter 9); cutaneous neoplasia has been reported to be the third most common tumor type in ferrets.21 While 141
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Fig. 11-1 Specialized surgical instruments. Top left to right: Right-angle forceps, neonatal Satinsky forceps, iris scissors, mosquito forceps, Bishop-Harmon forceps, vascular forceps. Bottom: Vascular clips and applicator, cotton-tipped applicators. Right: Surgical magnifying loupes: custom 4.5× magnification, adjustable 3.5× magnification.
some references have indicated that the most common dermal neoplasms are, in order, basal cell tumors, mast cell tumors, and squamous cell carcinomas, other retrospective studies have found fibromas to be the third most common tumor type, after basal cell tumors and mast cell tumors.31 Other less commonly identified cutaneous and subcutaneous tumors types include cutaneous hemangioma, hemangiosarcoma, lymphosarcoma, adenocarcinoma, piloleiomyosarcoma, rhabdomyosarcoma, neurofibroma, and cutaneous melanoma.18,31,36,47 Two cases of subcutaneous neoplasms histologically consistent with adrenocortical neoplasia have been reported; because these tumors were located along the ventral midline in young, spayed female ferrets, the authors speculated that ovarian or adrenal tissue had been seeded during ovariohysterectomy and later underwent malignant transformation.42 Signalment, history, and physical examination findings depend on tumor type. In one retrospective study, the mean age of presentation was 4.3 years31 with a range of 8 months to 9 years and slightly higher prevalence in females than in males (58% vs. 42%). Most cutaneous neoplasms in ferrets are not malignant. Basal cell tumors are generally well demarcated with minimal invasion of adjacent tissue. Ferret mast cell tumors are also benign tumors, more closely resembling the feline mast cell tumor rather than the more aggressive canine version. Preoperative treatment with corticosteroids and antihistamines may decrease mast cell tumor size and inflammation. Consequently wide surgical margins are generally not required. Despite the generally benign nature of most ferret cutaneous neoplasms, do not omit appropriate preoperative workup for potential neoplastic disease. Hematologic testing, plasma biochemical analysis, abdominal ultrasound, and thoracic radiographs to screen for concurrent diseases and anesthetic suitability should be considered, depending on the case and presentation. When appropriate, further diagnostic tests may include bone marrow aspiration as well as regional lymph node aspiration or biopsy, computed tomography, or magnetic resonance imaging. Obtain fine-needle aspirates of the mass
or masses and submit samples for cytologic evaluation to plan appropriate surgical margins and provide owners with prognostic information before surgery. While cytologic examination may not provide a definitive diagnosis, a general cell type as well as distinction between malignant and benign processes can often be appreciated.2,25 Fine-needle aspirates can generally be performed with sedation, restraint, and local blocks; to prevent tumor seeding, plan approaches such that the needle tract is through areas slated for removal. Full histologic analysis is recommended for all excised tumors to confirm the diagnosis. Surgery of cutaneous and subcutaneous neoplasia in ferrets is similar to that in other species. A variety of surgical modalities can be used to successfully remove cutaneous neoplasms. Scalpel, laser, and radiosurgical techniques have been described; a full review is beyond the scope of this chapter and the reader is directed to one of the excellent review articles for further information.25 Regardless of the technique used, pay attention to gentle tissue handling and rapid hemostasis. Take care during patient preparation not to spread or degranulate volatile masses such as mast cell tumors; therefore avoid aggressive palpation or scrubbing.43 Surgical margins depend on the tumor type and location. Cryosurgery has also been used for superficial tumors. Cryosurgery utilizes freezing/thawing cycles to destroy cancerous cells through the generation of ice crystals within the cells. Generally, two freeze/thaw cycles are recommended.17 Because of the extensive tissue necrosis adjacent to the mass, closure is usually not performed; for the same reason, make efforts to obtain a definitive diagnosis before cryosurgical treatment. Postoperative care, including pain control when required, is similar to that implemented in other species. As ferrets are not amenable to Elizabethan collars, we generally perform a subcuticular closure with a small-gauge (4-0) absorbable suture in resecting dermal masses. Overnight hospitalization is rarely required unless the mass was very large.
EXPLORATORY LAPAROTOMY For a complete exploratory laparotomy, make a ventral midline incision beginning at the xiphoid and extending to the pubis (in the female) or to just cranial to the prepuce and extending parapreputial if needed (in the male). Ferrets have relatively thin skin and little subcutaneous tissue, so a light touch with the scalpel blade is warranted. The linea alba of ferrets is a wide, thin structure that is easily identified (Fig. 11-2, A). Gently grasp the linea with forceps and lift it away from the abdominal contents before penetrating the abdomen with a No. 15 scalpel blade. Extend the initial incision in the linea with scissors or a scalpel blade (Fig. 11-2, B). Be extremely cautious in penetrating the abdominal cavity or extending the incision to avoid damaging dilated or enlarged organs (most commonly the spleen). Also, so as not to penetrate the diaphragm, use caution in extending the incision in the linea cranially. To help gain exposure to the abdominal cavity, use small Balfour retractors with moistened gauze sponges. As with any exploratory laparotomy, use a thorough and systematic approach to examine the entire abdominal cavity. Remember that hypothermia is a major intraoperative complication, especially when a large abdominal incision is required. Monitor body temperature and be prepared to utilize multiple methods of thermal support, including forced warm-air mechanisms, overhead light/heat support, and very warm abdominal saline lavage. After completing the procedure, close the linea with 3-0 or 4-0 absorbable monofilament suture material
CHAPTER 11 Soft Tissue Surgery
L
A
B Fig. 11-2 A, Approach to ferret exploratory laparotomy. Note the wide linea alba (L) and minimal subcutaneous fat. The adherent plastic sheeting is applied to the ventral abdominal skin to keep the ferret dry and warm during the surgical procedure. B, Approach to ferret exploratory laparotomy by extending the incision in the linea alba with fine scissors. Note that the abdominal wall is elevated to prevent damage to the underlying abdominal organs.
(for example, polydioxanone [PDS]) in a simple continuous or interrupted pattern. Close subcutaneous tissues with fine absorbable suture material. A subcuticular pattern may be used to complete the closure. Alternatively, cyanoacrylic tissue glue may be used by applying the material after opposing the skin edges. Do not place tissue glue within the subcutaneous tissues because it elicits a foreign-body reaction. Skin sutures may also be used.
GASTROINTESTINAL SYSTEM SALIVARY MUCOCELE RESECTION Ferrets have five pairs of salivary glands: parotid, mandibular, sublingual, molar (buccal), and zygomatic.16 Although mucoceles are uncommon, they have been reported in ferrets (see Chapter 3).3,26,28 The zygomatic and molar (buccal) glands are most commonly affected. Affected ferrets are presented with unilateral facial or periorbital swelling or exophthalmos, and a fine-needle aspirate of the mass reveals a thick, mucoid fluid. Aspiration may provide temporary relief from the swelling, but treatment should involve marsupialization or surgical removal of the involved gland or glands. A thorough knowledge of the regional anatomy
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is required so that all of the involved glandular material is removed. As described in dogs, the zygomatic arch can be partially resected to facilitate exposure of the zygomatic gland. Although there are few reports of mucoceles in ferrets, the prognosis appears to be good after surgical removal of the affected gland.3,26,28 Mucoliths are common in the parotid salivary gland of ferrets. Although these are generally asymptomatic, affected ferrets are used as models in studies of human salivary mucoliths.45,46
INTESTINAL SURGERY One of the most common surgeries performed in young ferrets is removal of ingested foreign material from the stomach and small intestine. Foam rubber sponges, rubber objects, and cork are the most common foreign bodies removed.27 Most ingested foreign material becomes lodged in the small intestine or stomach, but esophageal foreign bodies have been reported.9,27 Trichobezoars (hairball foreign bodies) may result from normal or excessive grooming behavior and tend to lodge in the stomach.27 Although vomiting is not as frequent in ferrets with gastrointestinal (GI) foreign bodies as in other domestic species,27 clinical signs in ferrets with foreignbody obstruction are similar to those of other small animal patients with GI obstruction. These signs include diarrhea, anorexia, lethargy, bruxism, and pawing at the mouth. Foreign bodies in the intestinal tract may also be discovered during exploratory laparotomy in ferrets that have no clinical signs attributable to intestinal obstruction. Ferrets presented with intestinal foreign bodies are commonly less than 18 months of age.20 Physical examination findings depend on the duration of illness and the location and consistency of the foreign material. The ferret may be lethargic and dehydrated, or it may simply present for anorexia or weight loss. On physical examination, the foreign body may be detected as a painful or nonpainful palpable intestinal mass. Holding the ferret upright during abdominal palpation may aid in the palpation of intestinal foreign bodies located in the stomach and proximal small intestine. Survey radiographs may show loops of small intestine dilated with gas or fluid, dilation of the stomach, or a radiodense foreign body. On abdominal ultrasound examination, fluid-filled loops of intestine may be seen proximal to the obstruction; often the object itself can be visualized as a hyperechoic intraluminal structure. As with other species, surgery to relieve an intestinal foreignbody obstruction is considered an emergency procedure and should be done as soon as the ferret is rehydrated and stabilized. If GI perforation has occurred, peritoneal effusion is often present. The fluid should be sampled and evaluated for the presence of bacteria, lower glucose, and/or higher lactate than the peripheral blood. Although the latter two diagnostic tests have not been validated for use in ferrets, they may assist in the diagnosis of septic abdomen and allow appropriate presurgical planning and prognostication. This fluid should also be submitted for culture and sensitivity testing. Esophageal foreign bodies are rare and should be treated as recommended in other small animal species. If possible, remove the foreign body by endoscopy or by advancing it into the stomach for retrieval by gastrotomy. An esophagotomy may
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SECTION I Ferrets
A
B Fig. 11-3 A, Gastric foreign body in a ferret. The object was palpated during a routine exploratory laparotomy for an unrelated problem. B, Removal of a gastric trichobezoar through a gastrotomy incision in a ferret. Note the stay sutures supporting the stomach and the sponges isolating the area to protect against contamination by intestinal contents.
be performed through a right lateral thoracotomy, median sternotomy, or midline cervical approach, but this procedure can be complicated by postoperative leakage or stricture formation.9 Most intestinal foreign bodies can be retrieved during an exploratory laparotomy. The ferret’s intestine lacks a distinct ileum; instead, the jejunoileum extends from the duodenojejunal flexure to the ascending colon.16 Evaluate the entire intestinal tract, as multiple foreign bodies may be present, and perform gastrotomy, enterotomy, or resection and anastamosis as in other small animals. Because the ferret’s intestinal tissue is very thin and fragile, gentle tissue handling is needed to prevent iatrogenic damage to the intestines. Isolate the intestinal surgery site from the abdominal cavity with moistened sponges. For a gastrotomy, stabilize the stomach with full-thickness stay sutures of 4-0 nylon before making an incision (Fig. 11-3). Close the gastrotomy incision in one or two layers by using a simple interrupted or continuous suture pattern with 4-0 or 5-0 monofilament absorbable suture. For an enterotomy, use a No. 15 or 11 scalpel blade to make a longitudinal incision in healthy tissue distal to the obstruction and then extend the incision with fine scissors. The foreign material should then be gently milked to the enterotomy site and out. Close the enterotomy site with 4-0 or 5-0 monofilament suture in a transverse simple interrupted pattern. The luminal diameter of the jejunum is very small, so careful tissue handling is required to minimize the chance of intestinal strictures. Resection and anastomosis is as described for other small animal species. The section of bowel to be removed should be isolated, occluded with Doyen clamps or sterile bobby pins, and resected. Take care to ensure an adequate mesenteric blood supply to the ends that are to be anastamosed. Place one suture at the mesenteric border
and a second simple interrupted suture on the antimesenteric border. As with other species, use full-thickness simple interrupted sutures with an absorbable small-gauge suture to complete the anastamosis. As with any intestinal surgery, carefully place laparotomy sponges and exteriorize the affected portion of the GI tract to minimize the chance of abdominal contamination. If contamination is suspected, lavage the abdominal cavity copiously with sterile saline solution. Change instruments, drapes, and gloves before abdominal closure. For a more complete description, the reader is referred to standard small-animal surgery texts. Water can be offered as soon as 2 hours after surgery, and small amounts of food should be offered after 2 to 4 hours, assuming that the ferret is awake and alert. Continue intravenous fluid therapy until the ferret is eating well. Overall, the prognosis is very good, but possible complications are intestinal leakage and peritonitis as well as stricture formation with possible reobstruction.
LIVER BIOPSY Ultrasound-guided fine-needle aspirates or biopsies are most commonly performed in ferrets to diagnose suspected liver disease. However, a liver biopsy may be performed during an exploratory laparotomy to help diagnose suspected or unsuspected liver disease such as hepatic lipidosis, lymphosarcoma, cholangiohepatitis, or neoplasia. If all lobes have a similar appearance and palpate similarly, a random liver biopsy is indicated. If a section of liver is protruding, a guillotine suture can be used. For this technique, place a preformed encircling ligature of 4-0 monofilament absorbable suture material around the protruding section of liver. Tighten the ligature until it has crushed through the hepatic parenchyma (Fig. 11-4). After completing several throws in the knot, excise the sample 1 to 2 mm distal to
CHAPTER 11 Soft Tissue Surgery
Fig. 11-4 The guillotine liver biopsy technique is performed by placing a ligature around the area of interest at the edge of a liver lobe and slowly tightening the knot. The biopsy specimen is then divided approximately 2 mm distal to the encircling ligature.
the ligature with Metzenbaum scissors or a scalpel blade. Take care to avoid crushing the sample and destroying architecture with forceps in handling it. If a specific area of the liver is of interest, the tissue sample can be obtained by the transfixion method or with a 6-mm skin biopsy punch. For the transfixion method, place a ligature through the liver lobe approximately 8 to 10 mm from its edge. Tighten the ligature to crush through the parenchyma of half of the desired biopsy specimen. Make an additional throw at a right angle to the first ligature, tightening the ligature to crush the parenchyma for the second half of the specimen. Remove the sample 1 to 2 mm distal to the crushed area with a scalpel blade or Metzenbaum scissors. If the area of interest does not lie near the edge of the liver lobe, a 6-mm biopsy punch can be used. Push the biopsy punch through the liver parenchyma in the desired area, making sure not to penetrate the opposite surface of the liver (Fig. 11-5). Use extra caution if the biopsy site is close to the hilum so that no more than one half of the thickness of the liver lobe is penetrated. Remove the biopsy punch and separate the biopsy sample from the liver with scissors, being careful not to crush the sample. If culture and sensitivity testing of a liver sample is warranted, it can be obtained at this time. Control bleeding either by filling the defect with gelatin sponge hemostatic material and applying digital pressure for 3 to 5 minutes or by suturing the liver capsule with a fine absorbable monofilament suture in a cruciate pattern.
GALLBLADDER SURGERY Choleliths are rare in domestic species and may occasionally be found in ferrets. Choleliths have been found incidentally in ferrets during exploratory abdominal surgery for other conditions and have been observed at surgery as an explanation for anorexia, fever, and depression. In four cases seen at the Animal Medical Center, choleliths were observed in older ferrets (mean age 6.5 years; range 5 to 8 years). Liver enzyme concentrations were high in only 1 of the 4 ferrets.
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The biliary system of ferrets is similar to that of other domesticated animals and is composed of three hepatic ducts feeding into a common bile duct (Fig. 11-6). The gallbladder empties into the cystic duct, which joins the central hepatic duct to become the common bile duct. The main pancreatic duct joins the common bile duct before it empties into the duodenum at the major duodenal papilla.33 Choleliths can be located in the gallbladder, or they can obstruct the common bile duct as choledocholiths. Therefore thoroughly examine the biliary tree during exploratory abdominal surgery for gallstones. If the obstruction is located in the common bile duct, attempt to retropulse the stones by gently milking them backward into the gallbladder with your fingertips. Alternatively, make an incision in the antimesenteric border of the duodenum at the level of the major duodenal papilla. Cannulate the common bile duct with a 25-gauge catheter and flush the stones into the gallbladder; the duodenal papilla is generally located 2 to 3 cm distal to the cranial duodenal flexure. Once the stones are in the gallbladder, check the common bile duct for patency by flushing the duct with a 3.5-Fr catheter. If it is patent, perform a standard cholecystectomy (as in other domestic species) to prevent recurrent cholecystitis.24 Gently elevate the gallbladder from the hepatic surfaces of the right medial and quadrate liver lobes with sterile cotton-tipped applicators (see Fig. 11-6). Use hemostatic sponges on the liver surface to control hemorrhage. After dissecting the gallbladder and cystic duct to the level of the central hepatic duct, double-ligate the cystic duct with fine absorbable or nonabsorbable suture material or vascular clips and transect it. Submit sections of the gallbladder for bacterial culture and histopathologic examination and submit the gallstones for crystallographic analysis. To date, all ferrets with gallstones have had a histologic diagnosis of cholecystitis, with 1 of 4 ferrets having a positive bacterial culture. All choleliths have been composed primarily of bile pigment and bile salts.
ENDOCRINE SYSTEM SURGERY OF THE ADRENAL GLAND Adrenal neoplasia is common in ferrets. Cortical hyperplasia, adenomas, and carcinomas secreting estrogen and other steroid hormones are the most common tumor types (see Chapter 7).4,28 Other types of adrenal gland neoplasia have been reported, although with a much lower frequency. These include spindle cell tumors, pheochromocytomas, teratomas, and myelolipomas.21 Adrenocortical disease in ferrets is not Cushing’s disease. Instead of secreting excess cortisol, affected adrenal glands produce estrogens and androgens.22,38,48 The most common clinical sign associated with adrenocortical disease is alopecia, which begins on the tail and progresses cranially along the dorsum, flanks, and body.19,49,50 Affected females often present with enlarged vulvas, and affected males may show sexual behavior, increased musky odor, or present with stranguria or urinary obstruction secondary to an enlarged prostate or paraurethral cysts. Ferrets typically present with clinical signs after 3 years of age (range 1 to 7.5 years). One retrospective study found that adrenocortical disease occurred more commonly in females; however, others have found an equal distribution between the
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SECTION I Ferrets
A
B
C Fig. 11-5 The punch liver biopsy technique allows a more selective biopsy. A, A 6-mm skin biopsy punch is pushed through the liver parenchyma without penetrating the capsule on the opposite side of the liver lobe. B, After removing the biopsy punch, the biopsy sample is separated from the liver with fine scissors, with care not to crush the sample. C, The defect is filled with absorbable hemostatic sponge and digital pressure is applied for 5 minutes. The hemostatic sponge may be removed or left in place to be absorbed. If bleeding continues, the liver capsule can be sutured with fine absorbable suture material in a cruciate pattern.
sexes.37,49,50 Most ferrets with adrenocortical disease have been neutered before 6 weeks of age; the disease occurs less commonly and at an older age in sexually intact ferrets.37 An association between adrenocortical disease in ferrets and neutering at an early age is suspected. This association is postulated to be caused by loss of negative gonadal feedback to the release of gonadotropin-releasing hormone (GnRH) from the hypothalamus, resulting in persistent stimulation of the adrenal cortex by luteinizing hormone (LH) and follicle-stimulating hormone (FSH). Both FSH and LH receptors have been isolated in the adrenal glands of affected ferrets; results of one study in ferrets showed a correlation between age at neutering and age at onset of adrenocortical disease.41 Genetic factors are also potentially involved in predisposition to this disease (see Chapter 7). However, more studies are needed to prove a cause-and-effect relationship. Adrenocortical disease in ferrets is often tentatively diagnosed by clinical signs. Dorsal symmetric bilateral alopecia with or without pruritus is a common presentation. Alternatively,
hair loss may begin on the tail. Differential diagnoses for these presentations include seasonal hair loss, external parasites, extended estrus, ovarian remnants, or severe metabolic disturbances.4 Physical examination may reveal a mass palpable cranial to the left kidney. The cranial aspect of the right kidney is generally not accessible to palpation. Abdominal ultrasound is used to identify an enlarged adrenal gland or glands and to determine whether the disease is unilateral or bilateral. Colorflow Doppler can be used to determine the extent of caudal vena cava invasion when a right adrenal gland is affected. Ultrasound also screens for the presence of other disease processes in abdominal organs. The diagnosis can be confirmed with an adrenal hormone panel that measures plasma concentrations of 17-hydroxyprogesterone, androstenedione, and estradiol.38 Before surgery, a complete blood count and plasma biochemical analysis should be performed on each patient to evaluate overall health. Commonly, hypoglycemia is present because of concurrent insulinoma. Whether or not the ferret exhibits hypoglycemia, always palpate the pancreas intraoperatively
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GB
B
A
Fig. 11-6 A, The biliary tree and gallbladder. A, Gallbladder; B, central hepatic duct; C, left hepatic duct; D, right hepatic duct; E, main pancreatic duct; F, major duodenal papilla entering the duodenum approximately 2.75 cm distal to the pylorus. B, Intraoperative view of the gallbladder (GB) and biliary tree of a ferret with gallstones. Note that the gallbladder is being elevated from the underlying liver lobes with a cotton-tipped applicator and is grasped with a Babcock forceps. The gallbladder is then ligated at its base, where the cystic duct joins the central hepatic duct.
VC AO
PH
Liver
KD
Fig. 11-7 Normal adrenal anatomy. The hepatorenal ligament
Fig. 11-8 Left adrenalectomy is usually simple because of the
has been incised to allow retraction of the caudate liver lobe and visualization of the right adrenal gland. The right adrenal gland is adhered to the wall of the caudal vena cava (VC). If neoplastic, it often extends dorsally along the vessel wall (indicated by dashes). AO, Aorta; KD, kidney; PH, phrenicoabdominal vein.
distance of the gland from the caudal vena cava. Hemostatic clips are useful for ligating the phrenicoabdominal vein as it courses from lateral to medial over the ventral surface of the gland.
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for detection of insulinomas; if encountered, these should be resected (see “Pancreatic Surgery,” below). Cardiac disease is common in middle-aged and older ferrets, and cardiac function should be evaluated before surgery. Thoracic radiographs, an ECG, and echocardiography are recommended. Aortic insufficiency, heartworm disease, mitral valve disease, and second- and third-degree heart block are all frequently diagnosed in ferrets and should be ruled out before surgery. Medical management for adrenal disease in ferrets is discussed in Chapter 7. If surgery is elected, the surgical approach is as follows: Make a ventral midline incision, extending it caudally from the xiphoid as needed to allow access to both adrenal glands. The left adrenal gland is often embedded in fat and is located cranial and medial to the left kidney (Fig. 11-7). The right adrenal gland lies craniomedial to the right kidney and adjacent to the caudal vena cava. Normal adrenal glands are whitish pink, 2 to 3 mm wide, and 6 to 8 mm long.28,30 Remember that not all diseased adrenal glands are enlarged, and the glands should always be palpated for abnormal texture. If the entire left adrenal gland is not visible, use mosquito hemostats and cotton-tipped applicators to carefully dissect the thin layer of peritoneum and fat surrounding the gland to free it. To expose the right adrenal gland, the hepatorenal ligament (which attaches the caudate lobe of the liver to the right kidney) must be incised. Yellowbrown discoloration, an overly firm or gritty texture, and the presence of cysts or enlargement are all indications of a diseased adrenal gland. Manipulation of either adrenal gland may cause a significant tachycardia and increased blood pressure because of catecholamine release. Esmolol (0.25-0.5 mg/kg slow IV bolus) may be used to control this affect. Left adrenalectomy is usually easily accomplished. In ferrets with significant perirenal fat, the gland may be difficult to visualize. Partial dissection may be necessary to fully examine the gland. Begin sharp blunt dissection caudal to the gland and continue in a craniolateral direction. Take care not to disrupt the capsule of the gland. If the gland is determined to be enlarged or otherwise abnormal, continue the dissection. Control hemorrhage of large vessels with cautery, ligation, a vessel-sealing device, or hemostatic clips (Fig. 11-8). The phrenicoabdominal (adrenolumbar) vein is first encountered at the craniolateral aspect of the gland as it courses over the gland’s ventral surface. Ligate this vessel with suture or small hemostatic clips. After dissecting the caudal, lateral, and cranial aspects of the gland, lift it by grasping tissue along its lateral aspect. Examine the area where the phrenicoabdominal vein enters the vena cava for soft tissue invasion. If no invasion is detected, ligate the vein with clips or suture and remove the gland. Right adrenalectomy is more difficult because of adherence of the gland to the wall of the vena cava and the greater potential for vascular invasion. At surgery, have magnifying loupes, microsurgical instruments, and vascular clamps available. Begin dissecting around the gland as described for left adrenalectomy. If the tumor is small and extends lateral to the cava, it often can be almost completely freed from the wall of the cava with gentle dissection. However, it may be impossible to determine if either the ligation or resection might cause disruption of the vena cava. If in doubt, place a vascular clamp (described below) before resection. Small hemostatic clips or small suture
ligations may be placed between the gland and the cava before the gland is resected. If the gland cannot be freed from the vena cava because of tumor invasion or if it is located mostly on the dorsal aspect of the vessel, this technique will often be unsuccessful for complete removal. In these cases, place a small (neonatal) atraumatic vascular clamp on the vena cava for either partial or total occlusion of the vessel (Fig. 11-9). Then remove the gland with a portion of the caval wall and suture the defect with 9-0 or 10-0 suture in a simple continuous pattern. If only a small defect is created, a simple interrupted pattern can be used. The time during which the cava is occluded should be limited. In addition to decreasing renal perfusion, prolonged occlusion of the vena cava may promote the formation of emboli. Before releasing the clamp, place a piece of gelatin sponge over the suture line. Mild oozing is usually observed from the incision line, and gentle pressure can be applied to the area with a cotton-tipped applicator. If hemorrhage is severe, replace the clamp and identify and suture the area of bleeding. With extensive vascular invasion of the cava, resection and anastomosis of the cava may be needed. Some have reported successful outcomes with complete ligation of the vena cava50; presumably these ferrets had enough collateral circulation to allow survival. However, renal compromise is common after ligation of the vena cava, and renal failure has been reported in 30% of ferrets in which this procedure has been performed.18 Venography is also being used to determine the extent of collateral circulation before surgery.18 If collateral circulation is evaluated before surgery and is deemed sufficient, ligation of the caudal vena cava would be less likely to impair renal function (although additional factors may be involved in the renal compromise that can follow adrenalectomy in ferrets).5 Cryosurgery, laser surgery, and ultrasonic and radiosurgical ablation of the right adrenal gland have been reported anecdotally. The Sonosurge ultrasonic surgical unit is widely used by veterinarians in Japan for adrenal gland resection (Watanabe T, Ishizaki I, personal communication, September 2010), but no published controlled studies are available on safety or efficacy of this modality. A disadvantage of these techniques is that the tissue is not available for histopathologic evaluation. In addition, it may be difficult to evaluate how completely the adrenal gland has been destroyed. Bilateral gland involvement has been reported in 16% to 68% of ferrets with adrenocortical disease.37,50 In one study, 32% of ferrets with unilateral adrenal disease presented an average of 11 months after the first surgery with disease of the contralateral gland.50 Subtotal adrenalectomy, with complete removal of one gland and partial removal of the other, is one option for bilateral disease.28,50 Often, the left adrenal is completely removed and the right is debulked by freeing it from surrounding tissue and placing a hemostatic clip or crushing suture across a portion of the gland to allow for removal of 50% to 75%.4 Another method involves incising the capsule and shelling out the contents of the gland.28 Subtotal adrenalectomy usually does not result in the need for long-term postoperative steroid therapy.50 Additionally, one retrospective study demonstrated no significant difference in prognosis for debulking versus total resection of affected adrenal glands.44 However, subtotal resection often results in recurrence of abnormal adrenal tissue and clinical signs of disease.
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B
A
D
C
Fig. 11-9 A, Large right adrenal mass attached to the caudal vena cava. B, Neonatal Satinsky clamp is applied to the vena cava around the base of the mass to result in partial occlusion of the caudal vena cava. C, Application of the Satinsky clamp. The wall of the caudal vena cava can be incised at this point to remove tumor that is invading the vessel, or it can be resected with the adherent right adrenal mass. D, Closure of the vena cava with a simple continuous suture pattern. The lumen of the vena cava has been preserved. Hemoclips have been used to ligate small vessels and the phrenicoabdominal vein.
Postoperative adhesions and rapid increase in size of the remaining adrenal tumor make a second surgery difficult. Complete bilateral adrenalectomy is often performed with good success. It is important to monitor electrolytes postoperatively for mineralocorticoid deficiency. After total bilateral adrenalectomy, most ferrets require glucocorticoid and mineralocorticoid therapy initially, and some may require it for life. Experimental techniques, including ring constrictors and celluloid placement, show promise.14 These techniques are designed to promote the development of collateral circulation, allowing ligation of the vena cava and total resection of the right adrenal gland to be performed during a second surgery. Intraoperative and postoperative complications include hypothermia, tachycardia, hypotension, hemorrhage, glucoand mineralocorticoid deficiency, and renal failure. If recovery from surgery is slow or the ferret appears depressed, collect a
blood sample to measure the packed cell volume (PCV) and blood glucose and electrolyte concentrations. Also, assess pulse quality and measure systolic blood pressure. Continue intravenous crystalloids, supplementing the fluids with dextrose as required. If hypotension is refractory to crystalloid therapy, a colloid bolus (hetastarch, 5 mL/kg) or continuous rate infusion (20 mL/kg/day) may be helpful. Vasopressors such as dopamine are a last resort. Body temperature should be monitored until it remains in the normal range without external thermal support. If hypoadrenocorticism is suspected, dexamethasone may be administered at 0.5 mg/kg initially, or begin therapy with a physiologic dosage of prednisone (0.2 mg/kg per day). If hyperkalemia and hyponatremia are present, mineralocorticoid supplementation may be necessary. In most cases, postoperative recovery is uneventful and the ferret is fed as soon as it is fully awake. In females, the vulvar
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swelling usually regresses within 2 weeks.52 Complete hair regrowth may take 1 to 6 months. Thirty-three to 40 days after surgery, the owner commonly reports that the ferret is bruised all over its body. This appearance results from the development of new hair under the skin, which gives a blue tint to the skin and is normal. Marked postoperative bruising around the incision is common in ferrets. Caution owners about this before surgery so that they will not be alarmed when they take the animal home. This bruising generally disappears 5 to 6 days after surgery and does not seem to cause undue discomfort. Histopathologic analysis usually reveals hyperplasia, adenoma, or adenocarcinoma. Two or even all three of these states of diseased tissue may be found within the same adrenal gland (D. Reavill, personal communication, March 2009). Metastasis of carcinomas is rare.
PANCREATIC SURGERY Insulin-secreting pancreatic islet cell tumors (also known as pancreatic beta-cell tumors or insulinomas) are among the most common neoplastic diseases of ferrets (see Chapter 7). Ferrets typically are presented between 3 and 7 years of age, but the disease can be seen as early as 2 years of age.10,15 Clinical signs associated with hypoglycemia include lethargy, weakness, hind-limb ataxia, nausea, and, in severe cases, seizures or a hypothermic, moribund presentation. Because adrenal disease commonly occurs in ferrets of this age, clinical signs such as hair loss and vulvar swelling may also be observed. A tentative diagnosis of insulinoma is often based on clinical signs associated with hypoglycemia. A resting or a 4- to 6-hour fasting blood glucose concentration lower than 60 mg/dL is indicative of an insulinoma. An elevated insulin level in conjunction with low blood glucose concentration supports the diagnosis, but a normal insulin level does not rule out disease. Abdominal ultrasonography is usually unsuccessful at detecting pancreatic nodules because of their small size but may be effective in ruling out other diseases that can result in hypoglycemia, such as neoplasia, sepsis or severe liver disease. Surgery is a recommended treatment for ferrets assuming that other health concerns are not prohibitive.10,51 Although surgery is often not curative, it allows for evaluation of concurrent disease and usually results in a prolonged medication-free and disease-free interval and increased survival time.15,51 The preoperative workup for insulinoma surgery is similar to that listed for adrenal surgery. Because of their low resting blood glucose concentration, insulinoma surgery patients should be placed on a continuous-rate infusion of fluids containing glucose before and during surgery. Monitor the glucose concentration and alter the concentration of glucose in the infusion as indicated. Preoperative glucocorticoids are often administered to help maintain blood glucose levels. The anatomy of the ferret’s pancreas is similar to that of cats. The common pancreatic duct empties into the duodenum at the major duodenal papilla with the bile duct. An accessory pancreatic duct is also occasionally present. The blood supply to the pancreas comes from three sources (Fig. 11-10): the right limb is supplied by the cranial and caudal pancreaticoduodenal vessels and the left limb is supplied primarily by a branch from the splenic artery. At surgery, assess any adrenal gland abnormalities, enlarged lymph nodes, or evidence of abnormalities of the liver or spleen.
Preoperative ultrasound will often indicate any organs or areas of concern. Evaluate each limb of the pancreas by visually inspecting and gently palpating both the ventral and dorsal surfaces. The body of the pancreas can be inspected only on its ventral surface. Insulin-secreting tumors can appear as visible masses that are discolored or as small nodules (less than 1-mm in diameter) that are not visible initially but are firm on palpation. Magnification loupes are ideal for locating small nodules. Multiple nodules involving more than one lobe of the pancreas are common. Partial pancreatectomy, when possible, is recommended and has been reported to increase survival times.51 Create a window on either side of the pancreatic limb to be removed and free the limb from the surrounding omentum or mesoduodenum. Tighten a 4-0 PDS or Monocryl (Ethicon, Johnson and Johnson Healthcare, Piscataway, NJ) suture around the base of the limb, allowing a margin of at least 5 mm from the last visible nodule (see Fig. 11-10). Transect the pancreatic limb distal to the ligature and close the rent in the omentum or mesoduodenum with small sutures in a simple continuous pattern. Take care during dissection to avoid the pancreaticoduodenal vessels; damage to these vessels can result in necrosis of the duodenum. A cottontipped applicator can often be used to bluntly free the pancreas from these vessels. Preserve the body of the pancreas to avoid damaging the pancreatic duct. Some nodules in this central area cannot be safely resected. If the nodules can be accessed without risking damage to the common bile duct, the nodules can be gently teased away from the surrounding pancreatic tissue. (nodulectomy). If more than one limb is affected, often a partial pancreatectomy is performed on the limb that contains most of the masses. The nodules on the remaining limb may be removed by nodulectomy. Use a mosquito hemostat and cotton-tipped applicators to bluntly free the nodules from the surrounding tissue. Hemorrhage is usually minimal and does not require suturing. The mesenteric lymph nodes in ferrets with adrenal disease and insulinoma are often enlarged and usually reactive, not neoplastic. If enlarged lymph nodes are detected and lymphoma is suspected, biopsy samples can be submitted for histopathologic evaluation. Maintain an intravenous catheter after surgery and check the blood glucose concentration during recovery. Hyperglycemia usually occurs 2 to 12 hours after surgery, but the blood glucose level generally normalizes within a few days. Occasionally insufficient affected pancreatic tissue has been removed, causing hypoglycemia to persist. Postoperative pancreatitis is uncommon, and food and water may be offered within 2 hours after surgery. Rarely, diabetes mellitus requiring insulin supplementation develops after a large pancreatic resection. Check the blood glucose concentration 10 to 14 days after surgery and every 3 to 4 months thereafter. Histopathologic analysis may reveal hyperplasia, adenoma, or adenocarcinoma, but these findings do not dictate prognosis. Recurrence of disease and clinical signs is common. Prognosis is affected by the duration of clinical signs and the presence of local invasion at surgery.15 The longer the duration of clinical signs before surgery, the worse the prognosis. Prognosis is not affected by the number of nodules present or by persistence of hypoglycemia after surgery.10,15 Mean disease-free intervals after surgery are reported to range from 284 to 365 days.15 If hypoglycemia recurs, ferrets can be successfully managed medically with frequent feedings and prednisone, diazoxide, or both for extended periods or a second surgery may be performed. Average survival times of
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A
151
B
Fig. 11-10 A, Pancreatic insulinoma at the tip of the left limb
C
ferrets treated surgically have been reported to range from 563 to 668 days.10,15,51
SPLENECTOMY Splenomegaly is a common finding on abdominal exploration. It is usually benign and represents extramedullary hematopoiesis.8 Neoplastic conditions associated with splenomegaly may include lymphosarcoma and occasionally mast cell tumor,
of the pancreas. B, Partial pancreatectomy. The surrounding mesentery has been dissected free from the left limb of the pancreas and a 4-0 PDS ligature used to ligate the pancreas proximal to the mass. C, Diagram of guillotine suture for partial pancreatectomy with normal pancreatic anatomy. Note the shared blood supply of the pancreas with the spleen and duodenum. Vc, Vena cava; Ao, aorta; SP, spleen; ST, stomach; Spa, splenic artery; Cra, cranial pancreaticoduodenal artery; Cda, caudal pancreaticoduodenal artery.
hemangiosarcoma, adrenal neoplasia, or insulinoma.8,53 Cardiomyopathy, Aleutian disease, and eosinophilic gastritis have also been reported as causes of splenic enlargement. Splenic sequestration of red blood cells, seen with isoflurane and sevoflurane anesthesia, can result in both splenomegaly and a significant reduction in the hematocrit and plasma protein concentration.23 Ferrets with splenomegaly should be evaluated for underlying disease conditions. If abnormal echogenicity is noted on
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UROGENITAL SYSTEM NEPHRECTOMY
Fig. 11-11 Splenectomy of an enlarged spleen in a ferret. Hemoclips are placed close to the spleen.
ultrasound, ultrasound-guided fine-needle aspiration or core needle biopsy can help determine whether primary splenic disease is present. Before splenic biopsy, determine clotting ability by evaluating the CBC, platelet count, prothrombin time, and partial thromboplastin time if possible. Splenectomy is indicated for primary splenic or metastatic neoplasia or severe splenic enlargement that interferes with abdominal visceral function. Because the spleen is a major site of erythropoiesis in ferrets, splenectomy should not be taken lightly and the potential consequences of surgery must be considered carefully. The approach to the spleen is comparable to that described for exploratory laparotomy. Splenic biopsy techniques are the same as for liver biopsy. Small absorbable suture material (e.g., 4-0 PDS) can be placed in a cruciate or mattress pattern to control hemorrhage. Sutures tend to hold better in the spleen than in the liver because of the relatively thick splenic capsule. Although partial splenectomy is rarely indicated, it can be done for benign processes that result in excessive splenic enlargement. For this procedure, ligate and divide the vessels supplying the portion of spleen to be removed. Place a clamp across the area to be removed, and close it to crush the tissue. Place a thoracoabdominal stapling device (TA-30, U.S. Surgical Corporation, Norwalk, CT) in the crushed area after the clamp is removed. After the staples are fired, transect the spleen distal to the stapler and release the stapler. Control any subsequent hemorrhage with hemoclips or mattress sutures. As an alternative to a stapling device, place mattress sutures through the spleen proximal to the crushing clamp to control hemorrhage. Close the splenic capsule in a simple continuous pattern before removing the clamp. Total splenectomy involves ligating all vessels supplying the spleen (Fig. 11-11). Individually double-ligate the splenic artery and vein, being careful to avoid damaging the vascular supply to the pancreas in this area. Ligate the short gastric vessels close to the spleen to avoid damaging the stomach. Vascular sealing devices and small hemoclips can be used instead of suture material to control hemorrhage, but stapling devices used in other small animals (Autosuture LDS, U.S. Surgical) are too large for use in ferrets.
Nephrectomy is indicated in cases of unresolved hydronephrosis, chronic bacterial nephritis, and neoplasia. The kidneys of ferrets, like those of other domestic species, are located in the retroperitoneal space. However, the large amount of retroperitoneal fat in ferrets often obscures their visibility. For nephrectomy, the surgical approach is through a ventral midline incision. Isolate the involved kidney with moistened sponges and open the peritoneum over the kidney to access the retroperitoneal space. Grasp the kidney and apply gentle traction while using cotton-tipped applicators to isolate the renal vessels and ureter. Although it is simpler to perform, do not massligate the renal artery and vein; instead, individually dissect and double-ligate the renal artery and vein close to the aorta and vena cava, respectively. This allows greater security in ligation and prevents the formation of an arteriovenous fistula.1 The vessels may be double-ligated with hemostatic clips or fine 4-0 or 5-0 suture material. Dissect the ureter along its retroperitoneal course all the way to the urinary bladder. Then ligate and transect the ureter as close to the urinary bladder as possible without damaging the nerves that run in the lateral ligament of the bladder. Lavage the abdominal cavity and close it routinely.
CYSTOTOMY Like other domestic species, ferrets can suffer from cystic and urethral calculi. Magnesium ammonium phosphate (struvite) is the most common type of urolith found in ferrets, although calcium oxalate and cysteine-based calculi have been reported.13 The history and clinical signs of urolithiasis in ferrets are similar to those in other small animal species and may be characterized by stranguria, hematuria, recurrent urinary tract infections, or urinary tract obstruction. Surgery to remove calculi is indicated to relieve clinical signs and for quantitative crystallographic stone analysis. Cystotomy in ferrets is similar to the procedure performed in other small animal species. Make a caudal midline abdominal incision (in females) or a parapreputial skin incision with a midline abdominal incision (in males). After locating the bladder, pack the abdominal cavity with moistened sponges to limit contamination. Choose a ventral location on the bladder that is relatively devoid of blood vessels as the incision site. Using fine suture material, place stay sutures on either side of the planned incision line. The wall of the ferret’s urinary bladder is very thin, so take care in applying traction to the stay sutures. After making the incision, remove the calculi and thoroughly inspect and lavage the bladder. Flush the urethra with a small, soft catheter to ensure that no calculi remain. Submit the calculi for stone analysis and submit a crushed specimen of the calculi and a small section of the bladder wall for bacterial culture and sensitivity testing. Close the cystotomy incision with fine (5-0) synthetic absorbable suture material in either a simple interrupted or a simple continuous pattern. Check the bladder for leakage by using a syringe and a 25-gauge needle to infuse sterile saline solution. If leakage is observed, add a continuous inverting suture layer over the initial sutures. Lavage the abdomen thoroughly and close routinely. Postoperative radiographs of the bladder and urethra are recommended to ensure removal of all calculi.
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Prostatic cyst
Fig. 11-12 Perineal urethrostomy in a ferret. A catheter is inserted in the urethra, and the first three sutures of 5-0 monofilament suture material have been placed.
PERINEAL URETHROSTOMY Because of urethral calculi, distal strictures, or neoplasia, a perineal urethrostomy is occasionally indicated in a ferret. The anatomy of the ferret’s penis and urethra most closely resembles that of dogs. The site of the urethrostomy should be located caudal (proximal) to the os penis and the site of obstruction and ventral to the anus. Magnification with surgical loupes greatly simplifies the procedure. Place the ferret either in dorsal recumbency with the pelvic area elevated or in ventral recumbency within a perineal stand. Aseptically prepare the perineal region and caudal abdomen. If possible, place a urethral catheter in the anesthetized ferret before surgery. This can be difficult because of the urethra’s small diameter, the hook-shaped os penis, and the relative difficulty of extruding the penis from the prepuce (see Chapter 4). Make a skin incision approximately 1 cm cranial (ventral) to the anus. This exposes the urethra and the cavernous tissue. If a urethral catheter is present, palpate the catheter and make a 1-cm longitudinal incision along the ventral aspect of the urethra. If a urethral catheter cannot be placed, make the incision ventrally, taking care not to stray laterally and incise the cavernous tissue. Open the urethra and a place a larger urethral catheter proximally to assist in locating the lumen and mucosa. Use simple interrupted sutures of 5-0 or 6-0 synthetic absorbable monofilament suture material to close the urethral mucosa to skin (Fig. 11-12). Remove the sutures after 2 weeks if possible; otherwise, the absorbable sutures may be left in place to dissolve.
PARAURETHRAL/PROSTATIC CYSTS Prostatic disease in ferrets is usually associated with adrenal gland neoplasia. Ferrets are presented with a history of stranguria and, potentially, urinary obstruction. Alopecia, pruritus, and other clinical signs associated with hyperadrenocorticism may also be observed. Differential diagnoses for stranguria include urinary calculi, infection, penile or preputial lesions, neoplasia of the urinary tract or prostate, and prostatic cysts or hyperplasia.39 Initial diagnostic tests should include a urinalysis and culture and an abdominal ultrasound examination to evaluate the urinary tract, prostate, and adrenal glands. Additional diagnostic tests, such as measurements of plasma concentrations of
Bladder
Fig. 11-13 Prostatic cyst in a male ferret with adrenal disease. Generally the prostatic cyst communicates with the bladder, and cystocentesis of either structure often reduces the obstruction and reestablishes urethral patency. Many of these cases can be managed medically after cystocentesis and catheterization. If medical management does not resolve the cyst, marsupialization of the cyst may be required.
adrenal hormones, may also be helpful. If the bladder is distended and cannot be expressed, pass a urinary catheter after the ferret is anesthetized. If urethral catheterization is unsuccessful, perform cystocentesis under anesthesia with a 25-gauge needle. Because of the communication between the prostate and the bladder, cystocentesis will significantly reduce the contents of a cystic prostate and often allow urine outflow to be reestablished (Fig. 11-13). Several drugs used to treat prostatic hypertrophy in men, such as finasteride, are useful in rapidly reducing both prostatic size and the secretions of prostatic cysts (see Chapter 7). Medical management is usually successful in treating urinary obstruction due to prostatic hypertrophy and cystic prostatic disease. If it is not successful, abdominal exploration and adrenalectomy may be indicated. At surgery, aspirate the prostatic or paraprostatic cyst with a small-gauge needle. The fluid contained within the cyst is often green and viscous; a sample should be submitted for bacterial culture and sensitivity testing. Because multiple cysts may be present, aspirate multiple sites. If the cysts are small, aspiration may be the only treatment required. For larger cysts (more than 2 cm in diameter), remove a portion of the cyst wall for biopsy and allow complete drainage of all cavitations. Dissection of the prostate should be restricted to the ventral two-third of the gland. Trauma to the dorsolateral aspect, between 10 and 2 o’clock, should be avoided at all costs to spare the neurovascular structures supplying the prostate in this region.34 The cyst can then be omentalized. Theoretically, omentalization provides continued drainage of the cyst, aids in adhesion formation, and enhances immune function to fight against infection. “Pack” a portion of the omentum into the cyst cavity and suture it to the cyst wall with 4-0 absorbable suture.7 Place three to four sutures, being careful to avoid disrupting the blood supply to the omentum. Express the bladder to check for leakage of urine from the cyst. If leakage is observed, place more omentum into the cyst and partially close the opening into the cyst with suture. If leakage is still seen, a transurethral catheter
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or tube cystostomy will have to be maintained for 1 to 2 days after surgery until the rents have sealed. Usually the prostate decreases in size and the cysts regress within 1 to 2 days after adrenalectomy. Histologic findings in ferrets with prostatic cysts have revealed squamous metaplasia and prostatitis caused by keratin, squamous cells, and cellular debris. These changes are believed to be caused by adrenocortical disease and associated high levels of circulating estrogens or androgens.11,39 Often secondary bacterial infection is present, which may result in prostatic abscesses.
to prevent contamination. Remove the sponge and lavage the stump before closing the abdomen. Treat with a broad-spectrum antibiotic to cover the organisms associated with pyometra in ferrets, including Staphylococcus, Streptococcus, and Corynebacterium species and Escherichia coli.28 Stump pyometra may be seen with ovarian remnants or adrenal gland disease. In these cases, remove the remnant or adrenal gland in addition to resecting the stump.
OVARIOHYSTERECTOMY
Ferrets are usually castrated to reduce aggressive behavior, decrease musky odor, and protect against testicular neoplasia. Castration at an early age, as routinely performed by ferret facilities in the United States, is associated with an earlier onset of adrenal disease. Delayed castration and use of GnRH receptor agonists in the interim may be considered in young male ferrets to mitigate testosterone related signs40 (see Chapter 7). Castration can be performed as it is in cats, with an incision in the scrotum over each testicle. An open or a closed technique can be used, and the spermatic cord can be ligated closed with an overhand tie, open with a “self-tie” technique (vas deferens to vessels), or ligated with 4-0 absorbable suture. Leave the incisions open to heal by secondary intention. Alternatively, make a prescrotal incision and exteriorize both testicles through the same incision. With this technique, close the subcutaneous tissue with 4-0 or 5-0 absorbable suture, and close the incision with either intradermal or skin sutures.
Because most pet ferrets in the United States are spayed at breeding farms before they are 8 weeks of age, ovariohysterectomy is not a common procedure in U.S. veterinary practice. Intact female ferrets remain in estrus with high circulating estrogen levels until they are stimulated to ovulate through breeding or artificial stimulation. Spaying of intact pet ferrets is recommended before 6 months of age to prevent life-threatening bone marrow suppression caused by chronically high estrogen levels. For ovariohysterectomy, make a ventral midline incision about 1 cm caudal to the umbilicus. The ferret’s uterus is bicornuate and can be found just dorsal to the bladder. Because of the large amount of body fat in female ferrets, the ovarian vessels may be difficult to locate. Completely ligate the ovarian pedicles and uterine body with 3-0 or 4-0 absorbable suture material. Close the linea routinely with 3-0 or 4-0 absorbable suture. Subcuticular sutures in a continuous or interrupted pattern may be used for skin closure.
OVARIAN REMNANT In a spayed female ferret, clinical signs of estrus (swollen vulva) are usually caused by adrenal neoplasia. Occasionally, these signs result from a remnant of ovarian tissue inadvertently left behind during prior ovariohysterectomy. Signs typically occur in ferrets younger than 2 years of age, which is younger than in ferrets with adrenal disease. Before abdominal exploratory surgery, evaluate the ferret for anemia and thrombocytopenia, which can result from estrogen toxicity. At surgery, the remnant is typically found just caudal to the kidneys.28 A thorough exploration is necessary because more than one remnant may be present. Ovarian remnants, which may have been dropped during the previous ovariohysterectomy and revascularized, can be found in any part of the abdominal cavity. Submission of the resected tissue for histopathologic examination is recommended to verify that it is ovarian. Vulvar swelling should resolve after removal of the remnant.
PYOMETRA Because most ferrets are spayed at an early age, pyometra is uncommon. Clinical signs suggestive of pyometra include vulvar discharge, lethargy, and anorexia. Polyuria and polydipsia are not commonly reported. Radiographs and ultrasound are usually diagnostic. The treatment for pyometra is ovariohysterectomy combined with fluid and antibiotic therapy. Before surgery, submit a blood sample for a CBC to evaluate for bone marrow suppression associated with hyperestrogenism. Ovariohysterectomy is routine. At surgery, exteriorize the uterus and place a moistened laparotomy sponge in the abdomen
CASTRATION
PREPUTIAL MASSES Masses and cystic structures involving the prepuce occur in male ferrets and are usually found incidentally on physical examination. However, occasionally they result in urethral obstruction. They often develop secondary to adrenal disease and secretion of androgenic hormones.35 These must be differentiated from adenomas, adenocarcinomas, and squamous cell carcinomas, which can also occur in this location.32 Adenomas can be removed simply by marginal excision and reconstruction of the preputial orifice with 5-0 suture material. Rotational or advancement rafts may be used if needed. Adenocarcinomas and squamous cell carcinomas must be resected more aggressively, possibly requiring surgical removal of the prepuce (Fig. 11-14). In these cases, a partial penile amputation is required to prevent exposure damage to the penis, with diversion of urine flow by a perineal urethrostomy. Subsequent radiation therapy may be required, and the prognosis is guarded.
MISCELLANEOUS SURGICAL PROCEDURES ANAL SACCULECTOMY Anal sacculectomy (descenting) is mainly performed in conjunction with early neutering at ferret breeding farms, but it may be done at any age. Removal of the anal sacs may decrease the musky odor in neutered ferrets, but some odor associated with the glands in the perianal region will remain.12 Also, body odor is primarily caused by sebaceous secretions of the skin, which increase during breeding season (see Chapter 1). Therefore neutering alone will significantly reduce the musky scent in male ferrets. However, anal sac removal is often desired.
CHAPTER 11 Soft Tissue Surgery
Fig. 11-14 The prepuce and catheterized urethra of a male ferret, prior to redraping and surgical preparation for reconstruction, after resection of a preputial adenocarcinoma. Preputial neoplasia, as in this case, may require surgical resection and subsequent radiation therapy to attempt control. Preputial cysts are also seen and may respond to treatment for the underlying adrenal production of testosterone.
The anal sac openings are located at the anocutaneous junction at the 4- and 8-o’clock positions. Magnification is of great assistance in locating the anal sac openings and throughout the surgical procedure. Identify the opening of the anal sac and grasp it with fine mosquito forceps. Incise the skin 1 to 2 mm around the anal sac opening with a No. 15 scalpel blade. Use a scraping motion with the blade to remove glandular tissue, skin, and subcutaneous tissue surrounding the duct and sac. Dissection of the gland is initially difficult because of the close association of the surrounding tissue, but after a few millimeters, a tissue plane exposing the yellow surface of the anal sac will become evident. Continue to dissect the anal sphincter muscle from the anal sac until the entire sac is freed. Ligate or clamp the duct and remove the intact sac. If any of the anal sac tissue remains after surgery, fistulous tracts may develop. If bleeding is observed that does not quickly stop with digital pressure, small blood vessels may need to be ligated or coagulated. Lavage the defect, and either close the opening with a single subcuticular suture using 5-0 synthetic absorbable suture material or leave it open to heal by secondary intention. Remnants of anal sacs are encountered from incomplete resection performed in young ferrets. These remnants may still be secretory, creating a typical strong ferret odor. They may also abscess; in some cases, the remnant may represent adenocarcinoma.29
HEARTWORM DISEASE—CAVAL SYNDROME One report of transvenous removal of heartworms in a 10-month-old ferret demonstrated the potential for this procedure despite the small size of the patient.6
References 1. Aiken SW, Jakovijevic S, Lantz GC, et al. Acquired arteriovenous fistula secondary to castration in a dog. J Am Vet Med Assoc. 1993;202:965-967. 2. Antinoff N, Hahn K. Ferret oncology: diseases, diagnostics, and therapeutics. Vet Clin North Am Exot Anim Pract. 2004;7:579-625. 3. Bauck LB. Salivary mucocele in two ferrets. Mod Vet Pract. 1985;66:337-339.
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4. Beeber NL. Surgery in pet ferrets. In: Bojrab MJ, ed. Current techniques in small animal surgery. Baltimore: Williams & Wilkins; 1998:763-769. 5. Bennett RA, Laraio L, Weisse C, et al. Collateral circulation during caval occlusion in ferrets. In: Proceedings. Assoc Exot Mammal Vet. 2008:105. 6. Bradbury C, Saunders AB, Heatley JJ, et al. Transvenous heartworm extraction in a ferret with caval syndrome. J Am Anim Hosp Assoc. 2010;46:31-35. 7. Bray JP, White RA, Williams JM. Partial resection and omentalization: a new technique for management of prostatic retention cysts in dogs. Vet Surg. 1997;26:202-209. 8. Brown SA, Rosenthal KL. Causes of splenomegaly in ferrets. Vet Med. 2000;95:599. 9. Caligiuri R, Bellah JR, Collins BR, et al. Medical and surgical management of esophageal foreign body in a ferret. J Am Vet Med Assoc. 1989;195:969-971. 10. Caplan ER, Peterson ME, Mullen HS, et al. Diagnosis and treatment of insulin-secreting pancreatic islet cell tumors in ferrets: 57 cases (1986-1994). J Am Vet Med Assoc. 1996;209:1741-1745. 11. Coleman GD, Chavez MA, Williams BH. Cystic prostatic disease associated with adrenocortical lesions in the ferret (Mustela putorius furo.) Vet Pathol. 1998;35:547-549. 12. Creed JE, Kainer RA. Surgical extirpation and related anatomy of the anal sacs of the ferret. J Am Vet Med Assoc. 1981;179:575-577. 13. Del Angel-Caraza J, Chávez-Moreno O, García-Navarro S, et al. Mixed urolith (struvite and calcium oxalate) in a ferret (Mustela putorius furo). J Vet Diagn Invest. 2008;20:682-683. 14. Driggers T. A Novel surgery for right-sided adrenalectomies inferrets (Mustelo putorious furo), Proceedings. Assoc Exot Mammal Vet. 2008:107-109. 15. Ehrhart N, Withrow SJ, Ehrhart EJ, et al. Pancreatic beta cell tumor in ferrets: 20 cases (1986-1994). J Am Vet Med Assoc. 1996;209:1737-1740. 16. Evans HE, An NQ. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:19-69. 17. Holmberg DL. Cryosurgery. In: Slatter DS, ed. Textbook of small animal surgery. Philadelphia: WB Saunders; 2003:222-227. 18. Kelleher S. Skin diseases of ferrets. Sem Av Exot Pet Med. 2002;11:136-140. 19. Lawrence HJ, Gould WJ, Flanders JA, et al. Unilateral adrenalectomy as a treatment for adrenocortical tumors in ferrets: five cases (1990-1992). J Am Vet Med Assoc. 1993;203:267-270. 20. Lennox AM. Gastrointestinal diseases of the ferret. Vet Clin North Am Exot Anim Pract. 2005;8:213-225. 21. Li X, Fox JG. Neoplastic diseases. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:405-447. 22. Lipman NS, Marini RP, Murphy JC, et al. Estradiol-17b- secreting adrenocortical tumor in a ferret. J Am Vet Med Assoc. 1993;203:1552-1555. 23. Marini RP, Callahan RJ, Jackson LR, et al. Distribution of technetium 99m-labeled red blood cells during isoflurane anesthesia in ferrets. Am J Vet Res. 1997;58:781-785. 24. Martin RA. Liver and biliary system. In: Slatter DH, ed. Textbook of small animal surgery. 2nd ed. Philadelphia: WB Saunders; 1993:645-659. 25. Mehler S, Bennett RA. Surgical oncology of exotic animals. Vet Clin North Am Exot Anim Pract. 2004;7:783-805. 26. Miller PE, Picket JP. Zygomatic salivary gland mucocele in a ferret. J Am Vet Med Assoc. 1989;194:1437-1438. 27. Mullen HS, Scavelli TD, Quesenberry KE, et al. Gastrointestinal foreign body in ferrets: 25 cases (1986-1990). J Am Anim Hosp Assoc. 1992;28:13-19. 28. Mullen HS. Soft tissue surgery. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders; 1997:131-144.
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29. Nakata M, Miwa Y, Nakayam H, et al. Localised radiotherapy for a ferret with possible anal sac apocrine adenocarcinoma. J Small Anim Pract. 2008;49:476-478. 30. Neuwirth L, Collins B, Calderwood-Mays M, et al. Adrenal ultrasonography correlated with histopathology in ferrets. Vet Radiol Ultrasound. 1997;38:69-74. 31. Parker GA, Picut CA. Histopathologic features and post-surgical sequelae of 57 cutaneous neoplasms in ferrets (Mustela putorius furo L.). Vet Pathol. 1993;30:499-504. 32. Pinches MDG, Liebenberg G, Stidworthy MF. What is your diagnosis? Preputial mass in a ferret. Vet Clin Pathol. 2008;37:443-446. 33. Poddar S. Gross and microscopic anatomy of the biliary tract of the ferret. Acta Anat. 1977;97:121-131. 34. Powers LV, Winkler K, Garner MM, et al. Omentalization of prostatic abscesses and large cysts in ferrets (Mustela putorius furo). J Exot Pet Med. 2007;16:186-194. 35. Protain HA, Kutzler MA, Valentine BA. Assessment of cytologic evaluation of preputial epithelial cells as a diagnostic test for detection of adrenocortical disease in castrated ferrets. Am J Vet Res. May 2009;70:619-623. 36. Rickman BH, Craig LE, Goldschmidt MH. Piloleiomyosarcoma in seven ferrets. Vet Pathol. 2001;38:710-711. 37. Rosenthal KL, Peterson ME, Quesenberry KE, et al. Hyperadrenocorticism associated with adrenocortical tumor or nodular hyperplasia of the adrenal gland in ferrets: 50 cases (19871991). J Am Vet Med Assoc. 1993;203:271-275. 38. Rosenthal KL, Peterson ME. Evaluation of plasma androgen and estrogen concentrations in ferrets with hyperadrenocorticism. J Am Vet Med Assoc. 1996;209:1097-1102. 39. Rosenthal KL, Peterson ME. Stranguria in a castrated male ferret. J Am Vet Med Assoc. 1996;209:63-64. 40. Schoemaker NJ, van Deijk R, Muijlaert B, et al. Use of a gonadotropin releasing hormone agonist implant as an alternative for surgical castration in male ferrets (Mustela putorius furo). Theriogenology. 2008;70:161-167. 41. Shoemaker NJ, Schuurmans M, Moorman H, et al. Correlation between age at neutering and age at onset of hyperadrenocorticism in ferrets. J Am Vet Med Assoc. 2000;216:195-197.
42. Smith M, Schulman FY. Subcutaneous neoplasms of the ventral abdomen with features of adrenocortical tumors in two ferrets. Vet Pathol. 2007;44:951-955. 43. Soderstrom MJ, Gilson SD. Principles of surgical oncology. Vet Clin North Am Small Anim Pract. 1995;25:97-110. 44. Swiderski JK, Seim 3rd HB, MacPhail CM, et al. Long-term outcome of domestic ferrets treated surgically for hyperadrenocorticism: 130 cases (1995-2004). J Am Vet Med Assoc. 2008;232(9):1338-1343. 45. Triantafyllou A, Fletcher D, Scott J. Histological and histochemical observations on salivary microliths in ferret. Arch Oral Biol. 2006;51:198-205. 46. Triantafyllou A, Harrison JD, Garrett JR. Microliths in the parotid of ferret investigated by electron microscopy and microanalysis. Int J Exp Pathol. 2009;90:439-447. 47. Tunev SS, Wells MG. Cutaneous melanoma in a ferret (Mustela putorius furo). Vet Pathol. 2002;39:141-143. 48. Wagner RA, Dorn DP. Evaluation of serum estradiol concentrations in alopecic ferrets with adrenal gland tumors. J Am Vet Med Assoc. 1994;205:703-707. 49. Weiss CA, Scott MV. Clinical aspects and surgical treatment of hyperadrenocortism in the domestic ferret: 94 cases (19941996). J Am Anim Hosp Assoc. 1997;33:487-493. 50. Weiss CA, Williams BH, Scott JB, et al. Surgical treatment and long-term outcome of ferrets with bilateral adrenal tumors or adrenal hyperplasia: 56 cases (1994-1997). J Am Vet Med Assoc. 1999;215:820-823. 51. Weiss CA, Williams BH, Scott MV. Insulinoma in the ferret: clinical findings and treatment comparison of 66 cases. J Am Anim Hosp Assoc. 1998;34:471-475. 52. Wheeler J, Bennett RA. Ferret abdominal surgical procedures: part I. Adrenal gland and pancreatic beta-cell tumors. Compend Contin Educ Pract Vet. 1999;21:815-822. 53. Wheeler J, Bennett RA. Ferret abdominal surgical procedures: part II. Gastrointestinal foreign bodies, splenomegaly, liver biopsy, cystotomy, and ovariohysterectomy. Compend Contin Educ Pract Vet. 1999;21:1049-1057.
SECTION TWO
Rabbits
CHAPTER
12
Basic Anatomy, Physiology, and Husbandry
David Vella, BSc, BVSc (Hons), Diplomate ABVP (Exotic Companion Mammal), and Thomas M. Donnelly, BVSc, Diplomate ACLAM
Etymology Taxonomy and Similarities to Rodents Breeds and Varieties Life Span Anatomy Skin and Hair Sense Organs and Nervous System Muscles and Skeleton Digestive System Respiratory System and Thymus Cardiovascular System Urinary System Puberty and Breeding Life Female Reproductive System Anatomy and Physiology Female Sexual Behavior Pregnancy and Nursing Behavior Hand-Rearing of Baby Rabbits Male Reproductive System Anatomy and Physiology Male Sexual Behavior and Reproduction Behavior Eating, Drinking, and Elimination Behavior Group Behavior Vocalization, Auditory and Visual Signals Husbandry Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
ETYMOLOGY The mammalian order Lagomorpha consists of two living families: Leporidae (rabbits and hares) and Ochotonidae (pikas) (Fig. 12-1). The scientific name for the Old World or European rabbit from which all domestic breeds originate is Oryctolagus cuniculus. The genus name is derived from the words orukter (Greek: a tool for digging) and lagos (Greek: a hare) and the species name is derived from cuniculus (Latin: a rabbit or an underground passage). In contrast to most other species in Lagomorpha, rabbits are burrowing animals. Through human mediation, wild European rabbits have been dispersed to all continents except Antarctica. This has led to some confusion with the common name rabbit. The family Leporidae splits broadly into two groups: the hares of the genus Lepus, containing 32 species, and the rabbits in the remaining 10 genera, which include Oryctolagus (Fig. 12-2). However, these vernacular names are often used synonymously and applied to the wrong animals. For example, the African red rock hares (Pronolagus species) and the endangered hispid hare (Caprolagus hispidus), although commonly called hares, are rabbits. Some hares are called rabbits—for example, the snowshoe rabbit (L. americanus) and, in North America, the several species of hare known as jackrabbits (shortening of jackass-rabbit, so called for its long ears). Even more confusing, in North America, cottontails, or New World rabbits, Syvilagus species (11 species), are commonly called rabbits. Both the European rabbit and the eastern cottontail (S. floridanus) are unique because of their great diversity of habitat: fields, farms, woodlands, deserts, swamps, and forests. However, in North 157
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America, the European rabbit Oryctolagus cuniculus has not become feral and is found only as a domesticated animal; all wild rabbits in North America are cottontails. The English word rabbit (from Middle English rabbet, from Middle French dialect rabbotte, from Walloon robète [baby rabbit], diminutive of Middle Dutch robbe) arose in the fourteenth century to mean the young of the species. The adult was a coney (from Anglo-Norman conis, plural of conil from Latin cunicula) until the eighteenth century and the name can be found in the King James Bible (Leviticus 11:5; Proverbs 30:26). Baby rabbits were an important source of food in the early Middle Ages, because Pope Gregory I (c. 540-604) officially classified laurices (unborn or newly born rabbits) as “fish” and thus permissible to eat during Lent and other Christian fasts. In sixteenth-century
Lagomorpha
Ochotonidae
Pikas Ochotona (30)
Leporidae
Jackrabbits and hares Lepus (32)
Rabbits Brachylagus (1) Bunolagus (1) Caprolagus (1) Nesolgaus (1) Oryctolagus (1) Pentalagus (1) Poelagus (1) Pronolagus (3) Romerolagus (1) Sylvilagus (16)
Fig. 12-1 Relationship of domestic rabbits (Oryctolagus cuniculus) within the family Leporidae (hares and rabbits). Approximately 80 living species of lagomorphs, placed in 2 families and 13 genera, are currently recognized.
British slang, a coney was someone to cheat or rob and doing so was known as “coney-catching”19; the Lancashire expression “coney-fogle” meant “to lay plots.”8 Although the pronunciation of coney (rhyming with honey and money) was changed from a short to long vowel (rhyming with boney),18 the word rabbit gradually pushed out coney in common usage. The similarity of present-day European words for rabbit to the English coney can be seen: coniglio [Italian], conejo [Spanish], konijn [Dutch], Kaninchen [German]. While there are specific words in English to mean the young of a species (e.g., dog/puppy, hare/leveret, cat/ kitten) there is no word for the young of rabbits, although they are often referred to as kits or by the hypocoristic term bunny.
TAXONOMY AND SIMILARITIES TO RODENTS Most veterinarians observe that rabbits and rodents show a similarity to each other. In these animals, the anterior incisors in the upper and lower jaws are modified to form chisel-like cutting organs.36 Enamel is deposited on the anterior surfaces only, the back surface being dentin. Because enamel is harder than dentin, the front surface wears down more slowly and the incisors remain in a permanently sharp condition from gnawing. This modification is associated with a loss of teeth between the last incisor and the first cheek tooth so that there is a toothless interval referred to as the diastema. Karl von Linne (also called Linnaeus, 1717-1738), who originated the systematic classification of plants and animals, recognized the likeness of rabbits and rodents to each other. In his Systema Naturae (10th ed., 1758) he assigned rabbits and rodents to a group called Glires (Latin glis = dormouse). Later naturalists designated Glires as the mammalian order Rodentia. Within this order rabbits, hares, and pikas were grouped in the suborder Duplicidentata because they possessed a second pair of incisor teeth in the upper jaw. Most rodents, which have only a single pair of upper incisors, were grouped in the suborder Simplicidentata. However, throughout the twentieth century there has been a strong consensus that the similarities of rabbits
Fig. 12-2 Diagram comparing the bodies of a rabbit (left) and a hare (right). Note the differences in size and length of legs and ears.
CHAPTER 12 Basic Anatomy, Physiology, and Husbandry and rodents exemplify convergent evolution (i.e., the development of a similar adaptive morphological trait by two or more unrelated species). Consequently naturalists now designate the Duplicidentata as the mammalian order Lagomorpha (to which rabbits belong) and restrict the order Rodentia to the larger group of mammals with only one pair of upper incisors, such as squirrels, rats, mice, and guinea pigs. Based on results of DNA work, in recent years the concept that the two orders are in fact related has resurged. The term Glires, which Linnaeus originally used, is now used to describe the infraclass that encompasses these two orders.
BREEDS AND VARIETIES Domestication of O. cuniculus was achieved between the fifth and tenth centuries in southern Europe. Monks kept rabbits in their monasteries as a food source long before forest clearance and agriculture altered the environment sufficiently to allow large numbers of rabbits to exist in the wild. Many of the domesticated forms bear little resemblance to the original wild stock. Some are small (dwarf rabbits), weighing 1 kg, but those bred for meat production may weigh as much as 7 kg. Rabbits are divided into over 60 fancy breeds and fur breeds. The fur group is divided into normal fur breeds, Rex breeds, and Satin breeds. The normal fur breeds have a coat made up of an undercoat and projecting guard hairs; the Rex breeds have short guard hairs that do not appear above the level of the undercoat; and the Satin breeds have an abnormal hair fiber that produces a sheen.71 The term variety describes a color (e.g., black, blue, steel gray, tortoiseshell) within a breed. Over 500 varieties are described. For a list and images of domestic rabbit breeds, refer to the American Rabbit Breeders Association (www.arba.net) or the British Rabbit Council (www.thebrc.org). Body conformation and ear size vary widely among breeds of rabbits, and rabbit fanciers have coined some unusual terms to describe lagomorph body shape and fall of ear. They refer to the small, chunky body of a dwarf rabbit, like a cobblestone, as cobby; they describe the long, lean body of a Belgian hare as racy; and they often describe giant rabbits as mandolin-shaped because of the high, curved top line over their hindquarters.81 Most breeds of rabbits have upright ears, which can be long or short. However, some breeds have soft, pliable ears that hang downward and are incapable of erection; we know these as lops.
LIFE SPAN Records for the life span of rabbits in captivity are 9, 10, and 18 years.13 Veterinarians often report seeing pet rabbits 9 to 10 years of age, and one of the authors has seen a 15-year-old pet rabbit. Wild rabbits in an enclosure and receiving supplementary winter feed were recorded as surviving a maximum of 7.7 and 8.7 years (female and male, respectively).76 The longest life span recorded for a female European rabbit in the wild is 7.6 years.65
ANATOMY Several books have been published on the anatomy of the rabbit. Unfortunately the two best atlases on rabbit anatomy are out of print. The more recent Colour Atlas of the Anatomy of Small Laboratory Animals: Rabbit, Guinea Pig has been republished three times since its first print in 1990.66 It is easily found in
159
many libraries. The other, Atlas of Rabbit Anatomy (in French and English), is found only in selected university libraries owing to its limited print pressing in 1973.5
SKIN AND HAIR Rabbit skin is very delicate compared with that of other exotic pet species, dogs, and cats. Unless care is taken in clipping fur for surgery, it is easy to tear or rip the skin. Because rabbit hairs are so fine, electric hair clippers require the use of thin blades (size 50, 1/125 in. [0.2 mm], or 40, 1/100 in. [0.25 mm]), which are normally not used with cats and dogs. Hair growth in rabbits occurs in periodic, orderly waves originating on the head and ventrum and spreading dorsally and caudally.80 This is most noticeable when hair grows back after clipping for surgery. Moulting in adult rabbits usually occurs twice a year (spring and fall). A female rabbit has a large fold of skin over the throat known as a dewlap. Breeding does pull fur from this area to line their nests before kindling. In older breeding does, the dewlap can be large and easily mistaken for an abscess. Moist dermatitis often develops in this area. Rabbits do not have footpads; instead, coarse fur covers the front and hind toes and metatarsal areas. When a rabbit is sitting undisturbed, the plantar surface of the lower hind limb from the toes to the hock is in contact with the ground. Rabbits housed on wire floors often develop an ulcerative pododermatitis of this area called sore hocks. Large breeds and obese, emaciated, or pregnant rabbits are more vulnerable. Guard hair provides the normal protective barrier between the feet, the plantar surface of the lower hind limb, and the substrate. Do not clip hair from the base of the feet or hocks. In Rex rabbits, the guard hairs are shortened. This makes Rex rabbits more vulnerable to developing pressure sores on their feet, especially at the points of the hocks or at the tips of the third phalanges. The claws of rabbits are very sharp, and a rabbit picked up without appropriate support of its hindquarters can inflict painful scratches on the handler. Tactile vibrissae present on the face aid in food discrimination and underground navigation. The only glabrous areas of the rabbit are the nose tip, scrotal sacs, and inguinal folds.
Scent Marking Glands Rabbits are strongly territorial, and both sexes have three glands used in scent-marking behavior: the chin glands, which are specialized submandibular glands opening onto the underside of the chin; the anal glands; and a pair of pocket-like perineal glands called the inguinal glands that are relatively large and often harbor a normal yellowish-brown deposit. The size of the glands and degree of marking are androgendependent and related to the level of sexual activity. Males mark more frequently than do females, dominants of both sexes mark more frequently than subordinates do, and dominants mark most in the presence of subordinate rivals. Under natural conditions, both bucks and does on their own territory, surrounded by their own odor and that of their clan, win two-thirds of all aggressive encounters.57 When a rabbit becomes dominant, a new compound appears in the chin gland secretion, 2-phenoxyethanol.38 This compound is a fixative used in the perfume industry and slows the release rate of compounds in chin gland secretions, enabling the scent to persist in the environment and not dissipate. The dominant
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rabbit thus reigns over the olfactory environment, just as it does the physical environment.
SENSE ORGANS AND NERVOUS SYSTEM Rabbits are subject to a high degree of predation; consequently their sense organs are well developed. They are sensitive to catecholamines and have evolved for flight rather than fight. Temperature, heart, and respiratory rates significantly increase in a frightened animal; therefore obtain these values early in the physical examination, as the stress of handling may alter these parameters. Stress from either transport or the veterinary hospital environment may alter these values, as well and washing one’s hands clean of “predator” animal odors (such as a dog, cat, or ferret) before starting the consultation can minimize stress.
Eye Prince has described the anatomy and physiology of the rabbit eye in great detail in the old but not outdated The Rabbit in Eye Research.68 The rabbit cornea is large, occupying 30% of the globe, and the eyes are directed more laterally than those of most mammals. These two features give rabbits a panoramic field of vision to detect predators readily. However, their eyes cannot visualize the small area beneath the mouth (blind spot), and rabbits depend on the sensitivity of the lips and vibrissae for food discrimination. During clinical examination, avoid touching the rostral aspect of the muzzle, as it involves contact with the blind spot and startles the rabbit. As in most rodents and other lagomorphs, the lens is spherical and large. The ciliary body is small and poorly developed. These two features suggest that rabbits have a limited need to alternate near and far focus (i.e., accommodation). The optic nerve is above the horizontal midline of the eye, and examination of the fundus involves looking upward into the eye. Retinal vessels spread out horizontally from the optic disk; also, the rabbit has a depression or physiologic cup in the optic disc, as does the dog. Rabbits do not have a tapetum lucidum.37 The nictitating membrane or third eyelid is prominent, found in the medial canthus of the eye and associated with deep (harderian) and superficial (nictitans) orbital glands. Behind it and separated from the deep part of the nictitating membrane cartilage is the harderian gland, which has a small white upper lobe and a large pink lower lobe. The lower lobe is known as the deep gland of the nictitating membrane. Both the deep and superficial glands of the third eyelid can potentially prolapse.42 The deep gland prolapses more frequently than the superficial gland.82 The clinical condition is similar to “cherry eye” in dogs. The harderian gland is wrapped behind the eyeball and is phylogenetically and anatomically associated with the nictitating membrane. It is believed that the harderian gland has a significant function in social behavior. Generally horseshoeshaped and situated deep within the orbit, the harderian gland consists of a smaller, almost white (upper) lobe connected to a large pink (lower) lobe by a narrow strand medial to the optic nerve. The single excretory duct opens at the base of the nictitating membrane, and the milky secretion is made up of a complex mixture of lipids, protein, and the pigment protoporphyrin, which provides lubrication for the edges of the eyelid.24 Epiphora in rabbits often presents as a white overflow of tears
because of impaired drainage or obstruction of the nasolacrimal drainage system—the white tears are a normal secretion of the harderian gland.45 Other orbital glands include the lacrimal and accessory lacrimal glands, located caudodorsal and ventral to the orbit respectively. A few millimeters posterior to the limbus and under the bulbar conjunctiva, the rectus dorsalis muscle is visible. Placement of an anchoring suture under or around this muscle serves to stabilize the globe during surgery; in other species, dissection is necessary to find the extraocular muscles. In rabbits, the primary channel for return of venous blood from the head, including that from the eye, is the external jugular vein.67 In contrast, the primary drainage of the eye and head in humans is via the internal jugular vein. In other species, such as dogs, significant anastomoses exist between the branches of the internal and external jugular veins. In rabbits, such anastomoses are minor, and ligation or chronic catheterization of the external jugular vein results in swelling and protrusion of the eyeball for about 24 hours, after which its normal appearance returns.41 The same pattern of vascularity also applies to the arterial blood supply of rabbit’s eyes, but ligation of the external carotid artery results in ipsilateral ocular necrosis. Rabbits, like rodents, have an extensive orbital venous plexus. Because of possible severe hemorrhage, enucleation of the eye is difficult compared with species such as dogs. Bilateral exophthalmos can occur in rabbits secondary to engorgement of the orbital venous sinuses (retrobulbar venous plexus). Rabbits with either compromised heart function or cranial thoracic masses,77 such as thymoma, thymic lymphosarcoma and thymic carcinoma, can present with marked “cranial vena cava” signs such as bilateral exophthalmos. Occasionally, bilateral exophthalmos can also be seen in rabbits exhibiting extreme fear. The nasolacrimal drainage system provides a conduit for tears from the lacrimal lake to the nasal cavity. In rabbits, a single ventral lacrimal punctum (about 3-4 mm ventral to the lid margin), canaliculus, sac, duct, and nasal meatus form the drainage system for each eye. The diameter of the nasolacrimal duct is small and narrows in two places where it changes course. These two sites, the proximal maxillary bend and the apex of the main upper incisor, are important in the development of obstruction50 (see Chapter 37). Dacryocystitis and nasolacrimal duct obstruction (dacryostenosis) often cause epiphora, one of the most common ocular problems seen in rabbits (see chapter on ophthalmology).
Ear The pinnae represent a large portion of the total body surface in rabbits, approximately 12% in New Zealand white rabbits.21 The pinnae are highly vascular and have the largest arteriovenous shunts in the body. At rest, the pinna of leporids is a thermoregulatory organ. Claude Bernard in 1852 implied that rabbits use their ears to lose heat6; at ambient temperatures above 30°C, there is a marked vasodilation of ears.69 The Australian population of the European rabbit has responded to its warmer environment by increasing the mean length of its pinna from 71 mm in England and France to 79 mm in Australia.62 Anatomists have hypothesized another role for the pinna of leporids as a part of a suspensory system for the greater portion of the head, absorbing shocks that might otherwise interfere with vision during high-speed locomotion.74 This is certainly the case with European hares. However, as the rabbit is semifossorial and does not rely on sustained fast running to avoid
CHAPTER 12 Basic Anatomy, Physiology, and Husbandry predation, as do hares, the significance of the ears in capital shock absorption in rabbits is less important. In large-eared rabbits, the vessels of the ear margins (medial and caudal auricular veins) are easily accessible for venipuncture. However, in small-eared rabbits such as dwarf breeds, venipuncture of the ear veins can lead to vasculitis, vascular necrosis, and sloughing of the ear pinna. During anesthesia, the central auricular artery and auricular veins are good sites to obtain noninvasive measurements such as pulse oximeter blood oxygen saturation and laser-based systolic and diastolic arterial blood pressure.39 The vertical ear canal has a natural diverticulum separated by a cartilaginous bridge called the tragus. Lop-eared breeds have a stenosis in their ear canals at the point of flexion that may predispose to otitis externa.
MUSCLES AND SKELETON Bones of rabbits are relatively delicate compared with their muscle mass. The skeleton represents only 7% to 8% of body weight in rabbits, whereas the skeletal muscle comprises more than 50% of the body weight.9,21 Fractures, especially of the tibia, are always a potential problem. In comparison, the skeleton of a cat constitutes 12% to 13% of body weight. However, the dry matter and percentage of calcium is higher in the bones of rabbits than those of cats.1 In countries where rabbit meat is commonly eaten, clients may present a headless carcass that they suspect to be that of a cat. The color of the muscles distinguishes the two: rabbit muscles are pale pink, whereas those of cats are deep red. There are also skeletal differences, most noticeably between the scapula and pelvis. The infraspinous fossa of the rabbit is sharply triangular, whereas that of the cat is more rounded; also, the suprahamate process of the acromion is truly hook-shaped in rabbits, whereas it is blunted in cats. Good drawings of these differences are found in Okerman’s Diseases of Domestic Rabbits.59 The acetabulum of rabbits is formed by the ilium, the ischium, and a small accessory bone, the os acetabuli, which excludes the pubis. In other animals, the acetabulum comprises the ilium, the ischium, and the pubis. The trochanteric fossa of the femur is a good site for intraosseous catheter placement and is easily located by palpating the prominent greater trochanter.10 Scientists have studied the architecture of skeletal muscle in different species to determine the force, velocity, and displacement properties of a muscle. The rabbit has a pelvic limb muscle mass of 13% (of total body mass), reflecting its need to evade predators by accelerating quickly and maintaining high-speed locomotion.83 Forelimb muscle mass accounts for 9% of total body mass, highlighting the predisposition toward hindlimb propulsion. The bulk of the fore- and hindlimb muscle mass is proximal; the distal limbs (e.g., carpi and tarsi) have little musculature; in the hindlimb, this adaptation decreases rotational inertia.46 Spinal musculature accounts for 9% of body mass—a substantial portion of the total locomotor muscle mass (~30%).83 Much of the back musculature extends the lumbar spine and increases the distance from the hindfoot to the rabbit’s center of mass. When normalized for body mass, the back muscles are much larger than those of the dog and horse. The powerful hindlegs and lumbar muscles mean that rabbits can kick violently. If they are not held securely when picked up, their kicking can result in a vertebral fracture (usually at the seventh lumbar vertebra) and damage to the spinal cord. Clinically,
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the large lumbar and proximal hindlimb muscles are the best sites for intramuscular injections. The number of thoracic (T) and lumbar (L) vertebrae varies from 12T, 7L in 44%; 13T, 6L in 33%; and 13T, 7L in 23% of rabbits.33 The spinal cord ends within the second sacral vertebra (S2) in 79%, within the first sacral vertebra (S1) in 19%, and within the third sacral vertebra (S3) in 2%.33 This feature is unusual, as the spinal cord ends in other species (e.g., dog, horse) within the caudal lumbar region.
DIGESTIVE SYSTEM The definitive references on the anatomy and physiology of the digestive tract of rabbits are Cheeke’s Rabbit Feeding and Nutrition14 and the more recent The Nutrition of the Rabbit by de Blas and Wiseman.22 Most of the information described in this section is from these books unless otherwise noted.
Teeth All the teeth of rabbits are classified elodont (continuously growing, with no anatomic “roots”) and hypsodont (long-crowned). The teeth can also be classified as aradicular. This leads to a dynamic state intrinsic to rabbit dentition; clinically, it serves to complicate dental disease. This type of dentition also allows for a concurrent increase in tooth size with growth of the animal.36 The rabbit dental formula is 2 (I2/1, C0/0, PM3/2, M3/3) = 28. It is useful clinically to divide rabbit dentition into two groups, the incisor teeth set and cheek teeth set (premolars and molars). Each cheek teeth arcade can be divided into quadrants (upper and lower on right and left). The rabbit mouth also features a relatively long diastema. The lower jaw is narrower than the upper jaw, a feature known as anisognathism. The maximum gape of the rabbit is only 20 to 25 degrees, compared with the rat’s 40 degrees.36 This, coupled with the long diastema, can make inspection of the oral cavity relatively difficult. Each tooth can be further divided into the clinical crown (region of tooth exposed above the gingival margin) and the reserve crown (region of tooth buried below the gingival margin). This reserve crown is often misnamed the root of the tooth. The growing portion at the tip of the reserve crown is the apex, which is open in the rabbit’s elodont hypsodont teeth (Fig. 12-3). Incisors have a single pulp cavity. The cheek teeth have a single pulp chamber at the tooth apex that diverges into two toward the clinical crown. Enamel is present only on the labial (facing toward the lips) side of the strongly curved maxillary
Fig. 12-3 Cross-section schematic of a rabbit skull showing dentition of mandible and maxilla and of an individual tooth. (Courtesy of Marise Watson.)
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SECTION II Rabbits
incisors, while the mildly curved mandibular incisors feature enamel on both the labial and lingual (facing toward the tongue) aspects of the teeth.20 Through normal wearing, this feature produces the characteristic chisel shape of the clinical crown tips (in occlusively normal incisor teeth). In contrast, the cheek teeth are longitudinally straighter and somewhat “folded” on the buccal (toward the cheek) side in cross section.36 This formation creates an increase in the proportion of enamel on the occlusal surface of the tooth (providing increasing grinding efficiency and helping to reduce the wear rate). The more heavily calcified alveolar bone surrounding each tooth socket is termed the lamina dura. Normally occluded mandibular incisor tips lie just caudal to the main maxillary incisors, in the space between the primary maxillary incisors and the peg teeth (the smaller second set of upper incisor teeth situated just caudal to the main maxillary incisors).20 The tips of the mandibular incisors in occlusively normal rabbits make contact with the peg teeth.36 Rabbits periodically “grind” their teeth to help shape their incisor tips to their characteristic chisel shape.36 At rest, the opposing cheek teeth quadrants are normally in contact.36 This observation contrasts to what many authors have previously described, where they stated that at rest, the rabbit’s opposing cheek teeth quadrants do not normally contact each other. In fact, in the normal resting closed mouth gape, both the opposing incisors and cheek teeth are in contact with each other.36 During mastication, each lower cheek tooth occludes with two upper cheek teeth except for the first and last lower cheek teeth.36 The rate of tooth growth varies between the different sets and can be influenced by age, pregnancy, and diet.36 For example, the upper incisors grow more slowly than the lower incisors, at rates of 2 and 2.4 mm/week, respectively.72 Rabbits primarily use a vertical action to “cut” foliage with their incisors.36 Once typical leaf material is present in the mouth, it is then masticated largely in a horizontal or lateral plane by the cheek teeth. This is important to consider, as dietary factors have been shown to affect a rabbit’s normal chewing process.78 Food is ground by only one side of the cheek teeth at a time.36 Natural vegetation such as grass complements a “normal” horizontal chewing action of the cheek teeth, while harder and thicker food items (such as pellets, grains, or carrots) tend to encourage more vertical and less horizontal movements in mastication.78 This vertical action can lead to a reduction in tooth wear and also potentially increase forces on teeth in the vertical plane, producing increased pressure on the growing apex of the tooth.12 Further discussion of rabbit dental anatomy and physiology may be found in Chapter 32.
Oral Cavity The mouth opening in rabbits is small. Muscles of the jaws extend both forward and backward. This confers a deceptively large appearance to the oral cavity, which is actually smaller than the size of the jaws suggests. Because the articular process forming the temporomandibular joint is elongated longitudinally, the mandible can move forward, backward, and vertically but less so from side to side.
Abdominal Cavity The abdominal cavity of rabbits is large. Rabbits are true herbivores and classified as hindgut fermenters. The gastrointestinal tract is relatively long, and its contents can make up 10% to
20% of the body weight. The two most striking organs because of their size are the stomach and cecum; their combined contents account for 10% (range 5%-19%) of the rabbit’s total body weight. Preanesthetic fasting is sometimes recommended to partly empty the intestinal tract, not to prevent vomiting.
Stomach The stomach serves as a reservoir for much of the ingested feed. It typically contains 15% of the alimentary tract ingesta, and food and fecal pellets are usually present. The stomach lacks specialized regions, is thin-walled, and often appears ruptured at necropsy because of gas distention during autolysis. However, the cardia and pylorus are well developed. The lower esophagus contains a massive muscular sphincter and a serrated mucosal rosette at the cardiac orifice; these two structures form a perfect closing mechanism between stomach and esophagus, and rabbits are unable to vomit. The pylorus is easily compressed by the duodenum, which exits at an acute angle. Gastric distention or compression due to a trichobezoar, gas, or hepatomegaly contributes to pyloric compression and prevents emptying. The rabbit’s stomach is extremely acidic (pH 2), which effectively kills bacteria and other microorganisms, so that the stomach and small intestine are essentially sterile. In suckling rabbits, the stomach’s acidity is higher (pH 5-6.5); after weaning, it drops (pH 2-3). This is why weanling rabbits are so susceptible to diarrhea, because the stomach pH is not low enough to kill ingested bacteria.
Small Intestine The small intestine is shorter in rabbits than in other species, making up about 12% of the total volume of the gastrointestinal tract. The duodenum and jejunum have a relatively small lumen. Peyer’s patches are absent from the duodenum and the first half of the ileum. No correlation exists between the number of Peyer’s patches and intestinal length; surgeons or pathologists may see only three to nine Peyer’s patches in the first quarter of the jejunum and last quarter of the ileum.54,70 The terminal portion of the ileum ends with the rounded sacculus rotundus at the ileocecocolic junction. The sacculus rotundus has a minute honeycombed external appearance owing to the presence of a large number of lymph follicles; it is sometimes referred to as the ileocecal tonsil. The terminal portion of the ileum is a common site of foreign-body impaction due to luminal narrowing.
Large Intestine Rabbits have a large, thin-walled, coiled cecum that displays 18 to 22 haustra-like bulges or pouches delineated by a spiral running constriction. The cecum holds about 40% of the ingesta (Fig. 12-4) and ends in a finger-like, thick-walled, pale vermiform appendix that is characterized, like the sacculus rotundus, by abundant lymphatic tissue. The cecum is the largest and most prominent organ in the abdominal cavity of rabbits. It folds onto itself three times as it constricts around most of the inner surface of the abdominal cavity wall. The cecal contents are generally semifluid. Sacculations and the presence of bands characterize the colon, which start at the bulbous cecal ampulla coli from which the proximal colon emerges. The fusus coli, a thickened section of colon heavily supplied with ganglion cell aggregates, separates the proximal colon from the distal. The fusus coli acts as a pacemaker that controls the contractions for excreting the two distinct types of feces that rabbits produce: a dry, hard
CHAPTER 12 Basic Anatomy, Physiology, and Husbandry
163
have a rapid turnover.32 Inflammation of the large intestine increases crypt populations of goblet cells; consequently mucus associated with feces is a normal inflammatory response in the rabbit. The presence of mucus is not pathognomic and the disease mucoid enteropathy is descriptive only (not etiologic) in its name.
7
8 1 6
Liver, Gallbladder, Pancreas, and Spleen
4 3
2 5
Fig. 12-4 Dissected large intestine of a rabbit. Key: 1, ileum; 2, sacculus rotundus; 3, body of cecum (note the long spiral fold along its length); 4, vermiform appendix; 5, ampulla coli; 6, proximal colon; 7, fusus coli; 8, distal colon.
fecal pellet, which is discarded, and a soft fecal pellet, which is contained within a strong mucous envelope (also known as soft feces, night feces or cecotropes). The soft feces appear as a cluster rather than as single pellets typical of hard feces and are produced during one or two periods each day. They attach to hairs around the anus and are ingested directly and swallowed without mastication. They remain within the mucoid membrane for up to 6 hours after ingestion, where their contents are maintained at a relatively neutral pH because of their high levels of phosphate buffers. The stimulus for ingestion of cecotropes may be short-chain fatty acids because of their strong odor and taste and high concentration in soft feces. Many owners mistake uneaten cecotropes for diarrhea, especially when they adhere to the fur around the anus. Germ-free cecotomized rabbits consume neither their hard nor soft fecal pellets. Cecotrophy can be extremely important to the nutrition of rabbits, as cecotropes contain high levels of short-chain fatty acids, microbial protein, B vitamins, sodium, potassium, and water. Cecotropes are also a rich source of nitrogen, providing up to 30% of the total nitrogen intake of rabbits, and the microbial protein shows a high content of essential amino acids. Therefore the practice of cecotrophy can greatly improve both the quantity and biological value of protein in the diet. The short-chain fatty acids in these fecal pellets also provides an additional source of energy, and the B vitamins provided can be in excess of the animal’s needs. It is estimated that B12 is synthesized at 100 times the daily requirement. Cecotropes also aid in the replenishment of cecal microflora. Thus the products of bacterial growth are made available to the rabbit either by direct absorption in the cecum and colon or in the small intestine by consumption of the cecal contents. The large intestine is rich in goblet cells that coat cecotropes with mucus. This mucilaginous membrane surrounding the cecotropes acts as a barrier to the low pH of the stomach and permits their reabsorption in the small intestine. Multiple zones of proliferating cells exist at multiple levels in the crypt column of the distal colon and rapidly differentiate as they migrate toward the surface epithelium. In contrast, only basally located stem cells in the small intestine proliferate and then differentiate only as they migrate toward the surface epithelium. Consequently, populations of proliferating cells in the distal colon
The liver lies caudal to the diaphragm, mostly within the caudal ribcage. It is divided into two major lobes (left and right separated by a deep median cleft). The left lobe is subdivided into lateral and medial lobes. Similarly, the right lobe has subdivisions of lateral and medial lobes, plus smaller quadrate and caudate lobes, giving a total of six lobes. The caudate lobe has a narrow attachment or stalk to the dorsal, hilar region of the liver. This stalk is prone to displacement, and torsion of the caudate lobe has been reported.79 The gallbladder is deceptively deep within the abdominal cavity and is located between the right medial and quadrate lobes. Like dogs, rabbits have separate openings for the bile duct and the pancreatic duct in the duodenum. The bile duct empties into the proximal part of the duodenum. Rabbits secrete mainly biliverdin, rather than bilirubin, in their bile, as do most nonmammalian species. Although the rabbit’s pancreas is closely associated with the duodenum, it is diffuse and often difficult to differentiate from the surrounding mesentery. The single pancreatic duct opens at the junction of the ascending and transverse loops of duodenum. Ligation of this duct does not result in pancreatic insufficiency. Gut-associated lymphoid tissue constitutes about 50% of the total lymphoid tissue mass. Therefore the rabbit’s spleen is relatively small compared with the spleen in other species. The spleen is elongated and flat and lies along the greater curvature of the stomach. The frequency of accessory spleens in rabbits is 9%.28
RESPIRATORY SYSTEM AND THYMUS The integument surrounding the snout tip is differentiated into the rhinarium, a hairless, somewhat protruding skin region. It is characterized by a midline cleft extending from the upper lip, which divides right and left to the nostrils. This feature, found only in lagomorphs and rodents, is known as the upper lip cleft, or by its common name, the harelip.3 Rabbits are obligate nasal breathers because of the close anatomic relationship of the larynx to the nasopharynx. The epiglottis normally lies dorsal to the soft palate. This characteristic has important clinical and anesthetic consequences. Mouth breathing is a poor prognostic sign, and inflammation of the upper respiratory tract increases the risk of anesthetic mortality in nonintubated animals. A rabbit can be challenging to intubate. Its small mouth, long narrow oral cavity, and large cheek teeth restrict direct visualization of the larynx. Traumatic or prolonged endotracheal intubation can cause mechanical abrasion, pressure necrosis, and subsequent ulceration of the laryngeal and tracheal mucosa. Average tracheal diameters of rabbits ranging from 2.3 to 5.1 kg are 5.8 mm (ventrodorsal) by 5.4 mm (lateral) at the level of the cricoid and 4.7 mm (ventrodorsal) by 5.9 mm (lateral) at the eighth tracheal ring.47 The dimensions of the rabbit’s subglottis appear consistent independent of weight, whereas tracheal size at the level of the eighth tracheal cartilage varies significantly by weight.
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SECTION II Rabbits
Endoscopic evaluation of bronchial morphology in rabbits has been described.43 Severe hyperemia of the trachea is a normal finding. The deep-red color of the epithelium does not involve the dorsal tracheal membrane and vanishes relatively abruptly at the level of the carina. In contrast, the mucosa of the trachea is pale pink in dogs and cats. In rabbits, there are three left lung lobes (cranial, middle, caudal) and four right lung lobes (cranial, middle, caudal, accessory). The left lung is about two-thirds the size of the right lung. The left lung of rabbits, unlike that in rats and mice, is lobed. In rabbits (and rodents), the lungs do not have connective tissue septa, which in other animals divide the lungs into the pyramidal portions of pulmonary tissue known as lobules. This is significant in relation to the pathology of the lower respiratory system. Most anatomic types of pneumonia in domestic animals are described as “lobular,” but “lobular pneumonia” is not described in rabbits and rodents. Lobular pneumonia is classically bronchogenic in origin compared with lobar pneumonia, which can be hematogenous or bronchogenic in origin. Pneumonia in rabbits and rodents is always lobar and therefore either hematogenous or bronchogenic in origin. The lungs of rodents and rabbits differ from those in humans and dogs as far as total lung volume changes are concerned. In humans and dogs, an increase in residual volume (air that remains in the lung following expiration) occurs with age at the expense of vital capacity (the maximal volume of air that can be expelled from the lungs). Short, shallow, sometimes labored breathing is seen in old humans and dogs. In rodents and rabbits, the lung volume changes with age and the ratio of residual volume:vital capacity does not change. Any signs of dyspnea in rabbits and rodents always point to a poor prognosis. The thoracic cavity of the rabbit is relatively small; in contrast, the abdominal cavity is large, and the rabbit breathes primarily by contraction of the diaphragm. These characteristics allow for an efficient method of artificial respiration: suspend the rabbit horizontally in midair, holding the fore limbs in one hand and back legs in the other hand, and gently rock the rabbit from a head-up to a head-down position every 1 to 2 seconds. The overall shape of the thymus is “sail shaped” similar to that in the dog, and it may persist into adult life. It consists of three lobes, one main left thoracic lobe and the dorsal and ventral right thoracic lobes (see Chapter 20). The thymus lies cranioventral to the heart and extends forward to the thoracic inlet. It is not unusual to have a large thymus occupying the cranial thorax (from ventral to the heart to the thoracic inlet) in rabbits.
CARDIOVASCULAR SYSTEM As with many other organs, the size of the rabbit’s heart is directly related to its body size. Because the size of the heart limits blood volume, the tissues cannot be supplied with oxygen by means of increased ventricular volume pumped out in one beat. Instead, the heart’s frequency of heartbeats remains the major modifying variable; consequently heart rates are higher in smaller animals than in larger ones. In rabbits, the heart rate can vary from 180 to 250 beats per minute. Despite the small size and rapid contraction rate of rabbit hearts, quantitative Doppler-echocardiographic methods have been validated to evaluate structural and functional abnormalities29,30 (see Chapter 20). The heart lies in the cranial thorax and is normally positioned between ribs 3 and 6. The rabbit has a relatively small heart, which accounts for about 0.3% of its body weight.
However, the small size of the thorax may paradoxically create the impression of an enlarged heart radiographically. The rabbits has a left and right cranial vena cava. The left cranial vena cava and coronary veins terminate in a coronary sinus, which may be misinterpreted as a congenital defect or early right-sided heart failure.63 A limited collateral coronary circulation can predispose the rabbit’s heart to ischemia induced by coronary vasoconstriction. It is a significant causative factor in ketamine/xylazine induced myocardial fibrosis.51 The right atrioventricular valve is unique because it is composed of two rather than three cusps. The rabbit aorta has a rhythmic contraction that is neurogenic in origin. The veins have thin walls and peripheral veins are susceptible to hematoma formation, which can be avoided through the judicious use of Teflon catheters and gentle pressure. Muscle swellings in the rabbit’s pulmonary artery make this vessel thicker and more muscular than it is in any other species. Pulmonary hypertension causes death from anaphylaxis; at necropsy, severe constriction of the pulmonary arteries and dilatation of the right side of the heart are observed.
Hematology The erythrocyte count varies between 5 and 8 million/mm3 and erythrocytes show marked anisocytosis. Erythrocyte life span is 60 to 70 days26; this relatively short span (e.g., humans, 120 days; dogs, 90 days) is associated with increased polychromasia (1%-2%) to replace senescent erythrocytes. A feature of rabbit blood is the occurrence of numerous thorn apple-shaped erythrocytes in the blood smear. The erythrocytes include a large number of reticulocytes (1%-7%) in adult animals and even higher numbers in young rabbits. Extramedullary hematopoiesis is possible in the liver and spleen. Leukocyte counts in rabbits vary considerably because of diurnal fluctuations, nutrition, and differences in sex, age, and breed. The differential count of one healthy rabbit can fluctuate considerably if repeated over a period of 1 month. The neutrophils of rabbits are comparable to those of large domestic animals and humans and are called heterophils or pseudoeosinophils because of their many small eosinophilic staining granules (see Chapter 36). The lymphocyte is the most common leukocyte in the blood of young animals (less than 12 months of age); after 13 months of age, heterophils and lymphocytes may be present in approximately equal numbers.11 There is a large variation in lymphocyte numbers according to age and nutritional condition. Overall, variations in hematologic values due to breed are not as pronounced as those due to the individual physiology, nutrition, age, and sex of the animal.
URINARY SYSTEM Most mammalian kidneys are multipapillate, but those of the rabbit and rodent are unipapillate. Only one papilla and one calyx enter the ureter directly. The plasticity of the rabbit urinary system has not been fully recognized. Rabbits from Australian desert zones have large kidneys with powerful urine-concentrating ability, small adrenal glands, and low levels of circulating aldosterone. Rabbits from Australian alpine zones have small kidneys (at least 25% less in weight than those of desert rabbits), large adrenal glands, and high levels of circulating aldosterone.56 At necropsy, the most striking difference is in the size of the renal medulla.
CHAPTER 12 Basic Anatomy, Physiology, and Husbandry In the desert-dwelling rabbits, which live on a high-fiber, highsalt, low-protein plant diet and have limited access to water, the medulla is long. In contrast, in alpine rabbits, which enjoy a high-protein diet of lush grasses that are low in sodium, the renal medulla is short. A major reason rabbits have been so preeminent in successfully colonizing diverse geographic regions is that their urinary system has the capacity to vary its developmental pattern under different environmental conditions and with different digestive strategies. The serum calcium level of rabbits is unusual because it is not regulated in a narrow range but reflects the level of dietary calcium. Urine is a major route of calcium excretion, which varies directly with the serum calcium level. Whereas the fractional excretion of calcium in most mammals is less than 2%, the range for rabbits is 45% to 60%. Increases in dietary calcium levels directly increase urinary calcium excretion (Fig. 12-5). The consistency of the urine is often thick and creamy because of a white calcium carbonate precipitate. Prolonged intake of a diet high in calcium can result in calcification of the aorta and kidneys. High vitamin D intake intensifies this effect.44 Biological pigments from the breakdown of endogenously synthesized compounds (e.g., bile pigments, porphyrins, flavins, melanins, pteridines) or ingested plant pigments (e.g., betacyanin, the red beet pigment; anthocyanin, a purple-red pigment; and lutein, a yellow pigment) are excreted in urine and,
with fluid volume (hydration status), give urine a characteristic color. Among the pigments is a group of yellow, brown, and red pigments that are generally designated as urochromes; they are considered metabolic breakdown products. The red pigment was first named uroerythrin by Simon in 184073 and was known to increase with certain metabolic states, such as high fever or tissue degradation.73 Uroerythrin is a breakdown product or a precursor of heme7 and is related to the bile pigment biliverdin. Urine pigments can be bile pigments such as bilirubin, biliverdin or bilicyanin; or they can be derivatives of bile pigments that appear singly, as compounds or as compounds with a peptide (e.g., urochrome B is a compound of urobilin or urobilinogen and a peptide). The color of normal rabbit urine varies from yellow to red. The red coloration is a normal variation of urine color, but rabbit owners often erroneously report the presence of blood in their pet’s urine. Certain types of feed (e.g., alfalfa, the tropical legume Leucaena, beet, carrot, spinach and cruciferous vegetables such as cabbage, radish, turnip, mustard, cauliflower, and broccoli) seem to increase the intensity of the red pigmentation.2,15,35 Feeds may contain plant pigments that directly color urine various shades of red. Alternatively, foods (e.g., cruciferous vegetables) may induce selective cytochrome P450 enzymes in the liver that will affect how bile pigments and their urinary derivatives (e.g., billirubin, urobilin, urobilinogen) are metabolized.
20
Item (%)
Rabbits
Rats
Ingested Ca in feces
19.8
92.7
Ingested Ca in urine
59.0
2.0
Apparent Ca digestibility
80.2
7.3
Plasma calcium (mg/100 mL)
19 Comparative calcium metabolism
165
18
17
16
15
14 1
2 3 4 Percent dietary calcium
5
Fig. 12-5 Table of calcium digestibility versus ingested calcium in urine and feces for rat and rabbit. Graph of plasma calcium concentration versus ingested (dietary) calcium for rabbit. In contrast to many species, the blood calcium level of rabbits readily reflects dietary calcium intake (see graph). Most mammals regulate intestinal calcium absorption to metabolic need. In a rat, little calcium is absorbed and most of it is passed in the feces, while a rabbit absorbs most of the calcium independent of metabolic needs (see table). When serum calcium exceeds the renal threshold in a rabbit, excess serum calcium is excreted in the urine, while other mammals excrete little calcium in the urine (see table). Rabbit kidneys have a high calcium fractional excretion (~2.25 mM) compared with other mammals (~1.5 mM). (Modified after Cheeke PR. Rabbit feeding and nutrition. Orlando, FL: Academic Press, 1987.)
SECTION II Rabbits
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Weaning
Puberty
Weight gain (grams/day)
1.0 0.8 Male
0.6
Female
0.4 0.2 0.0 0
10 20 30 40 50 60 70 80 90 100 110 120 Days postpartum
A
Fig. 12-6 Growth curve plot. Velocity curves for weight gain in mice. In both sexes, puberty occurs just after maximal rate of growth. Although this curve shows mouse weight gain, the underlying principle is the same in rabbits. The age at which rabbits attain sexual maturity varies considerably according to either the strain or breed. Sexual maturation occurs at the point on the growth velocity curve where growth is still taking place but the rate is decelerating rapidly. This means that body weight is more important than age in determining sexual maturity.
Left uterine horn
Right uterine horn Right vaginal cervix
Vaginal fornix
External uterine ostium Uterine broad ligament
Left ureter
Right ureter
Urinary bladder
PUBERTY AND BREEDING LIFE The age at which rabbits attain sexual maturity varies considerably according to the breed. However, if the biologic pattern of growth is plotted graphically, puberty occurs just after maximal rate of growth. On the growth velocity curve, sexual maturity occurs at the point at which growth is still taking place but its rate is decelerating rapidly (Fig. 12-6). Therefore body weight is more important than age in determining sexual maturity. Small breeds develop more rapidly and are mature at 4 to 5 months of age. Medium-sized breeds mature at 4 to 6 months, and large breeds reach maturity at 5 to 8 months of age.64 Does mature earlier than do bucks, which do not achieve optimal sperm production and reserves until 40 to 70 days after puberty. Among New Zealand white rabbits, females reach maturity at approximately 5 months and males at 6 to 7 months of age. The reproductive life of a rabbit depends on its breed but is about 5 to 6 years for the buck and up to 3 years for the doe.
FEMALE REPRODUCTIVE SYSTEM ANATOMY AND PHYSIOLOGY The reproductive tract of a doe lacks a uterine body; instead, each of two separate uterine horns has its own opening into the vagina. If a cesarean section is performed, the surgeon must remove the fetuses through a separate incision in each horn. The mesometrium is a major fat storage site, and identification and ligation of the uterine vessels can be difficult (Fig. 12-7, A). The vagina of the rabbit is long and flaccid. In neutering a female rabbit, the surgical procedure is actually an ovariohysterovaginectomy, as the incision is made through the vagina distal to the vaginal fornix. Do not manually express the urinary bladder before surgery, as urine will pool in the vagina; then,
Vagina (ventral floor) Urethra External urethral opening Vaginal vestibule Greater vestibular gland Corpus Clitoris Preputium Glans
Vestibular bulb Preputial gland Constrictor muscle of vestibule Lesser lip of pudenda Greater lip of pudenda
B Fig. 12-7 A, Dorsal surface of a dissected postparturient rabbit uterus. Note the abundant mesometrial fat, which makes identification and ligation of uterine vessels difficult. B, Diagram of rabbit vagina. The rabbit’s vagina is long and characterized by a well-developed, complex musculature. It forms a single canal by which does copulate, give birth, and urinate. In contrast to rodents, in which the vagina opens into the vulvar cleft on the external surface of the body, in the rabbit the vagina ends several centimeters beyond the caudal edge of the pubic bone, and the distal urethra opens midway along the length of the vagina (~4 cm from the vaginal orifice) into the anterior pelvic vagina. Never express the bladder during ovariohysterectomy, as urine will pool in the vagina; then, when the uterocervical tube is severed, urine can flow anterograde and contaminate the abdominal cavity.
when the vagina is cut, the urine may flow out and contaminate the abdominal cavity (Fig. 12-7, B). Like cats and ferrets, rabbits are induced ovulators and do not have an estrous cycle. However, does vary in their sexual receptivity, so that a certain rhythm can be detected.
CHAPTER 12 Basic Anatomy, Physiology, and Husbandry In domestic rabbits, this rhythm has been reported as having intervals of 4 to 6 days. Ovulation occurs after coitus or after the injection of luteinizing hormone. The time of ovulation in induced ovulators varies among species. In rabbits, it is approximately 10 hours after copulation compared with 30 hours in cats and ferrets.
FEMALE SEXUAL BEHAVIOR Sexual receptivity in a doe is characterized by lordosis—a reverse bending or flattening of the back, raising of the pelvis, or presenting of the perineum in response to attempts by a buck to mount. Similar behavior is seen in cats. Generally female rabbits are hyperactive and brace themselves when touched. When they are not receptive, does do not allow males to mount. Depending on cage space, nonreceptive behavior often takes the form of running away, cornering, biting, and vocalizing. In natural conditions, rabbits exhibit distinct breeding seasons that are influenced by both day length and temperature. In the northern hemisphere, rabbits in natural conditions exhibit their highest conception rate in spring and their lowest rate in autumn. When environmental conditions are controlled, male rabbits mate at any time. Maximal sexual receptivity in does is often accompanied by enlargement of the vulva, which also becomes reddish purple and moist. Although females will almost invariably mate in this condition, they also occasionally mate even when these changes have not occurred. Vaginal smears do not provide useful information. The most reliable indicator is lordosis, which occurs when the female is firmly clasped in the lumbar region. Ovulation occurs between 10 and 13 hours after copulation. To ensure successful ovulation, give a single intravenous injection of 100 IU of human chorionic gonadotropin to does after mating. Transportation of female rabbits may also result in spontaneous ovulation, with a resultant pseudopregnancy that lasts for approximately 18 days. Does will mate at the postpartum estrus. When the young from the previous pregnancy are removed, the willingness to mate persists for at least 36 days after parturition. If suckling of the young is permitted, however, sexual receptivity wanes. One study found that 100% of rabbits will copulate on the first day after parturition, 71% on the fourth day, 42% on the eighth day, and only 11% on the twelfth day.34 From the twelfth day onward, all of the does studied refused to copulate until they became receptive again, which occurred when the young began to feed themselves 50 to 60 days after parturition. Reproductive behavior is therefore minimal to absent as long as the mammary glands of the domestic rabbit are actively functioning. The lactating doe seldom conceives when mated before the eighth day after parturition. If a small litter of only 1 to 2 young is left suckling the doe, conception usually occurs (i.e., the percentage of fertile matings is inversely related to litter number).
PREGNANCY AND NURSING BEHAVIOR The gestation period in rabbits varies with the breed but is approximately 30 to 32 days. Litter size depends on the breed as well as on parity. Primiparous animals tend to produce smaller litters. Small breeds, such as the Dutch belted, produce small litters of 4 or 5 young, whereas larger breeds, such as the New Zealand white, produce large litters of 8 to 12 young. If pregnancy does not reach full term (for any or all of the fetuses), then the fetuses may be resorbed rather than aborted. There are two peaks
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for fetal resorption: days 11 to 15 and day 21. Fetal resorption is closely related to dominance and tends to be limited to subordinate does. Fetal resorption may have some advantageous role in energy preservation and masking signs to predators. Mummified fetuses, often floating in the peritoneal cavity or within the uterus, can be an incidental finding.60 Does have four pairs of mammary glands distributed within the fatty tissue of ventrolateral axillary, thoracic, abdominal, and inguinal areas.84 However, the number of teats can vary from 8 to 10. Suckling rabbits do not fix on any one teat. Consequently does are able to rear a larger number of offspring in comparison with the number of teats and rabbit producers selectively breed for supernumerary teats.27 Milk is stored mainly in the secretory alveoli. The milk ducts unite until there are 6 to 8 main ducts or galactophores, each draining a sector of the mammary gland, and these pass separately through the nipple. This arrangement allows for rapid secretion of milk by the doe during the brief daily nursing period. Several days to a few hours before parturition, the doe begins to collect hay, straw, or other similar material, which she carries to the nest site. In nature, this would be one of her burrows; in the domestic situation, it is usually the nest box. She then plucks hair from her abdomen, sides, and dewlap and interweaves it with the hay or straw to line the maternal nest.31 Normally the hair is tightly adhered to the skin but becomes loose near the time of parturition. Pregnant animals show a marked increase in hair loosening between 5 days prepartum and the day of parturition. The increase in hair loosening is also seen in a pseudopregnant rabbit at the termination of pseudopregnancy. The center of the nest is hollowed out, and this is where the young are generally born. In an excellent nest, the top is roofed over with nesting material, which effectively helps the neonates maintain their body temperature. Does usually give birth in the early morning. Normal parturition takes approximately 30 minutes. As the young are born, the mother usually eats the fetal placenta (placentophagia) and membranes and severs the umbilical cord.53 Placentophagia persists up to postpartum day 5 in about 50% of all rabbits. Occasionally the doe will inadvertently eat an offspring’s limb or ear during placentophagia, but the affected neonate usually survives. Rabbits are born altricial and weigh about 50 g (1.75 oz) at birth. They are sensitive to touch, temperature, and smell from birth. Their ears are functional from 7 days and their eyes open at 10 days. Neonatal rabbits recognize their mother by the scent of feces that she deposits in the nest. A gland in the region of the nipple produces a pheromone (2-methylbut-2-enal) that attracts the neonate.49 At birth, the neonates first suckle and then burrow deep into the nest material. Thereafter, a regular 22-hour cycle of behavior is followed. The doe feeds about once every 24 hours, with nursing occurring between the hours of 8:00 p.m. and 6:00 a.m., corresponding to the normal activity period of adults. Does nurse only once a day for 3 to 5 minutes. However, during this brief period, a young rabbit may drink 20% of its body weight. A healthy, well-fed litter bursts out of the nest like popcorn when it is examined. Within 15 minutes of nursing, all the young have reburied into the depth of the nest and group together for warmth. They conduct a continual slow circular movement to regulate differences in temperature at different levels of the nest. This is important, as neonatal rabbits are unable to thermoregulate on their own. After 22 hours, activity increases and they move to the surface of nest, sniffing the air for the doe’s return. Once attached to a nipple, they paddle with their forelimbs to stimulate milk let-down. Ejection of milk occurs toward the end
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of nursing period. Immediately after nursing, all the young urinate on the surface of the nest before reburrowing. Unique among altricial young, neonatal rabbits do not require stimulation to urinate. At about 8 days, they commence nibbling nest material, and by 12 days, they have often eaten most of it, including the doe’s fecal pellets. The ingestion of fecal pellets may aid in the colonization of the gut with important microflora. By day 18 of nursing, they begin leaving the burrow to explore as the doe “plugs” the entrance hole to the nest until the 18th day of nursing. After the 18th day, she ceases to perform this activity, thus encouraging the young to emerge. By 24 days, the pregnant doe abandons the young rabbits and they must fend for themselves. Olfactory cues are critical during the nursing period. Does mark their offspring with chin and inguinal gland secretions, and they are openly hostile to young rabbits that are not their own. They harass young from their own colony but hotly pursue and kill young rabbits from other colonies. A doe will attack and kill one of her own offspring if it is smeared with an odor from other rabbits.56 Successful cross-fostering requires that the young be healthy, that they have the energy to suckle, and that their scent is camouflaged by placing them on the bottom of the litter pile and rubbing them in the nest bedding.23
Box 12-1 Published Recipes for Rabbit Milk Replacer Cheeke’s Milk Replacer52 1 part evaporated milk 1 part water add to 1 cup of the mixture 1 egg yolk 1-tablespoon corn syrup
Taylor’s Milk Replacer 175 1 part Esbilac powder 0.25 parts heavy cream 1 part water
Taylor’s Milk Replacer 275 2 parts KMR liquid 1 part Multi-Milk powder
Taylor’s Milk Replacer 375 6 parts Esbilac 4 parts Multi-Milk (powder)
Taylor’s Milk Replacer 475
HAND-REARING OF BABY RABBITS Pet baby rabbits may require hand-rearing because of the death of the mother or because of poor nursing, which often occurs with a doe’s first litter. Most wild rabbits, including cottontail rabbits (Sylvilagus species) in North and South America, that someone brings into the clinic are probably not abandoned or orphaned. Unlike nursing cats and dogs, nursing does leave their nest, returning only once or twice a day to care for the young. Unless the wild neonate is injured, it should be left undisturbed in the nest. Immediate support and evaluation of the neonatal pet rabbit is a priority. Baby rabbits are very susceptible to hypothermia. Warming the neonatal rabbit is critical if its body temperature drops to less than 97°F (36°C). A rabbit can be warmed by immersing it in a warm (100°F [37.8°C]) bath and gently massaging it while keeping its head above the water. Because warming can worsen any dehydration, the neonate may require administration of isothermic (100°F [37.8°C]) crystalloids (10-15 mL/100 g body weight SC). Despite warming and fluid administration, weak neonates are often unable to nurse from a teat or syringe and require orogastric tube feeding. Use 5-Fr or smaller (human) infant feeding tubes and attempt to provide about 4 mL of rabbit milk replacement formula for newborns. The volume is then increased appropriately for weight gain and body size. Tube feeding permits control of how much milk the neonate receives and avoids the problem of aspiration of milk into the lungs. Numerous studies have looked at the quantity and composition of milk produced by lactating does. Neither breed nor pregnancy status affects milk composition. However, the stage of lactation significantly affects the major components of milk except lactose.25 Rabbit’s milk is higher in protein, fat, and ash than that of other species except rats. The milk of both rabbits and rats is high in fat and protein but low in carbohydrates. Several recipes for rabbit milk replacer have been published (Box 12-1). Milk replacer for neonatal and young rabbits is commonly a combination of Esbilac or KMR (Kitten Milk Replacer) and Multi-Milk (all available from PetAg Inc., Hampshire, IL; www.petag.com). Try to prepare enough fresh milk replacer
1 part Esbilac 1 part Multi-Milk powder 1.5 parts water
daily to allow for one day of feeding and keep it refrigerated between feeds. Rabbit breeders and wildlife rehabilitators often like to add a probiotic and infant multivitamin drops to the rabbit milk replacer; however, the effectiveness of this procedure has not been established. Although does feed their young only once a day, try to feed neonates three times a day to reduce bloat yet still provide adequate calories. Baby rabbits can take 2 to 3 days before they settle into a feeding pattern. Record the neonate’s body weight daily (or at every feeding time) as well as the amount of milk replacer consumed at each feeding. A neonate that drinks a small amount of milk replacer at one feeding may be greedy at the next. Although neonatal rabbits do not require stimulation of the perineum to urinate and defecate, we still recommend postfeeding stimulation by gently stroking the anogenital area with a warm, wet, cotton-tipped applicator or gauze sponge. Generally, this procedure is no longer necessary after 1 week of growth. Neonatal rabbits depend totally on milk up to day 10. They can digest a small amount (5%) of solid feed by day 15; by day 20, coprophagy is taking place and solid feed represents most of the feed intake.4 Hand-raised rabbits may take slightly longer to achieve these time points. By day 21, rabbits should be nibbling on hay and eating small amounts of solid feed. By day 28, rabbits should not be drinking milk replacement but can be offered water from a bottle.
MALE REPRODUCTIVE SYSTEM ANATOMY AND PHYSIOLOGY Unlike that of most other placental mammals but similar to that of marsupials, the rabbit’s penis is caudal to the testes. Rabbits do not have a scrotum but rather two hairless scrotal sacs; they
CHAPTER 12 Basic Anatomy, Physiology, and Husbandry do not have an os penis. Male rabbits have very small nipples that are not apparent under their fur. The testes descend at about 12 weeks of age, and the inguinal canals do not close. The technique of choice for castration must take into account measures to prevent inguinal herniation. Each testis is made up largely of closely packed seminiferous tubules. In many species, the tubules are arranged in lobules separated by bands of fibrous tissue. However, in rabbits (and many rodents), there are no subdivisions but tightly coiled tubules that apparently lie at random throughout the testis. When the testicular capsule is cut, the underlying tubules spill out like spaghetti. In species with lobular separation of tubules, testicular biopsies are possible without completely damaging the testis; this is not so in species without subdivisions, like the rabbit. The morphologic arrangement of the accessory sex gland complex of the male rabbit is unique. There is a lack of uniformity in the terminology used for the accessory glands: glandula vesicularis (glandula seminalis, vesicular seminalis), proprostata (glandula vesicularis, coagulating gland, prostata), prostata, paraprostata (glandula Cowperi superior), glandula bulbourethralis (glandula Cowperi inferior). The English equivalents are vesicular, proprostate, prostate, paraprostate, and bulbourethral glands. Holtz and Foote have described the anatomy of the male rabbit’s reproductive system in great detail.40
MALE SEXUAL BEHAVIOR AND REPRODUCTION Bucks show a constant libido after puberty. Initiation of copulation in rabbits is confined to basic patterns such as sniffing, licking, nuzzling, reciprocal grooming, and following the doe. Bucks may also exhibit tail-flagging and enurination, the emission of a jet of urine at a partner during a display of courtship. Experienced males usually initiate copulation within minutes or even seconds after a receptive female is introduced. Inexperienced males generally require longer. The species-specific copulatory patterns of the male are related to the ovulation and corpus luteal function of the female. Copulatory behavior of bucks may be understood by considering that the doe ovulates spontaneously after coitus. The stimulus of coitus is necessary only for ovulation and not for the maintenance of the corpus luteum, which always follows ovulation. Bucks rapidly mount receptive females and accomplish intromission after a series of rapid copulatory movements. Reflex ejaculation follows immediately on intromission. The copulatory thrust is generally so vigorous that the buck falls backward or sideways and may emit a characteristic cry. Vigorous bucks may attempt to copulate again within 2 to 3 minutes. A problem often encountered in male rabbits trained to use artificial vaginas is a reduced ability to induce lordosis during natural mating attempts. These conditioned bucks may become lazy. They fail to grasp and apply pressure to the flanks of the female in attempting to copulate and thus fail to induce lordosis. Bucks deposit semen into the anterior vagina, and the sperm pass individually through the cervical mucus. This pattern also occurs in sheep, cattle, and humans. If a rabbit is inseminated during an active corpus luteal phase (e.g., early pregnancy, pseudopregnancy), then sperm transport does not take place. During this period, when blood progesterone levels are increasing in rabbits (as in humans), the cervical secretions are thick and mucoid and inhibit sperm transport.
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BEHAVIOR The behavior of wild and domestic rabbits is very similar. The major difference is their response to confinement: wild rabbits do not adapt well to cages, often fail to breed, and exhibit abnormal behavior not seen under natural conditions. Other wild lagomorphs (e.g., hares) resist domestication and cannot be raised successfully in cages. In contrast, the domestication of rabbits has resulted in an animal that is not stressed by confinement while retaining most of the behavioral repertoires of its wild ancestors.
EATING, DRINKING, AND ELIMINATION BEHAVIOR Field studies of wild rabbits indicate that they are selective feeders with a wide food range. They prefer to eat tender, succulent plant parts as the major portion of their diet and consume small quantities of coarse roughage to stimulate gastrointestinal motility. Rabbits chew their food thoroughly, with highly organized tongue movements and up to 120 jaw movements per minute.17 Nutritional studies have shown that laboratory rabbits also adapt to a high-roughage diet. Wild rabbits can consume a large volume of fiber without the need for a large gut like ruminants because of the relatively rapid digestive transit time (19 +/- 10 h).22 Furthermore, their small, herbivorous gut does not interfere with the ability for swift evasion of carnivorous predators. The primary feeding times for rabbits are in the early morning and at night, with coprophagy commencing 3 to 8 hours after eating.14 Rabbits like sweet materials and select diets containing molasses or sucrose over similar diets without added sugar.14 This preference can be used to advantage when an anorectic rabbit is being encouraged to eat. Most pet owners feed their rabbits commercial pelleted diets because they are convenient and balanced in their formulation. Many of these diets are alfalfa-based and low in fiber. When pet rabbits are fed a diet of unlimited low-fiber pellets, they can become obese and develop chronically soft stools. Food restriction is practiced in some research laboratories to avoid these problems: New Zealand white rabbits weighing 2 to 3 kg are fed 100 to 120 g of pelleted diet per day. Food restriction in pet rabbits can lead to fur pulling and gnawing of carpets, furniture, shoes, and, sometimes fatally, electrical wires. Boredom and destructive behavior can be avoided by supplementing the diet with fresh grass, timothy or other hay, and a variety of vegetables and by providing gnawing toys such as a small log from an untreated fruit tree. Good-quality, high-fiber, timothy-based pellets formulated specifically for pet rabbits are available. Appropriate feeding of this type of pelleted diet helps to prevent obesity and ensures a high dietary fiber intake. Compared with other animals, rabbits have a high water intake. A rabbit’s average daily water intake is 50 to 150 mL/kg of body weight; a 2-kg rabbit drinks about as much water daily as does a 10-kg dog.16 When food is withheld from rabbits, they develop polydipsia; after 3 days of food deprivation, they can increase their water intake by 6.5 times.9 With water deprivation, food consumption declines; after 3 days of deprivation, anorexia results.16 The wild rabbit uses latrines or earth closets, which are special patches of ground near to the burrow to defecate. Latrines
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serve social functions, such as the recognition of their own or familiar odors, which may have a “comforting” or “confidence-enhancing” effect, and as loci for the indirect exchange of olfactory information between members of the same social grouping. Pet rabbits tend to void their droppings at one particular site, such as the corner of the cage. They can be trained to use a litter tray if they are constantly placed in the litter tray every few minutes when first acquired. However, adult bucks deposit strongsmelling feces in scattered places to mark their territory. Male house rabbits that are not castrated will mark their territory with a strong-smelling secretion by rubbing their chins on furniture, carpets, and household goods.
GROUP BEHAVIOR A unique aspect of rabbit behavior is the ability to form large groups or warrens. Rabbits are the only leporids that live in large, stable groups, sometimes numbering several hundred.61 The rabbit possesses no innate behavioral or physiologic mechanisms capable of regulating numbers at levels below the food ceiling. Although rabbits dig burrows that serve as primary havens and restrict their home ranges, they are active above ground, moving around, hopping, running, chasing, and playing. Rabbits also engage in “amicable” activities such as lying together, grooming, and nuzzling. These activities may occupy a considerable portion of the activity each day.48 Rabbits are fastidious groomers. They spend about 16% of their daily activity on grooming. Rabbits will lick their forepaws and wipe the secretions over their face and ears. They will also use their incisors and tongue to groom their backs, legs, haunches, and ventrum. Mutual grooming (allogrooming) is also performed. This usually occurs on the face, head, and ears of the companion. It may help to both relax and strengthen the social bond between rabbits.
VOCALIZATION, AUDITORY AND VISUAL SIGNALS Rabbits are considered silent animals with a limited vocal repertoire, in contrast to cats and dogs. However, a range of vocalizations and auditory signals may be produced to denote pleasure, pain, aggression, or fear. Some examples of these include soft clicking when contented; quiet teeth chatter while relaxing; loud teeth grinding when in pain; growl and/or hiss to denote aggression; soft humming by bucks before mating; loud shrill scream with intense pain or fear; and thumping of the hindfoot to signal a threat or danger. Visual signals are not a major part of a rabbit’s communication repertoire. This is primarily because rabbits spend much of their time in darkness. Some signs of visual communication are as follows: a relaxed rabbit may lie on its side with its hind limbs stretched out; a submissive rabbit may lie flat to the ground, adopting a small stature and flattening its ears to its body; a fearful rabbit may adopt a posture similar to that of a submissive rabbit; facial muscle contraction may also give the appearance of wide eyes; head shaking may denote an irritation or dislike of a smell or taste; head bobbing is an indicator of the response elicited by novel environmental stimuli; and displaying the white underside of the tail by raising the hindquarters while running signals to other rabbits to seek the safety of the burrow.
HUSBANDRY Rabbits require daily regular exercise to maintain good physical and mental health. Regular exercise may also stimulate normal defecation and urination, thus promoting gastrointestinal and urinary tract health. Access to unfiltered sunlight may also be beneficial to rabbits. Rabbits can be given free reign of a house, but if this is practiced, the home should be rabbit proofed. Rabbits like to chew and scratch objects found in a home; two of the greatest hazards are electrical cords and poisonous plants. Electrical wires should be placed out of reach of rabbits. The decorative houseplant dumbcane (Dieffenbachia seguinae) and the ornamental shrub oleander (Nerium oleander) are poisonous to rabbits.15 The alternative to rabbit proofing a home is to provide rabbits with supervised out-of-cage exercise/play time. Owners of house rabbits will attest to the fact that rabbits climb onto objects if they have the opportunity. In any housing or free-roam situation, rabbits should be provided with places to run and hide. Their natural tendency when frightened, especially from perceived predators, is to bolt underground into the safe haven of a burrow. These bolt-holes can be simulated by placing overturned boxes/containers or pipes around the area. If a pet rabbit is given free range in a house, it must have a cage or box into which it can escape. Divide the cage area that a rabbit occupies into two functional spaces: a space for lying and sleeping and a space for activities. A free-roam indoor rabbit requires only a cage that allows it to stretch out when lying on its side. A cage with a plastic bottom and a wire top is suitable because it can be easily cleaned and is well ventilated. If two or more pet rabbits are kept indoors, each animal should have its own cage, as fighting can result when two rabbits are kept in the same cage. An important feature of any rabbit enclosure is the substrate or flooring. The ideal substrate for rabbits simulates the compliant texture of earth. Many commercial hutches come with wire mesh flooring; the rabbit must be protected from this type of flooring. Rabbits are vulnerable to developing pododermatitis from wire-cage flooring and other noncompliant substrates. Provide rabbits with thick layers of soft bedding/substrates such as straw or grass hays on top of any flooring. These bedding materials should be changed regularly, especially if they become soiled or wet. Glass terrariums used for reptiles are not suitable for rabbits because they are poorly ventilated. Rabbits are sociable animals, so housing them with other rabbits is encouraged. Suitable groupings include mixes of neutered rabbits and/or intact females. Any attempted introductions should be monitored closely, as rabbits can inflict serious wounds upon each other and even fight to the death. Rabbits can be housed with other species; pet birds such as parakeets cohabitate well with rabbits. Take care in housing rabbits with some cats or dogs because the latter can potentially harm a rabbit. More often, rabbits reign supreme in mixed species households. Housing rabbits with guinea pigs is not recommended, as rabbits may subclinically harbor Bordetella bronchiseptica, which is pathogenic in guinea pigs. Some rabbits may also bully guinea pigs, and the nutritional requirements in these species are different. Outdoor rabbits should be housed in properly constructed hutches that provide shade and shelter from wind and cold below 39.2°F (4°C). Plans for the construction of hutches are available from libraries, feed manufacturers, and agricultural
CHAPTER 12 Basic Anatomy, Physiology, and Husbandry
Fig. 12-8 A grazing ark can be constructed from a solid frame and wire mesh. Alternatively, the metal lid from an indoor rabbit’s cage can be used as a small grazing ark. The idea of using an outdoor movable hutch in which rabbits could graze was first introduced by Major G. F. Morant in the United Kingdom in 1884. A grazing ark may also be known as a Morant hutch.
extension agents; assembled cages are available from farm supply stores or mail order houses that advertise in rabbit breeding journals. Professional and trade journals in laboratory animal science also advertise caging, but such caging is generally the most expensive. Space to move around in the hutch is important, and the space required for a rabbit to complete three hops is the minimum recommended length.48 A fully grown New Zealand white rabbit moves forward 1.5 to 2.0 m in three hops. The hutch should also be tall enough for a rabbit to stand up on its hind legs. Laboratory rabbits have been observed to climb onto raised platforms and shelves placed 2 m above the cage floor; these have been used successfully for rabbit housing.48 With the use of a grazing ark, indoor and hutch rabbits can feed on lawn.55 The ark can be a mesh top from an indoor rabbit cage or a solid frame with wire mesh (Fig. 12-8). Move the grazing ark every day to a fresh area of grass. Provide a shaded area within the ark, and peg the ark down so that the rabbit cannot tip it upward. Weed killers used on lawns can be poisonous to rabbits. If rabbits are housed outdoors or given free reign of a yard, measures must be taken to prevent their escape via digging, jumping, or climbing. Furthermore, predator proofing is essential. Subterranean wire mesh and adequately high fences with wire mesh “roofing” may be useful in some circumstances. Female rabbits more often perform digging than bucks. Does primarily dig burrows, and only pregnant or pseudopregnant females attempt to dig very deep tunnels.48,59 Behavioral enrichment can be provided by varying the environment. Penned rabbits often dig shallow holes in which they lie or sleep. Hide fresh, green food or treats in a heap of hay or a pile of soil for rabbits to root out. Cardboard boxes to explore and gnaw as well as varying straws, grass hays, tree branches, and wooden toys can be supplied for chewing purposes. Rabbits should ideally be protected from flies and mosquitoes. This may be difficult to achieve in some outdoor enclosures, but it is important because such insect vectors can spread the viral diseases myxomatosis and rabbit calicivirus. Rabbits tolerate cold better than heat. Because they do not possess brown fat, they shiver when exposed to cold. Shivering works well in the short term, and rabbits can tolerate cold weather if properly acclimatized and sheltered. Rabbits are unusually sensitive to elevated temperatures higher than 82°F (28°C) and have little protection against high ambient temperatures. They cannot sweat except through sweat glands confined to the lips; also, they pant ineffectively, and when sufficiently dehydrated, they stop panting. Rabbits do not increase
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water intake when the ambient temperature becomes high; heat actually seems to inhibit drinking. Rabbits can tolerate a water loss equivalent to 48% of their body weight. In contrast, when water loss reaches 11% to 14% of body weight in dogs, circulatory failure occurs. Although rabbits use their ears to dissipate heat, they actively seek shade and burrow to conserve water and shelter themselves from heat. The heart rate of rabbits increases only mildly in response to an increase in body temperature. Shelter from direct sunlight is essential in the design of any rabbit housing. The response of rabbits to high and low ambient temperatures is also important during transportation. The risk of mortality is greater in hot weather or in the presence of excessive indoor heating than in cold weather. A recent UK survey suggested that many rabbit owners were not aware of their pets’ needs.58 Over 50% of respondents believed that the average rabbit lives for only 3 to 4 years and 71% did not know that fiber, in the form of hay and grass, was a vital component of a rabbit’s diet.
References 1. Abdalla KEH, Abd El-Nasser M, Ibrahim IA, et al. Comparative anatomical and biochemical studies on the main bones of the limbs in rabbit and cat as a medicolegal parameters. Assist Vet Med J. 1992;26:142-153. 2. Ackerman S, Deeb B. Red urine: Blood or plant pigment? (Internet site). House Rabbit Society FAQs: House Rabbit Society. 3. Ade M. External morphology and evolution of the rhinarium of lagomorpha. With special reference to the Glires hypothesis. Mitt Mus Naturkunde Berl, Zoolog Reihe. 1999;75:191-216. 4. Alus G, Edwards NA. Development of the digestive tract of the rabbit from birth to weaning. Proc Nutr Soc. 1977;36:3A. 5. Barone R, Pavaux C, Blin PC, et al. Atlas of rabbit anatomy. [French, English]. Paris: Masson; 1973. 6. Bernard C. Experimental research on the sympathetic nervous system and especially on the influence that the section of this nerve exerts on animal heat. [French] Recherches expérimentales sur le grand sympathique et spécialement sur l’influence que la section de ce nerf exerce sur la chaleur animale. C R Soc Biol. 1853;5:77-107. 7. Beruter J, Colombo JP, Schlunegger UP. Isolation and identification of the urinary pigment uroerythrin. Eur J Biochem. 1975;56:239-244. 8. Birss JH. Some Americanisms of a hundred years ago. Am Speech. 1931;7:96-98. 9. Brewer NR, Cruise LJ. Physiology. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:63-70. 10. Briscoe JA, Syring R. Techniques for emergency airway and vascular access in special species. Semin Avian Exot Pet Med. 2004;13:118-131. 11. Campbell TW. Mammalian hematology: Laboratory animals and miscellaneous species. In: Thrall MA, ed. Veterinary hematology and clinical chemistry. Philadelphia: Lippincott Williams & Wilkins; 2004:211-224. 12. Capello V, Gracis M, Lennox AM. Rabbit and rodent dentistry handbook. Wiley-Blackwell; 2005. 13. Carey JR, Judge DS. Longevity records: life spans of mammals, birds, amphibians, reptiles and fish. Odense: Odense University Press; 2000. 14. Cheeke PR. Rabbit feeding and nutrition. Orlando: Academic Press; 1987. 15. Cheeke PR. Rabbit production. 6th ed. Danville, Ill: Interstate; 1987. 16. Cizek LJ. Relationship between food and water ingestion in the rabbit. Am J Physiol. 1961;201:557-566.
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17. Cortopassi D, Muhl ZF. Videofluorographic analysis of tongue movement in the rabbit (Oryctolagus cuniculus). J Morphol. 1990;204:139-146. 18. Coye DF. Orthoepic piracy: spelling pronunciations and standard English. Am Speech. 1998;73:178-196. 19. Cresswell J. Rabbit–Oxford dictionary of word origins. In: Cresswell J, ed. Oxford reference online. Oxford: Oxford University Press; 2002. 20. Crossley DA. Oral biology and disorders of lagomorphs. Vet Clin North Am Exot Anim Pract. 2003;6:629-659. 21. Cruise LJ, Brewer NR. Anatomy. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:47-61. 22. deBlas C, Wiseman J. The nutrition of the rabbit. Wallingford: CAB International; 1998;344. 23. Donnelly TM, Kelsey SF, Levine DM, et al. Control of variance in experimental studies of hyperlipidemia using the WHHL rabbit. J Lipid Res. 1991;32:1089-1098. 24. Eglitis I. The glands. In: Prince JH, ed. The rabbit in eye research. Springfield, IL: C.C. Thomas; 1964:38-56. 25. El-Sayiad GHA, Habeeb AAM, El-Maghawry AM. A note on the effects of breed, stage of lactation and pregnancy status on milk composition of rabbits. Anim Prod. 1994;58:153-157. 26. Fish W, Hyman GA, Ultmann JE. Survival studies on chromium-51-labeled erythrocytes in tumor-bearing rabbits. Cancer Res. 1956;16:885-889. 27. Fleischhauer H, Schlolaut W, Lange K. Influence of number of teats on rearing performance of rabbits. J Appl Rab Res. 1985;8:174-176. 28. Fox RR, Weisbroth SH, Crary DD, et al. Accessory spleens in domestic rabbits (Oryctolagus cuniculus). 1. Frequency, description, and genetic factors. Teratology. 1976;13:243-251. 29. Fontes-Sousa AP, Bras-Silva C, Moura C, et al. M-mode and Doppler echocardiographic reference values for male New Zealand white rabbits. Am J Vet Res. 2006;67:1725-1729. 30. Fontes-Sousa AP, Moura C, Carneiro CS, et al. Echocardiographic evaluation including tissue Doppler imaging in New Zealand white rabbits sedated with ketamine and midazolam. Vet J. 2009;181:326-331. 31. Gonzalez-Mariscal G, McNitt JI, Lukefahr SD. Maternal care of rabbits in the lab and on the farm: endocrine regulation of behavior and productivity. Horm Behav. 2007;52:86-91. 32. Grant TD, Specian RD. Proliferation of goblet cells and vacuolated cells in the rabbit distal colon. Anat Rec. 1998;252:41-48. 33. Greenaway J, Partlow G, Gonsholt N, et al. Anatomy of the lumbosacral spinal cord in rabbits. J Am Anim Hosp Assoc. 2001;37:27-34. 34. Hammond J, Marshall FHA. Reproduction in the rabbit. Edinburgh, London: Oliver and Boyd; 1925. 35. Harcourt-Brown F. Textbook of rabbit medicine. Oxford: Butterworth-Heinemann; 2002. 36. Harcourt-Brown FM. Metabolic bone disease as a cause of dental disease in pet rabbits. FRCVS thesis. London: Royal College of Veterinary Surgeons; 2005;174. 37. Hass GM, Brown DV, Eisenstein R, et al. Relations between lead poisoning in rabbit and man. Am J Pathol. 1964;45:691-727. 38. Hayes RA, Richardson BJ, Wyllie SG. To fix or not to fix: the role of 2-phenoxyethanol in rabbit, Oryctolagus cuniculus, chin gland secretion. J Chem Ecol. 2003;29:1051-1064. 39. Herrold EM, Goldweit RS, Carter JN, et al. Noninvasive laserbased blood pressure measurement in rabbits. Am J Hypertens. 1992;5:197-202. 40. Holtz W, Foote RH. The anatomy of the reproductive system in male Dutch rabbits (Oryctolagus cuniculus) with special emphasis on the accessory sex glands. J Morphol. 1978;158:1-20. 41. Hoyt Jr RF, Powell DA, Feldman SH. Exopthalmia in the rabbit after chronic external jugular catheter placement. Contemp Top Lab Anim Sci. 1994;33:A-19.
42. Janssens G, Simoens P, Muylle S, et al. Bilateral prolapse of the deep gland of the third eyelid in a rabbit: diagnosis and treatment. Lab Anim Sci. 1999;49:105-109. 43. Johnson LR, Drazenovich TL, Hawkins MG. Endoscopic evaluation of bronchial morphology in rabbits. Am J Vet Res. 2007;68:1022-1027. 44. Kamphues J, Carstensen P, Schroeder D, et al. Effects of an increasing supply of calcium and vitamin D on calcium metabolism in rabbits. [German] Effekte einer steigenden Calcium- und Vitamin D-Zufuhr auf den Calciumstoffwechsel von Kaninchen. J Anim Physiol Anim Nutr (Berl). 1986;56:191-208. 45. Landolt H. Ueber die Innervation der Thränendrüse. Pflugers Arch. 1903;98:189-216. 46. Lieber RL, Blevins FT. Skeletal muscle architecture of the rabbit hindlimb: functional implications of muscle design. J Morphol. 1989;199:93-101. 47. Loewen MS, Walner DL. Dimensions of rabbit subglottis and trachea. Lab Anim. 2001;35:253-256. 48. Love JA. Group housing: meeting the physical and social needs of the laboratory rabbit. Lab Anim Sci. 1994;44:5-11. 49. Luo MM. Got milk? A pheromonal message for newborn rabbits. BioEssays. 2004;26:6-9. 50. Marini RP, Foltz CJ, Kersten D, et al. Microbiologic, radiographic, and anatomic study of the nasolacrimal duct apparatus in the rabbit (Oryctolagus cuniculus). Lab Anim Sci. 1996;46:656-662. 51. Marini RP, Li X, Harpster NK, et al. Cardiovascular pathology possibly associated with ketamine/xylazine anesthesia in Dutch belted rabbits. Lab Anim Sci. 1999;49:153-160. 52. McNitt JI. Rabbit production. 8th ed. Danville, IL: Interstate Publishers; 2000. 53. Melo AI, Gonzalez-Mariscal G. Placentophagia in rabbits: incidence across the reproductive cycle. Dev Psychobiol. 2003;43:37-43. 54. Militaru M, Turcu D, Militaru D, et al. Lymphoid structures in the rabbit intestine: distribution, histology and ultrastructure. [Romanian] Formatiune limfoide din intestinul de iepure (Oryctolagus cuniculus), distributie, histologie si electronomicroscopie. Lucrari Stiintifice - Universitatea de Stiinte Agronomice Bucuresti Seria C, Medicina Veterinara. 1994;37:59-69. 55. Morant GF. Rabbits as a food supply, and how to fold them on our poor pastures. London: William Ridgway; 1883. 56. Myers K, Parer I, Richardson BJ. Leporidae. In: Walton DW, Richardson BJ, eds. Fauna of Australia: Mammalia. Canberra: Australian Government Publishing Service; 1989:917-931. 57. Mykytowycz R. Territorial marking by rabbits. Sci Am. 1968;218:116-126. 58. News and Reports. Survey reveals lack of knowledge of rabbit husbandry. Vet Rec. 2010;166:543. 59. Okerman L. Diseases of domestic rabbits. 2nd ed. Oxford: Blackwell Scientific; 1994. 60. Owensby T, Jackson K, Scharf B. Failure to deliver in a rabbit with intraabdominal masses. Lab Anim (NY). 2001;30:23-25. 61. Parer I. The population ecology of the wild rabbit (Oryctolagus cuniculus (L)), in a Mediterranean-type climate in New South Wales. Aust Wildl Res. 1977;4:171-205. 62. Parer I, Libke J. Distribution of rabbit, Oryctolagus cuniculus, warrens in relation to soil type. Wildl Res. 1985;12:387-405. 63. Pariaut R. Cardiovascular physiology and diseases of the rabbit. Vet Clin North Am Exot Anim Pract. 2009;12:135-144. 64. Patton NM. Colony husbandry. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:47-61. 65. Peacock DE, Sinclair RG. Longevity record for a wild European rabbit (Oryctolagus cuniculus) from South Australia. Aust Mammal. 2009;31:65-66. 66. Popesko P, Rajtova V, Horak J. A colour atlas of the anatomy of small laboratory animals: Rabbit, guinea pig. London: Saunders; 2002.
CHAPTER 12 Basic Anatomy, Physiology, and Husbandry 67. Prince JH. Anatomy and histology of the eye and orbit in domestic animals. Springfield, Ill: C.C. Thomas; 1960. 68. Prince JH. The rabbit in eye research. Springfield, Ill: C.C. Thomas; 1964. 69. Roberts MF, Zygmunt AC. Reflex and local thermal control of rabbit ear blood flow. Am J Physiol. 1984;246:R979-R984. 70. Sackmann W. Observations on the arrangement of Peyer’s patches in the small intestine of the rabbit (Oryctolagus cuniculus). Anat Histol Embryol. 1981;10:257-263. 71. Sandford JC. The domestic rabbit. 5th ed. Oxford: Blackwell Science; 1996. 72. Shadle AR. The attrition and extrusive growth of the four major incisor teeth of domestic rabbits. J Mammal. 1936;17:15-21. 73. Simon JF. Handbuch der angewandten medizinischen Chemie. Berlin: Albert Förstner; 1840. 74. Stott P, Jennings N, Harris S. Is the large size of the pinna of the ear of the European hare (Lepus europaeus) due to its role in thermoregulation or in anterior capital shock absorption? J Morphol. 2010;271:674-681. 75. Taylor KH. Orphan rabbits. In: Gage LJ, ed. Hand-rearing wild and domestic mammals. 1st ed. Ames, Iowa: Iowa State Press; 2002:5-12. 76. von Holst D, Hutzelmeyer H, Kaetzke P, et al. Social rank, stress, fitness, and life expectancy in wild rabbits. Naturwissenschaften. 1999;86:388-393.
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77. Wagner F, Beinecke A, Fehr M, et al. Recurrent bilateral exophthalmos associated with metastatic thymic carcinoma in a pet rabbit. J Small Anim Pract. 2005;46:393-397. 78. Weijs WA, Brugman P, Klok EM. The growth of the skull and jaw muscles and its functional consequences in the New Zealand rabbit (Oryctolagus cuniculus). J Morphol. 1987;194:143-161. 79. Wenger S, Barrett EL, Pearson GR, et al. Liver lobe torsion in three adult rabbits. J Small Anim Pract. 2009;50:301-305. 80. Whiteley HJ. Studies on hair growth in the rabbit. J Anat. 1958;92:563-567. 81. Williams CSF. Practical guide to laboratory animals. St. Louis: C.V. Mosby Co; 1976. 82. Williams DL. Laboratory animal ophthalmology. In: Gelatt KN, ed. Veterinary ophthalmology. 4th ed. Ames, Iowa: Blackwell Pub lishing; 2007:1336-1369. 83. Williams SB, Payne RC, Wilson AM. Functional specialisation of the pelvic limb of the hare (Lepus europeus). J Anat. 2007;210:472-490. 84. Yaldez K. Macroscopic-anatomical and electronical microscopic studies on the mammary gland of rabbits (Oryctolagus cuniculus). [German] Makroskopisch-anatomische und Rasterelektronenmikroskopische Untersuchungen an der Milchdruese des Kaninchens (Oryctolagus cuniculus). Dr Med Vet thesis Vienna: Veterinaermedizinische Universitaet Wien; 1996;105.
CHAPTER
13
Basic Approach to Veterinary Care
Jennifer Graham, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal), Diplomate ACZM, and Douglas R. Mader, MS, DVM, Diplomate ABVP (Canine and Feline)
Housing Handling and Restraint Physical Examination Sample Collection Blood Collection Collection of Urine and Feces Dermatologic Sampling Cerebrospinal Fluid Tap Treatment Techniques Catheterization and Fluid Therapy Injection Techniques Oral Medications Enteral Support Vaccinations Pain Control Miscellaneous Procedures Anesthetic Delivery Nasolacrimal Cannulation Ear Cleaning
Rabbits are popular companion animals that present to veterinary clinics for routine and emergency care. It behooves the veterinarian, therefore, to become familiar with basic techniques used on rabbits in a clinical setting. Clinics equipped for seeing dogs and cats can be easily adapted to accommodate rabbits. This chapter reviews common procedures specific to rabbits that may be performed by the clinician.
HOUSING Housing requirements can be readily met for rabbits. Hospitalized rabbits can be kept in caging designed for avian and exotic animals, stainless steel cages designed for dogs and cats, or 174
specially designed hutch cages. It is wise to place a thick towel on the bottom of the cage to prevent the rabbit from slipping on the smooth surface and injuring its back. A rubberized mat can provide the same traction and has the added benefit of allowing urine and feces to fall through, preventing soiling of the patient. A rabbit hutch can be easily and inexpensively constructed or purchased from a pet or feed store. Hutch cage units can also be adapted for use as tabletop cages with built-in catch pans or can be suspended with wire, as is commonly done in multianimal rabbitries. Cage floors should be constructed of 14-gauge wire mesh. The mesh openings should be rectangular and no greater in size than 1 cm by 2.5 cm. This facilitates cleaning and allows feces to drop through the floor but is not so large that a rabbit might accidentally get its foot stuck. A portion of the floor should be solid, giving the animal a place to rest and helping to prevent the rabbit’s hocks from becoming sore. Keep in mind that wood is difficult to sanitize and is not permitted in facilities regulated by the U.S. Department of Agriculture. Keep a supply of good-quality feed available for hospitalized rabbits. Rabbits can be finicky eaters, so check with the owner before hospitalizing a rabbit to find out what foods the rabbit prefers. If the diet is of poor quality, it may be necessary to offer the rabbit some of the food to which it is accustomed while gradually introducing a more appropriate diet. A rapid change of diet, even a change from a poor diet to a proper one, may cause gastrointestinal upset and anorexia (see Chapters 14 and 15).3 Fresh water should always be available. Consult the owner to learn what type of watering system the rabbit uses, although rabbits easily learn to drink from sipper bottles. If water crocks are used, they should be made of a heavy ceramic so that they will not easily be tipped over. Bowls with high sides are recommended because rabbits tend to hang their dewlaps in the water when they drink. If the sides of the bowl are too low, this chronic wetting can lead to “wet dewlap” disease, which is an easily preventable moist dermatitis most often associated with colonization by Pseudomonas species. All water containers should be cleaned daily. Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
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Fig. 13-2 Restrain a stressed rabbit by covering its eyes and supporting its hindquarters.
Fig. 13-1 Restrain the rabbit with one hand under its thorax and the other hand supporting its hindquarters.
If possible, hospitalize rabbits in an area separate from potential or perceived predators. Sounds and smells from hospitalized dog, cat, or ferret patients may stress sensitive rabbits.7 Care should be taken to avoid overheating hospitalized patients. Warm areas appropriate for sick birds or reptiles may cause heat stress in euthermic or hyperthermic small mammal patients.
HANDLING AND RESTRAINT The rabbit’s skeleton represents only 7% to 8% of its body weight (as opposed to 12%-13% in cats).8 With powerful musculature on the hind limbs and the delicate nature of the skeleton, rabbits are vulnerable to fractures of the back and hind limbs. Rabbits are easily stressed and care should be taken to reduce stress to the patient during examination. Proper handling and housing of patients is necessary to avoid injury. When a rabbit is being moved, support of the hindquarters is vital to prevent injury. Rabbits used to handling can be carried with one hand under the thorax or by holding the scruff, with the second hand supporting the hindquarters (Fig. 13-1). Fractious rabbits should be carried with the head under the handler’s arm so as to minimize stress by covering the eyes; the hindquarters are supported at the same time (Fig. 13-2). Place a rabbit in a cage with its rear facing the back of the cage while supporting the hindquarters to help reduce chances of injury from the rabbit kicking. In examining a rabbit or placing it in a cage, use a nonslip mat to avoid sliding and injury. Control of the rabbit should be maintained at all times during transport and examination. A towel can be used to wrap the patient and cover the head to prevent struggling when the rabbit is transferred from a carrier to a table for examination. In similar fashion, a towel can be wrapped around the rabbit if an assistant is not available to help facilitate physical examination or the administration of medicines (Fig. 13-3). It is important to avoid overheating when a towel is being used to facilitate handling or examination. As an alternative, place your hand over the eyes to calm the
Fig. 13-3 To help facilitate physical examination or the administration of medicines when an assistant is not available, a towel can be wrapped around the rabbit to restrain it.
rabbit. Rabbits are obligate nasal breathers, so care should be taken to avoid obstructing the nostrils with a towel or hand. Because some rabbits become calm when placed on their backs, the handler can sit on the floor with the rabbit in his or her lap, hindquarters toward the handler’s body. The incisors, abdomen, genitalia, and feet can then be examined (Fig. 13-4). It is preferable for the rabbit to be examined in a room away from “predator” species such as ferrets, dogs, and cats; noises or smells from these animals may stress the rabbit.
PHYSICAL EXAMINATION In general, performance of the rabbit’s physical examination is much like that of other species.6 Observe the rabbit at a distance prior to initiating examination. Take note of movement of the rabbit and document any neurologic or musculoskeletal abnormalities. Check for any difficulty breathing and monitor general stress level. Place the rabbit in a quiet, oxygenated cage prior to examination if dyspnea is noted. Thermal support should be provided if the rabbit is hypothermic, taking care to
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Fig. 13-4 Restraint of a rabbit while the handler sits on the floor with the rabbit in his or her lap, hindquarters toward the handler’s body. Examination of the incisors, abdomen, genitalia, and feet can be accomplished with this technique.
Fig. 13-5 To allow for examination of the ventrum, nail trimming, and obtaining body temperature, cradle the rabbit on its back in a C-shape with the hind end supported.
avoid overheating. Although rare, a rabbit may be too stressed to undergo an examination without sedation. A sedative such as midazolam, at a dosage of 0.5 to 1.0 mg/kg IM, is often very effective in calming the animal.4 A systematic approach is needed for the examination. Take the body temperature early because the body temperature may increase from the stress of the exam.1 A rabbit can be cradled on its back in a ‘C-shape’ with the hind end supported to allow for examination of the ventrum, trimming of the nails, and taking the animal’s temperature (Fig. 13-5). Avoid using a glass thermometer because it can break if the patient struggles. In examining the rabbit, use a nonslip surface to help keep the animal from slipping and injuring itself (Fig. 13-6). The heart and respiratory rates should be calculated before the rabbit is stressed. It can be challenging though, to differentiate heart and lung sounds on auscultation; respiratory sounds superimposed over the heartbeats may create the false impression of a heart murmur. Note any ocular or nasal discharge in examining the head. The tympanic membrane should be visualized on
Fig. 13-6 Use a nonslip surface to prevent the rabbit from slipping and injuring itself while it is being examined.
otic examination. Lymphadenopathy is rare in rabbits, whose lymph nodes are located as in other species. Skin and coat quality should be documented; the plantar surface of the feet should be examined for any signs of pododermatitis. A complete oral examination is an important part of the assessment. Palpate the face and ramus of the mandible for any evidence of swelling or tooth elongation. Although the mouth can be examined without using anesthesia, rabbits are often reluctant to allow access to the oral cavity. Therefore the examination may have to be performed using light sedation or general anesthesia. Lesions may be missed when otoscopic cones are used to examine the mouth of an unanesthetized patient. Nasal or vaginal specula with an attached light source can provide a better field of view than the otoscopic cone. Rabbit mouth gags and cheek dilators are commercially available; however, lesions are occasionally missed in live patients using these instruments. To examine the oral cavity in a sedated patient, place the mouth gag over the incisors to hold the mouth open and insert the cheek dilator. Dental cameras or endoscopy, good lighting, suction, and magnification can further aid in visualization of the oral cavity. Chapter 32 offers more details and sources of instrumentation. Palpate the abdomen for masses, distention, gas, or other abnormalities. Tensing of abdominal muscles or tooth grinding may be signs of discomfort. A firm, or dough-like stomach, increased gas or fluid within the intestinal tract, and absence of normal intestinal sounds on auscultation may be noted with gastrointestinal disorders.12 Document and investigate any organomegaly.
SAMPLE COLLECTION BLOOD COLLECTION Sites used for venipuncture in rabbits can include the marginal ear veins, central ear artery, jugular vein, cephalic vein, and lateral saphenous vein. Because hematoma formation, bruising, or vessel thrombosis and skin sloughing can result, use of the ear veins and, in some breeds, the central ear artery is not ideal. Use alcohol to part the fur and allow for visualization of the vessel; clipping or plucking of the fur may be useful in some
CHAPTER 13 Basic Approach to Veterinary Care
Fig. 13-7 The rabbit’s lateral saphenous vein.
cases. Avoid damaging the delicate skin of the rabbit if clippers are used. Cephalic veins are accessible in rabbits but can be difficult to locate and hold off in smaller breeds. Venipuncture of this vein, however, should be avoided so as to preserve the integrity of the vessels if catheter placement is necessary. Additionally, venipuncture of the cephalic veins may be more stressful for the animal as the handler is working near the rabbit’s head. The lateral saphenous vein is an ideal site for venipuncture in the rabbit (Fig. 13-7). If needed, restrain the rabbit in a towel with the head covered while a rear limb is gently extended. The restrainer holds off the vein with pressure across the proximal thigh. The vessel is readily accessible across the lateral surface of the tibia just proximal to the hock. Apply gentle pressure or a pressure wrap briefly over the venipuncture site to prevent hematoma formation after the sample is obtained. Rabbits have large paired jugular veins, which are often a preferred venipuncture site for very calm or sedated rabbits or when a larger amount of blood (e.g., for blood transfusion) is being collected. However, it is difficult to visualize the jugular vein in overweight rabbits or in females with large dewlaps. For jugular venipuncture, shave the neck over the midtrachea cranial to the thoracic inlet. Position the rabbit with the front legs held over the edge of the table and the head extended up, avoiding overextension of the head. Alternatively, wrap the rabbit in a towel and place it in dorsal recumbency, extending the head and neck to allow for visualization and sampling from the jugular vein (Fig. 13-8). While cardiocentesis in rabbits is commonly performed in research for obtaining large blood samples or terminal exsanguination under anesthesia, this technique is not appropriate for clinical settings. Tables 13-1 and 13-2 contain reference ranges for hematologic and plasma biochemical values in rabbits. Hormone levels in rabbits are presented in Table 13-3.
COLLECTION OF URINE AND FECES A rabbit’s bladder can be gently pressed to obtain a urine sample during examination. To decrease chances of iatrogenic bladder rupture, however, cystocentesis may be preferable to manual bladder expression. Cystocentesis in the rabbit is similar to that in other small animal species. Although sedation or anesthesia
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Fig. 13-8 Two-person jugular venipuncture.
Table 13-1 Reference Ranges for Hematologic Values in the Rabbit12 Erythrocytes Hematocrit Hemoglobin Mean corpuscular volume Mean corpuscular hemoglobin Mean corpuscular hemoglobin concentration Platelets Leukocytes Neutrophils Lymphocytes Monocytes Eosinophils Basophils
5.1-7.9 × 106/μL 33%-50% 10.0-17.4 g/dL 57.8-66.5 μm3 17.1-23.5 pg 29-37 g/dL 250-650 × 103/μL 5.2-12.5 × 103/μL 20%-75% 30%-85% 1%-4% 1%-4% 1%-7%
Table 13-2 Reference Ranges for Serum Biochemistry Values in the Rabbit12 Serum protein Albumin Globulin Glucose Blood urea nitrogen Creatinine Total bilirubin Cholesterol Total lipids Calcium Phosphorus Sodium Potassium Chloride Bicarbonate Amylase Alkaline phosphatase Alanine aminotransferase Aspartate aminotransferase Lacticate dehydrogenase
5.4-8.3 g/dL 2.4-4.6 g/dL 1.5-2.8 g/dL 75-155 mg/dL 13-29 mg/dL 0.5-2.5 mg/dL 0.0-0.7 mg/dL 10-80 mg/dL 243-390 mg/dL 5.6-12.5 mg/dL 4.0-6.9 mg/dL 131-155 mEq/L 3.6-6.9 mEq/L 92-112 mEq/L 16-38 mEq/L 166.5-314.5 U/L 4-16 U/L 48-80 U/L 14-113 U/L 34-129 U/L
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Table 13-3 Hormone Levels in 29 Spayed/Neutered Rabbits Hormones Progesterone 17-Hydroxyprogesterone Androstenedione Testosterone Cortisol
Range (ng/mL) 0.11-0.46 0.75-22.2 0.80-4.00 0.02-0.04 4.64-11.2
From Fecteau KA, Deeb BJ, Rickel JM, et al. Diagnostic endocrinology: blood steroid concentrations in neutered male and female rabbits. J Exot Pet Med. 2007;16:256-259.
Table 13-4 Reference Ranges for Urinalysis Values in the Rabbit Urine volume Large breeds Average breeds Specific gravity Average pH Crystals present
Casts, epithelial cells, or bacteria present Leukocytes or erythrocytes present Albumin present
20-350 mL/kg per day 130 mL/kg per day 1.003-1.036 8.2 Ammonium magnesium phosphate, calcium carbonate monohydrate, anhydrous calcium carbonate Absent to rare Occasional Occasional in young rabbits
From Quesenberry KE. Rabbits. In: Birchard SJ, Sherding RG, eds. Saunders manual of small animal practice. Philadelphia: WB Saunders; 1994:1346.
reduces the chance that the stressed rabbit will struggle and damage internal structures, tranquilizers are often unnecessary. With the rabbit in dorsal recumbency, stretch the animal by holding the hind limbs in one hand and the scruff in the other. Alternatively, wrap the rabbit in a towel, hold the animal firmly, and cover its eyes. Locate the bladder on the ventral midline just cranial to the pelvic brim. A small-diameter needle (22- to 25-gauge) attached to a sterile 6-mL syringe provides an adequate sample for complete urinalysis after preparation of the antepubic region. Ultrasound can help with bladder visualization and detection of any bladder abnormalities. Urinalysis reference values are presented in Table 13-4. Urethral catheterization can be performed in sedated or anesthetized patients to collect a urine sample or for therapeutic flushing of the bladder in cases of urinary sludging. In most rabbits, a well-lubricated 9-Fr sterile catheter can be used for cathe terization. In the female, place the animal in sternal recumbency and locate the urethral os on the floor of the vagina. Placing the male in a sitting position allows for extrusion and catheterization of the penis. Fresh feces can be acquired from the enclosure of the rabbit or from the animal during physical examination. A direct smear of fresh feces in saline should be examined microscopically for the presence of protozoal organisms; several nonpathogenic
protozoa may thus be found. Fecal flotation is used to diagnose coccidia, cryptosporidia, and helminths. Coccidiosis is a common cause of diarrhea in young rabbits.
DERMATOLOGIC SAMPLING Some rabbit mites may be visualized under low magnification of skin brushings. Cheyletiella, bacteria, and yeast can be found on acetate tape strips applied to the skin and examined microscopically.5 As in other mammalian species, skin scrapings can be used to diagnose some parasitic conditions including sarcoptic or demodectic mange. Fungal and bacterial cultures as well as biopsies can be performed on skin lesions. Dark-field microscopy is used to examine smears for Treponema paraluiscuniculi.
CEREBROSPINAL FLUID TAP The collection of cerebrospinal fluid (CSF) may be indicated to help diagnose neurologic disease. Anesthesia, not sedation, is mandatory. The techniques are similar to those used in cats. The best site to collect CSF is the cerebellomedullary cistern. Position the rabbit in lateral recumbency with the head flexed toward the chest at a 90-degree angle to the vertebral column. Shave the fur on the nape of the neck from the occipital protuberance to the level of the third cervical vertebra and laterally past the margins of the atlas. The cranial margins of the wings of the atlas and the occipital protuberance are the landmarks for needle placement. A 22-gauge, 1.0- to 1.5-in.-long spinal needle should enter the skin midway between these points and be directed perpendicular to the skin approximately toward the animal’s nose. A stylet is usually not necessary because of the relatively small size of most rabbits. After the needle has penetrated the dura and arachnoid membranes, watch carefully for the appearance of CSF. The fluid should be allowed to drip into empty glass or plastic tubes. Do not attach a syringe or manometer since the movements caused by syringe attachment will likely cause bleeding, contaminating the fluid (Allen Sisson, personal communication, 2009). If the fluid cannot be submitted to the laboratory right away, one drop of serum from the rabbit should be added to 10 drops of CSF to preserve the cells for cytological evaluation. If serum must be added for cell preservation to the other tube for delayed cytologic evaluation, another 10 drops must be collected in a separate tube for cell counts and protein analysis. Only 10 drops of fluid are needed in one tube if the fluid can be analyzed immediately. Normal values for constituents of CSF in rabbits are presented in Table 13-5. A lumbar puncture, not a cerebellomedullary injection, should be used if a myelogram is to be done, since the contrast media will generally not flow past an extradural obstruction if the injection is made cisternally.
TREATMENT TECHNIQUES CATHETERIZATION AND FLUID THERAPY While many hospitalized rabbits can be managed with subcutaneous fluids, critically ill animals require intravenous fluids via catheter in the cephalic (Fig. 13-9) or lateral saphenous vein. A 24- or 26-gauge catheter can be placed in smaller rabbits, while a 22-gauge catheter can be used in rabbits weighing more than 3 kg. Alternatively, a butterfly catheter can be taped in place for short-term or bolus fluid therapy. Although jugular catheters
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Table 13-5 Reference Ranges for Cerebrospinal Fluid in the Rabbit Constituent
Concentration
Glucose Urea nitrogen Creatinine Cholesterol Total protein Alkaline phosphatase Carbon dioxide Sodium Potassium Chloride Calcium Magnesium Phosphate Lactic acid Nonprotein N
75 mg/dL 20 mg/dL 17 mg/dL 33 mg/dL 59 mg/dL 5.0 U/dL 41.2-48.5 mEq/L 149 mEq/L 3.0 mEq/L 127 mEq/L 5.4 mEq/L 2.2 mEq/L 2.3 mEq/L 1.4-4.0 mg/dL 5.6-16.8 mg/dL
Fig. 13-10 Subcutaneous fluid administration in the loose skin located on the rabbit’s dorsum.
From Weisbroth SH, Flatt RE, Kraus AL. The biology of the laboratory rabbit. New York: Academic Press, 1974:65.
Fig. 13-11 Intramuscular injection into the large lumbar muscles on either side of the rabbit’s spine, just cranial to the pelvis.
Fig. 13-9 An intravenous catheter placed in the rabbit’s cephalic vein.
can be used, anesthesia is recommended for their placement to facilitate the procedure and reduce stress to the animal. Avoid placement of intravenous catheters in the marginal ear veins, because sloughing of the ear tips can occur. A rabbit’s normal water consumption is estimated to be 100 to 150 mL/kg per day. Care must be taken in administering intravenous fluids to rabbits to avoid volume overload; 50 to 70 mL/kg per day administered intravenously can usually be tolerated. Infusion pumps help to regulate fluid delivery. A combination of crystalloids and colloids can be used to treat hypovolemic shock. If peripheral vessels are collapsed from dehydration, intraosseous catheters can be placed in the proximal humerus, greater trochanter of the femur, or tibial crest. Administer fluids intraosseously until the patient is adequately rehydrated or an intravenous catheter is placed. Prior to inserting the intraosseous catheter, clip the fur and surgically prepare the selected site in the sedated rabbit. Using sterile gloves, palpate the site of insertion of and insert a spinal needle or intraosseous catheter ranging in size from 18- to 23-gauge and from 1 to 1.5 in. in length,
depending on the rabbit’s size, parallel to the long axis of the bone and into the medullary cavity. Flush the needle gently with sterile saline and attach a male adapter. Apply antimicrobial ointment to the insertion site and a light dressing applied to hold the catheter in place. The subcutaneous route of fluid administration can be used on most hospitalized rabbits that have normal blood pressure and are accepting oral feedings. Rabbits can easily tolerate 120 mL/kg per day of subcutaneous fluids divided into 2 or 3 treatments. The loose skin located on the dorsum of the rabbit is an ideal location for subcutaneous fluid administration (Fig. 13-10).
INJECTION TECHNIQUES Administer medications via the intraosseous, intravenous, intramuscular, subcutaneous, or oral route. Injection techniques in rabbits are similar to those used in cats. Intraosseous, intravenous, and subcutaneous routes are described in the discussion of catheterization and fluid therapy, above. Intramuscular injections can be given into the large lumbar muscles on either side of the spine, just cranial to the pelvis (Fig. 13-11). One person
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Fig. 13-12 Oral medication being administered to a rabbit
Fig. 13-13 Nasogastric tube in an obese adult male Flemish
wrapped in a towel in “burrito” fashion.
giant rabbit.
can restrain and administer an intramuscular injection in a rabbit by tucking the rabbit under an arm, as described under “Handling and Restraint.” To avoid damaging the sciatic nerve, the cranial aspect of the rear leg (i.e., the quadriceps) should be used if the hind limbs are used for injection. Because some medications, including enrofloxacin, may cause pain, necrosis, or abscessation when the agent is given undiluted into subcutaneous or muscular tissues, dilute such medications with fluids and administer them subcutaneously.
palatable. Because it will not easily pass through a nasogastric tube owing to its high fiber content, Oxbow Critical Care Fine Grind (Oxbow Animal Health) has recently been developed for use in feeding tubes. Syringe feeding is most often used in the anorectic rabbit but may be stressful to the animal. Several methods can be used to syringe feed a rabbit, depending on its size and demeanor.4 It is quickest to feed a rabbit directly out of a 60-mL syringe with a catheter tip if the animal will tolerate it. The tip of the syringe is introduced into the diastema and slowly depressed. In smaller patients or those rabbits that resist feeding, 1-mL syringes or small oral feeding syringes can be used. This method is more time-consuming but can be a very effective way to deliver the formula to some rabbits that would otherwise refuse feeding. Orogastric tubes can be used for single dosing but are inappropriate for chronic use. For orogastric feeding, premeasure and mark an 18- to 22-Fr round-tip rubber catheter for the distance from the mouth to the last rib. Keep the rabbit’s mouth open with a mouth gag, flex the rabbit’s neck, and pass the tube through the oropharynx into the stomach. Test the tube’s placement by auscultating as air is injected through the tube into the stomach or checking for negative pressure when the tube is backed into the esophagus. Unless the rabbit is very calm, sedation or anesthesia will be required to minimize stress. Nasogastric tubes are used in clinical settings but may be more stressful to maintain than syringe feeding. Measurement of the tube is as described for orogastric tubes, using the tip of the nose and the last rib as landmarks. A pediatric feeding tube (3.5 to 5 Fr) can be used for most rabbits. Place a topical anesthetic such as 2% lidocaine gel or several drops of pro paracaine (Ophthaine, Solvay Animal Health, Inc., Mendota Heights, MN) can be placed in the nasal opening several minutes prior to placement. Insert the tube through the ventromedial nasal meatus and pass it ventrally and medially with the head flexed. Check tube placement with a lateral radiograph or as described for orogastric tube placement. Secure the tube with a drop of cyanoacrylate glue on the furred area above the nose and with tape glued or sutured to the top of the head (Fig. 13-13).
ORAL MEDICATIONS Suspensions are preferable to tablets when medications are administered to rabbits orally. Tablets can be made into sus pensions by compounding pharmacists when needed, or, alternatively, some tablets can be crushed and placed into a favorite treat or jam. It is helpful to use a towel in which to wrap a fractious rabbit in “burrito” fashion when oral medications are being given (Fig. 13-12). Place the medication as far back as possible in the oral cavity to prevent the rabbit from spitting it out.
ENTERAL SUPPORT Anorectic rabbits are commonly presented for medical evaluation. Dental disease, gastrointestinal disease, neurologic disease, and other systemic diseases are often complicated by secondary anorexia.11 Failure to provide nutritional supplementation to an anorectic rabbit can result in hepatic lipidosis in as little as 2 to 3 days. Oxbow Critical Care (Oxbow Animal Health, Murdock, NE) is a timothy hay-based syringe-feeding formula that is mixed with water to provide an excellent high-fiber mixture for anor ectic herbivores. Reconstitution of the product is simple and results in a homogenous mixture. Soaking and blending pellets and greens with water is an alternative to this commercial diet but is more time-consuming to prepare and generally results in a less homogenous mixture. The Oxbow Critical Care diet has an excellent fiber level at 21% to 25% and is highly
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VACCINATIONS No routine vaccinations are recommended for pet rabbits.
PAIN CONTROL Pain control is important for successful recovery of the compromised rabbit (see Chapter 31). Pain assessment may be more difficult in rabbits than in other species because they typically do not vocalize when they are experiencing pain. The rabbit may sit very still in the back of the cage in a hunched position while grinding its teeth and being oblivious to its surroundings. Other clinical signs of pain in the rabbit may include decreased fecal production, head elevation, aggression, isolation, rapid shallow breathing.2 Preemptive analgesia is recommended whenever possible. Nonsteroidal anti-inflammatory drugs (NSAIDs) are estimated to be effective for 12 to 24 hours in rabbits, while the effects of opioid drugs may last for only a few hours. Buprenorphine is effective for 6 to 12 hours, while butorphanol is effective for 2 to 4 hours. Abdominal or visceral pain may respond better to opioid analgesia, while NSAIDs are generally more effective for somatic or integumentary pain. Multimodal analgesia, combining agents from different classes, should be more effective than a single agent alone and allows the agents to be used at lower dosages.
MISCELLANEOUS PROCEDURES ANESTHETIC DELIVERY There are a variety of ways to administer anesthetic agents to rabbits, including topical, injectable, inhalant, and combination protocols. Anesthesia involves many considerations, such as the stability of the patient, monitoring, anesthetic agents used, and others.9,10 Critically ill patients should be stabilized prior to anesthetic administration. Topical anesthesia may be useful prior to procedures such as catheter placement. A topical preparation containing 2.5% lidocaine and 2.5% prilocaine (lidocaine 2.5%/prilocaine 2.5% cream, Hi-Tech Pharmaceutical, Amityville, NY) enables percutaneous insertion of catheters into rabbit veins without causing any detectable pain or discomfort. After application, these topical preparations may take 45 to 60 minutes to become fully effective. A variety of injectable anesthetic/analgesic combinations have been used in rabbits (see Chapter 31). Injectable anesthetic protocols may include parasympatholytics, phenothiazines, benzodiazepines, alpha-2-adrenergic agonists, ketamine, propofol, and others. The veterinarian should be aware of the specifics of different anesthetic drugs used in rabbits; for example, the use of tiletamine/zolazepam has been associated with nephrotoxicity in rabbits. Epidural anesthesia/analgesia is becoming more commonplace with small mammals. Local anesthetics, alpha-2 adrenergic agonists, and opioid agonists have been injected into the epidural space of small mammals to control pain. The advantages of epidural anesthesia and analgesia include few or no systemic effects compared with intramuscularly or intravenously administered drugs, quicker recovery time because of decreased gas anesthetic needed, and postsurgical pain relief. Disadvantages may include inability to place a spinal needle because of
Fig. 13-14 Location of the nasolacrimal duct in a rabbit. The arrow indicates the opening of the single nasolacrimal duct medial to the lid margin in the conjunctiva of the lower eyelid.
the small size of the epidural and intervertebral space, potential for trauma to the spinal cord, and potential for death or serious complications if analgesia is administered incorrectly. Care must also be taken in calculating volumes of local anesthetic for infiltration so as to avoid toxicity. Inhalant anesthesia is the primary component of most anesthetic regimens in small mammals (see Chapter 31). Isoflurane and sevoflurane are commonly used inhalant anesthetic agents. Induction with an inhalant anesthetic is typically achieved either with an induction chamber or face mask. Rabbits, however, are generally premedicated to lower the excitatory response typically seen with induction. Restrain the animal carefully to prevent injury during this period. A variety of commercially made induction chambers and face masks are available for use with rabbits. Additionally, masks can be fashioned out of syringe cases or other materials. For maintenance of anesthesia, endotracheal intubation, using blind and direct techniques, is ideal to protect the upper airway and assist ventilation (see Chapter 31). To facilitate intubation, hyperextend the head and neck of the rabbit to allow for the alignment of the larynx and trachea with the oropharynx. Care should be taken to make sure that the rabbit is adequately premedicated and relaxed to allow for atraumatic intubation. In addition, it is important to avoid overextension of the neck, which can result in damage to the spine.
NASOLACRIMAL CANNULATION Nasolacrimal duct flushing is indicated to determine and/or restore patency of the nasolacrimal ducts when rabbits present with an ocular discharge. Common causes of nasolacrimal duct obstruction can include infectious agents and dental disease. The single nasolacrimal duct in a rabbit is located medial to the lid margin in the conjunctiva of the lower eyelid (Fig. 13-14). Most rabbits will allow flushing of the nasolacrimal duct without sedation after topical anesthetic drops are instilled. A lacrimal cannula or 24-gauge Teflon intravenous catheter can be used to flush the duct (Fig. 13-15).
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SECTION II Rabbits appropriate medical diagnostics to determine the cause of disease. In severe cases of otitis externa, rabbits may benefit from medical treatment to reduce inflammation prior to ear cleaning.
References
Fig. 13-15 Proper placement of nasolacrimal cannula for a nasolacrimal duct flush in a rabbit.
EAR CLEANING Ear cleaning may be indicated in cases of otitis externa. If a large amount of debris is present within the ear canal and the rabbit is anesthetized, insert a red rubber catheter into the ear canal and flush with warm saline. While using an otoscope, endoscope, or otoendoscope for visualization to minimize the chance of traumatizing the ear canal, an ear curette or cottontipped applicators can be used to carefully scoop out purulent debris. Alternatively, flush saline into the ear with gentle massage to soften the purulent debris, followed by removal of debris with a suction unit while visualizing the canal to avoid iatrogenic damage. Flushing should be avoided if the tympanic membrane is ruptured. Ear cleaning should be accompanied by
1. Antinoff N. Physical examination and preventive care of rabbits. Vet Clin North Am Exot Anim Pract. 1999;2:405-427. 2. Bradley T, Lightfoot T, Mayer J. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: WB Saunders; 2006;41–43. 3. Cheeke PR. Nutrition and nutritional diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:321-331. 4. Graham J. Common procedures in rabbits. Vet Clin North Am Exot Anim Pract. 2006;9:367-388. 5. Harcourt-Brown F. Textbook of rabbit medicine. Oxford: Butterworth-Heinemann; 2002:224–248. 6. Harkness JE, Wagner JE. The biology and medicine of rabbits and rodents. 4th ed. Baltimore: Williams & Wilkins; 1995:13–30. 7. Krauss AL, Weisbroth AH, Flatt RE, et al. Biology and diseases of rabbits. In: Fox JG, Cohen BJ, Loew FM, eds. Laboratory animal medicine. Orlando, FL: Academic Press; 1984:207-240. 8. Mader DR. Basic approach to veterinary care. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2004:147-154. 9. Meredith A, Crossley DA. Rabbits. In: Meredith A, Redrobe S, eds. BSAVA manual of exotic pets. 4th ed. Gloucester, Great Britain: British Small Animal Veterinary Association; 2002:76-92. 10. Meredith A, Flecknell P. Anaesthesia and perioperative care. In: BSAVA manual of rabbit medicine and surgery. 2nd ed. Gloucester, UK: British Small Animal Veterinary Association; 2006:154-165. 11. Oglesbee BL. The 5-minute veterinary consult: ferret and rabbit. Hoboken, NJ: Wiley-Blackwell; 2006:178–385. 12. Quesenberry KE. Rabbits. In: Birchard SJ, Sherding RG, eds. Saunders manual of small animal practice. 2nd ed. Philadelphia: WB Saunders; 2000:1493-1511.
CHAPTER
14
Gastrointestinal Physiology and Nutrition
Michelle L. Campbell-Ward, BSc, BVSc (Hons I), DZooMed, MRCVS
Rabbit Gastrointestinal Physiology Ingestion of Food Stomach Small Intestine Large Intestine Hindgut Flora and Fermentation Cecotrophy Motility Digestion and Absorption Energy Requirements Nutrition Protein Carbohydrate Fiber Fat Vitamins and Minerals Dietary Components Grass Hay Fresh Vegetables and Edible Plants (“Greens”) Commercial Mixes and Pellets Other Feed Items Water Summary of Dietary Recommendations
The rabbit is a monogastric, hindgut-fermenting herbivore with a complex and unique digestive physiology. Given the wild rabbit’s ecologic role as a small prey species, its digestive strategy has evolved to permit efficient digestion of fibrous vegetation without the need to store large volumes of food within the body. The gastrointestinal tract features a simple stomach, a well-developed cecum, and a particle-dependent separation mechanism within the proximal colon. Fine particles and Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
solutes are selectively retained as substrates for microbial fermentation within the cecum, while indigestible fiber is rapidly expelled.50 The production and ingestion of cecotrophs (packets of cecal contents) contributes significantly to the rabbit’s digestive efficiency. An appreciation of normal gastrointestinal physiology is essential to formulate appropriate diets, understand the pathophysiology of common diseases, interpret diagnostic tests, and aid formulation of effective treatment plans. The importance of nutrition in the maintenance of health in the pet rabbit cannot be overstated. Poor nutrition is a significant contributory factor in the development of many common dental, gastrointestinal, and behavioral disorders. Much of the published literature related to rabbit nutrition is derived from the production and laboratory rabbit industries. While the gastrointestinal physiology of all rabbits is similar, the nutritional needs of the average pet rabbit are vastly different from those of rabbits of commercial interest. This chapter summarizes the current understanding of rabbit gastrointestinal physiology and focuses on the dietary requirements of the pet rabbit.
RABBIT GASTROINTESTINAL PHYSIOLOGY INGESTION OF FOOD Rabbits are described as selective feeders; in feeding on natural vegetation, they select the most tender, succulent plant parts (i.e., the parts that are most nutrient-dense and lowest in available cell walls).12 Their natural diet consists of the preferred succulent buds and young leaves of bushes, and they routinely graze on grasses, weeds, and even the bark around bushes and trees.7 They have a relatively high metabolic rate and a fast feed-transit time (19 hours)32; moreover, the practice of selective feeding (often termed “concentrate selection”) allows them to meet their dietary requirements while minimizing the volume of food that must be eaten. Rabbits eat approximately 30 times per day (2-8 g of food per time) over 4- to 6-minute periods.48 The selection of food is based on olfactory cues and 183
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pH 5-6
Food intake
PC Consumption of solid foods
Milk and “milk oil” intake
1
2
3 4 5 Age (weeks)
SR
pH 1-2 6
7
IL
8
Fig. 14-1 Relationship between a rabbit’s age, type of food
CE
intake, and gastric pH.
tactile information obtained via the sensitive vibrissae around the nose and lips.27 The incisors have evolved to cut through vegetation with a vertical slicing motion, while the cheek teeth are responsible for grinding the food before it is swallowed. Jaw movements during mastication are reported to be up to 120 per minute5 and amylase-containing saliva is secreted continuously during the process. Jaw movements feature a lateral motion, which helps keep the constantly growing teeth worn down to maintain effective occlusal surfaces. Hunger is stimulated by a dry mouth and gastric contractions or a decrease in blood levels of metabolites such as glucose, amino acids, volatile fatty acids, or lactic acid.18
STOMACH The rabbit’s alimentary tract features a simple stomach comprising approximately 15% of total gastrointestinal volume. There is a well-developed cardiac sphincter that prevents vomiting. Hydrochloric acid and pepsin are secreted into the stomach and initiate the digestive process in much the same way as in other species. The gastric pH of the rabbit varies diurnally but is generally very acidic compared with that of other species; postprandially it can drop to as low as 1.0 to 2.0, whereas following the ingestion of cecotrophs it rises to 3.0.4 This low pH effectively sterilizes ingesta. Juvenile (preweaned) rabbits have a much higher gastric pH (5.0-6.5) (Fig. 14-1), which promotes the survival and passage of ingested bacteria, facilitating the establishment of the vitally important large intestinal flora.27 The stomach usually contains hair, food, and fluid even after 24 hours of fasting or anorexia. Gastric transit time is approximately 3 to 6 hours.10
SMALL INTESTINE The small intestine of the rabbit is the primary site of absorption of many nutrients, such as amino acids, lipids, monosaccharides, and electrolytes. A wild-type diet, however, contains very few of these nutrients in a form suitable for absorption as the ingesta pass from the stomach into the small intestine. From a nutritional perspective, vegetation is predominantly composed of lignin, cellulose, hemicelluloses, oligosaccharides, pectins, and plant proteins bound to cell wall constituents, and these pass through the small intestine largely unaltered. Conversely, nutrients present within ingested cecotrophs are available in a
Fig. 14-2 The ileocecocolic junction of the rabbit. IL, Ileum; SR, sacculus rotundus; PC, proximal colon; CE, cecum.
form that does permit absorption across the small intestinal epithelium. Transit time through the small intestine is fast: 10 to 20 minutes through the jejunum and 30 to 60 minutes through the ileum,10 which limits the time available for absorption at this site. Bicarbonate is secreted into the proximal small intestine and functions to neutralize the acidic digesta leaving the stomach. The bicarbonate is then absorbed in the jejunum. Unlike the situation in many other mammals, pancreatic amylase is not a significant contributor to digestion.27 To complement its role in the absorption of nutrients, the small intestine appears to have an important immune function. There are extensive aggregates of lymphoid tissue (Peyer’s patches) throughout the intestinal tract of the rabbit, most prominently in the distal jejunum. Additionally, the terminal ileum enlarges into a dilation known as the sacculus rotundus at the ileocecocolic junction (Fig. 14-2). The sacculus rotundus (also known as the ampulla ilei or ileocecal tonsil) is unique to the rabbit and is made up primarily of lymphoid tissue. Once ingesta leave the small intestine, retrograde movement from the large intestine back into the ileum is inhibited by a valve-type mechanism at the ileocecocolic junction.
LARGE INTESTINE The large intestine is highly developed in the rabbit; it comprises the cecum and the colon. In very basic terms, the cecum acts as a fermentation vat facilitating the breakdown of digestible fiber and starch, while the colon sorts and packs the excreta. The digestive process in the rabbit differs from the usual cecal and colonic fermentation process in many other herbivores in that fiber is eliminated as rapidly as possible. The colon is functionally divided into two parts: the proximal colon, which is grossly identified by the presence of distinct haustra (sacculations), and the distal part, which is devoid of haustra. The proximal colon is the site of separation of ingesta into the digestible and indigestible fractions. The separation process is facilitated by a combination of functional anatomy and colonic motility. The digestible particles (which tend to be shorter) settle near the mucosa and are subsequently propelled in a retrograde direction back into the cecum via a series of coordinated contractions. At the same
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B
C Fig. 14-3 Mechanism of selective retention of particles by the rabbit intestinal tract. A, Mixed ingesta move from the ileum into the large intestine. B, Contractions of the large intestine direct ingesta to the proximal colon, where the long fiber particles tend to accumulate in the lumen and the smaller digestible particles and fluid aggregate near the mucosa. C, Coordinated contractions direct the long fiber particles in the lumen aborally toward the distal colon to form hard fecal pellets; meanwhile the smaller particles and fluid are directed retrograde into the cecum, where they will act as a substrate for microbial fermentation. time, the longer, coarse, indigestible particles tend to remain in the center of the colonic lumen and are directed aborally to form hard fecal pellets (Fig. 14-3).29 The indigestible fraction (sometimes referred to as the “scratch factor”) is not broken down structurally and therefore does not contribute directly to the acquisition of energy or nutrients; nevertheless, it is absolutely crucial for the stimulating normal gastrointestinal motility and for maintaining the vital physiologic processes of cell regeneration, secretion, digestion, absorption, peristalsis, and excretion. At the junction between the proximal and distal colon there is an area of thickened circular muscle that is densely innervated and vascularized. This structure is termed the fusus coli and is unique to lagomorphs. It acts as an “intestinal pacemaker,” controlling segmental, peristaltic, and haustral colonic motility and is responsible for mechanically squeezing water and electrolytes from hard feces prior to their expulsion. Reabsorption of water and solutes removed in this fashion occurs in the distal colon. The fusus coli (and hence intestinal motility) is influenced by hormones such as aldosterone and prostaglandins.27 The cecum is the largest organ in the abdominal cavity and has 10 times the capacity of the stomach. It usually contains approximately 40% of the intestinal contents. It is very
thin-walled and coiled, ending in the blind-ended vermiform appendix.41 The appendix is rich in lymphoid tissue and also has a secretory function (water and bicarbonate). The cecum receives the short particles and fluid selectively retained by the proximal colon; therefore its contents are generally semifluid in consistency. Microbial fermentation is the primary mechanism by which nutrients are released from ingested food, and the retained particles directed from the proximal colon to the cecum provide the substrate for the indigenous population of cecal microorganisms. Some of the products of fermentation are absorbed directly through the cecal wall, while many others are expelled and reingested as cecotrophs.27 The nutrients present within cecotrophs are made available to the rabbit when they pass through the stomach into the small intestine. Excretion of feces follows a marked circadian rhythm, alternating between the hard-feces phase (which coincides with feeding activity) and the cecotroph phase (which usually coincides with a period of rest at least 4 hours after feeding); this rhythm is influenced by diet, age, and reproductive status.10 Like the small intestine, the large intestine features significant amounts of gut-associated lymphoid tissue (GALT). In fact, the large intestinal GALT accounts for over 50% of total lymphoid tissue in the rabbit.46
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HINDGUT FLORA AND FERMENTATION To facilitate the digestion of selectively retained materials, the rabbit’s cecum has a well-established autochthonous population of microorganisms. These microorganisms produce the volatile fatty acids acetate, butyrate, and propionate,23 which provide up to 40% of the rabbit’s maintenance energy requirement.38 The proportion of the three volatile fatty acids varies according to the time of day, the diet, and the rabbit’s developmental stage. The microbiology and ecology of the rabbit gastrointestinal tract has not been as extensively studied as that of ruminant species or humans,52 but it is widely agreed that across all mammalian species a healthy intestinal flora is vital for systemic health. The indigenous intestinal microflora is essential for digestion but also serves a protective role in relation to potential pathogens. Studies to date have identified an extensive list of organisms that may be present, including a variety of anaerobes,52 gram-positive and gram-negative facultative anaerobes,53 large ciliated protozoa, and a rabbit-specific ascosporogenous yeast (Cyniclomyces guttulatulus).20 It is generally agreed that the strict anaerobes of the genus Bacteroides are the predominant organisms within the cecum, accounting for 109 to 1010 per gram out of a total bacterial load of 1010 to 1012 per gram of cecal contents.45 Coliform bacteria and Clostridium species are occasionally isolated from normobiotic rabbits; if they are present, however, they represent a very small percentage of the total bacterial population.52 In contrast with other mammals, rabbits have very rarely been found to harbor lactobacilli.35,55 Energy is the limiting factor for the cecal microbial population36 and, as is the case in other species, the composition of the microflora in any one individual does not remain constant. The cecal production of carbohydrates and nitrogenous substances, along with cecal goblet cell mucin production, supports abundant microbial synthesis.7 The nitrogen sources for cecal bacteria are predominantly ammonia, urea, and biuret. Disruption of the normal balance of microflora in the gut, usually with overgrowth of known or potential pathogens, is termed dysbiosis and is a common and serious clinical problem in rabbit medicine.11,53 Such alterations in normal homeostatic mechanisms within the gut may occur secondary to inappropriate therapeutic antibiotic administration, exposure to pathogenic organisms or toxins, increased glucocorticoid levels (iatrogenic or secondary to stress), gastrointestinal hypomotility, and poor dietary composition (low fiber, high carbohydrate, and high protein levels) (see Chapter 15). Suckling rabbits are unique among nursing neonates in that they feed for only 3 to 4 minutes in every 24 hour period30 and in that they produce an antimicrobial fatty acid referred to as “milk oil.” This oil results from an enzymatic reaction that occurs in the stomach following the ingestion of doe’s milk, and it appears to be a factor in the control of the gastrointestinal microbial content of young rabbits.16 Rabbits fed milk from other species do not develop this antimicrobial factor and are more susceptible to infection.27 There is virtually no microflora in the rabbit gastrointestinal tract at 3 days of age, and although some bacteria can be found in the small and large intestines over the following 3 weeks, the stomach remains largely devoid of microflora during this time. The production of milk oil wanes as the weaning process progresses and solids replace milk.16 In this transitional period, as the production of stomach oil ceases but before gastric pH has decreased to adult levels, populations
A B
Fig. 14-4 Rabbits produce two types of feces. Cecotrophs, or “soft feces,” (A) are produced by the rabbit according to a circadian pattern of intestinal motility and are usually eaten directly from the anus. They are soft, arranged in clusters, and have a mucous coating. The appearance and composition of hard feces (B) is distinctly different from that of cecotrophs; hard fecal pellets are predominantly composed of indigestible fiber and are generally round and dry.
of bacteria pass from the stomach to colonize the small intestine, cecum, and colon.
CECOTROPHY Rabbits produce two types of feces over the course of a day: hard feces and cecotrophs (also termed “soft feces” or “night feces”) (Fig. 14-4).53 These two excretory products differ markedly in composition,29 and in a healthy rabbit the excretion of only one type occurs at any one time. Hard fecal pellets are composed of compressed indigestible fiber (dry matter, 52.7%) that is separated from the remainder of the ingesta in the proximal colon, whereas cecotrophs (dry matter, 38.6%) contain semiliquid cecal contents and are rich in essential amino acids, volatile fatty acids, enzymes such as amylase and lysozyme, vitamins B and K, and microorganisms including bacteria, yeasts, and protozoa.7,27,50 The protein content of cecotrophs varies between 24.4% and 37.8%, of which 81% is in the form of bacterial cells.25 Cecotrophs are excreted according to a complex circadian pattern, which opposes that of voluntary food intake and hard feces production.10 Initiation of cecotroph formation occurs when segmental and haustral contractions of the large intestine are replaced by mass peristaltic activity resulting in the expulsion of material from the cecum.15 The expelled cecal content is then packaged and covered by a mucous envelope in the colon and passed as soft pellets approximately 5 mm in diameter and arranged in clusters. The transit time for cecotrophs through the colon is 1.5 to 2.5 times faster than that for hard feces.19 They are consumed directly from the anus, a practice referred to as cecotrophy (Fig. 14-5), and are swallowed without mastication.11 The ingestion of cecotrophs is triggered by stimulation of rectal mechanoreceptors, the perception of the specific odor of cecotrophs, and blood concentrations of various metabolites and hormones.18 The percentage of cecotrophs eaten varies depending on feeding regime and dietary composition. For example, fewer cecotrophs are consumed on a high-protein diet compared with a high-fiber one.
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motilin is not active in the cecum but is active in the colon and rectum.5 The fusus coli plays an important role in controlling colonic motility; it is of paramount importance in driving the proximal colon’s sorting system. Motility in the large intestine varies from the hard-feces phase to the cecotroph phase, whereas small intestinal motility is unaffected by the phase of excretion.
DIGESTION AND ABSORPTION
Fig. 14-5 The rabbit in the foreground is demonstrating cecotrophy (i.e., the ingestion of cecotrophs directly from the anus). Given the posture required to perform this normal physiologic function, it is clear that rabbits with mobility problems or spinal issues or those that are overweight frequently have difficulty practicing cecotrophy.
Following consumption, cecotrophs remain in the stomach of the rabbit for 6 to 8 hours. They are preserved in an intact state because of their protective mucous coating, allowing the microorganisms within them to continue the fermentation process. As the mucous layer dissolves, most of the cecal bacteria are lost to the low-pH environment of the stomach. A cecotroph may occasionally pass intact into the intestine. Cecotrophs provide the rabbit with microbial protein, which accounts for between 15% and 25% of the daily total amino acid requirement and between 9% and 15% of the digestible energy requirement.34 Lysozyme (a bacteriolytic enzyme) is secreted by the colon onto the cecotroph, enabling this microbial protein to become available for absorption by the time the cecotroph reaches the small intestine.8 The small intestine is also the site of absorption of amino acids and vitamins present within cecotrophs. Amylase is produced by bacteria present within the cecotroph and, along with salivary amylase, converts glucose into carbon dioxide and lactic acid. These products of digestion are absorbed in both the stomach and the small intestine.18 Cecotrophy must be considered an integral part of the rabbit’s digestive physiology; it improves feed utilization by maximizing the digestibility of nutrients.32 It may be thought of as a “pseudorumination” process, as redigestion facilitates the absorption of previously undigested nutrients. This circadian behavior occurs only in healthy rabbits and may have evolved as an adaptation to thrive in marginal or extreme habitats as well as during periods of poor food availability.29
MOTILITY Normal gastrointestinal function relies on a complex and highly coordinated pattern of intestinal motility, which is under the influence of the autonomic nervous system, hormones, and nutritional content (especially indigestible fiber levels). The enterochromaffin cells of the duodenum and jejunum secrete the polypeptide hormone motilin, which stimulates gastrointestinal smooth muscle. Fat stimulates the release of motilin and carbohydrate inhibits it. Over the length of the small intestine, motilin activity decreases aborally. In the large intestine,
Pepsin and hydrochloric acid begin the process of protein digestion in the stomach. The microbial protein mucin, however, can be digested only by lysozyme, which is secreted in the colon and incorporated into cecotrophs. Degradation of proteins continues in the small intestine with the aid of the pancreatic enzymes trypsin, chymotrypsin, and carboxypeptidase A and B. The intestinal mucosa secretes a broad range of aminopeptidases, and amino acids cross the jejunal brush-border membrane. Residual proteins that reach the large intestine unaltered are used by the cecal microflora. Starch and simple sugars are digested and absorbed in the small intestine. Although cellulose, hemicelluloses, pectin, and lignin are primarily digested in the cecum, it is proposed that a degree of prececal digestion is achieved by pectinases and xylanases of microbial origin in the stomach and intestine.24 Volatile fatty acids produced as an end product of cecal fermentation are absorbed across the cecal wall. Propionic and butyric acid are mainly metabolized by the liver and butyric acid is the preferred substrate for colonocytes, while acetic acid is available for extrahepatic tissue metabolism. As in other monogastric species, pancreatic lipase, sterol ester hydrolase, phopholase A, bile salts, and a colipase are responsible for lipid digestion. Calcium metabolism in the rabbit is unusual in that absorption of calcium from the gastrointestinal tract is not thought to be under the control of vitamin D3. While calcium is absorbed by active transport that requires a carrier protein synthesized in the intestinal mucosa in response to 1,25-dihydroxyvitamin D3, passive absorption also occurs, for which vitamin D is not required if dietary levels are adequate. The kidney is responsible for preserving or excreting calcium, mediated by parathyroid hormone (PTH) and vitamin D3.
ENERGY REQUIREMENTS The energy requirement of the rabbit in various physiological states has been described in detail in the production literature.42 Estimates of the digestible energy requirement of the nonlactating, nonpregnant doe vary from 326 to 400 kJ day-1 kg-1 BW0.75, where BW is the body weight.44,54 On the basis of the digestible energy content of most commercially available foods, this equates to 24 to 27 g of concentrate per kilogram of body weight for a 2- to 3-kg rabbit. Factors such as environmental temperature and activity level will affect energy utilization11 and should be taken into account in assessing maintenance requirements. Growth, pregnancy, lactation, and disease states will increase energy and nutrient demands. For example, the energy requirements for lactation are three to four times maintenance needs; if food intake is reduced during this critical period, toxemia of pregnancy, abortion, and dystocia may occur. Conversely, sedentary, geriatric pet rabbits are likely to have below-maintenance energy requirements. Although production rabbits (generally young, growing animals) may eat to meet their energy
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requirements and have been shown to adjust their voluntary food intake in response to changes in the diet,1 this is not the case with many pet rabbits. When given unrestricted access to highly digestible pelleted feed, rabbits may consume more food than is required for maintenance. Chronic overfeeding predisposes to obesity and may result in gastrointestinal disturbances.7
NUTRITION An appropriate diet for a rabbit aims to provide sufficient fiber to support normal gastrointestinal motility while ensuring adequate amounts and appropriate types of digestible nutrients are made available for fermentation by the cecal microflora. Additionally, the diet should promote normal foraging behavior throughout the day. As for all animals, inappropriate diet is a predisposing factor for many diseases in this species, including those of the gastrointestinal tract.
PROTEIN Rabbits obtain amino acids directly from the foods they ingest and from cecotrophs. Essential amino acids in the rabbit include arginine, glycine, histidine, isoleucine, leucine, lysine, methionine, phenylalanine, threonine, tryptophan, and valine. Grasses tend to contain limited amounts of methionine and isoleucine but are rich in arginine, glutamine, and lysine. Cereals generally have low levels of methionine and lysine; therefore synthetic amino acids are often added to commercial mixes. Legumes are high in lysine and may be used to balance the deficiency of this amino acid in cereal-based production diets.27 Rabbits appear to be able to digest forage-based protein particularly well (e.g., they can digest 75%-85% of alfalfa protein).32 This may be because of the increase in protein digestibility that occurs as a result of cecotrophy.36 Urea is recycled in the large intestine51 but is not well utilized by cecal microbes if fed as a dietary component. Excessive dietary protein can increase cecal ammonia levels and result in alterations in cecal pH. Cecal dysbiosis and overgrowth of pathogenic microorganisms may result.12 A protein level of 12% to 16% is considered appropriate for pet rabbits.41 The optimal level may increase to 18% to 19% for lactating does, but this is considered excessive for nonbreeding animals.
vices by encouraging normal foraging behavior.2,3 Although fiber digestibility is relatively poor in the rabbit, the indigestible fraction of dietary fiber is essential for stimulating gut motility and controlling gut transit time. Given the size-dependent separation of particles in the colon, manufacturing processes can influence the digestibility of commercial foods: the finer the grinding, the longer the gut transit and cecal retention times and hence the greater the potential for cecal dysbiosis.32 A balance has to be struck between providing enough indigestible fiber to maintain normal motility, cell regeneration, secretion, absorption, and excretion while at the same time providing sufficient digestible fiber for adequate bacterial fermentation in the gut.41 Fiber levels have an effect on appetite and cecotrophy (e.g., lowfiber diets depress voluntary food intake).1,22,47 For pet rabbits, total dietary fiber levels of 20% to 25% are recommended. This can be achieved in practical terms by providing an ad libitum source of indigestible fiber (e.g., grass and/ or hay) and ensuring that the amounts of other dietary components are limited, so that the primary fiber source is actually consumed. Commercial foods used as a portion of the diet should preferably have a crude fiber content of >18%, with indigestible fiber at >12.5%.27
FAT Fat provides a noncarbohydrate source of energy and improves palatability. It may also help to decrease dustiness and crumbling of pellets.11 Pet rabbits, however, are prone to obesity and hepatic lipidosis; therefore high-fat diets must be avoided. Dietary fat levels of 2.5% to 4% are considered appropriate. Some laboratory strains of rabbit are prone to the development of arteriosclerosis when placed on diets high in fat.9
VITAMINS AND MINERALS Vitamin and mineral requirements of the rabbit are summarized in Table 14-1.
DIETARY COMPONENTS GRASS
Simple sugars and starches are utilized as an energy source, but care must be taken to ensure that excessive levels are not fed. Because of the rabbit’s rapid gut transit time, starch and simple sugars may be incompletely digested in the small intestine, resulting in these compounds being directed into the cecum, where they may be used as substrates for fermentation by the cecal microorganisms. Carbohydrate overload in the cecum predisposes to enterotoxemia, especially in young animals.41 Owing to this potential for incomplete starch digestion, lowenergy grains such as oats are preferred over corn or wheat,12 and they should not be processed too finely.
Grass provides a balanced source of digestible and indigestible fiber, protein, vitamins, and minerals. In addition, grass is highly abrasive because of silicates and other materials that are present to varying degrees, depending on the species of grass and growing conditions. This abrasive factor is important for normal tooth wear.13 Generally crude fiber of fresh grass varies from 20% to 40%, while the protein content tends to be 15% to 19%.41 As a rule, the fiber content is higher in lower-protein grass and vice versa. Where practical, it is recommended that pet rabbits be given access to grass for at least several hours a day. Grass must be either grazed or fed freshly cut. Obviously this is not possible in all situations (e.g., for rabbits living in apartments in urban environments). Lawn clippings should not be used, as they ferment rapidly and may cause digestive disturbances.
FIBER
HAY
Fiber is a vital component of the pet rabbit’s diet. It is essential for the maintenance of gastrointestinal health and to promote normal dental attrition. It is also believed to prevent behavioral
Hay is considered an essential part of the pet rabbit’s diet and should be available ad libitum to rabbits that do not have free access to grass for grazing. For those that do have access to grass
CARBOHYDRATE
CHAPTER 14 Gastrointestinal Physiology and Nutrition
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Table 14-1 Summary of Vitamin and Mineral Requirements of the Domestic Rabbita Vitamin/Mineral
Dietary Requirement
Comments
Vitamin A
10,000-18,000 IU/kg
Vitamin B complex
Pet rabbits consuming fresh grass and vegetables are unlikely to be deficient; rabbits housed indoors and fed a diet of cereal mixtures and poor-quality hay without fresh grass or greens are potential candidates for deficiency (as cereals other than corn are a poor source); deficiency is associated with enteritis, retarded growth, weight loss, neurologic symptoms, keratitis, iridocyclitis, hypopyon, blindness, abortion, low fertility, stillbirth, neonatal death, and fetal malformations; excessive levels can lead to abortion, low fertility, and fetal malformations as well as hyperostotic polyarthropathy. Produced in the cecum; supplied by the cecotrophs.
None if animal is capable of cecotrophy 10-50 ppm recommended if Synthesized endogenously, but requirements may increase during times of stressed stress. Ideally provided by exposure to Synthesized in skin following exposure to sunshine and/or provided by diet; sunlight rather than dietary dietary sources for rabbits include sun-dried hay and supplemented supplementation; published pellets; although rabbits are able to absorb calcium without vitamin D, requirement is 800-1200 vitamin D will improve absorption if dietary calcium levels are low; excess IU/kg levels (>2300 IU/kg) have been associated with fetal mortality, depressed appetite, diarrhea, ataxia, paralysis, calcification of soft tissues, and death; deficiency has been associated with hypophosphatemia and osteomalacia. 50 mg/kg Deficiency is associated with muscular dystrophy, infertility, abortions, stillbirths, and increased susceptibility to coccidiosis. Unknown, but dietary Deficiency unlikely in this species; synthesized by cecal bacteria; most supplementation should not commercial pellets/mixes contain 1-2 mg/kg; consider supplementation be necessary if animal is in cases of coccidiosis, concurrent use of sulfa drugs, especially in capable of cecotrophy pregnant does and in situations where cecotrophy is inhibited. 0.5%-1.0% Unlike most other mammals, rabbits do not control calcium uptake from the gut; calcium uptake is not dependent on vitamin D; therefore serum levels of calcium increase directly in relation to increased dietary intake and excess calcium is excreted in the urine; availability of calcium for absorption can be related to other compounds found in food (i.e., phytates, oxalates, and acetates form complexes with calcium that reduce availability); deficiency can lead to a loss of appetite, tetany, muscle tremors, and death, especially in breeding does, and is thought to contribute to poor bone and dental health; excess calcium can result in urolithiasis, renal disease, and calcification of soft tissues (e.g., aorta and kidneys). 0.17%-0.32% — 1.0 ppm — 5-20 ppm — 0.4-2 ppm — 30-100 ppm — 0.3% Deficiency results in poor growth, alopecia, hyperexcitability, convulsions, myocardial fibrosis, and fur chewing. 8-15 ppm — 0.4%-0.8% Calcium-to-phosphorus ratio of 1:1 to 2:1 or higher is recommended; inverse Ca:P ratios will result in decreased bone density if overall phosphorus levels rise above 1%; availability is influenced by the presence of plant phytates and phytases; deficiency leads to rickets (in growing animals) and osteomalacia (in adults). 0.6% — 0.05 ppm Does not seem to have a sparing effect on vitamin E requirements and does not seem to be involved in white muscle dystrophies, as in other species. 0.2%-0.25% — 0.3%-0.6% Deficiency leads to immunosuppression, failure to gain weight, increased mortality, and increased severity of induced herpes simplex keratitis in experimental animals.
Vitamin C Vitamin D
Vitamin E Vitamin K
Calcium
Chloride Cobalt Copper Iodine Iron Magnesium Manganese Phosphorus
Potassium Selenium Sodium Zinc
aData
drawn from references 6, 7, 11, 12, 14, 17, 21, 27, 33, 36, 39, and 41.
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on a daily basis, hay can still be used to support the diet, especially at times when grass availability is poor. Hay is produced from a variety of pasture species; its quality and availability vary markedly between regions, seasons, and even individual farms. Grass or meadow hays are preferred and may be derived from timothy, prairie, brome, ryegrass, fescues, meadow grass, and orchard grass. Timothy hay is packaged in small quantities by several feed companies (e.g., Oxbow Pet Products, Murdock, NE; Kaytee Products, Inc., Chilton, WI). The fiber content of grass hays is usually around 30% to 35% and protein ranges from 6.3% to 16.7%.40 Alfalfa (lucerne) and to a lesser extent other legumes such as clover are used widely in some regions for haymaking. The grain hays, primarily oats and barley, are also available in some areas. Leguminous hays tend to be high in protein (16.5%), calcium (1.5%), and energy and thus are very useful for growing rabbits; however, legume-rich diets are thought to predispose to obesity and urolithiasis in mature nonbreeding animals. Prolonged storage of hay (especially in warm conditions) can lead to loss of nutrients such as vitamins A and D. Good hay is sweet-smelling and should not be musty. Hay can be fed from racks or nets to increase time spent feeding and thereby reduce boredom. Straw is not recommended because, although rabbits will eat it, it is low in nutrients and will lead to deficiencies if it constitutes a major part of the diet.41 The feeding of silage is generally not practiced, although it has been investigated in some countries in farmed rabbits. The high moisture content of rye and timothy grass silage was found to restrict dry matter intake and lead to a lower rate of growth.43 There is little information available at present regarding the feeding of silage to pet rabbits.41 Anecdotal reports on the use of artificially dried grass in rabbits sound promising.41 The product is palatable and, provided that it is not stored inappropriately or for too long, the nutrient content should be superior to that of sun-dried hay except for vitamin D, which is produced via the irradiation of sterols following exposure to ultraviolet radiation in sunlight.
FRESH VEGETABLES AND EDIBLE PLANTS (“GREENS”) The provision of a variety of fresh vegetables (green leafy varieties) and edible plants is recommended for all rabbits. These may provide vital micronutrients and, because of their high water content, are unlikely to contribute significantly to caloric intake. Their fiber content alone is insufficient to meet the needs of the rabbit’s gastrointestinal tract. These feed items may be bought commercially or grown fresh/harvested from the wild. Fresh greens, especially novel items that have not been given before, should be introduced to a rabbit’s diet gradually to allow the gut microbes to adapt, thereby avoiding gastrointestinal upset. Suitable vegetables include collard, mustard, and dandelion greens; carrot, beet, and broccoli tops; alfalfa sprouts and clover; herbs such as parsley, cilantro, and basil; lettuce, broccoli, cauliflower, green peppers, chicory, chard, watercress, celery leaves, endive, raddichio, bok choy, dock, spring greens, kale, and cabbage. Wild plants that can be offered include bramble, dandelion, chickweed, plantain, sunflower, wild strawberry, dock, and yarrow.49 As a rough guide, 2 cups of varied fresh vegetables and/or edible plants daily is considered appropriate for a 2.3-kg (5-lb) rabbit. Fruit should be used sparingly if at all because of the high sugar content and potential for carbohydrate overload in the hindgut:
a small amount (e.g., up to 1 tablespoon for a 2.3-kg [5-lb] rabbit) as a treat several times a week is unlikely to be problematic. Perishable vegetables, greens, and fruits must be stored in a refrigerator. Rinsing in fresh water before feeding is advised to remove any contaminants (fertilizers, insecticides, feces, urine) from the outer surfaces of the food.
COMMERCIAL MIXES AND PELLETS Commercial rabbit food has traditionally been sold in one of two forms: pelleted diets and mixed rations. Pellets are homogenous in appearance and are usually forage and/or cereal-based with a number of nutritional additives. Mixed rations (sometimes referred to as “muesli-type” feeds) consist of a variety of ingredients such as flaked, micronized, or rolled cereals, extruded biscuits, grass-based pellets, and plant stems. These two broad types of commercially prepared diets have been used for many years for commercial production and in laboratory rabbit husbandry and more recently have been widely marketed for use in pet rabbits. Despite marketing claims, these commercial concentrate rations are not essential components of the adult pet rabbit’s diet. If ad libitum hay, grass, and a variety of greens are available and cecotroph intake is normal, the diet is essentially balanced and contains sufficient energy for maintenance requirements. Nevertheless, many owners like to feed these products for convenience and, if used correctly as a supplement to the dietary mainstays of grass/hay and greens, they can be useful in the provision of some micronutrients and extra energy when required. While they should be fed in only limited quantities to adult nonbreeding pet rabbits, concentrates may play an important role in the nutrition of growing, pregnant, lactating, and diseased rabbits. Similarly, they can be used to ensure that nutrient requirements are fulfilled in rabbits that are unable to consume significant amounts of hay and/or green vegetables or those that are unable to practice cecotrophy (e.g., rabbits with advanced acquired dental disease or debilitating spinal issues). The nutritional composition of commercial feeds varies enormously, and this is true of both pellets and mixed rations. Both are calorie-rich in comparison with grass, hay, and greens. If used as part of the diet, some attention should be paid to the fiber and protein content. Commercial feeds with high protein levels, used for maximal growth and weight gain in rabbits raised for meat production, often have a lower fiber content in order to increase palatability. These foods, especially when fed in large quantities, can be associated with the development of gastrointestinal diseases. Of the two broad types of commercial feeds, rabbits tend to be better nourished when fed pellets. This is because they are unable to feed selectively12; therefore they ingest a consistent ration. Pet rabbits offered mixed-ration diets tend to favor flaked peas and corn, which are high in starch and low in calcium and fiber.26 Because most owners tend to replenish the feed bowl regularly, discarding uneaten items, the complete mixture of ingredients is rarely consumed. Additionally, some ingredients in the mixed rations (e.g., locust beans) have been implicated in cases of intestinal obstruction.27 Owner perception is an important factor in pet nutrition and many owners become easily misguided by the appearance or even the packaging of certain concentrate foods. Pelleted feeds should be hard and durable32 and be relatively high in fiber (ideally > 18%). Extruded diets are now very popular for pet rabbits, incorporating long fiber particles without the pellet becoming friable. Pelleted diets formulated from timothy
CHAPTER 14 Gastrointestinal Physiology and Nutrition hay (e.g., Bunny Basic/T, Oxbow Pet Products; Forti-Diet Rabbit Timothy Blend Adult Maintenance Formula, Kaytee Products, Inc.) are commercially available for adult pet rabbits. Ideally the indigestible fiber particles within the pellet should be 0.5 mm in length to stimulate gut motility. Small particles (<0.3 mm) can lead to increased gut retention time, reduced gut motility, and enteritis.32 There is considerable debate regarding optimal pellet size (with recommendations ranging from 3-5 mm in diameter and from 6-10 mm in length).36,37 The feeding of excessive amounts of concentrates (pellets or mixed rations), because of their energy content and nutritional inadequacies as well as the ease with which they are eaten, frequently leads to obesity, dental disease due to lack of wear,13 gastroenteropathies, and/or metabolic issues28 and also boredom-associated problems such as stereotypic behavior and aggression. In situations where the feeding of concentrates in addition to the hay/grass-based diet of a particular rabbit is considered appropriate, offering a measured daily amount is the preferred feeding practice. A general guideline of 25 g (maximum 30 g) of a pelleted diet per kilogram of bodyweight per day is appropriate for a rabbit with a normal body condition. Depending on the brand and pellet size, this roughly equates to ¼ cup of pellets per 2.3-kg (5-lb) rabbit. For overweight rabbits, this amount should be reduced or concentrates eliminated entirely from the diet. If desired, the daily ration can be divided into two portions per day. Proper feed storage and quick use prevent rancidity. Synthetic antioxidants and vitamin E may be added to feeds to increase shelf life. Optimally, feed should be stored at 60°F (15.5°C) in a vermin-proof area and used within 6 months (or before the manufacturer’s date of expiration) to ensure the maintenance of nutritional quality. Some commercial rabbit feeds may contain coccidiostats or other antimicrobial agents; such additives are not necessary for the routine feeding of pet rabbits.7
OTHER FEED ITEMS Carbohydrate-rich or high-fat foods such as beans, peas, corn, bread, breakfast cereal, nuts, seeds, chocolate, and some commercial “treats” should not be fed to rabbits. Treat items such as carrots or other root vegetables or small pieces of fruit can be used as a source of environmental enrichment (e.g., suspending them from the cage roof to act as edible toys). In this way, occasional favored items can be used to increase the time spent feeding without significantly increasing the caloric content of the diet or posing a risk of gastrointestinal imbalance.
WATER Rabbits need access to water at all times. Water intake is approximately 10% to 12% of body weight daily.31 A rabbit can go for several days without feed, relying on coprophagy. However, it cannot endure a lack of water for longer than 24 hours, even less in hot weather. Potable water should be free of harmful contaminants and provided in a manner that minimizes contamination by urine and feces. Drinking bottles are easier to keep clean than water bowls, and they avoid wetting the dewlap, which can lead to a moist dermatitis. If bowls are used, heavy flat-bottomed ceramic varieties or bowls containing a heavy rock may reduce tipping. Watering devices, such as water bottles with drinking tubes and automatic waterers, should be checked daily to ensure
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proper operation and availability of water. It may be necessary to train some rabbits that are accustomed to a different watering system in the use of automatic devices. The application of a sweet, sticky molasses or corn syrup to the surface of the automatic water delivery system helps the animal to find and use it.7
SUMMARY OF DIETARY RECOMMENDATIONS The recommended diet for a pet rabbit should encourage foraging throughout the day on high-fiber materials (freely available grass and/or grass hay). Varied daily rations of fibrous (green and leafy) vegetables should also be offered (e.g., 2 cups per 2.3-kg [5-lb] BW). Concentrates are not essential, but limited amounts (<30 g/kg BW or ¼ cup per 2.3 kg BW per day) of a high-fiber homogenous pellet can be offered as a supplement to a grass/hay-based diet provided that the rabbit is not overweight. Concentrate pellets and leguminous hays (e.g., alfalfa) are a useful way to provide the added nutritional demands of lactation, pregnancy, growth, and recovery from illness. Carbohydrate-rich treat items must be avoided to prevent hindgut overload and dysbiosis. The nutritional composition of commercial diets varies considerably; varieties that are low in fiber and high in protein, fat, and carbohydrate should be avoided. Mixed rations may promote selective eating and result in unbalanced diets. The overfeeding of concentrated pellets or mixes is the most common mistake owners make in relation to dietary provision for their pet rabbits. This practice will reduce overall feeding time, reduce fiber intake, and increase caloric consumption. It is not, therefore, surprising that inappropriate diet is the most significant factor in the development of gastrointestinal and dental diseases in rabbits. Weanlings are particularly susceptible to gastroenteropathies when dietary provision (especially fiber content) is suboptimal. Any dietary changes must be instituted slowly to allow the gut microflora to adjust. Regardless of the presenting problem, dietary advice should be given to any owner of a rabbit with a history suggestive of malnutrition.
ACKNOWLEDGMENT I would like to thank Dr. Jeffrey R. Jenkins for reviewing the manuscript and providing helpful comments.
References 1. Bellier R, Gidenne T. Consequences of reduced fibre intake on digestion, rate of passage and caecal microbial activity in the young rabbit. Brit J Nutr. 1996;75:353-363. 2. Berthelsen H, Hansen LT. The effect of hay on the behaviour of caged rabbits (Oryctolagus cuniculus). Anim Welfare. 1999;8:149-157. 3. Beynen AC, Mulder A, Nieuwenkamp AE, et al. Loose grass hay as a supplement to a pelleted diet reduces fur chewing in rabbits. J Anim Physiol Anim Nutr. 1992;68:226-234. 4. Blas E, Gidenne T. Digestion of starch and sugars. In: de Blas C, Wiseman J, eds. The nutrition of the rabbit. Wallingford: CABI Publishing; 1998. 5. Brewer NR, Cruise LJ. Physiology. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. London: Academic Press; 1994:63-70. 6. Brommage R, Miller SC, Langman CB, et al. The effects of chronic vitamin D deficiency on the skeleton in the adult rabbit. Bone. 1998;9:131-139.
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7. Brooks DL. Nutrition and gastrointestinal physiology. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2004:155-160. 8. Camara VM, Prieur DJ. Secretion of colonic isozyme of lysozyme in association with cecotrophy of rabbits. Am J Physiol. 1984;247:G19-G23. 9. Campbell-Ward M, Raftery A. Arteriosclerosis. Vetstream Lapis Clinical Reference. http://vetstream.co.uk/lapis, 2007. 10. Carabaño R, Piquer J. The digestive system of the rabbit. In: de Blas C, Wiseman J, eds. The nutrition of the rabbit. Wallingford: CABI Publishing; 1998:1-16. 11. Cheeke PR. Rabbit feeding and nutrition. Orlando: Academic Press; 1987. 12. Cheeke PR. Nutrition and nutritional diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. New York: Academic Press; 1994. 13. Crossley DA. Oral biology and disorders of lagomorphs. Vet Clin North Am Exot Anim Pract. 2003;6:629-659. 14. DiGiacomo RF, Deeb BJ, Anderson RJ. Hypervitaminosis A and reproductive disorders in rabbits. Lab Anim Sci. 1992; 42:250-254. 15. Erhlein HJ, Reich H, Schwinger M. Colonic motility and transit of digesta during hard and soft faeces formation in rabbits. J Physiol. 1983;338:75-86. 16. Fann MK, O’Rourke D. Normal bacterial flora of the rabbit gastrointestinal tract: a clinical approach. Semin Avian Exot Pet. 2001;10:45-47. 17. Feiler LS, Smolin G, Okumoto M, et al. Herpetic keratitis in zincdeficient rabbits. Invest Ophthalmol Vis Sci. 1982;22:788-795. 18. Fekete S. Recent findings and future perspectives of digestive physiology in rabbits: a review. Acta Vet Hung. 1989;37:265-279. 19. Fioramonti J, Ruckebusch Y. Cecal motility in the rabbit. III. Duality of fecal excretion [French]. Ann Rech Vet. 1976; 7:281-295. 20. Forsythe SJ, Parker DS. Nitrogen metabolism by the microbial flora of the rabbit caecum. J Appl Bacteriol. 1985;58:363-369. 21. Frater J. Hyperostotic polyarthropathy in a rabbit—a suspected case of chronic hypervitaminosis A from a diet of carrots. Aust Vet J. 2001;79:608-611. 22. Garcia J, Carabaño R, Perez-Alba L, et al. Effect of fiber source on cecal fermentation and nitrogen recycled through cecotrophy in rabbits. J Anim Sci. 2000;78:638-646. 23. Gidenne T. Nutritional and ontogenic factors affecting the rabbit caeco-colic digestive physiology. In Proceedings. 6th World Rabbit Cong. Vol 1. Lempdes: Association Française de Cuniculture; 1996:13. 24. Gidenne T, Carabaño R, Garcia J, et al. Fibre digestion. In: de Blas C, Wiseman J, eds. The nutrition of the rabbit. Wallingford: CABI Publishing; 1998:69-88. 25. Griffiths M, Davies D. The role of the soft pellets in the production of lactic acid in the rabbit stomach. J Nutr. 1963;80: 171-180. 26. Harcourt-Brown FM. Calcium deficiency, diet and dental disease in pet rabbits. Vet Rec. 1996;139:567-571. 27. Harcourt-Brown FM. Textbook of rabbit medicine. Edinburgh: Butterworth-Heinemann; 2002. 28. Harcourt-Brown FM. The progressive syndrome of acquired dental disease in rabbits. J Exot Pet Med. 2007;16:146-157. 29. Hirakawa H. Coprophagy in leporids and other mammalian herbivores. Mammal Rev. 2001;31:61-80. 30. Hoy S, Selzer D. Frequency and time of nursing in wild and domestic rabbits housed outdoors in free range. World Rabbit Sci. 2002;10:77-84. 31. Hrapkiewicz K, Medina L. Clinical laboratory animal medicine: an introduction. 3rd Ed. Ames: Blackwell Publishing; 2007. 32. Irlbeck NA. How to feed the rabbit (Oryctolagus cuniculus) gastrointestinal tract. J Anim Sci. 2001;79:E343-E346.
33. Kamphues VJ, Carstensen P, Schroeder D, et al. Effect of increasing calcium and vitamin D supply on calcium metabolism in rabbits. J Anim Physiol Anim Nutr. 1986;56:191-208. 34. Lebas F. Nutrient requirements of various categories of rabbits. In Proceedings. First Internat Feed Prod Conf, Piacenza. 1989:297. 35. Linaje R, Coloma MD, Perez-Martinez G, et al. Characterization of faecal enterococci from rabbits for the selection of probiotic strains. J Appl Microbiol. 2004;96:761-771. 36. Lowe JA. Pet rabbit feeding and nutrition. In: de Blas C, Wiseman J, eds. The nutrition of the rabbit. Wallingford: CABI Publishing; 1998:309-332. 37. Maertens L, Villamide MJ. Feeding systems for intensive production. In: de Blas C, Wiseman J, eds. The nutrition of the rabbit. Wallingford: CABI Publishing; 1998:255-272. 38. Marty J, Vernay M. Absorption and metabolism of the volatile fatty acids in the hindgut of the rabbit. Brit J Nutr. 1984;51:265-277. 39. Mateos GG, de Blas C. Minerals, vitamins and additives. In: de Blas C, Wiseman J, eds. The nutrition of the rabbit. Wallingford: CABI Publishing; 1998:145-176. 40. McDonald P, Edwards RA, Greenhalgh JFD, et al. Animal nutrition. 5th ed. Essex: Longman; 1996. 41. Meredith A. General biology and husbandry. In: Meredith A, Flecknell P, eds. BSAVA manual of rabbit medicine and surgery. 2nd ed. Gloucester: British Small Animal Veterinary Association; 2006:1-17. 42. Parigi Bini R, Xiccato G. Energy metabolism and requirements. In: de Blas C, Wiseman J, eds. The nutrition of the rabbit. Wallingford: CABI Publishing; 1998:103-132. 43. Partridge GG, Allan SJ, Findlay M. Studies on the nutritive value of roots, cabbage and grass silage for growing commercial rabbits. Anim Feed Sci Technol. 1985;13:299-311. 44. Partridge GG, Lobley GE, Fordyce RA. Energy and nitrogen metabolism of rabbits during pregnancy, lactation, and concurrent pregnancy and lactation. Br J Nutr. 1986;56:199-207. 45. Penney RL, Folk GE, Galask RP, et al. The microflora of the alimentary tract of rabbits in relation to pH, diet and cold. J Appl Rabbit Res. 1986;9:152-156. 46. Percy DH, Barthold SW. Pathology of laboratory rodents and rabbits. 3rd ed. Ames: Blackwell Publishing; 2007. 47. Pinheiro V, Guedes CM, Outor-Monteiro D, et al. Effects of fibre level and dietary mannanoligosaccharides on digestibility, caecal volatile fatty acids and performances of growing rabbits. Anim Feed Sci Tech. 2009;148:288-300. 48. Prud’hon M, Cherubin M, Goussopoulos J, et al. Evolution, au cours de la croissance, des caracteristiques de la consommation d’aliments solide et liquide du lapin domestique nourri ad libitum. Ann Zootech. 1975;24:289-298. 49. Richardson V. Rabbits; health, husbandry and diseases. Oxford: Blackwell Science; 2000. 50. Sakaguchi E. Digestive strategies of small hindgut fermenters. Anim Sci J. 2003;74:327-337. 51. Stevens CE, Hume ID. Comparative physiology of the vertebrate digestive system. 2nd ed. Cambridge: Cambridge University Press; 1995. 52. Straw TE. Bacteria of the rabbit gut and their role in the health of the rabbit. J Appl Rabbit Res. 1998;11:142-146. 53. Ward ML. Microbial composition of caecotrophs and the effect of probiotic supplementation on the caecal microflora of the healthy adult rabbit, Oryctolagus cuniculus. RCVS Diploma in Zoological Medicine Thesis London: RCVS Trust Library; 2007. 54. Xiccato G. Nutrition of lactating does. In Proceedings. 6th World Rabbit Cong. Vol 1. Lempdes: Association Française de Cuniculture; 1996:29. 55. Yu B, Tsen HY. Lactobacillus cells in the rabbit digestive tract and the factors affecting their distribution. J Appl Bacteriol. 1993;75:269-275.
CHAPTER
15
Gastrointestinal Diseases
Barbara L. Oglesbee, DVM, Diplomate ABVP (Avian), and Jeffrey R. Jenkins, DVM, Diplomate ABVP (Avian)
Gastrointestinal Stasis Syndrome The Role of Fiber The Effect of Diet and Cecocolic Motility History and Clinical Signs Physical Examination Findings Diagnostic Testing Treatment Acute Gastrointestinal Dilation or Obstruction History and Physical Examination Findings Diagnostic Testing Initial Medical Treatment Surgical Treatment Cecotrophy and Intermittent Diarrhea Cecoliths Enteritis Complex and Enterotoxemia Mucoid Enteritis Dysbiosis Caused by Treatment with Antibiotics Treatment of Enteritis Prevention of Enterotoxemia Bacterial Enteritis Viral Diseases of the Digestive Tract Papillomatosis Rabbit Enteric Coronavirus Rotavirus Rabbit Hemorrhagic Disease Virus Parasitic Disorders of the Gastrointestinal Tract Coccidia Cryptosporidia Other Protozoa Helminths Neoplasia Liver Lobe Torsion Aflatoxicosis Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
GASTROINTESTINAL STASIS SYNDROME Gastrointestinal (GI) stasis is by far one of the most common disorders seen in pet rabbits. Very often, it is the consequence on an inappropriate diet. However, any illness, painful condition, or stressful event can trigger an episode of GI stasis. It is imperative that the practitioner understand the need for continued, uninterrupted intake of an appropriate diet to ensure the proper functioning of the rabbit’s GI tract. Left untreated, GI stasis can rapidly become life-threatening.
THE ROLE OF FIBER To understand the pathogenesis of diet-related and GI diseases of the rabbit, a thorough knowledge of the normal anatomic and physiologic aspects of rabbit digestion must be appreciated. Details of this are discussed in Chapter 14. In summary, rabbits have a unique digestive physiology. They are strict herbivores, but their digestive strategy differs from those of other hindgut or cecal fermenters (e.g., horses) and ruminants. Being a prey species of small size, rabbits rely on a high energy intake while at the same time having the ability to quickly eliminate fibrous waste that would otherwise have to be carried in the digestive tract. To accomplish this, the rabbit’s cecum and colon have a well-developed mechanism for separating out the digestible and readily fermentable components of the diet while allowing the crude fiber components to be purged. The main driving force for this mechanism is the presence of large quantities of indigestible fiber. Lack of this fiber, either due to dietary inadequacies or conditions that cause anorexia, is the major cause of GI disease in rabbits.12 Digesta are separated in the colon in a process of selective retention of fluid and small particles. Normal peristaltic movements propel the larger fiber particles through the colon, while contractions of the haustra in the colon move the fluid and small particles retrograde to the cecum. Small particles and fluid are retained in the cecum, allowing for extensive fermentation. 193
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Cecal contents are expelled at regular intervals and reflexively consumed directly from the anus. Fiber stimulates cecocolic motility, either by a distention effect of the bulk, or directly. For example, diets high in fiber promote the production of specific volatile fatty acids in the cecum that directly promote peristalsis.12 Inadequate ingestion of coarse, nondigestible fiber will inhibit normal GI peristalsis. Rabbits will ingest hair routinely in the process of grooming. However, unlike many other mammalian pets, rabbits cannot vomit to eliminate accumulated hair. If GI motility is normal, ingested hair moves along with food out of the stomach at regular intervals and is ultimately expelled in the feces. If GI motility is impaired, either as a result of inadequate fiber intake or ileus secondary to anorexia (stress, concurrent disease), hair and normal ingesta accumulate in the stomach. With this accumulation, fluid is absorbed from the stomach, further compacting the contents. Compacted ingesta may cause discomfort to the rabbit, contributing further to anorexia and exacerbating GI hypomotility. A vicious cycle can result, until large amounts of hair and compacted ingesta accumulate in the stomach. This accumulation is erroneously referred to as a “hairball,” “wool block,” or “trichobezoar.” These terms imply, incorrectly, that the hair accumulation is the cause of disease in the rabbit rather than simply being the consequence or a symptom of impaired intestinal motility.
THE EFFECT OF DIET AND CECOCOLIC MOTILITY Cecal bacteria are vital to health. The cecum acts as a fermentation chamber and contains large populations of anaerobic organisms, such as Bacteroides species, and large anaerobic metachromatic staining bacteria (LAMB).9,17,31 Other bacteria normally present include gram-negative oval and fusiform rods, along with several nonpathogenic species of protozoa and amoeba.31 A rabbit-specific ascosporogenous yeast (Cyniclomyces guttulatus), in the Saccharomyces family, resides in the normal rabbit cecum.20 Veterinarians unfamiliar with rabbit fecal flora commonly mistake this yeast for coccidia on fecal examinations. This combined microflora is responsible for the processing of food particles entering the cecum into digestible nutrients, which are then reingested as cecotrophs. An inappropriate diet or GI stasis can disrupt the balance of this complex cecal microflora and the environment in which it grows. Diets low in fiber cause cecocolic hypomotility, prolonging the retention of digesta in the cecum and ultimately producing changes in cecal microflora. Populations of potentially pathogenic bacteria, primarily Clostridium species and coliform species such as Escherichia coli, are normally present in small numbers in the cecum.9,17,21 A slowing of cecocolic motility leads to the production of abnormal cecal fermentation products and alterations in the cecal pH. Even mild alterations in cecal pH will cause an increase in these pathogens as the populations of normal organisms decrease. Similarly, a decrease in peristalsis in the small intestine will allow normally small populations of potentially pathogenic bacteria to proliferate. Overgrowth of these pathogens can cause a range of pathology from mild diarrhea to death from enterotoxemia. Other effects of fiber consumption are indirect. High-fiber diets have a low level of available carbohydrates and thus decrease the risk of enterotoxemia caused by carbohydrate overload of the hindgut. Carbohydrates provide an environment in which pathogens
such as E. coli and Clostridium species proliferate. Glucose, a byproduct of carbohydrate digestion, is necessary for the production of iota toxin by Clostridium species. Thus, diarrhea and enterotoxemia in pet rabbits is often caused by this disruption in microflora, commonly referred to as dysbiosis. Feeds high in fiber, such as long-stemmed hay, can also protect rabbits from infectious bacterial enteritis. Rabbits have highly acidic stomachs, with the normal pH of 1 to 2. This acidic environment destroys most ingested bacterial pathogens, resulting in nearly sterile gastric contents. Ingested large-particle fibers, such as hay, mix with other foodstuffs to form a latticelike ball of food in the stomach. The lattice-like structure of this ball of food allows gastric acid to fully penetrate the ingesta and destroy bacteria. However, if a rabbit is fed primarily pelleted foods, a dense, compacted mass of food is formed in the stomach. The compacted nature of this food protects ingested bacteria from degradation by gastric acid, thus allowing potentially pathogenic bacteria to enter the small intestine. The combination of ingestion of pathogenic bacteria and slowing of intestinal motility as described above can result in clinical bacterial enteritis.
HISTORY AND CLINICAL SIGNS Obtain a complete dietary history, including the type and amount of commercial pelleted ration, hay, fresh leafy greens, and treats. Rabbits whose regular diet consists primarily of pelleted rations are at higher risk of developing GI stasis. The pelleted diets fed exclusively to feeder rabbits are high in calories (high in digestible carbohydrates), low in fiber, high in protein, and highly digestible, designed to increase weight gain in growing rabbits raised for their meat or fur. Commercial pellet formulations available for pet rabbits vary significantly in their protein, carbohydrate, and fiber contents. Formulations that contain a mix of dried fruits, vegetables, seeds, nuts, grains, and pellets generally have the lowest fiber content and the highest content of fat and carbohydrates. As the previous discussion indicates, there is significant potential for GI complications in a rabbit fed these diets. Similarly, rabbits routinely fed large amounts of high-carbohydrate, high-fat treats such as nuts, seeds, baked goods, and fruits are also predisposed to GI stasis. Acute episodes of GI stasis and dysbiosis are common following ingestion of a large volume of these treats. Rabbits at lowest risk are fed diets of unlimited grass or timothy hay, a moderate amount of fresh leafy greens, minimal commercial pellets, and occasional treats (see Chapter 14). Question the owner about recent potentially stressful events or underlying disease processes that may cause anorexia. Most rabbits with episodes of GI stasis have such a history, even if fed an excellent diet. Examples of stressful events include changes in housing, introduction of new rabbits or other pets, recent illness, trauma, or surgery. Common underlying disorders that may cause anorexia include dental disease, chronic upper respiratory tract disease, neurologic disorders, lower urinary tract disease, and renal or hepatic disorders. Many affected rabbits will also have a history of little or no routine exercise. The most common presenting complaint in rabbits with GI stasis is a gradual decrease in appetite and subsequent decrease in fecal production. Appetite usually decreases over a period of 2 to 7 days. Most rabbits will first stop eating pellets, then greens, hay, and finally treats. Left untreated, they stop eating all foods. Water consumption is often decreased as well. Feces become
CHAPTER 15 Gastrointestinal Diseases scant, dry, and small, eventually ceasing altogether. Owners may report a corresponding decrease in activity, usually as a result of abdominal pain. Rabbits in pain are reluctant to move, appear less social, may grind their teeth, may dig or scratch, and sit in a hunched position. Weight loss may also be noted, either due to anorexia or underlying disease.
PHYSICAL EXAMINATION FINDINGS Affected rabbits generally appear alert and quiet, exhibiting little or no sign of depression. Palpate the abdomen, with careful attention to the stomach contents, intestines, and cecum. The size of the stomach can vary with the duration of disease, but it always contains ingesta. The stomach usually feels firm
Fig. 15-1 Lateral radiograph of a rabbit with gastrointestinal stasis secondary to ureteral and renal calculi. Note the distention of the stomach with ingesta (arrows) and large amounts of gas within the intestines and cecum (arrowheads), characteristic of gastrointestinal stasis.
A
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and doughy and remains pitted on compression. Occasionally, stomach contents are severely dehydrated and solid. No fluid and little or no gas is palpable within the stomach of affected rabbits. This contrasts sharply with rabbits suffering from acute GI dilation (bloat), discussed below. The intestines and cecum frequently contain large amounts of gas. Little or no feces are palpable in the colon. Auscultate the abdomen, listening for borborygmus. Rabbits with GI stasis have few or no gut sounds. The remainder of the physical examination is usually unremarkable except for those findings related to any underlying disorder. Perform a thorough oral examination in all anorectic rabbits to look for underlying dental disease.
DIAGNOSTIC TESTING In many cases, the history and physical examination are sufficient for a diagnosis of GI stasis. It is very important, however, to look for an underlying cause. As discussed above, GI stasis occurs secondary to dietary or husbandry deficiencies, stress, pain, or underlying disease and will quickly recur if these are not addressed. Obtain additional diagnostics as indicated to rule out suspected underlying disorders. Radiography may or may not be helpful for diagnosis because the mass of food and hair appears similar to normal ingesta, even with contrast radiography. Visualization of a large, ingesta-filled stomach in a rabbit that has been anorexic for several days is suggestive (Fig. 15-1). This contrasts sharply with the dilated gas and/or fluidfilled stomach observed radiographically in rabbits with acute GI dilation (bloat) (Fig. 15-2) and can be helpful to differentiate the two disorders. Other common radiographic features seen in rabbits with GI stasis include moderate to severe distention of the intestines and cecum and scant fecal pellets. Radiographs may also be helpful to identify underlying causes of GI stasis.
B
Fig. 15-2 Survey lateral (A) and ventrodorsal (B) radiographs of a rabbit with acute gastrointestinal obstruction. Note the severely distended gas- and fluid-filled stomach and the lack of gas within the gastrointestinal tract distal to the stomach. This is consistent with an acute proximal small intestinal obstruction.
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TREATMENT The key principles in the treatment of GI stasis are to rehydrate the patient and stomach contents, alleviate pain, provide nutrition, and treat any underlying disorders. Fluid therapy is essential to recovery. Administer fluids via either the intravenous or subcutaneous route, depending on the severity of dehydration. If they are anorexic for more than 1 to 2 days, affected rabbits are usually significantly dehydrated and generally require hospitalization for intravenous fluid therapy. Rehydrate the stomach contents by assisted feeding with a syringe or placing a nasogastric feeding tube (see Chapter 13) if the rabbit will not accept feeding slurries orally. A commercial product (Critical Care for Herbivores, Oxbow Pet Products, Murdock, NE) is available for this purpose, and most rabbits will readily accept this product when offered via a syringe. This formula is relatively high in fiber and will provide nutrition needed to stimulate GI motility in addition to rehydrating the stomach contents. If this formula is not available, blenderized pellets soaked with water or an oral electrolyte solution can be used. Feeding these slurries will also help to prevent hepatic lipidosis, which can develop rapidly in a rabbit with a negative energy balance. In addition to assisted feeding, offer rabbits water, plenty of fresh hay, and a variety of greens to provide every opportunity to self-feed. Ingestion of food is critical to reestablishing GI motility. Rabbits with GI stasis have mild to severe gut pain, especially if the intestines are distended with gas. Most will not begin to eat until this pain is alleviated. If the rabbit appears to be much in pain (reluctant to move, hunched posture, bruxism), administer buprenorphine (0.01-0.05 mg/kg SC, IV q6-12h) and a nonsteroidal anti-inflammatory drug (NSAID) such as meloxicam (Boehringer Ingelheim Vetmedica, Inc., St. Joseph, MO) (0.3-0.5 mg/kg SC q12-24h) or carprofen (1.0-2.2 mg/kg PO q12h). Rabbits feeling less pain may respond well to NSAIDs alone. Be certain that the rabbit is fully hydrated and no underlying renal disorders are present prior to administering NSAIDs. If intestinal motility is severely impaired, parenteral administration is indicated. Other treatments include GI prokinetics and antibiotics. The use of intestinal prokinetic agents is somewhat controversial; however, anecdotal evidence suggests that they may be beneficial. Cisapride (0.5 mg/kg PO q8h) is readily absorbed from the GI tract and is available through compounding pharmacies. Metoclopramide (0.2-1.0 mg/kg PO, SC, IM q6-8h) has also been used. Antibiotics may be indicated if evidence of dysbiosis is present. If an overgrowth of Clostridium species is suspected, administer metronidazole (20 mg/kg PO q12h). Antibiotics generally effective against other potential pathogenic bacteria include enrofloxacin (15-20 mg/kg PO q12h) and trimethoprim-sulfamethoxazole (30 mg/kg PO q12h). Continue treatment for 3 to 5 days. Most rabbits will begin to eat and pass stool within 24 to 48 hours of treatment. Feces passed initially may look abnormal in shape, size, and consistency and may contain mucus or hair. Rabbits not responding to treatment should be reevaluated to identify underlying disorders contributing to anorexia. When treatment is successful, instruct owners to feed a diet that contains large amounts of grass or timothy hay, fresh leafy greens, and limited pellets and treats. Other popular remedies of questionable value include the use of lubricants (e.g., petroleum laxatives), protein-digesting enzymes (e.g., pineapple for bromelain and papain), and simethicone. These treatments alone will have no beneficial
effect as they do nothing to return function to the intestinal tract. Many rabbit associations and websites promote the use of simethicone, and it appears to have no ill effects. Caution owners against the use of protein-digesting enzymes, as these can be very irritating to oral mucosa and potentially gastric mucosa. The risk of gastric ulceration is increased in anorexic rabbits, and use of these enzymes may exacerbate this. Very rarely, the mass of material in the stomach is so dehydrated that it forms a solid, immovable mass and the rabbit fails to respond to medical treatment. In this case, surgical intervention may be necessary. However, the prognosis for a successful outcome is greatly reduced in rabbits that are treated surgically. Anesthesia, pain, stress, and manipulation of the intestinal tract all exacerbate GI stasis. Complications of hepatic lipidosis are a common cause of death in these patients.
ACUTE GASTROINTESTINAL DILATION OR OBSTRUCTION GI obstruction (also referred to as GI dilation or “bloat”) is an acute, life-threatening condition. The pathogenesis, history, and clinical findings differ significantly from those of GI stasis. Confusion arises over the use of the term trichobezoar. As discussed above, trichobezoar is a common misnomer for GI stasis. In most cases of acute GI dilation, the obstruction is located in the small intestine and is a compact mat of hair (trichobezoar). It is not clear exactly where these mats of hair are formed. They may have formed while still on the coat of the animal and have been ingested whole during grooming. It has also been suggested that these trichobezoars were once large cecal pellets containing compacted hair. They may occasionally be ingested and swallowed whole with other cecal contents during normal cechotrophy.24,25 Other foreign objects reported to acutely obstruct the intestinal tract include carpet or other cloth fibers, locust beans, and plastic.24,25 The location of the obstruction is usually in the proximal duodenum, a short distance from the pylorus where the lumen narrows, or in the midduodenum (Fig. 15-3). The second most common location is the ileocecocolonic junction. Acute compressions of the intestinal tract by neoplasia, postsurgical adhesions, tapeworm cysts, and hernias have also been
Fig. 15-3 Postmortem examination of a 4-year-old rabbit that died from acute gastric rupture. Note the typical appearance of acute gastrointestinal obstruction caused by a hair pellet (arrow). The obstruction is located a few inches distal to the pylorus in the duodenum and consists of compacted hair. The location of this obstruction, along with its size and composition, are characteristic. Note the hyperemic appearance of the duodenum at the level of the obstruction (arrowhead).
CHAPTER 15 Gastrointestinal Diseases reported.24,25 These all create a complete physical obstruction of the intestines and are not related to functional GI stasis. Rabbits cannot vomit and have a well-developed cardiac sphincter. When outflow from the stomach is obstructed, gastric fluid and swallowed saliva quickly accumulate. This fluid may undergo fermentation to produce large volumes of gas, resulting in a rise in intragastric pressure and a severely dilated stomach. As with gastric dilation in other species, the combination of sequestered fluid, compression of the aorta and vena cava causing hypovolemic shock, electrolyte imbalances, and acid-base disturbances can lead to death within hours of obstruction. Death can also be due to peritonitis resulting from ischemic necrosis at the site of the obstruction or rupture of the stomach.
HISTORY AND PHYSICAL EXAMINATION FINDINGS A sudden onset of anorexia and depression is the hallmark feature of acute GI obstruction. Some rabbits are found acutely moribund or discovered dead with no premonitory signs. Unlike rabbits with GI stasis, there is no history of a recent stressful or painful event, and dietary history does not play a role. Affected rabbits suddenly refuse all food and quickly stop producing feces. Most demonstrate signs of severe pain, such as reluctance to move, hunched posture, and bruxism. As metabolic derangements and shock develop, affected rabbits become severely depressed, listless, laterally recumbent, and minimally responsive to external stimuli. With stomach rupture, they may suddenly cry out and die. On physical examination, a large, fluidfilled or tympanic stomach is palpable in the cranial abdomen. Pain is elicited on palpation of the stomach. If examined early, affected animals are alert but quiet, tachypneic, and tachycardic. Later, they become hypothermic, bradycardic, and hypotensive.
DIAGNOSTIC TESTING Obtain radiographs to confirm the diagnosis. The radiographic appearance of a rabbit with acute GI obstruction differs sharply from that of a rabbit with GI stasis or that of a normal rabbit. The stomach will appear severely distended with gas, fluid, or both (see Fig. 15-2). Gas distention of intestinal loops proximal to the obstruction are noted, especially with obstructions occurring in the midduodenum or ileocecocolonic junction. Pneumoperitoneum indicates that the stomach has ruptured and carries a grave prognosis. Obtain blood for a complete blood count (CBC) and biochemical profile. Results may demonstrate dehydration and a variety of acid-base and electrolyte disorders.
INITIAL MEDICAL TREATMENT Begin treatment immediately, as this is a life-threatening disorder. Initial treatment goals are to decompress the stomach, treat shock, correct any fluid and electrolyte imbalances, and control pain. To control pain, administer buprenorphine (0.02-0.05 mg/kg SC, IV q6-12h). Sedation is necessary to decompress the stomach in most cases. Sedate with midazolam (0.5-1.0 mg/kg IM, IV) and/or mask with isoflurane or sevoflurane. A welllubricated 16- to 18-Fr red rubber catheter works well as an orogastric tube. It may be necessary to cut additional holes in the end of the tube to allow larger volumes of gas and fluid to pass. Measure the distance from the nose to the last rib (the distance to the stomach) and mark the tube. Lubricate the tube well and
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pass it gently per os into the stomach. Remove all gas and air from the stomach. Percutaneous trocharization of the stomach is not recommended as a method of decompression, since this will likely cause rupture of the stomach. While the stomach is being decompressed, place an intravenous catheter and begin treatment with a shock dose (90 mL/kg per hour) of isotonic crystalloid fluids. When dehydration is corrected and shock is controlled, reduce administration to a maintenance rate. Correct any electrolyte imbalances. In most instances, surgery is immediately necessary to remove the obstruction. However, in a small number of cases, the foreign object will pass following medical treatment. Monitor these rabbits closely to determine if the obstruction is passing. Palpate the abdomen frequently, monitor for signs of pain, and repeat radiographs. The gas pattern will change, and gas will be visible in the distal intestines if the rabbit is passing the obstruction. These rabbits will appear comfortable and begin eating, drinking, and defecating.
SURGICAL TREATMENT Immediate surgery is required to remove the obstruction in the majority of cases. When possible, decompress the stomach and stabilize shock prior to attempting surgery. Most obstructions are found in the proximal or midduodenum or in the ileocecocolonic junction. When possible, manipulate proximal duodenal foreign bodies into the stomach and perform a gastrotomy rather than an enterotomy. Gastrotomies are generally better tolerated, with a lower probability of postoperative complications such as stricture, leakage, or GI stasis. If no intestinal foreign body is found, explore the abdomen for evidence of neoplasia, abscesses, or adhesions as the cause of obstruction. Provide postoperative supportive care, including fluid therapy, pain management, and antibiotic therapy. Even with prompt removal of the foreign body, the prognosis is guarded. Many patients die during surgery or within 48 hours postoperatively from peritonitis, postoperative GI stasis, endotoxemia, or acute renal failure.
CECOTROPHY AND INTERMITTENT DIARRHEA Cecotrophs are nutrient-rich pellets resembling feces that contain the products of cecal fermentation. They are produced several times a day, usually in the morning and evening, and consumed reflexively directly from the rectum, a behavior termed cecotrophy. Cecotrophs are swallowed whole without chewing and are covered with a mucous coating that protects them from gastric degradation. This allows them to be delivered intact to the small intestines for digestion and absorption. Cecotrophs differ in appearance to rabbit fecal pellets. They are dark in color, soft and sticky in texture, have a characteristic odor, and are covered in a mucous layer. If intact, they appear as many soft fecal pellets stuck together, resembling a blackberry. If cecotrophs are not consumed, they often stick to the fur around the perineum or are found smeared on the fur and flooring. This is often confused with intermittent diarrhea and is a common presenting complaint. Rabbits that do not consume their cecotrophs are either physically unable to do so or do not eat them because they are abnormally formed. A common cause of inability to consume cecotrophs is obesity, where the rabbit cannot reach the anal region. Other causes include musculoskeletal disorders, dental disease, pain, and physical barriers such as Elizabethan collars. Changes in normal cecal motility, pH, or flora result in
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the production of abnormal cecotrophs. These cecotrophs may be soft, malformed, pasty, or odiferous and are therefore not consumed. Dietary deficiencies, as discussed above, are a common cause; however, other factors such as stress, concurrent disease, or antibiotic usage may also contribute. Diagnosis is based primarily on history. Affected rabbits produce normal fecal pellets throughout most of the day. Soft feces are found on the fur or smeared on flooring. If the rabbit cannot reach the anus, the feces are pasted to the perineum, and secondary dermatitis often results. Obesity or signs of neuromuscular, dental, or other painful disorders are present on physical examination. Correction of the underlying disorder will allow a return to normal cecotrophy. If the rabbit can reach the perineum, question the owner about the diet. Insufficient fiber in the form of hay and/or excessive carbohydrate intake is a common cause. In this case, simple correction of the diet will usually correct the problem.
CECOLITHS Altered motility of the cecum, rate of transit in the colon, or abnormal diet (e.g., the feeding of diets consisting of very small fiber length or feeding indigestible fiber, such as psyllium) can result in compaction and dehydration of cecal or colonic material and subsequent formation of “cecoliths” or abnormally hard lumps of cecal contents. Cecoliths are the most common cause of lower bowel obstruction in the rabbit, lodging at the distal end of the sacculated portion of the large intestine, the fusus coli. Rabbits that form cecoliths often have a chronic history of cecal or large intestinal disorders including the production of large, malformed feces, recurrent cecal impaction, abdominal pain, and anorexia. Because affected rabbits are unable to form normal cecotrophs, they are often underweight and lack normal muscle mass. Many of these rabbits are serologically positive for Encephalitozoon cuniculi, and speculation exists that this parasite may play some role in the disorder. One of the present authors (JRJ) has also postulated that some of these rabbits may have neurologic damage associated with trauma to mesenteric nerves or spinal cord. A presumptive diagnosis of cecolith formation can be made by palpation of doughy to firm material in the cecum or colon. Radiography or ultrasound examination can be used to confirm the presence of cecoliths. If the intestine is completely obstructed, gas will accumulate in the sacculated large intestine. Obstructed patients are in severe pain and may present moribund. Treatment of cecoliths requires rehydration of inspissated cecal and colonic contents. Administer fluid therapy via the intravenous or subcutaneous route and feed foods with a high water content (leafy vegetables, vegetable baby food). Additionally, feed an appropriate fiber source, such as grass hay, to stimulate normal cecal motility and function. Short-term administration of an intestinal promotility agent may be of benefit. In the long term, the addition of canned pumpkin (1 tbsp q12h) to the rabbit’s diet may prevent recurrence. Rabbits with complete large intestinal obstruction are critically ill, in pain, and require immediate treatment. Begin intravenous fluid therapy and administer buprenorphine (0.03-0.05 mg/kg SC, IV q6-12h). In some cases the obstructing cecolith can be softened and moved retrograde with a gentle enema. Take great care in administering enemas, as the colon may have become necrotic at the point of obstruction. If the obstruction cannot be removed via medical therapy, surgical removal will be required once the patient is stable.
ENTERITIS COMPLEX AND ENTEROTOXEMIA In clinical practice, enteritis complex—with signs ranging from soft stool and diarrhea to enterotoxemia, sepsis, and death—is one of the most common diseases of rabbits. Pathogenic bacteria and the factors that allow them to proliferate are the usual causes. These factors involve diet, antibiotics, stress, and genetic predisposition to gut dysfunction. Simple cases of enteritis, resulting in a soft or pasty stool as the only clinical sign, may be caused by a minor disruption of cecal flora, pH, or motility. Simple correction of the diet, the addition of fiber in the form of hay, and removal of stress will often correct the problem. Enterotoxemia in rabbits, which is characterized by more significant dysbiosis than in the case of enteritis, is caused by the iota-like toxin from Clostridium spiroforme.41 Newly weaned animals (3-6 weeks of age) are most often affected, and they have the greatest mortality rate. These rabbits may develop enterotoxemia from simple exposure to C. spiroforme. This is likely because these young rabbits have an undeveloped population of normal GI flora and a high gastric pH, which allows the proliferation of C. spiroforme. Adult rabbits are more resistant and generally require some dietary, environmental, or other stress for the dysbiotic state to be induced and growth of the bacteria occur. Rapid multiplication of C. spiroforme results in significant alteration of the rabbit’s normal cecal flora. Nursing does with enterotoxemia can develop a so-called milk enterotoxemia that is thought to be caused by Clostridium endotoxin produced in the does’ cecum and passed to the bunnies in their milk. In acute disease, rabbits become anorexic and markedly depressed. The diarrhea is brown and watery and soils the perineum and rear legs. It may contain blood or mucus. As the disease progresses, affected rabbits become hypothermic and, moribund and die after 24 to 48 hours. Postmortem findings in these rabbits include petechial and ecchymotic hemorrhages on the serosal surface of the cecum. The appendix and proximal colon may also be involved. Various amounts of gas throughout the intestinal tract, cecum, and colon result from ileus. Hemorrhages, pseudomembranes, or mucus may be present on the mucosa of the cecum and proximal colon.
MUCOID ENTERITIS Mucoid enteritis is one of the major causes of morbidity and mortality in young rabbits 7 to 14 weeks of age. It is characterized by anorexia, lethargy, weight loss, diarrhea, cecal impaction, and excessive production of mucus by the cecum. Its cause is unknown; however, studies have convincingly established the relation between bacterial dysbiosis and hyperacidity of the cecum and the symptoms of mucoid enteritis.31 Alterations in cecal pH resulting from changes in the production or absorption of volatile fatty acids or from vigorous fermentation of carbohydrates can destabilize the cecal microbial population and stimulate mucus production within the cecum and colon. Feeding a diet high in fiber and low in simple carbohydrates is preventative.
DYSBIOSIS CAUSED BY TREATMENT WITH ANTIBIOTICS Other factors involved in the development of enteritis include antibiotic administration and stress. Some antibiotics suppress normal flora, allowing pathogens to proliferate. Clindamycin,
CHAPTER 15 Gastrointestinal Diseases lincomycin, ampicillin, amoxicillin, amoxicillin-clavulanic acid, cephalosporins, many penicillins, and erythromycin can induce enteritis in rabbits. Epinephrine-mediated inhibition of gut motility is believed to be the cause of stress-induced enteritis.
TREATMENT OF ENTERITIS The treatment of rabbits with severe enteritis, enterotoxemia and mucoid enteritis consists of aggressive supportive care and efforts aimed at increasing cecal and colonic motility, discouraging the growth of pathogenic bacteria and the production of toxins, and supporting the growth of normal flora. Administration of cholestyramine (Questran, Bristol Laboratories, Princeton, NJ), an ion-exchange resin capable of binding bacterial toxins, at a dosage of 2 g in 20 mL water q24h by gavage, has been reported to prevent death in rabbits with clindamycininduced enterotoxemia.33 Antimicrobial drugs have limited value in the treatment of the disease and are used primarily as supportive therapy. C. spiroforme has been shown to be sensitive to metronidazole and penicillin G.7 The use of metronidazole (20 mg/kg PO, IV q12h) has been reported to reduce the number of deaths from enterotoxemia. Correction of dehydration and maintenance of normal hydration are of paramount importance, and administration of intravenous or intraosseous fluids is indicated. If the rabbit is anorectic, assist feed and provide supportive care as described for treatment of GI stasis, above.
PREVENTION OF ENTEROTOXEMIA To prevent enterotoxemia, maintain optimal husbandry and minimize stress. Feed a good-quality grass hay and limit or remove pellets from the diet. If a pelleted diet is fed, it should contain no less than 18% to 20% fiber and should be limited to less than 1⁄3 cup per 5 lb (2.3 kg) of body weight. Avoid sudden changes in the diet. Make hay available to weanling rabbits from 3 weeks of age; avoid early or forced weaning.
BACTERIAL ENTERITIS Enteropathogenic E. coli Diarrhea and mortality caused by infection with enteropathogenic E. coli (EPEC) are major causes of economic loss in the commercial rabbit industry. Disease outbreaks have not been reported in pet rabbits. Rather than producing enterotoxins or invading intestinal mucosa, EPEC exert their virulence by attaching to enteric epithelial cells and inducing effacement of microvilli; they are thus referred to as attaching and effacing E. coli.3,32,42 Diarrhea, caused by the resultant villus atrophy and malabsorption, varies in severity depending on the age of rabbit and specific serogroup involved. E. coli-related diarrhea in postweaning commercial rabbits may be caused by a variety of different serotypes that belong to the rabbit EPEC group.3,32 Morbidity and mortality rates vary; signs range from mild diarrhea and weight loss to death, and the mortality rate can be 50% or greater. Those animals that recover may have retarded growth. EPEC-related diarrhea in neonatal rabbits is most common between 1 and 14 days of age. The diarrhea is typically watery and stains the abdomen and perineum yellow. Morbidity and mortality rates within a litter approach 100%. Subsequent litters of the doe may have passive immunity. The disease process is limited to the cecum and colon. The cecal wall may be inflamed with longitudinal
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“paintbrush” hemorrhages. In severe cases, intussusception and rectal prolapse may be present. Presumptive diagnosis may be based on isolation of E. coli from stool or tissue samples from affected animals; however, nonpathogenic E. coli routinely proliferates in any rabbit with dysbiosis. Confirmation of the diagnosis requires histologic examination of tissues and observation of E. coli attachment to the intestinal cells. Serotyping of E. coli isolated from rabbits is not available to clinical veterinarians and remains a tool of research only. Treat individual rabbits with appropriate antibiotics, guided by the results of culture and sensitivity testing. Use trimethoprim-sulfamethoxazole as a combination antibiotic (30 mg/ kg PO q12h) or enrofloxacin (10 mg/kg PO q12h) until culture and sensitivity test results are obtained. Positive results may be obtained with early treatment.
Proliferative Enteritis, Proliferative Enteropathy, Proliferative Enterocolitis The obligate intracellular bacterium Lawsonia intracellularis, previously referred to as an intracellular Campylobacter-like organism, has been reported as a cause of enterocolitis in rabbits both alone and in association with an EPEC strain of E. coli distinct from the prototypic rabbit diarrhea E. coli (RDEC-1) strain.27,44 This intracellular bacterium is gram-negative, curved to spiralshaped, and found free in the apical cytoplasm of intestinal epithelial cells. The disease is most often characterized as an acute diarrheal disease of rabbits 2 to 4 months of age (weanlings). Proliferative enteritis (PE) or enteropathy is an enteric disease that develops in many animals. Much of the literature focuses on the disease in swine and hamsters. In addition, PE has been reported in rats and guinea pigs; ungulates other than swine, including white-tailed deer, sheep, and horses; carnivores, including arctic foxes, dogs, and ferrets; nonhuman primates; and birds (ratites). The disease is not an important problem in these other species.44 Histologic findings in these cases most often show a proliferative ileitis, with or without proliferative colitis, characterized by epithelial hyperplasia and mucosal inflammation. Similar disease in pigs and ferrets has been shown to be caused by a similar but distinctively different bacterium, Desulfovibrio desulfuricans.21,36 Treatment of L. intracellularis in rabbits is challenging. Antibiotics used to treat L. intracellularis in other species include those of the macrolide family (e.g., tylocin, erythromycin, and lincomycin). These antibiotics are not recommended for use in rabbits. Chloramphenicol is generally efficacious and is administered at 30 to 50 mg/kg PO, SC q12h for 7 to 14 days. Florfenicol (NuFlor, Schering-Plough Animal Health Corp., Union, NJ) may be useful as an alternative antimicrobial agent in rabbits, but its efficacy and potential side effects in this species are yet to be evaluated.
Tyzzer’s Disease Tyzzer’s disease is caused by Clostridium piliforme (formerly Bacillus piliformis), a motile gram-variable spore-forming obligate intracellular bacterium.16 The disease occurs in many rodents and other mammalian species in addition to rabbits. Stress (produced by overcrowding, unsanitary conditions, high temperatures, or breeding) may be an important component of this disease. Clinical signs of Tyzzer’s disease include watery diarrhea, depression, and death. Morbidity and mortality rates may be especially high in weanling rabbits. Older rabbits can develop a more chronic form of the disease that results in chronic weight loss. Postmortem examination of rabbits with Tyzzer’s disease
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may show characteristic foci of necrosis in the liver and degenerative lesions of the myocardium. More often, the intestinal wall is edematous, with areas of necrosis in the mucosa of the proximal colon. Treatment is palliative once clinical signs have been observed. The intracellular location of the bacteria may contribute to the difficulty in treating affected animals. If exposed animals are treated early (if they are isolated from affected animals, good hygiene is promoted; supportive care and a high-fiber diet are provided), they may not develop the disease. Once symptoms of the disease develop, treatment may be unsuccessful. Prevention of the disease depends on good husbandry. Bacterial spores are killed with a 0.3% sodium hypochlorite solution or with heating to 173°F (80°C) for 30 minutes.
Other Bacterial Enteritides Other causes of enteritis include Salmonella and Pseudomonas species. Salmonellosis is not common but can cause disease with high rates of both morbidity and mortality. The disease is well studied in rabbits, and the rabbit is used as a model of salmonellosis in humans.23 The species and serovar most often associated with salmonellosis in rabbits is Salmonella typhimurium; however, other species and serovars have been reported.5 Transmission of the disease is most often associated with contaminated food or water. Affected rabbits usually develop sepsis, which quickly leads to death; however, diarrhea may occur as well. Postmortem findings are consistent with septicemia and include vascular congestion of organs and diffusely distributed petechial hemorrhages. Lymph nodes and gut-associated lymphoid tissue may be edematous and contain similar foci of necrosis. One of the authors (JRJ) has seen an outbreak of lethal diarrhea in rabbits associated with Pseudomonas aeruginosa, which was isolated from the watering system. The morbidity rate associated with this outbreak was low to moderate, but the mortality rate in affected animals was high.
VIRAL DISEASES OF THE DIGESTIVE TRACT PAPILLOMATOSIS Rabbit oral papillomatosis is a benign disease caused by a papillomavirus. The disease has been reported only in colonies of laboratory rabbits, especially New Zealand white rabbits.37,45 Lesions consist of small white growths on the ventral surface of the tongue but only rarely elsewhere in the mouth. Early lesions are sessile, later becoming rugose or pedunculated and ultimately ulcerated. The lesions can exceed 4 to 5 mm at their greatest dimension but are typically smaller (1-3 mm). Lesions may persist as long as 145 days, but they usually disappear within weeks.
RABBIT ENTERIC CORONAVIRUS In 1980, a coronavirus was found to be the cause of diarrhea in laboratory rabbits.30 Further research has shown that this virus affects rabbits 3 to 10 weeks of age, but it has also been found in clinically normal adult rabbits. Clinical signs in naturally occurring outbreaks include lethargy, diarrhea, abdominal swelling, and death. Pleural effusion and cardiomyopathy in rabbits have also been associated with coronavirus-like particles.39 The disease is associated with high rates of morbidity and mortality; in one described outbreak, 40% to 60% of rabbits were affected.
Death occurred in almost 100% of these animals within 24 hours of the onset of clinical signs.13 Necropsy findings include fluid cecal contents, and histopathologic examination reveals atrophy of intestinal villi. Tentative diagnosis of this disease is based on clinical history, clinical signs, necropsy findings, and results of histopathologic analysis. The virus agglutinates red blood cells; evidence of hemagglutination activity in the feces therefore supports a tentative diagnosis. The diagnosis is confirmed by demonstration of the virus in feces or cecal contents.
ROTAVIRUS Infections in animals caused by rotavirus alone are often only mildly pathogenic; in rabbits, however, the virus is associated with very high morbidity but variable mortality rates. Although poorly studied in pet rabbits, antibodies to rotavirus as well as the virus itself have been found in the feces of rabbits from commercial rabbitries throughout the world. Severity of diarrhea associated with rotavirus infection varies widely and is likely influenced by synergy with various microorganisms associated with the infection. Severe anorexia, dehydration, and mucoid or greenish-yellow watery diarrhea have been reported. Rabbits between 30 and 80 days of age are most often affected. The mortality rate in young rabbits with naturally occurring infections may be as high as 80%. High morbiditiy and mortality rates found in naturally occurring infections have not been well reproduced in experimental studies. In one study, rotavirus caused soft or fluid feces in some rabbits, but in most animals diarrhea did not develop at all.8 Another study showed that a strain of rotavirus induced diarrhea, depression, anorexia, and death; however, results of the experiment were not reproducible.14,15 The clinical signs of naturally occurring infections involving rotavirus and other agents include marked congestion and distention of the intestines and cecum and petechial hemorrhages in the small intestine and colon. Histologic lesions include moderate to severe villous atrophy, with the most severe lesions being found in the ileum. Apical enterocytes on the tips of villi are swollen, rounded, and desquamated, and the tips may be denuded. The lamina propria is usually infiltrated with lymphocytes and occasionally with neutrophils. Diagnosis is established on the basis of the results of histopathologic examination of the intestine, isolation of the virus, or demonstration of antibodies. Clinical signs and gross pathologic findings alone are not diagnostic.13 The prevention and control of rotavirus infection is complicated by its highly infectious nature. Reduction of stress (by cessation of breeding, reduction of crowding, removal of socially dominant animals, and the addition of fiber to the diet) along with appropriate treatment of concurrent disease and improved hygiene should reduce mortality rates.
RABBIT HEMORRHAGIC DISEASE VIRUS Viruses of the Lagovirus genus within the family Caliciviridae affecting rabbits include rabbit hemorrhagic disease virus (RHDV), European brown hare syndrome virus (EBHSV), and the nonpathogenic rabbit calicivirus (RCV).22 EBHSV affects European hares of the Lepus genus. RHDV specifically afflicts Oryctolagus cuniculus, the predominant species of domestic rabbits worldwide. RHDV does not cause disease in wild cottontail rabbits, jackrabbits, or hares. Since its first emergence in China in 1984, RHDV has become endemic in Europe, Cuba, Australia, and New Zealand. Limited outbreaks have occurred in the
CHAPTER 15 Gastrointestinal Diseases Middle East, South America, Mexico, and the United States. All outbreaks were traceable to the importation of live rabbits or rabbit products from China. The virus has been eradicated from Mexico. Sporadic outbreaks have taken place in the United States, the last of which occurred in Indiana in 2005. The disease was eradicated following each outbreak and as of this writing is not endemic in North America.35 Clinical disease is seen in rabbits older than 2 months of age; younger rabbits are clinically unaffected.18,35 Virus is shed in urine, feces, and respiratory secretions. Transmission is via direct contact, contact with carcasses or fur from affected rabbits, or fomites such as water, feed, utensils, clothing, or cages. Flies and other insects may also serve as vectors, and the virus can be found in feces from predators that have eaten infected rabbits. RHDV is highly infectious and has traditionally been associated with high rates of both morbidity (40%-100%) and mortality (approaching 100%). Higher rates of morbidity and mortality are seen in naive populations. The number of rabbits affected during outbreaks peaks in 2 to 3 days and the disease course may last only 7 to 13 days. The incubation period is 1 to 3 days. RHDV replicates in the liver, resulting in severe hepatic necrosis and death from disseminated intravascular coagulation.6,34,35 In peracute disease, rabbits become febrile, lethargic, and collapse and die within 12 to 36 hours of infection; they may be found dead with no premonitory signs. With acute disease, affected rabbits are febrile and show signs of depression, lethargy, anorexia, constipation, or diarrhea. Some may show neurologic signs such as ataxia, opisthotonos, excitement, or seizures. At the end stage of the disease, tachypnea, cyanosis, and a blood-tinged foamy nasal discharge are often seen. In some rabbits, the course of the disease is slower, with the animals exhibiting jaundice, depression, anorexia, and fever, eventually dying within 1 to 2 weeks. In the subacute form, milder signs are seen and many of these rabbits live. Persistent or latent infections may occur in asymptomatic rabbits.18,35 Hematologic testing often shows a lymphopenia and a gradual decline in the number of thrombocytes. In most moribund rabbits, prothrombin and thrombin times are prolonged, and fibrin degradation products can be detected.43 The most consistent gross postmortem changes are hepatic necrosis, splenomegaly, and evidence of disseminated intravascular coagulation. Congestion and hemorrhage may be seen in most organs but is most pronounced in the lungs. The liver is pale, and periportal necrosis with a fine reticular pattern is observed; the spleen is dark and thickened, and catarrhal enteritis is often identified.13,43 A presumptive diagnosis may be made on the basis of data in the history, clinical signs, and pathologic findings. Definitive diagnosis requires identification of the virus using a variety of diagnostic tests, including electron microscopy, reverse transcription polymerase chain reaction (RT-PCR), Western blot, and enzyme-linked immunosorbent assay (ELISA). Immediately contact state or federal regulatory agencies to report this disease, and send diagnostic samples only to authorized laboratories under secure conditions. A tissue-derived, inactivated vaccine is used to protect domestic rabbits in countries where the virus is endemic.2,13 However, this vaccine appears to be less effective in some an antigenic variant strains, which may be responsible for the spread of disease throughout Europe despite active vaccination programs. The virus can be inactivated by 0.5% sodium hypochlorite or 1% formalin.
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PARASITIC DISORDERS OF THE GASTROINTESTINAL TRACT COCCIDIA Coccidia are the most common parasites of the rabbit GI tract and are frequently a cause of illness in young rabbits (less than 6 months old). Adult rabbits are rarely clinically ill, and the identification of oocytes on fecal examination does not equate to disease. Twelve species, all members of the genus Eimeria, are reported to infect rabbits.40 Only one species, Eimeria stiedae, which infects the liver, is found outside the intestinal tract. Very often, two or more species of coccidia are present in diseased rabbits; the precise role of the different species as pathogens is therefore not clearly defined.
Hepatic Coccidia E. stiedae, the coccidium responsible for hepatic coccidiosis, is ubiquitous in open rabbitries in which rabbits are not treated preventatively with coccidiostats. Infection results from ingestion of sporulated oocysts that undergo excystation in the duodenum. Liberated sporozoites penetrate the intestinal mucosa and move to bile epithelial cells, where they undergo schizogony. Merozoites invade contiguous epithelial cells and undergo gametogeny, giving rise to microgametes and macrogametes. After being fertilized by a microgamete, the macrogamete develops into an oocyst. Oocysts rupture from the epithelial cells and are passed in the bile and eventually, in the feces.28 Many infections are asymptomatic; however, the disease may be fatal, especially in young rabbits. Heavily infected rabbits show signs related to decreased hepatic function and bile duct obstruction. These rabbits become anorexic and debilitated; diarrhea or constipation may be noted in the terminal stages of the disease. The abdomen is occasionally enlarged and icterus is observed. Biochemical testing reveals increased alanine aminotransferase (ALT), aspartate aminotransferase (AST), bile acids, and total bilirubin. On radiographs, hepatomegaly and ascites may be present. On postmortem examination, the liver is enlarged and has yellowish-white, nodular, abscess-like lesions of varying size, some of which are within a fibrous capsule. Diagnosis is based on the identification of oocysts in a sample of bile, by histologic examination, or by fecal examination.
Intestinal Coccidia The most important species of intestinal coccidia are Eimeria perforans, Eimeria magna, Eimeria media, and Eimeria irresidua, with E. perforans being the most common. Infection is by ingestion of sporulated oocysts. Although rabbits are cecotrophic, it is generally accepted that cecotrophs eaten from the anus do not contain infectious oocysts. Clinical signs vary widely depending on the age of the rabbit, the organism involved, the parasitic burden, and the relative susceptibility of the animal (determined by factors such as age, stress, and diet). Subclinical infection is common in both young and adult rabbits. The finding of oocysts in clinically normal rabbits does not warrant treatment. Clinical signs are most often associated with poor husbandry or overcrowding and generally occur in rabbits under 6 months of age. Severely immunosuppressed older rabbits may also become symptomatic. Mild intermittent to severe diarrhea that may contain mucus or blood, weight loss, and dehydration may be observed. Animals with severe diarrhea may develop intussusception. Death is most often attributed to dehydration and secondary intestinal dysbiosis. Postmortem examination reveals lesions in the
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small or large intestine, depending on the agent involved. The epithelium of the intestine may be ulcerated. The presence of the organism (or organisms) in fecal samples or intestinal scrapings in symptomatic animals supports a presumptive diagnosis. Definitive diagnosis is based on histologic findings. Numerous agents have been used to prevent and treat intestinal and hepatic coccidiosis. Sulfa drugs appear to be the most effective. The addition of sulfadimethoxine to the diet in an amount to ensure intake of 75 mg/kg for 7 days or 0.02% sulfamerazine sodium to the drinking water is efficacious for treating groups of rabbits.40 Amprolium 9.6% in drinking water (0.5 mL per 500 mL) also is effective. Treat individual pet rabbits with sulfadimethoxine (15 mg/kg PO q12h for 10 days) or trimethoprim-sulfamethoxazole (30 mg/kg q12h PO for 10 days). The major role of antiparasitic agents is to limit multiplication until immunity develops. Instruct rabbitry, shelter, and pet store personnel in the practice of good husbandry to control outbreaks. Most healthy rabbits kept in clean, stress-free environments show no clinical signs after infection and develop immunity that may be lifelong.40
CRYPTOSPORIDIA Cryptosporidium parvum may cause a discrete and transitory diarrhea in young rabbits, peaking at 30 to 40 days, which may lead to growth retardation. Clinical signs include diarrhea lasting 3 to 5 days, decreased appetite, depression, lethargy, exhaustion, and dehydration. C. parvum infects the intestinal tract, especially the ileum and the jejunum. The organism apparently does not cause disease in adults. Atrophy of villi of the ileum in young rabbits has been observed histologically.38 Currently no effective treatment for cryptosporidiosis is recognized.
OTHER PROTOZOA Several nonpathogenic flagellates may be found in the feces of rabbits. They occur more commonly in animals with diarrhea. Giardia duodenalis occurs rarely in the anterior region of the small intestine of rabbits and is not considered pathogenic. Other nonpathogenic protozoa found in the cecum and colon include Monocercomonas cuniculi and Retortamonas cuniculi, which are flagellates from the cecum; large ciliated protozoa found in the cecum that are similar to those of the genus Isotricha in ruminants; and Entamoeba cuniculi, which is commonly found in the cecum and colon of rabbits.40
HELMINTHS Nematodes Passalurus ambiguus is the common pinworm of domestic rabbits, although P. nonanulatus also is reported.26 Occurrence is widespread in both wild and domestic rabbits; however, the presence of even relatively large numbers of pinworms is nonpathogenic. The adult parasite is found in the anterior portion of the cecum and colon. Adult worms are grossly visible in the lumen of the cecum and large intestine and when they are passed with fresh feces. The life cycle is direct, with infection through the ingestion of infected eggs during cecotrophy. Juvenile stages are found in the mucosa of the small intestine and cecum. Pinworms are commonly seen during routine surgical procedures such as ovariohysterectomy. Diagnosis is made by identification of adult worms or by demonstration of the parasite’s eggs in the feces.
Pinworm infections, even those with heavy worm burdens, are usually asymptomatic and do not require treatment. However, owners may notice the worms in rabbit feces and desire treatment. Advise owners that pinworms are species-specific, and not zoonotic. The benzimidazoles are effective in greatly reducing if not eliminating pinworms. Thiabendazole (50 mg/kg PO repeated in 10-14 days) and fenbendazole (10-20 mg/kg PO repeated in 14 days) are generally effective. Piperazine (200 mg/kg PO repeated in 14 days), to treat individual rabbits, or in drinking water (100 mg/100 mL of water for 1 day repeated in 10 days), to treat large numbers of animals, may also be effective. Other helminths are extremely rare in pet rabbits.
Cestodes and Trematodes Clinical disease as the result of intestinal cestode or trematode infection has not been reported in pet rabbits. The rabbit’s GI tract can host up to five species of cestodes: Cittotaenia variabilis, Mosgovoyia pectinata americana, Mosgovoyia perplexa, Monoecocestus americana, and Ctenotaenia ctenoides. Only C. variabilis has been found in domestic rabbits, whereas the other species are most often found in wild rabbits in North America and Europe.1,4 Adult parasites are found in the small intestine. The life cycles of some species are not well known; however, oribatid mites or ants are thought to act as intermediary hosts. Treatment of cestode parasites consists of the administration of a single dose of praziquantel (5-10 mg/kg PO).
NEOPLASIA Neoplasms of the GI tract include adenocarcinoma and leiomyosarcoma of the stomach, leiomyoma and leiomyosarcoma of the intestine, papilloma of the sacculus rotundus, papilloma of the rectal squamous columnar junction, and bile duct adenoma and carcinoma. Metastatic neoplasia, most commonly uterine adenocarcinoma, often involves the GI tract. Surgical resection is the treatment of choice for many of these tumors. If diagnosed early, intestinal masses can be resected with good success. Rectal papillomas (cauliflower-like, fungating masses arising from the anorectal junction) appear to be benign and are not related to the papillomas of skin or the oral cavity. Removal of these lesions is often curative. Bile duct adenoma and adenocarcinoma occasionally occur in pet rabbits. These tumors are often multiple and consist of interlocking cysts filled with thick, viscous, myxoid fluid. A variety of noxious stimuli, particularly infection with E. stiedae, may be causative factors. Antemortem diagnosis in some rabbits is based on the results of radiography and ultrasound. Surgical removal is often not practical. Metastatic disease is most often miliary and carries a grave prognosis.47
LIVER LOBE TORSION Liver lobe torsion has been recognized as a problem in rabbits for some time, with published reports dating back to 1958; there have also been several reports since that time.11,49,50 The caudate lobe is most often affected; however, torsion of the right lobe, the quadrate lobe, and the posterior lobule of the left hepatic lobe have also been reported.49,50 Both acute and chronic forms of liver lobe torsion have been observed. Signs of acute liver lobe torsion generally progress over a 12- to 72-hour period. Initially, affected rabbits are anorexic, weak, and depressed they demonstrate cranial abdominal discomfort.
CHAPTER 15 Gastrointestinal Diseases Mucous membranes may initially appear jaundiced. The stomach may contain food and a small amount of gas, suggesting gastric stasis syndrome. However, affected rabbits deteriorate over a relatively short period of time; if not treated, they become obtunded, have decreased body temperature, dark mucous membranes, and prolonged capillary refill time. Death may occur 12 to 72 hours from the onset of signs. The most commonly reported hematologic and biochemical abnormalities include anemia (packed cell volume 16%-17%) and mild to severe elevation of the liver enzymes ALT, AST, and gamma glutamyl transferase (GGT).19,46,49 However, these values may be normal in some cases. Hepatomegaly or an increase in density of the liver may be observed on radiographs, although this change may be difficult to appreciate. Ultrasonographic examination may be normal early in the course of disease or may demonstrate a heterogeneous appearance to the affected liver lobe, free fluid in the peritoneum, and occasionally pleural fluid. Treatment of acutely ill rabbits consists initially of supportive care, including intravenous fluids, analgesia, and thermal support. Surgical removal of the affected liver lobe in these rabbits may be lifesaving; however, the postsurgical survival rate is low. Signs of chronic liver lobe torsion are often nonspecific. These rabbits are often described as “poor doers” with a past history of recurrent GI stasis. Liver lobe torsion has also been diagnosed as an incidental finding at necropsy in previously asymptomatic rabbits.48,50 On physical examination, a firm, nonpainful mass is sometimes palpable in the cranial abdomen. Hematologic and biochemical abnormalities may include mild to moderate increases in liver enzyme activity, anemia, and azotemia. Diagnosis is based on ultrasonic examination of the affected lobe. Surgical lobectomy has been successfully performed in rabbits with this chronic presentation.46 Anecdotal reports of rabbits surviving with supportive treatment alone also exist.
AFLATOXICOSIS Aflatoxins are secondary metabolites of fungi, produced primarily by Aspergillus flavus and Aspergillus parasiticus. The LD50 for aflatoxins in rabbits is among the lowest for any species studied.10 In one outbreak of aflatoxicosis in angora rabbits, affected animals had anorexia, dullness, and weight loss followed by jaundice in terminal stages. Death occurred within 3 to 4 days of the appearance of clinical signs. On postmortem examination, livers were moderately to severely congested, icteric, and hard to cut. Gallbladders were distended and had inspissated bile. Liver sections showed degenerative changes of hepatic cells along with dilatation and engorgement of sinusoids. Bile ducts had mild to severe periportal fibrosis. Focal areas of pseudolobulation and regenerative foci were also predominant. The level of aflatoxin B1 in feed samples from various farms submitted at the time of the investigation varied from 90 to 540 mg/kg of feed. Withdrawal of feed and supplementary therapy resulted in gradual disappearance of signs and deaths.29
References 1. Andrews CL, Davidson WR. Endoparasites of selected populations of cottontail rabbits (Sylvilagus floridanus) in the southeastern United States. J Wildl Dis. 1980;16:395-401. 2. Arguello Villares JL. Viral haemorrhagic disease of rabbits: vaccination and immune response. Rev Sci Tech. 1991;10:459-480. 3. Blanco JE, Blanco M, Blanco J, et al. O serogroups, biotypes, and eae genes in Escherichia coli strains isolated from diarrheic and healthy rabbits. J Clin Microbiol. 1996;34:3101-3107.
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4. Boag B. The incidence of helminth parasites from the wild rabbit Oryctolagus cuniculus (L.) in eastern Scotland. J Helminthol. 1985;59:61-69. 5. Bolton AJ, Osborne MP, Stephen J. Comparative study of the invasiveness of Salmonella serotypes typhimurium, choleraesuis and dublin for Caco-2 cells, Hep-2 cells and rabbit ileal epithelia. J Med Microbiol. 2000;49:503-511. 6. Capucci L, Scicluna MT, Lavazza A. Diagnosis of viral haemorrhagic disease of rabbits and the European brown hare syndrome. Rev Sci Tech. 1991;10:347-370. 7. Carman RJ, Wilkins TD. In vitro susceptibility of rabbit strains of Clostridium spiroforme to antimicrobial agents. Vet Microbiol. 1991;28:391-397. 8. Castrucci G, Frigeri F, Ferrari M, et al. Comparative study of rotavirus strains of bovine and rabbit origin. Comp Immunol Microbiol Infect Dis. 1984;7:171-178. 9. Cheeke PR, Patton NM, Lukefahr SD, et al. Rabbit production. 6th ed. Danville, IL: Interstate Printers and Publishers; 1987. 10. Clark JD, Jain AV, Hatch RC. Experimentally induced chronic aflatoxicosis in rabbits. Am J Vet Res. 1980;41:1841-1845. 11. Cruise LJ, Brewer NR. Anatomy. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:53. 12. Davies RR, Davies JA. Rabbit gastrointestinal physiology. Vet Clin North Am Exot Anim Pract. 2003;6:139-153. 13. DiGiacomo RF, Mare CJ. Viral diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:171-204. 14. DiGiacomo RF, Thouless ME. Age-related antibodies to rotavirus in New Zealand rabbits. J Clin Microbiol. 1984;19:710-711. 15. DiGiacomo RF, Thouless ME. Epidemiology of naturally occurring rotavirus infections in rabbits. Lab Anim Sci. 1986;36:153-156. 16. Duncan AJ, Carman RJ, Olsen GJ, et al. Assignment of the agent of Tyzzer’s disease to Clostridium piliforme comb. nov. on the basis of 16S rRNA sequence analysis. Int J Syst Bacteriol. 1993;43:314-318. 17. Fekete S. Recent findings and future perspectives of digestive physiology in rabbits: a review. Acta Vet Hung. 1989;37:265-279. 18. Ferreira PG, Costa-e-Silva A, Monteiro E, et al. Transient decrease in blood heterophils and sustained liver damage caused by calicivirus infection of young rabbits that are naturally resistant to rabbit haemorrhagic disease. Res Vet Sci. 2004;76:83-94. 19. Fitzgerald AL, Fitzgerald SD. Hepatic lobe torsion in a New Zealand white rabbit. Canine Pract. 1992;17:16-19. 20. Forsythe SJ, Parker DS. Nitrogen metabolism by the microbial flora of the rabbit caecum. J Appl Bacteriol. 1985;58:363-369. 21. Fox JG, Dewhirst FE, Fraser GJ, et al. Intracellular Campylobacter-like organism from ferrets and hamsters with proliferative bowel disease is a Desulfovibrio sp. J Clin Microbiol. 1994;32:1229-1237. 22. Green KY, Ando T, Balayan MS, et al. Taxonomy of the caliciviruses. J Infect Dis. 2000;181(Suppl 2):S322-S330. 23. Hanes DE, Robl MG, Schneider CM, et al. New Zealand white rabbit as a nonsurgical experimental model for Salmonella enterica gastroenteritis. Infect Immun. 2001;69:6523-6526. 24. Harcourt-Brown FM. Gastric dilation and intestinal obstruction in 76 rabbits. Vet Rec. 2007;161:409-414. 25. Harcourt-Brown TR. Management of acute gastric dilation in rabbits. J Exot Pet Med. 2007;16:168-174. 26. Hofing GL, Kraus AL. Arthropod and helminth parasites. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:231-257. 27. Horiuchi N, Watarai M, Kobayashi Y, et al. Proliferative enteropathy involving Lawsonia intracellularis infection in rabbits (Oryctlagus cuniculus). J Vet Med Sci. 2008;70:389-392.
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28. Kraus AL, Weisbroth SH, Flatt RE, et al. Biology and disease of rabbits. In: Fox JG, Cohen BJ, Loew FM, eds. Laboratory animal medicine. Orlando: Academic Press; 1984:207-240. 29. Krishna L, Dawra RK, Vaid J, et al. An outbreak of aflatoxicosis in angora rabbits. Vet Hum Toxicol. 1991;33:159-161. 30. LaPierre J, Marsolais G, Pilon P, et al. Preliminary report on the observation of a coronavirus in the intestine of the laboratory rabbit. Can J Microbiol. 1980;26:1204-1208. 31. Lelkes L, Chang CL. Microbial dysbiosis in rabbit mucoid enteropathy. Lab Anim Sci. 1987;37:757-764. 32. Licois D. Enteropathogenic Escherichia coli from the rabbit. Ann Rech Vet. 1992;23:27-48. 33. Lipman NS, Weischedel AK, Connars MJ, et al. Utilization of cholestyramine resin as a preventive treatment for antibiotic (clindamycin)-induced enterotoxaemia in the rabbit. Lab Anim. 1992;26:1-8. 34. Marcato PS, Benazzi C, Vecchi G, et al. Clinical and pathological features of viral haemorrhagic disease of rabbits and the European brown hare syndrome. Rev Sci Tech. 1991;10:371-392. 35. McIntosh MT, Behan SC, Mohamed FM, et al. A pandemic strain of calicivirus threatens rabbit industries in the Americas. Virol J. 2007;4:96. 36. McOrist S, Gebhart CJ, Boid R, et al. Characterization of Lawsonia intracellularis gen. nov., sp. nov., the obligately intracellular bacterium of porcine proliferative enteropathy. Int J Syst Bacteriol. 1995;45:820-825. 37. Mews AR, Ritchie JS, Romero-Mercado CH, et al. Detection of oral papillomatosis in a British rabbit colony. Lab Anim. 1972;6:141-145. 38. Mosier DA, Cimon KY, Kuhls TL, et al. Experimental cryptosporidiosis in adult and neonatal rabbits. Vet Parasitol. 1997;69:163-169. 39. Osterhaus AD, Teppema JS, Van Steenis G. Coronavirus-like particles in laboratory rabbits with different syndromes in The Netherlands. Lab Anim Sci. 1982;32:663-665.
40. Pakes SP, Gerrity LW. Protozoal diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:205-229. 41. Peeters JE, Geeroms R, Carman RJ, et al. Significance of Clostridium spiroforme in the enteritis-complex of commercial rabbits. Vet Micro. 1986;12:25-31. 42. Peeters JE, Geeroms R, Orskov F. Biotype, serotype, and pathogenicity of attaching and effacing enteropathogenic Escherichia coli strains isolated from diarrheic commercial rabbits. Infect Immun. 1988;56:1442-1448. 43. Plassiart G, Guelfi JF, Ganiere JP, et al. Hematological parameters and visceral lesions relationships in rabbit viral hemorrhagic disease. Zentralbl Veterinarmed B. 1992;39:443-453. 44. Schauer DB, McCathey SN, Daft BM, et al. Proliferative enterocolitis associated with dual infection with enteropathogenic Escherichia coli and Lawsonia intracellularis in rabbits. J Clin Microbiol. 1998;36:1700-1703. 45. Sundberg JP, Junge RE, el Shazly MO. Oral papillomatosis in New Zealand white rabbits. Am J Vet Res. 1985;46:664-668. 46. Taylor HR, Staff CD. Clinical techniques: successful management of liver lobe torsion in a domestic rabbit (Oryctolagus cuniculus) by surgical lobectomy. J Exot Pet Med. 2007;16:175-178. 47. Weisbroth SH. Neoplastic diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:259-292. 48. Weisbroth SH. Torsion of the caudate lobe of the liver in the domestic rabbit (Oryctolagus cuniculus). Vet Pathol. 1975;12:13-15. 49. Wenger S, Barett EL, Pearson GR, et al. Liver lobe torsion in three adult rabbits. J Sm Anim Pract. 2009;50:310-305. 50. Wilson RB, Holscher MA, Sly DL. Liver lobe torsion in a rabbit. Lab Anim Sci. 1987;37:506-507.
CHAPTER
16
Respiratory Disease and Pasteurellosis
Angela M. Lennox, DVM, Diplomate ABVP (Avian)
Anatomy of the Respiratory Tract Diseases of the Upper Respiratory Tract Infectious Noninfectious Diseases of the Lower Respiratory Tract Infectious Neoplastic Diseases Producing Secondary Respiratory Symptoms Diagnosis and Differentiation Physical Examination Laboratory Analysis Serology and Molecular Diagnostic Testing Diagnostic Imaging Treatment of Respiratory Disease Prevention and Control of Respiratory Disease
Respiratory disease is common in pet rabbits and, as in other species, is caused by a number of primary and secondary etiologies. Although the anatomy of the respiratory tract is discussed in more detail in Chapter 12, it should be kept in mind that rabbits are obligate nasal breathers because of the position of the elongated epiglottis engaged over the caudal margin of the soft palate. For this reason, mouth breathing is an indication of a severe abnormality. In general, disease of the upper respiratory tract is more severe and stressful in rabbits than in other species. Respiratory disease can be classified as affecting the upper or lower respiratory tract or both.
ANATOMY OF THE RESPIRATORY TRACT The nasal cavities contain the conchae; these structures create the ventral, middle, dorsal, and common meatus. The ventral meatus continues to the rhinopharynx. Two blind cavities inside the conchae are commonly termed sinuses and are often Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
reported as the conchal and maxillary sinuses. As these cavities are blind, the more correct term may be recesses. There is a single opening of each sinus into the nasal cavity. Each lung is divided into cranial, middle, and caudal lobes, with the right lung also possessing an accessory lobe. The volume of the thoracic cavity of the rabbit is small, especially in comparison with the volume of the abdominal cavity.17
DISEASES OF THE UPPER RESPIRATORY TRACT INFECTIOUS Numerous infectious diseases are associated with disease of the upper respiratory tract; they are due mainly to bacterial agents, but in rare cases, viral, fungal, or parasitic pathogens may be involved as well. The rabbit is a common research model for rhinitis and sinusitis. Sinusitis is established after mechanical blockage of the ostium connecting the nasal and sinus cavities, resulting in impaired mucociliary function and clearance of organisms from the sinus cavity.2 Ostial dysfunction appears to be an important requirement for the establishment of infection in laboratory models, especially for organisms not considered primary pathogens, and may have significance for naturally infected animals as well. Otitis media is associated with respiratory disease in rabbits, as infection can spread via the eustachian tube to the tympanic bulla and middle and possibly inner ear.11,17
Bacterial Pathogens While Pasteurella multocida is often implicated as a cause of respiratory disease, other pathogens must be considered. A recent epidemiologic study of 121 rabbits with symptoms of upper respiratory tract disease (nasal discharge and sneezing) indicated that the most common bacterial isolates (deep nasal samples) were P. multocida (54.8%), Bordetella bronchiseptica (52.2%), Pseudomonas species (27.9%), and Staphylococcus species (17.4%). Mixed infections were also seen.30 Pasteurellosis is associated with many disease processes in rabbits including rhinitis, sinusitis, conjunctivitis, nasolacrimal 205
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duct infection, otitis, tracheitis, pneumonia, and abscesses (dental, skin, other). While not identified as a pathogen in wild rabbit populations, Pasteurella is a significant pathogen in laboratory animals, leading to the development of “Pasteurella-free” animals for research use. Because of its ubiquitous nature, many pet rabbits are already infected with Pasteurella multocida. Many exhibit symptoms in response to stress, such as stress of weaning, transport, and purchase from the pet store as well as poor sanitation and ventilation and concurrent illnesses.11 Strains of Pasteurella vary in pathogenicity,27 with some being more likely to spread via the hematogenous route and causing acute septicemia, generalized disease, and death. P. multocida gains entry to the host primarily through the nares or wounds via aerosolization or direct contact with infected rabbits or fomites. Does with venereal infections may pass the organism to their young, who may remain apparently uninfected for several weeks until signs of infection develop. Infected rabbits can resist infection or become subclinical carriers, thus complicating determination of an incubation period. In experimental studies, rhinitis occurred 1 to 2 weeks after intranasal inoculation.24 After infection, the organism spreads to the sinuses, nasolacrimal duct, eustachian tube and middle ear, trachea, and lungs. Factors facilitating disease expression include exposure to ammonia through improperly cleaned enclosures, administration of corticosteroids, and the psychologic stress or stress of concurrent disease.10 Recent studies have indicated promise for the development of vaccines as an aid to management of this disease.33 B. bronchiseptica is a common inhabitant of the rabbit respiratory tract, and disease prevalence may increase with age.11 Experimental inoculation of organisms may or may not produce disease. This organism is pathogenic in guinea pigs, dogs, cats, and pigs. B. bronchiseptica is suspected to be a copathogen or predisposer to infection with P. multocida. However, Deeb has reported severe infection in the absence of P. multocida in a colony of inbred laboratory rabbits, suggesting that more pathogenic strains may exist (BJ Deeb and RF DiGiacomo, unpublished data). Pseudomonas species infections in rabbits are common. Rabbits are used as a laboratory model for human Pseudomonas infections in multiple body systems, including the cardiovascular system.21 Staphylococcus species is commonly isolated from the nares of rabbits and is thought to be a secondary invader. Pathogenicity depends on host susceptibility and bacterial virulence. Disseminated staphylococcosis has been linked with fibrinous pneumonia or abscesses in the lung or heart.10 While not a primary respiratory pathogen, Treponema cuniculi (rabbit syphilis) can produce crusting of the mucocutaneous borders of the nose, lips, and eyelids, in some cases resembling an upper respiratory infection. Marked crusting of the nares can produce mild to moderate respiratory distress. A number of other pathogens can produce upper respiratory disease in rabbits. Pasteurella species other than P. multocida can be cultured from rabbits but may be nonpathogenic. However, pure cultures associated with clinical disease should be treated as true pathogens. Other potential pathogens include Moraxella species, Yersinia pestis, and Escherichia coli. Mycoplasma pulmonis was isolated by Deeb in the rhinopharynx of rabbits with evidence of upper respiratory disease. Infected rabbits were housed in close proximity to rats, which may have been the source of the infection.10 S. Kelleher (personal communication) reported a case of nasal granuloma produced by Mycobacterium species.
Viral Pathogens Viral pathogens other than myxoma virus producing primary upper respiratory disease are uncommon. Myxoma virus can produce nasal and ocular disease as well as dyspnea as a feature of the disease; it has also been associated with acute hemorrhagic pneumonia.23,25 An outbreak of a herpesvirus was reported in a commercial rabbitry in Alaska in 2008. Respiratory signs and symptoms were a prominent feature.22
Fungal Pathogens Fungal granulomas of the sinuses have been described in pet rabbits and, as in other species, appear to require primary injury. Techniques used in rabbits to produce experimental models of fungal sinusitis include mucosal injury and blockage of the ostium.13 Simple introduction of pathogenic fungi into the nasal cavity usually does not produce disease.
NONINFECTIOUS Trauma Traumatic injury to the upper respiratory tract includes blunt force trauma, predator injury, and damage to the glottis and trachea after endotracheal intubation. There are numerous reports of inflammation of the glottis and stenosis of the trachea in the laboratory literature.15,29 In one report of three cases of tracheal stenosis secondary to intubation, evidence of respiratory disease appeared within 17 to 21 days postintubation. Risk factors could not be identified.29 The author has regularly performed blind endotracheal intubation in rabbits and taught the technique to students and support staff for over 10 years. The most severe complications have been rare cases of mild respiratory stridor that did not persist beyond 24 hours postprocedure. In normal position, the epiglottis is above the soft palate in the rabbit and other obligate nasal-breathing species, preventing both mouth breathing and access to the trachea via the oral cavity. Hyperextension of the head causes the epiglottis to disengage and reposition below the soft palate, allowing access to the trachea (Fig. 16-1). Other keys to successful intubation appear to be correct tube size (2-3 mm) and gentle technique without the use of pressure during placement. All mammalian species are susceptible to airway irritation due to chemical exposure, and mucosal damage may predispose to infection. Numerous studies have demonstrated upper airway and lung damage secondary to irritants such as tobacco and dried dung smoke.14 Other potential sources of respiratory irritants include household chemicals and ammonia from poorly cleaned enclosures. Foreign materials in the nares, pharynx, or trachea have been reported. In one rabbit with chronic sneezing, its clinical signs ceased following removal of hay from the nares.32
Dental Disease Certain expressions of dental disease can produce symptoms of respiratory disease, including epiphora and nasal discharge.6 The nasolacrimal duct in the rabbit courses close to the apex of first maxillary incisor tooth and the first maxillary cheek tooth (Fig. 16-2). Therefore abnormalities or infections of the apices and reserve crowns of these teeth can affect the natural course of the duct, resulting in epiphora. More severe disease can produce infection or abscess of the duct and subsequent purulent nasal discharge. Elongation, deformation, and infection of the roots
CHAPTER 16 Respiratory Disease and Pasteurellosis
A
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B Fig. 16-1 Endoscopic view of the epiglottis and glottis of a rabbit. A, The epiglottis is in the normal position engaged above the soft palate, a feature common in all obligate nasal-breathing species. B, The epiglottis is disengaged below the soft palate, allowing introduction of the endotracheal tube. (Used by permission from Vittorio Capello, DVM.) trachea was identified upon tracheoscopy; incomplete rings were confirmed at histopathology.9
DISEASES OF THE LOWER RESPIRATORY TRACT INFECTIOUS
Fig. 16-2 Radiograph of the head of a rabbit (lateral projection) with chronic bilateral epiphora and nasal discharge. Note the apical deformity of one of the apices of maxillary incisor teeth and abnormal curvature and dorsal margin of the reserve crown. These lesions are associated with impingement of the nasolacrimal duct. Radiographic abnormalities consistent with acquired dental disease of the cheek teeth are present as well, and the ventral nasal turbinates are denser than the dorsal nasal turbinates.
of maxillary cheek teeth (in particular the first two cheek teeth) can affect the nasal cavity, resulting in chronic nasal discharge and sinusitis.6
Neoplastic Carcinoma of the nasal turbinates has been reported anecdotally.
Miscellaneous Conditions An apparent congenital abnormality characterized by incomplete tracheal rings has been diagnosed in a rabbit with dyspnea and an appreciable honking noise upon inspiration. Collapsing
Infectious agents, as described previously, also produce lower respiratory disease, with classic clinical signs and presentation of pneumonia. P. multocida and sometimes other organisms can also produce pleuritis and pericarditis. A study of 66 rabbits with pulmonary lesions isolated the following pathogenic bacteria in this order of frequency: Pasteurella species including multocida, E. coli, B. bronchiseptica, and Pseudomonas aeruginosa.25 Chlamydia species have been isolated from the lungs of domestic rabbits with pneumonia. A mild interstitial pneumonia occurred when the agent was inoculated into the trachea of laboratory rabbits.10 Pneumocystis oryctolagi has been isolated from the lungs of newly weaned rabbits.12 Myxomavirus has been associated with acute hemorrhagic pneumonia.25
NEOPLASTIC A number of neoplasms produce metastasis to the lungs or, in the case of thymoma or thymic lymphoma, indirectly affect the lower respiratory tract. Neoplasms producing metastasis to the lungs include uterine adenocarcinoma, osteosarcoma, lymphoma, and mammary carcinoma.36 Primary carcinoma of the lungs of a 7-year-old rabbit has also been reported.19 Thymoma is a primary tumor of lymphoid or epithelial origin and is increasingly reported in the literature.26 Common clinical features include decreased activity, increased respiratory rate and effort, and bilateral exophthalmos, which can be intermittent and often increases in severity over time. Exophthalmos is thought to occur secondary to vascular compression
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and reduction of vascular return to the heart caused by the growing mass.26
DISEASES PRODUCING SECONDARY RESPIRATORY SYMPTOMS Cardiovascular disease can produce signs that suggest respiratory disease, including increased respiratory rate and effort. As the life span of pet rabbits increases, it can be expected that the incidence of cardiac disease will increase as well.17 Cardiomyopathy, endocardiosis, and congestive heart failure have been reported in rabbits.28 Diaphragmatic herniation with entrapment of the stomach, parts of the intestinal tract, and kidney has been identified, as well as pneumothorax secondary to trauma.7
DIAGNOSIS AND DIFFERENTIATION PHYSICAL EXAMINATION Symptoms of respiratory disease can be similar to those noted in other traditional pet species; they include nasal and/or ocular discharge and increased respiratory effort and rate, often worsening with exertion. Ocular and nasal discharge frequently produces wet or matted fur beneath the eyes and nares. Rabbits with ocular or nasal discharge often attempt to remove the material with the forefeet, resulting in an accumulation of debris in the fur of the medial aspect of the feet.17 In addition, rabbits with tracheitis will often cough when the trachea is gently palpated. Because rabbits are obligate nasal breathers, diseases producing nasal discharge or other occlusion of the nares can lead to severe respiratory distress.17 Other nonspecific symptoms that should arouse suspicion of respiratory disease include weight loss, decreased appetite, lethargy, and exercise intolerance. Auscultation in rabbits is extremely useful. Airway and lung sounds in the normal rabbit are similar to those in other small traditional pet species. However, small thoracic cavity and lungs can make identification of lung sounds challenging. Diseases affecting upper airways often result in increased airway noise or wheeze, which is often referred to the thorax. Pulmonary disease produces predictable changes in lung sounds, including decreased or increased sounds, which may be bilateral or unilateral. It should be noted that some rabbits under stress are particularly vocal and will produce a characteristic honking noise that should not be confused with a sign of respiratory disease. Breathing patterns are also important. As in other species, rabbits with pleural disease (pneumothorax, hydrothorax) may demonstrate an asynchronous breathing pattern, with the thorax and abdomen appearing to move opposite to each other. The author has noted a case of hydrothorax in a rabbit where removal of fluid resulted in immediate restoration of synchronous breathing movements.
LABORATORY ANALYSIS Hematology is often unremarkable in cases of respiratory disease. Laboratory animal references state that rabbits tend not to develop leukocytosis in the face of infectious disease25—a common observation reported by rabbit practitioners. Biochemical analysis does not directly support respiratory disease. The author has noted that many infectious/inflammatory disease
Fig. 16-3 Dacryocystorhinogram in a rabbit. Contrast medium depicts the abnormally dilated nasolacrimal duct (arrowhead) and the deformed apex of one of the maxillary incisors (white arrow). Despite the impingement, some of the contrast medium flows beyond the compression, showing partial patency (black arrow). These early and subtle cases of acquired dental disease (ADD) of the maxillary incisors benefit from extraction to prevent more serious sequelae (see Chapter 32).
processes in rabbits, including some respiratory diseases, produce elevations in globulins. Bacterial culture and sensitivity testing is useful for identification of specific bacterial pathogens. Common sources include deep nasal culture and samples from nasal or tracheal washes. It should be kept in mind that culture of the external nares is more likely to reveal environmental pathogens. Deep nasal cultures offer a much more reliable reflection of pathogens of the nasal cavity. Deep nasal cultures can be collected with very small culturettes introduced a distance of 2 to 3 cm into the ventral and common nasal meatus. Culturettes should be introduced into both sides, as disease may be unilateral. The author has occasionally encountered purulent material at the tip of the endotracheal tube after withdrawal, suggesting tracheitis. This material can be prepared for cytology and then submitted for culture and sensitivity. Tracheal/lung washes can be collected by introducing a sterile red rubber catheter through the lumen of a sterile endotracheal tube in place in the anesthetized patient. The author prefers 2.0- to 3.0-mm endotracheal tubes in rabbits; therefore catheter size is chosen based on the endotracheal tube’s interior diameter. Depending on patient size, 1-2 ml sterile preservative free saline is introduced, and then aspirated. Because the amount of fluid collected back into the syringe may be scant to none, it is helpful to withdraw the catheter, fill the syringe with air, and then flush catheter contents into a sterile tube for submission. Flushing of the nasolacrimal duct can be both diagnostic and therapeutic and is described under “Treatment of Respiratory Disease,” below.18 Patency is demonstrated by the appearance of fluid at the nasal puncta. Alternatively, corneal stain may be observed at the nasal puncta. Failure to flush may indicate permanent occlusion, rupture, or impingement. A dacrocystogram is useful in helping identify the duct radiographically (Fig. 16-3).18 Fluid collected via thoracocentesis or ultrasound-guided aspiration of thoracic masses can also be submitted for culture and
CHAPTER 16 Respiratory Disease and Pasteurellosis
B
A
209
C
Fig. 16-4 A, Normal radiograph of the thorax of a rabbit, lateral view. B, Radiograph of the thorax of a rabbit demonstrating a slightly increased respiratory rate at initial presentation and (C) severe respiratory distress 2 months later. Note multiple mineralized densities throughout the caudal lung fields. Metastatic adenocarcinoma (origin: uterus) was confirmed at necropsy. (Used by permission from Vittorio Capello, DVM.) sensitivity or cytology. In the case of thoracic abscesses, the submission of purulent material for culture is often unrewarding. Certain culture techniques may enhance isolation of P. multocida; therefore the reference laboratory should be consulted for instructions on sample collection and handling.
SEROLOGY AND MOLECULAR DIAGNOSTIC TESTING Many laboratories servicing the lab animal community offer serologic testing that can be used by the practitioner, including P. multocida and T. cuniculi. Each laboratory should be consulted regarding type of test available (enzyme-linked immunosorbent assay [ELISA] vs. immunofluorescence assay [IFA]) and sample submission requirements. These laboratories also provide insight into the interpretation of results. The polymerase chain reaction (PCR) can be a valuable diagnostic tool for rabbits with respiratory disease. At the time of publication, available testing included PCR for B. bronchiseptica and Pasteurella species from tracheal washes and nasal swabs.
DIAGNOSTIC IMAGING Radiography is extremely useful for the diagnosis of respiratory disease in rabbits but has a number of limitations, in particular for detection of lesions of the upper respiratory tract. As in all radiographic patients, excellent positioning and technique are essential to obtain images of diagnostic quality (see Chapter 35).7 This is particularly true for radiographs of the skull, where rotation and asymmetry of lateral and dorsoventral views greatly complicate interpretation. Lesions of the maxillary sinuses are often subtle; however, marked and aggressive changes (abscesses, masses, osteolysis) can be apparent radiographically. Radiographs of the skull are extremely useful as an aid to the diagnosis of dental disease and abnormalities of the tympanic bulla, which in some cases may produce symptoms of respiratory disease.7 The thoracic cavity is small in comparison to the abdomen in rabbits. Structures apparent in the thorax of the rabbit are the relatively large heart, trachea and bronchi, a small portion of the cranial lung lobes, and a larger portion of the caudal lung lobes. Changes in normal radiographic patterns are similar to those seen in other small domestic mammals and can include discrete nodules and interstitial and bronchial patterns. Other changes
can suggest pneumothorax, hydrothorax, mediastinal masses, and diaphragmatic hernia (Fig. 16-4).7 Radiographic quality is of critical importance. Radiographs are collected with sedation and careful manual restraint or full anesthesia, depending on the patient’s condition. Right and left lateral projections are collected with hyperextension of the forelimbs in order to reduce superimposition of the brachial muscles over the cranial portion of the thoracic cavity, which might be confused for a mass or effusion. The ventrodorsal projection is important as well but may be stressful for the conscious patient.7 Computed tomography (CT) is gaining acceptance as a useful and increasingly available tool for use in larger exotic companion mammals. The author and others have described the use of CT as an aid to the diagnosis of dental disease and diseases of the maxillary sinus and tympanic bulla in rabbits (Fig. 16-5).5 Other uses include diagnosis and characterization of pulmonary and mediastinal masses (Fig. 16-6). Magnetic resonance imaging (MRI) can also be a useful diagnostic tool in rabbits with respiratory disease. Rabbits with severe respiratory disease may not be good candidates for radiography or other diagnostic tests. Anesthesia may pose substantial risk, and manual restraint can produce significant stress. The author has had success with sedation (midazolam at 0.25 mg/kg IM with butorphanol at 0.2 mg/ kg IM) and careful manual restraint. Sedation reduces anxiety and stress, and many patients relax and demonstrate reduced dyspnea. Endoscopy also is gaining importance as a diagnostic tool for exotic companion mammals. In the rabbit with chronic upper respiratory disease, rhinoscopy is a diagnostic option that may help to identify granulomatous disease unlikely to respond to simple antibiotic therapy.20 Rhinoscopy may also identify nasal foreign bodies. The author prefers a 1.9-mm rigid endoscope with diagnostic sheath for rhinoscopy of rabbits weighing more than 2 kg. Thoracoscopy has also been described as an aid to diagnose pulmonary disease in the rabbit. See Chapter 34 for more information.
TREATMENT OF RESPIRATORY DISEASE Rabbits with respiratory disease may present in critical condition. Refer to Chapter 38 for more information on how to approach the rabbit in respiratory arrest or distress. In brief,
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210
Dorsal nasal concha
Middle nasal concha
Nasal bone Ventral nasal concha
Third endoturbinate
Maxillary cheek teeth Fourth endoturbinate
A
B
Dorsal nasal concha
C
G
E
I
Dorsal nasal concha
Nasal septum
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Fig. 16-5 Anatomy of the nasal cavities, turbinates (conchae), and meatus (sinuses, recess); computed tomography (CT) of a rabbit affected by bilateral rhinitis. A, Median section through the head, lateral view. (Modified from: Popesko P et al: A colour atlas of anatomy of small laboratory animals. Vol. I: Rabbit, guinea pig. London: Wolfe Publishing, 1992; with permission.) The scout view demonstrating the CT scanning planes has been adapted from a radiograph of the rabbit shown in G to J (B). Computed tomography (axial view, bone window) of the normal nasal cavities, and corresponding three-dimensional volume renderings (C-F). CT of a rabbit with bilateral rhinitis (axial view, soft tissue window) and corresponding three-dimensional volume renderings (G-J). Note the abnormal turbinates and empyema of the maxillary recesses. (C-F, H, and J used by permission from Vittorio Capello, DVM.)
rabbits in respiratory arrest require intubation and ventilation or positive pressure ventilation with a tight-fitting mask and initiation of cardiopulmonary-cerebral resuscitation (CPCR) (see Chapter 38). Rabbits in respiratory distress benefit from the administration of oxygen in a quiet, semidark environment. As mentioned previously, mild sedation often benefits rabbits in respiratory distress by eliminating anxiety. Rabbits with evidence of pneumothorax or hydro/hemothorax (dysynchronous breathing pattern) require a chest tap, which, depending on the patient’s condition, should be
performed prior to radiography or other diagnostic tests or treatments likely to produce respiratory compromise. See Chapter 38 for more information on distinguishing pulmonary from pleural disease. The chest tap is performed after preoxygenation and sedation using a 22- to 25-g butterfly needle with syringe and two-way stopcock. The area is clipped and prepared, after which a topical anesthetic is applied. Local infiltration of lidocaine into the muscle layers is also extremely useful. The technique is identical to that used in other traditional pet species, and the patient is maintained in a normal standing
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Fig. 16-6 Thoracic mass in a rabbit. Radiographs of the thorax, lateral (A) and ventrodorsal views (B). Computed tomography, axial view (C), and three-dimensional volume rendering (D). Ultrasound-guided fine-needle aspirate revealed purulent material, suggesting the presence of an abscess. Arrow, Mass; L, lungs; M, mass; H, heart.
position. The thoracic cavity of the rabbit is relatively small, and the heart occupies most of the entire length (second to fourth intercostal space). Therefore needle advancement is kept to a minimum. In most cases, the author prefers to place the needle at the fifth to sixth intercostal space. For relief of thoracic fluid in the cranial thorax associated with thymoma or other mediastinal masses, needle placement is at the second to third intercostal space. Treatment of bacterial disease is enhanced with accurate diagnosis, including pathogen identification and localization of disease. In the case of bacterial pathogens, antimicrobial selection based on culture and sensitivity is ideal. In the study of 121 rabbits with evidence of nasal discharge and sneezing mentioned above, marbofloxacin was demonstrated as most effective against most identified bacterial strains with the exception of B. bronchiseptica, where other antimicrobials were slightly more effective.30 Antibiotic selection, however, should also be made with species-specific contraindications in mind. Enteric
dysbiosis and potentially fatal enterocolitis or enterotoxemia are well-documented potential results of administration of inappropriate antibiotics such as oral penicillin, erythromycin, and similar drugs (see Chapter 41). Treponematosis (rabbit syphilis) responds well to three weekly injections of penicillin (see Chapter 41). Treatment and elimination of P. multocida has been the focus of much attention because of the economic impact of the disease on commercial and laboratory rabbit facilities. Treatment of acute cases is often associated with good outcomes, but numerous studies have demonstrated poorer long-term prognosis for treatment of chronic infection, with high rates of recurrence after discontinuation of drug therapy.10 Various treatment options are reported in the literature. Infection was eliminated in 7 of 8 rabbits treated with enrofloxacin (5 mg/kg SC q12h) for 14 days.3 Enrofloxacin given in the drinking water (50-100 mg/L) before and continuing for 48 hours after inoculation with a virulent strain of P. multocida protected rabbits against
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Fig. 16-7 A, Lacrimal punctum of a normal rabbit. It is located cranially in the ventral eyelid (arrow). B, In cases of dacryocystitis, the lacrimal punctum may be dilated (arrow) or may demonstrate retrograde reflux of mucopurulent material (C). Flushing of the nasolacrimal duct and dacryocystorhinography can be performed using a 22- or 24-gauge catheter or a 22-gauge blunt needle (D). After flushing, mucopurulent discharge can often be observed emerging from the distal opening of the nasolacrimal duct (E). In selected cases, flushing of the nasolacrimal duct can be also performed in a retrograde fashion (F). (A and C-F used by permission from Vittorio Capello, DVM.)
bacteremia provided that daily intake of the drug was greater than 5 mg/kg.10 Young from enrofloxacin-treated does were free of P. multocida infection, although the infection was not eliminated in the does.34 Ciprofloxacin (20 mg/kg PO q24h) for 5 days eliminated P. multocida infection in a group of diseased rabbits.16 High tissue concentrations of ciprofloxacin were found in kidney, lung, liver, spleen, and muscle. Parenteral penicillin (24,000 U/kg) penetrated easily and remained at high levels in the pleural space of rabbits with empyema caused by P. multocida.35 Deeb reported success in control of chronic cases of confirmed pasteurellosis using enrofloxacin (5-10 mg/kg PO q12h) or chloramphenicol (50 mg/kg PO q12h) for up to 2 to 3 months.10 Oral antibiotic therapy, however, may be unrewarding in some cases of respiratory disease in rabbits. Treatment failure can be a result of misidentification of the underlying etiology or bacterial disease in the form of nasal cavity or pulmonary granuloma or abscesses, where oral medications are often only partially to poorly effective. In some cases, treatment is enhanced with the addition of antibiotic therapy in the form of nebulization. The addition of mucolytic agents can be helpful as well.18
Flushing of the nasolacrimal duct and instillation of antibiotics into the duct is also useful. To perform a nasolacrimal flush, instill a topical ocular anesthetic, and identify the lacrimal punctum. With a feline nasolacrimal cannula (or a small blunt needle or catheter), gently dilate the punctum and advance the cannula carefully (Fig. 16-7). Successful flushing results in the appearance of fluid from the nasal puncta. Multiple flushing attempts may help resolve infection and occlusion of the duct with infectious/inflammatory debris. Long-term inflammation, rupture of the duct, or dental disease may result in permanent occlusion and failure to flush.7,18 Rhinotomy with surgical debridement and infusion of topical antibiotics may be the most effective treatment option for granulomatous disease of the nasal cavity and sinuses that is unresponsive to antibiotic therapy. In a laboratory rabbit model of Pseudomonas sinusitis, a decrease in bacterial count was achieved with a catheter embedded in the nasal cavity for irrigation with saline and tobramycin. It should be noted that while experimental models decreased bacterial counts in the nasal cavity, complete elimination of organisms in the mucosa was not achieved in this study.1
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C Fig. 16-8 Unilateral rhinotomy in a rabbit with chronic purulent nasal discharge unresponsive to medical therapy. A, The skin and periosteum have been reflected over the nasal bone. A highspeed dental drill with a fine burr is used to perform temporary rhinostomy. Purulent material and necrotic bone are then carefully removed. B, Placement of a catheter is shown. Periosteum is reopposed if possible and skin closed routinely. C, Postsurgical appearance of the rabbit with nasal catheter sutured above the fenestration and skin incision and behind the ears. The catheter is used to deliver small amounts of antibiotic in saline for several weeks after surgery and then removed. Healing of the soft tissues and closure of the rhinostomy site will occur by secondary intention.
The author achieved significant improvement in rabbits with chronic nasal bacterial infection with rhinostomy of the nasal bone (unilateral or bilateral), debridement and flushing, with frequent irrigation via an indwelling nasal catheter. The rabbits tolerated the procedure and postprocedural care extremely well. Rhinotomy is performed after incision of the skin over the nasal bones and deflection and retraction of the periosteum. The nasal bone is perforated with a high-speed precision drill. The nasal cavity is carefully debrided and flushed, and a small-gauge catheter is implanted into the nasal cavity and sutured in place (Fig. 16-8). The choice of antibiotic is based on culture and sensitivity, and the nasal cavity is flushed daily with a small amount of sterile fluid and antibiotics. V. Capello (personal communication, 2009) described a case of bilateral rhinotomy in another patient with more severe disease involving end-stage acquired dental disease (Fig. 16-9). Otitis can contribute to respiratory disease via the spread of organisms through the eustachian tube. Therefore resolution
of respiratory disease may rely on the management of otitis. In cases of inner- and middle-ear infection, lateral resection of the ear canal and/or ostectomy of the bulla may be useful. Both procedures have been described and successfully used in pet rabbits.4 A number of options have been described for treatment of thymoma or thymic lymphoma in rabbits; these include thoracotomy and surgical excision and radiation therapy (Fig. 16-10).8,26,31 Both treatments have specific advantages and disadvantages. Surgery is likely the treatment of choice for single masses without involvement of the great vessels because it offers the best chance for complete removal and cure. Descriptions of the surgical approach via ventral thoracotomy have been published. Successful treatment with radiation therapy has also been reported. The largest risk of the surgical approach is that the tumor lies in close proximity to critical normal thoracic structures.8,26,31 N. Antinoff (personal communication, December
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Fig. 16-9 Bilateral rhinostomy in a rabbit with a chronic purulent nasal discharge unresponsive to medical therapy, following end-stage acquired dental disease and involvement of the alveolar bullae and maxillary recesses. A, Clinical appearance of bilateral nasal discharge and facial dermatitis. Severe respiratory distress was present. Surgical access to nasal cavities is achieved through osteotomy of the nasal bones on their lateral margins. B, The flap is reflected caudally, still attached to the frontal bones. The wide rhinostomy access allows flushing and cleaning of both maxillary recesses. C, The nasal bones are then repositioned. D, Bilateral rhinostomy access is maintained for several days after surgery through two drainage tubes (black arrows), allowing thorough flushing of the nasal cavities (clear arrow). E, The nasal bone flap is surgically removed 10 days after surgery due to necrosis and a permanent rhinostomy is created. F, Follow-up 2 weeks after surgical excision. Healing of soft tissues by secondary intention is almost complete. Breathing has dramatically improved, and the permanent rhinostomy allows frequent flushing of the nasal cavities and maxillary recesses. Note that the nasal septum is still intact. (Used by permission from Vittorio Capello, DVM.)
2008) reported a survival time of 25 months in a patient postradiation therapy. Treatment of dental disease is described in Chapter 32 and elsewhere.7 Treatment of cardiac disease in the rabbit is similar to that in traditional pet species. Drugs reported as useful for cardiac disease in the rabbit include furosemide, enalapril, digoxin, and pimobendam.28
PREVENTION AND CONTROL OF RESPIRATORY DISEASE A large part of prevention of infectious disease is optimal husbandry, including ideal diet, exercise, reduction of stress, and proper sanitation. Potential rabbit owners should be advised
against purchasing rabbits from groups exhibiting nasal or ocular discharge, sneezing, or other signs or symptoms of respiratory disease. Quarantine of new purchases from existing pets is essential. Animals with evidence of respiratory disease should be housed away from other rabbits. Handwashing and disinfecting helps to prevent spread via fomites. Pasteurella-free rabbits are available in the laboratory animal community through Webster’s methods of barrier housing and testing, cesarean derivation, and fostering of kids onto Pasteurella-free does or possibly early weaning with or without antibiotic treatment of does. These animals are unlikely to be found in pet stores.10 Examination of new rabbits immediately after purchase is beneficial, as early identification and treatment are more likely to result in cure.
CHAPTER 16 Respiratory Disease and Pasteurellosis
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Fig. 16-10 Thymoma in pet rabbits. A, Bilateral dynamic exophthalmos is the typical prodromal sign. Dyspnea may or may not be present, depending on the degree of tracheal compression. In the same rabbit, lateral (B) and ventrodorsal (C) radiographic views. B, The cardiac silhouette is displaced caudally; the cranial margin of the heart and the cranial lung lobes are obscured; the trachea is displaced dorsally and compressed. C, The heart is displaced to the left; an abnormal radiodensity of the right side is visible as well. D, Color Doppler ultrasonography of the thorax. Ultrasonography is important to help define the size and relationship of the mass with the heart and great vessels. Thymomas may be solid or, as in this case, contain fluid, which can be drained via ultrasonographic guidance as a palliative treatment. E, Repeat radiograph after ultrasound-guided drainage of fluid. Note that tracheal compression has been significantly reduced. F, Computed tomography (CT) (axial view). Even at the first onset of recognizable symptoms, the mass is often very large, in many cases larger than the heart. G, Three-dimensional rendering of CT image. This view is particularly useful in considering surgical excision in order to study the exact topographic anatomy of the mass. Note the cranial superimposition of the mass over the heart (dotted line). H, Surgical treatment in another rabbit. The most commonly reported surgical approach for thymoma is ventral, after median sternotomy or paramedian distal costocondrotomy, as in this case. Appearance at necropsy of the mass shown in B to G and its relationship with the heart (I). T, Thymoma; H, heart. (Used by permission from Vittorio Capello, DVM.)
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References 1. Antunes MB, Feldman MD, Cohen NAA, et al. Dose-dependent effects of topical tobramycin in an animal model of Pseudomonas sinusitis. Am J Rhinol. 2007;21:423-427. 2. Berglof A, Norlander T, Feinstein R, et al. Association of bronchopneumonia with sinusitis due to Bordetella bronchiseptica in an experimental rabbit model. Am J Rhinol. 2000;14:125-130. 3. Broome RL, Brooks DL. Efficacy of enrofloxacin in the treatment of respiratory pasteurellosis in rabbits. Lab Anim Sci. 1991;41:572-576. 4. Capello V. Lateral ear canal resection and ablation in pet rabbits. Proceedings. N Am Vet Conf. 2006;1711-1713. 5. Capello V, Cauduro A. Clinical technique: application of computed tomography for diagnosis of dental disease in the rabbit, guinea pig, and chinchilla. J Exot Pet Med. 2008;17:93-101. 6. Capello V, Gracis M, Lennox L. Rabbit and rodent dentistry handbook. Lake Worth, FL: Zoological Education Network; 2005;165-186. 7. Capello V, Lennox AM. Clinical radiology of exotic companion mammals. Ames, IA: Wiley-Blackwell; 2008. 8. Clippinger TL, Bennett RA, Alleman AR, et al. Removal of a thymoma via median sternotomy in a rabbit with recurrent appendicular neurofibrosarcoma. J Am Vet Med Assoc. 1998;213:1140-1143. 9. Deeb B. The dyspneic rabbit. Exot DVM. 2005;7:39-42. 10. Deeb BJ. Respiratory disease and pasteurellosis. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. Philadelphia: WB Saunders; 2004:172-182. 11. Deeb BJ, DiGiacomo RF, Bernard BL, et al. Pasteurella multocida and Bordetella bronchiseptica infections in rabbits. J Clin Microbiol. 1990;28:70-75. 12. Dei-Cas E, Chabe M, Moukhlis R, et al. Pneumocystis oryctolagi sp. nov, an uncultured fungus causing pneumonia in rabbits at weaning: review of current knowledge and description of a new taxon on genotypic, phylogenetic and phenotypic bases. FEMS Microbiol Rev. 2006;30:853-871. 13. Dufour X, Kauffman-Lacroix C, Goujon JM, et al. Experimental model of fungal sinusitis: a pilot study in rabbits. Ann Otol Rhinol Laryngol. 2005;114:162-172. 14. Fidan F, Unlu M, Sezer M, et al. Acute effects of environmental tobacco smoke and dried dung smoke on lung histopathology in rabbits. Pathology. 2006;38:53-57. 15. Grint NJ, Sayers IR, Cecchi R, et al. Postanaesthetic tracheal strictures in three rabbits. Lab Anim. 2006;40:301-308. 16. Hanan MS, Riad EM, el-Khouly NA. Antibacterial efficacy and pharmacokinetic studies of ciprofloxacin on Pasteurella multocida infected rabbits. Deutsch Tierarztl Wochenschr. 2000;107:151-155. 17. Harcourt-Brown F. Cardiorespiratory disease. In: Textbook of rabbit medicine. Philadelphia: Elsevier Science Limited; 2002:324-334.
18. Harcourt Brown F. Ophthalmic diseases. In: Textbook of rabbit medicine. Philadelphia: Elsevier Science Limited; 2002:292-306. 19. Heatley JJ, Smith AN. Spontaneous neoplasms of lagomorphs. Vet Clin North Am Exot Anim Pract. 2004;7:561-577. 20. Hernandez-Divers SJ. The rabbit respiratory system: anatomy, physiology, and pathology. Proceedings. Annu Conf Assoc Avian Vet/Assoc Exot Mam Vet. 2007:61-68. 21. Hu XJ, Wang Q, Tong SP, et al. Comparative study of ceftazidime administered in continuous versus intermittent infusion in a rabbit Pseudomonas aeruginosa penumonia model. Chin J Antibiotics. 2008;33:363-368. 22. Jin L, Valentine BA, Baker RJ, et al. An outbreak of fatal herpesvirus infection in domestic rabbits in Alaska. Vet Pathol. 2008;45:369-374. 23. Kritas SK, Dovas C, Fortomaris P, et al. A pathogenic myxoma virus in vaccinated and nonvaccinated commercial rabbits. Res Vet Sci. 2008;85:622-624. 24. Manning PJ, DiGiacomo RF, DeLong D. Pasteurellosis in laboratory animals. In: Adlam C, Rutter JM, eds. Pasteurella and pasteurellosis. London: Academic Press; 1989:263-302. 25. Marlier D, Mainil J, Linde A, et al. Infectious agents associated with rabbit pneumonia: isolation of amyxomatous myxoma virus strains. Vet J. 2000;159:171-178. 26. Morrisey JK, McEntee M. Therapeutic options for thymoma in the rabbit. Sem Avian Exot Pet Med. 2005;14:175-181. 27. Okerman L, Spanoghe L, DeBruycker R. Experimental infections of mice with Pasteurella multocida strains isolated from rabbits. J Comp Pathol. 1979;89:51-55. 28. Pariaut R. Cardiovascular physiology and diseases of the rabbit. Vet Clin North Am Exot Anim Pract. 2009;12:135-144. 29. Phaneuf LR, Barker S, Groleau MA, et al. Tracheal injury after endotracheal intubation and anesthesia in rabbits. J Am Assoc Lab Anim Sci. 2006;45:67-72. 30. Rougier S, Galland D, Boucher S, et al. Epidemiology and susceptibility of pathogenic bacteria responsible for upper respiratory tract infections in pet rabbits. Vet Microbiol. 2006;115:192-198. 31. Sanchez-Migallon DG, Mayer J, Gould J, et al. Radiation therapy for the treatment of thymoma in rabbits (Oryctolagus cuniculus). J Exotic Pet Med. 2006;15:138-144. 32. Sjoberg JG. Foreign body in the nose of a rabbit. Exot DVM. 2005;6:19. 33. Suckow MA, Haab RW, Miloscio LJ, et al. Field trial of a Pasteurella multocida extract vaccine in rabbits. J Am Assoc Lab Anim Sci. 2008;47:18-21. 34. Suckow MA, Martin BJ, Bowersock TL, et al. Derivation of Pasteurella multocida-free rabbit litters by enrofloxacin treatment. Vet Microbiol. 1996;51:161-168. 35. Teixeira LR, Sasse SA, Villarino MA, et al. Antibiotic levels in empyemic pleural fluid. Chest. 2000;117:1734-1739. 36. Weisbroth SH, Hurvitz A. Spontaneous osteogenic sarcoma in Oryctolagus cuniculus with elevated serum alkaline phosphatase. Lab Anim Care. 1969;19:264-268.
CHAPTER
17
Disorders of the Reproductive and Urinary Systems
Eric Klaphake, DVM, Diplomate ACZM, Diplomate ABVP (Avian), and Joanne Paul-Murphy, DVM, Diplomate ACZM
Disorders of the Reproductive System Uterine Adenocarcinoma Endometrial Hyperplasia or Uterine Polyps Pyometra and Endometritis Pregnancy Toxemia Pseudopregnancy Dystocia or Retained Fetuses Abdominal Pregnancy Abortion and Resorption Reduced Fertility Prolapsed Vagina Endometrial Venous Aneurysms Hydrometra Uterine Torsion Uterus Unicornis and Uterine Atresia Cryptorchidism Orchitis and Epididymitis Testicular Neoplasms Venereal Spirochetosis Disorders of the Mammary Glands Septic Mastitis Cystic Mastitis, Mammary Dysplasia, and Mammary Tumors Disorders of the Urinary System Urolithiasis and Hypercalciuria Renal Failure Hypervitaminosis D Nephrotoxicity Renal Adipose Deposition Renal Cysts Renal Agenesis Encephalitozoonosis Urinary Incontinence Psychogenic Polyuria and Polydipsia Urinary Bladder Eversion Tumors of the Urinary Tract Red Urine Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
DISORDERS OF THE REPRODUCTIVE SYSTEM Rabbit does are prolific breeders. They reach sexual maturity by 4 to 6 months of age, are induced ovulators, and have serial cycles that last 7 to 14 days until conception. A mildly swollen and congested vulva often accompanies receptivity and should not be mistaken for a vulvitis or vaginitis. Bucks are precocious and may attempt to breed as early as 3½ to 4 months of age. Spraying is a normal sexual behavior of intact bucks and sometimes of does and should not be confused with inappropriate elimination because of urinary tract infection or inflammation. In an evaluation of wild European hares (Lepus europaeus), reproductive abnormalities were found in 20.8% (51 of 245) of the does examined. Cystic endometrial hyperplasia was relatively common and often accompanied by hydrosalpinx. Extrauterine fetuses, neoplasms, pseudopregnancies, and resorptions also were also found. However, although pseudopregnancies and resorptions were found in young adults (<12 months) as well as older hares, conditions possibly causing infertility were almost always seen in older hares, with prevalences up to 46.2%. Only hares with access to known sources of estrogens (mixed pastures including legumes known to produce phytoestrogens and mycoestrogens) exhibited pathologic conditions, but sympatric European rabbits (Oryctolagus cuniculus) did not, which is consistent with known difference in responses between the corpora lutea of the two species to exogenous estrogen. Cystic ovarian tumors seem to be relatively common in hares, and tumors of the mammary gland and uterus have also been reported.76 Disorders of the reproductive tract in female pet rabbits are seen less commonly because educated owners typically elect to have their rabbits spayed to prevent uterine disease. In the second edition of this text, it was reported that less than 6% of all rabbits presented to the University of Wisconsin Veterinary Medical Teaching Hospital were diagnosed with reproductive disorders.62 A review of the data from the University of California-Davis Veterinary Medical Teaching Hospital reported a 4% incidence of uterine disorders (not including healthy rabbit ovariohysterectomies) from January 1999 through January 2009 in presenting rabbits (JPM). However, older intact females must be carefully and regularly monitored for reproductive tract disease, especially neoplasia. Careful abdominal palpation 217
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allows for the detection of ovarian or uterine enlargement. Any perceived abnormality should be investigated fully and aggressively.
UTERINE ADENOCARCINOMA Uterine adenocarcinoma is the most common neoplasia of female rabbits.85 Age is the most important factor in its development and occurrence is independent of breeding history. Rabbits of certain breeds (tan, French silver, Havana, and Dutch) that are more than 4 years old have an incidence of 50% to 80%.4,37,85 With age, the endometrium undergoes progressive changes, a decrease in cellularity, and an increase in collagen content. These changes are associated with the development of uterine cancer.4 Adenocarcinoma of the uterus is a slowly developing tumor. Local invasion of the myometrium occurs early and may extend through the uterine wall to adjacent structures in the peritoneal cavity; hematogenous metastasis to the lungs, liver, brain, and bones may occur within 1 to 2 years.85 Early clinical signs, such as decreased fertility, small litter size, and an increased incidence of fetal retention, fetal resorption, and stillbirths may be recognized in a breeding doe. The first signs observed in many pet rabbits are hematuria or a serosanguineous vaginal discharge. Frank blood in the urine is most pronounced at the end of urination. Cystic mammary glands can develop concurrently with uterine hyperplasia or adenocarcinoma.32,45 Clinical signs of late-stage adenocarcinoma include depression, anorexia, and dyspnea if pulmonary metastasis has occurred. Ascites may be present. The diagnosis relies on palpation of an enlarged uterus, uterine nodular masses (1-5 cm in diameter), or both in the caudal abdomen. Adenocarcinomas are often multicentric, involving both uterine horns (Fig. 17-1).4,85 Other causes of uterine enlargement include pregnancy, pyometra, metritis, hydrometra or mucometra, endometrial venous aneurysms, endometrial hyperplasia, and other tumors such as leiomyosarcoma. In one pet rabbit, both a leiomyoma and an adenocarcinoma of the endometrium were found, with the latter extending into the leiomyoma.48
The expression of estrogen receptor-alpha and progesterone receptor in normal, hyperplastic, and neoplastic endometrium in rabbits was evaluated by immunohistochemistry. Sixteen of 27 cases of normal uteri (59.3%) and 13 out of 19 hyperplasias (68.4%) stained positive with both receptor types. Adenocarcinomas were further subdivided into 26 papillary and 16 tubular/solid adenocarcinomas. Papillary adenocarcinoma infiltrated the myometrium late in the disease and caused attenuation of the myometrium. In contrast, tubular/solid adenocarcinoma invaded the deep myometrium early in the disease without thinning of the myometrium. Twenty-one out of 26 (80.8%) cases of papillary adenocarcinoma were receptor negative for both types, whereas 15 out of 16 (93.8%) of the tubular/ solid adenocarcinomas were positive for one or both, suggesting that there may be two different developmental pathways for uterine adenocarcinomas in the rabbit.2 Radiography and ultrasound imaging assist in establishing the diagnosis when a caudal abdominal soft tissue mass or uterine enlargement can be identified. Imaging may further allow for differentiation between potential causes of uterine enlargement, the measurement of masses, or scanning for multiple nodules. Perform thoracic radiographs to screen for the presence of pulmonary metastasis. Pulmonary involvement carries a grave prognosis. If abdominal masses are identified in the early stages of disease before metastasis occurs, surgical excision with ovariohysterectomy is the treatment of choice. Ovariohysterectomy is curative if the tumor is contained within the uterus. Metastatic disease or local invasion of peritoneal structures may not be macroscopically apparent at laparotomy, and a guarded prognosis is always warranted. Reexamine the rabbit every 3 months for a period of 1 to 2 years after surgery for evidence of abdominal or pulmonary metastases. Successful chemotherapy for this tumor has not been reported. Prevention is the key to management of this disease. Client education begins at the time of the initial visit for routine health evaluation of the patient. Many veterinarians recommend ovariohysterectomy for pet rabbits before they reach 2 years of age. Spaying rabbits between the ages of 6 and 9 months is preferred because they have less abdominal fat than older rabbits. Alternatively, discuss early clinical signs of the disease with the client and recommend semiannual health examinations for intact female rabbits 3 years of age and older. Other reported uterine neoplasias in rabbits include spontaneous choriocarcinoma and a homologous malignant mixed müllerian tumor.29,43
ENDOMETRIAL HYPERPLASIA OR UTERINE POLYPS
Fig. 17-1 Surgically removed uterine adenocarcinoma with secondary pyometra from a 4-year-old doe.
Endometrial changes may occur along a continuum—from polyp formation, to cystic hyperplasia, to adenomatous hyperplasia, to adenocarcinoma—as it does in humans.20,37,45 Uterine hyperplasia is associated with aging, as are cystic and hyperplastic changes in endometrial glands. Other reports, however, have found no association between cystic hyperplasia and uterine adenocarcinoma in rabbits because adenocarcinomas are associated with senile atrophy of the endometrium.4 Cystic endometrial hyperplasia is the major indication of phytoestrogenism in sheep. Hares are particularly sensitive to the luteotrophic effect of exogenous estrogen, which can double the length of pseudopregnancy in that species; however this did not seem to be the case in sympatric rabbits.75
CHAPTER 17 Disorders of the Reproductive and Urinary Systems Clinical signs of endometrial hyperplasia can mimic those associated with uterine adenocarcinoma, including intermittent hematuria, anemia, and a decrease in activity. A firm, irregular uterus can sometimes be detected by palpation. Cystic mammary glands and cystic ovaries can occur concurrently with this condition.32,45 Ultrasonography is the most efficient diagnostic tool to image soft tissue changes in the uterus although other imaging modalities can provide the diagnosis of uterine changes. Ovariohysterectomy is the recommended treatment, and a thorough exploration of the abdomen is warranted.
PYOMETRA AND ENDOMETRITIS Vaginal discharge, anorexia, lethargy, weakness, and an enlarged abdomen are clinical signs that frequently accompany endometritis or pyometra. Clinical signs of mild endometritis can be subtle, making the condition difficult to diagnose. Rabbits with chronic disease may have no overt clinical signs. The history of an affected breeding doe often includes a recent parturition, pseudocyesis, or an inability to rebreed. Rabbits with mild endometritis may kindle successfully or have fetal resorptions and stillbirths. Pyometra and endometritis can also develop in nulliparous does. Pyometra can be secondary to uterine adenocarcinomas, indicating the importance of histopathology on all grossly abnormal uteruses. The main reasons for culling adult rabbit does on two Spanish rabbit farms over 1 year included pyometra (8.7%). Pasteurella species were more prevalent, although two strains of Staphylococcus aureus were identified by using polymorphism of the coagulase gene as the criterion. One of these strains was responsible for the majority of the staphylococcal infections and was isolated from several pathological processes.72 S. aureus was found to be a cause in another study.81 Diagnosis relies on palpation of a doughy, enlarged uterus, best demonstrated on radiographs or ultrasound. If the uterus is greatly enlarged, use caution in palpating the abdomen, because the uterine wall becomes very thin and may be friable. Ultrasound is advantageous because it can often rule out other uterine conditions such as polyps, masses, or cystic changes. Results of a complete blood count (CBC) may be normal or may show a slight leukocytosis due to a heterophilia. Evaluate serum or plasma biochemical values, because chronic inflammation of the uterus has been reported to induce amyloid deposition in the kidneys.35 Cytologic assessment and a Gram’s stain of cervical mucus or drainage can assist diagnosis. Exploratory laparotomy and ovariohysterectomy are the procedures of choice to confirm the diagnosis. Uterine vessels may be engorged and prominent. Multiple adhesions to adjacent viscera often complicate the procedure. Obtain an intraoperative sample for culture. Begin broad-spectrum antibiotic therapy as soon as a sample for culture has been collected. Intravenous fluids and pain management are important components of therapy. Pasteurella multocida and S. aureus are frequently isolated from rabbits with pyometra or metritis. Venereal transmission occurs when infected does breed with uninfected bucks, or vice versa. P. multocida can localize in the genital tract by hematogenous spread from another location, or a retrograde infection can occur as a result of vaginitis.17 Ovarian abscesses may occur concurrently with P. multocida pyometra.39 Rare cases of naturally occurring metritis or pyometra have been associated with Chlamydia species, Listeria monocytogenes, S. aureus, Moraxella
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bovis, Actinomyces pyogenes, Brucella melitensis, and Salmonella species.34,59,74,84 Postpartum metritis can occur in conjunction with hypervitaminosis A; the delivery of stillborn young and metritis have been associated with uterine torsion.34 For a breeding rabbit with mild endometritis, appropriate antibiotic treatment, the administration of nonsteroidal anti-inflammatory drugs (NSAIDs), and fluid therapy may be sufficient, but the tenacious and caseous nature of inflammatory exudates in rabbits makes it impossible to achieve adequate drainage of the uterus. The use of prostaglandins to assist uterine contraction and drainage has not been reported. Ovariohysterectomy is the best choice for the treatment of pyometra.
PREGNANCY TOXEMIA Pregnancy toxemia usually occurs during the last week of gestation, when nest-building behavior begins. Rabbit fetuses show considerable development during the second half of pregnancy.23 The exact causes leading to metabolic disturbances and pregnancy toxemia are unknown. It is more common in obese rabbits. Inadequate caloric intake predisposes pregnant rabbits to toxemia, and environmental changes or stress can precipitate the disease. Hair pulling for nesting can contribute to hairball formation and inappetence.63 Weakness, depression, incoordination, anorexia, abortion, convulsions, and coma are common clinical signs. Signs can progress over 1 to 5 days, or death may occur acutely. Some rabbits are dyspneic and may have ketotic breath. The urine becomes acidic and clear because the lower pH decreases the concentration of calcium carbonate crystals. Clinical pathology findings include acidic urine (pH 5-6), proteinuria, ketonuria, hyperkalemia, ketonemia. hyperphosphatemia, and hypocalcemia. Hepatic enzymes may be elevated and lipidosis is commonly noted at necropsy. There is no consistently effective treatment for pregnancy toxemia. Keep the animal warm and administer intravenous or intraosseous fluids. Calcium gluconate can be administered parenterally to treat hypocalcemia. Analgesics may be indicated; however, concerns regarding the gastrointestinal effects of opioids and renal/hepatic effects of NSAIDs should be evaluated on a case-by-case basis. When the rabbit is stabilized, nutritional support must be provided, either by syringe feeding, esophagostomy, or nasogastric or percutaneous endoscopically placed gastrostomy tubes. Pregnancy toxemia carries a very grave prognosis; treatment is usually unrewarding by the time it is recognized. The best approach is prevention. Avoid fasting or underfeeding during late pregnancy and prevent obesity and sudden stress at all times. If feeding timothy hay-based pellets, consider switching to alfalfa-based pellets until postweaning.
PSEUDOPREGNANCY Pseudopregnancy (pseudocyesis, false pregnancy) can occur in rabbits, even in pet does kept singly. Pseudopregnancy typically lasts 16 to 17 days and may be followed by hair pulling and nesting behavior.64 The corpus luteum secretes progesterone, causing uterine and mammary development.60 Mammary development is most pronounced in the first 10 days of false pregnancy, after which mammary involution typically follows by day 16.60,64 The condition resolves spontaneously but may recur or may lead to hydrometra or pyometra. Pseudopregnancy strongly depresses fertility, and its cause is still unknown.79 A lower kindling rate was found in group-housed does with one
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buck versus those housed in a one-to-one ratio, with pseudopregnancies found in 23% of the does in the group-housing system against 0% in the control does housed individually.69 Ovariohysterectomy is the treatment of choice and ideally would be performed once the mammary tissue had involuted. Hormonal therapy with either progestins or androgens is of unproved effectiveness and should be considered only in rare cases where pseudopregnancy is protracted or refractory.
timely parturition.28 New husbandry systems in rabbits, such as artificial insemination, are factors to be considered.28 Surgery is the treatment of choice for the individual animal to remove the fetuses, and ovariohysterectomy is indicated if the pregnancy is identified early enough. Euthanasia may be required in advanced cases of abdominal pregnancy.
DYSTOCIA OR RETAINED FETUSES
Fetal death before 21 days of gestation typically resolves as resorption, whereas fetal death after 21 days results in abortion. A critical period in gravid does occurs at 21 days of gestation because of a temporary reduction in blood flow to the uterus and the changing size and shape of the fetuses. When abortion or fetal resorption is suspected, a thorough history is extremely important. Ask questions such as: “Is this the first litter? Is there a prior history of abortion? Have any medications been administered recently? Has there been a recent change in environment? Are any other does aborting?” Always check the doe for remaining fetuses by radiographs and/or ultrasound. Submit fetuses and placentas for anerobic and aerobic bacterial culture and histopathologic examination. Fetal resorption is not regarded as a disease condition in lagomorphs, since it occurs in response to social or environmental stress.76 Possible causes of fetal abortion are numerous and include infection, stress, genetic predisposition, trauma, drug use, and dietary imbalances (e.g., deficiencies of vitamin E, vitamin A, and protein). Listeria species have a predilection for the gravid uterus; this should be considered in rabbits with late-term abortion.84 A herpesvirus infection affecting mini Rex and crossbred meat rabbits was identified in a rabbitry in Alaska, with one of the symptoms being abortion.38
Gestation in rabbits averages 31 to 32 days but may be as short as 28 or as long as 36 days. Normal delivery is usually complete within 30 minutes after its onset, although it is rare for the young to be delivered several hours apart. Litters range from 4 to 12 kits. Anterior and breech positions are normal for rabbits. Palpate does 24 hours after delivery to determine whether any fetuses have been retained. Dystocia is rare in rabbits. A rabbit may be predisposed to dystocia by obesity, large fetuses, a small pelvic canal, or uterine inertia. Signs of dystocia include persistent contractions, straining, and bloody or greenish-brown vaginal discharge. Take radiographs to determine abnormalities in the fetuses or in the width of the pelvic canal. Assistance usually requires analgesia, sedation, gentle manual removal of the fetuses, and rapid removal of fetal membranes from them. Oxytocin (1-3 U IM) can assist uterine contraction. If uterine inertia is suspected, give 5 to 10 mL of 10% calcium glubionate orally 30 minutes before administering the oxytocin. Place the doe in a quiet, dark room for 30 to 60 minutes after administration. Perform a cesarean section if there is no response to the injection of oxytocin and the prognosis is guarded. Focus should be placed on saving the doe unless adequate staff is present to try also to revive the kits.
ABDOMINAL PREGNANCY Abdominal pregnancy sometimes occurs in does and is usually subclinical.1,6,7,16 Extrauterine implantation of fertilized ova, often on the parietal peritoneum, can lead to the development of near-term fetuses, with subsequent mummification.7 It is classified as a primary abdominal pregnancy if there is no evidence of uterine rupture, with presumed regurgitation of early embryos from the uterine tube. The abdominal pregnancy is secondary if there is evidence of uterine rupture. A study of 550 adult fertile female New Zealand white rabbits culled from two rabbit farms in Valencia (Spain) found that 5% had abdominal pregnancies, with 25% of those having normal reproductive tracts.28 Mummified fetuses were found free in the abdominal cavity of a healthy doe presented for a routine examination.6 The uterus and ovaries appeared normal, and the rabbit made an uneventful recovery after surgical removal of the fetuses. False abdominal pregnancy refers to cases in which fetal implantation occurred in the uterus and fetuses were later expelled into the abdominal cavity as a result of a traumatic event.7 One report describes a rabbit mortality associated with three partially mummified, full-sized fetuses free in the abdominal cavity.1 On postmortem examination, one uterine horn showed evidence of a relatively recent tear. The doe had delivered three kits 3 weeks previously, suggesting that the uterine rupture had occurred at parturition. Primary abdominal pregnancy or false abdominal pregnancy should be considered on the differential when palpable fetuses are present in does that have not undergone
ABORTION AND RESORPTION
REDUCED FERTILITY One or more factors can contribute to reduction in fertility, including malnutrition (e.g., excess vitamin A, deficiencies of vitamins A, D, or E), heat stress, systemic illness, nitrate contamination of food or water, environmental disturbances, a decrease in daylight, endometrial carcinoma, metritis, or pyometra. Old age, sexual exhaustion, or breeding of rabbits that are too young are additional causes of infertility. Vitamin E deficiency causes myodystrophy, which can lead to abortions, stillbirths, and neonatal deaths. An increased level of creatine phosphokinase occurs with hypovitaminosis E and indicates a need for diet supplementation.59 Hypervitaminosis A can cause fetal resorptions, abortions, and stillbirths.18,75 A suppurative metritis may follow the delivery of dead fetuses.18 Hypovitaminosis A can cause similar reproductive disorders, resulting in poor fertility and weak, hydrocephalic young. The National Research Council recommends vitamin A levels of 1,160 IU/kg of diet for gestation, or approximately 20 mg/kg of body weight per day.57 Administration of vitamin E for 2 weeks lowered the serum vitamin A levels and increased the vitamin E serum and liver levels. It was concluded that vitamin E therapy appears to be an effective treatment for hypervitaminosis A.75
PROLAPSED VAGINA A prolapsed vagina manifests itself as a blood-covered mass of swollen and fragile tissue protruding from the vulva. The prolapse may be full of clotted blood. Affected rabbits are depressed or recumbent with an increased respiratory rate or are in shock
CHAPTER 17 Disorders of the Reproductive and Urinary Systems
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with cool extremities. Pale mucous membranes or cyanotic ears and mucous membranes indicate severe shock. The hematocrit in such rabbits may be as low as 9%.82 Treatment is directed at correcting hypovolemic shock and blood loss. Opioid analgesia is recommended and NSAIDs can help to reduce inflammation as well as providing analgesia. Reduce the prolapse under anesthesia or surgically amputate the tissue if it is necrotic. Prolapses start from the proximal circular part of the vaginal vault just distal to the urethral opening.82 Eight cases of vaginal prolapse were described in closely related rabbits during periods of increased sexual activity or receptivity, suggesting a genetic susceptibility.82
ENDOMETRIAL VENOUS ANEURYSMS Multiple endometrial venous aneurysms can cause hematuria because of episodic bleeding in the lumen of the uterus. Cylindrical blood clots molded within the uterine horns are typically passed with the urine and are highly suggestive of this condition. Affected does are at high risk for fatal exsanguination from uterine hemorrhage, and ovariohysterectomy should be performed as soon as the animal is stabilized. Endometrial venous aneurysms occur in young does of larger breeds. The condition was reported in three New Zealand white rabbits and was confirmed by exploratory laparotomy and histopathologic examination of the uterus.9 In rabbits with this condition, the uterine horns have multiple blood-filled endometrial varices (veins) that periodically rupture into the uterine lumen, causing the clinical hematuria. Although the causes have not been elucidated, venous aneurysms in other species are related to congenital defects of the adventitia, increased intraluminal pressure, or trauma.9
HYDROMETRA Hydrometra is the accumulation of watery fluid in the uterus. It has been described in four unbred sandy half-lop rabbits from the same research colony55 and in New Zealand white rabbits.34 Clinical signs include an enlarged fluid-filled uterus, increased respiratory rate, anorexia, and weight loss. Transabdominal uterocentesis yields clear fluid with a low specific gravity, a low cell count, and a moderate amount of protein.34 Diagnosis can be supported by radiography and ultrasonography. The rabbits in the published reports were all euthanatized or found dead, and no anatomic abnormalities could be correlated with this condition.55 Ovariohysterectomy and supportive care are indicated if hydrometra is diagnosed in a pet rabbit.
UTERINE TORSION Torsion of the uterus is rare in rabbits but has been reported in association with pregnancy, hydrometra, and endometritis.34 Clinical signs of uterine torsion include shock, cachexia, and abdominal distention with hydrometra, or a bloody vaginal discharge when seen with endometritis.34 The cause of uterine torsion is difficult identify and the prognosis is grave. Following critical support, pain management, and stabilization of the doe, ovariohysterectomy is the treatment of choice.
UTERUS UNICORNIS AND UTERINE ATRESIA Two cases of congenital anomalies seen in domestic rabbits have been described. Uterus unicornis was reported in a 4-year-old intact female Rex rabbit presenting with some
Fig. 17-2 Uterine atresia in a healthy, nonclinical 9-month-old laboratory New Zealand white rabbit undergoing surgical implantation for a research study. Note surgically exteriorized and caudally reflected reproductive tract. The left uterine tube is normal, while the right side is engorged with fluid at distal aspect with a blind proximal end as well as a small fluid-filled solitary uterine tube segment between larger structure and vagina. The right cervix is absent. (Courtesy of Dr. Matthew Johnston.)
urinary and behavioral issues involving two ovaries but only one uterine horn, cervix, and oviduct. There was no evidence of previous abdominal surgery. After a modified ovariohysterectomy, the rabbit recovered uneventfully. Uterine atresia was incidentally found in a 9-month-old intact female white New Zealand rabbit. The left portion of the uterus was normal but the right lacked a cervix and the caudal portion of the right uterus. A large and a second smaller cystic structure midway between the vagina and ovary in the mesometrium was noted (Fig. 17-2). The oviduct was traced to a normalappearing right ovary. There have been anecdotal reports of uterus unicornis on the Veterinary Information Network (VIN, www.vin.com, accessed February 19, 2009) in other rabbits. The congenital etiology of uterine anomalies makes it important to assess the affected animal for renal and ureteral abnormalities.80
CRYPTORCHIDISM In bucks, the testicles have usually descended by 12 weeks of age. Failure of one or both testicles to descend into the scrotal sacs by 4 months of age is defined as unilateral or bilateral cryptorchidism. Rabbits, however, do have the ability to retract their testicles intra-abdominally. Cryptorchid rabbits retain their sexual drive but fertility is impaired. In true cryptorchid bucks, the scrotal sac of the retained testicle does not develop (Fig. 17-3). If the scrotal sac is present but the testicle cannot be palpated, it has most likely been retracted into the inguinal canal or the abdominal cavity. Neither the causes nor the medical implications of cryptorchidism for rabbits have been adequately investigated. Based on other species, it appears prudent to assume a hereditary pattern for cryptorchidism in at least some rabbits as well as a potential for the development of testicular neoplasia in untreated animals. Cryptorchid rabbits should be castrated using a standard midline abdominal approach. Incorrect age history may lead to unnecessary laparotomy, so waiting several more months beyond the estimated 16-week age estimate would be prudent.
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SECTION II Rabbits Collagenous hamartomas are benign proliferations of connective tissue that occur in rabbits in two forms: as a solitary nodule or in a disseminated form with multiple nodules. In one report, the average age was 5.6 years and all but one were middle-aged males, with nodules most commonly located on the abdomen or the thorax. These rabbits also had testicular tumors.83 The etiology of collagenous hamartomas remains unclear but is suggestive of a hormonal etiology or an X-linked recessive genetic disease. The disseminated form of this disease must be differentiated from malignant lymphoma.
VENEREAL SPIROCHETOSIS
A
B Fig. 17-3 A, Unilateral cryptorchid rabbit with lack of scrotal development (arrows). B, Image shows both testes exteriorized to emphasize the discrepancy in testicular size. (Courtesy of Dr. Stephen Barten.)
ORCHITIS AND EPIDIDYMITIS Clinical signs of orchitis include fever, intermittent appetite, and weight loss. The testicles may be enlarged with obvious abscesses, or abscesses may be small and internal, with minimal swelling of the testicles. The epididymis rather than the testis may be infected. An affected breeding buck has low conception rates. Treatment is castration, antibiotic therapy, and NSAIDs. Pasteurella multocida is often isolated from exudate or abscessed tissue on bacterial culture. Specific culture or testing for Treponema species should also be requested. Male rabbits should be housed separately to prevent bite wounds to the testes or scrotum and other fighting injuries that lead to abscesses.
TESTICULAR NEOPLASMS Testicular tumors are rare in rabbits but include seminomas, interstitial cell tumors, Sertoli cell tumors, and teratomas.58,85 A tentative diagnosis is typically based on palpation of an enlarged, firm or nodular, nonpainful testicle. A definitive diagnosis is achieved with surgical castration and submission of the affected testicle for histopathologic examination. Rarely, tumors are incidentally found on the cut section of an otherwise normal-looking testicle.88
Treponema paraluiscuniculi is the spirochete responsible for rabbit syphilis or vent disease. This is not a zoonotic disease. Transmission between rabbits is by direct and venereal contact. Bucks can spread the disease to several does, and young rabbits can be infected. It is a self-limiting disease; carriers may be asymptomatic until stress occurs. Lesions first appear on the skin of the perineum and genitalia and begin as areas of redness that progress to edema, vesicle formation, ulcerations, and scabs. The lesions can be painful and can impair breeding activity. Autoinfection can lead to facial lesions around the chin, lips, nostrils, and eyelids. Inguinal lymph nodes may be enlarged. Colony epidemics result in a decreased rate of conception and an increased incidence of metritis, placental retention, and neonatal deaths. Clinical signs and the distribution of lesions are often diagnostic. Few other skin problems resemble those of rabbit syphilis, but bacterial dermatitis, dermatophytosis, acariasis, and myxomatosis are possibilities. Lesions on the nose and lips are often proliferative and scaly and are commonly mistaken for dermatophyte lesions. For a definitive diagnosis, submit a skin biopsy sample and request silver staining. Examine skin scrapings of the lesions by dark-field microscopy to identify the organism. Skin biopsies and cytology, though, may not contain Treponema species bacteria, leading to false-negative results (T. Donnelly, personal communication). Large rabbitries can benefit from a serologic survey with the microhemagglutination test to screen and verify infected animals. Other serologic tests are available, such as the rapid plasma reagin card test, which is very specific.19,70 Two nontreponemal serological antigen tests are available, the Venereal Disease Research Laboratory (VDRL) test, which is more sensitive, and the rapid plasma reagin (RPR) test, which is more specific.19,70 Nontreponemal tests look for the presence of nonspecific reagin antibodies that appear and rise in titers following infection. The fluorescent treponemal antibody-absorption test is a serologic test used as a confirmatory treponemal antigen test. The RPR is the most common test, and many labs confirm with fluorescent treponemal antigen preparation. Serology is more sensitive and less expensive than biopsies and is usually performed by specific diagnostic labs dealing with laboratory animals. A minimum of 3 months is needed for antibody levels to be measurable, even with clinical lesions. Titers fall rapidly with successful treatment.54 In rabbitries, the prevalence of T. paraluiscuniculi infection increases with parity: often females that have had six or more litters are seropositive; bucks that have been in a breeding program for 6 to 12 months are also likely to be seropositive.19 Bucks are often asymptomatic carriers and may have small starshaped scars on their scrotum.
CHAPTER 17 Disorders of the Reproductive and Urinary Systems Rabbits with venereal spirochetosis are effectively treated with parenteral (never oral) penicillin. Administer penicillin G benzathine-penicillin G procaine at 42,000 to 84,000 IU/kg SC at 7-day intervals for three injections15 or penicillin G procaine at 40,000 to 60,000 IU/kg IM q24h for 5 to 7 days. Tetracyclines and chloramphenicol can also be effective. Treat all exposed rabbits.
DISORDERS OF THE MAMMARY GLANDS Does usually possess 8 but may have up to 12 mammary glands.64 Disorders of the mammary glands are inflammatory, infectious, hormonal, or neoplastic and are often accompanied by ovarian or uterine abnormalities or both.7
SEPTIC MASTITIS Mastitis can occur in a lactating doe or in a rabbit in pseudocyesis. Abscesses can develop in the mammary glands independent of lactation, but mastitis is rare in ovariohysterectomized rabbits. Heavy lactation, poor sanitation, abrasive bedding or caging, or injury to the gland or teat predisposes the doe to mammary infection. Mastitis can occur in conjunction with metritis. Clinical signs may include depression, fever, anorexia, polydipsia, septicemia, and death of the doe or the young. The mammary glands are firm, hot, and swollen, and the skin is discolored red to dark blue. Infection can begin in one gland and spread to others. The initial discharge may not be purulent. Mastitis was the most common reason for culling adult rabbit does on two Spanish rabbit farms, with a 33.3% incidence in the culled population. Staphylococcus aureus infections were associated with severe mastitis, the organism being isolated from 69.2% of infected animals. Two strains of S. aureus were identified by using polymorphism of the coagulase gene as the criterion, with one strain responsible for the majority of the staphylococcal infections, and it was isolated from several pathologic processes.72 Streptococcus species, S. aureus, and Pasteurella species are most frequently isolated. Suckling kits may die of peracute septicemia, particularly with S. aureus mastitis.72,81 Cytology and bacterial culture samples are obtained by expressing exudates from the gland. Choose systemic antibiotic therapy based on the results of aerobic and anaerobic culture and sensitivity testing. Common antibiotic choices are enrofloxacin or a trimethoprim-sulfamethoxole combination. Supportive care includes fluid therapy, pain management with NSAIDs or opioids, application of hot packs, and massaging the tissue to promote drainage if not caseous. Surgical excision or mastectomy is often necessary for severe infections. Force-feeding may be necessary if the doe is anorexic. Remove young rabbits but do not foster them onto another doe; this is known to transmit infection. Bottle-raising young rabbits is a challenging process, particularly when weaning; thus euthanasia can be a more humane option for the rabbits and owners. Disinfect the environment.
CYSTIC MASTITIS, MAMMARY DYSPLASIA, AND MAMMARY TUMORS Noninfectious cystic mastitis occurs in both breeding and nonbreeding does. The affected glands are swollen and firm, and a clear to serosanguineous discharge can be expressed from the distended nipples. The glands do not seem painful, and the doe is not depressed. Epithelial hyperplasia, adenosis, papillomatous changes, and cystic mammary glands have been associated
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with uterine hyperplasia and adenocarcinoma.32,85 Cystic mammary glands may continue to progress and coalesce, with fibrous connective tissue accumulating around the cysts.85 Eventually, malignant cellular changes may occur, leading to invasive mammary adenocarcinoma. Metastasis to the regional lymph nodes, lungs, or other organs can occur with adenocarcinoma. Mammary adenocarcinomas are not uncommon in 3- to 4-year-old multiparous does. Evaluate rabbits with clinical signs of cystic mastitis for infectious mastitis and uterine disease. Fine-needle aspiration and cytology will assist with the diagnosis of mammary mass etiology. The treatment for noninfectious cystic mastitis is ovariohysterectomy; clinical signs usually resolve within 3 or 4 weeks after surgery. Partial mastectomy or wide excision of mammary tumors is indicated if malignant changes have occurred. The correlation between mammary and uterine disorders suggests that routine ovariohysterectomy of young, healthy does may reduce the risk of mammary neoplasms at a later age. One should also consider the fact that many antineoplastic medications are tested on rabbits and may provide a starting framework for dosages and regimens. Several cases of mammary dysplasia, associated with pituitary adenomas, have been described in older, primiparous New Zealand white rabbits.65 One or more mammary glands were swollen and firm, with enlarged and discolored teats. Histologically, changes were dysplastic in nature. At necropsy, prolactinproducing acidophil pituitary adenomas were found in affected does.65 Another case of hyperprolactinemia with mammary dysplasia (and the first report of cystic mammary adenocarcinoma associated with a prolactin-secreting pituitary adenoma) in a rabbit was also reported.73
DISORDERS OF THE URINARY SYSTEM Disorders of the urinary tract are relatively common in rabbits. Ten percent of pet rabbits presented to the University of Wisconsin Veterinary Medical Teaching Hospital over a 5-year period were diagnosed with urinary tract disease.62 A review of the data from the University of California-Davis Veterinary Medical Teaching Hospital reported a 3% incidence of renal insufficiency, acute renal failure, or chronic renal failure from January 1999 through January 2009 and a 2% incidence of nonhypercalciuria urolithiasis in presenting rabbits (JPM). Polyuria, polydipsia, incontinence, inappropriate elimination behavior, perineal urine scalding, hematuria, stranguria, and pollakiuria are common presenting complaints. Physical examination and careful abdominal palpation may reveal abnormalities in size, contour, and texture of the kidneys. Uroliths or excess granular material may be palpated within the urinary bladder. Results of a CBC and a serum biochemical profile may disclose changes consistent with acute or chronic renal insufficiency, but normal results do not rule this out. Urine voided onto a clean surface may yield useful information, but ideally urine should be collected by cystocentesis, taking care to avoid the voluminous intestinal tract. Lateral radiographs or ultrasound imaging may help to assess for the presence and size of a full urinary bladder. Catheterization of the urethra is easily performed in both male and female rabbits (Figs. 17-4 and 17-5). A 2% sterile lidocaine gel may reduce pain associated with passing the catheter and a low dose of midazolam is recommended for sedation. Alternatively, gentle manual pressure to the bladder often leads to micturition, and a midstream urine sample can be collected.
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Urinalysis is useful in assessing renal function and the presence of bacteria, proteinuria, glucosuria, ketonuria, and occult hematuria. Normal urine excretion is approximately 130 mL/kg/day and is the normal excretion route for calcium and magnesium.60 Creatinine is a more reliable test of renal function than blood urea nitrogen (BUN). Prerenal azotemia can be due to
Fig. 17-4 Female rabbit positioned for urinary catheter placement. Sedation may be helpful in some cases.
dehydration because of the limited ability of rabbits to concentrate urine. Hyperphosphatemia usually indicates chronic renal failure.54 The first urine of the morning is the best sample to assess. Clear urine indicates low calcium excretion, which may be pathologic if renal failure is present or physiologic in the growing or lactating rabbit. Glucosuria can occur with stress hyperglycemia and should be interpreted in light of the collection situation. Normal pH in rabbits tends to be basic (7.5-9.0). Normal rabbit urine has a low specific gravity (1.003-1.036), with prerenal azotemia suspected when associated with a higherthan-normal specific gravity and renal azotemia suspected when associated with a lower-than-normal specific gravity. Traces of protein can be found in normal urine; however, protein with isothenuria is significant.54 Triple phosphate and calcium carbonate crystals can normally be found in large quantities.60 Histologic lesions were observed in the kidneys of 77 (32.5%) of 237 rabbits that were either found dead or were euthanatized because of illness and in 19 (25%) of 75 apparently healthy adult rabbits.33 This suggests that renal disease is likely underdiagnosed and underreported. Lesions associated with an infectious process such as renal abscesses, staphylococcal nephritis, pyelonephritis, and pyelitis were the principal findings in rabbits up to 5 months of age, while renal fibrosis with or without dystrophic calcification was the most common lesion observed in rabbits aged over 10 months.33 Spontaneous amyloidosis was seen in rabbits, with amyloidosis being found only in the kidneys.33
Bladder 1
2 Urethra
A 3
Opening of urethra
Vagina
A Fig. 17-5 Illustration of the anatomic location of the urethral opening in the female rabbit. The rabbit’s vagina is long and the urethral opening is located at about half the length. The insert illustrates that the opening lies along the midline of the ventral aspect of the vagina under a thin flap of tissue. A catheter must be advanced along the floor of the vagina.
CHAPTER 17 Disorders of the Reproductive and Urinary Systems
UROLITHIASIS AND HYPERCALCIURIA Urolithiasis refers to the presence of calculi in the urinary system. Rabbits can have any combination of cystic, urethral, renal, and ureteral calculi. The cause of urolithiasis in rabbits is not fully understood, but several factors are involved, including nutrition, anatomy, and, rarely, infection. It is believed that rabbits have evolved to maximize absorption of dietary calcium, with the excess being normally excreted as calcium carbonate in the urine. Hypercalciuria is a clinical condition seen frequently in pet rabbits. Affected rabbits have a large amount of amorphous, pasty to slightly gritty calcium “sand” or “sludge” in their bladders (Fig. 17-6). Rabbits have an unusual calcium metabolism in that intestinal absorption of calcium is independent of vitamin D3 levels. Although the fractional urinary excretion of calcium is less than 2% in most mammals, the range for rabbits is 45% to 60%. Increases in dietary calcium directly increase its urinary excretion.10,13 This hypercalciuria also reflects the 30% to 50% higher calcemia seen in rabbits compared with other mammals.60 Phosphorus, however, is excreted in the feces.60 There are a number of theories as to why problematic hypercalciuria is common in the domestic rabbit, but little scientific investigation has been done to define the problem. Rabbits with either urolithiasis or hypercalciuria often have limited exercise, are fed a free-choice diet of pellets and alfalfa hay, and tend to be obese. Affected rabbits may have a history of vitamin or mineral dietary supplementation. Clinical signs of urolithiasis include depression, anorexia, weight loss, lethargy, hematuria, anuria, stranguria, a hunched posture, grinding of teeth, and urine scald of the perineum. Urolithiasis in rabbits may also be subclinical. Rabbits with hypercalciuria usually have thick, creamy urine. Often, voided urine appears only slightly turbid in these rabbits; however, with manual bladder expression, copious amounts of pasty urine are passed. Frequently, a
A
dough-like mass is palpated in the caudal abdomen, or a turgid bladder is evident if urethral obstruction is present. An enlarged kidney or ureter may be palpated and imaged with ultrasonography if hydroureter or hydronephrosis has occurred.49 There is a report of a female rabbit with a large cystic calculus wedged in the pelvic inlet that compressed and indirectly obstructed the bowel.78 Confirm the diagnosis of urolithiasis radiographically. A discrete calculus or the presence of homogenous radiopacity can be seen in the dependent portion of the bladder or completely filling a distended bladder. A small amount of sand in the bladder is a common incidental radiographic finding, especially in older animals, because the presence of amorphous calcium carbonate crystals is normal in rabbits. Close inspection may be needed to identify stones in the kidneys, ureters, or urethra. Perform an ultrasonographic examination to detect discrete calculi in a bladder that is distended and diffusely opaque radiographically. Multiple renal cysts may be confused with hydronephrosis; use ultrasonography to differentiate. If renal calculi are present (Fig. 17-7A), intravenous pyelography or renal scintigraphy (Fig. 17-7B) may be performed to evaluate renal function. Urinalysis may include crystalluria. Numerous calcium oxalate crystals are common, but ammonium phosphate, calcium carbonate, and monohydrate crystals are also frequently observed. Proteinuria and hematuria are common additional findings. If bacteria are noted on urine cytology, submit urine obtained by cystocentesis for culture. Pseudomonas species and Escherichia coli are bacteria known to cause cystitis. Results of a CBC and a serum biochemical profile assist in assessing renal function and in developing a prognosis. Treatment of urolithiasis depends on the location and severity of the lesion. Uroliths obstructing the urethra may be dislodged by catheterization of the urethra, performed carefully with a 3.5- or 5.0-Fr soft catheter of sufficient length to pass
B Fig. 17-6 Ventrodorsal (A) and lateral (B) radiographic views of a rabbit with a diffusely opaque urinary bladder filled with urinary sand (arrows).
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A
B
Fig. 17-7 A, Ventrodorsal radiograph of a 2-year-old female spayed rabbit with a large left renal calculus (arrow) conforming to the contour of the dilated renal pelvis. The calculus extends down the proximal ureter on the left. There is evidence of thinning of the adjacent renal cortex due to chronic obstruction of the left ureter. B, Scintigraphic image of the same rabbit. There is a photopenic area in the caudal aspect of the left kidney—also cumulatively more radiopharmaceutical uptake in the right kidney than the left (arrow). Relative filtrations were calculated. The left kidney contributes approximately 28% of total filtration and the right kidney approximately 72%. The rabbit underwent left nephrectomy based on the diagnostic information provided by radiology and nuclear imaging.
the length of the vagina (see Fig. 17-4) or penile urethra. As mentioned previously, catheterization is a painful procedure; therefore use topical 2% lidocaine gel to reduce pain associated with passing the catheter. Midazolam may provide sedation as well as relaxation of the urethral sphincter. Following catheterization and unblocking of the urethra, the critical supportive treatment for hypercalciuria is aggressive fluid therapy. Rarely is the urolith firmly lodged in the urethra, but in those cases surgery is required. Cystotomy is the treatment of choice for large cystic calculi. The procedure may be difficult because the neck of the bladder is flaccid and extends into the pelvic canal. Preanesthetic drugs such as benzodiazepines induce muscle relaxation and facilitate the passage of surprisingly large calculi into the urethra unintentionally. The location of the calculus should, therefore, be ascertained immediately preoperatively on radiographs. Attempts to force small stones out of the urethra during surgery may result in their becoming lodged within the neck of the bladder or in the urethra. Use a surgical spoon to retrieve small stones from the bladder. Flushing and gentle
surgical suction can aid in the removal of fine granular sand. Submit the calculi for analysis and obtain a swabbed sample of the bladder wall or biopsy a piece of bladder wall for bacterial culture. Perioperative support includes intravenous fluid diuresis, analgesia, and systemic antibiotics. Loop diuretics are not recommended. Subsequent radiographic monitoring is helpful because this condition often recurs. One of the present authors (EK) treated an older male rabbit with a 5-cm-diameter urolith lodged in a urethral diverticulum in the inguinal region, just cranial to the scrotal sacs. Surgical removal of the urolith and excision of the diverticula resolved the problem. No renal or cystic mineralization was noted on radiographs and the urine was normal both visually and on urinalysis. Nephrectomy may be indicated if a calculus in the renal pelvis has caused chronic obstruction and hydronephrosis. Only one case report describing nephrectomy has been published, though it references several personal communications of other nephrectomies.68 Contrast material (0.6 mL/kg) (Hypaque-76, Amersham Health, Princeton, NJ) can be given intravenously as a bolus to
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calcium concentration and the amount of calcium excreted in the urine.13,41 To reduce calcium intake, feed grass hay (not alfalfa), greens, and timothy hay-based pellets. Discontinue any vitamin or mineral supplementation; such supplements are not recommended in rabbits at any time. One investigation determined that a level of 0.22 g of calcium per 100 g of food was necessary for maximal growth12; commercial alfalfa-based rabbit pellets contain 0.9 to 1.6 g of calcium per 100 g of food.10 Consideration should be given to the hardness of drinking water, especially in polydipsic animals. Because many rabbits that develop urolithiasis are overweight, a decrease in total caloric intake and an increase in exercise may be helpful. Acidifiers are ineffective because rabbits are herbivores with naturally alkaline urine.
RENAL FAILURE
Fig. 17-8 Demonstration of the proper positioning of a catheterized rabbit for manual expression with gentle caudal pressure on the bladder. The urinary bladder is catheterized and sterile saline is retropulsed into the urinary bladder. It is mixed with the calcium “sand” material and finally the bladder contents are manually expressed to discharge the slurry. This procedure may have to be repeated several times. If the sandy urine flows around the catheter, administer a nonsteroidal anti-inflammatory agent.
perform a pyelographic evaluation.68 Radiographs are taken immediately after injection, with dye appearing quickly in the urinary bladder.68 A pyelolithotomy or nephrotomy may be considered if function of the affected kidney is still adequate and renal parenchymal damage is minimal. Extracorporeal shock wave lithotripsy has been experimentally applied to rabbit kidneys, but no information is available on a clinical application.42 Prognosis is guarded for rabbits with either unilateral or bilateral renal calculi. If hypercalciuria or nonobstructive calculi in the kidney or ureter are identified, consider a nonsurgical approach. Fluids administered either intravenously or subcutaneously are necessary to increase diuresis. Manually express the bladder daily for 2 to 4 days to encourage the passage of crystals or residual calcium sand that may not be voided during normal micturition. A technique similar to voiding that is used in dogs and cats, termed urohydropropulsion,52 can be used in rabbits for the nonsurgical removal of fine granular or small, smooth cystic calculi. Rabbits must be heavily sedated or anesthetized for this procedure. Administer preanesthetic benzodiazpines to help relax the urethralis muscle; additionally, opioids, such as butorphanol or buprenorphine, are recommended because bladder expression is painful. Pass a sterile catheter to administer sterile saline into the bladder, then hold the rabbit in an upright position so that the vertebral column is vertical and apply steady pressure to the bladder (Fig. 17-8). Hematuria is expected to occur for 1 to 2 days after urohydropropulsion.52 Dietary changes are an important part of treatment and prevention. The alkaline pH of rabbit urine and the high concentration of calcium in the urine increase the risk of precipitation of solutes.40 Decreasing dietary calcium levels directly lowers serum
Both acute and chronic renal failure can occur in older rabbits. Clinical signs include lethargy, depression, anorexia, polyuria, polydipsia, and perineal urine scalding. Acute renal failure is characterized by a sudden onset of filtration failure by the kidneys, accumulation of uremic toxins, and dysregulation of fluid, electrolytes, and acid-base balance. It is potentially reversible if diagnosed quickly and treated aggressively. In chronic renal failure there is a history of chronic loss of body condition, anemia, polydipsia, polyuria, and isosthenuria. One or both kidneys may be palpably small, with an irregular surface in chronic conditions. Serum creatinine and BUN levels are elevated in severe cases, as are serum concentrations of calcium, phosphorus, and potassium. Urine specific gravity can be isosthenuric, but this can also be seen in normal rabbits. Other findings from urinalysis may include proteinuria, hematuria, pyuria, and cast formation. Submit a urine sample for aerobic bacterial culture to screen for infectious causes; blood cultures are indicated if clinical signs of septicemia are concurrent. Indicators of inflammation in the urine (pyuria, proteinuria, hematuria) occur more frequently with acute renal failure and nephritis than with chronic renal failure. Acute renal failure has a better potential for response to treatment, but the prognosis remains guarded. Pyelonephritis in rabbits is often caused by P. multocida or Staphylococcus species Encephalitozoon cuniculi is a common cause of subclinical, chronic, interstitial nephritis. Noninfectious causes of chronic renal disease include hypercalcemia,27 renal calcinosis resulting from hypervitaminosis D,67 mineralization of the kidneys with interstitial fibrosis due to the excessive presence of vitamin D in the diet,87 and fatty degeneration in overweight animals. In rabbits with lymphosarcoma, renal involvement is common, often causing marked renomegaly and eventually resulting in renal failure from infiltrative destruction of the cortices.85 Treatment focuses on diuresis and diminishing the consequences of uremia; it is supportive only for chronic renal failure. Treatment of acute renal failure involves aggressive fluid therapy divided into three parts: (1) correction of perfusion, (2) correction of dehydration deficits, and (3) diuresis to correct azotemia, electrolyte, and acid-base status. Once the animal is normotensive and rehydrated, this is the polyuric or diuresis phase of acute renal failure. Measure the volume of urine produced every 4 hours by placing preweighed absorptive “pads” under the vulva or penis, weighing the pad every 4 hours, and estimating urine voided by assuming that 1 g equals 1 mL. In acute renal failure, urine production can be as high as 5 to 10 mL/kg per hour. The volume of fluid to be administered in each 4-hour period is the sum of the calculated maintenance requirements (2 mL/kg per hour)
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and voided urine volume from the previous interval. Additional ongoing losses (such as diarrhea) are added to the volume of administered fluids. Most mammals with acute renal failure become 3% to 5% dehydrated each day as a result of ongoing losses. Fluids are gradually discontinued when hydration and urine production are restored, blood urea nitrogen (BUN) and creatinine are normal, and the patient is eating and drinking. Tapering fluids by 50%/day will prevent medullary washout. Chronic renal insufficiency requires long-term subcutaneous fluid support. Rabbits with chronic renal failure usually develop anemia; therefore frequent reevaluation of CBC, chemistry profiles, and serum electrolytes is recommended. Dietary changes previously discussed for urolithiasis are also indicated. The calcium and phosphorus content of diet markedly influences parathyroid hormone (PTH) secretion in the uremic rabbit; when placed on a low calcium-high phosphorus diet, uremic rabbits develop secondary hyperparathyroidism.5 Masugi nephritis is an inflammatory nephritis seen in laboratory rabbits induced by continuous immunization with large doses of antigen. The condition results in elevations of BUN and creatinine. Tea polyphenols were suggested to be an efficacious treatment for this induced condition.24,86
HYPERVITAMINOSIS D Rabbits are sensitive to vitamin D toxicosis, and levels as low as five times normal may result in toxicity. Adults are more sensitive, though clinical signs of toxicity may take weeks to several years to become manifest. Pathologic changes consist of mineral deposition in various tissues, with arteries (especially the aorta) and kidneys being the most sensitive. Gross lesions in the kidneys consist of multifocal tan-to-gray areas located throughout the cortices. The exact pathophysiology of the soft tissue mineralization is unknown. Clinical signs are nonspecific, making early clinical diagnosis difficult. Since rabbits have high serum calcium concentrations normally, radiographic evidence of soft tissue mineralization, especially of the aorta and kidneys, is most helpful in making the diagnosis.24
NEPHROTOXICITY Several compounds are nephrotoxic in rabbits. Therapeutic use of gentamicin can result in acute tubular necrosis.21,44 Diuresis and supplemental doses (10 mg/kg SC) of vitamin B6 during gentamicin therapy may help protect the kidneys.21 The tiletamine in the tiletamine/zolazepam (Telazol, Fort Dodge Laboratories, Fort Dodge, IA) anesthetic combination is documented to cause nephrotoxicity in rabbits, inducing nephrosis at low doses and severe unreversible nephrosis at high doses.8,22 Rabbits have been used as laboratory animal models to study the renal responses to NSAIDs. Chronic exposure to therapeutic doses of an NSAID (diclofenac) produced significant ultrastructural renal alterations.77 Another study documented increased renal artery vasoconstrictor activity after NSAID administration.46 Rabbit studies support the premise that preexisting renal insufficiency increases the effect of NSAID-induced interstitial nephritis.14
RENAL ADIPOSE DEPOSITION Fatty infiltration of the kidney has been reported in does with pregnancy toxemia during the last week of pregnancy in primiparous, obese animals on high planes of nutrition that suddenly
become anorexic. Clinical pathologic abnormalities include ketosis, hypocalcemia, hyperphosphatemia, and fluctuating blood glucose levels. Fatty infiltration of the kidney with subsequent acute renal failure may also occur in conjunction with anorexia and the development of hepatic lipidosis.24,31
RENAL CYSTS The occurrence of multiple small, subcapsular cysts in the kidneys is an inherited condition of rabbits. The cysts are either of tubular origin or primitive ductules; the condition is similar to renal cortical dysplasia in humans.50 Although renal cysts do not create any clinical signs or alter the results of renal function tests, they may be detected on imaging studies or at necropsy. Out of 203 records with documented renal histopathology, seven cases of polycystic kidney syndrome (PKS) were found in a collection of white New Zealand rabbits by three morphologic criteria: (1) cysts or microcysts derived from tubules, glomeruli, or both; (2) loose mesenchymal expansion of cortical and/or medullary interstitium; and (3) irregular thickening, thinning, and splitting of basement membranes. PKS was associated with hypercalcemia, hypercreatinemia, and arterial mineralization. In the liver, mild chronic cholangitis with cholangiodysplasia and fibrosis was common. Anorexia and lethargy were the clinical signs most often reported. This condition had similarities to both autosomal dominant and autosomal recessive polycystic kidney diseases of humans.53
RENAL AGENESIS Renal agenesis, or congenital absence of one kidney, has been reported as an autosomal recessive mutation in one colony of laboratory rabbits.56 A high incidence of this condition has been reported in the Havana breed.50 The condition is seen in both sexes. In males, the ipsilateral testicle is often missing as well.50
ENCEPHALITOZOONOSIS Encephalitozoonosis, or nosematosis, is a common disease of rabbits caused by E. cuniculi, a microsporidian obligate, intracellular protozoan parasite. Transmission is by urine-oral passage, usually from doe to young. The organism is absorbed from the intestines into mononuclear cells and then distributed to other organs. Spores have a predilection for kidney, brain, and spinal tissue, where the most common lesions are found. Spores appear in the kidney 31 days after inoculation and are excreted in the urine up to 3 months after inoculation.65 Infections typically result in nonsuppurative, granulomatous nephritis that may progress to interstitial fibrosis, but the disease is chronic and usually subclinical.25 At necropsy, numerous small pits and stellate scars may be found on the cortical surface of the kidneys in rabbits with chronic encephalitozoonosis. Clinical signs of incontinence result from central nervous system infection rather than renal lesions. Diagnostic tests and therapeutic options for encephalitozoonosis are discussed in detail with diseases of the central nervous system. At the Animal Hospital of the Veterinary University of Vienna (Austria), among 144 serologically positive rabbits of a total of 184 with suspected encephalitozoonosis, 3.5% were found to have clinical signs of renal failure; 87.5% of these were moribund or euthanized. Polymerase chain reaction could not detect parasite DNA in urine or cerebrospinal fluid.47
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URINARY INCONTINENCE
TUMORS OF THE URINARY TRACT
Urinary incontinence can be caused by lumbosacral vertebral fractures and dislocations or by central nervous system lesions from E. cuniculi infection. Rabbits with urinary calculi or hypercalciuria often exhibit urinary incontinence and urine scalding. Clinical signs include a urine-soiled perineum and ulcerations of the vaginal mucosa and intertrigonal pouches as well as sticky, strong-smelling urine. A positive titer to E. cuniculi and additional central nervous system signs may suggest a protozoal infection. Other neurologic diseases, such as toxoplasmosis and larva migrans, should also be considered. Concurrent diarrhea with fecal caking may lead to urinary scalding. Urolithiasis or hypercalciuria can be identified radiographically. Vertebral fractures can be ruled out through neurologic examination and radiography. Ovariohysterectomized rabbits can develop urinary incontinence; this is responsive to diethylstilbestrol.11 Estrogen supplementation mediates a functional hypertrophy characterized by increased contractile responses to all forms of stimulation in both young and old rabbits that have undergone ovariohysterectomy. The increased contractile responses might be explained by the increases in vascular density and smooth muscle/collagen ratio.61 A positive response to 0.5 mg of diethylstilbestrol given orally once or twice weekly to spayed females suggests a hormone-responsive urinary incontinence similar to that seen in dogs. Treatment of rabbits with diethylstilbestrol or phenylpropanolamine is still under debate and investigation, with published results reflecting a scientific study not aimed at therapeutic relief. Additional differentials for urinary incontinence include ectopic ureter, urinary tract infection, neoplasia, and pyoderma. Initial supportive care includes daily cleaning of the perineum and topical treatment for dermatitis with a drying agent such as Domeboro astringent solution (Bayer HealthCare, Morristown, NJ) in addition to treatment of the primary problem. NSAIDs can be used if there is no evidence of renal insufficiency; they will decrease inflammation and provide analgesia. Older, obese, and/or arthritic rabbits may choose not to urinate in their litter box and appear suddenly “incontinent” to their owner. Using a litter box with easier access (low entry point) and keeping the box clean may be beneficial in these cases.
Benign embryonal nephromas are common in rabbits of all ages and are usually incidental findings at necropsy. In one report, an extremely large, palpable embryonal nephroma caused obliteration of a kidney and polycythemia.51 Lymphosarcoma in rabbits often involves the kidneys. Renal carcinoma and leiomyoma have also been reported.62 A triphasic nephroblastoma was diagnosed in a 1-year-old Angora rabbit.3
PSYCHOGENIC POLYURIA AND POLYDIPSIA Normal daily water intake for a rabbit is 120 mL/kg or more; urine output is approximately 130 mL/kg of body weight. Psychogenic polyuria/polydipsia has been observed by one of the authors (JPM) and been reported in four laboratory New Zealand white rabbits by excluding other potential causes through a set of diagnostic tests.66 The causes were not clearly identified, but in some affected rabbits, water consumption decreased after environmental enrichment measures were added, suggesting that boredom was a contributing factor.66
URINARY BLADDER EVERSION Transurethral eversion and prolapse of the urinary bladder has been documented in postparturient does shortly after kindling.30 Multiple kindlings at a young age, along with other undetermined factors, may predispose to the occurrence of this condition.30 Surgical reduction by laparotomy was successful in one Hotot doe.
RED URINE Rabbits can excrete porphyrin-pigmented urine that often incorrectly suggests hematuria. The urine may be dark brown, dark orange, or red. The unusual urine color is probably caused by a plant pigment and does not affect the rabbit’s health. Pigmented urine tends to be intermittent and lasts only 3 or 4 days. It is speculated that dietary compounds, ingestion of pine needles, or antibiotic administration may cause increased pigment levels in rabbit urine. Porphyrin pigments, but not hemoglobin, fluoresce when urine is examined under a Wood’s lamp. True hematuria is determined on examination of a urine sediment and the finding of more than five red blood cells per high-power field; a positive reaction for blood with the use of a urine dipstick also indicates hematuria or hemoglobinuria. Hematuria can originate from the genital tract or from the urinary tract. Blood from the reproductive tract can be associated with adenocarcinoma, polyps, abortion, or endometrial venous aneurysms. Blood originating from the urinary tract can be the result of cystitis, bladder polyps, pyelonephritis, renal infarcts, urolithiasis, or disseminated intravascular coagulation.26 Lead toxicosis can also cause hematuria, along with renal damage.36 Blood clots in the urine most likely originate from the genital tract and are particularly suggestive of endometrial venous aneurysms. History, signalment, physical examination, laboratory tests, radiography, and ultrasound examination are helpful in determining the cause. Hyperpigmented urine has also been associated with urobilinuria, which may appear similar to hematuria except that the test results are negative for blood and positive for urobilinogen.26 There is one documented report of a New Zealand white rabbit with porphyria.71 Porphyrias are a group of inherited or acquired disorders of certain enzymes in the “heme” biosynthetic pathway. No clinical signs were noted in the case reported, but necropsy findings included a pink tinge to the teeth, ultraviolet fluorescence of teeth and femurs, and increased uroporphyrin levels in the urine.71
References 1. Arvidsson A. Extra-uterine pregnancy in a rabbit. Vet Rec. 1998;142:176. 2. Asakawa MG, Goldschmidt MH, Une Y, et al. The immunohistochemical evaluation of estrogen receptor-alpha and progesterone receptors of normal, hyperplastic, and neoplastic endometrium in 88 pet rabbits. Vet Pathol. 2008;45:217-225. 3. Atasever A, Beyaz L, Deniz K. A case of triphasic nephroblastoma with lung metastases in an Angora rabbit. Revue Med Vet. 2007;158:303-308. 4. Baba N, von Haam E. Animal model: spontaneous adenocarcinoma in aged rabbits. Am J Pathol. 1972;68:653-656. 5. Bas S, Bas A, Estepa JC, et al. Parathyroid gland function in the uremic rabbit. Domest Anim Endocrinol. 2004;26:99-110.
230
SECTION II Rabbits
6. Beddow BA. Ectopic pregnancy in a rabbit. Vet Rec. 1999;144:624. 7. Bergdall VK, Dysko RC. Metabolic, traumatic, mycotic, and miscellaneous diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. New York: Academic Press; 1994:335-353. 8. Brammer DW, Doerning BJ, Chrisp CE, et al. Anesthetic and nephrotoxic effects of Telazol in New Zealand white rabbits. Lab Anim Sci. 1991;41:432-435. 9. Bray MV, Weir EC, Brownstein DG, et al. Endometrial venous aneurysms in three New Zealand white rabbits. Lab Anim Sci. 1992;42:360-362. 10. Buss SL, Bourdeau JE. Calcium balance in laboratory rabbits. Miner Electrolyte Metab. 1984;10:127-132. 11. Caslow D. Hormone responsive perineal urine soiling in two female ovariohysterectomized rabbits. Comp Anim Pract. 1989;19:32-33. 12. Chapin RE, Smith SE. Calcium requirement of growing rabbits. J Anim Sci. 1967;26:67-71. 13. Cheeke PR, Amberg JW. Comparative calcium excretion by rats and rabbits. J Anim Sci. 1973;37:450-454. 14. Chen CY, Pang VF, Chen CS. Pathological and biochemical modifications of renal function in ibuprofen-induced interstitial nephritis. Ren Fail. 1996;18:31-40. 15. Cunliffe-Beamer TL, Fox RR. Venereal spirochetosis of rabbits: eradication. Lab Anim Sci. 1981;31:379-381. 16. David C, Grimminger S, Wehrend A. Ectopic pregnancy in a miniature rabbit. Tieraerz Praxis. 2006;34:197-200. 17. DeLong D, Manning PJ. Bacterial diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. New York: Academic Press; 1994:131-170. 18. DiGiacomo RF, Deeb BJ, Anderson RJ. Hypervitaminosis A and reproductive disorders in rabbits. Lab Anim Sci. 1992;42:250-254. 19. DiGiacomo RF, Talburt CD, Lukehart SA, et al. Treponema paraluis-cuniculi infection in a commercial rabbitry: epidemiology and serodiagnosis. Lab Anim Sci. 1983;33:562-566. 20. Elsinghorst TA, Timmermans HJE, Hendriks HG. Comparative pathology of endometrial carcinoma. Vet Q. 1984;6:200-208. 21. Enriquez Sr JI, Schydlower M, O’Hair KC, et al. Effect of vitamin B6 supplementation on gentamicin nephrotoxicity in rabbits. Vet Hum Toxicol. 1992;34:32-35. 22. Evans KD, Dillehay DL, Huerkamp MJ, et al. Diagnostic exercise: azotemia in a rabbit. (Oryctolagus cuniculus). Lab Anim Sci. 1996;46:442-443. 23. Fekete SG, Hullar I, Romvari R, et al. Study of the energy and protein balance of pregnant rabbit does using two comparative methods. Acta Vet Hung. 2005;53:435-447. 24. Fisher PG. Exotic mammal renal disease: causes and clinical presentation. Vet Clin North Am Exot Anim Pract. 2006;9:33-67. 25. Flatt RE, Jackson SJ. Renal nosematosis in young rabbits. Pathol Vet. 1970;7:492-497. 26. Garibaldi BA, Fox JG, Otto G, et al. Hematuria in rabbits. Lab Anim Sci. 1987;37:769-772. 27. Garibaldi BA, Goad ME. Hypercalcemia with secondary nephrolithiasis in a rabbit. Lab Anim Sci. 1988;38:331-333. 28. Gil PS, Palau BP, Martinez JM, et al. Abdominal pregnancies in farm rabbits. Therio. 2004;62:642-651. 29. Goto M, Nomura Y, Une Y, et al. Malignant mixed mullerian tumor in a rabbit (Oryctolagus cuniculus): case report with immunohistochemistry. Vet Pathol. 2006;43:560-564. 30. Greenacre CB, Allen SW, Ritchie BW. Urinary bladder eversion in rabbit does. Compend Contin Educ Pract Vet. 1999;21:524-528. 31. Harcourt-Brown FM. Urogenital disease. In: Textbook of rabbit medicine. Oxford (UK): Butterworth Heinemann. 2002:335-351. 32. Hillyer EV. Pet rabbits. Vet Clin North Am Small Anim Pract. 1994;24:25-65.
33. Hinton M. Kidney disease in the rabbit: a histological survey. Lab Anim. 1981;15:263-265. 34. Hobbs BA, Parker RF. Uterine torsion associated with either hydrometra or endometritis in two rabbits. Lab Anim Sci. 1990;40:535-536. 35. Hofmann JR, Hixson CJ. Amyloid A protein deposits in a rabbit with pyometra. J Am Vet Med Assoc. 1986;189:1155-1156. 36. Hood S, Kelly J, McBurney S, et al. Lead toxicosis in 2 dwarf rabbits. Can Vet J. 1997;38:721-722. 37. Ingalls TH, Adams WM, Lurie MB, et al. Natural history of adenocarcinoma of the uterus in the Phipps rabbit colony. J Natl Cancer Inst. 1964;33:799-806. 38. Jin L, Valentine BA, Baker RJ, et al. An outbreak of fatal herpesvirus infection in domestic rabbits in Alaska. Vet Pathol. 2008;45:369-374. 39. Johnson JH, Wolf AM. Ovarian abscesses and pyometra in a domestic rabbit. J Am Vet Med Assoc. 1993;203:667-669. 40. Kamphues J, Carstensen P, Schroeder D, et al. Effects of increasing calcium and vitamin D supply on calcium metabolism of rabbits. J Anim Physiol A Anim Nutr. 1986;56:191-208. 41. Kamphues J. Calcium metabolism of rabbits as an etiological factor for urolithiasis. J Nutr. 1991;121:S95-S96. 42. Karalezli G, Gogus O, Beduk Y, et al. Histopathologic effects of extracorporeal shock wave lithotripsy on rabbit kidney. Urol Res. 1993;21:67-70. 43. Kaufmann-Bart M, Fischer I. Choriocarcinoma with metastasis in a rabbit (Oryctolagus cuniculus). Vet Pathol. 2008;45:77-79. 44. Kojima T, Kobayashi T, Iwase S, et al. Gentamicin nephrotoxicity in young rabbits. Exp Pathol. 1984;26:71-75. 45. Kraus AL, Weisbroth SH, Flatt RE, et al. Biology and disease of rabbits. In: Fox IE, Cohen B, Loew FM, eds. Laboratory animal medicine. New York: Academic Press; 1984:207-237. 46. Kristova V, Djibril NM, Fackovcova D, et al. Comparison of vasoconstrictor responses to selected NSAIDs in rabbit renal and femoral arteries. Bratisl Lek Listy. 2002;103:50-53. 47. Künzel F, Gruber A, Tichy A, et al. Clinical symptoms and diagnosis of encephalitozoonosis in pet rabbits. Vet Parasitol. 2008;151:115-124. 48. Kurotaki T, Kokoshima H, Kitamori F, et al. A case of adenocarcinoma of the endometrium extending into the leiomyoma of the uterus in a rabbit. J Vet Med Sci. 2007;69:981-984. 49. Lee KJ, Johnson WD, Lang CM, et al. Hydronephrosis caused by urinary lithiasis in a New Zealand white rabbit. (Oryctolagus cuniculus). Vet Pathol. 1978;15:676-678. 50. Lindsey JR, Fox RF. Inherited diseases and variations. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. New York: Academic Press; 1994:293-319. 51. Lipman NS, Murphy JC, Newcomer CE. Polycythemia in a New Zealand white rabbit with an embryonal nephroma. J Am Vet Med Assoc. 1985;187:1255-1256. 52. Lulich JP, Osborne CA, Carlson M, et al. Nonsurgical removal of urocystoliths in dogs and cats by voiding urohydropropulsion. J Am Vet Med Assoc. 1993;203:660-663. 53. Maurer KJ, Marini RP, Fox JG, et al. Polycystic kidney syndrome in New Zealand white rabbits resembling human polycystic kidney disease. Kidney Int. 2004;65:482-489. 54. Melillo A. Rabbit clinical pathology. J Exot Pet Med. 2007;16:135-145. 55. Morrell JM. Hydrometra in the rabbit. Vet Rec. 1989;125:325. 56. Nath A, Juyal R, Venkatesan R, et al. Renal agenesis in New Zealand white rabbit. Scand J Lab Anim Sci. 2006;33:197-200. 57. National Academy of Sciences. The National Research Council: Nutrient requirements of domestic animals: nutrient requirements of rabbits. 2nd ed. Washington, DC: National Academy of Sciences; 1992. 58. Ness RD. Neoplasia in rabbits and guinea pigs. Proceedings. North Am Vet Conf. 1998:853-854.
CHAPTER 17 Disorders of the Reproductive and Urinary Systems 59. Okerman L. Diseases of domestic rabbits. 2nd ed. London: Blackwell Scientific Publications; 1994. 60. O’Malley B. Rabbits. Clinical anatomy and physiology of exotic species. Philadelphia: Elsevier Saunders; 2005:173-195. 61. Onal B, Levin RM, Kogan BA, et al. The effect of maturation and age on oestrogen-induced functional hypertrophy of the female rabbit bladder. BJU Int. 2007;99:674-679. 62. Pare JA, Paul-Murphy J. Disorders of the reproductive and urinary systems. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis, MO: Elsevier; 2003:183-193. 63. Patton NM. Colony husbandry. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. New York: Academic Press; 1994:27-45. 64. Patton NM, Holmes HT, Cheeke PR. Hairballs and pregnancy toxemia. J Appl Rabbit Res. 1983;6:99. 65. Percy DH, Barthold SW. Pathology of laboratory rodents & rabbits. 2nd ed. Ames, lA: Iowa State University Press; 2001. 66. Potter MP, Borkowski GL. Apparent psychogenic polydipsia and secondary polyuria in laboratory housed New Zealand white rabbits. Contemp Top Lab Anim Sci. 1998;37:87-89. 67. Quimby F, Foote R, Profit-Olstad M, et al. Hypercalcemia, hypercalcitoninism, and arterial calcification in rabbits fed a diet containing excessive vitamin D and calcium. Lab Anim Sci. 1982;32:415. 68. Rhody JL. Unilateral nephrectomy for hydronephrosis in a pet rabbit. Vet Clin North Am Exot Anim Pract. 2006;9:633-641. 69. Rommers JM, Boiti C, De Jong I, et al. Performance and behaviour of rabbit does in a group-housing system with natural mating or artificial insemination. Repro Nutr Dev. 2006;46: 677-687. 70. Saito K, Tagawa M, Mimura M, et al. Experimental transmission of rabbit syphilis. J Vet Med Sci. 2005;67:79-81. 71. Samman S, Fussell SH, Rose CI. Porphyria in a New Zealand white rabbit. Can Vet J. 1991;32:622-623. 72. Segura P, Martinez J, Peris B, et al. Staphylococcal infections in rabbit does on two industrial farms. Vet Rec. 2007;160: 869-872. 73. Sikoski P, Trybus J, Cline JM, et al. Cystic mammary adenocarcinoma associated with a prolactin-secreting pituitary adenoma in a New Zealand white rabbit (Oryctolagus cuniculus). Comp Med. 2008;58:297-300.
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74. Soave OA, Dominguez J, Doak RL. Moraxella bovis-induced metritis and septicemia in a rabbit. J Am Vet Med Assoc. 1977;171:972-973. 75. St Claire MB, Kennett MJ, Besch-Williford CL. Vitamin A toxicity and vitamin E deficiency in a rabbit colony. Contemp Top Lab Anim Sci. 2004;43:26-30. 76. Stott P, Wight N. Female reproductive tract abnormalities in European hares (Lepus europaeus) in Australia. J Wild Dis. 2004;40:696-703. 77. Taib NT, Jarrar BM, Mubarak MM. Ultrastructural alterations in renal tissues of rabbits induced by diclofenac sodium (Voltaren). Saudi Med J. 2004;25:1360-1365. 78. Talbot AC, Ireton VJ. Unusual cause of intestinal blockage in the female rabbit. Vet Rec. 1975;96:477. 79. Theau-Clement M. Preparation of the rabbit doe to insemination: A review. World Rabbit Sci. 2007;15:61-80. 80. Thode HP, Johnston MS. Probable congenital uterine developmental abnormalities in two domestic rabbits. Vet Rec. 2009;164:242-244. 81. Tirpude RJ, Jain R, Tuteja U, et al. Isolation, identification and characterization of Staphylococcus aureus from systemic infection in New Zealand white rabbits. Indian J Anim Sci. 2007;77:207-210. 82. Van Herck H, Hesp APM, Versluis A, et al. Prolapsus vaginae in the IIIVO/JU rabbit. Lab Anim. 1989;23:333-336. 83. von Bomhard W, Mauldin EA, Pfleghaar S. Disseminated collagenous hamartomas in rabbits—a new entity. Kleintierpraxis. 2008;53:224-230. 84. Watson GL, Evans MG. Listeriosis in a rabbit. Vet Pathol. 1985;22:191-193. 85. Weisbroth SH. Neoplastic diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. New York: Academic Press; 1994:259-292. 86. Xiu-Fang H, Ji-shuang C, Xian-qiang Y. Effect of tea polyphenols on masugi nephritis of rabbit. Pakistan J Biol Sci. 2002;5:784-788. 87. Zimmerman TE, Giddens WE, DiGiacomo RF, et al. Soft tissue mineralization in rabbits fed a diet containing excess vitamin D. Lab Anim Sci. 1990;40:212-215. 88. Zwicker GM, Killinger JM. Interstitial cell tumors in a young adult New Zealand white rabbit. Toxicol Pathol. 1985; 13:232-235.
CHAPTER
18
Dermatologic Diseases
Laurie Hess, DVM, Diplomate ABVP (Avian), and Kathy Tater, DVM, Diplomate ACVD
Bacterial Infections Subcutaneous Abscesses Mastitis Methicillin-Resistant Staphylococcal Infection Cellulitis Moist Dermatitis Ulcerative Pododermatitis Rabbit Syphilis Necrobacillosis Fungal Infections Dermatophytosis Parasitic Infections Ear Mites Fur Mites Fleas Myiasis Ticks Lice Pinworms Tapeworm Cysts Viral Infections Myxomatosis Rabbit (Shope) Fibroma Virus Rabbit (Shope) Papillomavirus Oral Papillomavirus Rabbitpox Cutaneous Neoplasia Behavioral Causes of Skin Disease Barbering Self-Mutilation after Intramuscular Injection Skin Disease of Unknown Cause Sebaceous Adenitis Ehlers-Danlos-Like Syndrome Dermal Fibrosis Eosinophilic Granuloma
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BACTERIAL INFECTIONS SUBCUTANEOUS ABSCESSES Subcutaneous abscesses commonly occur in rabbits from traumatic wounds or bacteremia secondary to tooth root infection, oral foreign bodies, or upper respiratory tract or urinary tract infections. Frequently, no direct cause is identifiable.22,27,28,33 Abscesses usually are soft to firm swellings that gradually enlarge over days to weeks. Although abscesses most commonly develop on the head and limbs, they may occur anywhere on the body (Fig. 18-1). They usually are not painful, are frequently immovable, are minimally inflamed, contain caseous exudate, and are typically walled off by a thick capsule.19 Abscesses may be confined to the subcutaneous space, or they may extend to underlying dermis and bone. Facial abscesses are often associated with underlying dental or nasolacrimal duct disease. Periapical abscesses are common and are typically associated with elongated cheek teeth roots. Superficial abscesses may occur as a result of fight wounds or penetrating injuries. Affected rabbits may have no other clinical signs, or they may be inappetant, lose weight, drool (with oral abscesses), or become lame (with limb abscesses). Diagnose subcutaneous abscesses by palpation and oral examination. Aspirate subcutaneous swellings with a 22-gauge or larger needle to obtain samples for cytologic evaluation, Gram’s stain, and both aerobic and anaerobic bacterial culture and sensitivity testing.27,28 Organisms most commonly isolated from rabbit abscesses include Staphylococcus aureus, Pasteurella multocida, Pseudomonas aeruginosa, Proteus species, Fusobacteria species, Bacteroides species, and Actinomyces species.19,20,32,41,58 Samples for bacterial culture and sensitivity testing are always recommended for recurrent abscesses because bacteria can become resistant to antibiotics.22 Despite the presence of bacteria within abscesses, cultures of abscess aspirates are sometimes negative for bacterial growth. Swabs taken from the inner lining of an abscess cavity are less likely to yield negative bacterial culture results than those taken from the purulent contents. Negative growth on aerobic culture may also occur if the primary pathogen is an anaerobic bacterial species. In addition Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 18 Dermatologic Diseases
Fig. 18-1 Cutaneous abscess on the forelimb of a rabbit. (Courtesy of Jan Declercq, DVM.)
to bacterial cultures, obtain radiographs to determine if the underlying bone is affected. For abscesses on the head, both thoracic and skull radiographs should be obtained because thoracic radiographs can reveal pneumonia or pulmonary abscessation. Ultrasonography and computed tomography may help delineate abscess margins and are especially useful with retrobulbar abscesses (see Chapter 35). A blood sample for a complete blood count (CBC) and plasma biochemical analysis and a urine sample for urinalysis are all indicated in the diagnostic workup of patients with subcutaneous abscesses. Treatment depends on the location and extent of the abscess. Complete surgical excision of the abscess en bloc, followed by at least 2 weeks of antibiotic administration based on results of culture and sensitivity testing, is ideal.* If joints are affected, limb amputation may be necessary. With retrobulbar abscesses, enucleation may be warranted (see Chapter 33). All inciting causes, such as foreign bodies or dental disease, should be addressed to minimize the recurrence of the abscess. If en bloc excision including the abscess capsule is not possible, debride all infected soft tissue and bone. Because the purulent content is usually thick, rabbit abscesses cannot be treated effectively with drains. If any infected tissue remains after abscess debridement, recurrence is likely because antibiotics do not penetrate either the thick capsule or its caseous contents easily. If complete excision is impossible, abscessed tissue should be debrided, flushed with sterile saline solution twice a day, and the remaining pocket allowed to heal by second intention. Any part of the capsule that can be removed should be removed to minimize contamination of unaffected tissue with infected material. Abscesses have been successfully treated by packing the completely debrided abscess cavity with antibiotic-impregnated polymethylmethacrylate beads (PMMA), bone cement, or a synthetic polymer.16,19,33,41 Beads slowly release antibiotics over days to weeks, providing high local tissue concentrations of drugs with low systemic levels and few systemic side effects (see Chapter 32). Commonly used heat-stable antibiotics in beads include cephalothin (2 g/20 g PMMA), cefazolin (2 g/20 g PMMA), amikacin (1.25 g/20 g PMMA), gentamicin (1 g/20 g PMMA), and tobramycin (1 g/20 g PMMA).74 While rabbits should not be given most cephalosporins orally owing to the risk of developing fatal enterotoxemia, they generally tolerate these drugs in beads because of the slow release and localized
*References 16, 19, 22, 27, 28, 32, 33, 80.
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concentration.16,19 Be careful not to place beads in abscess pockets that communicate with the oral cavity, or rabbits may ingest them. Beads may be left in place for weeks and removed at a subsequent surgery. Beads may be left intact permanently, but they may eventually act as a nidus for further abscess formation and have to be removed later. Large abscesses may require multiple surgeries to treat. If contaminated abscess cavities must be temporarily closed to retain packed beads, use only nonabsorbable monofilament sutures because they are less likely to induce an intense tissue response than absorbable or multifilament suture. Close uncontaminated wounds with absorbable monofilament suture. Older references mention the application of calcium hydroxide powder to abscess pockets after debridement to alkalinize the local wound environment and limit bacterial growth. Calcium hydroxide powder is no longer recommended owing to the severe soft tissue necrosis it can induce.16,19 Some authors advocate application of honey or 50% dextrose solution with or without topical antibiotics to the abscess cavity after debridement.16,74 This hyperosmotic solution may dry out the affected area, provide antibacterial effects through acidification of local tissue, and stimulate wound healing. It may also encourage the rabbit to lick the area and thus promote drainage. Jaw abscesses are particularly difficult to treat. These abscesses may result from periapical disease causing abnormal tooth growth and destruction of surrounding bone.58 Dental abscesses often fistulate through underlying tissue to teeth roots. Bacteria commonly isolated from these abscesses are Fusobacterium, Prevotella, Peptostreptococcus, Actinomyces, and Arcanobacterium species.19,67,68 To treat dental abscesses, curette and flush necrotic tissue (including infected bone), extract infected teeth, and pack the remaining pocket with antibiotic-impregnated PMMA beads or a synthetic bone graft particulate (Consil, Nutramax Laboratories, Baltimore, MD) (see Chapter 32). Alternatively, marsupialize the debrided abscess pocket open to surrounding skin to allow daily flushing until a clean wound pocket is achieved. In addition, CO2 laser ablation has been used successfully in cases of recurrent mandibular abscessation.19 Regardless of how the wound is treated, choose systemic antibiotic therapy based on results of culture and sensitivity testing and administer antibiotics postoperatively for at least 2 weeks and up to 6 weeks or longer. While cultures are pending, administer a penicillin G benzathine/penicillin G procaine combination at a dosage of 75,000 U SC every other day for rabbits weighing less than 2.5 kg and 150,000 U SC every other day for rabbits weighing more than 2.5 kg.61 Combination treatment with enrofloxacin (5 mg/kg PO q12h) and metronidazole (30 mg/kg PO q12h) also provides broad-spectrum treatment until culture results are obtained.74 For rabbits with recurrent abscesses, lifelong antibiotic treatment and repeated surgeries may be necessary.28 The long-term prognosis for rabbits with abscesses is improved with proper husbandry, including good sanitation and a high-fiber diet.27
MASTITIS Rabbit mastitis is most common in heavily lactating does.28,41 It may be caused by trauma to the teats with secondary bacterial infection from environmental contamination. The most commonly isolated organism in rabbits with suppurative mastitis is coagulase-positive Staphyloccus aureus. Other bacteria
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that cause mastitis include Pasteurella and Streptococcus species. Mammary glands are typically swollen, indurated, and sometimes abscessed. Teats initially are pink from hyperemia and then turn blue from vascular stasis (hence the term, blue breast).19 Does become depressed, anorectic, septicemic, and febrile; they often die. Rabbits with mastitis should be treated with intravenous fluids and antibiotics, hot packing of infected teats, and surgical debridement of abscessed mammary glands Any affected animals should be isolated to prevent the spread of infection. Remove kits to prevent them from starving or contracting bacterial enteritis.65 Restrict contact with other carriers of S. aureus, including people, to minimize the risk of infection.55 Nonseptic cystic mastitis can develop in nonbreeding does (see Chapter 17).28 Mammary glands are swollen and firm, with discolored, distended nipples exuding clear to brown discharge. Glands usually are not painful and does usually are not systemically ill. This condition is often associated with uterine hyperplasia and adenocarcinoma. Treatment requires an ovariohysterectomy. In young rabbits, S. aureus also causes exudative dermatitis with multiple raised pustules on the legs, head, back, and sides. One virulent biotype can cause high mortality of juvenile rabbits in rabbitries due to septicemia.19,55 Diagnosis and treatment is as for suppurative mastitis.
CELLULITIS Cellulitis in rabbits usually occurs acutely and may develop secondary to respiratory tract infection.3,27,32,33 Affected rabbits have fevers of 104° to 108°F (40°-42.2°C), and the skin around the head, neck, and chest becomes painful, inflamed, and edematous. Bacteria commonly isolated from these lesions include S. aureus, P. multocida, and Bordetella bronchiseptica. In these cases, treat affected rabbits with parenteral antibiotics (enrofloxacin, beta lactams, and aminoglycosides) based on the results of culture and sensitivity testing, topical antiseptics (1% chlorhexidine or 10% povidone-iodine), and cool baths. In rabbits that survive, lesions may become necrotic eschars or abscesses requiring surgical debridement.
MOIST DERMATITIS
Drug-resistant staphylococcal infections have been reported in rabbits. Methicillin-resistant S. aureus (MRSA) became a serious concern in human medicine during the late 1970s and has been reported frequently in animals over the past 10 years.45,60,69 MRSA strains are resistant to all penicillins, cephalosporins, and carbapenems; many are also resistant to aminoglycosides, macrolides, lincosamide, streptomycins, tetracycline, chloramphenicol, fluorquinolones, and rifampicin.76 Methicillin resistance also occurs in other staphylococcal species such as Staphylococcus intermedius and Staphylococcus schleiferi. Numerous studies indicate potential transfer of these bacteria between humans and animals and vice versa, and evidence increasingly demonstrates that isolates from dogs and cats are indistinguishable from those found in human health care facilities.40,43,70 Research also suggests that veterinary hospital staff and hospitalized pets can transit MRSA back and forth.17,64,66,78,79 MRSA has been isolated in companion animals and livestock; pigs, in particular, have found to be a source of human infection.* Few studies have focused on the prevalence of MRSA in exotic pets. MRSA has been isolated in chinchillas and parrots, and another report cites MRSA in a pet rabbit and a guinea pig.77 MRSA was also isolated from a rabbit with bilateral facial dermatitis and conjunctivitis (K. Quesenberry, personal communication, 2010). The widespread nature of these “superbug” infections and their potential transmission between humans and animals necessitates awareness of this organism as another possible cause of dermatitis in rabbits. In any case of dermatitis in a rabbit that does not respond to empiric antibiotic therapy, submit a sample for bacterial culture and sensitivity testing.
In rabbits, moist dermatitis usually develops on the chin, dewlap, or ventral neck (“slobbers”) or perianally as urine scald (“hutch burn”).* Chin or neck lesions can result from excessive drooling secondary to dental disease or from a constantly wet dewlap in rabbits that drink out of dirty water bottles or bowls. In addition, some rabbits chew their dewlaps as a displacement behavior for pain. Urine scald can be associated with excessive urination or urinary incontinence from renal disease (including renal infection with Encephalitizoon cuniculi), cystitis, urolithiasis, or urine “sludge.” Urine scald may also occur with immobility from posterior paresis, ulcerative pododermatitis (see below), obesity, genital infection with Treponema cuniculi (see below), vaginal discharge from uterine disease, or limited cage space/damp bedding. Rabbits that do not groom themselves may develop perianal and inguinal fur mats that absorb urine and trap feces, resulting in a moist dermatitis. With both perianal and ventral neck dermatitis, the skin may be inflamed, alopecic, ulcerated, necrotic, and fly-infested. Commonly, P. aeruginosa causes secondary dewlap infection and produces pyocyanin pigment that turns the skin blue-green; hence this condition is sometimes referred to as “blue fur disease.”19,80 To determine any underlying cause for moist dermatitis, take skull or abdominal radiographs and submit samples for urinalysis, a CBC, and a plasma biochemical analysis. Submit swab samples of exudate or skin for bacterial culture and sensitivity testing. Do a cytologic examination of a skin swab sample to look for yeast and the type of bacteria present (cocci or rods) to guide treatment pending culture results. Treat underlying diseases and correct associated environmental factors. Clip the hair over the affected skin and allow the area to dry. Drying agents, such as Domeboro Powder (Bayer Inc., Elkhart, IN) or dilute topical chlorhexidine, may help. Administer a systemic antibiotic while awaiting the results of culture and sensitivity testing and provide analgesics and/or nonsteroidal anti-inflammatory drugs (NSAIDs) for pain. Compounded topical preparations containing antibiotics, antifungal agents, and NSAIDs in combination, based on cytologic and bacterial culture results, can be helpful; consult with a veterinary dermatologist for recommendations. If obesity is a factor, ensure that the diet is high in fiber and low in calcium and encourage exercise. Keep affected rabbits indoors to prevent myiasis.16 Urine scald in grossly obese rabbits may require surgical removal of excess skin folds
*References 35, 36, 43, 52, 71, 84.
*References 3, 20, 27, 28, 32, 33, 41, 55, 65, 74.
METHICILLIN-RESISTANT STAPHYLOCOCCAL INFECTION
CHAPTER 18 Dermatologic Diseases that interfere with urination; a more conservative approach is to restrict food and encourage exercise to promote weight loss.
ULCERATIVE PODODERMATITIS Ulcerative pododermatitis (sore hocks) is a chronic, granulomatous, ulcerative dermatitis of the plantar metatarsal and, occasionally, volar metacarpal and phalangeal surfaces of the feet. This condition usually results in avascular necrosis of the plantar foot surfaces from trauma due to rough, dirty floors or frequent thumping.19,73 It may also develop in rabbits with thin plantar fur pads (such as Rex rabbits, which lack protective guard hairs), in rabbits with altered weight bearing due to spondylosis, or in large-breed or obese rabbits as a result of ischemia and pressure necrosis from confinement in small cages with concrete or wire floors.* The plantar surfaces of rabbits housed on rough carpet, tile, or vinyl also are exposed to increased frictional forces, predisposing them to pododermatitis.19 Rabbits kept on inappropriate bedding shift their weight off their claws and the plantar surfaces of their feet and onto their metatarsi and hocks. This shift causes lesions to develop that start as erythematous decubital ulcers and then become secondarily infected, usually with S. aureus or P. multocida, from contaminated bedding. Infected ulcers progress to abscesses covered with raised, dry, hyperkeratotic fibrotic scabs. Infection can spread to underlying bone and ligaments, causing osteomyelitis and sepsis. Osteomyelitis and synovitis may lead to superficial flexor tendon displacement. This forces rabbits to shift weight permanently onto their hocks and away from their digits, thus exacerbating the condition and worsening the long-term prognosis. In addition, chronic bleeding from ulcerated feet can lead to anemia. To treat pododermatitis, submit samples of abscessed tissue for bacterial culture and sensitivity testing and take radiographs of the legs to check for osteomyelitis.22 Clip ulcerated areas free of hair but be careful not to clip away hair surrounding ulcers, as the hairs may actually relieve pressure on the ulcer itself.19 Clean wounds with warm saline, lactated Ringer’s solution, or dilute antiseptics such as 0.05% chlorhexidine and debride necrotic tissue. Consider packing severe abscesses with antibiotic-impregnated PMMA beads or honey (see above). Surgical debridement is often very difficult or impossible because very little extra skin is present on the plantar metatarsus for wound closure.16,74 Postoperatively, bandage open wounds left to heal by second intention or delayed primary closure with light dressings. A variety of nonadherent semi-occlusive to occlusive dressings are available for wound treatment in small animals; Tegaderm (3M Medical-Surgical Division, St. Paul, MN), Telfa pads (The Kendall Co., Mansfield, MA), and other wound dressings have been used successfully in several rabbits. Protect superficial wounds with liquid bandage preparations (New Skin, Prestige Brands, Irvington, NY), surgical glue painted on topically, or cyanoacrylate skin protectants (Marathon Liquid Skin Protectant, Medline Industries Inc., Mudelein, IL). Alleviate pressure at ulcerated sites by applying donut-shaped bandages with the ulcer in the center of the donut.19 Regardless of the type of dressing used, change bandages daily to prevent reinfection from feces and urine. Additionally, a variety of topical agents that promote wound healing and epithelialization may be applied to aid in wound closure.16 Application of water-based,
*References 15, 16, 19, 28, 32, 33, 74, 80.
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topical antibiotics, such as silver sulfadiazine (Silvadene Cream 1%, Monarch Pharmaceuticals, Bristol, TN) or a nontoxic antimicrobial product (Vetericyn, Innovacyn, Inc., Rialto, CA), and systemic antibiotics, selected based on results of culture and sensitivity testing, usually is warranted. Avoid using petroleumbased topical antibiotics because these are more likely to cause local allergic reactions and rabbits may ingest them. In addition, avoid using topical steroids in rabbits, as they may cause immunosuppression and delayed wound healing. Change the environment to nonabrasive, solid cage floors and clean, soft, dry bedding such as towels, cellulose fiber bedding (CareFRESH, Absorption Corp., Ferndale, WA), or clean hay.16 Provide analgesia with NSAIDs and/or opioids. Increase the fiber and lower the carbohydrates in the diets of obese rabbits and encourage exercise. Unilateral pododermatitis that is unresponsive to treatment may require amputation at the midfemur. Bilateral severe pododermatitis involving osteomyelitis and infection of underlying tendons or ligaments may warrant euthanasia.
RABBIT SYPHILIS Rabbit syphilis, also called venereal spirochetosis or “vent disease,” is a nonzoonotic, contagious venereal disease caused by the spirochete Treponema paraluiscuniculi (see also Chapter 17).16,54,55,63,80 Transmission is through direct contact with infected skin or from infected dam to kits at birth.3,22,28 Intrauterine transmission is not reported and there is an age-related susceptibility, with some young rabbits seeming relatively resistant to certain strains.53 Incubation periods are long, lasting up to 10 to 16 weeks.65 Lesions in affected, typically juvenile rabbits may appear as nonpruritic, erythematous, edematous papules and crusty ulcers at the mucocutaneous junctions of the lips, eyelids, nostrils, philtrum, vulva, prepuce, and perineum (Fig. 18-2).41 Affected rabbits also may experience metritis, abortion, or neonatal death. Lesions usually wax and wane over months. Recovered bucks may have star-shaped scrotal scars and can remain
Fig. 18-2 Periocular erythema and erosions in a rabbit infected with Treponema cuniculi (rabbit syphilis). (Courtesy of Stephen White, DVM. Picture from the case files of the Veterinary Medical Teaching Hospital, School of Veterinary Medicine, University of California. Used with permission.)
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carriers.65 Asymptomatic carrier rabbits may develop lesions if exposed to stress (e.g., overcrowding, poor sanitation). Rabbit syphilis is diagnosed from clinical signs and identification of the characteristic corkscrew-shaped causative organisms with dark-field microscopic examination of wet-mounted skin scrapings.55 Serologic testing can also help in diagnosis but lesions may be present before animals are serologically positive; thus false negatives can occur.* Because of low sensitivity in comparison with other methods, demonstration of spirochetes in silver-stained histologic sections is a less reliable diagnostic tool.55 Rabbit syphilis is usually self-limiting; however, autoinfection from perineum to face is possible and rabbits with syphilis may be more susceptible to other infections. Treatment is with parenteral benzathine penicillin G/procaine penicillin G (42,000-84,000 IU/kg SC q7d for three treatments) or penicillin G procaine (40,000-60,000 IU/kg IM q24h for 5-7 days). Treat all exposed rabbits and remove nursing kits from dams during treatment to decrease the risk of their developing penicillin-associated enterotoxemia.19 Breeding rabbits should be inspected regularly for lesions.
NECROBACILLOSIS Necrobacillosis, or Schmorl’s disease, is an uncommon skin infection with Fusobacterium necrophorum, an anaerobic, filamentous gram-negative bacterium normally found in the gastrointestinal tract and feces.22,33 Infection occurs commonly from wound contamination. Abscessed, ulcerated, necrotic lesions may be found around the head, neck, and feet.3,19 Does with large dewlaps that salivate profusely or rabbits with malocclusion may be prone to necrobacillosis due to excessive moisture in skin.55 The causative bacterium may be isolated from anaerobic culture of affected tissue. Debride and flush lesions and administer antibiotics effective against anaerobic bacteria, such as penicillin G procaine.
FUNGAL INFECTIONS DERMATOPHYTOSIS Dermatophytosis, or ringworm, is a potentially zoonotic and often asymptomatic infection of rabbit skin, most commonly caused by Trichophyton mentagrophytes.† Rabbits are less often infected with Microsporum species, but this species may be more common in pet rabbits.81 Dermatophytosis occurs more often in young rabbits, possibly because of their incompletely developed immune systems and low levels of fungistatic fatty acids in sebum. Asymptomatic carriers may develop lesions in response to overcrowding, malnutrition, or other infections. Rabbits also may acquire infection from humans or other pets. Dermatophytes cause alopecic skin lesions around the head, legs, feet, and nail beds that may be dry, crusty, erythematous, or pruritic. Histopathologic examination shows a hyperkeratotic and acanthotic epidermis with an acute to chronic inflammatory cell infiltrate in the underlying dermis. Diagnose dermatophytosis based on growth of the infecting organism on dermatophyte culture medium from hair or skin samples. Speciating the
*References 3, 18, 22, 27, 32, 40, 79. †References 3, 6, 11, 14, 22, 27, 28, 33, 55, 74, 81, 82.
dermatophyte may determine the origin of the dermatophyte infection. Fungal mycelia and arthrospores on hair shafts also may be seen on skin scrapings or hair plucks mounted in 10% potassium hydroxide or in skin biopsy samples stained with a periodic acid-Schiff stain, Gridley fungal stain, or Gomori’s methenamine silver stain. Fluorescence of mycelia under ultraviolet light from a Wood’s lamp is not a reliable method of testing in rabbits because T. mentagrophytes does not fluoresce and debris, bacteria, and keratin plugs can cause false-positive white to blue fluorescence.6 Although most dermatophyte infections in rabbits are self-limiting, treatment is recommended because of potential zoonosis.6,57 Several methods can be used to treat dermatophytosis in rabbits. Localized, topical therapy for dermatophytosis is not recommended because dermatophytes can also be present in nonlesional areas. Effective treatments for dermatophytosis include dips with lime sulfur solution (LymDyp, IVX Animal Health, Inc., St. Joseph, MO) 1:32 dilution with water twice weekly, or 0.2% enilconazole (Immaveral, Janssen; not available in the United States) twice weekly. There have been anecdotal reports of stress-induced death from bathing rabbits; thus rabbits may have to be tranquilized beforehand. In addition, treat rabbits with multiple lesions systemically with griseofulvin (15-25 mg/kg PO q24h or divided q12h for 30 days; Grifulvin V Suspension, Ortho Pharmaceutical, Raritan, NJ).3,27,41,74 If griseofulvin is in an ultramicrosized form, decrease the dose by 50%, and administer with a high-fat meal for better absorption.6 Use griseofulvin cautiously, because at high doses it can cause bone marrow suppression and panleukopenia.11 Also, griseofulvin is teratogenic and cannot be given to pregnant does. Owners administering griseofulvin should wear gloves. Other drugs used to treat dermatophytosis include itraconazole (Sporanox, Ortho-McNeil-Janssen Pharmaceuticals, Titusville, NJ), 5 to 10 mg/kg PO q24h, and terbinafine (Lamisil, Novartis Pharmaceuticals Corp, St. Louis, MO), 10 mg/kg PO q24h. Anecdotal reports exist of treatment of rabbit dermatophytosis with lufenuron (Program, Novartis Animal Health, Greensboro, NC), an insect growth regulator that inhibits chitin synthesis in fungal cell walls.6 However, use of lufenuron in rabbits is off label; controlled studies of this drug in rabbits have not yet been performed, and this treatment is not considered effective in the management of dermatophytosis in other animals. The clipping lesions of before applying topical medications is controversial. Some feel that clipping helps to make medications penetrate the skin; others warn against clipping because of the risk of inducing skin abrasions through which infection may become generalized.6 If hair is clipped, clip only the hair surrounding the wound and dispose of infected hair carefully. Continue systemic antifungal treatment until monthly fungal cultures are negative twice.11,28,33 Most treatments last a minimum of 3 to 4 months. Disinfection of the environment also is essential. Vacuum contaminated areas, and clean all surfaces with a 1:10 solution of bleach and water. Foggers containing enilconazole or formaldehyde are preferred to steam cleaning for carpets because steam cleaning does not reach a temperature high enough to kill infectious spores. Commercial steam cleaning services are preferable, because they use higher-temperature steam to clean carpets. Be careful not to use oral cisapride concurrently with azole drugs as they may potentially interact adversely.19
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Fig. 18-4 Scaling in a rabbit with fur mites (Cheyletiella parasi-
Fig. 18-3 Severe otic debris in a rabbit with ear mites (Psoroptes
tovorax). (Courtesy of Stephen White, DVM. Picture from the case files of the Veterinary Medical Teaching Hospital, School of Veterinary Medicine, University of California. Used with permission.)
cuniculi). (Courtesy of Jan Declercq, DVM.)
Dermatophytes may be hard to eliminate from large rabbit colonies. Large numbers of infected rabbits can be sprayed with a 2% to 3% lime sulfur solution weekly for 4 weeks. In large outbreaks of dermatophytosis, also treat with a systemic antifungal such as griseofulvin-medicated diets (0.825 g griseofulvin per kilogram of diet).41
PARASITIC INFECTIONS EAR MITES The rabbit ear mite, Psoroptes cuniculi, is a large, obligate, nonburrowing parasite with a 3-week life cycle and the ability to survive off the host for up to 21 days. Transmission occurs through direct contact with affected rabbits or with fomites.74 Psoroptes typically causes inflammation and reddish-brown, bran-like crusting of one or both external ear canals, head shaking, ear drooping, and pruritus (Fig. 18-3).* Crusty exudates develop as a result of hypersensitivity to the mite.74 In debilitated rabbits, inflammation and crusting may extend to the face, dewlap, neck, trunk, legs, feet, and perineum. Mites may spread down the ear canal, leading to otitis media and neurologic signs due to perforation of the tympanum.80 Rabbits with subclinical infections that are otherwise healthy may be only mildly pruritic for years. Recent immunohistochemical studies of the serum of rabbits with repeated Psoroptes infection suggest that rabbits may acquire long-lasting immunity after multiple infections.62 Psoroptes mites may be identified with the naked eye, otoscope, or microscope. Mites have 2 to 3 pairs of legs with jointed pedicles ending in a sucker-shaped bell. Microscopic examination of wet mounts of the crust reveals mites, mite feces, mite eggs, inflammatory cells, desquamated epithelial cells, and serum. Ear mites can be treated in an off-label fashion with various parasiticides including ivermectin (Ivomec, Merck, AgVet Division, Rahway, NJ), 0.4 mg/kg SC q7-14d for three treatments), or selamectin (Revolution, Pfizer Animal Health, New York, NY), 6 or 18 mg/kg topically q4wk for 1-2 treatments) (see “Fleas,” below).† Imidacloprid and moxidectin (Advocate/Advantage Multi, Bayer, Agriculture Division, Shawnee Mission, KS) are also
*References 3, 8, 19, 22, 28, 32, 33, 41, 55, 74. †References 3, 4, 8, 9, 19, 22, 28, 32, 33, 48.
effective for ear mites when applied topically three times 30 days apart.18 Subcutaneous eprinomectin (200-300 μg/kg once) can be used for Psoroptes, but this treatment is not effective topically.53 Less effective treatments are topical mineral oil, acaricides, and flea powder.27,41 Severe pruritus may be treated with a combination of topical thiabendazole, dexamethasone, and neomycin sulfate (Tresaderm, MSD-AGVET, Rahway, NJ). Meloxicam (Metacam, Boehringer Ingelheim Vetmedica, St. Joseph, MO), 0.2 mg/kg PO q12-24h for 5 days, may be administered to reduce pain and inflammation from the otitis.74 Aural crusts should not be debrided before treatment, as they usually resolve afterwards; they may be painful to remove, and removing them may lead to damage of the ear canal.16,80 Also, use any insecticide dips or baths (such as carbaryl, pyrethrin, and lime sulfur-based products) cautiously in rabbits, because shock and death have occurred in association with these products.22 The act of bathing or dipping rabbits may induce stress that is exacerbated by overheating, chilling, or the presence of hepatic dysfunction. The toxicity of topical drugs used safely in cats and dogs may be related to the thinness of rabbit skin, ingestion of topical products from excessive grooming, hypersensitivity to the drug’s vehicle, or absolute overdosage.3 To prevent reinfection, clean the environment and treat contaminated areas with flea products. Fipronil spray (Frontline, Merial Limited, Duluth, GA) has been used in rabbits every 2 months as a preventative measure in case of poor environmental disinfection.10 However, both topical and spray forms of fipronil have been associated with death in rabbits, and the product’s manufacturer does not recommend its use in this species.19,47
FUR MITES Cheyletiella parasitovorax, the rabbit fur mite, is zoonotic to people, dogs, and cats.17,27,31,50 In humans, it causes mild pruritic dermatitis. This nonburrowing mite lives on the keratin layer of the epidermis and is often called “walking dandruff” because it resembles a large, white, mobile dandruff flake (Fig. 18-4).19,33,74 Although Cheyletiella can cause subclinical disease, more often it causes a scaly, dry, sometimes pruritic dermatitis with patchy alopecia or broken hairs over the dorsal neck, trunk, hind end, and abdomen.32,37,41,55,80 Severe infestations occur more commonly in young, immunosuppressed, or obese rabbits that have difficulty grooming.19,74 In severe infestations, mites may be visible to the naked eye as clear to white saddle shapes with inward-curving claws and hook-like mouth parts.22 Their eggs adhere to hair shafts. Histopathologic analysis of
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affected skin shows hyperkeratosis with an exudate of inflammatory cells (mononuclear phagocytes, plasma cells, lymphocytes, and eosinophils). Diagnose Cheyletiella infection from clinical signs and microscopic identification of mites in skin scrapings, acetate tape preparations, or skin and hair debris that sticks to a flea comb. Treat infected rabbits with ivermectin (0.4 mg/kg SC q7-14d for three treatments) or with selemectin (12 mg/kg applied topically to skin at the base of the neck q2-4wk for one to three treatments).28,38,44,49 Other treatments include weekly lime sulfur dips for 3 to 6 weeks or application of carbaryl flea powder appropriate for cats twice weekly for 6 weeks. Dips and flea powder have successfully eliminated mites, but both treatments have been associated with toxicity in rabbits. In addition, comb out skin debris and dead mites and disinfect the environment with flea products (i.e., carbaryl 5% dust). Address any underlying causes such as obesity. Another mite that causes dermatitis in rabbits but is not zoonotic is Leporacarus gibbus (formerly Listrophorus gibbus).* Like Cheyletiella, this mite attaches to hair shafts; however, its eggs stick more distally on the shaft and are more loosely attached. Rabbits can carry Leporacarus asymptomatically until they become stressed and develop clinical signs.30 Leporacarus may cause a moist, sometimes pruritic alopecic dermatitis affecting the back, groin, ventral abdomen, and tail.3 Diagnose Leporacarus infection by the same methods as those used in rabbits infected with Cheyletiella. Leporacarus mites are brown and laterally compressed, with short legs and single projections extending from the head. Males have long clasping organs projecting caudally from their bodies. Empty larval cuticles persist on hair shafts, making the fur look as though it were covered with salt and pepper.19 Treatment is the same as for Cheyletiella infection. Dead mites may persist on hair shafts even after treatment. Sarcoptes scabiei and Notoedres cati occur in both laboratory and pet rabbits.27,30,41,55,74 These mites bury and cause a yellow, crusty, pruritic alopecic dermatitis around the head, neck, trunk, feet, and genitals.19 Infected, pruritic rabbits may self-mutilate. Like Cheyletiella, Sarcoptes mites are zoonotic. Diagnosis and treatment are the same as for Cheyletiella infection.33 Demodex cuniculi may occur in rabbits without clinical signs.33 This mite normally inhabits the epidermis and hair follicles; rarely, it causes nonpruritic alopecia in rabbits that are immunosuppressed by other diseases or subject to environmental stress. As a follicular mite, Demodex may be treated systemically in clinically affected rabbits with ivermectin, as described above for cheyletiellosis, or topically with a 0.05% solution of amitraz, until two negative skin scrapings are achieved.23,27,80 Laboratory rabbits are commonly affected by another mite, Ornithonyssus bacoti, which causes pruritus, alopecia, dermal crusting, and anemia and may be associated with secondary bacterial infection.80 Eradication of this mite is difficult and involves treatment of both the animals and the surrounding environment.
FLEAS Flea infestation most commonly affects rabbits housed with dogs and cats.27,28,32,34,59 Pet rabbits are often infected with the cat flea, Ctenocephalides felis, or the dog flea, Ctenocephalides canis. Other fleas found on domestic rabbits, especially those
*References 33, 39, 41, 55, 56, 80.
exposed to wild rabbits, include Cediopsylla, Odontopsyllus, Spilopsyllus, and Hoplopsyllus species, and Echnidnophaga gallinacea (the “sticktight flea”). In rabbit colonies infested with Spilopsyllus, the life cycle of the flea is determined by the host rabbit’s hormonal cycle, with outbreaks occurring most often in pregnant does and the young.80 Flea infection may manifest itself as a dull coat, easily epilated hair, and patchy alopecia with pruritus, skin erythema, and crusting, especially on the pinnae and face (with Spilopsyllus) and along the dorsum and tail (with Ctenocephalides).3,19,22,74 Fleas not only cause dermatitis but also transmit myxoma virus (discussed later). Diagnose flea infestation from clinical signs and by identifying fleas, their debris, and their eggs on the rabbit. Treat flea dermatitis with a carbaryl-based flea powder safe for cats once or twice a week. Imidacloprid (Advantage, Bayer Animal Health, Agriculture Division, Shawnee Mission, KS) and the insect growth regulator lufenuron (Program, Novartis Animal Health Canada, Mississauga, Ontario) have been used in rabbits to treat fleas,19,30,31,33 although neither is labeled for use in rabbits. The 10% spoton topical formulation of imidacloprid (Advantage 40 for cats, 0.4 mL topical solution) has been used safely and effectively to treat flea infestations in rabbits.36 For rabbits more than 10 weeks old and weighing less than 4 kg, 0.4 mL is applied to the skin at the base of the neck. Adult rabbits weighing more than 4 kg should receive 0.8 mL at the same site. Rabbits treated with fipronil have had adverse reactions and some have died; thus use of this product in rabbits is not recommended.80 A study to determine the efficacy and pharmacokinetics of selamectin against flea (C. felis) infestations in rabbits demonstrated that while selamectin at both 10 and 20 mg/kg was >91% effective in eliminating existing fleas infestations in rabbits, neither dose provided prolonged residual activity.7 The short duration of selamectin activity in rabbits is believed to be due to its short half-life (approximately 1 day) after topical administration as compared with its duration in dogs (11.1 days) and cats (8.25 days). These results suggest that selamectin is more rapidly absorbed, metabolized, and eliminated in rabbits than in dogs and cats. Thus rabbits require topical administration of selamectin every 7 days to effectively treat flea infestations long term, as compared with every 30 days in other domestic species. Although a dose as high as 20 mg/kg was suggested in this study, no toxicologic studies on selamectin use in rabbits have been done. Transient (<24-hour) ataxia was anecdotally reported in a 2-year-old healthy angora rabbit after topical administration of selamectin at 20 mg/kg (K. E. Quesenberry, personal communication). The environment also must be treated with insect growth regulators and insecticidal sprays and rabbits must be removed until the environmental products have dried. Borate powder may be used on infected rugs. Permethrin and pyrethrin-based environmental treatments may be ineffective because of flea resistance. Directed sprays are more effective than aerosolized flea treatments (“flea bombs”).
MYIASIS Larvae of Cuterebra species, or bot flies, and maggots of dipterid flies may infect rabbits housed outdoors in warm weather.16,27,28,33 Cuterebra flies are large and have three larval stages that commonly infect wild rabbits and rodents. Adult flies live only a short time to breed. Larvae pupate in the subcutis, causing multiple swellings especially over the dorsum and in the axillary, inguinal, and ventral cervical regions. Each 1- to 3-cm subcutaneous swelling encapsulates a single larva and has
CHAPTER 18 Dermatologic Diseases a breathing hole visible at the skin surface.22,30,32,74 Although some rabbits are unaffected by these swellings, others suffer pain and become weak, anorectic, dehydrated, and lame and can go into shock. Cuterebra larvae also can migrate aberrantly from the nasal passages, eyes, sinuses, and ear canals through the central nervous system, causing neurologic signs.25,27 Treatment involves removal of larvae. Affected rabbits typically must be tranquilized for this often painful procedure.17,30 After preparing the affected skin area for surgery, enlarge the breathing holes with hemostats. Gently remove the larvae through the enlarged openings, ensuring that they are not crushed, because damage to larvae can cause anaphylaxis. After removing the larvae and debriding necrotic tissue, the swelling usually resolves. If it does not, or if the skin is abscessed, surgically excise the affected tissue. Administer antibiotics to treat secondary bacterial infection and NSAIDs to reduce pain. Prevent cuterebrid infection through fly control and the use of protective screens. Noncuterebrid maggots, including those of the flesh fly Wohlfahrtia vigil, cause moist dermatitis and matted hair around the perineum, face, and rump, particularly during warm summer months in rabbits housed outdoors.* Hundreds of larvae may colonize a single skin area. Maggots cause extensive lesions by feeding on dead tissue.22,32 Preexisting wounds are not necessary for maggot infection. Perineal dermatitis (due to adherence of caecotrophs, or night feces), urine scald, or skin-fold dermatitis secondary to obesity may predispose to maggot infection. Inability of rabbits with dental disease to groom properly also may predispose to infection. Maggot infestation initially may not be obvious because once eggs hatch, maggots may be concealed within matted fur. Initially, affected rabbits may be inappetant and restless but may subsequently go into shock from extensive skin necrosis and secondary bacterial infection. Treat infested rabbits first with supportive care, then sedate them to clean affected skin with a dilute antiseptic solution (such as chlorhexidine), remove the maggots, and debride necrotic tissue. Administer ivermectin (0.4 mg/kg SC q14d for two treatments) to kill larvae, an antibiotic with good skin activity such as trimethoprim-sulfamethoxazole to treat secondary bacterial infection, and analgesics and NSAIDs for pain. Clean the surgical site daily, apply a topical antimicrobial preparation such as silver sulfadiazine cream (Silvadene Cream 1%, Monarch Pharmaceuticals, Bristol, TN) or Silvasorb gel (Medline Industries Inc.), and allow the wound to heal by secondary intention. Rabbits with extensive infestations may initially appear stable after surgery but then die, possibly from secondary infection of necrotic wounds with Clostridium species. Penicillin G procaine (30,000-60,000 IU/kg SC q24h for up to 5 days) may be administered to prevent secondary anaerobic bacterial infection. Blackflies of the Simuliidae family bite rabbits around lips, ears, and nares and transmit viral infections, including myxomatosis (see below).33 Bites are painful and may become inflamed. Fly control and protective screens prevent blackfly bites.
TICKS Ticks affect both domestic and wild rabbits. The most common hard (ixodid) tick of rabbits, the continental rabbit tick or Haemaphysalis leporis-palustris, has three stages in its life cycle, and
*References 19, 27, 28, 33, 41, 74, 80.
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rabbits may host each stage.33,41,80 Other ixodid ticks that feed on rabbits include Amblyomma, Boophilus (cattle ticks), Rhipicephalus, and Dermacentor species. Soft (argasid) ticks, such as Otobius lagophilus, Ornithodoros turicata, and Ornithodoros parkeri, also may parasitize rabbits. Severe tick infestation causes blood loss with macrocytic normochromic anemia. Rabbit ticks may also transmit viral infections, such as myxomatosis and papillomatosis (see later discussion), and zoonotic diseases, including tularemia, Lyme disease, and Rocky Mountain spotted fever. Remove visible ticks with forceps and give ivermectin (0.4 mg/ kg SC) to kill any remaining feeding ticks.
LICE The rabbit louse, Haemodipsus ventricosus, rarely occurs on domestic rabbits.3,20,41,71 With severe infestation, this bloodsucking parasite causes anemia, weight loss, erythematous papules, hair loss, and pruritus. Haemodipsus also may transmit tularemia (see Chapter 40).80 Lice and their eggs (nits) are visible with the naked eye, especially on the dorsolateral aspects of the body and perineum.19 Infected rabbits may be treated with ivermectin (0.4 mg/kg SC q14d for three treatments).
PINWORMS Passalurus ambiguus, the common rabbit pinworm, is hostspecific.3 It inhabits the small intestine, cecum, and colon, passing eggs into the feces without causing clinical signs. This nematode may be a commensal that aids in plant digestion.74 Heavy worm burdens cause weight loss, anorexia, perianal pruritus, fecal impaction, self-trauma, and occasionally rectal prolapse.21,74 Diagnose pinworm infection by clinical signs and by identifying ova or adult worms in a fecal smear or fecal float. Treatment is difficult, because eggs passed in contaminated feces often are reingested. Usually infection is only transiently eliminated and reinfection is common. Clean pinworms from the perianal area and give thiabendazole (50 mg/kg PO) or fenbendazole (10-20 mg/ kg PO) in two doses 10 to 14 days apart. Alternatively, administer piperazine directly to individual rabbits (200 mg/kg PO, repeat in 14 days) or give it in food to adult rabbits (0.5 g/kg per day for 2 days) or to juvenile rabbits (0.75 g/kg per day for 2 days).21,41 Piperazine may also be given in drinking water (100 mg/100 mL of water for 1 day, repeat in 10 days). Environmental cleanup to kill pinworm eggs is difficult.
TAPEWORM CYSTS Coenurus serialis, the parasitic larval stage of the dog and fox tapeworm Taenia serialis, can cause multiple fluctuant subcutaneous and intramuscular nodules in rabbit connective tissues.19,80 Lesions occur in cheeks, axillae, and the retrobulbar space. Cysts typically do not affect the rabbit. Individual lesions may be excised; however, there is no effective treatment for multiple lesions.
VIRAL INFECTIONS MYXOMATOSIS Myxomatosis is caused by myxoma viruses of the pox family.27,42 This disease is transmitted through arthropod bites (mosquito, gnat, fly, flea, fur mite), on spiny thistles and birds’
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feet, and by direct contact.1,22,27,33 Clinical signs depend on the virulence of the viral strain and on the genus and species of the infected rabbit. In wild rabbits (Sylvilagus species), myxomatosis appears as benign skin tumors at the site of viral entry (usually at the ear base). Wild rabbits serve as a reservoir host for this disease in domestic rabbits (Orytolagus species). Myxomatosis is enzootic in the western United States, where outbreaks have occurred in rabbitries.55 Domestic rabbits may exhibit lethargy, fever, anorexia, skin hemorrhages, and seizures with a high mortality rate.* The exact cause of death is not known but is hypothesized to be virus-induced gram-negative septicemia.74 Rabbits that survive initial infection may develop mucopurulent blepharoconjunctivitis and erythematous, edematous nodules on the face, ears, eyelids, and perineum.3,27,33,41 Occasionally lesions spontaneously regress. In large outbreaks, the virus may be inhaled and may lead to pulmonary infection and pneumonia.65 Diagnose myxomatosis by clinical signs, histopathologic analysis of affected tissue, and viral isolation. Histopathologic examination of biopsy samples of affected skin shows undifferentiated mesenchymal cells within a matrix of mucin, inflammatory cells, and edema. Intracytoplasmic inclusions may be seen in the epidermis and in conjunctival epithelium.55 Treatment is supportive and often unsuccessful. Give antibiotics for secondary bacterial infection. Eliminate insect vectors and wild rabbit reservoirs, and euthanatize severely affected rabbits.74 Vaccines, including one made from a genetically engineered recombinant virus, have been developed for high-risk rabbitries but are not commercially available.1 Cross-immunity may be attained with a live attenuated Shope fibroma virus (see below) vaccine, but this vaccine does not provide complete immunity to myxoma virus, must be boosted repeatedly, and can be administered only to healthy, nonpregnant animals.74
RABBIT (SHOPE) FIBROMA VIRUS Rabbit (Shope) fibroma virus is a Leporipoxvirus that infects many European wild rabbits and cottontails (Sylvilagus species) in the United States and Canada; other rabbit species are resistant, but infection of domestic rabbits has been reported.3,33,42,55,74 The natural host of the poxvirus is the Eastern cottontail (Sylvilagus floridanus). Other Sylvilagus species (Sylvilagus bachmani, Sylvilagus nuttalli, and Sylvilagus audoboni) are refractory to infection. Malignant rabbit fibroma virus, a recombinant virus of Shope fibroma and myxoma virus, is a distinct virus that causes generalized infection, tumors, and immunosuppression that is usually fatal. Rabbit (Shope) fibroma virus is transmitted by biting arthropods, including fleas and mosquitoes. In cottontails, clinical signs of fibroma virus infection are worse in young rabbits and include large, flat, wart-like subcutaneous tumors of the face, feet, legs, and perineum. Diagnosis is based on gross and histopathologic analysis of tumors and on isolation of virus from affected skin. Tumors start as acute inflammation and progress to localized fibroblastic proliferation with infiltration of mononuclear and polymorphonuclear inflammatory cells. Large intracytoplasmic, eosinophilic inclusions may be seen in mesenchymal cells and in the epidermis overlying the tumors.55 The epidermis may become necrotic and slough or lesions may regress over several months. Treat with supportive care,
*References 27, 28, 32, 41, 74, 80.
administer antibiotics for secondary bacterial infection, and eliminate insect vectors.
RABBIT (SHOPE) PAPILLOMAVIRUS The rabbit (Shope) papillomavirus, also called cottontail rabbit papillomavirus, is an oncogenic DNA virus of the Papovaviridae family that is transmitted by biting arthropods (especially continental rabbit ticks, reduvid bugs, and mosquitoes).33,55 This virus causes rough, red, wart-like, keratinized, often pigmented lesions on the ears, eyelids, neck, shoulders, abdomen, and thighs. Lesions in wild cottontail rabbits (Sylvilagus species) are often found around the neck and shoulders and in domestic rabbits (Orytolagus species) around the hairless areas of the ears and eyelids.41,42,55,74 This virus is distinct from the rabbit oral papillomavirus (see below), which causes tongue and oral cavity papillomas in domestic rabbits; however, dermal lesions induced by the oral papillomavirus provide cross-immunity to the Shope papillomavirus.74 In rabbits infected with Shope papillomavirus, lesions may undergo immune-mediated resolution and disappear after months, or they may progress to squamous cell carcinoma that often metastasizes to axillary lymph nodes.3,55 Experimental inoculation of domestic rabbits with this virus frequently leads to the development of squamous cell carcinoma at the inoculation site.80 The incidence of carcinoma development from viral infection is threefold lower in cottontail rabbits than in domestic rabbits. Diagnose Shope papillomavirus by histopathologic analysis of typical gross lesions and viral isolation. In domestic rabbits, affected tissue is hyperkeratotic but rarely contains the inclusion bodies that are characteristic of papillomaviruses. Differential diagnosis must discriminate Shope papillomas from spontaneous, nonviral papillomas. In contrast, in the natural host (Sylvilagus species), papillomas usually contain infectious viral particles. Treat with surgical removal of horny papillomas and control arthropod vectors to help prevent spread of the disease.
ORAL PAPILLOMAVIRUS Rabbit oral papillomavirus causes oral papillomatosis in domestic rabbits, most commonly between 2 and 18 months of age.55 The virus is spread by direct contact and enters oral abrasions, leading to the development of slowly growing verrucous, sometimes pedunculated lesions ventral to the tongue that typically regress spontaneously within weeks.19 Histopathologic analysis reveals findings typical of squamous papillomas with basophilic intranuclear inclusions.
RABBITPOX Rabbitpox virus, an orthomyxopoxvirus, is a very rare, highly contagious and often fatal infection that is usually seen only among research populations in the United States and the Netherlands.33,74 This virus is transmitted by ingestion or inhalation of infected tissue.40 Affected rabbits die without clinical signs or develop enlarged lymph nodes, fever, and an erythematous rash that progresses to papules and crusty nodules. Papules may extend to the oral cavity, respiratory tract, spleen, and liver.55 There may be edema of the face, oral cavity, scrotum, and vulva.3 Some rabbits develop blepharitis, keratitis, and conjunctivitis. Diagnose rabbitpox through the recognition of characteristic skin lesions and virus isolation or fluorescent antibody
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if they are pregnant, pseudopregnant, or in heat. These rabbits pull hair from the dewlap, ventrum, legs, and thorax.80 Occasionally, rabbits groom excessively as a dissociative behavior in response to stress (i.e., overcrowding, boredom, aggression from cage mates, sexual frustration, or trauma).74 Generally, barbering causes alopecia without other signs of dermatitis.55 Occasionally, barbering can progress to self-mutilation.19 Treatment for barbering depends on its cause. Separate dominant and subordinate rabbits, provide a high-fiber diet, create a less crowded enclosure as needed, neuter if sexual frustration is suspected, and ensure adequate social stimulation. A change in light cycle or intensity may help. If no other treatment stops self-mutilation, try administering haloperidol (0.2-0.4 mg/kg PO q12h).19 Fig. 18-5 Patchy alopecia, scaling and erythema in a rabbit with cutaneous T-cell lymphoma. (Courtesy of Stephen White, DVM. Picture from the case files of the Veterinary Medical Teaching Hospital, School of Veterinary Medicine, University of California. Used with permission.)
identification of viral antigen in infected tissue. Histopathologic analysis of lesions shows lymphoid necrosis and mononuclear cell infiltration. Treat supportively.
CUTANEOUS NEOPLASIA Skin tumors unassociated with viral infection are uncommon in rabbits.22,33 Nonviral skin tumors include squamous cell carcinoma, basal cell tumor, trichoepithelioma, trichoblastoma, sebaceous cell carcinoma, nonviral squamous papilloma, apocrine carcinoma, meibomian adenoma, sebaceous carcinoma, lipoma, liposarcoma, myxosarcoma, nerve sheath tumor, fibrosarcoma, leiomyosarcoma, leiomyoma, rhabdomyosarcoma, anaplastic sarcoma, and malignant melanoma.51,75 One retrospective study found that trichoblastomas are the most common non-viral-associated cutaneous neoplasm.75 Uterine adenocarcinoma and connective tissue tumors can metastasize to skin.19,46 Lymphosarcoma occurs in rabbit skin as erythematous, crusty, alopecic plaques confined to the epidermis or extending through the dermis (Fig. 18-5).29,73,82 Often, cutaneous lymphosarcoma in rabbits is associated with neoplastic lymphocytes in organs besides the skin (i.e., lymph nodes and lungs). Other cutaneous growths in rabbits include collagenous hamartoma, a nodular, nonneoplastic, tumor-like lesion composed of excessive collagen that causes hair follicle compression and atrophy. In addition, rectal papillomas occur in rabbits as cauliflower-like lesions extending from the anus that either cause no clinical signs or mild, frank rectal bleeding.19,74 Diagnose neoplasia by microscopic examination of biopsy samples. Treat tumors with surgical excision or debulking. Chemotherapy has generally been unsuccessful in treating cutaneous lymphoma in rabbits (see Chapter 20). Complete surgical removal of rectal papillomas is usually curative.
BEHAVIORAL CAUSES OF SKIN DISEASE BARBERING Barbering, or hair pulling, occurs when a dominant rabbit pulls a subordinate’s hair; it can also be self-induced in rabbits fed low-fiber diets.27,32,33 Female rabbits self-barber to build nests
SELF-MUTILATION AFTER INTRAMUSCULAR INJECTION Rabbits commonly develop injection site reactions. Lesions appear within a few days of injection; when they are severe, the overlying skin can slough. Carprofen and enrofloxacin injections are frequently implicated in injection site reactions in rabbits.19 Intramuscular injection of ketamine and xylazine into the caudal thigh can result in mutilation of the digits on the ipsilateral leg 2 to 4 days later.2 Self-mutilation is thought to be caused by perineural drug infiltration around the sciatic nerve, resulting in axonal degeneration and dysthesia. If this occurs, clean and bandage damaged digits and administer antibiotics and anti-inflammatory agents such as meloxicam (0.2 mg/kg PO q12-24h for 5 days). To prevent this syndrome, give ketamine and xylazine separately in different sites and away from the sciatic nerve, such as in the lumbar muscles. Self-mutilation can occur following hypersensitivity reactions to any injection. Giving injections subcutaneously may help limit the incidence of these reactions.74
SKIN DISEASE OF UNKNOWN CAUSE SEBACEOUS ADENITIS Sebaceous adenitis has been reported in several different breeds and ages of rabbits. All affected animals had nonpruritic, scaly, flaky dermatitis that began around the face and neck and progressed to diffuse skin disease (Fig. 18-6).83 Histopathologic examination of affected tissue showed absence of sebaceous glands, hyperkeratosis, follicular keratosis, follicular dystrophy, perifollicular fibrosis, and replacement of sebaceous glands with perifollicular lymphocytic infiltrate. No infectious organisms were identified. Sebaceous adenitis in rabbits is histopathologically similar to sebaceous adenitis in dogs. But because sebaceous adenitis in rabbits can be associated with systemic disease, some cases of rabbit sebaceous adenitis are more similar clinically to another condition, thymoma-associated exfoliative dermatitis in cats. This is a paraneoplastic condition that also results in loss of the sebaceous glands and scaling.12 Sebaceous adenitis in the rabbit has also been seen with hepatitis.13 Various treatments for sebaceous adenitis—including isotretinoin (Accutane, Hoffman-LaRoche Inc., Nutley, NJ), etretinate (Tegison, Hoffman-LaRoche Inc.), prednisone, azathioprine (Imuran, Faro Pharmaceuticals, Bedminster, NJ), griseofulvin, essential fatty acids, and topical propylene glycol—have been tried with limited success. Rabbits with sebaceous adenitis usually die or are euthanatized because of the severity of the
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Fig. 18-7 Dermal fibrosis in an adult male intact rabbit. Note the excessive thick skin in the dorsal lumbosacral region extending over the rabbit’s back like a shell.
Fig. 18-6 Alopecia and scaling in a rabbit with sebaceous adenitis. Note the follicular casts visible at the margins of the pinnae. (Courtesy of Nico Schoemaker, DVM, PhD.)
skin condition. One report describes successful treatment of a 4-year-old male rabbit diagnosed by skin biopsy with sebaceous adenitis. The rabbit was given cyclosporine (Neoral, Novartis Pharmaceuticals, East Hanover, NJ), 5 mg/mg PO q24h, dissolved in Miglyol 812 (SASOL Germany GmbH, Witten, Germany), a mixture of triglycerides administered to facilitate gastrointestinal uptake of cyclosporine, in combination with oral essential fatty acids and topical 50% propylene glycol.72 After 2 months of therapy, the skin lesions had regressed and hair had regrown. In addition to this case, one of the present authors (LH) successfully treated two rabbits diagnosed with sebaceous adenitis with a combination of oral acitretin (Soriatane, Stiefel Laboratories, Coral Gables, FL), 1 mg/kg PO q24h for 8 weeks, then q48h for 8 weeks, then q96h maintenance, and pentoxifylline (Trental, Sanofi-Aventis Pharma, Paris, France), 10 mg/kg PO q24-48h for 6 weeks) for over a year. These rabbits were also bathed twice weekly with shampoo compounded from tranilast 0.5% (Rizaben, Kissei Pharmaceutical, Nagano, Japan) plus glycolic acid 1.5% and with a leave-on conditioner compounded from vitamins A and E plus ammonium lactate cream (Lac-hydrin cream; Ranbaxy Pharmaceuticals, Princeton, NJ). One of these two rabbits was also treated concurrently with radiation therapy for thymoma, a neoplasm that has been associated with exfoliative dermatitis in other companion animals.
shaped dermal fibrils and high variability in dermal cell diameter. Treatment for this syndrome in rabbits is not yet known.
DERMAL FIBROSIS Dermal fibrosis, a thickening of skin over the dorsal thorax and lumbar region, has been reported in two male rabbits without any other clinical signs (Fig. 18-7).19 No treatment was necessary, and the condition was hypothesized to be a hormonally induced secondary sexual characteristic.
EOSINOPHILIC GRANULOMA Eosinophilic granulomas appear as raised erythematous and erosive-ulcerative plaques and have been reported rarely in rabbits.26,81 This lesion may represent a hypersensitivity reaction to various causes, similar to eosinophilic granuloma lesions in cats. One case was associated with self-mutilation and concurrent Cheyletiella mites. Cytologic examination of impression smears shows large numbers of eosinophils. Confirm the diagnosis through biopsy. Treat with a glucocorticoid and manage the underlying condition.
ACKNOWLEDGMENT We thank Dr. Tom Donnelly for his review and helpful comments.
EHLERS-DANLOS-LIKE SYNDROME
References
Ehlers-Danlos syndrome is an inherited skin disorder in humans that is associated with abnormal collagen structure.67 It results from mutations in collagen-forming genes or collagensynthesizing enzymes that cause joint laxity and skin hyperextensibility. An Ehlers-Danlos-like syndrome has been reported in young rabbits with markedly fragile skin and slow-healing wounds5,24,80 The cause of this syndrome is not yet known, but a genetic defect is suspected because of the affected rabbits’ young ages and the occurrence of the condition in siblings. Electron microscopic examination of affected tissue revealed irregularly
1. Bárcena J, Morales M, Vásquez B, et al. Horizontal transmissible protection against myxomatosis and rabbit hemorrhagic disease by using a recombinant myxoma virus. J Virol. 2000;74:1114-1123. 2. Beyers TM, Richardson JA, Prince MD. Axonal degeneration and self-mutilation as a complication of the intramuscular use of ketamine and xylazine in rabbits. Lab Anim Sci. 1991;41:519-520. 3. Bourdeau PJ. Dermatology of small mammals. I. Parasitic and infectious skin diseases in rodents and rabbits. Proceedings. Fourth World Cong Vet Derm. 2000:195-200.
CHAPTER 18 Dermatologic Diseases 4. Bowman DD, Fogelson ML, Carbone LG. Effect of ivermectin on the control of ear mites (Psoroptes cuniculi) in naturally infested rabbits. Am J Vet Res. 1992;53:105-109. 5. Brown PJ, Young RD, Cripps PJ. Abnormalities of collagen fibrils in a rabbit with a connective tissue defect similar to Ehlers-Danlos syndrome. Res Vet Sci. 1993;55:346-350. 6. Canny CJ, Gamble CS. Fungal diseases. Vet Clin North Am Exot Anim Pract. 2003;6:429-433. 7. Carpenter JW, Dryden M, KuKanich B. Efficacy and pharmacokinetics of selamectin in the pet rabbit. Proceedings. Assoc Avian Vet Annual Conf. 2010:47. 8. Curtis SK, Brooks DL. Eradication of ear mites from naturally infested conventional research rabbits using ivermectin. Lab Anim Sci. 1990;40:406-408. 9. Curtis SK, Housley R, Brooks DL. Use of ivermectin for treatment of ear mite infestation in rabbits. J Am Vet Med Assoc. 1990;196:1139-1140. 10. Cutler SL. Ectopic Psoroptes cuniculi infestation in a pet rabbit. J Small Anim Pract. 1998;39:86-87. 11. Donnelly TM, Rush EM, Lackner PA. Ringworm in small exotic pets. Semin Avian Exot Pet Med. 2000;9:82-93. 12. Florizoone K. Thymoma-associated exfoliative dermatitis in a rabbit. Vet Dermatol. 2005;16:281-284. 13. Florizoone K, van der Luer R, van den Ingh T. Symmetrical alopecia, scaling and hepatitis in a rabbit. Vet Dermatol. 2007;18:161-164. 14. Garner MM. Cytologic diagnosis of disease of rabbits, guinea pigs, and rodents. Vet Clin North Am Exot Anim Pract. 2007;1:2549, v-vi. 15. Gentz EJ, Carpenter JW. Neurologic and musculoskeletal disease. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders Co; 1997:220-226. 16. Graham JE. Rabbit wound management. Vet Clin North Am Exot Anim Pract. 2004;7:37-55. 17. Hanselman BA, Kruth SA, Rousseau J, et al. Methicillin-resistant Staphylococcus aureus colonization in veterinary personnel. Emerg Infect Dis. 2006;12:1933-1938. 18. Hansen O, Gall Y, Pfister K, et al. Efficacy of a formulation containing imidacloprid and moxidectin against naturally occurring acquired ear mite infections (Psoroptes cuniculi) in rabbits. Intern J Appl Res Vet Med. 2005;3:281-286. 19. Harcourt-Brown F. Textbook of rabbit medicine. Woburn, Butterworth Heinemann. 2002. 20. Harkness JE, Wagner JE. The biology and medicine of rabbits and rodents. 4th ed. Baltimore: Williams & Wilkins; 1995: 143-151. 21. Harkness JE, Wagner JE. The biology and medicine of rabbits and rodents. 3rd ed. Baltimore: Williams & Wilkins; 1989:168-171. 22. Harvey C. Rabbit and rodent skin diseases. Semin Avian Exot Pet Med. 1995;4:195-204. 23. Harvey RG. Demodex cuniculi in dwarf rabbits (Oryctolagus cuniculus). J Small Anim Pract. 1990;31:204-207. 24. Harvey RG, Brown PJ, Young RD, et al. A connective tissue defect in two rabbits similar to the Ehlers-Danlos syndrome. Vet Rec. 1990;126:130-132. 25. Hendrix CM, DiPinto MN, Cox NR, et al. Aberrant intracranial myiasis caused by larval Cuterebra infection. Comp Contin Educ Pract Vet. 1989;11:550-559. 26. Henriksen P. Eosinophilic granuloma like lesion in a rabbit. Nord Vet Med. 1983;35:243-244. 27. Hillyer EV. Dermatologic diseases. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders Co; 1997:212-219. 28. Hillyer EV. Pet rabbits. Vet Clin North Am Small Anim Pract. 1994;24:25-65. 29. Hinton M, Regan M. Cutaneous lymphosarcoma in a rabbit. Vet Rec. 1978;103:140-141.
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30. Hughes JE. Diagnosis and treatment of selected rabbit dermatologic disorders. Exot DVM. 2004;5:18-20. 31. Hutchinson MJ, Jacobs DE, Bell GD, et al. Evaluation of imidacloprid for the treatment and prevention of cat flea (Ctenocephalides felis felis) infestations on rabbits. Vet Rec. 2001;148:695-696. 32. Jenkins JR. Skin disorders of the rabbit. J Small Exot Anim Med. 1991;1:64-65. 33. Jenkins JR. Skin disorders of the rabbit. Vet Clin North Am Exot Anim Pract. 2001;4:543-563. 34. Jenkins JR. Soft tissue surgery and dental procedures. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders; 1997:227-239. 35. Juhász-Kaszanyitsky E, Jánosi S, Somogyi P, et al. MRSA transmission between cows and humans. Emerg Infect Dis. 2007;13(4):630-632. 36. Khanna T, Friendship R, Dewey C, et al. Methicillin resistant Staphylococcus aureus colonization in pigs and pig farmers. Vet Microbiol. 2008;128:298-303. 37. Kim SH, Jun HK, Song KH, et al. Prevalence of fur mites in pet rabbits in South Korea. Vet Dermatol. 2008;19:189-190. 38. Kim SH, Lee JY, Jun HK, et al. Efficacy of selamectin in the treatment of cheyletiellosis in pet rabbits. Vet Dermatol. 2008;19:26-27. 39. Kirwan AP, Middleton B, McGarry JW. Diagnosis and prevalence of Leporacarus gibbus in the fur of domestic rabbits in the UK. Vet Rec. 1998;142:20-21. 40. Kottler S, Middleton JR, Weese JS, et al. Prevalence of Staphylococcus aureus methicillin-resistant Staphylococcus aureus carriage in three populations. J Vet Int Med. 2010;24:132-139. 41. Kraus AL, Weisbroth SH, Flatt RE, et al. Biology and diseases of rabbits. In: Fox JG, Cohen BJ, Loew FM, eds. Laboratory animal medicine. San Diego: Academic Press; 1984:207-240. 42. Krogstad AP, Simpson JE, Korte SW. Viral diseases of the rabbit. In: Greenacre C, ed. Vet Clin North Am Exot Anim Pract. 2005;8:123-138. 43. Kuehn BM. Antibiotic-resistant “superbugs” may be transmitted from animals to humans. JAMA. 2007;298:2125-2126. 44. Kurtdede A, Karaer Z, Acar A, et al. Use of selamectin for selemectin in the treatment of psoropticpsorptic and sarcoptic mite infestation in rabbits. Vet Dermatol. 2007;18:18-22. 45. Leonard FC, Markey BK. Methicillin-resistant Staphylococcus aureus in animals: a review. Vet J. 2008;175:27-36. 46. Li X, Schlafer DH. A spontaneous skin basal cell tumor in a black French minilop rabbit. Lab Anim Sci. 1992;42:94-95. 47. Malley D. Use of Frontline spray on rabbits [letter]. Vet Rec. 1997;140:664. 48. McTier TL, Hair JA, Walstrom DJ, et al. Efficacy and safety of topical administration of selamectin for treatment of ear mite infestation in rabbits. J Am Vet Med Assoc. 2003;223:322-324. 49. Mellgren M, Bergvall K. Treatment of rabbit cheyletiellosis with selamectinselemectin or ivermectin: a retrospective case study. Acta Vet Scand. 2008;50:1-6. 50. Merchant SR. Zoonotic diseases with cutaneous manifestation: part I. Compend Contin Educ Pract Vet. 1990;12:371-377. 51. Miwa Y, Mochiduki M, Nakamaya H, et al. Apocrine adenocarcinoma of possible sweat gland origin in a male rabbit. J Small Anim Pract. 2006;47:541-544. 52. Morgan M. Methicillin-resistant Staphylococcus aureus and animals: zoonosis or humanosis? J Antimicrob Chemother. 2008;62:1181-1187. 53. Pan B, Wang M, Xu F, et al. Efficacy of an injectable formulation of eprinomectin against Psoroptes cuniculi, the ear mange mite in rabbits. Vet Parasitol. 2006;137:386-390. 54. Paul-Murphy J. Reproductive and urogenital disorders. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders Co; 1997:202-211.
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55. Percy D, Barthold SW. Pathology of laboratory rodents and rabbits. 3rd ed. Ames: Blackwell Publishing; 2007. 56. Pinter L. Leporacarus gibbus and Spilopsyllus cuniculi infestation in a pet rabbit. J Small Anim Pract. 1999;40:220-221. 57. Quesenberry KE. Rabbits. In: Birchard SJ, Sherding RJ, eds. Saunders manual of small animal practice. Philadelphia: WB Saunders; 1994:1345-1362. 58. Remeeus PG, Verbeek M. The use of calcium hydroxide in the treatment of abscesses in the cheek of the rabbit resulting from a dental periapical disorder. J Vet Dent. 1995;12:19-22. 59. Reuter K, Pospischil R, Endepols S, et al. Flea infestation in exotic pet animals. Suppl Compend Contin Educ Pract Vet. 2002;24:10-13. 60. Rich M. Staphylococci in animals: prevalence, identification and antimicrobial susceptibility, with an emphasis on methicillin-resistant Staphylococcus aureus. Br J Biomed Sci. 2005;62:98-105. 61. Rosenfield ME. Successful eradication of severe abscesses in rabbits with long-term administration of penicillin G benzathine/penicillin G procaine. Available at www.unix.oit.umass. edu/~jwmoore/bicillin/bicillin.htm. Accessed on July 20, 2010. 62. Rossi G, Donadio E, Perrucci S. Immunohistochemistry of Psoroptes cuniculi stained by sera from naïve and infested rabbits: preliminary results. Parasitol Res. 2007;100:1281-1285. 63. Saito K, Tagawa M, Hasegawa A. Rabbit syphilis diagnosed clinically in household rabbits. J Vet Med Sci. 2003;65:637-639. 64. Sasaki T, Kikuchi K, Tanaka Y, et al. Methicillin-resistant Staphylococcus pseudointermedius in a veterinary teaching hospital. J Clin Microbiol. 2007;45:1118-1125. 65. Saunders RA, Davies RR. Notes on rabbit internal medicine. Oxford: Blackwell Publishing; 2005. 66. Seguin JC, Walker RD, Caron JP, et al. Methicillin-resistant Staphylococcus aureus outbreak in a veterinary teaching hospital: potential human-to-animal transmission. J Clin Microbiol. 1999;37:1459-1463. 67. Sinke JD, van Dijk JE, Willemse T. A case of Ehlers-Danlos-like syndrome in a rabbit with a review of the disease in other species. Vet Q. 1997;19:182-185. 68. Tyrrell KL, Citron DM, Jenkins JR, et al. Periodontal bacteria in rabbit mandibular and maxillary abscesses. J Clin Microbiol. 2002;40:1044-1047. 69. van Duijkeren E, Box AT, Heck ME, et al. Methicillin-resistant staphylococci isolated from animals. Vet Microbiol. 2005;105:313-314. 70. van Duijkeren E, Wolfhagen MJ, Box AT, et al. Human-to-dog transmission of methicillin-resistant Staphylococcus aureus. Emerg Infect Dis. 2004;10:2235-2237.
71. van Loo I, Huijsdens X, Tiemersma E, et al. Emergence of Staphylococcus aureus of animal origin in humans. Emerg Infect Dis. 2007;13:1834-1839. 72. van Zeeland YRA, Lee AJ, Schoemaker NJ. Successful treatment of a rabbit with sebaceous adenitis. Proceedings. Ann Conf Assoc Avian Vet. 2008:81-82. 73. Vangeel I, Pasmans F, Vanrobaeys M, et al. Prevalence of dermatophytes in asymptomatic guinea pigs and rabbits. Vet Rec. 2000;146:440-441. 74. Vennen KM, Mitchell M. Rabbits. In: Mitchell M, Tully T, eds. Manual of exotic pet practice. St. Louis: Saunders-Elsevier; 2009:375-405. 75. von Bomhard W, Goldschmidt MH, Shofer FS, et al. Cutaneous neoplasms in pet rabbits: a retrospective study. Vet Pathol. 2007;44:579-588. 76. Walther B, Friedrich AW, Brunnberg L, et al. [Methicillinresistant Staphylococcus aureus (MRSA) in veterinary medicine: a “new emerging pathogen”?] [article in German]. Berl Munch Tierarztl Wochenschr. 2006;119:222-232. 77. Walther B, Wieler LH, Friedrich AW, et al. Methicillin-resistant Staphylococcus aureus (MRSA) isolated from small and exotic animals at a university hospital during routine microbiological examinations. Vet Microbiol. 2008;127:171-178. 78. Weese JS, Dick H, Willey BM, et al. Suspected transmission of methicillin-resistant Staphylococcus aureus between domestic pets and humans in veterinary clinics and in the household. Vet Microbiol. 2006;115:148-155. 79. Weese JS, Faires M, Rousseau J, et al. Cluster of methicillinresistant Staphylococcus aureus colonization in a small animal intensive care unit. J Am Vet Med Assoc. 2007;231:1361-1364. 80. White SD, Bourdeau PJ, Meredith A. Dermatologic problems of rabbits. Semin Avian Exot Pet Med. 2002;11:141-150. 81. White SD, Bourdeau PJ, Meredith A. Dermatologic problems of rabbits. Comp Cont Ed for the Practicing Veterinarian. 2003;25:90-103. 82. White SD, Campbell T, Logan A, et al. Lymphoma with cutaneous involvement in three domestic rabbits (Oryctolagus cuniculus). Vet Dermatol. 2000;11:61-67. 83. White SD, Linder KE, Schultheiss P, et al. Sebaceous adenitis in four domestic rabbits (Oryctolagus cuniculus). Vet Dermatol. 2000;11:53-60. 84. Wulf M, Voss A. MRSA in livestock animals—an epidemic waiting to happen? Clin Microbiol Infect. 2008;14:519-521.
CHAPTER
19
Neurologic and Musculoskeletal Diseases
Peter G. Fisher, DVM, and James W. Carpenter, MS, DVM, Diplomate ACZM
Parasitic Encephalitozoonosis Cerebral Larva Migrans Cuterebra Species Toxoplasmosis Bacterial Otitis Interna Bacterial Infections of the Central Nervous System Viral Rabies Herpes Simplex Virus Traumatic Vertebral Fracture or Luxation Degenerative/Developmental Spondylosis and Osteoarthritis Splay Leg Toxic Lead Toxicosis Fipronil Toxicosis Pyrethrin/Permethrin Toxicosis Metabolic Toxemia of Pregnancy Heat Stroke Nutritional Issues Other Causes Neoplastic Vascular Idiopathic Miscellaneous
Neurologic and musculoskeletal diseases are common in the rabbit, and recognition of these conditions is becoming more frequent, in part because many pet rabbits are living longer. Several rabbit neurologic diseases, in particular encephalitozoonosis, have zoonotic potential and therefore have been investigated Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
more thoroughly in recent years, increasing our confidence in diagnosis and possible treatment. The most common causes of neurologic and musculoskeletal diseases in rabbits include otitis interna, pasteurellosis and other bacterial infections, encephalitozoonosis, cerebral larva migrans, cranial or vertebral trauma, spondylosis, heat stress, and toxemia. Signs of neurologic disease in rabbits include behavioral changes, head tilt (labyrinthine torticollis or wry neck), nystagmus, tremors, paresis, paralysis, and seizures. Head tilt, usually an indication of vestibular dysfunction, can be central (cerebellum, brainstem) or peripheral (inner ear) and was the most common clinical sign noted in a retrospective study of rabbits with neurologic disease (Table 19-1).22 Ataxia, paresis, and paralysis can also be caused by central (brain or spinal cord) or peripheral nerve disease. Subtle or overt behavioral changes, such as hyperesthesia, may be caused by central or peripheral disease, while seizures and rolling indicate brain lesions. Differentiation of the causes of encephalomyelitis is challenging, and sometimes more than one cause is present in the same animal.22,24 Along with signalment and history, it is essential to perform a thorough neurologic examination to localize the underlying lesion and determine whether the disease is local or multifocal/diffuse. The neurologic examination will help the clinician form a differential diagnosis list and aid in directing appropriate diagnostic procedures. Detailed information on rabbit neuroanatomy, performing a rabbit neurologic examination, and neurodiagnostics has been previously published.48 Signs of musculoskeletal disease in the rabbit may be associated with pain or decreased function; they include diminished activity, a hunched posture, varying degrees of paresis, paralysis or a stilted gait, and angular limb deformities. The most common musculoskeletal diseases in rabbits are developmental, degenerative, or related to trauma. Diagnostic procedures used to help in narrowing the differential diagnoses for neurologic and musculoskeletal disease include clinical pathology (Fig. 19-1), imaging, and serology. Perform a complete blood count (CBC), serum chemistries, and urinalysis to help identify systemic diseases that may manifest themselves with neurologic signs. Take radiographs in cases of lameness, gait abnormalities, head tilt, and spinal abnormalities. Magnetic resonance imaging (Fig. 19-2) is useful in looking for fine detail, including central nervous system soft tissue, and computed tomography is preferred for evaluation of the inner 245
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SECTION II Rabbits
Table 19-1 Etiology and Clinical Signs in a Retrospective Study of 118 Rabbits Presented with Neurological Diseasea Category Inflammatory Idiopathic Toxic/metabolic Neoplastic Traumatic Vascular Degenerative Totalb Number of patients
Vestibular Other Signs Paresis Seizures Signs 48 2 4 2 2 1 1 60 57
15 6 5 1 2 1 1 31 29
4 1 1 1 1 1 0 9 8
12 7 3 3 0 2 0 27 24
aConducted
at the Institute of Pathology, University of Veterinary Medicine, Vienna.22 bThe total number of patients does not match with 118 animals examined because some animals were classified in two categories simultaneously.
Fig. 19-2 A magnetic resonance image in a rabbit with neurologic disease. This scan was performed because results of more routine diagnostic procedures were nondiagnostic.
Fig. 19-1 A cerebrospinal fluid sample being obtained for culture and cytology from a rabbit with neurologic disease.
ear, skull, and vertebral column. The determination of antibody titers can be useful in looking for infectious etiologies such as encephalitozoonosis, toxoplasmosis, or pasteurellosis.
PARASITIC ENCEPHALITOZOONOSIS Encephalitzoon cuniculi infection is often associated with neurologic disease in pet rabbits. Encephalitzoonosis seems to be a widespread disease in rabbits, with reports of infection found in 50% to 75% of conventional rabbit colonies.36 Signs of neurologic disease caused by E. cuniculi include behavioral changes, head tilt, nystagmus, ataxia, rolling, or seizures and often follow a stressful event in the rabbit’s life. E. cuniculi has shown zoonotic potential especially in immunocompromised humans such as transplant recipients or those infected with human
immunodeficiency virus (HIV) as well as children, travelers, contact lens wearers, and the elderly.13 It may, therefore, be important to know the serologic status of many pet rabbits. E. cuniculi is a microsporidium—an obligate intracellular protozoan parasite. Postnatal transmission often occurs within 6 weeks from an infected dam or contact with other infected animals36; however, vertical transmission is possible.6,12,50 A spore, ingested or inhaled, is the infectious stage of E. cuniculi, with oral ingestion of spores from infected rabbit urine being the most common source of infection. Spores can be found in the urine 1 month after infection and are excreted in large numbers up to 2 months postinfection.23 E. cuniculi spores can survive outside the host for up to 6 weeks at 72°F (22°C). Shedding of spores is essentially terminated by 3 months postinfection. The spore possesses a polar filament that it extrudes into host intestinal mucosal cells, injecting spore contents and initiating infection. Multiplication of the E. cuniculi organism takes place in host alimentary cell vacuoles, with eventual cell rupture and spore invasion of the reticuloendothelial system and systemic circulation by infected macrophages. Initial target organs include those with high blood flow, such as the lungs, liver, and kidney,43 with infection of nervous tissue occurring later in the course of the disease. Further organism multiplication occurs via ordinary fission or schizogony within vacuoles or pseudocysts (schizonts) found in reticuloendothelial cells of target organs. Spores eventually develop and, with time, the pseudocyst becomes overcrowded and ruptures. Cell rupture is associated with a chronic inflammatory response, and most immunocompetent rabbits develop chronic, subclinical infections in a balanced host-parasite relationship associated with granulomatous lesions primarily affecting the brain, kidney, or eyes. A wide range of symptomatology is possible and E. cuniculi has been associated with phacouveitis secondary to lens rupture, myocarditis, vasculitis, and spinal nerve root inflammation in addition to encephalitis, pneumonitis, hepatitis, nephritis, and splenitis.39 Abortion and neonatal deaths have also been attributed to encephalitozoonosis.43
CHAPTER 19 Neurologic and Musculoskeletal Diseases
A
247
B
Fig. 19-3 A, This 8-year-old female spayed rabbit was presented with signs of vestibular disease, including a head tilt and nystagmus. There was no evidence of tympanic bulla disease on skull radiography. The rabbit’s E. cuniculi enzyme-linked immunosorbent (ELISA) test was positive, and it was treated with fenbendazole (20 mg/kg PO × 30 days). B, The vestibular signs resolved with time. With encephalitozoonosis infections in the rabbit, it is difficult to determine whether resolution of clinical signs is the result of specific medical therapy or the body’s natural ability to heal. The most commonly recognized neurologic sign in rabbits infected with E. cuniculi is vestibular disease, with infected rabbits showing varying degrees of head tilt, ataxia, circular movements, and nystagmus (Fig. 19-3). Paresis alone is the second most common neurologic sign. In one study determining the extent of histologic lesions in the brain of naturally infected rabbits, the cerebrum was most frequently affected, followed by the leptomeninges (Fig. 19-4).8 The vestibular cores, cerebellum, and spinal cord were less commonly affected in spite of the fact that vestibular disease is the most common clinical sign. In this study, animals were considered infected with E. cuniculi if spores were present; the use of Ziehl-Neelsen stain was slightly more sensitive for spore detection than acid-fast trichome stain. On histopathology, most rabbits showed perivascular inflammatory infiltrates and, less frequently, granulomas. This study found no significant difference for the presence of central nervous system (CNS) granulomas in rabbits with or without neurologic symptoms and concluded that histologic findings of inflammatory lesions are not always indicative of overt encephalitozoonosis as the cause of neurologic signs. As E. cuniculi organisms spread to various organs, antibodies develop and encapsulation occurs, thus limiting tissue damage and spore excretion. Antibodies become detectable 3 to 4 weeks after infection, with maximum titers occurring 6 to 9 weeks postinfection. A healthy immune system prevents the organisms from multiplying, but the spores remain viable. Immunosuppression, as a result of illness, stress, or aging, may result in overt disease many years after initial infection. Antibodies contribute to resistance by inducing opsonization by macrophages and complement-mediated killing.9 Currently, clinical means of diagnosing definitive antemortem encephalitozoonosis are limited. However, since E. cuniculi infection is persistent, antibodies continue to be produced and, as a general rule, the validity of antibody assays for the detection of E. cuniculi compares favorably with histology in rabbits.20,42,49
Fig. 19-4 Section of cerebrum from a rabbit with encephalitozoonosis. A granulomatous inflammatory focus is shown. Hematoxylin and eosin; bar, 100 μm. The inset shows a parasitophorous vacuole located within the parenchyma without any associated inflammatory reaction. Acid-fast trichrome stain; bar, 17.7 μm. (From Gruber A, Pakozdy A, Weissenböck H, et al. A retrospective study of neurological disease in 118 rabbits. J Comp Pathol. 2009;140:31-37.)
Several methods for detecting antibodies against E. cuniculi are available from commercial laboratories in the United States (Table 19-2). A study comparing two indirect immunofluorescence (IIF) assays, two enzyme-linked immunosorbent assays (ELISA), and the carbon immunoassay (CIA, or India-ink assay) for determination of antibodies to E. cuniculi revealed that any of the assays would be suitable for monitoring rabbits for evidence of infection.2 There was no difference in the ability of any of these assays to detect antibodies in serum, although a
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SECTION II Rabbits Table 19-2 Laboratories That Provide Encephalitozoon cuniculi Testing in the United Statesa Laboratory
Serology
Sound Diagnostics, Enzyme-linked Woodinville, WA immunosorbent assay (ELISA) Taconic Labs, ELISA Rockville, MD Charles River Labs, Multiplexed Wilmington, MA fluorometric immunoassay (MFIA) with either ELISA or immunofluorescent antibody (IFA) as confirmation as needed Comparative ELISA (to determine Pathology Lab, IgG titers) University of and protein Miami, Miami, electrophoresis FL
Specimen Requirements
Shipping Requirements
0.25 mL serum
Does not have to be shipped on ice
0.5 mL serum or whole blood 150 μL diluted serum (1 part serum diluted with 4 parts phosphate buffered saline)
Does not have to be shipped on ice Overnight on ice
0.1 mL serum or heparinized plasma
Does not have to be shipped on ice
aSeveral
methods for detecting antibodies against E. cuniculi include indirect immunofluorescence (IIF), ELISA, and carbon immunoassay (CIA or India-ink assay). Any of these assays is suitable for routine health monitoring of rabbits.
Table 19-3 Percentage of Rabbits Showing a Seropostive Status to E. cuniculi in Three Separate Studiesa Signs of Disease
Deeb, 20049 (n = 1279)
Harcourt-Brown, 200325 (n = 180)
Kunzel, 200833 (n = 224)
Asymptomatic Vestibular signs Paresis, paralysis Renal signs Intraocular lesions
49% 78% 63% 61% 75%
37% 88% 71% 86% 100%
35% 90% 44% 72% 84%
aNote the statistically significant difference between asymptomatic rabbits and those showing neurologic, renal, and ocular signs.
comparison of techniques showed that IIF, rather than ELISA, is best for quantitative measurement of antibodies. These results, however, may be related to the investigators’ reliance on optical density readings from only one serum dilution.7 A positive titer with detection of antibodies does not differentiate between rabbits with an active infection, those with a latent infection, or rabbits that developed an antibody response and are no longer infected. Positive results, therefore, indicate exposure to the organism but do not confirm E. cuniculi as a cause of disease. Follow-up samples may clarify equivocal results where early-stage infection antibody levels will be considerably higher in the 3- to 4-week follow-up sample. The presumptive diagnosis of encephalitozoonosis is made on the basis of signs of neurologic disease together with demonstration of high levels of serum antibodies or of spores in affected tissues. Previous studies profiling the serostatus of rabbits have found a higher incidence of seropositive results in E. cuniculi-suspect
rabbits versus clinically normal rabbits2,7,9,25,33 Asymptomatic rabbits have a variable degree of seropositive status, with two studies from the United Kingdom showing 52%31 and 37%25 positive serostatus, one United States study showing 49% seropositivity,9 and an Austrian study33 revealing 35% seropositive results in clinically normal rabbits (Table 19-3). E. cuniculisuspect rabbits with vestibular signs (head tilt, ataxia, circling, nystagmus, and torticollis) showed the greatest degree of consistent antibody production against E. cuniculi, with 90%,33 78%,9 and 88%25 of rabbits tested being seropositive. Rabbits with other neurologic signs—such as ataxia, paresis, and paralysis— demonstrated 44%,33 63%,9 and 71%25 seroprevalence to E. cuniculi. Rabbits with renal signs (polydipsia, polyuria, perineal scalding, and weight loss plus elevated BUN and creatinine) also demonstrated a significant antibody response with 72%,33 61%,9 and 86%25 being seropositive to E. cuniculi. Finally, rabbits with intraocular lesions (cataracts, phacoclastic uveitis)
CHAPTER 19 Neurologic and Musculoskeletal Diseases demonstrated 84%,33 75%,9 and 100%25 seroprevalence to E. cuniculi (Table 19-3). Other methods advocated for antemortem diagnosis of E. cuniculi include urine antibody analysis and cerebrospinal fluid (CSF) analysis. One study demonstrating an increase in anti-E. cuniculi IgG antibodies in urine samples from asymptomatic seropositive rabbits suggests that the measurement of urine antibodies may be useful in the detection of latent infections in rabbits.17 CSF analysis showed a higher concentration of CSF proteins in rabbits with suspect encephalitozoonosis (based on vestibular disease and/or paresis) compared with normal rabbits.28 Cytologic evaluation of the CSF in these suspect rabbits was characterized by a lymphomonocytic pleocytosis compared with healthy rabbits, which showed low CSF leukocyte counts.28 In human medicine, where E. cuniculi is not an uncommon opportunistic pathogen in immunocompromised patients such as transplant recipients or those infected with human immunodeficiency virus (HIV), polymerase chain reaction (PCR)-based studies have been very successful in identifying E. cuniculi DNA in stool specimens of infected individuals with clinical diarrhea.40 However, studies in rabbits looking for microsporidial DNA to E. cuniculi by PCR in urine samples from those with clinical signs consistent with encephalitozoonosis, significantly high antibody titers to E. cuniculi, and gamma globulinemia on protein electrophoresis did not show statistically significant evidence of the parasite on PCR (C. Cray, personal communication). Another study also found that CSF and urine from rabbits with suspect neurologic and renal encephalitozoonosis, respectively, were negative by PCR for E. cuniculi DNA but did find positive PCR in 4 out of 5 samples of liquefied lens material from rabbits with supposed phacoclastic uveitis.33 These studies confirm that naturally infected rabbits with renal disease or neurologic signs are usually chronically infected, that they either excrete spores for a short period of time postinfection or in low concentrations intermittently, and that PCR-based tests will not prove to be a good antemortem diagnostic tool. In one study involving four rabbits with phacoclastic uveitis, immunohistochemistry was used to identify spores reacting with antiE. cuniculi antiserum.19 Therefore, in cases of E. cuniculi-induced phacoclastic uveitis where enucleation is performed, histology, PCR analysis, and immunohistochemistry may be used to confirm a diagnosis of antemortem E. cuniculi infection. In the absence of controlled studies, it is difficult to assess the efficacy of therapeutic agents against E. cuniculi, because latent infections occur and some clinical cases may improve spontaneously without treatment, presumably as a result of the host’s immune response.23 In addition, clinical signs may not be associated with presence of the protozoan itself but rather with the inflammatory response that persists after the organism has been eliminated.25,47 Treatment protocols for rabbits showing clinical signs suspicious for E. cuniculi infection have been based on fundamental principles of therapy for granulomatous inflammation, on studies demonstrating efficacy against human encephalitozoon infections, and on in vitro susceptibility studies of E. cuniculi organisms to various pharmacologic agents. However, there is still no universal agreement on how to effectively treat encephalitozoonosis in rabbits. Several benzimidazole derivatives including albendazole, oxibendazole, and fenbendazole have been used to treat presumptive E. cuniculi infections in rabbits based on their anti-inflammatory actions and their in vitro antiprotozoal activity, including bioenergetic disruptions of
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membranes and microtubular (tubulin) inhibition of E. cuniculi.15,37 In vitro studies suggest that albendazole can kill the organism and spare the host cell (M. Hawkins, personal communication). In vitro studies have demonstrated that nitazoxide, an antiprotozoal used in equine medicine and advocated on the Internet as an effective treatment option was not effective against the E. cuniculi organism (M. Hawkins, personal communication). Some clinicians advocate the administration of one dose of a short-acting corticosteroid (i.e., dexamethasone, 0.1 mg/kg SC) to control the inflammation associated with the organism when neurologic signs appear acutely. Treatment success of encephalitozoonosis is based on resolution or improvement in clinical signs. The use of albendazole is partially based on its effectiveness in treating and eliminating Encephalitozoon species from human AIDS patients along with relief of clinical symptoms associated with infection.11 One study demonstrated the in vitro efficacy of fumegillin, thiabendazole, oxibendazole, and albendazole in inhibiting E. cuniculi proliferation in rabbit kidney tissue culture cells.15 Another study showed the efficacy of albendazole in reducing serum creatinine levels in rabbits experimentally infected with E. cuniculi microsporidia (however, the study did not measure creatinine levels in a group of untreated control animals, nor did it do histopathology to prove renal infection with the E. cuniculi organism).5 Some veterinarians have had success with albendazole (30 mg/kg PO q24h for 30 days). Albendazole is embryotoxic and teratogenic in rabbits and has been associated with anecdotal reports of pancytopenia, fever, and death in rabbits15; evaluation of an intratreatment CBC is, therefore, recommended. Fenbendazole has also been used as a potentially successful chemotherapeutic and prophylactic treatment of rabbit E. cuniculi infections.46 Administration of fenbendazole at 20 mg/kg PO daily for 30 days resulted in the apparent elimination of E. cuniculi spores from the CNS of infected rabbits and has been used with apparent success by the author (PGF) (Fig. 19-1). Oxibendazole (30 mg/kg PO q24h for 7-14 days, then 15 mg/kg PO q24h for 30-60 days) has been used against rabbit encephalitozoonosis with apparent safety9; however, bone marrow suppression has also been reported anecdotally with oxibendazole; therefore an intratreatment CBC is recommended. In some patients, signs abated during oxibendazole treatment and then recurred after the drug was withdrawn. These patients were then managed on long-term oxibendazole (15-30 mg/kg PO q24h).9 Recent work in mice has shown that a drug designed to block human p38 mitogen-activated protein kinase (MAPK) was successful in treating mice infected with E. cuniculi, suggesting that MAPK inhibitors may represent a novel class of agents with a broad spectrum of antiparasitic activity that may be applied to infection with the Encephalitozoon species.51 Additional therapeutic strategies for human microsporidial infections may someday be applied in the domestic rabbit; these include agents that target the microsporidial polyamines (e.g., polyamine analogs), methionine aminopeptidase 2 (e.g., fumagillin-related compounds), chitin inhibitors (e.g., nikkomycins), topoisomerases (e.g., fluoroquinolones), and tubulin (e.g., benzimidazolerelated compounds).13 Control of encephalitozoonosis may be most effective in rabbitries, where transmission usually occurs. The acute phase of the disease usually occurs in very young rabbits, and spores are shed in urine until antibodies are produced. A simple test
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to determine whether rabbits are shedding spores in the urine is needed to clarify potential transmission duration time. This information could then guide recommendations of quarantine periods when new rabbits are introduced. It may then be possible to select E. cuniculi antibody-negative breeders and to develop encephalitozoon-free rabbit colonies. It must be kept in mind that even if transmission from rabbit to rabbit is prevented, the possibility exists of transmission of spores from exposure to rodents, including mice, hamsters, and guinea pigs. Cleaning and sanitation are essential to limit transmission. Most disinfectants are effective at inactivating spores, including quaternary ammonium compounds, amphoteric surfactants, phenolic derivatives, alcohols, iodophors, and hydrogen peroxide.
CEREBRAL LARVA MIGRANS Cerebral larva migrans due to Baylisascaris species has been reported in rabbits.10,16,18 The history includes an outdoor living environment with possible exposure to eggs of Baylisascaris procyonis or Baylisascaris columnaris, the roundworms carried, respectively, by raccoons and skunks. Raccoons are indigenous to North America, but B. procyonis larva migrans has been reported in small animals kept in close proximity to raccoons in zoo collections in Germany, Ireland, and Japan.16 Raccoons deposit feces in or around edible vegetation, feed, hay, and bedding; these materials, contaminated with Baylisascaris eggs, are ingested by rabbits. Eggs in the feces may remain infective for 1 year or longer. After eggs are ingested, larvae are released in the intestine of the aberrant host and migrate through the body, where they have a predilection for the CNS, particularly the cerebral cortex, midbrain, and brainstem, causing encephalomalacia. Destruction of brain tissue may be extensive, even from only a few larvae, and may be progressive as the larvae continue to migrate in the brain, grow rapidly, and produce metabolic wastes and enzymes, resulting in a severe inflammatory response.4 Signs of disease associated with cerebral larva migrans are tremors, ataxia, head tilt, circling, and vertical nystagmus progressing to falling, rolling, paralysis, seizures, and recumbency. Swaying and falling are usually pronounced with cerebral larva migrans, whereas they are seen less often with encephalitozoonosis or otitis interna. Typically an affected rabbit shows intermittent improvement, followed by more severe signs. Slowing the progression of presumptive cerebral larva migrans by treatment with oxibendazole (60 mg/kg PO q24h) indefinitely has shown some success.9 Signs may recur and progress when the drug is discontinued. Rabbits with severe posterior paresis can be fitted with an apparatus to permit movement. Rabbits that are most severely affected are probably best euthanatized. Advise rabbit owners to prevent cerebral larva migrans by eliminating access to feed and bedding that is potentially contaminated by raccoon feces.
CUTEREBRA SPECIES Larvae of Cuterebra species, or bot flies, commonly pupate in the rabbit subcutis but have also been reported to migrate aberrantly through the ear canals and CNS, causing neurologic signs.27 No specific clinical or clinicopathologic tests are diagnostic for cuterebriasis. This aberrant migration has been reported more
commonly in cats, where clinical abnormalities tend to be progressive and include blindness, depression, and behavioral changes.54 Histopathologic changes reported in the feline suggest entry from the nasal cavity; they also point to a toxic factor elaborated by the parasite as well as potential vascular compromise.54 Could a similar pathogenesis be possible in the rabbit?
TOXOPLASMOSIS Toxoplasmosis is an uncommon cause of neurologic disease in rabbits, and infections are usually subclinical.22 One study measuring serum antibodies to Toxoplasma gondii—by indirect ELISA in domestic rabbits from three rabbit farms in Mexico— demonstrated a seroprevalence in 26.9% of animals tested.14 Signs of CNS infection may include ataxia, tremors, posterior paresis, paralysis, and tetraplegia.18 In most species, clinical toxoplasmosis is often associated with immunosuppression, and although the role of the domestic rabbit in the epidemiology in humans has not been established in detail, some studies suggest potential zoonosis.45 Toxoplasma may induce granulomatous meningoencephalitis similar to encephalitozoonosis, but foci of necrosis as well as tachyzoites may be found in many organs including skeletal muscle, spleen, heart, lung, and lymph nodes.22 This disease can be differentiated from encephalitozoonosis by serologic testing and histologically by demonstration of E. cuniculi spores in brain tissue and immunohistochemical labeling.22 If toxoplasmosis is diagnosed, treat with trimethoprim- sulfamethoxazole and pyrimethamine or doxycycline. Clindamycin should not be used because it causes gastrointestinal dysbiosis and death in rabbits. Toxoplasma gondii may be transmitted in herbivores congenitally or by ingesting oocysts from infected cat feces. Prevent toxoplasmosis in rabbits by avoiding exposure to outdoor grazing areas, feed, or bedding contaminated by cat feces.
BACTERIAL OTITIS INTERNA Vestibular signs resulting from otitis interna, including nystagmus and a head tilt of varying severity with loss of balance and rolling, have often been described and historically associated with Pasteurella multocida infection. However, one of the authors (PGF) has more commonly cultured other bacterial agents from cases of otitis media or otitis interna in rabbits, including S. aureus, Streptococcus species, Bordetella bronchiseptica, Pseudomonas aeruginosa, Escherichia coli, and Proteus mirabilis. A primary otitis media may be caused by infection spreading from the upper respiratory tract through the auditory or eustacian tubes; a concurrent sinusitis/rhinitis is suggestive of this pathogenesis. Infection may also spread from the middle ear to the ear canal (if the tympanic membrane ruptures) or to the inner ear, causing vestibular labyrinthitis; it can also sometimes spread to the brain, causing severe neurologic signs, including seizures. (For a more detailed discussion of pasteurellosis and otitis media, see Chapter 16.) Diagnosis of otitis media/interna is based on clinical signs, aural exam, and imaging, including skull radiography and computed tomography (CT) or magnetic resonance imaging (MRI) where available. Bacterial culture and sensitivity of deep aural or nasal swabs taken under anesthesia are indicated when physical exam supports infection. Endoscopic exam performed under general anesthesia aids in
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VIRAL RABIES Rabies has been reported in rabbits, albeit rarely. Between 1971 and 1997, a total of 30 cases of rabies in rabbits were reported to the Centers for Disease Control and Prevention.30 Most of these were privately owned domestic rabbits located in states where the raccoon variant of rabies was enzootic or epizootic. Infected rabbits usually develop varying degrees of paralysis or paresis in one or more limbs but also may develop head tremors. Early signs of rabies in rabbits may be nonspecific and include anorexia, fever, and lethargy.30 No rabies vaccine is approved for use in rabbits in the United States; therefore pet rabbits housed outdoors should be protected from contact with wildlife, especially in areas where rabies is endemic or epizootic.
HERPES SIMPLEX VIRUS Fig. 19-5 Normal endoscopic appearance of the tympanic membrane in a 1.5-kg dwarf rabbit. (With permission from Vittorio Capello, DVM.)
visualizing the distal ear canal and tympanic membrane (Fig. 19-5). In some cases, rabbits with severe otitis media may have soft tissue swelling at the base of the ear canal, apparently a result of the buildup of purulent material within the ear. The diagnosis of encephalitis related to bacterial extension from the middle ear is made histologically. Antimicrobial treatment of otitis interna should be given over the long term, 4 to 6 weeks or longer, as antibiotics do not penetrate well into pus-filled tympanic bullae. Systemic antibiotics, preferably based on the results of a bacterial culture and their ability to penetrate the CNS, are recommended. Quinolone antibiotics such as enrofloxacin, ciprofloxacin, and marbofloxacin are common choices. In addition to antibiotic therapy, affected rabbits often benefit from nutritional supplementation, environmental support to minimize the rolling and severe ataxia associated with this disease, and medical therapy with meclizine hydrochloride, an antihistamine that aids in the control of the vestibular disease-associated dizziness and nausea. In cases of severe disease, anesthetize the rabbit and gently flush the ear canal with warm saline solution by passing a small, 3.5-Fr red rubber catheter into the ear canal. For cases of severe otitis interna/media that are not responsive to medical therapy and where inspissated pus accumulation in the tympanic bulla is suspected, surgical treatment via total ear canal ablation and ostectomy of the tympanic bulla has been described.3
BACTERIAL INFECTIONS OF THE CENTRAL NERVOUS SYSTEM Primary bacterial suppurative encephalomyelitis has been reported in the rabbit, with both Staphylococcus species22 and P. multocida38 being cultured as causative agents. Bacteria may reach the CNS via hematogenous extension or some other focus of infection, such as the nasal cavity. However, in one retrospective study,22 over half the cases of rabbits with suppurative encephalomyelitis had concurrent otitis media, making extension of bacterial otitis to the CNS a common underlying etiology.
Herpes simplex virus (HSV) encephalitis is rare in the rabbit. Published reports of spontaneous infections21,37a,53 have been associated with close contact between affected rabbits and humans with herpes labialis. This observation suggests that transmission from humans to rabbits is the most likely source of infection. Histologic examination of neural tissues from affected rabbits revealed nonsuppurative meningoencephalitis with multifocal neuronal degeneration and necrosis, mainly located in the gray matter of the cerebrum and cerebellum. Infected rabbits present with acute conjunctivitis followed by signs of CNS dysfunction such as circling, ataxia, seizures and/or ospthisthotonus. Diagnoses have been confirmed by the histologic presence of numerous intranuclear inclusions in neurons and glial cells. Immunohistochemistry and in situ hybridization identified these viruses with human HSV, and in one case HSV-1 infection was proven by PCR analysis.22
TRAUMATIC VERTEBRAL FRACTURE OR LUXATION The most common cause of acute posterior paralysis is vertebral fracture or luxation (Fig. 19-6). Fractures are more common than dislocations. The most common site of fracture (or luxation) is the lumbosacral region (L6-L7). This injury can occur in domestic rabbits that are startled or frightened and the heavily muscled hindquarters are allowed to twist about the lumbosacral junction, which acts as a fulcrum in applying leverage to the vertebral column.18 This traumatic injury has often been associated with inappropriate handling, but the authors have seen even properly restrained rabbits suddenly flinch and fracture their lumbar vertebrae. In addition to paraplegia, neurologic signs may include loss of deep pain reflexes and skin sensation as well as loss of motor control of the urinary bladder and anal sphincter, depending on the amount of compromise to the spinal cord. The clinical diagnosis of posterior paresis or paralysis due to vertebral trauma is confirmed radiographically. Prognosis depends on the site and severity of the lesion and clinical signs. High-dose glucocorticosteroid therapy, such as methylprednisolone sodium succinate, for treatment of CNS trauma is controversial. The current clinical and pharmacologic evidence in humans and dogs suggests that there is no benefit to glucocorticosteroid administration in cases of acute CNS trauma and that the potential
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SECTION II Rabbits cause spinal cord compression and paresis and can mimic spinal fractures.
DEGENERATIVE/DEVELOPMENTAL SPONDYLOSIS AND OSTEOARTHRITIS Age-related osteoarthritis (degenerative joint disease) and spondylosis (degenerative spinal defects) are now more commonly encountered as improved husbandry and nutrition have extended the lives of pet rabbits. Owners of affected rabbits typically report an abnormal gait, lameness, or inability to hop. The signs of musculoskeletal pain may be more subtle, with the quiet and immobile nature of affected rabbits being attributed to age. Still other rabbits demonstrate lack of proper grooming owing to a reluctance or inability to reach the caudal body and perineum; this results in an unkempt coat and/or perineal soiling and dermatitis. Radiographs of joints with degenerative disease may show capsular distension, osteophytosis, narrowing joint spaces, or surrounding soft tissue thickening or mineralization. Spondylosis is identified radiographically by spinal exostoses, degenerative changes, and spinal defects such as kyphosis, lordosis, or scoliosis (Fig. 19-9). Rabbits with pain and inflammation due to degenerative musculoskeletal conditions often show remarkable improvement when they are treated with analgesics such as nonsteroidal anti-inflammatory drugs or a centrally acting opiate agonist such as tramadol. Perineal scald treatment involves long-term hygiene maintenance with cleansing of affected areas as well as topical and systemic antibiotics to control infection.
A
SPLAY LEG
B Fig. 19-6 Lateral (A) and ventrodorsal (B) radiographs of an 8-year-old mixed-breed rabbit showing a fracture and luxation of the spine at L3-L4 (arrow). Intervertebral spaces L2, L3, L4, and L5 are compressed. The injury resulted from a fall. harmful effects (e.g., gastric hemorrhage, intestinal necrosis, sepsis) are serious. The same can probably be presumed of the rabbit. Moderately affected rabbits may respond to conservative medical management, including cage rest for several weeks, if the spinal cord is not transected (Fig. 19-7). Medical therapy depends on the severity of the condition and usually includes administration of analgesics and nonsteroidal anti-inflammatory drugs. Supportive therapy must include assessment of bladder function and control and manual expression as indicated. Decubitus ulcers often develop, and the perineum may become soiled with urine and feces. Many rabbit owners are very dedicated to their rabbits, and as long as quality of life issues are addressed, some animals with severe posterior paresis can be fitted with an apparatus to permit movement (Fig. 19-8). In rabbits with paralysis and lack of bladder function, euthanasia is often indicated. The extrusion of intervertebral disc material into the vertebral canal, particularly in the area of the lumbar spine, can also
Splay leg, a developmental musculoskeletal condition, occurs in young rabbits, ranging in age from a few days to a few months, or in sedentary senior rabbits. The condition may be related to housing on smooth, slippery flooring that results in the inability to adduct one or more limbs and the subsequent inability to ambulate effectively. The condition may be relatively mild, allowing some clumsy movement, or so severe that the animal is completely unable to walk. The hind-limb anatomy is more commonly affected in juvenile rabbits, resulting in various degrees of flattening and reduction of the femoral head, subluxation of the hip, valgus deformity, and patellar luxation.41 If this condition is diagnosed early in a young, growing rabbit, the author (PGF) has been able to reverse the splay deformity through splinting and hobbling of the affected limbs into a more appropriate anatomic position. Success requires frequent splint changes and anatomic assessment along with improved traction of bedding or flooring. Because this condition may also have a genetic component in a simple autosomal recessive pattern, breeding from the affected animal and its parents should be discouraged. Abduction or splaying of the front limbs may also be seen in sedentary senior rabbits. This may occur as the result of inactivity and consequent muscle wasting, so that over time the front limbs are not adducted under the body and splay out. Obesity and osteoarthritis probably contribute to this condition, which, once started, is hard to reverse. Prevention or diminishing progression involves trying to improve muscle tone through increased activity, housing on a surface that allows for firm footing, and treatment of any underlying predisposing conditions.
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B
A
Fig. 19-7 This 5-month-old male rabbit was presented for acute onset of severe rear limb paresis after being dropped. Lateral (A) and ventrodorsal (B) spinal radiographs revealed a compression fracture of L5-6 (arrow). Treatment included nonsteroidal anti-inflammatory therapy (meloxicam), manual expression of the urinary bladder for several days along with cage rest, and padding to keep the rabbit in a normal resting position with legs tucked under the body. Within 3 weeks the rabbit made a good recovery, with mild ataxia and rear gait abnormalities remaining.
Fig. 19-8 A rabbit with posterior paralysis resulting from luxation of the spine. The patient was fitted with a cart, permitting daily exercise periods.
TOXIC LEAD TOXICOSIS Lead toxicosis can cause neurologic signs in rabbits, including seizures, torticollis, and blindness,29 although anemia, nonspecific anorexia, depression, loss of condition, and gastrointestinal stasis are more characteristic. Signs result from oxygen deprivation and CNS edema. Potential sources of lead contamination
include household paint (in houses built prior to 1974, after which lead paint was no longer available), improperly glazed ceramics, linoleum, golf balls, and metallic objects. Lead poisoning should be suspected in rabbits that have the tendency to chew at baseboards, flooring, plasterboard, or metallic objects. Radiographs showing gastrointestinal metallic densities, blood smears with evidence of erythrocytes that are nucleated or have basophilic stippling, and the measurement of blood lead levels aid in diagnosis, with levels greater than 10 μg/dL considered diagnostic for lead poisoning. Treat with calcium ethylenediaminetetraacetic acid (CaEDTA) (30 mg/kg SC q12h for 5-7 days). Although no veterinary-labeled product is available, CaEDTA is available as a human-labeled product (Calcium Disodium Versenate, 3M Pharmaceuticals, St. Paul, MN) and may be available from compounding pharmacies. Blood lead levels should be repeated posttreatment, as two courses of treatment 1 week apart may be required. Adjustable dog fencing works as a portable barrier within which to exercise house rabbits when they are not chaperoned and thereby to prevent exposure to lead-containing objects in the home.
FIPRONIL TOXICOSIS Fipronil (Frontline, Bayer Animal Health, Shawnee Mission, KS) is a topical ectoparasiticide approved for use in dogs and cats. Multiple reports exist of rabbits being intentionally dosed with fipronil and within 24 hours showing signs of anorexia and lethargy, with or without seizures, leading to possible death. The prognosis for recovery is poor in rabbits already showing clinical signs. Bathing along with oral activated charcoal should
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A
B Fig. 19-9 A 6-year-old female spayed rabbit presented with a history of progressive rear limb weakness and a recent onset of perineal soiling. Lateral (A) and ventrodorsal (B) spinal radiographs showed severe thoracic lordosis at T9-11 (arrow). Spondylosis can contribute to gait abnormalities and the inability to flex the spine in order to groom the caudal body, resulting in an unkempt fur coat and perineal soiling.
be used to limit any additional absorption of the drug from topical application or grooming ingestion, and midazolam or diazepam should be administered to control seizures. Supportive care including intravenous fluid administration and syringe feeding is used as needed.
PYRETHRIN/PERMETHRIN TOXICOSIS Pyrethrin or permethrin topical ectoparasiticides are widely available over the counter; they may cause anorexia, lethargy, muscle twitching, and seizures when intentionally applied to rabbits. Control of further toxicity and treatment of clinical signs are similar to the measures outlined above for fipronil toxicity.
METABOLIC TOXEMIA OF PREGNANCY Pregnancy toxemia can produce neurologic signs in rabbits. Although toxemia is primarily a problem of late gestation, it may also occur in postpartum and pseudopregnant does.9 Neurologic signs may include weakness, depression, incoordination, convulsions, and coma. Death may occur within a few hours after the signs are first noted. Obesity and diminished dietary intake are predisposing factors. Therapy involves monitoring and treating electrolyte disturbances as well as the associated ketosis with intravenous administration of appropriate crystalloids with 5% glucose solution. Prevent toxemia by avoiding fasting and obesity in pregnant does and by providing a highenergy, palatable diet during late gestation.
HEAT STROKE Rabbits are particularly susceptible to heat stroke or heat stress. Decreased cerebral blood flow (as reflected by prolonged cerebral circulation time) and decreased cerebral perfusion pressure
are seen during the onset of heat stroke, resulting in cerebral ischemia and edema.34 Neurologic signs may include weakness, depression, incoordination, convulsions, and coma and are accompanied by an elevation in rectal temperature to greater than 105°F (40.5°C). Treatment includes slowly reducing the body temperature by spraying the rabbit or immersing it in tepid water or wrapping it in cool wet towels, being careful to monitor body temperature and discontinue cooling as the rectal temperature approaches 102°F (38.9°C). Treatment may also involve the control of seizures and mannitol to reduce cerebral edema. Give intravenous fluids and supportive care as needed, with special attention to monitoring for secondary gastrointestinal stasis, metabolic abnormalities, or renal failure. Pet rabbits housed outdoors during the summer, when the ambient temperature can exceed 85°F (29.4°C), require shade, good ventilation, and an adequate supply of cool drinking water.
NUTRITIONAL ISSUES Nutritional muscular dystrophy with clinical paresis in rabbits is associated with hypovitaminosis E and is characterized by degeneration and necrosis of skeletal muscle myofibers. Rabbits with vitamin E deficiency have increased plasma concentrations of creatine phosphokinase and cholesterol. Prolonged storage of feed adversely affects the vitamin E content; therefore feeding pet rabbits fresh feed or supplementing the diet with an alternative vitamin E source such as wheat germ and leafy green vegetables should prevent this problem. The restriction of dietary calcium, phosphorus, or vitamin D has been associated with deleterious effects on the skeletons both of young, growing rabbits and of adult females as a result of nutritional hyperparathyroidism.44 Young rabbits exhibit enlarged joints, crooked legs, arched backs, and enlarged rib costochondral junctions. Adults may experience demineralization of the bones, resulting in fragile bones and pathologic fractures. Hypocalcemia in lactating does has been associated
CHAPTER 19 Neurologic and Musculoskeletal Diseases with generalized musculoskeletal weakness, paresis, and lateral recumbency. Other neurologic and musculoskeletal diseases in rabbits may also have a nutritional cause. Hypovitaminosis A may cause a neurologic disturbance in rabbits characterized by circling, convulsions, opisthotonos, and paralysis. Hydrocephalus has been observed in young rabbits born to does with hypo- or hypervitaminosis A. Convulsions can occur in rabbits maintained on a diet that is deficient in magnesium; ataxia or muscular weakness may be a manifestation of hypokalemia. With the current rigid quality-control standards for commercial feed production, confirmed nutritional problems are relatively rare.
OTHER CAUSES NEOPLASTIC Lymphoma is reported to be the second most commonly occurring tumor in rabbits after uterine adenocarcinoma and is the most common cause of CNS neoplasia in rabbits.52 Although no specific treatment protocols have been published, various chemotherapy drugs have been studied and used in the rabbit, but clients should understand that side effects, including death, may occur.26 Other tumors that have been reported within the CNS of rabbits include one teratoma1 and one ependymoma.32 Both skeletal and extraskeletal osteosarcoma have been reported in rabbits but are rare.26 No treatment protocols have been described, but—as with other species with osteosarcoma of the long bones—limb amputation can extend a good quality of life in affected rabbits.
VASCULAR Neurologic signs may be reported in rabbits secondary to CNS hypoxia resulting from primary disease of the vascular system or heart. Cerebrovascular accidents have not been documented but have been suspected in older rabbits that develop sudden unilateral loss of motor function. If other causes of neurologic disease have been ruled out, a diagnosis of stroke may be considered. Supportive care allows recovery of function in many cases.
IDIOPATHIC In one study of 118 rabbits with varying signs of neurologic disease, including vestibular signs, paresis, and seizures, specific testing failed to identify an etiology for the neurologic signs in 16 rabbits.22
MISCELLANEOUS A variety of other conditions can cause generalized muscular weakness, incoordination, and other neuromusculoskeletal signs in rabbits. Ulcerative pododermatitis is a relatively common musculoskeletal problem and is discussed in Chapter 18. Neurologic signs in rabbits have been associated with listeriosis. In addition to the causes mentioned in this chapter, seizures in the rabbit may also be caused by hypoxia secondary to empyema, pneumonia, terminal stages of hepatic lipidosis, or metastatic tumors or be associated with the azotemia and electrolyte disturbances of renal disease.4 A variety of heritable disorders causing neuromuscular or skeletal diseases in rabbits have been reported35; they include hereditary
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ataxia (a glycogen storage disorder characterized by neurologic signs such as nystagmus, opisthotonos, and paddling), tremors, syringomyelia, and various skeletal deformities (e.g., achondroplasia).
ACKNOWLEDGMENT The authors would like to dedicate this chapter to the memory of Dr. Barbara J. Deeb (1937-2005), one of the pioneers of original and scientific research involving exotic mammals. A compassionate practicing veterinarian, she touched the lives of all who were blessed and honored to know her.
References 1. Bishop L. Intracranial teratoma in a domestic rabbit. Vet Pathol. 1978;15:525-530. 2. Boot R, Hansen AK, Hansen CK, et al. Comparison of assays for antibodies to Encephalitozoon cuniculi in rabbits. Lab Anim. 2000;34:281-289. 3. Capello V. Surgical treatment of otitis externa and media in pet rabbits. Exot DVM. 2004;6:15-21. 4. Carpenter JW. Diagnosing and treating common neurologic diseases in rabbits. Vet Med. 2006:728-736. 5. Conkova E, Cellarova, Neuschl J, et al. The dynamics of creatinine and urea concentrations in the blood serum of rabbits infected by Encephalitozoon cuniculi microsporidium and treated with albendazole. Acta Veterinaria (Beograd). 1999;49:321-326. 6. Couzinet S, Cejas E, Schittny J, et al. Phagocytic uptake of Encephalitozoon cuniculi by nonprofessional phagocytes. Infect Immun. 2000;68:6939-6945. 7. Cray C, Arcia G, Schneider R, et al. Evaluation of the usefulness of an ELISA and protein electrophoresis in the diagnosis of Encephalitozoon cuniculi infection in rabbits. Am J Vet Res. 2009;70:478-482. 8. Csokai A, Gruber A, Künzel F, et al. Encephalitozoonosis in pet rabbits (Oryctolagus cuniculus): pathohistological findings in animals with latent infection versus clinical manifestation. Parasitol Res. 2009;104:629-635. 9. Deeb BJ, Carpenter JW. Neurologic and musculoskeletal diseases. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2004:203-210. 10. Deeb BJ, DiGiacomo RF. Cerebral larva migrans caused by Baylisascaris sp. in pet rabbits. J Am Vet Med Assoc. 1994;205:1744-1747. 11. De Groote MA, Visvesvara G, Wilson ML, et al. Polymerase chain reaction and culture confirmation of disseminated Encephalitozoon cuniculi in a patient with AIDS: successful therapy with albendazole. J Infect Dis. 1995;171:1375-1378. 12. Didier ES. Microsporidiosis. Clin Infect Dis. 1998;27:1-8. 13. Didier ES, Maddry JA, Brindley PJ, et al. Therapeutic strategies for human microsporidia infections. Expert Rev Antiinfect Ther. 2005;3:419-434. 14. Fiqueroa-Castillo JA, Duarte-Rosas V, Juarez-Acevedo M, et al. Prevalence of Toxoplasma gondii antibodies in rabbits (Oryctolagus cuniculus) from Mexico. J Parasitol. 2006;92:394-395. 15. Franssen FF, Lumeij JT, van Knapen F. Susceptibility of Encephalitozoon cuniculi to several drugs in vitro. Antimicrob Agents Chemother. 1995;39:1265-1268. 16. Furuoka H, Sato H, Kubo M, et al. Neuropathological observation of rabbits (Oryctolagus cuniculus) affected with raccoon roundworm (Baylisascaris procyonis) larva migrans in Japan. J Vet Med Sci. 2003;65:695-699. 17. Furuya K, Asakura T, Igarashi M, et al. Microsporidian Encephalitozoon cuniculi antibodies in rabbit urine samples. Vet Rec. 2009;165:85-86.
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18. Gentz E, Carpenter JW. Neurologic and musculoskeletal disease. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders; 1997:220-226. 19. Giordano C, Weight A, Vercelli A, et al. Immunohistochemical identification of Encephalitozoon cuniculi in phacoclastic uveitis in four rabbits. Vet Ophthalmol. 2005;8:271-275. 20. Greenstein G, Drozdowicz CK, Garcia FG, et al. The incidence of Encephalitozoon cuniculi in a commercial barrier-maintained rabbit breeding colony. Lab Anim. 1991;25:287-290. 21. Grest P, Albicker P, Hoezle L, et al. Herpes simplex encephalitis in a domestic rabbit (Oryctolagus cuniculus). J Comp Pathol. 2002;126:308-311. 22. Gruber A, Pakozdy A, Weissenböck H, et al. A retrospective study of neurological disease in 118 rabbits. J Comp Pathol. 2009;140:31-37. 23. Harcourt-Brown FM. Infectious diseases of domestic rabbits. Textbook of rabbit medicine. Oxford, UK: ButterworthHeinemann; 2002:361-385. 24. Harcourt-Brown FM. Neurological and locomotor disorders. Textbook of rabbit medicine. Oxford, UK: ButterworthHeinemann; 2002:307-323. 25. Harcourt-Brown FM, Holloway HKR. Encephalitozoon cuniculi in pet rabbits. Vet Rec. 2003;152:427-431. 26. Heatley JJ, Smith AN. Spontaneous neoplasms of rabbits. Vet Clin North Am Exot Anim Pract. 2004;7:561-577. 27. Hendrix CM, DiPinto LN, Cox NR, et al. Aberrant intracranial myiasis caused by larval Cuterebra migration. Comp Cont Ed. 1989;11:550-559. 28. Jass A, Matiasek K, Henke J, et al. Analysis of cerebrospinal fluid in healthy rabbits and rabbits with clinically suspected encephalitozoonosis. Vet Rec. 2008;162:618-622. 29. Johnston MS. Clinical toxicoses of domestic rabbits. Vet Clin North Am Exot Anim Pract. 2008;11:315-326. 30. Karp BE, Ball NE, Scott CR, et al. Rabies in two privately owned domestic rabbits. J Am Vet Med Assoc. 1999;215:1824-1827. 31. Keeble EJ, Shaw DJ. Seroprevalence of antibodies to Encephalitozoon cuniculi in domestic rabbits in the United Kingdom. Vet Rec. 2006;158:539-544. 32. Kinkler RJ, Jepsen PL. Ependymoma in a rabbit. Lab Anim Sci. 1979;29:255-256. 33. Künzel F, Gruber A, Tichy A, et al. Clinical symptoms and diagnosis of encephalitozoonosis in pet rabbits. Vet Parasitol. 2008;151:115-124. 34. Lin MT, Lin SZ. Cerebral ischemia is the main cause for the onset of heat stroke syndrome in rabbits. Experientia. 1992;48:225-227. 35. Lindsey JR, Fox RR. Inherited diseases and variations. In: Weisbroth SH, Flatt RE, Kraus AL, eds. The biology of the laboratory rabbit. New York: Academic Press; 1974:379-382. 36. Lyngset A. A survey of serum antibodies to Encephalitozoon cuniculi in breeding rabbits and their young. Lab Anim Sci. 1980;30:558-561. 37. McCracken RO, Stillwell WH. A possible biochemical mode of action for benzimidazole anthelmintics. Int J Parasitol. 1991;21:99-104.
37a. Muller K, Fuchs W, Heblinski N, et al. Encephalitis in a rabbit caused by human herpesvirus-1. J Am Vet Med Assoc. 2009;235:66-69. 38. Murray KA, Hobbs BA, Griffith JW. Acute menigoencephalomyelitis in a rabbit infected with Pasteurella multocida. Lab Anim Sci. 1985;35:169-171. 39. Nast R, Middleton DM, Wheler CL. Generalized encephalitozoonosis in a Jersey wooly rabbit. Can Vet J. 1996;37:303-305. 40. Notermans DW, Peek R, de Jong MD, et al. Detection and identification of Enterocytozoon bieneusi and Encephalitozoon species in stool and urine specimens by PCR and differential hybridization. J Clin Microbiol. 2005;43:610-614. 41. Owiny JR, Vandewoude S, Painter JT, et al. Hip dysplasia in rabbits: association with nest box flooring. Comp Med. 2001;51:85-88. 42. Pakes SP, Shadduck JA, Feldman DB, et al. Comparison of tests for the diagnosis of spontaneous encephalitozoonosis in rabbits. Lab Anim Sci. 1984;34:356-359. 43. Percy DH, Barthold SW. Rabbit. In: Pathology of laboratory rodents and rabbits. Ames: Blackwell Publishing; 2007:253-307. 44. Redrobe S. Calcium metabolism in rabbits. Semin Avian Exot Pet Med. 2002;11:94-101. 45. Sroka J, Zwolinski J, Dutkiewicz J, et al. Toxoplasmosis in rabbits confirmed by strain isolation: a potential risk of infection among agricultural workers. Ann Agric Environ Med. 2003;10:125-128. 46. Suter C, Müller-Doblies UU, Hatt JM, et al. Prevention and treatment of Encephalitozoon cuniculi infection in rabbits with fenbendazole. Vet Rec. 2001;148:478-480. 47. Valencakova A, Balent P, Petrovova E, et al. Encephalitozoonosis in household pet Nederland dwarf rabbits (Oryctolagus cuniculus). Vet Parasitol. 2008;153:265-269. 48. Vernau KM, Osofsky A, LeCounteur RA. The neurological examination and lesion localization in the companion rabbit (Oryctolagus cuniculus). Vet Clin North Am Exot Anim Pract. 2007;10:731-758. 49. Waller T, Morein B, Fabiansson E. Humoral immune response to infection with Encephalitozoon cuniculi in rabbits. Lab Anim. 1978;12:145-148. 50. Wasson K, Peper RL. Mammalian microsporidiosis. Vet Pathol. 2000;37:113-128. 51. Wei S, Daniel BJ, Brumlik MJ, et al. Drugs designed to inhibit human p38 mitogen-activated protein kinase activation treat Toxoplasma gondii and Encephalitozoon cuniculi infection. Antimicrob Agents Chemother. 2007;51:4324-4328. 52. Weisbroth SH. Neoplastic diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. New York: Academic Press; 1994:259-292. 53. Weissenbock H, Hainfellner JA, Berger J, et al. Naturally occurring herpes simplex encephalitis in a domestic rabbit (Oryctolagus cuniculus). Vet Pathol. 1997;34:44-47. 54. Williams KJ, Summer BA, de Lahunta A. Cerebrospinal cuterebriasis in cats and its association with feline ischemic encephalopathy. Vet Pathol. 1998;35:330-343.
CHAPTER
20
Cardiovascular Disease, Lymphoproliferative Disorders, and Thymomas
Sharon M. Huston, DVM, Diplomate ACVIM (Cardiology), Pamela Ming-Show Lee, DVM, MS, Katherine E. Quesenberry, DVM, MPH, Diplomate ABVP (Avian), and Anthony A. Pilny, DVM, Diplomate ABVP (Avian)
PART I CARDIOVASCULAR DISEASES Cardiac Disease Normal Cardiovascular Structure Examination of the Rabbit with Cardiovascular Disease Diagnostic Methods Radiography Electrocardiography Echocardiography Diseases and Management Congestive Heart Failure Congenital Heart Disease Arrhythmia Myocardial Disease Valvular Disease Vascular Disease PART II LYMPHOPROLIFERATIVE DISORDERS AND THYMOMAS Etiology Genetic Factors Infectious Factors Types of Lymphoproliferative Disorders Multicentric Lymphoma Cutaneous Lymphoma Leukemia Thymic Masses: Thymoma/Thymic Lymphoma/ Thymic Carcinoma Diagnosis Treatment Chemotherapy Treatment Options for Thymomas Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
PART I CARDIOVASCULAR DISEASES
CARDIAC DISEASE Cardiac disease has become increasingly recognized in domestic rabbits. Despite their frequent use as laboratory models for cardiac disease in humans, little is known about the pathogenesis and treatment of naturally occurring heart disease in rabbits. Reports of spontaneous heart disease are sporadic and case numbers are often low. Despite this, a complete cardiac evaluation and systematic approach can lead to a correct diagnosis and successful treatment of rabbits with cardiovascular disease.
NORMAL CARDIOVASCULAR STRUCTURE The rabbit heart is unique in several ways: the tricuspid valve is composed of two rather than three cusps; the aortic nerve is not associated with chemoreceptors but only with baroreceptors; the rabbit pulmonary artery and its branches are heavily muscular6; and a persistent left cranial vena cava is also normally found and drains into the coronary sinus.25 Additionally, the myocardium has limited collateral circulation and is therefore predisposed to ischemia mediated by coronary vasoconstriction.36
EXAMINATION OF THE RABBIT WITH CARDIOVASCULAR DISEASE Information gathered from a thoughtful history and complete physical examination comprises the most important part of the cardiac evaluation.
History A thorough general history, including husbandry, diet, and past or present illnesses, should be obtained for rabbits suspected of having cardiac disease. Tachypnea, dyspnea, syncope, 257
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A
B Fig. 20-1 Normal thoracic radiographs of a 6-year-old rabbit. A, Right lateral projection. B, Ventrodorsal projection.
anorexia, weight loss, and malaise may be signs of heart disease in rabbits.
Physical Examination Take care to minimize the handling of a rabbit in cardiac distress; a complete physical examination and further diagnostic testing must sometimes be delayed until the rabbit is clinically stable. Stress can be minimized by examining the rabbit in a quiet room and moving slowly. Rabbits are a prey species and thereby very sensitive to external stimuli. If necessary, sedatives may also be given to minimize the risk of self-injury. Sedation can be achieved with midazolam 0.5 to 1 mg/kg intramuscularly45 or, for heavier sedation, a combination of ketamine 5 to 20 mg/kg and midazolam 1 to 2 mg/kg subcutaneously13 with minimal cardiovascular depression (see Chapters 31 and Appendix). Provide flow-by oxygen as needed if a rabbit is dyspneic or becomes tachypneic with handling. In rabbits suspected of having cardiac disease or respiratory distress, focus the initial examination on observing the respiratory rate and pattern, obtaining the heart rate and rhythm, auscultating the thorax, examining the mucous membranes, and palpating the pulses. Normal heart rate is 180 to 250 beats per minute and normal respiratory rate is 30 to 60 breaths per minute. Peripheral pulses can be palpated at the central artery of the ear and should be strong and synchronous with the heartbeat. Two heart sounds (S1 and S2) are normally heard. Rabbits are obligate nose breathers; therefore do not occlude the nares. Rabbits with heart disease may have cyanotic or pale mucous membranes, arrhythmias, or heart murmurs. Rabbits with congestive heart failure often have tachycardia, tachypnea, and labored breathing. Pulses may be irregular or weak. Systematic auscultation of the thorax is the most important part of the cardiac examination. The entire thorax should be auscultated to localize heart murmurs and detect extra heart sounds (gallop sounds), arrhythmias, and abnormal lung sounds. A pediatric stethoscope allows better localization of heart sounds in rabbits and is preferred for cardiac auscultation; a larger-diaphragm stethoscope enhances auscultation of the lungs. Auscultatory findings vary among rabbits with congestive heart failure, and these findings are not pathognomonic. The examiner may hear muffled heart and lung sounds with
pleural effusion and increased bronchial sounds or crackles with pulmonary edema.
DIAGNOSTIC METHODS Diagnosis of cardiovascular disease is based on a complete history and physical examination findings complemented by appropriate diagnostic tests. Thoracic radiographs, electrocardiography, echocardiography, and routine blood tests are useful in reaching a definitive diagnosis and treatment plan.
RADIOGRAPHY Thoracic radiography provides critical information in the patient with cardiopulmonary disease—namely cardiac shape and size, pulmonary pattern, vascular pattern, and other thoracic lesions. Congestive heart failure and respiratory disease can be differentiated by evaluating thoracic radiographs. Radiographic findings supporting a diagnosis of cardiac disease are similar to those in other species and include cardiac enlargement, pulmonary vascular enlargement, pulmonary interstitial and alveolar pulmonary pattern of pulmonary edema, and pleural effusion. Figures 20-1 and 20-2 show thoracic radiographs of a normal adult rabbit and a rabbit with heart disease, respectively.
ELECTROCARDIOGRAPHY Electrocardiography (ECG) is a simple and practical diagnostic test in rabbits with suspected or confirmed cardiac disease. An ECG is critical to diagnose and manage arrhythmias or syncope. The ECG may also be a helpful addition to the cardiac database. However, one should not use an ECG to assess or detect chamber enlargement or hypertrophy. Record the ECG with the rabbit in sternal recumbency. File the alligator-style ECG clips to minimize skin trauma, and place medical-grade alcohol or electrode gel on the clips to enhance conduction. Record tracings with a vertical calibration of 2 cm/mV and a horizontal paper speed of 50 mm/s. Normal rabbit rhythm is sinus and does not include respiratory sinus arrhythmia.44 Spontaneous changes in the QRS complex have been observed in normal rabbits in serial ECG recordings. Normal ECG values for a variety
CHAPTER 20 Cardiovascular, Lymphoproliferative Disorders, and Thymoma
A
259
B Fig. 20-2 Severe generalized cardiomegaly with congestive heart failure in a 5-year-old male castrated angora rabbit. A, In the right lateral projection, the cardiac silhouette is generally enlarged. The carina is elevated owing to left atrial and ventricular enlargement. A marked bronchointerstitial pattern is present in the caudal dorsal lung lobes compatible with pulmonary edema. The lung lobes are mildly retracted and the ventral border of the cardiac silhouette is obscured, indicating mild pleural effusion. B, In the ventrodorsal projection, a markedly enlarged left auricle is present (black arrowheads). An enlarged right atrium is also present (white arrowhead). An increased bronchointerstitial pattern is present in the caudal lung lobes, worse on the right than the left. Retraction of the lung lobe indicates pleural effusion (black arrows).
Table 20-1 Electrocardiographic Values in Clinically Normal Pet Rabbits ECG Parameter Heart rate Measurements (lead II) P wave Amplitude (height) Duration (width) P-R interval Duration QRS complex R-wave amplitude Duration Q-T interval Duration T wave Amplitude Electrical axis (frontal plane) Body weight
Values34
Values45
198-330 (mean 264) beats/minute
240 beats/minute (mean)
0.04-0.12 mV 0.01-0.05 s
0.04-0.07 mV 0.02-0.04 s
0.04-0.08 s
0.05-0.07 s
0.03-0.039 mV 0.02-0.06 s
0.12-0.2 mV 0.03-0.04 s
0.08-0.16 s
—
0.05-0.17 mV 43-80 degrees 1.1-7.9 (mean 2.57) kg
— — —
of pet rabbit breeds have been reported.34,45 In the authors’ experience, R-wave amplitudes in lead II of 0.1 to 0.2 mV seem most common. Reference ranges for ECGs are summarized in Table 20-1.
ECHOCARDIOGRAPHY Echocardiography provides a sensitive, accurate, noninvasive means of assessing the heart. Most rabbits tolerate echocardiography easily, making it a practical diagnostic tool. However,
in rabbits that are tachypneic or dyspneic, give oxygen by face mask during restraint. The echocardiographic exam may be performed in lateral or sternal recumbency. Standard views used to evaluate dogs and cats can also be obtained with the rabbit. Because of the rabbit’s rapid heart rate and small size, optimal evaluation requires a high-frequency transducer and highframe-rate ultrasound machine. Two-dimensional and M-mode echocardiography assess cardiac structure, chamber size, wall thickness, and motion as well as extracardiac structures, masses, and pleural effusion. Color-flow, spectral, and tissue Doppler
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Table 20-2 Echocardiographic Values in Clinically Normal Rabbitsa
Parameter
Dutch Belted Rabbits36 (n = 6)
Japanese White Rabbits50 (n = 4)
New Zealand White Rabbits54 (n = 20)
New Zealand White Rabbits13 (n = 26)
Body weight (kg) Age (months) LVEDD (cm) LVESD (cm) %FS IVSD (cm) LVPWD (cm) LA (cm) Ao (cm) LA/Ao RADs (cm) RA/Ao EPSS (cm) MVEFS (mm/s) RVOT velocity (m/s) LVOT velocity (m/s) LVET (s) VCF (circumference/s) MV E (m/s) MV A (m/s) MV E/A MV DT (m/s) TDI E LW (m/s) TDI A LW (m/s)
2.32 ± 0.36 7 (mean) 1.17 ± 0.19 0.70 ± 0.09 39.50 ± 5.39 0.25 ± 0.05 0.31 ± 0.08 — 0.67 ± 0.10 1.38 ± 0.32 0.61 ± 0.08 0.88 ± 0.17 0.05 ± 0.05 70.17 ± 31.82 0.83 ± 0.10 0.65 ± 0.14 0.08 ± 0.01 4.74 ± 0.45 — — — — — —
3.0 (mean) >13 1.69 ± 0.05 1.15 ± 0.05 — 0.33 ± 0.03 0.33 ± 0.03 1.05 ± 0.25 1.07 ± 0.12 — — — — — — — — — 0.44 ± 0.12 0.46 ± 0.17 1.0 ± 0.2 41.3 ± 2.5 — —
2.92 (mean) 4 1.54 ± 0.112 1.009 ±0.091 34.5 ± 4.9 0.217 ± 0.056 0.274 ± 0.041 — — — — — — — — 0.749 ± 0.195 0.126 ± 0.014 — 0.715 ± 0.138 0.514 ± 0.145 1.44 ± 0.28 — 0.067 ± 0.019 0.039 ± 0.007
2.3 ± 0.4 4-5 1.351 ± 0.105 0.864 ± 0.082 36.01 ± 4.31 0.265 ± 0.031 0.225 ± 0.029 0.749 ± 0.114 0.657 ± 0.046 1.15 ± 0.19 — — 0.141 ± 0.025 — 0.78 ± 0.12 0.86 ± 0.12 0.096 ± 0.010 — 0.78 ± 0.15 0.55 ± 0.11 1.44 ± 0.16 — 0.16 ± 0.05 0.09 ± 0.03
aValues
are mean ± SD. Ao, aorta; EPSS, E point to septal separation; IVSD, intraventricular septal thickness at end-diastole; IVSS, intraventricular septal thickness at end-systole; LA, left atrium; LA/Ao, left atrium-to-aorta ratio; LVEDD, left ventricular end-diastolic dimension; LVESD, LV end-systolic dimension; LVET, left ventricular ejection time; LVOT, left ventricular outflow tract; LVPWD, LV posterior wall thickness at end-diastole; MV A, mitral valve A wave; MV E/A, mitral valve E-to-A ratio; MV DT, mitral valve deceleration time; MV E, mitral valve E wave; MVEFS, mitral valve E-F slope; %FS, percent LV fractional shortening; RA/Ao, right atrium-to-aorta ratio; RADs, right atrial dimension in systole; RVOT, right ventricular outflow tract; TDI A LW, tissue Doppler imaging A’ wave from the left ventricular free wall; TDI E LW, tissue Doppler imaging E’ wave from the left ventricular free wall; VCF, velocity of circumferential fiber shortening.
echocardiography assess direction and velocity of blood flow, further defining cardiac conditions with more insight on systolic and diastolic function. Normal echocardiographic values have been published for several breeds of rabbits (Table 20-2). Sedative drugs used for restraint may affect cardiac measurements, especially alpha2-agonists such as dexmedetomidine and xylazine. Reduced percent fractional shortening, E/A reversal, decreased heart rate, and other changes have been documented with intramuscular ketamine at dose of 50 mg/kg and xylazine at 4 mg/kg.54 Sedation with ketamine 20 mg/kg and midazolam 2 mg/kg given subcutaneously has been shown to be less cardiodepressive.13
DISEASES AND MANAGEMENT CONGESTIVE HEART FAILURE In rabbits, congestive heart failure (CHF) is the clinical condition in which pulmonary edema, pleural effusion, or hepatomegaly develops as a result of structural or functional cardiac disease. The goal of therapy is to relieve congestion, control future retention of sodium and fluids, and improve cardiac performance. To this end, numerous management strategies are used during the acute stage. Place the patient in a quiet cage
with supplemental oxygen. Administer parenteral furosemide (1-4 mg/kg IV or IM q4-12h) and nitroglycerin 2% ointment (1⁄8 inch applied transdermally q6-12h). In a rabbit with pleural effusion, perform therapeutic pleurocentesis if the rabbit is dyspneic. Long-term therapy of CHF should include a diuretic (furosemide 1-2 mg/kg PO q8-24h) combined with treatment directed at the underlying precipitating cause. Knowledge of the cardiac disease process is the basis of specific treatment of the underlying condition. No cardiac drugs are approved for use in rabbits by the U.S. Food and Drug Administration. Drug dosages for rabbits are not available for all cardiac medications, but drugs and dosages published for cats or ferrets may be successfully used on a milligram-per-kilogram basis. Angiotensin-converting enzyme inhibitors such as enalapril maleate (0.25-0.5 mg/kg PO q24-48h, begun q24h) may be beneficial in treating rabbits with congestive heart failure. Pimobendan (0.1-0.3 mg/kg PO q12-24h) may also be used for treatment of systolic dysfunction and myocardial failure.45 During acute and chronic management, it is critical that clinical and radiographic signs, hydration status, appetite, and body weight as well as serum or plasma blood urea nitrogen, creatinine, and electrolyte concentrations be monitored. For drugs and dosages, refer to Chapter 41.
CHAPTER 20 Cardiovascular, Lymphoproliferative Disorders, and Thymoma
RV
LV
RA
261
RV LV
LA
B
A
RV
RV Ao
MV
LA
C
D
LV
E
Fig. 20-3 Standard echocardiographic views in rabbits with cardiac disease. A, Right parasternal long-axis four-chamber view. Notice the left and right ventricular dilation in this rabbit with mitral and tricuspid insufficiency. B, Right parasternal short-axis left ventricle papillary muscles view. Notice the prominence of the left ventricular papillary muscles (arrows) in this rabbit with left ventricular hypertrophy. C, Parasternal short-axis M-mode echocardiogram in which the M-mode beam is directed across the right ventricle, aortic valve, and left atrium. Note the dilation of the left atrium. D, Parasternal short-axis M-mode view of the left ventricle in which the M-mode beam is directed across the right ventricle (top) and left ventricle (bottom). Notice the dilated left ventricle and the decreased excursion of the septum compared with the posterior wall in this rabbit with cardiomyopathy and ventricular tachycardia. E, Parasternal short-axis M-mode echocardiogram of the left ventricle and mitral valve. Note the irregular rhythm. LV, Left ventricle; LA, left atrium; RV, right ventricle; RA, right atrium; Ao, aorta; MV, mitral valve.
CONGENITAL HEART DISEASE
MYOCARDIAL DISEASE
Congenital heart disease in rabbits is rarely reported. Ventricular septal defect, diagnosed with echocardiography, has been described.47 A ventricular septal defect, pulmonary hypertension, and valvular cyst identified at necropsy have been described in a New Zealand white rabbit.31
Numerous myocardial diseases have been reported in rabbits, and cardiomyopathy is a common postmortem finding in older rabbits.47 Idiopathic hypertrophic cardiomyopathy and dilated cardiomyopathy have been diagnosed by echocardiography (Fig. 20-3).44 Vitamin E deficiency produces a muscular dystrophy in which the myocardium may be affected.3 In experimental studies, myocardial disease has been created through inoculation of Trypanosoma cruzi49 and administration of doxorubicin.59 Infectious myocardial diseases are rare in pet rabbits. Known infectious organisms include Pasteurella multocida, Salmonella species, Encephalitozoon cuniculi,36 and coronavirus.8 The alpha-agonist drug detomidine has been associated with myocardial necrosis and fibrosis in New Zealand white rabbits.23 A similar ischemia-mediated process is suggested in association with ketamine/xylazine administration.36
ARRHYTHMIA Arrhythmias such as atrial fibrillation and ventricular premature complexes have been identified in pet rabbits with underlying cardiomyopathies and congestive heart failure.35 We have also diagnosed arrhythmias in several rabbits that exhibited syncope and an irregular heartbeat. In a rabbit with an arrhythmia, the treatment protocol should be based on ECG findings and clinical signs. For syncope associated with bradycardia, oral theophylline or pacemaker implantation may be indicated. Glycopyrrolate may be more effective than atropine sulfate in increasing heart rate.43 Atropine may not be as effective because some rabbits produce atropinesterases. Supraventricular tachycardias may be treated with oral digoxin or diltiazem. Ventricular tachyarrhythmias may respond to intravenous lidocaine. Dosages are published for lidocaine, verapamil, atropine, and glycopyrrolate. Other antiarrhythmic drugs may be used at dosages published for cats or ferrets.
VALVULAR DISEASE Mitral and tricuspid insufficiencies are not uncommon and have been identified in pet rabbits.44 Valvular insufficiencies may be associated with primary valve degeneration, cardiomyopathy, or infection. Progression of the condition leads to volume overload and potential congestive heart failure. A focal murmur is the most common clinical finding. Valvular disease is diagnosed with two-dimensional and Doppler echocardiography (see Fig. 20-3). Echocardiographic findings most often include
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thickening of one or both atrioventricular valves, dilation of cardiac chambers, and turbulent regurgitation of blood detected by Doppler. Valvular endocarditis, caused by Staphylococcus aureus, has been reported in rabbits.53
recessive gene, termed Is. In the WH strain, affected rabbits typically die at 5 to 13 months of age, usually with generalized lymphoma involving visceral organs and lymph nodes similar to the distribution of organ involvement seen in other domestic animals.14
VASCULAR DISEASE
INFECTIOUS FACTORS
Spontaneous arteriosclerosis of the aorta and other arteries has been observed in nearly all rabbit breeds. Clinical signs, if present, may include lethargy, anorexia, and weight loss. The cause is unknown. In rabbits with spontaneous arteriosclerosis, arterial walls and other soft tissues mineralize; the aortic arch and descending thoracic aorta are most commonly affected. Radiopaque vessels, caused by calcification, may be visible on radiographs.32,51 Pulmonary hypertension associated with high altitude has been reported in a rabbit.20 Lesions included right ventricular hypertrophy and pulmonary artery proliferation from hypoxia. Based on basic research, rabbits with pulmonary hypertension may respond to phosphodiesterase type-5 inhibitors, particularly vardenafil.55
Generalized lymphoma in strains of rabbits has been speculated as compatible with vertical transmission of an oncogenic virus, as occurs with feline leukemia virus. Some have speculated that lymphoma in rabbits may be caused by an oncogenic C-type tumor virus, similar to viruses causing this disease in rodents. In an early study attempting to evaluate this theory, tissues samples from an adult New Zealand white rabbit with generalized lymphoma were examined. Results of electron microscopic examination, tissue culture, immunodiffusion studies with FeLV antiserum, and immunofluorescent tests were negative for a virus.57 However, virus-like particles were demonstrated by electron microscopy in the kidneys of a 7-month-old New Zealand white rabbit with generalized lymphoma.18 In this case, the significance of the virus-like particles was not determined. To date, the hypothesis that a retrovirus (oncogenic virus) may be involved in the pathogenesis of lymphoid disease in rabbits has not been confirmed.
PART II LYMPHOPROLIFERATIVE DISORDERS AND THYMOMAS Lymphoproliferative disorders are uncommonly seen in pet rabbits, but clinicians treating rabbits should be able to properly recognize and diagnose these diseases when presented and offer treatment options when appropriate. Before the 1960s, rare cases of lymphoproliferative disease in domestic rabbits had been reported in the European literature, and lymphoma (malignant lymphoma or lymphosarcoma) was first reported in a domestic rabbit in the United States in 1968. In that report, a New Zealand white rabbit from a research colony had generalized lymphoid neoplasia involving the lymph nodes, liver, spleen, lungs, and gastrointestinal tract.58 Since then, lymphoproliferative diseases have been reported in both laboratory and pet rabbits, whereas most of these cases were diagnosed as generalized or multicentric lymphoma; however, cutaneous lymphoma, and lymphoid leukemia, thymoma, and thymic lymphoma have also been described.
ETIOLOGY The cause of lymphoid neoplasia is largely unknown and likely multifactorial in rabbits. Owing to their use in biomedical research, much is known about disease pathogenesis of induced lymphoid neoplasia in rabbits; however, few studies have investigated the cause of naturally occurring lymphoid disorders.
GENETIC FACTORS In one of the first reports, lymphoma was described in a series of rabbits from a breeding farm,33 leading to speculation that a breed or strain susceptibility exists or an infectious agent is associated with disease.14,33,58 A strain susceptibility has been shown in Wirehair (WH) rabbits associated with a single autosomal
TYPES OF LYMPHOPROLIFERATIVE DISORDERS MULTICENTRIC LYMPHOMA Multicentric lymphoma is the most common type of lymphoproliferative disease in rabbits and has been reported in several rabbit breeds, including New Zealand white, Japanese white, Dutch, and Netherland dwarf rabbits.5,19,52,56,57 Clinically, lymphoma has been observed in a variety of rabbit breeds, including satin, mini lop, and tan breeds. Lymphoma can occur in rabbits of all ages, from animals less than 1 year of age to geriatric animals.* In pet rabbits with lymphoma seen at the Animal Medical Center in New York City, ages have ranged from 2 to 9 years, with most averaging 4 to 5 years. Lymphoma of both T- and B-cell origin has been documented in rabbits. In a domestic rabbit with lymphoma and lymphocytic leukemia, the neoplasia was of T-cell origin.56 In a pet Dutch dwarf rabbit, T- and B-cell infiltrates were observed in the skin, lung, kidneys, liver, intestine, and lymph nodes. In this rabbit, the diagnosis was multicentric, T-cell–rich, B-cell lymphoma with cutaneous involvement.16 A 22-month-old rabbit that presented with acute unilateral exophthalmos was diagnosed with retrobulbar lymphoma.61 In this rabbit, histopathologic examination identified the mass as a B-cell lymphoma of the Harder’s gland, and the mesenteric lymph nodes, cecum, and both kidneys were also affected. In a rabbit that presented with pelvic paralysis, the diagnosis was spinal lymphoma of the sixth lumbar vertebra and concurrent pulmonary filariasis.48 In this rabbit, the neoplasm was immunophenotyped as B-cell lymphoma and the filariasis was considered an incidental finding. In another case, the authors described a rare lymphoma that developed in the cecum of a 6-year-old pet rabbit.24 Rabbits with multicentric lymphoma may exhibit nonspecific general signs, such as anorexia, lethargy, emaciation, pallor, *References 5, 14-16, 18, 19, 38, 52, 56-58, 61, 65.
CHAPTER 20 Cardiovascular, Lymphoproliferative Disorders, and Thymoma diarrhea, and rhinitis, depending on organ involvement and location. In a 2-year-old rabbit seen at The Animal Medical Center in New York City, the presenting clinical sign was severe upper respiratory stridor. At necropsy, lymphoma was present in the nasal turbinates and sinus, stomach, liver, spleen, kidneys, lymph nodes, and bone marrow. Another rabbit with multicentric lymphoma presented with an acute onset of hind-limb paresis and gastrointestinal stasis. At necropsy, neoplastic cells compatible with large-cell lymphoma (immunoblastic) were found in the spleen, kidneys, lungs, cecum, intestines, lymph nodes, and adrenal glands; no lesions were found in the spinal cord. Laboratory findings often depend on the organs involved. Results of plasma biochemical analysis may be unremarkable or reveal increases in concentrations of aspartate aminotransferase, creatine phosphokinase, blood urea nitrogen, and creatinine. Rabbits may be moderately to severely anemic14,65; in young rabbits, fluctuating and depressed hematocrit values were considered the best diagnostic tool for early identification of lymphoma.14 In rabbits with multicentric lymphoma seen at The Animal Medical Center in New York City, most had hematocrit levels in the low normal range (30%-33%; reference interval, 30%-50%).2 The white blood cell (WBC) count is often within reference intervals; however, some rabbits with lymphoma have shown a leukemic phase (see below). In young rabbits with multicentric lymphoma, high WBC counts were less frequent than a relative predominance of lymphoid cells, including immature and atypical cells, representing 80% to 90% of total WBCs. In an 18-month-old rabbit seen at the Animal Medical Center, the WBC count was 10,000/mL, with 63% lymphocytes. Lymphoma was present in the bone marrow of this rabbit. At necropsy, neoplastic lesions are commonly found in the lymph nodes, gastrointestinal tract, kidneys, liver, spleen, adrenal glands, gonads, and bone marrow.* Less common sites of neoplastic infiltrates are the auditory meatus, vertebrate, eye, and heart.5,48
CUTANEOUS LYMPHOMA Cutaneous lymphoma is usually primary but can be secondary to multicentric involvement. Two forms have been distinguished histologically and immunohistochemically: the epitheliotrophic form (mycosis fungoides) is composed of T lymphocytes, whereas the nonepitheliotropic form is composed of B lymphocytes (sparing the epidermis affecting the middle and deep portions of the dermis) (see Chapter 18, Fig. 18-5). Cutaneous lymphomas have the potential to metastasize to visceral organs, or cases may be diagnosed as visceral lymphomas with cutaneous involvement. Several cases of cutaneous lymphoma have been reported in rabbits.21,62,65 In an 18-month-old Netherland dwarf rabbit with multiple subcutaneous swellings over the shoulders, cutaneous lymphoma was diagnosed by histologic examination of biopsy samples.21 At necropsy, no gross or histologic evidence of lymphoma was found in any other organ system, and no viral particles were seen on electron microscopic examination of neoplastic tissue. In another report, three domestic rabbits were diagnosed with cutaneous lymphoma.65 Two rabbits were young (7 months and 1 year) and the third was 9 years old. One young rabbit had erythematous alopecia and hemorrhagic crusts of the chin and ventral neck. At necropsy, neoplastic lymphocytes were found in the skin, lymph nodes, and lungs. In skin sections, the lymphocytes infiltrated the entire dermis and into *References 5, 18, 24, 57, 58, 65.
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the epidermis. The second rabbit had bilateral blepharitis that was unresponsive to treatment. At necropsy, superficial and deep lymph nodes were markedly enlarged, and lungs had reddened areas. Lymphocytic infiltrates were found in the skin, lungs, liver, kidneys, and heart. In the skin sections, lymphoid infiltrates were primarily in the subcutis and deep dermis. The third rabbit had nonpruritic alopecia of the left lateral thorax. Cutaneous lymphoma was diagnosed by biopsy of a skin sample; in this rabbit, lymphocytes infiltrated the superficial dermis and epidermis. The rabbit lived for an additional year after diagnosis, with no response to treatment with interferon alpha-2b (see “Treatment,” below). A necropsy was not performed. In all three rabbits, immunologic staining of tissue sections confirmed the lymphoma to be of T-cell origin. In a large retrospective study of cutaneous neoplasms in pet rabbits over 16 years, the authors reported a single case of lymphoma in the flank of a 4-year-old female rabbit. In this case, epitheliotropism was not noted.62 In three clinical cases seen by one of the present authors (KQ), rabbits presented because of one or more subcutaneous masses. In a 7-year-old rabbit, cutaneous lymphoblastic lymphoma was diagnosed by excisional biopsy of a subcutaneous nodule on its dorsal neck. Three more masses developed within 1 month and were excised, but the rabbit died 2 months after the first biopsy. At necropsy, lymphoma was found in the cervical, mesenteric, and thoracic nodes. In another rabbit, a large, hemorrhagic mass was present on the ventral thorax. This rabbit was euthanatized. On histologic examination, lymphoma involving the skin mass and spleen was diagnosed. A 9-year-old rabbit presented with multifocal subcutaneous masses on the dorsum and ventral abdomen. Fine-needle aspirates of the masses revealed large-round-cell neoplasia. At necropsy, lymphoma was widely disseminated to the skin and subcutis, all abdominal organs, lungs, diaphragm, heart, adipose tissues, mandible, and incisor pulp cavity with marked intratumoral necrosis. Because of the extensive involvement of multiple sites, the primary source was unclear.
LEUKEMIA Lymphoid leukemia is the proliferation of neoplastic lymphocytes that typically originate in the bone marrow and occasionally in the spleen. The neoplastic cells may or may not be found circulating in peripheral blood. Three cases of lymphoblastic leukemia and one case of myeloid leukemia have been documented in rabbits.5,10,37,56 In rabbits with lymphoid leukemia, WBCs have ranged from 30,000 to more than 100,000/μL. In all rabbits with lymphoblastic leukemia, neoplastic cells were present in bone marrow, lymph nodes, and other organs typical of stage V lymphoma.
THYMIC MASSES: THYMOMA/THYMIC LYMPHOMA/THYMIC CARCINOMA Rabbits normally have a persistent large thymus that lies cranioventral to the heart and extends into the thoracic inlet (Fig. 20-4).29 In adult rabbits, the thymus can become hyperplastic and enlarge up to three or four times normal size, grossly resembling a tumor but with no neoplastic characteristics on histologic examination.64 A thymoma is a tumor derived from the epithelial components of the thymus and is composed of a mix of lymphoid and reticuloepithelial cells. Thymic lymphoma denotes that the neoplasm is of T-lymphocytic origin, possibly with other organ and systemic involvement. In some cases the lymphoid cells are small mature cells; in others, lymphocytes
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Fig. 20-4 Anatomy of the thymus of a normal adult rabbit. A, left lateral view; B, right lateral view. The thymus remains large in adult rabbits and is located cranioventral to the heart; it extends into the thoracic inlet. 1, Left thoracic lobe, thymus; 2, right ventral thoracic lobe, thymus; 3, right dorsal thoracic lobe, thymus; 4, left ventricle, heart; 5, right ventricle, heart; 6, left auricle, heart; 7, right auricle, heart; 8, thoracic aorta; 9, left subclavian artery; 10, left internal thoracic artery; 11, thymic branches; 12, thymic branch of the left subclavian artery; 13, costocervical trunk; 14, left pulmonary artery, pulmonary vein; 15, caudal vena cava; 16, right cranial vena cava; 17, right internal thoracic vein; 18, thymic veins; 19, right azygos vein; 20, right subclavian vein; 21, right external jugular vein; 22, right axillary vein. (Adapted from Popesko P, Rajtova V, Horak J, eds. Colour atlas of the anatomy of small laboratory animals. Vol I: Rabbit and guinea pig. St. Louis: Elsevier, 1990/1992.)
are pleomorphic with prominent nucleoli. Thymic lymphomas are unique because they reflect the function of the thymus gland as an organ involved in two-cell generation and differentiation. Diagnosis is made from cytologic examination of fine-needle aspirates or histopathologic examination of biopsy specimens. Clinically and cytologically, distinguishing thymoma from thymic lymphoma can be difficult. Tissue samples are often needed, since these tumors are classified based on the cells undergoing neoplastic transformation (epithelial and/or lymphocytic). Because of the difficulty in distinguishing hyperplasia, thymoma, thymic lymphoma, and carcinoma on the basis of gross appearance or imaging results, histologic examination is important. Thymoma, thymic lymphoma, and thymic carcinoma have been reported in rabbits.* Thymic lymphoma and thymic carcinoma are rare. Although thymomas are seen more frequently clinically, the incidence of thymoma appears to be low. In an early study of 1,100 female rabbits more than 2 years of age submitted for necropsy over a 17-year period, 234 rabbits had tumors. Of those 234 rabbits, 55 had multiple primary tumors, and 4 of these had thymomas.17 In a study of thymomas in 19 rabbits, incidence in males (9) and females (10) was similar, and mean age at presentation was 6.7 years.1
*References 1, 4, 28, 46, 60, 63, 64.
The presence of the cranial mediastinal mass usually causes difficulty breathing (hyperpnea, open-mouth breathing), which is commonly the presenting complaint. Physical examination findings typically relate to respiratory symptoms with nasal flaring evident, secondary to increased respiratory rate and effort.28,46 Auscultation of the thoracic cavity may reveal decreased or muffled lung sounds over the anterior mediastinum.46 Although judged subjectively, decreased compressibility of the thoracic cavity in smaller breeds of rabbits may be detected as well. On thoracic radiographs, a mediastinal mass is identified (Fig. 20-5, A) and pleural effusion may be present. In a 6-year-old Netherland dwarf rabbit with pleural effusion, chylothorax was diagnosed and considered secondary to the presence of the mediastinal mass.46 Uniquely, rabbits with thymic masses may have exophthalmos or present with prolapse of the third eyelids.28,60,63 In rabbits with exophthalmos, the eyes can be retropulsed without evidence of pain, suggesting that the exophthalmos is not caused by the presence of a space-occupying retrobulbar mass. These signs are consistent with a diagnosis of cranial vena caval syndrome (precaval syndrome) caused by the space-occupying mass compressing the vessels of the anterior thorax and impeding vascular return to the heart.4 The signs of paraneoplastic syndrome have also been described in rabbits and include hypercalcemia and exfoliative dermatitis
CHAPTER 20 Cardiovascular, Lymphoproliferative Disorders, and Thymoma
A
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B Fig. 20-5 A, Right lateral thoracic radiographs of a lop rabbit with a thymic mass. Ultrasoundguided fine-needle aspirates were diagnostic of thymoma. At presentation, preoperative radiographs show loss of cardiac silhouette, with tracheal elevation on the lateral radiograph. B, Postoperative lateral radiograph with surgical staple present show a more defined cardiac silhouette and return of the trachea to its normal anatomic position.
(see Chapter 18).12,26,60 In one rabbit, the hypercalcemia (14.7 mg/dL) was described as a paraneoplastic syndrome similar to that seen in dogs with thymoma.60 However, because of the unique calcium homeostasis in rabbits and the fact that the influence of diet and calcium metabolism on the plasma calcium concentration was not considered in this case, this conclusion is likely erroneous. Rabbits with thymoma may present with generalized scaling and sebaceous adenitis associated with thymoma. In one case, a paraneoplastic syndrome of thymoma-associated exfoliative dermatitis similar to the syndrome in cats was described.12 In another rabbit that responded to treatment with cyclosporine and oral tryglycerides, an autoimmune reaction directed at the sebaceous glands and a defect in lipid metabolism was suspected.26 Metastasis of thymic carcinoma in rabbits has been reported.63 Rabbits with thymic masses may have a normal complete blood count (CBC), a high WBC count, or anemia. Heterophilia with monocytosis, eosinophilia, basophilia, and thrombocytosis was described in one rabbit.1 In two clinical cases of thymoma, the WBC counts were 42,000 and 18,000/μL, with more than 70% lymphocytes. On microscopic interpretation of a blood smear from the first rabbit, the cells were characterized as small lymphocytes with cleaved nuclei and scant cytoplasm. A bone marrow aspirate showed no marrow infiltration. Both rabbits were eventually euthanized and no bone marrow involvement was found on postmortem examination in either one. On histopathologic examination, the masses in both rabbits were identified as thymoma.
DIAGNOSIS The diagnostic workup of a rabbit with a suspected lymphoproliferative disease is similar to that of other small animals. If the CBC results reflect either a high WBC count with mature lymphocytosis or a normal WBC count with an inverse lymphocyte/heterophil ratio, repeat the test to confirm the findings. If anemia is found on a routine CBC, consider ruling out possible lymphoma and perform additional tests. Submit a blood sample for a plasma biochemical analysis to look for evidence of multiorgan involvement. Obtain both thoracic and abdominal radiographs and an abdominal ultrasound to
identify abdominal masses or enlarged lymph nodes. Obtain fine-needle aspirates or do a biopsy of any enlarged peripheral lymph nodes or subcutaneous masses and submit a bone marrow sample for evaluation if appropriate. Submit full-thickness skin biopsy samples to a dermatohistopathologist if cutaneous lymphoma is suspected. If a thoracic mass is present, perform ultrasound of the thorax to determine the architecture of the mass. Do an ultrasound-guided fine-needle aspirate of the mass if possible and submit the sample for cytologic examination. Because thymomas are frequently cystic and do not exfoliate well, aspiration results may be nondiagnostic. Cytologic evaluation of cases with thymomas usually yields mostly mature lymphocytes as opposed to lymphoblasts, which would support a diagnosis of lymphoma. Thoracic needle core biopsy can be a diagnostic option but may yield necrotic and/or cystic material that might prohibit a definitive diagnosis. Computed tomography with contrast is indicated to determine the location and extent of a thoracic mass, particularly if surgery or radiation therapy are treatment options (see Chapter 35).
TREATMENT In general, little information is available on treating rabbits with lymphoproliferative diseases. Much information is anecdotal, with protocols based on those used in other small animals. At best, the long-term prognosis with treatment is guarded to poor. Consultation with an oncologist is indicated when clients are interested in pursuing chemotherapy or radiation treatments. Advise clients of the risks, benefits, and potential side effects of treatment options.
CHEMOTHERAPY Although much information has been published about chemotherapeutic agents in experimental studies in rabbits, few reports describe the use of chemotherapy in clinical cases of lymphoproliferative disease. As described above, a 9.5-year-old rabbit with cutaneous lymphoma was treated with recombinant human interferon alpha-2b at 1.5 million U/m2 administered
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subcutaneously three times weekly.65 After 2 months, no response was seen, and isotretinoin (4 mg/kg q24h on food) was added to the treatment for 2.5 weeks. No change was seen in the lesions and all treatments were discontinued; the rabbit died suddenly 1 year after diagnosis. Anecdotal information is available on the use of chemotherapeutic agents in pet rabbits. The CVP/COP (cyclophosphamide, vincristine, prednisolone) protocol, with or without doxorubicin and l-asparaginase, has also been recommended.42 Because of the lack of information available on the benefits of chemotherapy in rabbits, the potential risks must be considered before treatment. In one study, rabbits that were intraperitoneally infected with spores of E. cuniculi were treated with cyclophosphamide (50 mg/kg first dose, then 15 mg/kg weekly during the 12-week experimental period).22 In these rabbits, clinical signs of encephalitozoonosis developed between weeks 4 and 6, and all died during week 6. No signs of infection were seen in control rabbits. These results indicate that immunosuppression induced by cyclophosphamide gave rise to lethal encephalitozoonosis. Therefore consider serologic testing to determine if a rabbit is seropositive for E. cuniculi before beginning treatment with immunosuppressive drugs (see below). Other side effects of chemotherapy can include severe anemia, enteritis, typhlitis, and nephrotoxicity. In an experimental study of rabbits given daunorubicin or doxorubicin at 3 mg/ kg weekly for 10 weeks, cardiotoxicity was documented with daunorubicin but not with doxorubicin.27 Both drugs produced hemotoxicosis manifesting as aplastic anemia. Rabbits treated with doxorubicin exhibited more weight loss and had higher mortality rates than those treated with daunorubicin. A single dose of l-asparaginase at 10,000 IU/kg when given intravenously can induce a hyperinsulinemic, insulin-resistant, diabetic syndrome in rabbits.30 However, in small animals, l-asparaginase is currently given intramuscularly or subcutaneously; therefore route of administration may be a factor in toxicity. The neurotoxic effects of vincristine have been well studied in rabbits.11,40,41 Rabbits with thymomas have been treated with immunosuppressive therapy.1,39 Prednisone (0.5-2.0 mg/kg q12h) has been used successfully as adjuvant therapy in rabbits with thymoma undergoing radiation therapy.39 Prednisone is used both for its antineoplastic and anti-inflammatory effects, potentially helping to ameliorate the side effects of radiating thoracic structures, in particular radiation pneumonitis.39 However, in one clinical case of a rabbit treated with radiation therapy and prednisone (0.5 mg/kg q12h for 28 days), although the tumor regressed, the rabbit was euthanized 3 months after diagnosis because of lethargy and severe pleural edema. On histologic examination, the mediastinum was markedly fibrotic, with sterile granulomatous inflammation and thrombi found in mediastinal vessels. Chronic active pyelonephritis was also found, with numerous E. cuniculi organisms in renal tubular cells and lumens. In another clinical case of thymoma, the rabbit was treated with a low dose of prednisone (0.5 mg/kg q24h), with no other therapy. After 1 month, the tumor had not changed in size. The rabbit was euthanized 5 months later because of labored breathing and poor clinical condition. In one rabbit treated with radiation therapy and cyclophosphamide, severe bilateral renal fibrosis was found at necropsy, with death attributed to renal failure. The renal changes were attributed to repeated cyclophosphamide administration.1
TREATMENT OPTIONS FOR THYMOMAS Radiation Radiation therapy (RT) appears to be a good treatment option in rabbits with thymomas. In a recent study of 19 rabbits with thymomas that were treated with radiation, median overall survival was 313 days; when 3 rabbits that died acutely during the first 14 days of treatment were excluded, median survival was 727 days.1 Total radiation dose ranged from 28.8 to 48 Gy in rabbits receiving definitive fractionated protocols (≤4 Gy delivered three or more times weekly) and 24 to 32 Gy in rabbits receiving coarsely fractionated protocols (defined as any other RT protocol). Complications associated with radiation were uncommon and included radiation-induced myocardial failure, radiation pneumonitis, and alopecia. Tumors may be considerably smaller after only a few treatments (see Chapter 35, Fig. 35-9). Rabbits have been used in research studies regarding effects of irradiation on soft tissues.7,9 Risk factors to consider with radiation therapy include the number of anesthetic episodes and radiation-associated side effects, including late-term side effects. Cost is also a factor, with definitive radiation therapy being the most expensive because of the number of treatments necessary.
Surgical Excision Surgery is the treatment of choice for thymoma in all species when there is a solitary mass, as it provides the best chance of a cure by removal of the entire tumor (Fig. 20-5, B). Surgical excision of thymomas is done successfully in rabbits, but the risk of surgical or anesthetic-related complications and death is high. In three rabbits treated with surgery alone, postoperative survival ranged from 8 months to 3 years.1 Two rabbits treated with radiation therapy were also treated with surgical cytoreduction after completion of radiation.1 In a series of rabbits with mediastinal masses treated surgically by median sternotomy and mass excision, 7 of 14 rabbits survived 6 months or longer. One rabbit died during surgery, 6 died within 10 days after surgery, 1 survived 6 to 12 months, 4 survived 12 to 24 months, and 2 survived more than 24 months (F. Harcourt-Brown, personal communication, 2010). In a clinical report, a right fourth intercostal thoracotomy was performed in one rabbit with thymoma and the mass was excised. A chest drain was placed, but pneumothorax persisted after surgery and the rabbit was euthanized.60 In another rabbit, the mass was removed by median sternotomy.4 A chest drain was kept in place for 24 hours, after which it was removed and the rabbit recovered uneventfully. When the rabbit was euthanized 9 months later because of recurrent appendicular neurofibrosarcoma, no evidence of thymoma recurrence was present. In a clinical case seen by one of the present authors (AP), the mass was excised successfully but the patient went into cardiac arrest and died approximately 28 hours after surgery. Tumor-related cardiac dysfunction, anesthesia-related perfusion abnormalities, stress, and analgesia are important considerations in these cases. When a rabbit is positioned in dorsal recumbency for surgery, the mass may impinge on normal lung tissue and decrease ventilation and perfusion. Therefore intubation and a short anesthesia time are extremely important.
Therapeutic Aspiration of Cystic Thymomas Thymomas characteristically are cystic. In rabbits with thymomas that have large cystic components, removal of cystic fluid may help in relieving clinical signs of dyspnea and improving quality of life. In four clinical cases seen by one of the authors
CHAPTER 20 Cardiovascular, Lymphoproliferative Disorders, and Thymoma (KQ), periodic aspiration of the fluid by ultrasound-guided aspirate was done at varying intervals over 8 to 12 months. In three rabbits, the owners declined surgery or radiation therapy. Serial ultrasound-guided aspiration of cystic fluid was the only treatment done, and all rabbits improved clinically immediately after each aspiration. The procedure was repeated at varying intervals, usually at 3 to 6 months, as clinical signs of dyspnea or tachypnea recurred. In one rabbit treated with radiation therapy, the rabbit continued to develop large amounts of cystic fluid after treatment. Cystic fluid was aspirated periodically over a 12-month period to provide symptomatic relief. The rabbit subsequently died during surgery to excise the remaining thymoma.
References 1. Andres K, Kent M, Seidlecki C, et al. The use of megavoltage radiation therapy in the treatment of thymomas in rabbits: 19 cases. Vet Comp Oncol. 2011. In press. 2. Carpenter JW, Mashima TY, Rupiper DJ. Exotic animal formulary. 2nd ed. Philadelphia: WB Saunders; 2001. 3. Cheeke PR. Nutrition and nutritional diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:321-333. 4. Clippinger TL, Bennett RA, Alleman AR, et al. Removal of a thymoma via median sternotomy in a rabbit with recurrent appendicular neurofibrosarcoma. J Am Vet Med Assoc. 1998;213:1131,1140-1143. 5. Cloyd GG, Johnson GR. Lymphosarcoma with lymphoblastic leukemia in a New Zealand white rabbit. Lab Anim Sci. 1978;28:66-69. 6. Cruise LJ, Brewer NR. Anatomy. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:47-51. 7. Danielsson M, Engfeldt B, Larsson B, et al. Effects of therapeutic proton doses on healthy organs in the neck, chest, and upper abdomen of the rabbit. Acta Radiol Ther Phys Biol. 1971;10:215-224. 8. DiGiacoma RF, Maré CJ. Viral diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:171-204. 9. Engfeldt B, Larsson B, Naeslund C, et al. Effect of single dose or fractionated proton irradiation on pulmonary tissue and Vx2 carcinoma in lung of rabbit. Acta Radiol Ther Phys Biol. 1971;10:298-310. 10. Finnie JW, Bostock DE, Walden NB. Lymphoblastic leukaemia in a rabbit: a case report. Lab Anim. 1980;14:49-51. 11. Fiori MG, Schiavinato A, Lini E, et al. Peripheral neuropathy induced by intravenous administration of vincristine sulfate in the rabbit. An ultrastructural study. Toxicol Pathol. 1995;23:248-255. 12. Florizoone K. Thymoma-associated exfoliative dermatitis in a rabbit. Vet Dermatol. 2005;16:281-284. 13. Fontes-Sousa AP, Moura C, Carneiro CS, et al. Echocardiographic evaluation including tissue Doppler imaging in New Zealand white rabbits sedated with ketamine and midazolam. Vet J. 2009;181:326-331. 14. Fox RR, Meier H, Crary DD, et al. Lymphosarcoma in the rabbit: genetics and pathology. J Nat Cancer Inst. 1970;45:719-729. 15. Fox RR, Norberg RF, Meier H. Clinical hematological progression of hereditary lymphosarcoma in rabbits. J Hered. 1976;67:376-380. 16. Gómez L, Gázquez A, Roncero V, et al. Lymphoma in a rabbit: histopathological and immunohistochemical findings. J Small Anim Pract. 2002;43:224-226. 17. Greene HSN, Strauss JS. Multiple primary tumors in the rabbit. Cancer. 1949;2:673-691.
267
18. Gupta BN. Lymphosarcoma in a rabbit. Am J Vet Res. 1976; 37:841-843. 19. Hayden DW. Generalized lymphosarcoma in a juvenile rabbit: a case report. Cornell Vet. 1970;60:73-82. 20. Heath D, Williams D, Rios-Dalenz J, et al. Pulmonary vascular disease in a rabbit at high altitude. Int J Biometeorol. 1990;34:20-23. 21. Hinton H, Regan M. Cutaneous lymphoma in a rabbit. Vet Rec. 1978;103:140-141. 22. Horváth M, Leng L, Stefkovic M, et al. Lethal encephalitozoonosis in cyclophosphamide-treated rabbits. Acta Vet Hung. 1999;47:85-93. 23. Hurley RJ, Marini RP, Avison DL, et al. Evaluation of detomidine anesthetic combinations in the rabbit. Lab Anim Sci. 1994;44:472-478. 24. Ishikawa M, Maeda H, Kondo H, et al. A case of lymphoma developing in the rabbit cecum. J Vet Med Sci. 2007;69:1183-1185. 25. James TN. Anatomy of the cardiac conduction system in the rabbit. Circ Res. 1967;20:638-648. 26. Jassles-van der Lee A, van Zeeland Y, Kik M, et al. Successful treatment of sebaceous adenitis in a rabbit with ciclsproin and triglycerides. Vet Dermatol. 2009;20:67-71. 27. Klimtová I, Simunek T, Mazurová Y, et al. Comparative study of chronic toxic effects of daunorubicin and doxorubicin in rabbits. Hum Exp Toxicol. 2002;21:649-657. 28. Kostolich M, Panciera RJ. Thymoma in a domestic rabbit. Cornell Vet. 1992;82:125-129. 29. Kozma C, Macklin W, Cummins LM, et al. Anatomy. In: Weisbroth SH, Flatt RE, Krause SE, eds. The biology of the laboratory rabbit. New York: Academic Press; 1974:50-72. 30. Lavine RL, Dicintio DM. l-Asparaginase-induced diabetes mellitus in rabbits. Diabetes. 1980;29:528-531. 31. Li X, Murphy JC, Lipman NS. Eisenmenger’s syndrome in a New Zealand white rabbit. Lab Anim Sci. 1995;45:618-620. 32. Lindsey JR, Fox RR. Inherited diseases and variations. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:293-319. 33. Loliger H. Ueber das vorkommen von leukosen beim kaninchen. Berlin u Munchen Tierarztl Wchnschr. 1966;79:192. 34. Lord B, Boswood A, Petrie A. Electrocardiography of the normal domestic pet rabbit. Vet Rec. 2010;167:961-965. 35. Lord B, Devine C, Smith S. Congestive heart failure in two pet rabbits. J Small Anim Prac. 2011;52:46-50. 36. Marini RP, Li X, Harpster NK, et al. Cardiovascular pathology possibly associated with ketamine/xylazine anesthesia in Dutch belted rabbits. Lab Anim Sci. 1999;49:153-160. 37. Meier H, Fox RR, Crary DD. Myeloid leukemia in the rabbit (Oryctolagus cuniculus). Cancer Res. 1972;32:1785-1787. 38. Meier H, Fox RR. Hereditary lymphosarcoma in WH rabbits and hereditary hemolytic anemia associated with thymoma in strain X rabbits. Bibl Haematol. 1973;39:72-92. 39. Morrisey JK, McEntee M. Therapeutic options for thymoma in the rabbit. Sem Avian Exot Pet Med. 2005;14:175-181. 40. Muzylak M, Maslinska D. Neurotoxic effect of vincristine on ultrastructure of hypothalamus in rabbits. Folia Histochem Cytobiol. 1992;30:113-117. 41. Ogawa T, Mimura Y, Kato H, et al. The usefulness of rabbits as an animal model for the neuropathological assessment of neurotoxicity following the administration of vincristine. Neurotoxicology. 2000;21:501-511. 42. Ogilvie G, Bennett A, Bergman P. Cutaneous lymphoma in rabbits. Retrieved December 30, 2009, from http://www.vin.com/ Members/SearchDB/Boards/B0047500/B0046942.htm. 43. Olson ME, Vizzutti D, Morck DW, et al. The parasympathetic effects of atropine sulfate and glycopyrrolate in rats and rabbits. Can J Vet Res. 1994;58:254-258.
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44. Orcutt CJ. Cardiac and respiratory disease in rabbits. Proceedings. Autumn Meet Brit Vet Zoo Soc. 2000:68-73. 45. Pariaut R. Cardiovascular physiology and diseases of the rabbit. Vet Clin North Am Exot Anim Pract. 2009;12:135-144. 46. Pilny AA, Reavill DR. Chylothorax and thymic lymphoma in a pet rabbit (Oryctolagus cuniculus). J Exot Pet Med. 2008; 17:295-299. 47. Redrobe S. Imaging techniques in small mammals. Sem Avian Exot Pet Med. 2001;10:195. 48. Reed SD, Shaw S, Evans DE. Spinal lymphoma and pulmonary filariasis in a pet domestic rabbit (Oryctolagus cuniculus domesticus). J Vet Diag Invest. 2009;21:253-256. 49. Rossi MA. Microvascular changes as a cause of chronic cardiomyopathy in Chagas’ disease. Am Heart J. 1990;120:233-236. 50. Saku K, Fujino M, Yamamoto K, et al. Cardiac function of WHHL rabbit, an animal model of familial hypercholesterolemia. Artery. 1990;17:271-280. 51. Shell LG, Saunders G. Arteriosclerosis in a rabbit. J Am Vet Med Assoc. 1989;194:679-680. 52. Shibuya K, Tajima M, Kanai K, et al. Spontaneous lymphoma in a Japanese white rabbit. J Vet Med Sci. 1999;61:1327-1329. 53. Snyder SB, Fox JG, Campbell LH, et al. Disseminated staphylococcal disease in laboratory rabbits (Oryctolagus cuniculus). Lab Anim Sci. 1976;26:86-88. 54. Stypmann J, Engelen MA, Breithardt AK, et al. Doppler echocardiography and tissue Doppler imaging in the healthy rabbit: differences of cardiac function during awake and anaesthetised examination. Int J Cardiol. 2007;115:164-170. 55. Toque HA, Teixeira CE, Priviero FB, et al. Varendafil, but not sildenafil or tadalafil, has calcium-channel blocking activity in rabbit isolated pulmonary artery and human washed platelets. Br J Pharmacol. 2008;154:787-796.
56. Toth LA, Olson GA, Wilson E, et al. Lymphocytic leukemia and lymphosarcoma in a rabbit. J Am Vet Med Assoc. 1990; 197:627-629. 57. Ubertini TR. Brief communication: etiological study of a lymphosarcoma in a domestic rabbit. J Natl Cancer Inst. 1972; 48:1507-1511. 58. Van Kampen KR. Lymphosarcoma in the rabbit. A case report and general review. Cornell Vet. 1968;58:121-128. 59. Van Vleet JF, Ferrans VJ. Clinical and pathologic features of chronic adriamycin toxicosis in rabbits. Am J Vet Res. 1980;41:1462-1469. 60. Vernau KM, Grahn BH, Clarke-Scott HA, et al. Thymoma in a geriatric rabbit with hypercalcemia and periodic exophthalmos. J Am Vet Med Assoc. 1995;206:820-822. 61. Volopich S, Gruber A, Hassan J, et al. Malignant B-cell lymphoma of the Harder’s gland in a rabbit. Vet Ophthalmol. 2005;8:259-263. 62. von Bomhard W, Goldschmidt MH, Shofer FS, et al. Cutaneous neoplasms in pet rabbits: a retrospective study. Vet Pathol. 2007;44:579-588. 63. Wagner F, Beinecke A, Fehr M, et al. Recurrent bilateral exophthalmos associated with metastatic thymic carcinoma in a pet rabbit. J Small Anim Pract. 2005;46:393-397. 64. Weisbroth SH. Neoplastic diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. New York: Academic Press; 1994:259-292. 65. White SD, Campbell T, Logan A, et al. Lymphoma with cutaneous involvement in three domestic rabbits (Oryctolagus cuniculus). Vet Dermatol. 2000;11:61-67.
CHAPTER
21
Soft Tissue Surgery
Jeffrey R. Jenkins, DVM, Diplomate ABVP (Avian)
Evaluation of the Rabbit as a Surgical Patient Equipment Presurgical Treatment Postsurgical Monitoring Blood Loss Pain and Analgesics Surgical Techniques Adhesion Formation Choice of Suture Material Skin Closures Common Procedures Ovariohysterectomy Orchidectomy (Castration) Gastrointestinal Surgery Gastrotomy Intestinal Resection, Anastomosis, and Enterotomy Surgery of the Large Bowel Anorectal Papilloma Removal Urinary Tract Surgery Pyelolithotomy Nephrotomy Nephrectomy Abscesses Dermoplasty and Tail Amputation to Correct Problems of Urine Scald Surgery of the Ear Canal and Auditory Bulla Lateral Ear Canal Ablation Osteotomy of the Ventral Bulla Thoracotomy
There are substantial physiologic and anatomic differences between rabbits and species that are more familiar to veterinarians. Rabbit behavior—including reaction to stress and pain and poor acceptance of sutures, dressings, and coaptation devices— is unlike that of other pets. The surgeon should have a thorough Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
knowledge of rabbit physiology and behavior prior to performing surgery in these patients.
EVALUATION OF THE RABBIT AS A SURGICAL PATIENT Before surgery, take a thorough history, including signalment, diet, and appetite, and perform a complete physical examination, including state of hydration and presence of infection; address preexisting problems to improve the patient’s prognosis for a successful surgery. Make an attempt, through discussion with the owner, observation of the animal, and evaluation of clinical data, to assess the level of stress that the animal is experiencing. Sometimes it is advantageous to postpone surgery until the animal has adapted to its new situation.
EQUIPMENT Little specialized equipment is required for surgery in rabbits. Appropriately sized Cole-style endotracheal tubes and a short, narrow laryngoscope or an open-style otoscope and a rigid plastic catheter are helpful for intubating the rabbit (see Chapter 31 for other intubation techniques). Equipment to monitor heart rate should be capable of measuring rates of 250 to 300 beats per minute. A heated surgery table or circulating water blanket, a source of radiant heat at locations where the patient is anesthetized, and a device to accurately monitor body temperature are necessary during long procedures on small patients. Small (human infant) Balfour retractors, flexible hook retractors (Lone Star Retractor, Lone Star Medical Products, Stafford, TX), or chess piece retractors (CHESS Surgical System, Canica, Ontario, Canada) enhance abdominal exposure, and malleable retractors serve well to hold viscera out of the way without removing them from the abdomen. An assortment of sterilized stainless steel spoons of various sizes is helpful in removing the stomach contents during gastric exploratory surgery; vascular clamps, hair clips (hair-control clips), or bobby pins substitute for Doyen forceps, and mechanical suction is helpful for removing pus from dental abscesses and for surgery of the gastrointestinal tract. Skin staples work well for skin closure and cannot be removed by most rabbits. 269
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PRESURGICAL TREATMENT The author provides food (grass hay) and water to rabbits up until the time of surgery, although some authors have suggested that a large volume of food in the rabbit’s stomach can cause variations in anesthetic doses.12 Begin antibiotic therapy in rabbits with systemic or localized bacterial infections, such as those with upper respiratory tract infections caused by Pasteurella species (“snuffles”) or with dental abscesses or infected wounds. Prophylactic antibiotics may be given if there is a significant chance of bacterial contamination during surgery. Quinolones, azithromycin, chloramphenicol, trimethoprim-sulfa combinations, and sulfa drugs generally do not negatively affect the normal cecal-colic microflora of rabbits. Use caution with beta-lactams, other macrolides, and other antibiotics that target gram-positive or anaerobic bacteria. Parenteral fluids are not necessary in routine procedures. When supportive fluid therapy or vascular access is needed, a 20to 26-gauge indwelling catheter is placed in the cephalic or lateral saphenous vein. Peripheral catheters work well during surgery and in the immediate recovery period. However, some rabbits do not tolerate a catheter when awake and must have the catheter and intravenous line covered with split-loom tubing or heavily bandaged to protect them. Alternatively, an intraosseous catheter may be placed within the greater trochanter. Moving the intravenous line away from the face results in better tolerance in some rabbits. Rabbits are considered steroid-sensitive species,4 and steroids are administered only if indicated by the underlying disease process; they usually are not given for routine or elective surgery. Give atropine when indicated (0.1-0.2 mg/kg SC, IM; see Chapter 31) or glycopyrrolate (0.01-0.02 mg/kg SC) to control bradycardia, salivation, or respiratory secretions. These problems are rare when isoflurane anesthesia is used. Some rabbits produce atropine esterase; in such cases the dose of atropine may have to be repeated if signs recur. The combination of thin skin and dense, fine fur makes it easy to cut a rabbit when clipping hair or shaving. Keep the skin spread flat in front of the blade and clip with the flat surface of the blade held parallel and close to the skin to minimize nicks and cuts. A fine No. 40 blade and an unhurried approach can help to prevent the fine hair from accumulating between the clipper blades, causing them to jam or cut poorly.
anthropomorphic evaluation of the injury or surgery the animal has experienced. See Chapters 31 and 41 for suggested dosages of analgesics.
SURGICAL TECHNIQUES ADHESION FORMATION Problems resulting from adhesions are regularly found in rabbits; these are most often reflected in postoperative problems involving the cecum, colon and bladder and result from adhesions to the uterine stump, broad ligament, or ovarian pedicle. Adhesions associated with gastrotomy, enterotomy, and a variety of other types of gastrointestinal surgery may also occur. Symptoms include recurrent cystitis, cystic calculi, and microurinary calculi.14 Prevention of adhesions has been the subject of much study, often involving rabbits as animal models of human disease.6,7 A large number of substances have been used to combat adhesion formation, but the majority of these have proven to either be too toxic for use, to possess a high a complication rate, to be too difficult in application, or simply not to work. Promising substances include hyaluronic acid solutions10 and sodium carboxymethyl cellulose.5 Chondroitin sulfate20 has also been reported to be effective. These agents act by mechanical separation of surfaces as well as adding a protective “finish” to the viscera. Both mechanisms being a feature of their inert nature. Thrombolytics, such as streptokinase,13 have been reported to be effective in adhesion prevention, but they present difficulties with local and systemic administration and the risk of hypersensitivity. There have been several studies using recombinant tissue plasminogen activator (rt-PA) in rabbits.3,19 In a rabbit adhesion model, rt-PA was found to reduce the primary and recurrent adhesion rates by 80%.3 Use of rt-PA was shown to be safe in the presence of colonic anastomoses and did not alter abdominal wound strength or increase postoperative hemorrhage. Other studies have evaluated the use of endoscopy to correct adhesions as compared with traditional laparotomy.23 These have shown that laparoscopic adhesiolysis is associated with a significantly reduced formation of new postoperative adhesions as compared with laparotomy.
CHOICE OF SUTURE MATERIAL
The blood volume of the rabbit is reported to be approximately 57 mL/kg body weight.16,18 Most mammalian species experience a drop in arterial pressure and cardiac output with moderate blood loss. Loss of 15% to 20% of the total blood volume causes massive cholinergic release, with tachycardia and intense arterial constriction; thus blood is redistributed away from the gut and skin. In a 4-kg rabbit, this amounts to 34 to 45 mL of blood. An acute blood loss of 20% to 30% of total blood volume, or 45 to 68 mL in a 4-kg rabbit, is critical.
The biologic and physical characteristics of suture material influence wound healing. New suture material made from polymers that are removed by hydrolytic degradation are much less reactive and cause fewer and weaker adhesions. For example, monofilament polyglyconate (Maxon, Davis & Geck, Manati, PR) or other similar monofilament synthetic suture for closure of gastrotomy, enterotomy, or colotomy sites, cesarean sections, or other major abdominal surgery are excellent choices. Stainless steel or tantalum clips (Hemoclip, Weck, Research Triangle Park, NC) are excellent for vessel and small pedicle ligation with minimal tissue reaction. A study of laser anastomosis for sutureless closure of the colon of rabbits has been published,15 and this technique holds potential for the future.
PAIN AND ANALGESICS
SKIN CLOSURES
The very nature of rabbits is the foremost argument for the use of analgesics. A rabbit in pain is inactive, anorectic, and poorly responsive, and it may grind its teeth. Pain is assumed based on
Rabbits commonly remove skin sutures. All but the smallest rabbits will chew sutures of nylon or even stainless steel monofilament. Intradermal closures work very well but are
POSTSURGICAL MONITORING BLOOD LOSS
CHAPTER 21 Soft Tissue Surgery time-consuming to place because of the tenuity of rabbit skin. Cyanoacrylate tissue cement (Vet-Bond, 3M Medical-Surgical Division, St. Paul, MN) works well to hold rabbit skin but is often removed by the rabbit. Skin staples are both reliable and well accepted by rabbits and their owners.
COMMON PROCEDURES OVARIOHYSTERECTOMY The reproductive tract of the female rabbit is unusual compared with that of the dog or cat (Fig. 21-1). The uterus is bicornate. Each uterine cornua possesses a cervix (there is no uterine body). At maturity it is coiled in the caudal abdomen, cranial and just dorsal to the urinary bladder. Long uterine (tuba uterina) and infundibular tubes (infundibulum tubae) extend between the cornua and the ovary.21 The uterus is easily exteriorized but is more fragile than that of other species. In a healthy doe, the caudal portion of the broad uterine ligament (ligamentum latum uteri), the mesometrium, is a principal fat-storage site that makes identifying and ligating uterine vessels difficult. The urethra of the female rabbit empties into the proximal end of a deep vaginal vestibule. Expression of the bladder with the animal in dorsal recumbency often leads to retrofilling of the vaginal vault. This can be a source of contamination of the peritoneal cavity during uterine surgery. It is also important not to confuse the vaginal vault with the bladder. With the rabbit anesthetized before surgery, empty the rabbit’s bladder by gentle palpation. Shave and prepare the abdominal area and restrain the rabbit on the surgery table in dorsal recumbency, draped for surgery. Make a 2- to 3-cm midline incision centered over the cranial pole of the bladder, about half the
c ut/it o
u
c
ut/it
u bl
bl vv
Fig. 21-1 Uterus of a 4-month-old nulliparous rabbit at surgery for ovariohysterectomy. The ovarian vessels have been clamped with mosquito hemostatic forceps, ligated, and incised and the uterus exteriorized. The top of the illustration is cranial. The rabbit’s ovary (o) is often coiled in the uterine/infundibulum tubules (ut/it). The bicornate uterus (cornua) (u) end in a paired cervix (cc). The broad ligament (bl) of this young, well-fed rabbit is not so filled with fat that the uterine vessels cannot be easily identified. The arrowheads mark the point where the uterine vessels are transfixed and ligated. The broad ligament is trimmed from the vagina to the level of the cervix, as marked by the orange lines. Finally the cranial end of the vagina is double ligated and transfixed just caudal to the cervix at the level of the black lines.
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distance between the umbilicus and the cranial rim of the pubis. Lift the narrow linea alba from the abdominal contents as you make a stab incision into the abdomen; be very careful in entering the abdominal wall, because the thin-walled cecum and bladder are often pressed firmly against the ventral abdomen. In the reproductively mature rabbit, the uterus can typically be seen as it lies cranial and dorsal to the cranial pole of the bladder and may be lifted through the incision with forceps. A spay (snook) hook is not necessary and may cause damage to the cecum. Follow the uterus to the uterine/infundibular tubes. The uterine/infundibular tubes are coiled in a large loop. They are several times longer than those of a dog or cat; be careful not to leave any portion of them. Multiple vessels are associated with the ovary, but they are smaller than those of many mammals. Carefully identify each and double ligate them with transfixing sutures of chromic gut or synthetic absorbable suture. Hemorrhage is seldom a problem with these vessels. The uterine vessels stand as much as a centimeter off the uterus and may be of significant size in mature does. Double ligate these vessels with transfixing ligatures to the vaginal serosa. The mesometrium may then be stripped away from the cervix and vagina. Ligate the uterus just caudal to the cervices. Avoid contaminating the abdomen with urine or vaginal contents if a caudal ligature is used. Ligate each uterine horn if removed cranial to the cervix, or carefully ligate at the dorsal vagina if the uterus is removed caudal to the cervices. Closure of the abdomen is routine. Close the skin with surgical staples, an intradermal suture pattern, or tissue cement.
ORCHIDECTOMY (CASTRATION) Sexually active male rabbits (bucks) have obnoxious sexual mounting and urine-marking behaviors that generally lead their owners to want to have them neutered. Furthermore, bucks may become territorial and possessive about their environment and owners, leading to aggressive behavior. The rabbit’s testes are similar to those of the cat but may move freely from the scrotum to the abdomen through an open inguinal canal. Soft tissue herniation and strangulation of bowel loops is prevented by a large mass of fat associated with the epididymis, which rests in the inguinal canal when the testicle is in the scrotum. For castration, anesthetize and restrain the buck in dorsal recumbency. Carefully shave the hair from the scrotum and surrounding area and surgically prepare and drape the area to minimize contamination. Make a 1- to 1.5-cm incision with a No. 15 scalpel blade through the skin and vaginal tunic on the ventral surface of both sides of the scrotum (Fig. 21-2). Remove the testis from the tunic and carefully tear the ligament of the testicle from the tunic with a dry gauze sponge. Pull the testis caudally to expose a section of the vas deferens and the vascular structures of the spermatic cord, then tie them in an overhand knot with a small Mayo needle holder or mosquito forceps. Alternatively, ligate the duct and vasculature with 2-0 to 3-0 synthetic absorbable suture. Cut the duct and vessels distal to the knot or ligature and return the spermatic cord to the inguinal canal in such a way that it can be recovered if bleeding occurs. Return the tunic to the scrotum and repeat the process for the remaining testis. Observe the rabbit for several hours after the surgery for hemorrhage. Because complications most often result from overactivity or sexual activity, castrated rabbits are often hospitalized
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T
VT
A
B G L
SC T T
VT
D
C
SC
T
E
F Fig. 21-2 A technique for castration of a rabbit. A, The scrotum and surrounding skin is prepared for sterile surgery. B, The skin and vaginal tunic (VT) on both sides of the scrotum are incised (arrowhead). C, The testes (T) are removed from the vaginal tunic and the ligament of the testicle (L) is torn from the tunic using a dry gauze sponge (G). D, The vas deferens and vessels of the spermatic cord (SC) are tied with an overhand knot using a needle holder. F, The tunic is returned to its correct position within the scrotum.
CHAPTER 21 Soft Tissue Surgery
R
S
S P
Fig. 21-3 Herniation of rectum (R) through the right orchidectomy site in an underweight rabbit 48 hours after surgery. The scrotum (S) and the penis (P) are seen below the exposed rectum. Herniation can be prevented by postponing surgery on exceptionally thin rabbits or closing the tunic or inguinal canal after removal of the testis. Because of the fragile nature of the rabbit rectum and colon, returning the herniated tissue should not be attempted without enlarging the inguinal canal.
overnight. Herniation is unusual but may be associated with the castration of exceptionally thin rabbits, where there is no fat to fill the inguinal canal (Fig. 21-3). Surgery should be postponed in these rabbits until such time as they obtain normal body weight. An alternative closed technique for castration may also be used (A. Bürgos, personal communication). Make a single prescrotal skin incision with caution to incise only the skin. A curved hemostat is gently tunneled through the subcutaneous tissues to the caudal aspect of the testicle. The testicle, within the tunic, is gently pulled through the skin incision, breaking the attachments between the subcutis and the tunic. Transfix and double ligate the tunic and spermatic cord using encircling ligatures. Incise and remove the testicle distal to the point of ligation. Repeat the procedure for the opposite testicle using the same skin incision. Closure can be made with tissue glue or skin suture. Advantages of this procedure are avoidance of opening of the vaginal tunic, thus decreasing the risk of herniation. Disadvantages include increased surgery time, increased tissue manipulation, and the use of ligature.
GASTROINTESTINAL SURGERY GASTROTOMY Because of the success of medical management of gastric stasis, gastrotomy is rarely necessary. See Chapter 15 for treatment of gastric stasis. Surgery is indicated for rabbits with suspected or known gastric foreign bodies or complete gastric or pyloric obstruction or for those that fail to show clinical signs of improvement after 3 to 5 days of intensive medical therapy for signs related to gastric stasis syndrome. Rabbits with delayed gastric emptying should receive a thorough workup, and every effort should be made to return the rabbit to a positive energy balance and to correct fluid and electrolyte imbalances before surgery. A common accompanying lesion with prolonged
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gastric obstruction is hepatic lipidosis, presumably caused by starvation. Correct any negative energy balance, acidosis, and ketosis to reduce complications in these rabbits.9 Prior to surgery, address pain with appropriate analgesic medications and place an intravenous catheter for fluid administration and vascular access during surgery and the postoperative period. Anesthetize the rabbit and restrain it on a circulating water blanket in dorsal recumbency. Shave the rabbit from the inguinal to the midthoracic area and prepare it for aseptic surgery. Make a midline incision that is long enough for exploration of the entire gastrointestinal tract, taking care not to damage the stomach or cecum, which may be pressed tightly against the abdominal wall. Drape lap pads moistened with normal saline along the incision line and use a Balfour or other retractor to fully expose the abdomen. Explore the abdomen before the gastrotomy to determine whether additional lesions are present. Examine the liver carefully; if it is abnormally pale or yellow, do a liver biopsy and submit the sample for histopathologic examination. Place stay sutures in the greater curvature of the stomach and elevate it into the surgical field. Place additional moistened lap pads around the stomach to prevent contamination of the abdomen with gastric contents. Make an incision in the avascular area between the lesser and greater curvatures and carefully remove the stomach contents or foreign body with a sterile spoon. Rinse the lumen with a small volume of warm saline and examine it for abnormalities, then gently palpate the pylorus for patency. Close the stomach in a two-layer inverting pattern with 3-0 to 4-0 synthetic monofilament suture. Extend the sutures into but not through the gastric mucosa. Thoroughly lavage the abdomen with warm isotonic fluids and close the incision routinely. Postoperative management is equally critical to the successful outcome of the procedure.22,24 Supportive therapy to minimize pain and support wound healing, prevent further hepatic damage, and promote hepatic regeneration is essential. Furthermore, care must be taken to support the normal gut microflora to prevent the complications of stress-induced enteritis complex.
INTESTINAL RESECTION, ANASTOMOSIS, AND ENTEROTOMY Intestinal resection, or enterotomy, is most often indicated in rabbits with foreign-body ingestion or trauma to the intestine. Neoplasia and infiltrative intestinal diseases are uncommon and generally consist mostly of metastatic uterine adenocarcinomas. Successful intestinal surgery in rabbits demands attention to surgical principles. Special care to preserving the blood supply and luminal diameter must be taken to compensate for the small size of the lumen in relation to the thickness of the visceral wall in the rabbit intestine.1 The bowel must be handled gently to prevent shock and postoperative ileus. Preparation of the rabbit for surgery and the abdominal incision are the same as for gastrotomy. Care must be taken not to damage the cecum while making the incision. Examine the entire gastrointestinal tract before the enterotomy. If resection and anastomosis are necessary, ligate the mesenteric and arcade vessels and apply crushing and noncrushing clamps, as is done in other small animals. Vascular clamps, including bulldog clamps, or serrefines (noncrushing clamps) can be used on the small-diameter intestines of rabbits. For the last few years, I have also used gas-sterilized hair clips or bobby pins as noncrushing clamps in some cases (Fig. 21-4). Incise the intestine at an acute
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B
A
D
C
Fig. 21-4 Intestinal occluders used in rabbit surgery. A variety of small clamps may be used for intestinal occlusion in small patients. Pictured clockwise from the upper left are (A) Acland microvascular clamps; (B) small microvascular clips, which work well on small intestine; (C) bobby pins; and (D) hair clips, which work better for the cecum, colon, and descending colon. Not shown are bulldog vascular clamps.
angle to augment the small luminal diameter. I prefer an appositional suture technique with 5-0 to 6-0 synthetic monofilament suture. Occasionally linear foreign bodies may be accompanied by bowel plication or intussusception. Multiple enterotomy sites may be required to remove these foreign bodies. Longitudinal incisions with transverse closures are sometimes used if the luminal diameter is small (Fig. 21-5). If the abdomen is contaminated during the procedure, lavage with warm saline for several cycles of irrigation and suction before closure.
SURGERY OF THE LARGE BOWEL Surgery of the rabbit colon is performed primarily to correct obstructions at the level of the fusus coli. Other indications for surgery of the large bowel include injury resulting from bite wounds, intraluminal trauma resulting from diagnostic or clinical instrumentation, or trauma secondary to accidents during surgery. Many rabbits with these injuries are seen as referral cases, and the condition may have been present for several days before examination. With the exception of rectal polyps and metastatic uterine adenocarcinoma, neoplasia of the lower bowel is rare. Several areas of the colon and cecum are thin-walled and easily torn; their suturing characteristics are poor at best. Surgery at the fusus coli is challenging because of its location at the root of the mesentery. If possible, massage the material causing the obstruction (typically hard feces, fecoliths, or cecoliths) past the tight spot and into the descending colon. If bowel must be resected, incise it at an acute angle, as with the small intestine. In the descending colon, make longitudinal incisions and close them transversely to increase luminal diameter. Use an interrupted suture pattern in an appositional technique for anastomosis of the colon. An inverting suture technique may be used to close lesions of the cecum; use a 4-0 to 6-0 synthetic monofilament suture placed at intervals of 2 to 3 mm. A segment of omentum can be used to reinforce the anastomosis or incision line. Consider medical treatment to reduce the risk of adhesions (see “Adhesion Formation,” above). Before closure, lavage the abdomen with several cycles of warmed saline containing 10% povidone-iodine,
Fig. 21-5 A longitudinal incision (A) of the intestine with a transverse closure (B) may help to prevent stricture formation in the rabbit if the luminal diameter is small. followed by suction. Remove any grossly evident ingesta in the abdominal cavity. If the abdominal cavity becomes contaminated, take samples for both aerobic and anaerobic bacterial culture and treat the rabbit aggressively with antibiotics.
ANORECTAL PAPILLOMA REMOVAL Anorectal papillomas are cauliflower-like, fungating masses that arise from the anorectal junction. They are benign and are not related to the papillomas of the skin or oral cavity. Papillomas arise from the rectal squamous columnar junction and protrude from the anus once they have grown to a sufficient size. Removal of these lesions is usually successful. The tissue of the papilloma is friable and has a tendency to bleed. The mucosal attachment may be stalk-like or broadbased. Removal is facilitated by good exposure of the lesion. Position the rabbit in dorsal recumbency with the pelvis slightly elevated. Have an assistant place and hold a human nasal speculum (veterinary canine vaginal speculum) in the anus. Alternatively, place several stay sutures around the circumference of the anus so that an assistant can provide gentle traction. Remove the papilloma by sharp dissection or by laser or electro/radiosurgery, taking care to remove all of the mass to prevent recurrence. Suture the mucosa with a fine, 5-0 to 6-0 absorbable suture material in a simple continuous pattern. Large papillomas may be removed in sections to simplify closure. As an alternative, cryosurgery yields good results with these masses. Take care to protect and not freeze healthy tissues. Three freeze-thaw cycles with the ice ball extending 2 to 3 mm into the mucosa at the base of the papilloma stalk results in successful treatment.
URINARY TRACT SURGERY The kidneys are mobile, accessible, and convenient for biopsy. The urinary bladder of the rabbit is tough but thin-walled. When greatly distended, it may rupture easily.
CHAPTER 21 Soft Tissue Surgery Renal calculi are a common problem in rabbits and a common cause of renal failure. Pyelolithotomy, nephrotomy, or nephrectomy is indicated in these cases. Like a dog or cat, a rabbit with renal calculi must be thoroughly evaluated before surgical correction of the problem is attempted. If renal calculi have resulted in dilation of the proximal ureter and renal pelvis, they may be removed through an incision made at the junction of these structures (pyelolithotomy). This approach to removal of renal calculi does not require occlusion of the renal vasculature and avoids trauma to the renal parenchyma, thereby minimizing deleterious effects on postoperative renal function. In rabbits with substantial renal disease, nephrectomy or nephrotomy is indicated. Anesthetize the rabbit, shave the ventral abdomen from pubis to midthorax, and prepare the rabbit for aseptic surgery in dorsal recumbency. Make a midline incision that is long enough to explore the abdomen. Take care not to damage the stomach or the cecum, which may be pressed tightly against the abdomen. Drape lap pads moistened with normal saline solution along the incision and place a retractor for better visibility of the abdomen. Explore the abdomen to determine whether additional lesions are present. To facilitate exposure of the kidney, gently remove the cecum, colon, and intestines from the abdomen and wrap them in moistened gauze.
PYELOLITHOTOMY Dissect the kidney free of its peritoneal attachments and rotate it medially, exposing the renal pelvis and proximal ureter. Make an incision over the pelvis and proximal ureter and remove the calculi. Flush the renal pelvis and calyces with saline to remove any small calculi that may remain. Pass a 3.5- to 5-Fr catheter into the bladder through the ureter to ensure patency. Close the incision in the renal pelvis and ureter with 5-0 or 6-0 continuous absorbable suture, lavage the abdomen, and close normally.
NEPHROTOMY The author has performed sagittal nephrotomy in rabbits; however, complications, including renal failure, have occurred in some rabbits with this technique. Thoroughly evaluate renal function before performing this surgery. Sagittal nephrotomy is performed as in the dog or cat. Isolate the kidney as described previously. Use serrefines or vascular clamps of appropriate size for temporary vascular occlusion. Divide the kidney along its sagittal plane. Remove renal calculi and flush the renal pelvis and calyces with saline to remove any small calculi that may remain. Pass a 3.5- to 5-Fr catheter into the bladder through the ureter to ensure patency. Press the kidney together and hold it in place for several seconds to allow the two halves to “stick together.” Suture the capsule, using 4-0 to 6-0 suture and a simple interrupted pattern and placing sutures every 2 to 4 mm. Remove the vascular clamps and observe the kidney for bleeding. Control minor bleeding by covering the site with absorbable gelatin sponges (Gelfoam, Pharmacia & Upjohn, Kalamazoo, MI). Close the abdomen routinely.
NEPHRECTOMY Nephrectomy is indicated in rabbits with a diseased or damaged kidney if adequate function remains in the opposite kidney. Expose the kidney as described previously. Use vascular clips
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to ligate the renal artery, vein, and ureter. Close the abdomen routinely.
ABSCESSES In reaction to most bacterial infections, rabbits form thickwalled abscesses that contain caseous, purulent discharge. If these abscesses are merely opened and drained, they frequently recur. Therefore abscesses should be excised with a substantial margin, so that all contaminated tissue is removed. Ligate blood vessels with a small-diameter absorbable suture and close the skin with surgical staples. Excision of large abscesses may require the use of skin flaps to cover the defect. Complete excision is often not possible with foot or dental abscesses; aggressive surgical debridement must be used. After debridement, the wound is allowed to granulate in, but the prognosis is guarded to poor for a complete cure. The use of antibiotic-impregnated polymethylmethacrylate (AIPMMA) beads in these abscesses has proved to be highly successful when total excision is not an option. Pododermatitis and the resulting osteomyelitis can cripple a rabbit; these lesions are painful, and the rabbit may be reluctant or unable to stand or walk. In these rabbits, consider amputation as an alternative to more conventional treatment (see Chapter 33).
DERMOPLASTY AND TAIL AMPUTATION TO CORRECT PROBLEMS OF URINE SCALD Dermatologic problems that result from urine scald or chronic diarrhea are common in pet rabbits (see Chapter 18). Underlying causes of obesity, dietary indiscretion, urinary tract disease, and spinal cord disease must be addressed; however, correction of the dermatologic problem may depend on preventing urine contamination of the perineal tissues. These problems result from the repeated contamination by urine or diarrhea of the perineal skin and medial surfaces of the hind legs. In obese rabbits, a large fold of skin and fat may partially cover the genital area, interfering with the passage of urine. Alternatively, the rabbit may be unable to rotate its pelvis during urination or defecation to direct the stream of urine caudally, resulting in contamination of the skin of the perineum and legs. These problems are compounded because the urine scald causes increased immobility, which further contributes to the likelihood of urine contamination. Several techniques can be used to correct these problems: dermoplasty of the caudal abdomen to remove the fold of skin that interferes with the passage of urine and a combination of tail amputation and dermoplasty to lift the genital area dorsally, resulting in passage of urine in a more caudal direction. It must be emphasized that these surgeries are adjunct and salvage procedures; every effort should be made to correct the underlying cause of the problem before surgery is undertaken. However, excellent results have been obtained with these corrective procedures in rabbits. Use systemic as well as topical antibiotics and protectants before surgery to decrease inflammation and infection in the area. For the skin fold resection, position the rabbit in dorsal recumbency and shave and prepare the area from the midabdomen to the tail, including the medial thigh region to the stifle. Identify a crescent area of skin cranial to the genital area that, when removed, will eliminate any excessive tissue that protrudes over the genital area (Fig. 21-6). Incise along this
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Fig. 21-6 Procedure for ventral skin fold resection in a rabbit. A large fold of abdominal skin may interfere with urine passage in an obese rabbit, causing contamination of the surrounding skin and urine scald. A ventral skin fold resection of the caudal abdomen removes a crescent-shaped area of skin, allowing unimpeded passage of urine.
crescent of skin, avoiding damage to the lateral abdominal vein, which lies lateral to the nipple and deep to the glandular tissue of the mammary gland. There may be an advantage in removing some or all of the inguinal adipose body, which extends from just cranial to the inguinal mammary gland to the genital area and is a primary location for the deposition of adipose tissue. Tissue removal should result in taut skin (but without tension on the incision) over the caudal ventral abdomen when the rabbit is in dorsal recumbency. A two-layer closure is preferred, with an absorbable suture in the deep layer and staples in the skin. The second procedure (B. Loudis, personal communication) is more difficult to perform but very effective for rabbits that urinate on their legs because of an inability to lift the pelvis and direct the stream of urine elsewhere. Position the rabbit for surgery in ventral recumbency with the rear legs extended behind the rabbit. Shave an area extending 3 to 5 inches from the anus and tail and prepare it for surgery. Delineate a crescent-shaped section of skin that, when removed, will lift the anus and urethral opening to a dorsal position at the caudalmost extreme of the rabbit. The crescent should extend from a point dorsal to the anus, just beyond the dorsal limits of the external anal sphincter muscle on its ventral curvature to dorsal of the tail and lateral and ventral to the urethral opening (Fig. 21-7). Identify the coccygeal and lateral ventral sacrocaudal muscles of the coccygeal vertebrae and remove them from their insertion. In male rabbits, the retractor penis muscle must also be identified and, in some cases, carefully dissected from its origin. Amputate the tail at the third or fourth coccygeal vertebra and ligate the medial sacral artery. The coccygeal and retractor penis muscles (if removed) are reattached at or about the dorsocaudal origin of the semitendinosus muscle with polypropylene or strong absorbable suture. Place vertical mattress sutures to position the closure and relieve tension and close the incision in two layers, as described previously.
SURGERY OF THE EAR CANAL AND AUDITORY BULLA Otitis secondary to infections of the upper respiratory tract and as an extension of infections and abscesses of the upper cheek teeth is common in rabbits.8 From their origins in the tympanic bulla, these infections can but do not always spread to affect the vestibular apparatus or may track through the acoustic meatus and along the vestibular nerve to form abscesses within the cranium.11 In many cases, the tympanic membrane ruptures and a thick suppurative exudate accumulates in the horizontal auditory canal. Pressure against the auditory canal at the intersection of the horizontal and vertical canals will cause a herniation of the canal between the cartilage of the acoustic meatus and the scutiform and auricle cartilage, forcing a pus-filled pocket of the canal into the subcutaneous space. The hernia is palpable at the point where the horizontal canal turns to the vertical canal. Interestingly, the majority of rabbits with otitis media show no outward clinical signs of the infection and only the most astute owner is aware that the rabbit has a problem. Because of the difficulty of diagnosing otitis in rabbits and the high rate of recurrence, it has been recommended “not to embark on treatment unless clinical signs are present.”11 I have found that many of these infections will respond to a combination of debridement of the material in the auditory canal and topical and long-term systemic medical treatment using azithromycin. In cases where these infections cause clinical signs (erythema, pruritus, headshaking, malodor, etc.) and do not respond to more conservative measures, surgery may be indicated.
LATERAL EAR CANAL ABLATION Anesthetize the rabbit and restrain it with the lateral side of an ear or the face at a 45-degree angle to the surgery table. Shave the ear and head surrounding the ear and the ear canal.
CHAPTER 21 Soft Tissue Surgery
A
277
B
C Fig. 21-7 Procedure to lift the perineal area and prevent urine contamination of the legs and perineum. A, The rabbit is positioned in ventral recumbency and a crescent-shaped area of skin around the tail is incised. B, The coccygeal and lateral ventral sacrocaudal muscles of the coccygeal vertebrae are removed at their origin, and the tail is amputated at the third or fourth coccygeal vertebra. C, The incision is closed in a two-layer closure, resulting in lifting of the anus and urethral opening dorsally.
Flush the ear canal to remove infected material. Make an incision over the ear canal exposing both its vertical and horizontal portions. Bluntly dissect the skin away from the canal. No skin is removed in order to simplify closure. Dissect the canal from surrounding tissue, sparing the musculature (the nuchal and parotic portions of the superficial sphincter muscles of the neck, the dorsal and ventral superficial scutuloauricular muscles, the auricular portion of the zygomatic muscle, and the parotidoauricular muscle)21 and the parotid salivary gland and remove this as one long, intact, curved cylinder. Incise the canal from the inside of the pinna 2 to 3 cm deeper in the canal than the upper limit of the skin incision to allow skin to close the incision. Expose the lateral and ventral side of the external auditory meatus and lateral tympanic bulla and remove the meatus and the lateral side of the bulla using a small ronguer. Remove any infected material within the bulla by repeatedly flushing and suctioning until all is removed. Remove the epithelial lining of the bulla using a small curette; then flush and suction repeatedly. Any material left in the bulla may result in recurrence and chronic drainage from the area of the incision. AIPMMA or absorbable sponges (Gelfoam absorbable gelatin compressed sponge, Pfizer, New York, NY) soaked in antibiotic
may be placed within the bulla. Close using absorbable sutures and skin staples.
OSTEOTOMY OF THE VENTRAL BULLA Osteotomy of the ventral bulla is indicated for the management of otitis media without otitis externa that is not responsive to medical management; it is also used in the treatment of neoplasia of the bulla. Osteotomy allows removal of caseous debris, collection of diagnostic samples, and may provide a route for the administration of topical treatment. Ingrowth of tissue into the bulla’s lumen increases blood supply to the area and facilitates treatment of the infection. AIPMMA beads may be placed within the bulla to provide long-term local antibiotic therapy.2 The rabbit is anesthetized and restrained in dorsal recumbency. Shave and prepare the rabbit for sterile surgery from the affected ear to beyond the midline of the larynx. Palpate the right mandibular angle and make a skin incision medial and parallel to the mandibular angle to approximately the level of the wing of the atlas. Incise the superficial sphincter muscles of the neck along the same plane as the skin incision medial to the mandibular salivary gland and lymph node. Bluntly dissect the
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SECTION II Rabbits
digastric muscles away from the muscles (the medial pterygoid and the mandibular retractor muscles) on the medial side of the mandible to expose the linguofacial artery and vein and the hypoglossal nerve. Continue dissection lateral to the styloglossal muscle along the division marked by the lingual nerve to expose the bulla.21 Great care must be taken not to damage the important nerves and vessels in this dissection. The bulla is palpable as a spherical protrusion of the skull rostral to the jugular process. Place a Gelpi retractor to maintain exposure. Penetrate the tympanic bulla on the medial aspect with a small Steinmann pin and enlarge the opening laterally using a small pair of rongeurs. Take samples for culture and cytology. Use a small curette to remove caseous material and the lining of the bulla. The dorsomedial compartment of the bulla is avoided so the inner ear is not damaged.17 Flush and suction the bulla repeatedly. AIPMMA or absorbable sponges soaked in antibiotic may be placed within the bulla. Close subcutaneous tissues using absorbable sutures and skin staples. Postoperative considerations include evaluating for signs of Horner’s syndrome, hypoglossal nerve deficits, progressive head tilt, and torticollis. Analgesics and antibiotics are continued as indicated.
THORACOTOMY Rabbit thoracotomy is routinely performed in medical research; however, it is uncommon in private practice. Indications for thoracotomy in practice include perforating injuries, biopsy or treatment of thoracic masses, surgical drainage of abscesses, and lung lobectomy. Many of these procedures may be performed via thoracoscopy or transthoracically when aided by ultrasound.
References 1. Booth HW, Hartsfield SM. Use of the laboratory rabbit in the small animal student surgery laboratory. J Vet Med Educ. 1990;17:16-18. 2. Chow EP, Bennett RA, Dustin L. Ventral bulla osteotomy for treatment of otitis media in a rabbit. J Exot Pet Med. 2009;18:299-305. 3. Doody KJ, Dunn RC, Buttram Jr VC. Recombinant tissue plasminogen activator reduces adhesion formation in a rabbit uterine horn model. Fertil Steril. 1989;51:509-512. 4. Edita J, Lenka L, Zoran J, et al. Dexamethasone-induced immunosuppression: a rabbit model. Vet Immunol Immunopathol. 2008;122:231-240. 5. Elkins TE, Bury RJ, Ritter JL, et al. Adhesion prevention by solutions of sodium carboxymethyl cellulose in the rat. I. Fertil Steril. 1984;41:926-928. 6. Ellis H. Internal overhealing: the problem of intraperitoneal adhesions. World J Surg. 1980;4:303-306.
7. Ellis H. The causes and prevention of intestinal adhesions. Br J Surg. 1982;69:241-243. 8. Flatt RE, Deyoung DW, Hogle RM. Suppurative otitis media in the rabbit: prevalence, pathology, and microbiology. Lab Anim Sci. 1977;27:343-347. 9. Gillett NA, Brooks DL, Tillman PC. Medical and surgical management of gastric obstruction from a hairball in the rabbit. J Am Vet Med Assoc. 1983;183:1176-1178. 10. Goldberg EP, Sheets JW, Habal MB. Peritoneal adhesions: prevention with the use of hydrophilic polymer coatings. Arch Surg. 1980;115:776-780. 11. Harcourt-Brown F. Cardiorespiratory disease. Textbook of rabbit medicine. Oxford: Butterworth Heinemann; 2002:324-334. 12. Harkness JE, Wagner JE. The biology and medicine of rabbits and rodents. 4th ed. Media, PA: Williams & Wilkins; 1995. 13. James DCO, Ellis H, Hugh TB. The effect of streptokinase on experimental intraperitoneal adhesion formation. J Pathol Bacteriol. 1965;90:279-287. 14. Jenkins JR. Clinical pathology. In: Meredith A, Flecknell P, eds. BSAVA manual of rabbit medicine and surgery. 2nd ed. Gloucester: BSAVA; 2006:45-51. 15. Kawahara M, Kuramoto S, Ryan P. First experimental sutureless laser anastomosis of the large bowel: long-term study. Dis Colon Rectum. 1992;35:792-798. 16. Kozma C, Macklin W, Cummins LM, et al. Anatomy, physiology, and biochemistry of the rabbit. In: Weisbroth SH, Flatt RE, Kraus AL, eds. The biology of the laboratory rabbit. New York: Academic Press; 1974:50-72. 17. Mehler SJ. Common surgical procedures. In: Meredith A, Flecknell P, eds. BSAVA manual of rabbit medicine and surgery. 2nd ed. Gloucester: BSAVA; 2006:166-183. 18. McGuill MW, Rowan AN. Biological effects of blood loss: implications for sampling volumes and techniques. ILAR News. 1989;31:5-18. 19. Menzies D, Ellis H. Intra-abdominal adhesions and their prevention by topical tissue plasminogen activator. J R Soc Med. 1989;82:534-535. 20. Oelsner G, Graebe RA, Pan SB, et al. Chrondroitin sulphatea new intraperitoneal treatment for postoperative adhesion prevention in the rabbit. J Reprod Med. 1987;32:812-814. 21. Popesko P, Rajtová V, Horák J. A colour atlas of anatomy in small laboratory animals. Vol. 1. Rabbit, guinea pig. London: Wolfe Publishing; 1992. 22. Sebesteny A. Acute obstruction of the duodenum of a rabbit following the apparently successful treatment of a hairball. Lab Anim. 1977;11:135. 23. Tittel A, Treutner KH, Titkova S, et al. New adhesion formation after laparoscopic and conventional adhesiolysis: a comparative study in the rabbit. Surg Endosc. 2001;15:44-46. 24. Wagner JL, Hackel DB, Samsell AG. Spontaneous deaths in rabbits resulting from gastric trichobezoars. Lab Anim Sci. 1974;24:826-830.
SECTION THREE
Guinea Pigs and Chinchillas
CHAPTER
22
Biology, Husbandry, and Clinical Techniques of Guinea Pigs and Chinchillas
Katherine E. Quesenberry, DVM, MPH, Diplomate ABVP (Avian), Thomas M. Donnelly, BVSc, Diplomate ACLAM, and Christoph Mans, MedVet
Biology and Husbandry of Guinea Pigs Anatomy and Physiology Behavior Husbandry Biology and Husbandry of Chinchillas Anatomy, Physiology, and Behavior Husbandry Clinical Techniques for Guinea Pigs and Chinchillas Handling and Restraint Physical Examination Blood Collection Urethral Catheterization and Cystocentesis Clinical Laboratory Findings Treatment Techniques
Guinea pigs (Cavia porcellus) and chinchillas (Chinchilla laniger) are hystricomorph rodents from South America (Fig. 22-1). They share many anatomic and physiologic characteristics and the approach to their veterinary care is similar. Both species are monogastric herbivores with a large cecum, and both produce precocious young after a relatively long gestation period. There are also some important differences between the two species. For example, guinea pigs require a dietary source of vitamin C but chinchillas do not; dystocia is common in guinea pigs but not in chinchillas; and chinchillas require dust baths but guinea pigs do not. Both guinea pigs and chinchillas are small, gentle, Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
lively species that make good pets because they are docile and relatively easy to care for. This chapter summarizes the information on basic biology, husbandry, and clinical techniques that is relevant to the medical care of these species as pets. Readers who are interested in more detailed descriptions of anatomy and physiology of guinea pigs can find these in the references.6,46,52 Guinea pigs have been used as laboratory animals for more than 500 years, and abundant information is available about them; until recently, less has been published about chinchillas.
BIOLOGY AND HUSBANDRY OF GUINEA PIGS Guinea pigs, also known as cavies, were domesticated in South America between a.d. 500 and 1000 and possibly as early as 1000 b.c.66 At the time of the Spanish invasion of South America, guinea pigs, known as cuy, were raised by the Incas for food and for use in religious ceremonies.66 Guinea pigs were brought to Europe about 500 years ago. Although they never became popular as a food source outside of South America, in Europe and North America they have been raised as pets and laboratory animals ever since. Domestic guinea pigs remain a very important food source with a very high protein content (19%) for many people in the Altiplano region of South America.19 They are raised by families and often left uncaged to forage for food about the dwellings.61,66 A larger breed, weighing up to 2.5 lb (1.1 kg), was recently developed by researchers at La Molina National University to promote exportation as a food product to Peruvian immigrant populations in the United States, Japan, and Europe.14 Guinea pigs have been introduced into the Democratic Republic of the Congo as “microlivestock” to serve as a food source in that impoverished country.53 279
SECTION III Guinea Pigs and Chinchillas
280
Fig. 22-2 Hairless “skinny” guinea pig. Many different breeds, such as the hairless, and varieties of guinea pigs have become popular in recent years both in the United States and in Europe.
A
B Fig. 22-1 A, Normal guinea pig. B, Normal chinchilla. Several species of wild cavy are described (Cavia aperea, Cavia intermeida, Cavia fulgida, Cavia magna, and Cavia tschudii)71 and are found today in Colombia, Venezuela, Brazil, Argentina, Paraguay, and Peru.61,66 They inhabit a wide variety of habitats, including grasslands, the forest edge, swamps, and rocky areas. The American Rabbit Breeders Association originally recognized three standard breeds of domestic guinea pigs: the American (or English), the Abyssinian, and the Peruvian. The American guinea pig has short smooth hair, the Abyssinian has relatively short coarse hair that grows in whorls or rosettes, and the Peruvian has very long (up to 15 cm) silky hair. Currently, the American Cavy Breeders Association (www.acbaonline.com) recognizes at least 13 breeds, and the British Cavy Council (www.britishcavycouncil.org.uk) recognizes at least 22 breeds, with published breed standards. New breeds and varieties are constantly being added. Newer recognized breeds include the Texel, which has long hair in ringlets and curls all over its body; the Teddy, which has a short dense coat with a kinked hair shaft; and the Silkie (or Sheltie), which is similar to the Peruvian except that long hair does not cover its face. Additionally, several hairless breeds have been developed by cavy breeders, such as the Skinny, which is hairless except for the head and lower
legs (Fig. 22-2), and the Baldwin, which is born with hair but becomes totally bald at weaning. Each breed has different varieties of color and markings, including white, red, tan, brown, chocolate, and black; coats can be monochromatic, bicolored, or tricolored. A solid color is described as self; for example, black self is monochromatic black. The Himalayan color variety has a white body with black or chocolate nose, ears, and feet. Roan describes a coat of mixed black and white hairs. Agouti guinea pigs have an undercolor that is ticked throughout with another color, similar to wild guinea pigs. Dalmatian guinea pigs have a white body with black spotting. Albino guinea pigs are common in a laboratory setting. Guinea pigs are lively, responsive, gentle pets, particularly if they are handled frequently while young. They have some peculiarities, however, of which all persons working with them should be aware. First, their response to perceived danger is freeze or flight. If frightened, they tend to become immobile or, alternatively, make an explosive attempt to escape. Second, guinea pigs do not tolerate dietary or environmental changes well. Their food preferences are established early in life, and they often refuse to eat if their food is changed in type or presentation. Moreover, they may become depressed or go off feed when hospitalized; therefore the attitude and food consumption of all hospitalized animals should be monitored carefully. Third, guinea pigs require a dietary source of vitamin C. Under conditions of good husbandry, guinea pigs are hardy animals with few disease problems. Conversely, inadequacies in diet or husbandry can lead to illnesses that, if not managed early, are difficult to reverse. Sick guinea pigs do not tolerate clinical procedures well and have been known to go into cardiac and respiratory arrest secondary to the stress of restraint or a diagnostic procedure that would be routine in species such as ferrets. Handle very sick guinea pigs carefully. Concentrate on providing good supportive care and maintaining caloric intake in a low-stress environment while working toward a diagnosis and therapy specific to that diagnosis.
ANATOMY AND PHYSIOLOGY General Characteristics Guinea pigs have stocky bodies, delicate short limbs, rounded hairless pinnae, and no tails. Males are larger than females, weighing 900 to 1,200 g compared with 700 to 900 g for
CHAPTER 22 Biology, Husbandry, and Clinical Techniques
281
Fig. 22-3 Normal genitalia of an adult male guinea pig. The
Fig. 22-4 A female guinea pig with dystocia prepared for surgery.
scrotal sac is large and the penis is partially extruded. Males have a small os penis. Both male and female guinea pigs have one pair of inguinal nipples. The two nipples of the mammary glands are visible.
The Y-shaped urethral-vaginal orifice is visible above the anus. The two mammary glands are visible. A cephalic catheter is placed in the right front leg.
females. Obesity is relatively common in pet animals. The life span of pet guinea pigs is typically 5 to 6 years. Wild cavies feed at dawn and dusk; they live in small groups (5 to 10 individuals) in burrows or crevices.61 The hair coat of guinea pigs is composed of large guard hairs surrounded by an undercoat of fine hairs. Androgen-dependent sebaceous glands are abundant along the dorsum and around the anus. The sebaceous glands around the anal area are important for marking, and the guinea pig is frequently seen rubbing or pressing its rump against a surface.6 In older males, excessive accumulation of sebaceous secretions occurs in the skin around the base of the spine, resulting in thick, matted, greasy fur. Breeders may call this a “grease gland.” Both male and female guinea pigs have one pair of inguinal nipples (Figs. 22-3 and 22-4). Guinea pigs have large tympanic bullae. They have 32 to 36 vertebrae; the vertebral formula is C7, T13(14), L6, S2(3), Cd4(6). There are 13 to 14 pairs of ribs, of which the last one or two are cartilaginous. The small cylindrical clavicle attaches laterally to the coracoid process of the scapula and medially to the manubrium. The pelvic symphysis generally remains fibrocartilaginous,6 and a gap in the symphysis can be palpated at the time of impending parturition. Guinea pigs have four digits on the front feet and three on the rear feet; each has a short claw that may require periodic clipping in some individuals. The spleen is relatively broad in guinea pigs. The thymus in immature animals is located within the ventral cervical area and the cranial mediastinum; in adults, a thymic remnant may be present in the cranial mediastinum.6 Guinea pigs, like monkeys, ferrets, and humans, are considered to be corticosteroid-resistant species because steroid administration is not associated with marked changes in thymic physiology or peripheral lymphocyte counts. Guinea pigs have no laryngeal ventricles, and their vocal folds are small; nonetheless, they command a wide range of vocalizations. The right lung is composed of four lobes (cranial, middle, caudal, and accessory), whereas the left lung is composed of three lobes (cranial, middle, and caudal). In the heart, as in other mammals, the right atrioventricular valve is tricuspid and the left is bicuspid. There is usually one septomarginal trabecula
(moderator band) within the lumen of the right ventricle; there is rarely such a band within that of the left ventricle.6
Gastrointestinal System The dental formula of guinea pigs is 2(I11, C00, PM11, M33) = 20. All teeth are elodont and grow throughout life. In animals with dental malocclusion, the maxillary cheek teeth tend to overgrow laterally into the buccal gingival; the mandibular cheek teeth tend to overgrow in a medial direction, entrapping the tongue (see Chapter 32). The incisors are normally white, unlike those of many other rodents. Between the incisors and the cheek teeth (premolars and molars) is a gap called the diastema. Guinea pigs have large tongues and relatively small, narrow oral cavities. The soft palate is continuous with the base of the tongue. The oropharynx communicates with the remainder of the pharynx through a hole in the soft palate called the palatal ostium.58 Take care in attempting to pass any instrument (such as a feeding tube) that the instrument does not slip to either side of the ostium, where it can damage the vascular soft palate. Guinea pigs have four pairs of salivary glands—parotid, mandibular, sublingual, and molar—the ducts of which empty into the oral cavity near the molars. The entire alimentary tract measures approximately 2.3 m from pharynx to anus.30 The stomach is lined with glandular epithelium; unlike the stomachs of rats, mice, and hamsters, there is no nonglandular portion.6 The small intestine is located on the right side of the abdominal cavity, and the cecum occupies the central and left portions. The cecum is a large, thin-walled sac with many lateral pouches formed by the action of three taeniae coli (thick, longitudinal muscular bands that run the length of the large intestine). Smooth muscle cells harvested from the taeniae coli of guinea pigs were commonly used in physiologic studies because they could be obtained easily without damaging the gut. The cecum is 15 to 20 cm long and contains up to 65% of the gastrointestinal (GI) contents.34 The liver has six lobes—right, medial, left lateral, left medial, caudate, and quadrate. The gallbladder is well developed. The normal gastric emptying time in guinea pigs is 2 hours. Total GI transit time is approximately 20 hours (range 8- 30 hours); however, when coprophagy is factored in, the total GI transit time is 66 hours.30
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The mean gastric pH in laboratory guinea pigs is 2.9, whereas the small intestinal pH is in the range of 6.4 to 7.4.35 Water content in the GI tract in relation to body weight is higher in the guinea pig than in the rabbit, but water content is less than in the rabbit when normalized to unit length of the gut. As herbivorous hindgut fermenters, guinea pigs are coprophagic and may ingest feces from the anus many times per day.17 Obese or pregnant animals may eat fecal pellets from the floor, and young, unweaned guinea pigs can be seen eating the dam’s droppings. Coprophagy appears to be an important function, although its contribution to the nutritional needs of guinea pigs has not been fully characterized. As in rabbits, coprophagy may be a source of B vitamins and a means of optimizing protein utilization.8 However, unlike rabbits, guinea are not cecotrophic; whereas cecotrophs provide a rich source of B vitamins for rabbits, guinea pigs require a dietary source of 7 out of 10 B vitamins, while rabbits require a dietary source for only 3.57 If coprophagy is prevented, guinea pigs lose weight, digest less fiber, and excrete more minerals in the feces.8,17 Small herbivores have one of two colonic separation mechanisms necessary for coprophagy.7,21 Guinea pigs and chinchillas exhibit a “mucus trap” strategy, in which bacteria from the cecum are trapped in mucus in the colon with few to no food particles and returned to the colon by antiperistalsis. Rabbits and other lagomorphs exhibit a “wash-back” strategy, in which bacteria, solutes, and small food particles are returned to the cecum by antiperistalsis in a stream of water from the proximal colon. The mucus-trap strategy is less efficient than the wash-back strategy in extracting bacteria from the colonic digesta.21 Therefore the colon of guinea pigs is comparatively larger and heavier than that of rabbits, which may be a factor in the ability of rabbits to run faster than other similarly sized mammals.21 Guinea pigs and chinchillas have a colonic furrow in the ascending colon, in which the concentration of bacteria and nitrogen is twice as high as in the lumen. Bacteria in the proximal colon are transported in the furrow into the cecum as part of the separation mechanism.26 Geriatric guinea pigs may develop fecal impactions within the anus, perhaps because of a loss of muscle tone or inability to eat feces directly from the anus. These impactions can be relieved by gentle manual expression, which may have to be repeated weekly. Like the GI flora of rabbits, that of guinea pigs is primarily gram-positive.39 Anaerobic lactobacilli are the predominant bacterial species in the large intestine.8
Urogenital System The accessory sex glands of male guinea pigs (boars) include the vesicular glands, prostate gland, coagulating glands, and bulbourethral glands. The vesicular glands are long, coiled, blind sacs that lie ventral to the ureters and extend 10 cm into the abdominal cavity.6 Do not mistake these for uterine horns! The testes are located in the open inguinal canals. Guinea pigs have an os penis. A pouch containing two horny styles (slender projections) is located caudoventral to the urethral opening. During erection, the pouch is everted and the styles project externally. Female guinea pigs (sows) have paired uterine horns, a short uterine body (12 mm long), and a single os cervicis opening into the vagina. Guinea pigs have a vaginal closure membrane that opens at estrus, at parturition, and, in many animals, at day 26 or 27 of gestation.65 The renal pelvis is relatively large and has a single longitudinal renal papilla. The alkaline urine is normally thick and
Table 22-1 Physiologic Values for Guinea Pigs Usual life span as pet Adult weight Sexual maturity Type of estrous cycle Length of estrous cycle Ovulation Gestation period Litter size Normal birth weight Weaning age Rectal temperature Average blood volume Heart rate
5-6 years Males, 900-1200 g; females 700-900 g Males, 3 months; females, 2 months Nonseasonally polyestrous 15-17 days Spontaneous 59-72 days (average, 68 days) 1-13 (2-4 is usual) 45-115 g (70-110 g is usual) 21 days (or at 180 g body weight) 99.0-103.1°F (37.2-39.5°C) 70 mL/kg 240-310 beats per minute
cloudy white or yellow. Like the urine of other herbivores, it contains many crystals.
Sexing Guinea pigs are easily sexed. Boars have obvious scrotal pouches and large testes (Fig. 22-3). A flat area of tissue lies between the urethral opening and the anus, in which a longitudinal shallow slit may be present, marking the junction of the two scrotal pouches on the midline. The penis can be everted from the prepuce by placing gentle pressure at its base. Sows have a Y-shaped depression in the perineal tissues. The top branches of the Y point cranially and surround the urethral opening (Fig. 22-4). The vulvar opening lies at the intersection of the branches, and the anus is located at the base of the Y. In immature males, the penis can be prolapsed out of the prepuce; in immature females, the Y-shaped tissue depression is evident.
Reproductive Behavior and Breeding Reproductive values for guinea pigs are presented in Table 22-1. Many reported features of the reproductive cycle (e.g., length of estrus and the estrous cycle, timing of postpartum estrus, litter size) vary according to the source, presumably because of variations among strains of guinea pigs. The numbers reported here are compiled from several different sources and are referenced accordingly. Puberty, defined as age at first conception, occurs at 2 months of age in females and at 3 months in males.65 Males begin to mount at 1 month of age, and ejaculation is evident by the time they are 2 months old.23 The peak reproductive period for females is from 3 or 4 months to 20 months of age; pet animals may reproduce until they are 4 or 5 years old.34 Guinea pigs are polyestrous and breed year-round in a laboratory setting. The estrous cycle in most females is 15 to 17 days (range 13-21 days), and ovulation is spontaneous.34 Fertile postpartum estrus occurs in most females from 2 to 10 hours after parturition.52 Females show distinct signs of proestrus and estrus. During proestrus, they become more active and may chase their cage mates; they may sway their hindquarters and utter a distinct guttural sound. Estrus lasts 6 to 11 hours, during which time females show lordosis, or the copulatory reflex—an arching and straightening of the back with elevation of the rump and dilation of the vulva. In mature
CHAPTER 22 Biology, Husbandry, and Clinical Techniques sows, the vaginal membrane is open for approximately 2 days during estrus; it closes after ovulation. Copulation in guinea pigs (and chinchillas) can be confirmed by finding the vaginal (copulatory) plug, a solid mass of coagulated ejaculate that falls out of the vagina several hours after mating. Rodent copulatory plugs are typically hard and rubbery or waxy in consistency and are the exclusive product of male secretions. Copulatory plugs are not restricted to rodents and have been reported in some bats, insectivores, primates, and marsupials.16 There are several possible functions of rodent copulatory plugs.60 They may (1) store sperm, (2) prevent sperm leakage, (3) induce pseudopregnancy, (4) effect sperm transport, or (5) prevent later fertilization of the female by other males. Current hypotheses suggest that the primary function of the copulatory plug is the enforcement of chastity in polygamous breeding rodents. The other functions, especially sperm transport, are regarded as incidental but necessary effects. In guinea pigs, the copulatory plug prevents competing ejaculate from reaching the site of fertilization. This is also the case with chinchillas. Like the placenta of humans, that of guinea pigs is hemochorial, meaning that the trophoblasts are in contact with maternal blood. The duration of gestation is 59 to 72 days (average, 68 days), depending on the strain of guinea pig, parity of the sow, and litter size. Gestation is typically shorter in primiparous sows and in those with small litters. The fetuses can be palpated as early as 15 days of gestation, although they are more evident at 28 to 35 days. Impending parturition is signaled by separation of the pubic symphysis; a gap of 15 mm is palpable about 2 days before parturition and increases in width (up to 25 mm or more) at the time of parturition.23 This separation may be inadequate in sows that are bred for the first time after 7 or 8 months of age, and dystocia commonly results. Other causes of dystocia include obesity and large fetal size. Suspect dystocia in gravid sows that show depression or a bloody or discolored vaginal discharge; an emergency cesarean section is indicated in most cases (see Fig. 22-4). Normal parturition is typically rapid, with only a few minutes between births. Guinea pigs do not build nests. The average litter size varies according to guinea pig strain and management practices but is typically 2 to 4,52,65 with a range of 1 to 13 young reported.65 Birth weights range from 45 to 115 g and are inversely related to litter size21; young weighing less than 60 g rarely survive.23 Although otherwise completely herbivorous, guinea pigs are placentophagic. In addition to the mother, males and other females may also consume the placenta. Newborn guinea pigs (typically called pups or young but not piglets) are precocious, meaning that they are fully furred, their eyes are open, and they are able to stand shortly after birth. They are not, however, able to fend for themselves at this time.65 The pups ideally should receive sow’s milk for a minimum of 5 days, and the normal lactation period is 3 weeks.65 Pups often do not survive if they fail to receive sow’s milk for the first 3 to 4 days of life.23 Guinea pig sows are not very “motherly.” They passively allow nursing to occur rather than seeking out the young. Lactating sows permit the young of other females to nurse. Because voluntary micturition does not occur until the second week of life, the sow must lick the pup’s anogenital region to stimulate urination and defecation. The young are weaned at 21 days, or at a weight of 180 g (15-28 days of age).34
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Orphaned guinea pigs should be fostered to a lactating sow if feasible. If there is no suitable foster mother, the young can be fed from a dropper or pet nurser beginning at 12 to 24 hours after birth. One author recommends feeding the pups every 2 hours until 5 days of age, after which feeding every 4 hours becomes sufficient. The hand-rearing formula should approximate guinea pig milk, which contains 4% fat, 8% protein, and 3% lactose.23 Evaporated milk mixed with an equal amount of water can be used. Guinea pigs begin nibbling on solid food at 2 days of age, and guinea pig pellets moistened with water or formula can be offered at that time.
BEHAVIOR Guinea pigs are social animals that seek physical contact with other guinea pigs when housed together. They often stand side by side when resting and crowd together at feeders. However, there is little mutual grooming. Hair pulling can be a form of aggression, and hair pulling and ear nibbling of subordinate animals is seen in crowded or stressful environments.24 The vocalizations of guinea pigs have been well characterized. Recognized call types include the chutt, chutter, whine, tweet, whistle (single or in long bouts), purr, drr, scream, squeal, chirp, and grunt.5,24 Many guinea pig owners are familiar with the excited squeals emitted by their pets when a refrigerator door is opened or feeding is imminent.
HUSBANDRY Housing Housing for guinea pigs should be set up with the knowledge that healthy guinea pigs produce prodigious amounts of feces, often defecate in food and water containers, turn over any unstable container, and are known to inject a premasticated slurry of pellets into the tubes of their sipper bottles. That said, guinea pigs require relatively simple housing. In a laboratory setting in the United States, the minimum required floor space is 652 cm2 (101 in.2) per adult animal; however, guinea pigs in a laboratory are often group-housed and each animal has more than this amount of space. Pets should be provided with at least twice as much floor space. Cages may be constructed of plastic, metal, or wire. Good ventilation is important. If a solid-sided cage, such as an open glass aquarium, is used, change the bedding frequently to minimize ammonia levels in the cage. Guinea pigs do not jump or climb; therefore the top of the cage does not have to be enclosed. The cage walls should be at least 25 cm (10 in.) in height. Solid cage flooring is preferable to wire mesh: foot and leg injuries are more common in guinea pigs that are kept on wire. Guinea pig breeders often use 12 × 38 mm wire mesh to minimize the potential for leg injuries. Cellulose fiber bedding is excellent for use on solid floors, although newspaper, shredded paper, wood shavings, and straw may be used. A small upside-down cardboard box provides shelter within the cage, although some guinea pigs prefer chewing the box to hiding in it. Place the cage in a quiet area out of direct sunlight. The recommended temperature range for guinea pigs is 65° to 79°F (18° to 26°C).23 Because of their susceptibility to hyperthermia, guinea pigs are better able to tolerate cool than warm environments and should not be exposed to high levels of temperatures and humidity.
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For breeding purposes, guinea pigs are usually housed in a harem-style arrangement with a single boar and 1 to 10 sows in a pen.23 They can also be housed in pairs. In intensive breeding systems, the sow and young are left in the pen so that the sow can be rebred at the postpartum estrus. However, removal of the sow and young to a nursery area shortly after parturition minimizes trampling and ear chewing of the young by other adults.
Nutrition and Feeding Guinea pigs develop dietary preferences early in life and do not adapt readily to changes in type, appearance, or presentation of their food or water. Even a change in the brand of pelleted feed can result in refusal of food. It may be a good idea to expose pets, while they are still young, to small amounts of different guinea pig chows and vegetables so that they become accustomed to variety. Always teach clients about this characteristic of guinea pigs to prevent a potentially dangerous, self-imposed fast by a pet guinea pig that is fed new food. Wild cavies eat many different types of vegetation. Domestic guinea pigs are also completely herbivorous (with the exception of placentophagy). They digest fiber more efficiently than rabbits do.8 Interestingly, they do not increase their food intake, as do rabbits and many other species, when cellulose or other fiber is used to dilute the diet. This suggests that satiety in guinea pigs is governed more by distention of the GI tract than by metabolic energy need.8 A crude protein level of 18% to 20% is adequate for growth and lactation, and the recommended minimum level of crude fiber is 10%.7 Guinea pigs require a dietary source of vitamin C (ascorbic acid) because they lack l-gulonolactone oxidase, an enzyme involved in the synthesis of ascorbic acid from glucose. Nonbreeding adult guinea pigs require 10 mg/kg daily of ascorbic acid. Higher levels should be provided for growing and pregnant animals; 30 mg/kg daily is recommended during pregnancy.23 The recommended diet for pet guinea pigs consists of guinea pig pellets and grass hay, supplemented with fresh vegetables. Usually the pellets are offered by free choice, although some clinicians believe that, as with rabbits, a limited quantity of this nutritionally rich food source is best for sedentary adult guinea pigs. Good-quality grass hay should be available at all times. Guinea pigs enjoy a variety of leafy greens, and these can be offered in handfuls. All fresh foods should be washed and prepared as though for human consumption, and they should be removed from the cage after a few hours if not eaten. Fruits, rolled oats, and dry cereals should be offered only in very small quantities, if at all, as treats. Any additions or changes to the diet should be made gradually. Commercially available guinea pig pellets usually contain 18% to 20% crude protein and 10% to 16% fiber.23 Pellets are fortified with ascorbic acid; however, approximately half of the initial vitamin C content may be oxidized and lost 90 days after the diet has been mixed and stored at 22°C. Many commercial diets are now available with stabilized vitamin C; these diets should be stored in a cool, dry area (<70°F [22°C]). Foods can be refrigerated or frozen but should be protected from condensation and increased storage temperature and humidity in vegetables and fruits or in the drinking water. Foods that contain high levels of ascorbic acid are red and green peppers, broccoli, tomatoes, kiwi fruit, and oranges. Many types of leafy greens (kale, parsley, beet greens, chicory, spinach) are high in vitamin C, but many contain high levels of calcium or oxalates; these should be offered in only small amounts. Vitamin C can be
added to the water at 1 g/L.13 In an open container, water with added vitamin C loses more than 50% of its vitamin C content in 24 hours. Aqueous solutions of vitamin C deteriorate more rapidly in the presence of metal, hard water, or heat. Vitamin C is more stable in neutral to alkaline solutions. Water must be changed daily to ensure adequate activity of the vitamin.
BIOLOGY AND HUSBANDRY OF CHINCHILLAS Chinchillas, like guinea pigs, originated in South America. The common name chinchilla may derive from the Quechua words chin, “silent”; sinchi, “strong” or “courageous”; and lla, a diminutive. The Spaniards in the sixteenth century copied the word because Quechua Indians used chinchilla pelts to decorate their ceremonial dress.43 Chinchillas are nearly extinct in the wild because of extensive hunting for their pelts. In the wild, chinchillas inhabit cool, semiarid, rocky slopes at elevations in relatively barren areas of the Andes from 3,000 to 5,000 m (10,000-16,600 ft) above sea level.55 Consequently they have a higher hemoglobin oxygen affinity than other pet rodents and rabbits.44 Chinchillas live in burrows or rock crevices but are well adapted for running. They dust bathe, are vegetarian, and are active throughout the year. They are gregarious, living in groups of several hundred. The small-bodied, large-eared, and long-tailed form, Chinchilla lanigera, from central Chile, was domesticated in the United States from 13 individuals brought to California in 1927 by Matthew Chapman.13 (The name laniger is from Latin, meaning “woolly”). It took nearly 12 months to capture them. These animals are the ancestors of all North American domestic chinchillas. The large-bodied, small-eared, and short-tailed form, Chinchilla brevicaudata, from the Altiplano (the highlands of Peru, Bolivia, northern Chile, and Argentina) was domesticated in Chile around 1931.22 Comparison of the mitochondrial cytochrome-b gene sequence shows that both chinchilla species differ at 22 sites and the average genetic distance is approximately 6%.54 A few reports suggest that some crosses between C. lanigera and C. brevicaudata have occurred because of captive breeding.22 Hybrid males are infertile. Hybrid females are fertile, but when backcrossed, two-thirds of the second-generation hybrids are sterile.22 Although no traces of C. brevicaudata mitochondrial cytochrome-b gene variants, wild or domestic, were found in any of five domestic C. lanigera, sequences indicated a small difference between domestic C. lanigera and wild C. lanigera.54
ANATOMY, PHYSIOLOGY, AND BEHAVIOR General Characteristics The chinchilla has a slender body with short forelimbs and long muscular hind limbs that give it a rabbit-like appearance. The head, eyes, and ears are relatively large and the bullae are greatly expanded. Chinchillas have long whiskers and a bushy tail. They usually weigh between 400 and 600 g, and females tend to be larger than males (Table 22-2). Chinchillas have four toes on both front and rear feet, all with small, weak claws. There is no fur on the palmar and plantar regions of the feet. They have very large, thin-walled auditory bullae, which are readily visible on radiographs Chinchillas are quiet, shy, and agile. In the wild, chinchillas live in burrows and rock crevices; they have long hind limbs and feet adapted for leaping. The long plumed tail acts as a balance when the animal is sailing through the air. Chinchillas require
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Table 22-2 Physiologic Values for Chinchillas Usual life span as pet Adult weight Sexual maturity Type of estrous cycle Length of estrous cycle Ovulation Gestation period Litter size Normal birth weight Weaning age Rectal temperature Heart rate
10 years (up to 20 years reported) Males, 400-500 g; females 400-600 g 8 months Seasonally polyestrous (November to May) 30-50 days Spontaneous 105-118 days (average, 111 days) 1-6 (2 is usual) 30-50 g 6-8 weeks 98.6-100.4°F (37-38°C) 100-150 beats per minute
larger cages than do their less active cousins the guinea pigs. In their natural habitat, chinchillas are active at dusk and at night; however, in captivity, they can be active during the day. They readily habituate to humans if handled frequently while young. Flight is their defense mechanism; rarely, they bite. Chinchillas are virtually odorless, although one author reported that frightened animals produced secretions that gave off an odor, similar to that of scorched almonds, from glands inside the anus.37 Females reportedly may stand up on their hind legs and spray urine at a presumed attacker. Anecdotally, wild chinchillas are described as living to 10 years of age, with 11.3 years as the longest reported life span. Captive chinchillas are claimed to live longer than 20 years and some are claimed to have bred at 15 years. Documented life span is different. The maximum recorded life span in the wild is 6 years.31 The documented mean life span in zoos is 4 years (the maximum is 7 years). However, these are data taken from 16 individuals in the London Zoological Gardens, 1848 to 1894, which averaged a life of 4 years, 3 months, 24 days, the maximum being 6 years, 10 months, 5 days.20 The maximum recorded life span for a chinchilla in research is 15 years.65 The hair coat is luxurious, soft, and very dense. As many as 60 hairs grow from a single hair follicle.70 Chinchillas come in a variety of colors. The natural wild-type color is bluish-gray with yellow-white underparts. Through selective breeding, the most common color seen is dark blue gray (the dominant fur color gene).29 Other colors have emerged as mutations of the original standard color and include the dominant colors of beige, white, and ebony; recessive colors include sapphire, violet, charcoal, and velvet, which are varying shades of gray. Eye color may be black or pink to red due to coat color genes. Homozygous white or homozygous black combinations are lethal. Frequent dust baths are necessary to maintain the health of the fur (see “Dust Baths,” below). When frightened, chinchillas can shed patches of fur, a condition known as “fur-slip.” The hairless patches take 6 to 8 weeks to fill in; it may take several months for the patches to become indistinguishable from surrounding fur. The vocal repertoire of chinchillas comprises 10 different sounds.51 These are attributed to the behavioral contexts of exploratory behavior, predator avoidance, sexual behavior,
Fig. 22-5 Radiograph of the skull of a normal chinchilla. Chinchillas have very large tympanic bullae and are commonly used laboratory species for auditory research.
and social behavior, including social contact and antagonistic behavior (defensive and offensive). Chinchillas can raise and lower the tones of the calls they make. All chinchillas have a basic cry that will be used commonly from birth. The eyes are large and sit in a shallow bony orbit. The iris is densely pigmented with a vertical pupil, both features consistent with the chinchillas’ habit of basking in the sun in their high-altitude habitat.13 A large and exposed corneal segment, a vertical slit pupil, and a heavily pigmented iris characterize the eyes of chinchillas.45 A third eyelid is only rudimentarily developed as a small conjunctival fold at the medial canthus.45 The lacrimal drainage system consists of two lacrimal punctae at the medial canthus of the upper and lower eyelids; these drain into a single lacrimal duct. The lacrimal duct runs in a similar course as in other rodents and lagomorphs.9,11 Because of their very large tympanic bullae, chinchillas are common laboratory species for auditory research (Fig. 22-5).
Gastrointestinal System The dental formula of chinchillas is the same as that of guinea pigs: 2 (I11, C00, PM11, M33) = 20. Chinchillas fall into the small group of caviomorph rodents (i.e., South American hystricomorphs such as guinea pigs, chinchillas, and degus) whose teeth are all elodont. Their teeth have adapted to a more voluminous abrasive diet and have large chewing surfaces. All the teeth are open-rooted and grow continuously throughout life to compensate for the wear that occurs during prolonged chewing.10,59 The incisors grow 5 to 7.5 cm (2-3 in.) per year and are normally yellow in adult animals. Upper incisors grow faster than lower incisors.9 The oral cavity is small and narrow. Like guinea pigs, chinchillas have a palatal ostium—an opening in the soft palate through which the oropharynx communicates with the rest of the pharynx.58 Captive-bred chinchillas tend to have significantly longer cheek teeth compared with their wild counterparts, and reduced chewing of a less abrasive diet in captivity is suspected
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to diminish tooth wear and therefore lead to elongation of the continuously growing (elodont) cheek teeth (see Chapter 32).10 Chinchillas have a long GI tract. The small and large intestines in one adult animal measured 3.5 m (11.5 ft).72 In a wild specimen with a total body length of nearly 21 cm, total length of the small intestine was 117 cm and of large intestine, without cecum, 145 cm.55 Compared with guinea pigs, the jejunum and descending colon are long.12 The cecum is relatively large and coiled; the colon is highly sacculated. The cecum of a chinchilla holds less of the contents of the large intestine than that of a rabbit or guinea pig. According to one study, the cecum of a chinchilla held 23% of the dry matter content from the large intestine; that of a rabbit held 57% and that of a guinea pig 44%.26 Chinchillas eat mainly at night, ingesting more than 70% of their total daily intake in the dark. Fecal excretion is predominantly at night.27 Chinchillas produce two types of fecal pellets. One is nitrogen-rich, intended for cecotrophy; the other is nitrogen-poor, excreted as fecal pellets.26 Chinchillas defecate between 0300 and 0600 hours (3 a.m. and 6 a.m.) and consume cecotropes between 0800 and 1400 hours (8 a.m. and 2 p.m.).26 Mean transit time of food in the GI tract is 12 to 15 hours, similar to that of other rodents. However, in contrast to other pet rodents, transit time in chinchillas is not affected by reducing the dietary fiber level.27 The insulins of chinchillas and other hystricomorph rodents (e.g., guinea pigs, degus) are highly divergent from those of other mammals. Hystricomorph insulins exhibit a low biological potency relative to pig insulin, yet the receptor-binding affinity is significantly higher, indicating that the efficacy of these insulins on receptors is about twofold lower than that of pig insulin.41,42 Hypoglycemia is always a great risk in treating diabetes with recombinant human insulin or porcine insulin.
A
Urogenital System Chinchillas produce concentrated urine. In 10 chinchillas deprived of water for up to 8 days, the mean urine osmolality was 3.505 mOsm/kg (range 2.345-7.599 mOsm/kg).68 Females have two uterine horns and two cervices.63 Chinchillas have three pairs of mammary glands: one inguinal pair and two lateral thoracic pairs.65 Several reports have erroneously stated that chinchillas have two pairs of mammary glands. Male chinchillas do not have a true scrotum. Instead, the testes are contained within the inguinal canal or abdomen, and there are two small movable sacs (the postanal sacs) next to the anus into which the caudal epididymis can drop.63 The external appearance of the scrotal sacs is similar to that of the nonpendulous scrotum of pigs and cats. Testes are relatively large. Those from four wild mature males were 21 to 29 mm long (mean 26.2 mm), with a mass of 3.0 to 5.1 g (mean 4.3 g).55 The penis is readily apparent below the anus, from which it is separated by an expanse of bare skin. The penis can be manually extruded 1 to 2 cm when flaccid. The tip of the erect penis extends to the level of the axilla, a distance of about 11 cm. Male chinchillas possess unusually well developed and elaborate male accessory reproductive glands. The secretions of the male accessory reproductive glands form a hard plug that remains in the female tract after copulation. In chinchillas, the vesicular gland provides the bulk of the accessory gland secretions and the fluid hardens or gels when mixed with prostatic secretions. A 2- to 3-in.-long 1-in.-diameter irregularly shaped firm waxy plug is often found in the cage after mating.63 This is normal.
B Fig. 22-6 A, Normal external genitalia of an adult male chinchilla. The penis is withdrawn into the prepuce. The urethral orifice is seen at the top of the picture. Below the anus are the postanal or scrotal sacs. The epididymis can be dropped into each scrotal sac. However, the testes remain within the body. B, Male chinchilla with penis protruding from prepuce. The anus is obscured by the caudally directed penis.
Sexing As in other rodents, the anogenital distance gives the best initial indication to the animal’s sex. In males, the distance is greater. Extrusion of the penis from the urethral orifice will confirm the sex of the chinchilla as long as the clitoris is not mistaken for a penis. There are major differentiating features: the penis is significantly larger than the clitoris, and the extruded penis can be separated and distinguished from the prepuce, whereas the extruded clitoris tends to evaginate and the clitoral prepuce is not apparent (Figs. 22-6, 22-7). This difference between the
CHAPTER 22 Biology, Husbandry, and Clinical Techniques
A
B Fig. 22-7 A, Normal external genitalia of an adult female chinchilla in anestrus. The urethral orifice is located on the protruding urethral papilla (at the top of the picture) and separated from the vaginal orifice, which is sealed by a membrane (and not apparent in this picture) in anestrus and pregnancy. The anus is seen at the bottom of the picture. B, External genitalia of a female chinchilla shortly after giving birth. The urethral orifice is located on the protruding urethral papilla. In contrast to (A), the vaginal orifice is open, and blood-stained postpartum fluid is oozing out. The vaginal orifice is open only at estrus and during birth. The anus is located below the vaginal orifice.
sexes is evident even at birth: the urinary papilla is adjacent to the anus in females, whereas the penis is separated from the anus by a narrow band of tissue in males. Eversion of the penis from the urethral orifice confirms the sex of the chinchilla.
Reproductive Behavior and Breeding Female chinchillas are seasonally polyestrous and the breeding season in the northern hemisphere is approximately from November to May.4 Estrus lasts 3 to 4 days and the entire estrus
287
cycle about 28 to 35 days.32,64 Age at puberty in males, defined as age at successful conception, is 8 months or older; in females, the average age at puberty is 8.5 months (range 2-14 months).63 In females, a vaginal closure membrane seals the vulva at all times except during estrus and parturition and in cases of underlying disease. The vaginal closure membrane is open only during parturition and for 2 to 4 days during estrus (Fig. 22-7).65 The vulva (often referred to in books as the vaginal orifice) is U-shaped and situated between the anus and the moundshaped urethral orifice; it is difficult to distinguish when closed, and is “indicated” by a slightly, raised semicircular area. When its closure membrane covers the vaginal orifice, the urethral orifice can be mistaken for a genital opening. The well-developed clitoris of female chinchillas and guinea pigs can be manually extruded through the urethral orifice and mistaken for a penis. The vagina is open during estrus. During these times, the vaginal closure membrane dissolves and is then repaired. During estrus a mucoid vaginal discharge occurs; however, there is no vulval swelling. Rather, there is a change in perineal color, which goes from a dull flesh color to a deep red. The color of the perineum increases dramatically at the time of vaginal perforation and remains intense throughout most of luteal phase of cycle. Cytologic characterization of vaginal smears can be helpful to differentiate between physiologic and pathologic conditions causing vaginal discharge.4 The predominant cells during estrus are cornified superficial epithelial cells. Neutrophils are absent during estrus but common during proestrus and metestrus.4 Chinchillas can be housed in pairs or in polygamous units, with a single male and two to six females. The polygamous units used by breeders are set up with separate cages for the females, each with a rear door onto a common runway used by the male. The male can go through any open door at will, but the females wear collars that prevent exit from their cages. In a polygamous setting, the male is kept out of the female’s cage during parturition and raising of young; however, in a pair setting, the male can often remain with the female during this time if she tolerates his presence. Breeders facilitate mating by observing changes in the vaginal closure membrane and performing vaginal cytology. Pregnancy is detectable by palpation at 90 days’ gestation and may be determined by regular weighing. After 6 weeks, weight gain in pregnant chinchillas will increase rapidly. Gestation averages 111 days (see Table 22-2), and there are usually two young in each litter (range 1-6).65 Pet chinchillas have smaller litters and more males per litter than do chinchillas from fur ranches or laboratory colonies.65 Parturition typically occurs in the early morning (before 8 a.m.) and only rarely late at night.63 Dystocia is uncommon in well-managed breeding establishments. Although chinchillas do not build nests,72 the females can learn to use a nest box that, when heated, prevents the firstborn young from becoming hypothermic while the rest of the litter are being born.37 Chinchillas, like guinea pigs, are placentophagic. Blood on the nose and front paws of the female indicates that she has eaten the placenta and the birthing process is over.37 Fetal reabsorption occurs frequently and may take place at any stage of pregnancy. Even in late stages, when the skeletal tissue of the fetus has formed, reabsorption rather than mummification or spontaneous abortion occurs. Neither placental nor fetal tissue can be recognized in the necrotic mass, but a central blood-filled cavity is usual.50 Chinchilla young are precocious, weighing 30 to 50 g at birth, and are fully furred with teeth and open eyes. They are able to
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walk within 1 hour after birth. The dam stands rather than lying down while they nurse, so infants often lie on their backs during suckling.37 If the mother dies after birth, another lactating female will usually accept the newborn young, especially if they are close in age to her own. According to one author, a lactating guinea pig may be an appropriate foster mother in some cases.62 The growth rate is 3.6 g per day during the first month, decreasing to 1.56 g per day from 2 to 6 months, and to 0.65 from 6 to 12 months.40 Weaning is normally at 6 to 8 weeks, and the minimal period of suckling necessary for survival is 25 days.40 Hand-feeding is necessary if a foster mother is not available or if supplemental nutrition is needed for litters of four or more. A formula of equal parts evaporated milk and water can be administered with an eye dropper or pet nurser. One author recommends adding glucose (1g/15 mL) to this formula.63 For the first 3 or 4 days, the young should be hand-fed as often as possible during the day, with no more than 4 hours between feedings, and once or twice at night. After this time, the night feedings can be dropped and the intervals between daytime feedings gradually lengthened.63 Chinchillas begin to eat solid foods at 1 week of age. In family groups, fathers are tolerant and fairly friendly, sitting with mother and young in a protective manner.33 Juveniles display frisky-hop playing, which includes vertical leaps, body twisting, head tossing, racing and pivoting, and prancing with kicking back of the hind feet.33 The young of large litters may fight over access to the teats, and it may be necessary to clip the incisor teeth to prevent serious injury to the teats or to siblings. To minimize fighting, food bowls should be large enough to accommodate the entire litter simultaneously.
HUSBANDRY Housing Chinchillas are very active, acrobatic animals and require a lot of space. According to one author, the enclosure should be at least 2 × 2 × 1 m (6.6 × 6.6 × 3.3 ft), with a wooden nesting box measuring 30 × 25 × 20 cm (12 × 10 × 7 in.).62 Large multilevel cages that provide sufficient space for climbing and jumping are excellent for housing pet chinchillas.67 Because chinchillas chew wooden cages, the cage should be made of 15 × 15 mm welded wire mesh, with or without an area of solid flooring. Drop pans below the cage facilitate cleaning. Chinchillas are shy animals, and in captivity they need a place to hide. In the wild, chinchillas conceal themselves in rock crevices. Polyvinyl chloride (PVC) plumbing pipes, especially elbows and Y and T sections, make ideal hiding places and can be sanitized in a dishwasher. The pipes should be 10 to 13 cm (4-5 in.) in diameter. Alternatively, clay pipes of a similar diameter can be used. Chinchillas are easily housed in either mesh- or solid-bottom cages, although solid-bottom cages are recommended for pregnant females about to have young. Ensure that mesh spacing in cages is narrow, as tibial fractures commonly occur in young animals when a chinchilla catches its leg in a cage bar. Chinchillas can be housed in pairs, colonies, or polygamous units, although colony housing is not advised for breeding chinchillas.37 The cage setup for polygamous units was described earlier. Chinchillas are very tolerant of cold but sensitive to heat. The ambient temperature range to which chinchillas are adapted is 18.3° to 26.7°C (65°-80°F). Chinchillas do best in
a dry environment at relatively cool temperatures. One recommended temperature range for housing chinchillas is 50° to 68°F (10°-20°C).62 Temperatures lower than 65°F (18°C) and relative humidity lower than 50% promote good fur growth.62 Chinchillas do not tolerate dampness and are prone to heat stroke at environmental temperatures greater than 82° to 86°F (28°-30°C). Chinchillas will develop matted fur if kept in a warm (greater than 26.7°C [80°F]), humid environment.
Dust Baths Access to a dust bath should be provided daily, if possible, or at least several times per week. Dust bathing reduces fur lipids of chinchillas.3 Sanitized chinchilla dust is available commercially at pet stores. Two commercial dust baths, Blue Cloud and Blue Sparkle, can be obtained from various web-based chinchilla supply stores. Alternatively, a 9:1 mixture of silver sand and Fuller’s earth can be used.28 Fuller’s earth is a variety of kaolin that contains aluminum magnesium silicate. The name is derived from the ancient process of cleaning, or fulling, wool to remove the oil and dirt particles with a mixture of water and earth or clay. Beach or playground sand is not suitable for dust baths. Some people are allergic to the commercially available powders. It is possible to make a homemade dust bath preparation consisting of perfume-free talc powder (also known as talcum or French chalk) and food-grade cornstarch. Food-grade cornstarch (marketed as Maizena or Mondamin) is best. Avoid using soluble starch, which is potato starch or cornstarch treated with dilute hydrochloric acid. Some breeders eliminate or reduce the amount of cornstarch used for nursing mothers because the babies get it in their noses and develop rhinitis.49 More recently, volcanic ash from Mt. St. Helens and other locations has become popular as a dust bath.49 This is a very fine powder that can cause conjunctivitis and rhinitis; therefore, limit access to this type of dust bath for 3 to 4 minutes.49 The dust is placed at a depth of 2 to 3 cm (1 in.) in a pan, such as a plastic dishpan, that is big enough for the chinchilla to roll around in.36 A chinchilla in the wild may spend up to an hour dust bathing, rolling, and fluffing its fur.56 The dust bath can be kept clean and free of feces by removing it from the cage after use. Excessive use of dust baths can lead to conjunctivitis, especially in young chinchillas.
Nutrition and Feeding In their natural habitat, the relatively barren areas of the Andes Mountains, chinchillas reportedly feed on any available vegetation, eating in the early morning and the late evening by holding the food with their forepaws while sitting on their haunches.63 Chinchillas eat mainly at night. Studies done more than 50 years ago showed the importance of grasses and hays in the diet,18 and it is recommended that chinchillas be given a diet high in fiber27; however, the specific nutrient requirements for chinchillas are still unknown. Urinary calculi, urolithiasis, metastatic renal calcification, and nephritis are reported occasionally. Calculi are typically composed of calcium carbonate. The conditions are associated with feeding a diet high in calcium, such as alfalfa hay. Commercial chinchilla diets are available (e.g., Chinchilla Deluxe, Oxbow Hay Co., Murdock, NE, www.oxbowhay.com; Mazuri Chinchilla Diet, Mazuri, St. Louis, MO, www. mazuri.com). Some diets marketed for chinchillas are in reality mixtures of rabbit, guinea pig, and rodent pellets. As such, they provide a diet supplemented with vitamin C that is lower
CHAPTER 22 Biology, Husbandry, and Clinical Techniques in protein and fat than standard rodent chow and has the same fiber content as a rabbit maintenance diet. However, the pellets are longer than rabbit or guinea pig pellets and therefore are easier for the chinchilla to hold. The accepted formula for chinchilla pellets is 16% to 20% protein, 2% to 5% fat, and 15% to 35% bulk fiber.28,33,64 Although very little has been published about the nutritional needs of pet chinchillas, it is safe to assume that growing animals and breeding females require more calories and higher levels of calcium, protein, and fat than do nonbreeding chinchillas. Nonbreeding animals do well on a diet of good-quality grass hay supplemented with small amounts of chinchilla or rabbit pellets, fresh vegetables, and grains.28 One to two tablespoons of pellets daily should be sufficient for a nonbreeding adult animal. A pellets-only diet has insufficient roughage and can predispose the chinchilla to enteritis. Chinchillas consume their food more slowly than rabbits and guinea pigs do. Limit treats such as grains, dried apples, raisins, figs, hazelnuts, and sunflower seeds to not more than 1 teaspoon per day. As with guinea pigs, clean and prepare all food as if it were for human consumption. Any change in diet should be instituted gradually. Abrupt dietary changes will lead to temporary but dramatic (up to 50%) decreases in food intake.27 Anorexia often follows any change in diet. Have clean, fresh drinking water available at all times. Chinchillas can be trained to use automated watering devices in a laboratory setting, or they can do equally well with cage-mounted water bottles. Water in a bowl tends to get dirty quickly and can spill. Hard foods for gnawing can be offered. These may include porous stones such as pumice; young branches of trees such as elm, ash, maple, and birch; pieces of bark from apple, pear, and peach trees; and young grapevines. Advise the owner to avoid branches from poisonous trees, such as cedar, plum, redwood, cherry, and oleander.
CLINICAL TECHNIQUES FOR GUINEA PIGS AND CHINCHILLAS HANDLING AND RESTRAINT Guinea pigs are docile animals that usually need minimal restraint during a physical examination. Most will sit quietly on the examination table while the owner or an assistant places a hand on the rump so that the animal does not back away. To auscultate the heart and lungs and palpate the abdomen, gently pick up the animal in one hand. Turn it over on its back while supporting it in your hand to examine the perineal area and genitalia. Carry a guinea pig by supporting its weight in one hand and cupping its dorsum with the other. If the guinea pig is nervous or not used to handling, keep it in a carrier as much as possible and avoid excessive handling. Chinchillas should be handled calmly and gently. Docile, nonpregnant animals can be removed from a cage by grasping and lifting the base of the tail while using the opposite hand to support the body. Routine restraint can be accomplished by wrapping a towel around the body. Small chinchillas may be grasped gently around the thorax, taking care not to restrict breathing. Pregnant females should not be handled unless necessary. A protective reaction from capture in chinchillas, known as fur slip, results in the release of a large patch of fur revealing
289
Fig. 22-8 A chinchilla with fur biting, a behavioral problem, on its flank. The inset shows fur slip, a protective reaction that results in the release of a large patch of fur revealing smooth, clean skin underneath.
smooth, clean skin underneath. It may also occur with improper handling, fighting, or anything that overexcites the animal (Fig. 22-8). The fur can take several months to regrow and frequently comes up in a different shade. To prevent this, be gentle and minimize stress during capture or restraint.
PHYSICAL EXAMINATION The initial examination should involve observing the guinea pig or chinchilla in its cage. Focus on the animal’s movement, mentation, and rate and rhythm of breathing. Healthy guinea pigs have an alert demeanor with clear eyes. The animal should react to stimuli by moving or vocalizing; some animals will move very quickly. Healthy guinea pigs usually eat readily when offered treats or greens. Healthy chinchillas have a spirited curiosity and a curled tail that is carried high. Sick animals are indifferent, appear hunched, and have a dull coat; often the perianal area is stained or covered with feces. A chinchilla that flies around the cage in a frenzy when the owner attempts to capture it has not been socialized to people or other chinchillas and will be difficult to examine without sedation. Begin the physical examination by measuring the animal’s weight; this is also a good time to obtain the animal’s temperature before it becomes excited or stressed. Next, examine the fur, skin, and mucous membranes. Follow this by auscultating the heart and lungs, and then palpate the abdomen. Check the rectal area for impaction of feces in guinea pigs, and check the penis of male chinchillas for fur-ring. Observe the genitalia and mammary glands and note any abnormalities. Overgrown nails are common in guinea pigs. Often a horny growth is present and extends from the footpads, especially
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SECTION III Guinea Pigs and Chinchillas
in older animals. Trim the nails of guinea pigs and chinchillas with fingernail clippers or with cat claw clippers. The horny overgrowth can be trimmed back carefully, but avoid causing bleeding. Examine the oral cavity last; this can be stressful for the guinea pig or chinchilla, and the animal may become excited. Guinea pigs will object to examination of their teeth by squealing, and both guinea pigs and chinchillas may struggle or try to escape. To examine a guinea pig, have an assistant place one hand on the animal’s rump and the other around the shoulder and thoracic area. Use a speculum or otoscope to examine the cheek teeth, similar to the method used in rabbits. Chinchillas can be held by an assistant; use both hands to encircle the thoracic area while the animal is restrained on the table, or support the animal’s weight in one hand and restrain the forelimbs with the other. Healthy chinchillas have yellow incisors because of iron deposition on the enamel, whereas the incisors of guinea pigs are white.
BLOOD COLLECTION Venipuncture in guinea pigs and chinchillas can be difficult. The lateral saphenous and cephalic veins are the most accessible, but they are very small, and only small amounts of blood can be collected from each vein. Shave the fur from the area and wet the skin with alcohol to enhance visibility of the vein. Use an insulin or tuberculin syringe and a small (25- to 27-gauge) needle to prevent collapse of the vein. Venipuncture of multiple peripheral veins is often necessary to collect an adequate volume of blood for analysis. The jugular vein can be used to collect large blood samples; however, manually restraining a guinea pig or chinchilla for jugular venipuncture can be very stressful for these animals. Tranquilization or anesthesia for collecting blood samples may be preferable to minimize stress (see Chapter 31). Guinea pigs and chinchillas have short, thick, compact necks, and it is often difficult to locate the jugular vein. For venipuncture, restrain the animal with the forelegs extended down over a table edge and the head and neck extended up (Fig. 22-9). If necessary, shave the fur from the area to enhance visibility of the vein and use a small (22- to 25-gauge) needle and a 1- to 3-mL syringe for blood collection. If the animal shows obvious signs of stress or becomes dyspneic during jugular venipuncture, abort the procedure immediately. Observe the animal closely for several minutes after restraint is removed to see that it recovers. If the animal still appears to be stressed or dyspneic after several minutes, abandon further venipuncture attempts with the animal awake. Venipuncture of the cranial vena cava is often used for blood collection in guinea pigs. However, because of the close proximity of the cranial vena cava to the major vessels of the thoracic cavity and the heart, there is some risk of subsequent traumatic bleeding into the thoracic cavity or pericardial sac.47 For this technique, anesthesia or deep sedation of the animal is mandatory.1,48 Place the anesthetized animal on its back. Use a small (25-gauge) needle attached to a 1- or 3-mL syringe. Palpate the manubrium and insert the needle lateral to the manubrium under the first right rib at an approximate 30-degree angle to the horizontal axis of the body. Insert the needle about ½ in. (1 cm), then withdraw slowly with negative pressure until blood starts to fill the syringe. If venipuncture is unsuccessful, do not redirect the needle because of the risk of lacerating surrounding vessels. Instead, withdraw the needle and try again.1
Fig. 22-9 Restraint for jugular venipuncture in a guinea pig. Venipuncture of the jugular vein can be difficult because of the short, thick neck of guinea pigs and the stress caused by restraint. Tranquilization may be necessary.
The blood volume in guinea pigs averages 7 mL/100 g body weight.52 Approximately 7% to 10% of the blood volume (0.50.7 mL/100 g) can be safely collected from a healthy, nonanemic guinea pig. Similar guidelines can be used in chinchillas.
URETHRAL CATHETERIZATION AND CYSTOCENTESIS Guinea pigs have relatively large urethras. Catheterization of male guinea pigs is sometimes necessary to retropulse urethral calculi into the bladder. For catheterization, a 5- to 8-Fr red rubber catheter can be used with sterile technique. Extrude the penis by placing gentle pressure on the scrotum at the base of the penis. Minimize handling of the penis itself to prevent irritation and trauma, which can lead to temporary partial prolapse. Cystocentesis is sometimes necessary in animals with urethral obstruction. The method is similar to that used with other small animals, and a small (25-gauge) needle is used. Anesthesia or sedation is necessary for catheterization and cystocentesis.
CLINICAL LABORATORY FINDINGS Reference intervals of laboratory values for guinea pigs and chinchillas are available from several sources, some of which are listed.25a,34,36,38,51a,57a Representative hematologic and biochemical values are listed in Tables 22-3 and 22-4. Ideally, as for other species, each veterinary diagnostic laboratory should have established reference intervals. Clinical laboratory values can vary according to the physiologic state of the animal, sex, and the laboratory techniques used. In guinea pigs, alanine aminotransferase activity is low in hepatocytes; therefore it is not sensitive or specific as a marker of hepatocellular injury.69 Hypercholesterolemia is common in
CHAPTER 22 Biology, Husbandry, and Clinical Techniques Table 22-3 Approximate Hematologic Values for Guinea Pigs and Chinchillas; Values Are Expressed as Mean, Range, or Mean and Rangea
291
Table 22-4 Approximate Biochemical Values for Guinea Pigs and Chinchillas; Values Are Given as Range or Mean and Rangea
Analyte
Guinea Pig Chinchlla
Analyte
Guinea Pig Chinchilla
Hemotcrit/PCV Hemoglobin (g/dL) Red blood cells (×106/μL) Reticulocytes (% RBC) Mean corpuscular volume (MCV) (fL) Mean corpuscular hemoglobin (MCH) (pg) Mean corpuscular hemoglobin concentration (MCHC) (g/dL) White blood cells (×103/μL) Lymphocytes (%) Neutrophils, segmented (%) Monocytes (%) Eosinophils (%) Basophils (%) Platetlets (×103/μL)
39-55 11.6-16.9 4.5-6.4 — 80-89
37 (35-40) 13.1 (11.6-14.5) 3.8 (3.4-4.2) 0-2.8 101 (87-116)
0-61
40 (31-49)b
2.6-4.1 0-418 0-3159 0-90
4.5 (3.7-5.4)b 44 (37-51)b
24-27
—
29-32
35 (33-38)
Alanine aminotransferase (U/L) Albumin (g/dL) Alkaline phosphatase (U/L) Amylase (U/L) Aspartate aminotransferase (U/L) Bile acids (μmol/L) Bilirubin (mg/dL) Blood urea nitrogen (mg/dL)
0-84.5 0-0.09 9.4-28.9
2.9-14.4 28-84 12-62
5.2 (4.4-6.1) 35.0 (29.6-40.5) 40.0 (32.7-47.2)
0-9 0-14 0-2 250-850
11.5 (7.8-15.2) 5.8 (3.9-7.6) 5.3 (3.2-7.4) 276
Calcium (mg/dL) Chloride (mEq/L) Cholesterol (mg/dL) Creatine kinase (U/L) Creatinine (mg/dL) Fructosamine (μmol/L) γ-Glutamyl transferase (U/L) Globulin (g/dL) α1 globulin (g/dL) α2 globulin (g/dL) β globlulin (g/dL) γ globulin (g/dL) Glucose (mg/dL)
9.6-12.4 94-111 12-65 0-2143 0-0.87 134-271 0-13 1.7-2.6 0.10-0.36 0.79-1.48 0.25-0.68 0.17-0.78 89-287
Lactate dehydrogenase (U/L) Lipase (U/L) Potassium (mEq/L) Phosphorous (mg/dL) Protein, total (g/dL) Sodium (mEq/L) Triglycerides (mg/dL)
0-515 0-152 4.5-8.8 3.2-21.6 4.4-6.6 130-150 29-208
— 0.2 (0.1-0.3)b 57 (46-69)b 35 (29-42)c 9.5 (7.4-11.5)b 105-115 106 (78-134)b 0.8 (0.6-1.0)b — — — — — — — — 180 (163-197)b 136 (108-165)c — — 5.0-6.5 4-8 5.6 (5.3-6.0)b 130-155 173 (117-229)b
aReferences
25a, 34, 51a, 57a.
guinea pigs, often in conjunction with fatty infiltration of many tissues, including the liver. If guinea pig serum is stored in plastic, potassium levels are lower than if glass containers are used.69 A unique leukocyte of the guinea pig is the Kurloff cell (see Chapter 36). This mononuclear cell resembles a lymphocyte but contains round or ovoid inclusions termed Kurloff bodies.52 The origin of Kurloff cells (thymus or spleen) is controversial. The number of circulating cells is variable: Kurloff cells are rare in very young animals, numbers are low in males, and numbers in females are related to the estrous cycle. Kurloff cells are highest in females during pregnancy and may play a role in creating a physiologic barrier between the fetus and the mother.52 The cellular distribution of bone marrow in the guinea pig is 26.7% erythroblasts, 63.3% myeloid cells, 4.6% lymphocytes, and 5.4% reticulum cells.2 The myeloid/erythroid (M/E) ratio is approximately 1.5:125 to 1.9:1.8.15 In chinchillas, the M/E ratio is approximately 0.9-1.1:1, and the distribution is approximately 27% to 30% lymphocytes, 23% metarubricytes, 12% rubricytes, and 20% segmented and band neutrophils.51a The normal pH of guinea pig urine is 9.0.39 The normal pH of chinchilla urine is 8.5; specific gravity often exceeds 1.045.36
TREATMENT TECHNIQUES Intravenous and Intraosseous Catheters Peripheral intravenous catheters are commonly used in guinea pigs and chinchillas but sometimes can be difficult to place because of the small size and fragility of the veins, especially in chinchillas. Use a small (24- to 26-gauge) indwelling catheter and place the catheter while the animal is under anesthesia or tranquilized. Intraosseous catheters can be placed; these are commonly placed in the cranial tibia or femur (Fig. 22-10); see Chapter 2 for a description of the technique in ferrets. Monitor
249 (202-297)b
aReferences
23, 25a, 34, 51a. from normal males (n = 16), ages 8-12 months, samples obtained from cardiac puncture immediately after death by cervical rupture. cValues from normal males (n = 8) anesthetized with ketamine and acepromazine. bValues
animals with indwelling catheters closely in case they attempt to chew at the catheter or intravenous line. Placing neck collars on these animals to prevent chewing usually causes stress and animals become inappetant; avoid placing these, but if necessary, use a soft collar.
Fluid Therapy Normal daily water intake in the guinea pig is estimated to be approximately 100 mL/kg.34 Supplemental fluid requirements should be calculated based on this estimation plus additional fluid to compensate for dehydration or fluid loss. Supplemental fluids are commonly given subcutaneously into the loose skin of the dorsal neck and upper back areas. The total volume can be divided into two to three daily treatments. Volumes of 25 to 35 mL can be given into each subcutaneous injection site with a 22- to 25-gauge butterfly catheter. Many guinea pigs react to the pain caused by subcutaneous
292
SECTION III Guinea Pigs and Chinchillas Guinea pigs especially do not adapt well to changes in their environments or routines and should be hospitalized only if necessary. Hospitalized guinea pigs should be given supportive care in anticipation of decreased appetite and water consumption.
References
Fig. 22-10 An intraosseous catheter placed in the cranial tibia of a guinea pig. A 20-gauge, 1.5-inch spinal needle can be used as a catheter.
fluid administration and become very stressed. To avoid unnecessary stress, animals that are drinking water can be given oral fluids unless they are azotemic or moderately to severely dehydrated. Use an injection pump or a Buretrol for either continuous or intermittent infusion of intravenous fluids. Monitor fluid volumes closely to avoid overhydration.
Antibiotic Therapy Because they are hindgut fermenters that depend almost entirely on the bacterial production of volatile fatty acids for energy, guinea pigs and chinchillas are very susceptible to changes in enteric microbial flora.57 Although all antibiotics have the potential to affect normal gut microflora, classes of drugs such as the fluoroquinolones, tetracyclines, chloramphenicol, and aminioglycosides have less impact than drugs such as the penicillins, cephalosporins, lincosamides, and older macrolides. Therefore choose antibiotic therapy cautiously, especially oral antibiotics, to lessen the risk of enteric dysbiosis and antibioticassociated enterotoxemia (see Chapter 23).
Administration of Medications Give parenteral medications by subcutaneous or intramuscular injection. The upper back is a common site for subcutaneous injection. The skin in this area is thick in guinea pigs, especially in intact males, and is sometimes difficult to penetrate with a 25-gauge or smaller needle. Give intramuscular injections in the lumbar muscles. Give oral medications and nutritional supplements by squeezing them from a syringe into the side of the mouth. Oral medications in tablet form must be compounded into liquid formulations so that they can be easily administered by the owners. Administration of tablets to chinchillas is sometimes possible; because chinchillas are inquisitive, they will usually eat tablets that are hidden in raisins. Force-feed guinea pigs and chinchillas that are anorectic or are consuming less food with supplements such as Critical Care (Oxbow Hay Co.) or softened guinea pig pellets. Give hospitalized guinea pigs parenteral vitamin C (25-50 mg) daily as needed.
1. Animal Research, Institutional Animal Care and Use Committee. University of Iowa website. Blood collection and administration of fluids and drugs (guinea pig). Available at http:// research.uiowa.edu/animal/?get=g_pig_t#Cranial vena cava. Accessed Feb 21, 2011. 2. Baranski S. Effect of chronic microwave irradiation on the blood forming system of guinea pigs and rabbits. Aerosp Med. 1971;42:1196-1199. 3. Barber N, Thompson RL. Sandbathing reduces fur lipids of chinchillas, Chinchilla laniger. Anim Behav. 1990;39: 403-405. 4. Bekyurek T, Liman N, Bayram G. Diagnosis of sexual cycle by means of vaginal smear method in the chinchilla (Chinchilla lanigera). Lab Anim. 2002;36:51-60. 5. Berryman JC. Guinea-pig vocalizations: their structure, causation and function. Z Tierpsychol. 1976;41:80-106. 6. Breazile JE, Brown EM. Anatomy. In: Wagner JE, Manning PJ, eds. The biology of the guinea pig. New York: Academic Press; 1976:53-62. 7. Björnhag G, Snipes RL. Colonic separation mechanism in lagomorph and rodent species – a comparison. Zoosystematics Evolution. 1999;75:275-281. 8. Cheeke PR. Rabbit feeding and nutrition. Orlando: Academic Press; 1987. 9. Crossley DA. Dental disease in chinchillas. Thesis. School of Dentistry. Manchester: University of Manchester; 2003;263. 10. Crossley DA, Miguelez MM. Skull size and cheek-tooth length in wild-caught and captive-bred chinchillas. Arch Oral Biol. 2001;46:919-928. 11. Crossley DA, Roxburg G, Miguelez Vidales MM. Anatomy of the chinchilla (Chinchilla lanigera) lacrimal drainage system and its obstruction in dental disease. Proceedings. 8th Europ Vet Dental Society. 1999:21-22. 12. de Casto TF, Dummer RJ, Rickes EM, et al. Morphological, morphometric and topographical description of the digestive tract in Chinchilla lanigera. Brazilian J Vet Res Animal Sci. 2010;47:86-94. 13. Detwiler SR. The eye of the chinchilla. J Morphol. 1949;84: 123-144. 14. De Vries L. Peru pushes guinea pigs as food in hopes of developing more robust export business. CBS News World. Available at http://www.cbsnews.com/stories/2004/10/19/world/ main650148.shtml. Accessed Feb 20, 2011. 15. Dineen JK, Adams DB. The effect of long-term lymphatic drainage on the lympho-myeloid system in the guinea pig. Immunology. 1970;19:11-30. 16. Donnelly TM. Behaviour and reproduction. Rabbits and rodents: laboratory animal science. Proceedings 142. Sydney: Post-Graduate Committee in Veterinary Science. 1990:81–388. 17. Ebino KY. Studies on coprophagy in experimental animals. Jikken Dobutsu. 1993;42:1-9. 18. Farmer FA. A study of the dietary requirements of chinchillas. Natl Chinchilla Breeder Can. 1951;5:11-18. 19. Fiedler LA. Rodents as a food source. Proceedings. 14th Vert Pest Conf. 1990. Available at http://digitalcommons.unl.edu/ vpc14/30/. Accessed Feb 20, 2011. 20. Flower SS. Contributions to our knowledge of the duration of life in vertebrate animals. V. Mammals. Proceedings. Zool Soc London. 1931;101:145-234.
CHAPTER 22 Biology, Husbandry, and Clinical Techniques 21. Franz R, Kreuzer M, Hummel J, et al. Intake, selection, digesta retention, digestion and gut fill of two coprophageous species, rabbits (Oryctolagus cuniculus) and guinea pigs (Cavia procellus), on a hay-only diet. J Anim Physiol Anim Nutr (Berl). 2010 Nov 22. Epub ahead of print. 22. Grau J. La chinchilla: su crianza en todos los climas [Spanish]. The chinchilla: its breeding in all the climates. 3rd ed. Buenos Aires: El Ateneo; 1986. 23. Harkness JE, Turner PV, VandeWoude S, et al. Harkness and Wagner‘s biology and medicine of rabbits and rodents. 5th ed. Hoboken: Wiley-Blackwell; 2010. 24. Harper LV. Behavior. In: Wagner JE, Manning PJ, eds. The biology of the guinea pig. New York: Academic Press; 1976:31-48. 25. Harris RS, Herdan G, Ancill RJ, et al. A quantitative comparison of the nucleated cells in the right and left humeral bone marrow of the guinea pig. Blood. 1954;9:374-378. 25a. Hein J, Harmann K. Reference ranges for laboratory parameters in guinea pigs. Tierarztliche Praxis Ausgabe Kleintiere. 2003;31:383-389. 26. Holtenius K, Bjornhag G. The colonic separation mechanism in the guinea-pig (Cavia porcellus) and the chinchilla (Chinchilla laniger). Comp Biochem Physiol A Physiol. 1985;82:537-542. 27. Huet F, Fargeas MJ, Fioramonti J, et al. Fecal excretion and digestive transit in the chinchilla: physiological values, effect of dietary fiber intake and changes in diet. [French] Excretion fecale et temps de transit digestif chez le chinchilla: valeurs physiologiques et effets de la teneur en fibres et des transitions de regime alimentaire. Rev Med Vet (Toulouse) 1998;149:739-744. 28. Jenkins JR. Husbandry and common diseases of the chinchilla (Chinchilla laniger). J Small Exot Anim Med. 1992;2:15-17. 29. Jezewska G, Tarkowski J, Niezgoda GA, et al. Results of crossbreeding different colour varieties in chinchillas. [Polish] Wyniki krzyzowania roznych odmian barwnych u szynszyli. Ann Univ Mariae Curie Sklodowska Sect EE Zootech. 1997;15:197-201. 30. Jilge B. The gastrointestinal transit time in the guinea-pig. Z Versuchstierk. 1980;22:204-210. 31. Jiménez JE. Bases biológicas para la conservación y manejo de la Chinchilla Chilena silvestre: Proyecto de conservación de la Chinchilla Chilena (Chinchilla lanigera). Final report. Santiago, Chile: Corporación Nacional Forestal-World Wildlife Fund (CONAF; the Chilean Forest Service); 1990. 32. Jordan WJ. Sterility in Chinchilla lanigera. [German] Unfruchtbarkeit beim Chinchilla lanigera. Kleintierpraxis. 1965;10:243-244. 33. Kleiman DG. Patterns of behaviour in hystricomorph rodents. Symp Zool Soc London. 1974;34:171-209. 34. Manning PJ, Wagner JE, Harkness JE. Biology and diseases of guinea pigs. In: Fox JG, Cohen BJ, Loew FM, eds. Laboratory animal medicine. Orlando: Academic Press; 1984:149-177. 35. Merchant HA, McConnell EL, Liu F, et al. Assessment of gastrointestinal pH, fluid and lymphoid tissue in the guinea pig, rabbit and pig, and implications for their use in drug development. Eur J Pharm Sci. 2011;42:3-10. 36. Merry CJ. An introduction to chinchillas. Vet Tech. 1990;11: 315-322. 37. Mösslacher E. Breeding and caring for chinchillas. Neptune City: T.F.H. Publications; 1986. 38. Mitruka BM, Rawnsley HM. Clinical biochemical and hematological reference values in normal experimental animals and normal humans. 2nd ed. Chicago: Year Book Medical Publishers; 1981. 39. Navia JM, Hunt CE. Nutrition, nutritional diseases, and nutrition research applications. In: Wagner JE, Manning PJ, eds. The biology of the guinea pig. New York: Academic Press; 1976:235-261.
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40. Neira R, García X, Scheu R. Descriptive analysis of the reproductive behavior and of growth of chinchillas (Chinchilla laniger Gray) in confinement. [Spanish] Análisis descriptivo del comportamiento reproductivo y de crecimiento de chinchillas (Chinchilla laniger Gray) en confinamiento. Avances en Producción Animal (Chile). 1989;14:109-119. 41. Neville RWJ, Weir BJ, Lazarus NR. Hystricomorph insulins. Symp Zool Soc London. 1974;34:417-433. 42. Opazo JC, Soto-Gamboa M, Bozinovic F. Blood glucose concentration in caviomorph rodents. Comp Biochem Physiol A Mol Integr Physiol. 2004;137:57-64. 43. Osgood WH. The technical name of the chinchilla. J Mammal. 1941;22:407-411. 44. Peichao W, Hogee L, Helin S, et al. Influence of environment temperature on oxygen consumption and heat production of adult chinchilla (Chinchilla lanigera). Scientifur. 1980;4:16-17. 45. Peiffer RL, Johnson PT. Clinical ocular findings in a colony of chinchillas (Chinchilla laniger). Lab Anim. 1980;14:331-335. 46. Popesko P, Rajtová V, Horák J. A colour atlas of anatomy in small laboratory animals. Vol. 1. Rabbit, guinea pig. London: Wolfe Publishing; 1992;48–240. 47. Reuter RE. Venipuncture in the guinea pig. Lab Anim Sci. 1987;37:245-246. 48. Riggs S. Guinea pigs. In: Mitchell MA, Tully TN, eds. Manual of exotic pet practice. St. Louis: Saunders an imprint of Elsevier; 2009:456-473. 49. Ritchey L, Cogswell EC, Beemam R. The joy of chinchillas. 6th ed. Menlo Park: Privately printed; 2004. 50. Roberts CM, Perry JS. Hystricomorph embryology. Symp Zool Soc London. 1974;34:333-360. 51. Schneider M, Bartl J, Erhard M. Vocalizations of chinchillas living in sociable groups. Proceedings. Conference of the ACVBAVSAB (American College of Veterinary Behaviorists-American Veterinary Society of Animal Behavior). 2006. 51a. Silva TdO, Kreutz LC, Barcellos LJG, et al. Reference values for chinchilla (Chinchilla laniger) blood cells and serum biochemical parameters. Ciência Rural. 2005;35:602-606. 52. Sisk DB. Physiology. In: Wagner JE, Manning PJ, eds. The biology of the guinea pig. New York: Academic Press; 1976: 63-92. 53. Smith D. Guinea pigs as food source. The Hindu. Feb 18, 2010. Available at http://www.hindu.com/2010/02/18/stories/ 2010021862021600.htm. Accessed Feb 20, 2011. 54. Spotorno AE, Valladares JP, Marin JC, et al. Molecular divergence and phylogenetic relationships of chinchillids (Rodentia: Chinchillidae). J Mammal. 2004;85:384-388. 55. Spotorno AE, Zuleta CA, Valladares JP, et al. Chinchilla laniger. Mammal Species. 2004;758:1-9. 56. Stern JJ, Merari A. The bathing behavior of the chinchilla: effects of deprivation. Psychon Sci. 1969;14:115-116. 57. Stevens CE, Hume ID. Contributions of microbes in vertebrate gastrointestinal tract to production and conservation of nutrients. Physiol Rev.1998;78:393-427. 57a. Strike TA. Hemogram and bone marrow differential of the chinchilla. Lab Anim Care. 1970;20:33-38. 58. Timm KI, Jahn SE, Sedgwick CJ. The palatal ostium of the guinea pig. Lab Anim Sci. 1987;37:801-802. 59. Townsend KEB, Croft DA. Enamel microwear in caviomorph rodents. J Mammal. 2008;89:730-743. 60. Voss R. Male accessory glands and the evolution of copulatory plugs in rodents. Occasion Papers Museum Zool. No. 689. Ann Arbor: University of Michigan; 1979. 61. Walker EP. Mammals of the world. 3rd ed. Vol. II. Baltimore: Johns Hopkins University Press; 1975. 62. Webb RA. Chinchillas. In: Beynon PH, Cooper JE, eds. Manual of exotic pets. Cheltenham: British Small Animal Veterinary Association; 1991:15-22.
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63. Weir BJ. Chinchilla. In: Hafez ESE, ed. Reproduction and breeding techniques for laboratory animals. 1st ed. Philadelphia: Lea & Febiger; 1970:209-323. 64. Weir BJ. The managment and breeding of some more hystricomorph rodents. Lab Anim. 1970;4:83-97. 65. Weir BJ. Reproductive characteristics of hystricomorph rodents. In: Rowlands IW, Weir BJ, eds. The biology of hystricomorph rodents. London: Zoological Society of London-Academic Press; 1974:265-301. 66. Weir BJ. Notes on the origin of the domestic guinea-pig. Symp Zool Soc Lond. 1974;34:437-446. 67. Weiss S. Ethological studies of chinchillas in climbing cages. [German] Verhaltensuntersuchungen an Chinchillas in ausgestalteten Kletterkäfigen. Institut für Tierschutz, Verhaltenskunde und Tierhygiene, Faculty of Veterinary Medicine. Munich: Ludwig-Maximilians-Universität München; 2005;143.
68. Weisser F, Lacy FB, Weber H, et al. Renal function in the chinchilla. Am J Physiol. 1970;219:1706-1713. 69. White EJ, Lang CM. The guinea pig. In: Loeb WF, Quimby FW, eds. The clinical chemistry of laboratory animals. New York: Pergamon Press; 1989:27-30. 70. Wilcox HH. Histology of the skin and hair of the adult chinchilla. Anat Rec. 1950;108:385-397. 71. Wilson DE, Reeder DM. Mammal species of the world: A taxonomic and geographic reference. 3rd ed. Baltimore: Johns Hopkins University Press; 2005. 72. Williams CSF. Practical guide to laboratory animals. St. Louis: C.V. Mosby; 1976.
CHAPTER
23
Disease Problems of Guinea Pigs
Michelle G. Hawkins, VMD, Diplomate ABVP (Avian), and Cynthia R. Bishop, DVM
Gastrointestinal and Hepatic Diseases Dental Disease Gastrointestinal Hypomotility or Stasis Dysbiosis and Antibiotic-Associated Enterotoxemia Enteritis and Diarrhea Fecal Impaction Hepatic Lipidosis Neoplasia Respiratory Diseases Bacterial and Viral Pneumonia Noninfectious Respiratory Diseases Urinary Diseases Urolithiasis Cystitis and Urinary Tract Infections Chronic Interstitial Nephritis and Chronic Renal Failure Other Uropathies Reproductive Diseases Ovarian Cysts Uterine Prolapse Dystocia Toxemia of Pregnancy Mastitis Pyometra and Metritis Vaginitis and “Scrotal Plugs” Orchitis and Epididymitis Neoplasia Dermatologic Diseases Dermatophytosis Ectoparasites Cervical Lymphadenitis Pododermatitis Alopecia Neoplasia Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
Musculoskeletal Diseases Vitamin C Deficiency (Scurvy) Osteoarthritis and Osteoarthrosis Fibrous Osteodystrophy Metastatic Mineralization Iatrogenic Muscle Necrosis Nutritional Muscular Dystrophy Neurologic Diseases Otitis Media/Otitis Interna Mites Rabies Lymphocytic Choriomeningitis Virus Ophthalmologic Diseases Corneal Ulceration “Pea Eye” Conjunctivitis Miscellaneous Diseases Ototoxicity Diabetes Mellitus Heat Stress Lymphosarcoma/Cavian Leukemia
GASTROINTESTINAL AND HEPATIC DISEASES DENTAL DISEASE Dental disease is one of the most common veterinary presentations of guinea pigs. The anatomy of the guinea pig’s dentition, common clinical presentations of guinea pigs with dental disease, diagnosis, and dental treatment are discussed in detail in Chapter 32. Diets deficient in fiber or vitamin C, infection, and trauma are common reasons for malocclusion in guinea pigs; genetic predisposition while not proven, is also strongly suspected. A thorough oral examination should be part of the routine physical, as disease of the cheek teeth is easily overlooked. 295
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A
B
Fig. 23-1 Right lateral (A) and ventrodorsal (B) radiographs of a guinea pig with gastrointestinal stasis. Gas is normally found within the guinea pig’s GI tract and, in most cases of GI stasis, remains within normal limits. But GI stasis can also result in gas accumulation that may become life-threatening. Because guinea pigs with dental disease often have concurrent disease processes, a thorough systemic evaluation is indicated before dental treatment is initiated. Perioperative supportive care is just as critical to a good outcome as the dental treatment itself; pain, hydration, nutrition (including vitamin C supplementation), and secondary infection must be thoroughly considered. Prevention is aimed at ensuring an appropriate high-fiber diet and vitamin C support.
S
C
GASTROINTESTINAL HYPOMOTILITY OR STASIS Gastrointestinal (GI) hypomotility or stasis occurs primarily or as a sequel to virtually any other disease process in guinea pigs. Inadequate dietary fiber is a significant predisposing factor, but any disease process causing pain or anorexia can lead to GI hypomotility, dehydration of GI contents, and GI stasis. Gastric or intestinal torsion or volvulus, intestinal intussusception, or ingested foreign bodies can result in partial or complete gastrointestinal obstruction and signs of GI stasis, but these are infrequently reported. Malabsorption occurs as GI contents become dehydrated; hypovitaminosis C can occur if the syndrome is protracted. A thorough history is invaluable in determining the inciting cause and should include complete dietary and husbandry information, duration of clinical signs (specifically anorexia), health status, and any recent antibiotic use. Dental disease is very commonly identified in guinea pigs presenting with signs of GI stasis. Hepatic lipidosis can occur secondary to anorexia, especially in obese guinea pigs. Clinical signs of GI stasis include decreased/absent fecal material, anorexia, bruxism, gas or fluid-distended stomach, cecum, and bowel loops, pain on abdominal palpation, decreased gastrointestinal sounds, and respiratory or cardiovascular compromise. The GI contents become dehydrated during stasis, exacerbating GI pain and anorexia and possibly leading
Fig. 23-2 Right lateral radiograph of a guinea pig with gastric dilatation/volvulus. Identification of the normal anatomic locations of the stomach and cecum are critical to diagnosis. S, Stomach; C, cecum.
to partial or complete GI obstruction. GI stasis can also result in the accumulation of gas within the intestinal tract (“bloat”), which can become life threatening (Fig. 23-1). Gastric dilatation-volvulus (GDV) has been reported in only two pet guinea pigs to date.11,33 A complete physical examination and appropriate diagnostics to evaluate for underlying disease are performed. Abdominal radiographs and ultrasound evaluate GI anatomy and motility. The clinical signs and physical exam findings of gastric/cecal dilatation/torsion are similar to those of GI stasis; imaging is required to distinguish the two conditions (Fig. 23-2). Blood work should be performed to evaluate for evidence of inflammation, infection, and organ dysfunction. Regardless of the cause or severity, medical management of GI stasis consists of aggressive supportive care with replacement
CHAPTER 23 Disease Problems of Guinea Pigs fluid therapy, pain management, and assisted nutrition. Hydrate the GI tract first to facilitate its motility and function during nutritional therapy. Currently several timothy hay-based critical care feeding formulas for small herbivores are commercially available. Supplement with vitamin C. Most guinea pigs tolerate syringe feeding very well. Some guinea pigs will not accept syringe feeding with a 60-mL catheter-tip syringe; patience is often required to feed small boluses of food with a 1- or 3-mL syringe. Nasogastric tube placement is difficult in guinea pigs; although finely ground critical care formulas that will pass through these small tubes are commercially available, it is difficult to provide the appropriate fiber particle size necessary to stimulate GI motility with these products. Pain management is an essential component of therapy. GI pain is visceral and often too severe to respond to nonsteroidal anti-inflammatory drugs (NSAIDs) alone. The risks and benefits of using opioids are considered for each case, but guinea pigs with GI stasis usually respond well to buprenorphine or butorphanol. Prokinetics (metoclopramide, cisapride) should not be used initially in these cases, since partial or complete obstruction is common prior to hydration of GI contents. The use of commercially available probiotics or transfaunation has been advocated with prolonged stasis and when dysbiosis is suspected. Gastric decompression is performed on an emergency basis in cases of gastric tympany and is accomplished by passing a large-bore red rubber tube into the stomach through the oral cavity, being careful to not obstruct the glottis; however, gas lower in the GI system will not be affected. Gastric decompression can carry a high risk in the already debilitated patient, and often the volume of gas yielded is disappointing. Trocarization carries significant risk of gastric or cecal rupture or peritonitis and should be used only as a last resort. Surface tension relaxants such as simethicone anecdotally decrease the volume of accumulated gastric gas. Antibiotics should not be administered empirically in guinea pigs with GI stasis because of the guinea pig’s GI tract is highly sensitive to some antibiotics. Bacterial culture and sensitivity results or suspicions that the GI disease is caused by specific bacteria should be carefully weighed in deciding on antibiotic use. Surgery is rarely required for GI hypomotility/stasis, but if GDV is highly suspected, this is a surgical emergency. No information is available regarding prognosis and surgical management in guinea pigs with GDV, as all published reports have been based on diagnosis at necropsy.11,33
DYSBIOSIS AND ANTIBIOTIC-ASSOCIATED ENTEROTOXEMIA Guinea pigs possess a predominantly gram-positive GI flora and are exquisitely sensitive to some antibiotics, which eradicate that flora. Penicillin, ampicillin, chlortetracycline, clindamycin, erythromycin, and lincomycin can destroy the most susceptible gram-positive organisms, permitting overgrowth of gram- negative bacteria and Clostridium difficile, with elaboration of its toxin.10,30 Toxin production causes hyperactivity of secretomotor neurons, resulting in secretory diarrhea and hemorrhagic typhlitis.55 This occurs most commonly when these antibiotics are given orally; however, it can also occur in some animals when given parenterally as well as with appropriate but prolonged antibiotic therapy. Non-antibiotic-induced dysbiosis also occurs and is generally associated with abrupt dietary changes, ingestion of contaminated foods, and stress. Anorexia, dehydration, and hypothermia are the most common clinical signs; diarrhea
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may or may not be present.30 Diagnosis is usually based on history, clinical signs, and histopathologic lesions. C. difficile is difficult to isolate, but polymerase chain reaction (PCR) and enzyme immunoassay for its toxin are commercially available. Antibiotic-associated enterotoxemia is treated symptomatically. Provide thermal support and administer intravenous or subcutaneous crystalloid fluids to restore hydration. Commercially available probiotics (Lactobacillus species) live-culture yogurt, or cecotrophs transfaunation from other guinea pigs can be used to try to reestablish normal microflora. Chloramphenicol (50 mg/kg PO q8h) may be somewhat effective in suppressing further clostridial overgrowth.10 Antibiotic-associated enterotoxemia is prevented by treating diseases in guinea pigs with appropriate antibiotics. Trimethoprim-sulfa, chloramphenicol, and enrofloxacin are generally safe in guinea pigs.
ENTERITIS AND DIARRHEA Diarrhea is uncommon in adult cavies, but soft stools are frequently reported due to excess carbohydrates or inadequate dietary fiber and even sometimes with vegetable consumption. Some animals have intermittent soft stools without identification of etiology. Bacterial enteritis is also a common cause of soft stools/ diarrhea. Tyzzer’s disease, caused by Clostridium piliforme, is transmitted by the fecal-oral route. Young, stressed, or immunocompromised animals are particularly affected. Clinical signs include lethargy/anorexia, diarrhea, unthrifty appearance, and acute death.30 Lesions observed at necropsy include intestinal inflammation and focal hepatic necrosis. C. piliforme is an intracellular bacterium and does not grow on routine culture media. Definitive diagnosis is by histopathologic identification of organisms from liver or intestinal sections. Treatment is generally unrewarding. The disease can be prevented by good husbandry practices and stress reduction, particularly at weaning. Salmonella typhimurium and Salmonella enteritidis are less frequently reported causes of bacterial enteritis in pet guinea pigs, but mortality can be high during an outbreak. Transmission is usually due to fecal contamination of feed or water. Weanlings, pregnant sows, aged animals, and those with nutritional deficiencies are particularly susceptible. Signs include scruffy hair coat, weight loss, weakness, conjunctivitis, and abortion. Diarrhea may or may not be present. At necropsy, the spleen and liver are often enlarged, and yellow necrotic foci may be present in the viscera.41 Diagnosis is by fecal or intestinal culture and sensitivity testing. Treatment is usually not recommended, as affected animals can become asymptomatic carriers and salmonellosis can be zoonotic. Prevention is aimed at proper disinfection/sanitization of the environment, storing of food in airtight containers, and thorough washing of all fresh fruits and vegetables offered. Other causes of bacterial diarrhea include Yersinia pseudotuberculosis, Clostridium perfringens, Escherichia coli, Pseudomonas aeruginosa, Citrobacter freundii, and Listeria monocytogenes.41 Like Salmonella species, these are usually contracted through food contamination. Y. pseudotuberculosis can cause abscesses of the intestine, liver, and regional lymph nodes.41 E. coli causes wasting, depression, and death in weanlings. Intestines may contain yellow fluid. Guinea pigs are hosts to Eimeria caviae, Balantidium caviae, and Paraspidodera uncinata. In juvenile animals, E. caviae causes diarrhea, but this is much less common in adults.
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HEPATIC LIPIDOSIS Hepatic lipidosis occurs quickly secondary to anorexia, especially in obese guinea pigs. Significant hepatic changes can occur in as little as 48 hours. Anorexia of as little as 12 hours should be considered an emergency and nutritional support initiated immediately. The prognosis is guarded once this condition develops.
NEOPLASIA Tumors of the intestinal tract and liver are not common, but lymphosarcoma and gastric/cecal adenocarcinomas have been identified in pet guinea pigs in the authors’ practices.
Fig. 23-3 Fecal impaction of unknown etiology is commonly seen in older guinea pigs, especially boars. The guinea pig is usually presented for straining to defecate, constipation, or passing large amounts of foul-smelling soft stool. Mild bilateral pododermatitis is also present.
Cryptosporidium wrairi causes failure to gain weight, weight loss, diarrhea, and death, especially in weanlings and immunosuppressed animals.9 Transmission is via the fecal-oral route. Immunocompetent animals recover within 4 weeks and are resistant to reinfection. Organisms can be identified on fecal examination or by histopathology of the brush border of mucosal epithelial cells. No treatment has proved effective. Oocysts of C. wrairi can be destroyed in the environment with a 5% ammonia solution, freezing to below 32°F (0°C), or heating to above 149°F (65°C). Cryptosporidiosis is a potentially zoonotic disease. Regardless of the underlying etiology, diarrhea is a serious problem, because hypoglycemia, dehydration, hypothermia and electrolyte imbalances quickly occur. Administer warm fluids (crystalloids ± dextrose; colloids, if hypoalbuminemia or decreased oncotic pressure) and supplemental thermal support. Dietary correction with adequate fiber is of utmost importance. Antibiotic treatment is based on culture and sensitivity results. With the recent E. coli outbreaks associated with green vegetables in the United States, it is extraordinarily important to thoroughly wash all vegetables and fruits prior to offering them.
FECAL IMPACTION Fecal impaction is most commonly identified in older guinea pigs, especially boars, but the etiology is unknown (Fig. 23-3). Soiled bedding combined with inguinal sebaceous secretions can become adhered to the scrotum and anus, potentially resulting in inguinal gland infections and fecal impaction. The guinea pig is usually presented for straining to defecate, constipation, or passing large amounts of foul-smelling soft stool. Affected animals may appear to be uncomfortable when straining to push out the accumulated feces. Often the only physical exam abnormalities detected will be an enlarged rectum impacted with normal, soft feces and a flaccid anus. Suggested therapy has included a diet change to increase fiber, warm-water enemas, and manual evacuation of stools using a cotton-tipped applicator (often long-term therapy is required).
RESPIRATORY DISEASES Guinea pigs have a relatively small thoracic cavity compared with body mass. They seem particularly sensitive to airborne pollutants and respiratory infections; thus husbandry issues such as proper airflow, sanitation, and choice of bedding are of utmost importance. Insufficient airflow in combination with ammonia buildup from soiled cages can predispose to respiratory disease. Bedding made of recycled paper products is recommended, rather than wood chips; cedar chips are often associated with respiratory disease.
BACTERIAL AND VIRAL PNEUMONIA Bacterial pneumonia is one of the most important diseases of guinea pigs. Poor husbandry conditions mentioned above, along with humid environments, predispose to pneumonia.38 The most common etiologic agents are Bordetella bronchiseptica, Streptococcus pneumoniae, Streptobacillus moniliformis, and Haemophilus species.14,38 Regardless of etiology, the general progression of clinical signs is as follows: ocular/nasal discharge, dyspnea, increased respiratory sounds, sneezing, coughing, depression, anorexia, weight loss, and unthrifty appearance of the coat.14,38 Diagnosis includes clinical signs, physical exam findings, thoracic radiographs, and culture and sensitivity of respiratory secretions. Bacterial culture of the nasal and/or ocular discharge is often inconsistent with tracheal or bronchial cultures (causative organisms). Diagnostic samples are best retrieved from tracheal secretions, bronchoalveolar lavage, or fresh necropsy specimens from the bronchopulmonary region in colony outbreaks.21,38 Opacity within the tympanic bullae or lungs may be seen on radiographs in guinea pigs with respiratory disease.38 Thoracic auscultation may reveal crackles, rales, and wheezing or minimal air movement, depending on the severity of the disease. Systemic antibiotic therapy based on culture and sensitivity results and supportive care (fluid and nutritional support) should be instituted. Nebulized antibiotics can be considered for upper respiratory therapy. Supplemental oxygen, saline nebulization, bronchodilators, and decongestants should be considered as needed. Vitamin C supplementation is always a prudent addition to infectious disease treatment. B. bronchiseptica is a gram-negative rod that is an important cause of respiratory disease. Rabbits, dogs, and nonhuman primates may be asymptomatic carriers. The disease is characterized by purulent bronchopneumonia often involving consolidated lung lobes and fibrinosuppurative pleuritis
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NONINFECTIOUS RESPIRATORY DISEASES Noninfectious causes of respiratory disease may include foreign body inhalation, choke, bloat, molar malocclusion, thoracic trauma, and electrocution. Pulmonary adenoma is common and sometimes multicentric, with firm, variably sized white nodules that may be apparent on radiographs. This should not be confused with metastatic disease; clinical signs are usually not associated with it.
URINARY DISEASES UROLITHIASIS
Fig. 23-4 Bronchopneumonia and fibrinosuppurative pleuritis (arrows) in a guinea pig with Bordetella bronchiseptica. Guinea pigs are very sensitive to disease with this organism. Rabbits and dogs can be asymptomatic carriers, so contact with these animals should be discouraged to minimize the potential for transmission to cavies.
(Fig. 23-4). B. bronchiseptica infection may also cause exudates in the tympanic bullae, metritis, and abortions. Treatment is with antibiotics based on culture and sensitivity results. Prevention with an injectable B. bronchiseptica vaccine (Bronchicine, Bicor Animal Health, Omaha, NE; 0.2 mL SC with a booster 2-3 weeks later) has been used with moderate success in the face of an outbreak (C. Bishop, personal observation). It is recommended to prevent exposure of guinea pigs to rabbits and dogs that might be asymptomatic carriers of Bordetella.38 S. pneumoniae is a gram-positive coccus that may be transmitted by asymptomatic carriers of many species, including guinea pigs. Serotypes III, IV, and XIX cause disease in guinea pigs.18 Disease is predisposed by poor husbandry and is most common in young cavies. Bronchopneumonia, fibrinopurulent pleuritis, and pericarditis are common manifestations of this infection. One of the present authors (CB) has experienced an outbreak of S. pneumoniae in a caviary of 90 to 100 animals in which symptoms were not limited to the respiratory system. Metritis, encephalitis, and pericarditis were also identified in necropsy specimens. Animals affected by this outbreak had intermittent respiratory signs, infertility, abortion, death of young and sows after parturition, as well as neurologic signs including blindness, ataxia, and paralysis. Treatment with various antibiotics (based on culture and sensitivity results) led to resolution in only some individuals. Some animals initially improved from respiratory signs but succumbed to neurologic disease a few weeks later. Outbreaks of adenoviral pneumonia have been reported in laboratory colonies,14 but the incidence in pet or exhibition animals is unknown. Adenoviral pneumonia causes a necrotizing bronchopneumonia with an incubation period of 5 to 10 days.38 Although morbidity is low, mortality is high and cavies may die acutely.
Urolithiasis is a common problem in guinea pigs. Historically middle-aged to older (>2.5 years old) female guinea pigs were overrepresented in the literature,13,40 but a recent study of 75 guinea pigs with calculi found an equal distribution of males and females >2 years.20 The etiopathogenesis is unclear, although the alkaline pH and high mineral content of normal guinea pig urine may favor crystal formation and precipitation. Of the few published reports of this disease in guinea pigs, the majority have been single case reports.15,48,50,52 Historical information suggested that calcium oxalate calculi were most common,37,48 but recent data have more commonly identified calcium carbonate calculi.13,15,20,50 Urinary tract infections involving E. coli, Streptococcus pyogenes, and Staphylococcus species were suggested to be associated with the presence of urinary calculi in laboratory guinea pigs.37,40 Pure cultures of Corynebacterium renale and Facklamia species and mixed bacterial cultures including C. renale, the Streptococcus bovis/equinus group, and Staphylococcus species were found from urine of pet guinea pigs with urolithiasis, although a direct association was not confirmed.13,20 Also, Streptococcus viridans, Proteus mirablis, and mixed growth with S. viridans, Staphylococcus species, E. coli, or Enterococcus species were reported from cultures from some of the calculi.20 The majority of calculi are located in the bladder or urethra, but they are also found in the kidneys, ureters, or vagina or sometimes in the seminal vesicles in males. Stones located in the ureter have been primarily identified from male guinea pigs.20 Affected guinea pigs were more likely to be fed a diet high in overall percent pellets, low in percent hay, and a lesser variety of vegetables and fruits.19 Alfalfa-based pellets and hay contain higher concentrations of calcium; it has been suggested this may also contribute to urinary calculi in guinea pigs. Clinical signs are commonly associated with the size and location of the calculi. Bladder or urethral calculi are often associated with micturition abnormalities such as hematuria, stranguria, or dysuria and vague clinical signs such as lethargy and anorexia. If the calculus is located higher in the urinary tract, micturition abnormalities may still be present but lethargy, anorexia, weight loss, and a hunched posture may be the only clinical signs. Diagnosis is based upon clinical signs, physical examination findings, imaging studies including radiographs, ultrasonography, excretory intravenous pyelograms (IVPs), computed tomography (CT), and urinalysis. Urinary calculi in guinea pigs are generally radio-opaque, allowing for ease of identification using survey radiography (Fig. 23-5). However, if multiple calculi or significant GI gas is present, the anatomic locations of the calculi using survey radiography alone may be obscured. Ultrasound is useful for anatomic location of the calculi and for
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A
B
Fig. 23-5 Calcium-based urinary calculi are generally radio-opaque, allowing for visualization of the calculi in most cases. Two orthogonal views (A, B) are necessary to help determine the anatomic location of the calculi within the urinary tract (arrows).
evaluating anatomic changes in the kidneys or ureters, such as hydronephrosis or hydroureter, ureteral mucosal inflammation, or perforation. Excretory IVPs are useful to further elucidate relative functional abnormalities in the kidneys or ureters. In some institutions, CT has virtually replaced excretory IVP, as images can be obtained more rapidly using significantly less contrast material. However, even though the time to image collection is longer, the excretory IVP can be performed at a reduced cost in any veterinary practice. In suspected cases of urolithiasis, perform a urinalysis, including both a standard dipstick and microscopic examination of the sediment. Urinalyses data from 44 guinea pigs with urinary calculi found that the mean ± SD urine specific gravity was 1.015 ± 0.008 (range 1.004-1.046) and urine pH was 8.4 ± 0.5 (range 6.5- > 9). Hematuria was the most commonly reported abnormality on urine sediment, followed by mucus and lipid droplets.20 Calcium carbonate, calcium oxalate, and struvite crystals are all commonly seen on sediment examinations but, as in other small animals, crystal type(s) may not predict the mineralogy of the calculi.20 Prior to antibiotic use, culture the urine or bladder wall if high numbers of red or white blood cells, bacteria, or a combination of these are present on examination of the sediment. Collect urine via cystocentesis, as free-catch samples are commonly contaminated. Even gentle expression of the bladder can lead to hematuria. Calculi containing calcium carbonate require specific methodologies to differentiate calcium carbonate crystals from calcium oxalate monohydrate (COM) crystals; therefore the laboratory chosen for analysis must be able to perform these differentiating methods.20 Calcium carbonate calculi are not found in humans; thus human and some veterinary (those that do not commonly evaluate large/exotic animal samples) laboratories do not use techniques for differentiating calcium oxalate and calcium carbonate. Confirm the lab’s ability to perform these techniques in advance to ensure that the calculus is appropriately evaluated for the guinea pig patient.
Medical treatment of urolithiasis has been unrewarding to date, and surgical removal of the stones is most often required (see Chapter 25). In females, with the patient under deep sedation and analgesia, gently flush the bladder using a 3.5-Fr red rubber catheter to attempt to remove the calculi. Catheterizing boars is much more dangerous because of the small size of the urethra. Submit the calculi for analysis and culture and submit the bladder wall for culture and sensitivity. Postoperative management includes assisted feeding, fluids, analgesics, and antibiotics based on culture and sensitivity results. However, recurrence of the disease is common. Prevention is targeted at increasing water intake and reducing (but not eliminating) dietary calcium. We have seen metabolic bone disease (fibrous osteodystrophy; see “Musculoskeletal Diseases,” below) induced in guinea pigs with severe dietary calcium restriction for prevention of urinary calculi. For most animals, water is the cornerstone of any prevention protocol. Hydrochlorothiazide (2 mg/kg q12h) is a thiazide diuretic that reduces urinary Ca2+, K+, and citrate. It is unknown if diuresis would be of benefit to guinea pigs whose urine is already considered isosthenuric.20 Do not use hydrochlorothiazide with severe renal disease or fluid imbalances. Avoid alfalfa-based diets. Diets containing a high percentage of timothy, oat, or grass hays, a lower overall percentage of pellets, and a wide variety of vegetables and fruits decrease the risk of urolith development in pet guinea pigs.19 It is possible that dietary inhibitors of calcium are found in greater concentration in hays than in pellets. Urinary acidifiers were historically recommended based upon an assumed diagnosis of calcium oxalate calculi, but the normally alkaline urine of guinea pigs making the use of dietary acidifiers concerning. Potassium citrate (30-75 mg/kg PO q12h) binds urinary Ca2+, reduces ion activity, and alkalinizes the urine. Because guinea pigs have alkaline urine even with disease, the efficacy of this treatment is unclear. Hyperkalemia occurs, so monitor plasma K+ closely during treatment. According to anecdotal reports,
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potassium citrate and hydrochlorothiazide have been used together with some clinical success.
CYSTITIS AND URINARY TRACT INFECTIONS It has been reported that female guinea pigs are more vulnerable to bacterial cystitis because the urethral opening is closer to the anus, potentially allowing gastrointestinal bacteria to ascend the urethra into the urinary bladder.40 Bacterial cystitis is frequently identified concomitantly with urolithiasis, but with an equal gender distribution.20 Clinical signs mimic those of urolithiasis, with vocalizing/straining during urination, pollakiuria, dysuria, hematuria, anorexia, and depression. Base your diagnosis and treatment on urinalysis, urine (obtained via cystocentesis) culture and sensitivity results, and radiographs to rule out urolithiasis. Provide analgesics. Gently flush the sow’s bladder using a 3.5-Fr red rubber catheter under deep sedation and analgesia if warranted. Catheterization is not recommended for boars because of the small size of the urethra.
CHRONIC INTERSTITIAL NEPHRITIS AND CHRONIC RENAL FAILURE Chronic interstitial nephritis is commonly found in guinea pigs more than 3 years of age. Guinea pigs more than 1 year of age can develop some degree of renal segmental fibrosis, progressing in the aged animal to chronic renal failure. The pathogenesis of this disease is still unclear. Obstructive urolithiasis and pyelonephritis can lead to interstitial nephritis. It has been reported in guinea pigs with diabetes mellitus and with staphylococcal pododermatitis with chronic renal amyloidosis and nephritis as sequelae.38 Encephalitozoon cuniculi has been shown to cause subacute interstitial nephritis in laboratory animals, but its significance in pet guinea pigs is unknown. Contrary to previous literature, advanced cases present clinically with evidence of renal compromise such as polyuria/polydipsia (PU/PD), azotemia, chronic weight loss, cardiac compromise including dyspnea and secondary heart failure due especially to uremia, and other vague clinical signs such as anorexia or diarrhea/loose stool. Elevations in blood urea nitrogen (BUN) and creatinine, changes in electrolytes and minerals, isosthenuria, and nonregenerative anemia have all been recorded antemortem.22 Typical histopathological changes include interstitial fibrosis, glomerular ectasia, and sclerosis with variable numbers of mononuclear inflammatory cells. The renal parenchyma can be completely distorted, often with no recognizable normal tissue. Cardiac lesions such as epicarditis, myocarditis and fibrosis, pericarditis, and ventricular dilation are often identified concurrently with renal lesions. While chronic interstitial nephritis may be prevented by proper husbandry and sanitation to reduce the guinea pig’s susceptibility to pododermatitis,22,38 appropriate hydration throughout the guinea pig’s lifetime is also prudent.
OTHER UROPATHIES Renal cysts are not uncommon and are usually identified incidentally on necropsy. There is one report of renal failure in a guinea pig following ingestion of peace lily, with similar clinical signs as are seen in other mammals.24 Klossiella cobayae, a renal coccidian, lives in the epithelial cells lining the renal tubules, and sporocysts are shed in the urine. Clinical disease rarely results from infection, and treatment is generally not
Fig. 23-6 Ovarian cysts are identified in up to 75% of female guinea pigs and usually occur in both ovaries.
necessary,30 but sulfadimethoxine or trimethoprim-sulfa have been used. Segmental nephrosclerosis is reported as an incidental finding at necropsy and is often associated with autoimmune disease, high-protein diet, infections, and vascular diseases.41 Cytomegalovirus inclusion bodies have been identified in the kidneys as incidental findings at necropsy.41 Neoplasia of the urinary system is not common, but transitional cell carcinoma of the bladder, renal cell carcinoma, and renal fibrosarcoma have been identified in pet guinea pigs in the practice of one of the present authors (MGH).
REPRODUCTIVE DISEASES OVARIAN CYSTS Nonfunctional serous cysts (cystic rete ovarii) are extraordinarily common and have been identified in 66% to 75% of sows between 3 months and 5 years of age.8,46 Middle-aged (2- to 4-year-old) sows are most commonly affected. Serous cysts develop spontaneously throughout the estrous cycle. Follicular cysts also occurred in 22.4% of one study population and always coincided with serous cysts.46 They may be single or multilocular and are usually filled with clear fluid (Fig. 23-6). Cysts range in diameter from 0.5 to 7 cm and increase in size and prevalence as the animal ages.34 No significant correlation has been identified between reproductive history and the prevalence of cysts,34 but other problems reported concurrently with ovarian cysts include leiomyomas, granulosa cell tumors,8 cystic endometrial hyperplasia, and endometritis. In most cases, both ovaries are affected; however, if the cysts are unilateral, the right ovary is usually affected. The incidence of serous ovarian cysts increased following passive immunization against inhibin, suggesting that serous cysts are a normal component of the cyclic guinea pig ovary and that alterations in the inhibin-folliclestimulating hormone system may modulate the incidence of serous ovarian cysts in cavies.46 Affected animals present with abdominal distention and sometimes with anorexia, weakness, depression, and hunching in pain.4 If cysts are functional (follicular cysts), bilateral symmetric hair loss can be seen in the flank region. The most consistent sign of cysts in breeding sows is a decline in fertility after approximately 15 months of age. Diagnosis is best via ultrasound (Fig. 23-7), but abdominal radiography can identify
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SECTION III Guinea Pigs and Chinchillas after 1 year of age may deliver normally, the majority of older primiparous sows have difficulty with parturition. Sows that are at risk must be monitored closely and may require cesarean section. Signs in full-term sows include contractions and straining that produce a bloody or green discharge but no pups. Pressure from the gravid uterus or pups may also lead to temporary paresis or paralysis of the rear legs. Palpation of the pelvic area for relaxation of the symphysis will help determine the need for a C-section. If the symphysis is separated approximately 2.5 to 3 cm, palpation of the vaginal canal may reveal a pup that must be manually removed or assisted. Sterile water-based lubricating gel placed in the vaginal canal helps ease pup removal. If the symphysis is relaxed and uterine inertia is suspected, treatment with oral or injectable calcium and/or oxytocin may facilitate contractions.
Fig. 23-7 Diagnosis of ovarian cysts is best made via ultra sonography.
larger cysts. The treatment of choice for ovarian cysts is ovariohysterectomy, but space-occupying cysts complicate the procedure. Postoperative care requires assisted feeding, fluids, and analgesics. If GI stasis secondary to pain is present, continue supportive care until the GI tract is functioning appropriately (see “Gastrointestinal Hypomotility or Stasis,” above). Palliative therapy with ultrasound-guided percutaneous drainage of the fluid can be performed, but cysts commonly return, sometimes within days. Gonadotropin-releasing hormone and human chorionic gonadotropin have been used to induce luteinization of ovarian follicular cysts43; but with nonfunctional serous cysts, temporary or no response to treatment is seen.
UTERINE PROLAPSE Uterine prolapse is most commonly associated with parturition.31 Acutely, the prolapse may be pink or red and smooth; however, depending on the duration of the prolapse, the tissues may be dry, darker, and covered with bedding or fecal material. The sow should be stabilized with fluids, analgesics, and antibiotics. The prolapsed tissue is rinsed thoroughly with sterile saline or dilute chlorhexidine solution. Then liberal amounts of a sterile lubricating gel are placed on the tissues to keep them moist until they have been reduced. If the prolapse is acute and the tissues are in good condition, cold hypertonic saline or sugar solutions are used to reduce swelling; then manual reduction of tissues can be performed. If prolapse occurs after reinsertion, ovariohysterectomy should be performed. Future breeding of the sow is not recommended. If the tissues are in poor condition, ovariohysterectomy is recommended after stabilization.
DYSTOCIA Guinea pigs are more predisposed to dystocia than other rodents or rabbits. This may be because of the large size of their pups, narrow pelvic canals, or fusion of the pubic symphysis.6 Other causes suggested are uterine torsion, obesity, nutritional (including vitamin C) deficiencies, and uterine inertia.5,16 Most cases occur in sows first bred after 8 to 12 months of age. Many breeders do not believe that the “fusing” of the pelvic symphysis always occurs, instead blaming dystocia of mature sows on obesity and vitamin C deficiency. Although some sows first bred
TOXEMIA OF PREGNANCY Toxemia of pregnancy is most commonly seen in pregnant sows 2 weeks prepartum to 2 weeks postpartum. There are probably two forms of pregnancy toxemia that may affect the guinea pig. The more familiar form, pregnancy ketosis, is similar to that seen in sheep and is caused by a negative energy balance of the sow due to the heavy demand of the growing fetuses. Predisposing factors include obesity, lack of exercise, large fetal loads, change in diet and/or environment, heat stress, and primiparity.6,18 The second form may be similar to that seen in pregnant women, which is caused by the gravid uterus compressing either its own vascular supply or that of the kidneys or GI tract, leading to tissue ischemia, and hypertension. In some cases, this condition may initiate disseminated intravascular coagulation (DIC).6,18 Hemorrhagic syndrome, which occurs during or shortly after parturition and usually results in fatal hemorrhage in the sow, may be related to this form of toxemia.6 Proposed etiologies for this syndrome in sows include vitamin K deficiency due to poorquality feed, hepatic dysfunction secondary to pressure from the gravid uterus, calcium deficiency affecting the clotting cascade, and other causes of DIC.6,18 Signs of pregnancy ketosis include complete or partial anorexia, lethargy, depression, uncoordinated movements, and dyspnea; this may progress to muscle spasms, paralysis, and death. Some sows die acutely and others deteriorate progressively over several days. The smell of ketones on the breath may occur with ketonemia. Laboratory findings include ketonuria, proteinuria, aciduria, hypoglycemia, acidosis, hyperlipemia, and hyperkalemia.6,18 Proteinuria can occur as a result of proximal tubular necrosis and subcapsular renal hemorrhage.41 Ultrasound imaging of the liver and necropsy findings often identify hepatic lipidosis. Treat ketosis with intravenous (IV) or intraosseous (IO) isotonic fluids with dextrose and oral glucose; calcium gluconate and magnesium sulfate have been suggested to be useful. Immediate emergency treatment of 1 to 2 mL of 50% dextrose administered in 3 to 5 mL saline IV or IO has been recommended. Nutrition is a critical component of treatment and is accomplished by syringe or gavage feeding via gastric, nasogastric, or esophagostomy tube. Herbivore critical care formulas should be used for maintenance feedings. Pregnancy-related blood pressure and ischemia issues associated with the heavily gravid uterus are significantly different from the ketosis type of pregnancy toxemia. Indirect blood pressure measurement will help determine whether the patient is
CHAPTER 23 Disease Problems of Guinea Pigs hypertensive (e.g., compression of the renal vessels) or hypotensive (e.g., shock). If the patient is hypertensive, an emergency cesarean section is required to relieve vascular compression. If the patient is hypotensive, treatment should commence with IV or IO crystalloid or colloid fluids, but it is likely that a cesarean section will be required as well. Treating pregnancy toxemia is often unsuccessful; therefore prevention is essential. Avoiding stress, obesity, and changes in the diet or environment during late pregnancy reduces the potential for pregnancy toxemia. Increase carbohydrate supplementation during the last 2 weeks of gestation and early postpartum period and make sure that food and water are readily available. Encourage exercise to minimize obesity before breeding, as the fetal load may exceed the sow’s weight and she may not be able to exercise sufficiently while pregnant. Postpartum breedings should be avoided as they are more likely to lead to large litters and thus larger fetal loads.
MASTITIS Bacteria cultured from infected mammary glands include E. coli, Pasteurella species, Klebsiella species, Staphylococcus species, Streptococcus species, and Pseudomonas species.6,38 Husbandry-related causes include dirty, wet cages, sharp objects, abrasive bedding, wire cage bottoms, and trauma from pups.6,18,38 The gland may become infected secondary to neoplasia. In acute mastitis the mammary glands are swollen, inflamed, reddened, and warm. The animal is often in pain, reluctant to move or eat, and will not nurse.6,38 In more chronic cases the glands become cool and cyanotic.18,38 There may or may not be a mucopurulent or bloody discharge from the teat or in the milk,6 and some may become abscessed and rupture. Diagnosis is based on clinical signs, culture and sensitivity, and cytology of the discharge or milk. Treat mastitis with antibiotics based on culture and sensitivity results, antiinflammatory medications, analgesics, hot-packing of the glands, and supportive care.6,38 Unresponsive cases may require surgical draining or removal of abscessed glands with histopathology to evaluate for neoplasia.18,38
PYOMETRA AND METRITIS Bacterial infections of the uterus may manifest as metritis or pyometra and are identified in breeding and nonbreeding animals.6 The most commonly isolated bacteria are B. bronchiseptica and hemolytic Streptococcus species, but other possible pathogens include E. coli, Corynebacterium pyogenes, and Staphylococcus species.6,18 Clinical signs include bloody or purulent vaginal discharge, abdominal distention, depression, anorexia, and fever.6 In more chronic cases, sows may be polydipsic and hypothermic. Diagnosis is made based on clinical signs, imaging, and culture and sensitivity results from the discharge. While survey radiographs can show uterine enlargement, ultrasound is the best modality for the diagnosis of this disease. Treatment involves stabilization with supportive care (e.g., IV, IO, SC fluids and nutrition), analgesics and/or NSAIDs, and broad-spectrum antibiotics while awaiting culture and sensitivity results.6,38 Ovariohysterectomy is the treatment of choice in nonbreeding animals.38 Long-term antibiotic therapy may be used in breeding sows, but the authors recommend ovariohysterectomy as the best treatment in all cases of uterine infection.
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VAGINITIS AND “SCROTAL PLUGS” Wet, soiled bedding combined with inguinal sebaceous secretions can become adhered to the penis, scrotum, vulva, or vagina, resulting in secondary infections or urinary/fecal obstruction. Older, intact boars may accumulate a scrotal plug of bedding, feces, or sebaceous material in the skin fold of the bilateral perineal sacs. Gently soak the affected part in a dilute chlorhexidine solution or in warm saline and carefully remove the debris; administer topical or systemic antibiotics as warranted. These conditions are easily prevented by appropriate sanitation and husbandry practices.
ORCHITIS AND EPIDIDYMITIS Infections of the male reproductive tract occur in breeding and nonbreeding boars via sexual transmission, bite wounds, and hematogenous spread.6 The most common pathogens isolated are B. bronchiseptica and Streptococcus species, but other bacterial causes occur, therefore culture any discharge identified.6 Signs include unilateral or bilateral swelling of the scrotal area, preputial discharge, anorexia, weight loss, and fever.6 Animals that recover may become carriers and therefore should not be used for breeding. Treatment involves appropriate antibiotics, supportive care, and analgesics and NSAIDs as indicated. In cases that do not resolve, castration is recommended.
NEOPLASIA There are many reports of reproductive tract tumors in the guinea pig. Leiomyomas and leiomyosarcomas of the uterus, ovarian adenocarcinomas, fibroadenomas, and adenocarcinomas of the mammary glands have all been reported.6,18 Mammary gland tumors are common in boars as well as sows. Approximately 50% are malignant tumors, but metastasis is not common. Hormonal abnormalities and viral etiologies have been proposed. Differentiate this condition from mastitis by cytology and/or biopsy. Clinical signs of mammary tumors include swelling of one or both glands with or without serous or bloody discharge. Ovarian and uterine neoplasms can be induced by exposure to estradiol, diethylstilbestrol, or testosterone.42,47 Clinical signs of uterine and ovarian tumors may be mistaken for pregnancy and include a hemorrhagic vaginal discharge, abdominal distention, and abdominal pain.6 One enlarged testicle with or without testicular atrophy in the opposite gonad is the most common clinical sign of a testicular tumor. The treatment of choice is surgical removal with histopathologic evaluation.6 Prevention of all reproductive neoplasms is best accomplished by ovariohysterectomy or castration of young animals.
DERMATOLOGIC DISEASES To evaluate dermatologic disorders in guinea pigs, it is important to recognize what is normal for a specific breed or for cavies in general. There are 13 recognized cavy breeds in the United States, and many mixed breeds are found in the pet trade. All guinea pigs have minimal or no hair between the nose and lips, around the lips, on the outer ear pinnae, and behind the ears. The Abyssinian and Abyssinian Satin breeds have many rosettes (central hairless areas with hair radiating in a circle) that should not be confused with alopecia or skin disease. The Teddy, Teddy Satin, and Texel breeds have a terrier-like coarse, kinky coat with
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SECTION III Guinea Pigs and Chinchillas
curly whiskers. Occasionally hair and/or eyelashes may curl into the eyes of newborn babies, causing irritation. The Teddy and Texel also seem predisposed to dry, flaky skin. The coronet and white crested breeds have one rosette on top of the head. Peruvian and Peruvian Satin breeds normally have one rosette on the forehead (creating a frontal flow of hair) and one rosette over each hip (causing hair to be directed from the rear toward the head).
DERMATOPHYTOSIS Guinea pigs, like most mammals, are susceptible to fungal infection caused by dermatophytes. Young and immunosuppressed animals are more susceptible.38 Purebred Teddy and Teddy Satin guinea pigs appear to be the most susceptible to fungal infections. Dermatophytosis presents with scaly, patchy lesions on the face, feet, and dorsum.14 These skin lesions are usually circular areas of alopecia with inflamed and sometimes crusty edges; they are also pruritic.14,38 Although both Trichophyton mentagrophytes and Microsporum canis have been isolated, the majority of clinical cases are caused by T. mentagrophytes. Diagnosis is based on clinical signs, ultraviolet fluorescent lamp evaluation (Wood’s lamp), cytology, and fungal culture. T. mentagrophytes does not fluoresce with Wood’s lamp evaluation; thus this test is often negative.14,38 Fungal culture is required for accurate diagnosis. Treatment includes oral antifungal medications (itraconazole, 5 mg/kg q24h × 4-6 weeks; griseofulvin, 25 mg/kg q24h × 14-28 days; fluconazole, 16 mg/kg q24h × 14 days), antifungal shampoos (ketoconazole/chlorhexiderm), and topical antifungal lotions or sprays (miconazole, enilconazole, butenafine).14,38 Dermatophytosis is potentially zoonotic and the organisms may survive in the environment, leading to reinfection.38
ECTOPARASITES The most common ectoparasites found in guinea pigs include mites (Trixacarus caviae, Chirodiscoides caviae), lice (Gliricola porcelli, Gyropus ovalis), and rarely Demodex caviae.14,18,25,38 The most severe ectoparasitic dermatitis is caused by T. caviae. Infection can be caused by direct or indirect contact. Severe pruritus is a hallmark sign of infection; animals can scratch so intensely that they appear to be having a seizure; this may lead to severe self-induced trauma with secondary fungal or bacterial infections.14,18,38 Lesions include white-yellow crusty areas with inflammation and abrasions from self-induced trauma (Fig. 23-8).14,38 Louse infestation usually causes a less severe dermatitis and, because lice spend their entire life cycle on the host, requires direct contact for transmission. In heavy infestations, guinea pigs may have alopecia, crusty lesions, and an unthrifty-looking the coat. Flea (Ctenocephalides felis) infestations are also reported in guinea pigs.38 Cavies housed with rabbits, other rodents, or birds may become transiently infected with other ectoparasites (e.g., Sarcoptes muris, Notoedres muris, Mycoptes musculinus).25 Diagnose the condition by direct visualization of some ectoparasites such as lice and fleas, microscopic examination of skin scrapings, acetate tape preparations, and trichograms for specific organisms or parasite eggs.14,18,38 A microspatula with a flat-ended blade is preferable for skin scrapings, but the dull edge of a scalpel blade can also be used. Perform bacterial and fungal cultures of skin if secondary infections are suspected.
Fig. 23-8 Dermatitis (sarcoptic mange) caused by Trixacarus caviae. Severe infestation is seen, with crusty lesions covering the body.
Treatment is based upon the organism identified, but all products available are used off label in the guinea pig. Ivermectin (0.2-0.5 mg/kg PO, SC q7-10d × 3-4 treatments) has been the treatment of choice for T. caviae, but anecdotally topical selamectin treatment (6 mg/kg q14d × 3 treatments) has been successful. Treatment for lice and fleas may be accomplished with flea shampoos and powders that are safe for kittens. Ivermectin (0.2-0.5 mg/kg PO, SC q7-10d × 2 treatments) has been successful against lice. Use of monthly dog and cat topical flea products in guinea pigs is common for fleas and lice, but controversy exists as to the efficacy of some, and there are no data regarding the safety of any of these products in this species. Treat severe pruritus with antihistamines (diphenhydramine, hydroxyzine) and/or NSAIDs.14 Treat animals with secondary bacterial or fungal infections based upon culture and sensitivity. Asymptomatic animals should also be treated for ectoparasites and the environment disinfected.14 Lice are usually species-specific.25 T. caviae can be zoonotic and survive in the environment, leading to reinfection.38
CERVICAL LYMPHADENITIS Cervical lymphadenitis is usually caused by Streptococcus zooepidemicus and occasionally by Streptobacillus moniliformis, which is potentially zoonotic.14,25,38 S. zooepidemicus is considered part of the normal oropharyngeal/nasal flora of guinea pigs. Oral abrasions caused by overgrown teeth, abrasive feed, or bite wounds lead to invasion of the bacteria into deeper tissues and cervical lymph nodes, which become abscessed.14,25,38 Guinea pigs are presented with swellings in the neck region that are purulent and occasionally rupture on their own. In rare cases S. zooepidemicus can spread systemically and cause pneumonia, metritis, and septicemia.38 One of the present authors (CB) has observed an outbreak of S. zooepidemicus in a herd of approximately 500 guinea pigs. Animals were identified with typical signs of cervical lymphadenitis but also with acute hind-end paralysis, enlarged lymph nodes in the caudal region, metritis, abortion, dermatitis, and pneumonia. In all animals tested, the organism recovered was S. zooepidemicus. Diagnosis is based upon clinical signs, Gram’s stain, culture and sensitivity results from purulent material or lymph node tissue, and biopsy results of tissues.14,25,38 Treatment may require surgical removal or lancing of the abscessed lymph nodes and systemic antibiotics based on culture and sensitivity results.
CHAPTER 23 Disease Problems of Guinea Pigs
Fig. 23-9 Pododermatitis in the guinea pig. Swollen soft tissues and ulceration, as seen here, should alert the clinician to potential underlying osteomyelitis; radiographs are warranted.
PODODERMATITIS Pododermatitis is commonly seen in pet and laboratory cavies. Lesions may originally develop from irritation due to improper husbandry (e.g., wire bottom cages, abrasive or soiled bedding, sharp pieces of wood chips) and secondary bacterial invasion most often involves Staphylococcus aureus.25,38 The condition is most common in obese adult guinea pigs and results in a significant amount of pain and disability. Affected animals are reluctant to move around or eat and frequently are more vocal than normal.38 Vitamin C deficiency may be an important predisposing factor in animals with pododermatitis and supplementation is indicated as part of treatment protocols.14,25 Clinical signs vary from mild-to-severe inflammation, erythematous lesions with or without ulceration of primarily the plantar surfaces of the feet (Fig. 23-9), to granulomatous, callous-like swellings that, in severe cases, may result in bacterial invasion into the tendons, joints, and bone (e.g., osteomyelitis).14,25,38 Chronic inflammation may lead to amyloidosis of the kidney, liver, spleen, adrenal glands, and pancreas.25 Radiographs are indicated to diagnose osteomyelitis.38 In less severe cases, soak the lesions with dilute chlorhexidine or iodine solutions, maintain the animals on soft substrates, bandage, and administer systemic antibiotics based upon culture and sensitivity results. More severe cases may require long-term systemic antibiotics, surgical debridement ± antibiotic-impregnated polymethylmethacralate beads, bandaging with regular changes, and, as a salvage procedure, limb amputation.14,25,38 Analgesic and anti-inflammatory medications are an important part of treatment.14,25,38 The prognosis is poor in severe cases and prevention is essential. Proper diet, husbandry, cleanliness, and prevention of obesity are all helpful in the prevention of pododermatitis.
ALOPECIA Alopecia without inflammation is often related either to husbandry or hormonal conditions.14,25 Nutritional deficiencies as well as poor sanitation and bedding material reactions can cause alopecia in the guinea pig. Hormonal causes of alopecia include follicular ovarian cysts (bilaterally symmetric, nonpruritic; see
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Fig. 23-10 One of the most common masses seen in guinea pigs, the trichofolliculoma (arrow).
“Reproductive Diseases,” above) and hormonal changes common in late pregnancy or in lactating sows in poor condition or with large litters.14,38 The hairs are not completely epilated with barbering; close examination of the skin reveals broken hair shafts. Self-inflicted barbering may occur out of boredom or poor nutrition. The addition of hay and toys to the environment may resolve the problem. Dominant guinea pigs will barber their cage mates and may have to be separated. Examination of the barbering pattern will reveal whether it is self-inflicted (in which case the head and neck are spared).14,38
NEOPLASIA Trichofolliculomas are the most common tumors of guinea pigs. They are benign skin tumors seen predominantly in males and often arise on the dorsal rump, incorporating the scent glands (Fig. 23-10). They can be large, malodorous, exudative, and ulcerated, frequently with secondary infections and/or myiasis. Complete excision is usually curative. They are also considered incidental in most cases. Liposarcoma has a relatively high prevalence and is generally found in the skin or subcutis. Lymphoma is perhaps the most common malignancy, usually multicentric, and may appear in skin and viscera. Typically, lymphoma in cavies is a high-grade malignancy with a poor prognosis. Fibropapillomas of the ear canal are occasionally found in cavies. The etiology is undetermined; they are benign and usually resolve spontaneously.
MUSCULOSKELETAL DISEASES VITAMIN C DEFICIENCY (SCURVY) Guinea pigs are incapable of endogenous synthesis of vitamin C because they possess a mutated gene for l-gulono-g-lactone oxidase, which prevents the conversion of l-gulonolactone to l-ascorbic acid.35 Lack of dietary vitamin C results in defective type IV collagen, laminin, and elastin; this compromises blood vessel integrity and results in joint and gingival hemorrhages.29 Collagen is necessary to anchor teeth tightly; without it, teeth loosen and malocclusion occurs. In addition, vitamin C is necessary for appropriate retention of vitamin E.27 Young, growing animals are more susceptible to scurvy, and clinical disease can
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A
B Fig. 23-11 Postmortem findings of vitamin C deficiency in guinea pigs include periarticular hemorrhage (arrow), swollen joints (especially the stifle) (A), and enlargement of the costochondral junctions of the ribs (B) (arrows).
develop after as little as 2 weeks of ascorbic acid deprivation.10 Guinea pigs require 10 to 25 mg/kg per day of vitamin C added to their diet; pregnant animals require 30 mg/kg per day.17 Signs of vitamin C deficiency include rough hair coat, anorexia or difficulty prehending food, diarrhea, teeth grinding, vocalizing from pain, delayed wound healing, lameness, swollen joints (especially the stifle; Fig. 23-11A), GI stasis, and increased susceptibility to bacterial infections.30 Radiographically, long bone epiphyses and costochondral junctions of the ribs are enlarged (Fig. 23-11B). Pathologic fractures may also be evident. Postmortem examination may reveal hemorrhage into joints, skeletal muscle, gingiva, intestine, and subcutaneous tissues from abnormal collagen production. The diagnosis of vitamin C deficiency is based on history, clinical signs, and radiographic and pathologic lesions. Serum ascorbic acid levels can be used to confirm the diagnosis. Treat vitamin C deficiency with parenteral ascorbic acid at a dose of 50 to 100 mg/day IM or SC, but use caution with intramuscular medication because of the musculoskeletal pain associated with this disease. Once response is noted, administer vitamin C orally at the same dose. After recovery, ensure adequate supplementation of vitamin C daily in the diet. Fresh, good-quality guinea pig pellets provide adequate vitamin C if used within 90 days of the production date, but dietary recommendations have recently been made to reduce the percentage pellets in the diet because of possible urolithiasis.19 Fresh cabbage, kale, and oranges provide good sources of vitamin C. Vitamin C tablets (50 mg) are now available and can be freshly crushed and sprinkled over vegetables. Vitamin C added to the drinking water at a concentration of 200 to 400 mg/L daily has been advocated; but the vitamin is unstable in light and is most likely inactivated quickly.
OSTEOARTHRITIS AND OSTEOARTHROSIS Spontaneous osteoarthritis is clinically seen in guinea pigs but can also be secondary to ulcerative pododermatitis (see “Dermatologic Diseases,” above). Obesity and improper exercise and/ or substrate are predisposing factors. Treatment is palliative: soft clean bedding, pain management, increased exercise and/ or physical therapy of the limbs to increase range of motion, and prevention of obesity. Spontaneous cartilage degeneration and osteoarthrosis of the femorotibial joints of young guinea pigs have been described but no cause was identified.3
FIBROUS OSTEODYSTROPHY While there is only one report in the literature to date,44 anecdotal reports suggest that fibrous osteodystrophy is being documented in pet guinea pigs with increasing frequency. This condition is caused by primary or secondary hyperparathyroidism and results in increased osteoclastic resorption of bone and replacement by fibrous tissue. An imbalanced Ca:P ratio was found in the guinea pigs from the one report,44 and we have also seen guinea pigs presented with this disease that were purposely placed on low-calcium diets to prevent urolithiasis. Breeders in Europe and the United States also report this disease in the Satin type, suggesting an inherited factor in this line of guinea pigs. Diets with a low Ca:P ratio cause reduced growth rates, stiffness of joints, and calcification of soft tissues; they may also affect the uptake of magnesium and potassium.36 In addition, vitamin D deficiency, calcium malabsorption, or other calcium metabolism disturbances may play a role in the onset of disease. Clinical signs include anorexia or difficulty eating, lethargy, difficulty walking or unwillingness to move, or pathologic fractures. Diagnosis is based upon dietary history, physical examination findings, plasma total and ionized calcium concentrations, serum 25-hydroxyvitamin D, and parathyroid hormone concentrations. This disease must be differentiated from hypovitaminosis C, as the clinical signs can appear similar. Radiographs often show extensive changes to all bones in the body and skull, including osteopenia and pathologic fractures.44 Necropsy findings include severely thinned trabecular bone, marked osteoclastic activity, resorption of cortical bone and extensive replacement with fibrous connective tissue, and hyperplastic parathyroid glands (M. G. Hawkins, personal observation). No renal lesions have been identified to date. Treatment is aimed at normalizing Ca:P ratios via the diet. Initially treat with calcium gluconate intramuscularly, then continue with oral calcium glubionate therapy, in some cases this will be required for months before resolution is seen. Supplement vitamin D and increase exposure to sunlight empirically. Osteoporosis has also been reported in guinea pigs, so vitamin D supplementation should be used with caution.23 Pain management should include opioid medications for skeletal pain as well as anti-inflammatory medications. Provide supportive care, including assisted
CHAPTER 23 Disease Problems of Guinea Pigs feeding and fluids as needed. Prognosis is guarded without aggressive supportive care.
METASTATIC MINERALIZATION Metastatic mineralization occurs in guinea pigs generally above 1 year of age. Often disease is subclinical, but clinical signs include unthriftiness, muscle stiffness, and renal dysfunction. The etiology is unclear and is possibly related to subclinical dietary imbalances such as an imbalanced Ca:P ratio, low magnesium and potassium, oversupplementation of dietary vitamin D3 or minerals, or dehydration.22,23,36,41 Soft tissue mineralization, including mineralization of kidneys, heart, vessels, brain, and GI tract, may be identified on radiographs.41 Lesions are irreversible once evident; prevention is by providing an appropriately balanced diet.23
IATROGENIC MUSCLE NECROSIS Ketamine, diazepam, and fentanyl/droperidol combinations have been implicated in nerve damage, self-mutilation, and muscle necrosis at injection sites.38
NUTRITIONAL MUSCULAR DYSTROPHY Myopathies have been reported to be associated with vitamin E/ selenium deficiencies. Clinical signs include lethargy, hind-limb weakness, decreased reproduction in breeding sows and boars, and conjunctivitis.41 Creatine kinase (CK) may be elevated. Severely affected animals may die within 1 week of presentation. As in other animals with this disease, the muscles may be pale and streaked at necropsy. Animals may respond to supplementation with vitamin E.
NEUROLOGIC DISEASES OTITIS MEDIA/OTITIS INTERNA While less frequent than in rabbits, otitis media occurs in pet guinea pigs and can progress to otitis interna. B. bronchiseptica, S. zooepidemicus, and S. pneumoniae have been associated with otitis media.23 It is unclear whether oral flora can become opportunistic with dental disease and ascend via the eustachian tube in guinea pigs, but otitis media and dental disease are often found together and cultures often isolate oral bacteria. Also aural polyps may occur within the tympanic bullae, with secondary infection causing clinical signs. Clinical signs include head tilt, ataxia, circling, torticollis, and facial nerve paralysis with secondary ulcerative keratitis due to exposure. Take skull radiographs to evaluate the bullae (Fig. 23-12A,B) and dentition; computed tomography can also be useful to determine the extent of the disease (Fig. 23-12C). Treat with appropriate antibiotics, anti-inflammatories, analgesics, and dental treatments, but these measures are often not curative. If needed, flush the ears with warm sterile saline under deep sedation/analgesia to help break up caseous material. Surgery (bullous osteotomy) has not been reported for guinea pigs.
MITES Severe infestations with T. caviae can cause such severe pruritus that guinea pigs are often presented for seizing (see “Dermatologic Diseases,” above).
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RABIES Although rodents are not natural reservoirs for rabies virus, rabies has nevertheless been reported in guinea pigs.12 Clinicians should always consider rabies in guinea pigs with neurologic signs and husbandry conditions that could allow exposure.
LYMPHOCYTIC CHORIOMENINGITIS VIRUS Lymphocytic choriomeningitis virus (LCMV) is an arenavirus that can cause meningitis and hind-limb paralysis in guinea pigs, although it is more commonly reported in mice, hamsters, and chinchillas. Lesions include lymphocytic infiltrates in the choroid plexus, ependyma, and meninges.30 The virus is transmitted through inhalation, ingestion, or direct contact with contaminated urine, saliva, and feces. Biting insects can transmit LCMV, and transplacental transmission also occurs. LCMV can be transmitted to humans. Signs of LCMV infection in humans include headache, vomiting, and fever; fatalities are rare.
OPHTHALMOLOGIC DISEASES CORNEAL ULCERATION Corneal ulceration is most commonly caused by trauma and is diagnosed by fluorescein staining. Disseminated T-cell lymphosarcoma has been reported in a guinea pig that was presented with a unilateral corneal opacity.49
“PEA EYE” Adult guinea pigs, especially purebred American shorthairs, develop a protrusion from the inferior conjunctival sac in one or both eyes, termed “pea eye” by fanciers (Fig. 23-13). Histopathology of these tissues has demonstrated lesions in the lacrimal or zygomatic glands.26 Some lesions cause ventral ectropion, lagophthalmos, and secondary axial corneal degeneration.26 However, this condition does not appear to be painful and usually resolves without treatment.
CONJUNCTIVITIS Conjunctivitis is very common in pet guinea pigs and is generally caused by infection or hypovitaminosis C (scurvy). Chlamydophila caviae causes guinea pig inclusion conjunctivitis (GPIC) in laboratory and pet guinea pigs.28,51 Juvenile animals are most commonly affected. Some infected animals remain asymptomatic, but clinical signs range from mild to severe keratoconjunctivitis with serous to purulent ocular discharge, conjunctival chemosis, follicular hypertrophy, and uveitis.28 Diagnose GPIC by identifying intracytoplasmic inclusions in conjunctival scrapings and by performing C. caviae PCR on conjunctival swabs or scrapings.28,30,51 In a recent study, 48 of 75 symptomatic and 11 of 48 asymptomatic adult guinea pigs were positive via PCR for C. caviae. Conjunctival samples submitted from one owner and his cat and rabbit were also positive by PCR for C. caviae, and C. caviae was identified in rabbits housed with guinea pigs in another study.39 While the route of transmission is still unclear for this disease, nonetheless C. caviae GPIC could pose zoonotic
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A
B
C Fig. 23-12 (A) Normal (arrows) and (B) abnormal (arrows) radiographic appearances of the tympanic bullae of the guinea pig and (C) computed tomography (CT) imaging of bilateral tympanic bullae disease in the guinea pig. While survey radiography can provide evidence of sclerosis of the bullae, CT can provide evidence for osteoproliferative or osteolytic changes (arrows) as well as changes to the ear canals.
potential. It is often self-limiting within 3 to 4 weeks, but the potential zoonoses warrant treatment consideration with a topical tetracycline ophthalmic ointment. Conjunctivitis is one of the many clinical signs seen with hypovitaminosis C. Neoplasia of the guinea pig eye is uncommon38,49; lymphosarcoma affecting the conjunctival lymphoid tissue is most commonly reported.38
MISCELLANEOUS DISEASES OTOTOXICITY Guinea pigs seem quite sensitive to antibiotics; they may develop antibiotic-induced ototoxicity, especially when the agent is applied topically.1,7,32,56 Cisplatin induces ototoxicity when administered at human clinical doses,45 making guinea pigs a common model for study of this condition.
DIABETES MELLITUS Spontaneous diabetes mellitus similar to adult-onset diabetes in humans has been described in laboratory colonies of guinea pigs. Shortened life span (≤5 years), bladder hypertrophy, and voiding dysfunction have been reported.2 A pet guinea pig was diagnosed with diabetes mellitus after the animal presented with cystitis and urination of small, frequent amounts; it responded to insulin therapy.53,54 Diabetes mellitus may be transient in the guinea pig, and insulin therapy is generally not necessary. A low-fat, high-fiber diet is most important in treatment and prevention.54
HEAT STRESS Guinea pigs are susceptible to heat stress. Those housed outdoors can develop heat stress in ambient temperatures as low as 75°F (24°C). Guinea pigs will salivate profusely in an attempt
CHAPTER 23 Disease Problems of Guinea Pigs
Fig. 23-13 Adult guinea pigs, especially purebred American shorthairs, develop a protrusion from the inferior conjunctival sac in one or both eyes, termed “pea eye” by fanciers.
to thermoregulate; they will exhibit shallow, rapid respirations, pale mucous membranes, and elevated rectal temperature, which may be followed by coma and death. Treatment is supportive and includes cool water baths and parenteral fluids. Prognosis is very guarded.
LYMPHOSARCOMA/CAVIAN LEUKEMIA Lymphosarcoma is the most common malignancy of guinea pigs. Typically, lymphosarcoma in guinea pigs is a highgrade malignancy with poor prognosis. Clinical signs include anorexia, lethargy, unkept coat, and peripheral lymphadenopathy. Hepatomegaly, splenomegaly, and mediastinal masses are occasionally identified. It is sometimes seen concurrently with lymphoblastic leukemia.10 Cavian leukemia caused by a type-C retrovirus was reported in the 1960s in laboratory guinea pigs; leukemic animals may have a total white blood cell count of 25,000 to 500,000/mL.10 Diagnosis is based on the results of a complete blood count and cytologic examination of aspirates of enlarged nodes or abdominal or pleural fluids. At necropsy, lymph nodes and visceral organs may be enlarged, with infiltration by proliferating lymphoblasts. The course of the disease can be short, as little as 2 to 5 weeks. The prognosis is generally poor, although some animals have responded initially to chemotherapy. To date, there are no published reports regarding the use of chemotherapeutics in guinea pigs.
References 1. Aquino TJ, Oliveira JA, Rossato M. Ototoxicity and otoprotection in the inner ear of guinea pigs using gentamicin and amikacin: ultrastructural and functional aspects. Braz J Otorhinolaryngol. 2008;74:843-852. 2. Belis JA, Curley RM, Lang CM. Bladder dysfunction in the spontaneously diabetic male Abyssinian-Hartley guinea pig. Pharmacology. 1996;53:66-70.
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3. Bendele A, McComb J, Gould T, et al. Animal models of arthritis: relevance to human disease. Toxicol Pathol. 1999;27:134-142. 4. Beregi A, Zorn S, Felkai F. Ultrasonic diagnosis of ovarian cysts in ten guinea pigs. Vet Radiol Ultrasound. 1999;40:74-76. 5. Bishop C. Emergency medicine and surgery of rabbits and rodents. In: Proceedings. Ontario Vet Med Assoc. 1999:115-124. 6. Bishop CR. Reproductive medicine of rabbits and rodents. Vet Clin North Am Exot Anim Pract. 2002;5:507-535. 7. Blakley BW, Hochman J, Wellman M, et al. Differences in ototoxicity across species. J Otolaryngol Head Neck Surg. 2008; 37:700-703. 8. Burns RP, Paul-Murphy J, Sicard GK. Granulosa cell tumor in a guinea pig. J Am Vet Med Assoc. 2001;218:726-728. 9. Chrisp CE, Suckow MA, Fayer R, et al. Comparison of the host ranges and antigenicity of Cryptosporidium parvum and Cryptosporidium wrairi from guinea pigs. J Protozool. 1992;39:406-409. 10. Collins B. Common diseases and medical management of rodents and lagomorphs. In: Jacobson E, Kollias G, eds. Exotic animals. New York: Churchill Livingstone; 1988:261-316. 11. De Voe R. Clinical snapshot: gastric bloat with possible volvulus in a guinea pig. Compend Contin Educ Pract Vet. 2001; 23:543:571. 12. Eidson M, Matthews SD, Willsey AL, et al. Rabies virus infection in a pet guinea pig and seven pet rabbits. J Am Vet Med Assoc. 2005;227:932-935. 13. Fehr M, Rappold S. Urolithiasis in 20 guinea pigs (Cavia porcellus). Tierarztl Prax. 1997;25:543-547. 14. Flecknell P. Guinea pigs. In: Meredith A, Redrobe S, eds. BSAVA manual of exotic pets. 4th ed. Quedgeley: BSAVA; 2002:52-64. 15. Gaschen L, Ketz C, Lang J, et al. Ultrasonographic detection of adrenal gland tumor and ureterolithiasis in a guinea pig. Vet Radiol Ultrasound. 1998;39:43-46. 16. Hardesty D. The effect of obesity on fertility in the cavy. In: The American Rabbit Breeders Association. Official guide book raising better rabbits and cavies. Bloomington, IL: ARBA; 2000: 196-200. 17. Harkness JE, Turner PV, Vande Woude S, et al. Biology and husbandry-guinea pig. In: Harkness and Wagner’s biology and medicine of rabbits and rodents. Ames: Wiley-Blackwell; 2010:45-57. 18. Harkness JE, Turner PV, Vande Woude S, et al. Specific diseases and conditions. In: Harkness and Wagner’s biology and medicine of rabbits and rodents. Ames: Wiley-Blackwell; 2010:249-396. 19. Hawkins MG, Drazenovich TL, Kass PH, et al. Risk factors associated with the development of urolithiasis in pet guinea pigs (Cavia porcellus). Proceedings. Annu Assoc Avian Vet Assoc Exot Mam Vet. 2008:59. 20. Hawkins MG, Ruby AL, Drazenovich TL, et al. Composition and characteristics of urinary calculi from guinea pigs. J Am Vet Med Assoc. 2009;234:214-220. 21. Hawkins MG, Vernau W, Drazenovich TL, et al. Results of cytologic and microbiologic analysis of bronchoalveolar lavage fluid in New Zealand white rabbits. Am J Vet Res. 2008;69:572-578. 22. Hoefer H, Latney L. Rodents: urogenital and reproductive system disorders. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester: BSAVA; 2009:150-160. 23. Hollamby S. Rodents: neurological and musculoskeletal disorders. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester: BSAVA; 2009:161-168. 24. Holowaychuk MK. Renal failure in a guinea pig (Cavia porcellus) following ingestion of oxalate containing plants. Can Vet J. 2006;47:787-789. 25. Huerkamp MJ, Murray KA, Orosz SE. Guinea pigs. In: LaberLaird K, Swindle MM, Flecknell P, eds. Handbook of rodent and rabbit medicine. New York: Elsevier Science Inc; 1996:91-95. 26. Kern TJ. Rabbit and rodent ophthalmology. Sem in Avian Exot Pet Med. 1997;6:138-145.
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27. Liu JF, Lee YW. Vitamin C supplementation restores the impaired vitamin E status of guinea pigs fed oxidized frying oil. J Nutr. 1998;128:116-122. 28. Lutz-Wohlgroth L, Becker A, Brugnera E, et al. Chlamydiales in guinea-pigs and their zoonotic potential. J Vet Med A Physiol Pathol Clin Med. 2006;53:185-193. 29. Mahmoodian F, Peterkofsky B. Vitamin C deficiency in guinea pigs differentially affects the expression of type IV collagen, laminin, and elastin in blood vessels. J Nutr. 1999;129:83-91. 30. Manning P, Wagner JE, Harkness J. Biology and diseases of guinea pigs. In: Fox J, Cohen B, Loew F, eds. Laboratory animal medicine. Orlando: Academic Press; 1984:149-181. 31. Mehler SJ, Bennett RA. Surgical oncology of exotic animals. Vet Clin North Am Exot Anim Pract. 2004;7:783-805. 32. Migirov L, Himmelfarb M. Methodology for studying the effects of topically applied ear drops on otoacoustic emissions in guinea pigs. J Laryngol Otol. 2003;117:696-699. 33. Mitchell EM, MacLeod A, Hawkins MG. Gastric dilatation- volvulus in a guinea pig (Cavia porcellus). J Am Anim Hosp Assoc. 2010;46:174-180. 34. Nielsen TD, Holt S, Ruelokke ML, et al. Ovarian cysts in guinea pigs: influence of age and reproductive status on prevalence and size. J Small Anim Pract. 2003;44:257-260. 35. Nishikimi M, Kawai T, Yagi K. Guinea pigs possess a highly mutated gene for l-gulono-gamma-lactone oxidase, the key enzyme for l-ascorbic acid biosynthesis missing in this species. J Biol Chem. 1992;267:21967-21972. 36. O’Dell BL, Morris ER, Pickett EE, et al. Diet composition and mineral balance in guinea pigs. J Nutr. 1957;63:65-77. 37. Okewole PA, Odeyemi PS, Oladunmade MA, et al. An outbreak of Streptococcus pyogenes infection associated with calcium oxalate urolithiasis in guinea pigs (Cavia porcellus). Lab Anim. 1991;25:184-186. 38. O’Rourke DP. Disease problems of guinea pigs. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2004:245-254. 39. Pantchev A, Sting R, Bauerfeind R, et al. Detection of all Chlamydophila and Chlamydia spp. of veterinary interest using species-specific real-time PCR assays. Comp Immunol Microbiol Infect Dis. 2010;33:473-484. 40. Peng X, Griffith JW, Lang CM. Cystitis, urolithiasis and cystic calculi in ageing guinea pigs. Lab Anim. 1990;24:159-163. 41. Percy DH, Barthold SW. Guinea pig. In: Percy DH, Barthold SW, eds. Pathology of laboratory rodents and rabbits. 2nd ed. Ames: Blackwell Publishing; 2001:209-247.
42. Porter KB, Tsibris JC, Porter GW, et al. Use of endoscopic and ultrasound techniques in the guinea pig leiomyoma model. Lab Anim Sci. 1997;47:537-539. 43. Purswell BJ, Parker NA, Bailey TL, et al. Theriogenology question of the month. Persistent estrus caused by functional granulosa cell tumor of the left ovary. J Am Vet Med Assoc. 1999;215:193-195. 44. Schwarz T, Stork CK, Megahy IW, et al. Osteodystrophia fibrosa in two guinea pigs. J Am Vet Med Assoc. 2001;219:63-66. 45. Sepmeijer JW, Klis SF. Distribution of platinum in blood and perilymph in relation to cisplatin induced ototoxicity in the guinea pig. Hear Res. 2009;247:34-39. 46. Shi F, Petroff BK, Herath CB, et al. Serous cysts are a benign component of the cyclic ovary in the guinea pig with an incidence dependent upon inhibin bioactivity. J Vet Med Sci. 2002;64:129-135. 47. Silva EG, Tornos C, Deavers M, et al. Induction of epithelial neoplasms in the ovaries of guinea pigs by estrogenic stimulation. Gynecol Oncol. 1998;71:240-246. 48. Spink RR. Urolithiasis in a guinea pig (Cavia porcellus). Vet Med Small Anim Clin. 1978;73:501-502. 49. Steinberg H. Disseminated T-cell lymphoma in a guinea pig with bilateral ocular involvement. J Vet Diagn Invest. 2000; 12:459-462. 50. Stieger SM, Wenker C, Ziegler-Gohm D, et al. Ureterolithiasis and papilloma formation in the ureter of a guinea pig. Vet Radiol Ultrasound. 2003;44:326-329. 51. Strik NI, Alleman AR, Wellehan JFX. Conjunctival swab cytology from a guinea pig: it’s elementary! Vet Clin Pathol. 2005;34:169-171. 52. Stuppy DE, Douglass PR, Douglass PJ. Urolithiasis and cystotomy in a guinea pig (Cavia porcellanus). Vet Med Small Anim Clin. 1979;74:465-467. 53. Vannevel J. Diabetes mellitus in a 3-year-old, intact, female guinea pig. Can Vet J. 1998;39:503. 54. Vannevel J. Diabetes in the guinea pig–not uncommon. Can Vet J. 1999;40:613. 55. Xia Y, Hu HZ, Liu S, et al. Clostridium difficile toxin A excites enteric neurons and suppresses sympathetic neurotransmission in the guinea pig. Gut. 2000;46:481-486. 56. Xu M, Hu HT, Jin Z, et al. Ototoxicity on cochlear nucleus neurons following systemic application of gentamicin. Acta Otolaryngol. 2009;129:745-748.
CHAPTER
24
Disease Problems of Chinchillas
Christoph Mans, MedVet, and Thomas M. Donnelly, BVSc, Diplomate ACLAM
Disorders of the Digestive System Gastroenteritis and Dysbacteriosis Constipation Diarrhea and Soft Feces Tympany Rectal Tissue Prolapse and Intussusception Esophageal Disorders Dental Disorders Eye Disorders Epiphora Conjunctivitis Corneal Disorders Other Eye Disorders Ear Disorders Respiratory System Disorders Reproductive System Disorders Endometritis and Pyometra Dystocia Penile Disorders Urinary System Disorders Neurologic Disorders Seizures Heat Stroke Lead Poisoning Dermatologic Disorders Dermatophytosis Fur Chewing Fur Slip Matted Fur Foot Disorders Miscellaneous Disease Problems Cardiac Disease Hepatic Lipidosis and Ketosis Diabetes Mellitus Fractures Neoplasia Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
Infectious Diseases Viral Infections Fungal Infections Parasitic Infections Bacterial Infections
DISORDERS OF THE DIGESTIVE SYSTEM GASTROENTERITIS AND DYSBACTERIOSIS In chinchillas, any systemic disease or a painful or stressful condition may result in secondary gastrointestinal problems with nonspecific clinical signs such as anorexia, lack of fecal output, and lethargy. Obtaining a thorough clinical history and physical examination is critical for formulating a sound diagnostic and therapeutic plan. A variety of infectious and noninfectious causes of gastroenteritis and dysbacteriosis may affect chinchillas and result in a range of clinical syndromes including constipation, tympany, diarrhea, intussusception, and rectal prolapse. Gastrointestinal disorders were the major cause of morbidity and mortality in farmed chinchillas in the 1960s to 1990s. In pet chinchillas, noninfectious causes such as sudden dietary changes or inappropriate oral antibiotic therapy (e.g., cephalosporins, penicillins, clindamycin, erythromycin) are more frequent and important. However, a few secondary infectious causes such as giardiasis, coccidiosis, and Pseudomonas or Enterobacter overgrowth can be seen in pet chinchillas. Identifying the underlying cause of gastroenteritis and dysbacteriosis is important to improve the therapeutic outcome and reduce the chance of recurrence. Consider performing whole-body radiographs, fecal parasite examination, fecal cytology, and fecal culture for enteric opportunistic pathogens (e.g., Escherichia coli, Pseudomonas aeruginosa) in the initial diagnostic workup. Laboratory tests such as urinalysis, plasma biochemical analysis, and a complete blood count (CBC) can aid in diagnosing non-alimentary and concurrent metabolic disorders (e.g., hepatic lipidosis, ketosis, renal disease) that will influence 311
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the prognosis and therapy. Besides specific treatment for the primary underlying cause of gastroenteritis and dysbacteriosis, general treatment guidelines for all cases of gastrointestinal disease should include replacing fluid deficits and maintaining normovolemia by parenteral and enteral fluid therapy, nutritional and caloric support, and managing pain (buprenorphine 0.03-0.05 mg/kg SC q8h) if a painful condition is suspected. Consider systemic parenteral antimicrobial therapy (enrofloxacin 10 mg/kg SC diluted q12h) (Baytril Animal Health, Shawnee Mission, KS) for the treatment of predominately gramnegative opportunistic pathogens in chinchillas with severe dysbacteriosis when an infectious cause is suspected but unconfirmed and the animal is in a compromised general condition. Avoid oral drug administration because the absorption and effectiveness of oral drugs may be decreased when gastrointestinal function is abnormal. Parenteral administration is the preferred initial route for most drugs. Once an animal is eating and gastrointestinal function is improved, switch to the oral route.
CONSTIPATION Constipation is a common problem in chinchillas.16,61 Reduced output or absence of fecal pellets that are smaller and irregular in size is commonly seen. Sudden changes in diet, an inappropriate diet, or infectious causes can lead to dysbacteriosis, gastroenteritis, and ileus and consequently constipation. Underlying causes include anorexia, dental disease, and gastroenteritis.16,61 Abdominal palpation may reveal firm ingesta in the cecum and a tense abdomen. Animals often become anorexic and progressively lethargic. An important differential diagnosis for complete lack of fecal output in chinchillas is intestinal intussusception. General guidelines for the treatment of gastrointestinal disease apply for constipation. The aim is to rehydrate the gut by fluid therapy. Consider enteral fluid therapy (100 mL/kg PO q24h divided in 4-5 doses) in chinchillas with constipation to stimulate a gastrocecal reflex and rehydrate dehydrated ingesta.50,77 Abdominal massage and regular exercise may also be helpful.
DIARRHEA AND SOFT FECES Diarrhea and soft feces are common presentations in chinchillas. Besides infectious causes (e.g., parasites, bacteria), inappropriate feeding or sudden changes in diet with resultant dysbacteriosis commonly cause soft feces. Overfeeding of fresh green feed or items high in simple carbohydrates can result in dysbacteriosis and soft feces. Feces smeared on the cage resting board of a pet with or without matted, fecal-stained perianal fur are often the first signs the owner notices. The animal might be otherwise normal or, in chronic and severe cases, it can be anorexic, dehydrated, and depressed. Rule out infectious causes based on the history and by appropriate diagnostic testing. Intestinal secondary yeast overgrowth, caused by Cyniclomyces guttulatus (previously Saccharomycopsis guttulata), which lines the stomach, is often seen in chinchillas with soft feces.145 However, increased numbers of this yeast in chinchillas with diarrhea or soft feces is considered secondary rather than a cause, usually promoted by an underlying disease process.38 Offer well-dried, high-quality grass hay if the animal is still eating. Consider treatment with nystatin (100,000 IU/kg PO q8h for 5 days) if C. guttulatus overgrowth is very high or no response to initial treatment is seen.
TYMPANY Tympany of the stomach and intestine is less common in chinchillas compared with other rodents or rabbits. Tympany is often secondary to gastroenteritis, dysbacteriosis, ileus or luminal obstruction or, very rarely, intestinal torsion.86 The affected animals usually has a distended and tense abdomen. In severe cases, depressed and dyspneic chinchillas may lie on their sides. Signs of shock may be present. Prognosis depends on the severity and duration of tympany but is usually poor in severe or chronic cases. Institute treatment based on general guidelines for the management of gastroenteritis. Although gastric decompression is recommended for severe cases of tympany, this might result in collapse and death in a decompensated patient. Do not use motility-enhancing drugs (e.g., cisapride) if an infectious or obstructive cause cannot be ruled out.
RECTAL TISSUE PROLAPSE AND INTUSSUSCEPTION Rectal tissue prolapse and intestinal intussusception frequently occur together, secondary to dysbacteriosis, enteritis, constipation, or diarrhea (Fig. 24-1).82,128 Intussusception of the descending colon and rectum is associated with most cases of rectal prolapse; however, the small intestine can also be affected.82,94,128 Abdominal palpation might reveal a turgid cylindrical mass reflecting the intussuscepted portion of the intestine.82 The amount of intestine involved in the intussusception can be extensive and the affected portion is usually cyanotic, congested, and, in advanced cases, often nonviable (see Fig. 24-1, B).82 Besides treatment of the primary underlying cause, surgically correcting the intussusception is critical: intestinal resection and anastomosis may be necessary. Simple replacement or resection of prolapsed rectal tissue is insufficient. Assess the prolapsed tissue for viability and trauma. If an intussusception is ruled out, carefully clean and soak the edematous prolapsed rectal tissue in a concentrated sugar solution (50% dextrose). Replace the prolapsed tissue and place a perianal purse-string suture.94,128 Successful outcome after laparotomy and manual correction of more proximally located intussusceptions has been reported.82 However, the prognosis remains poor in most cases and the outcome will depend on the location and duration of the intussusception, viability of the prolapsed and intussuscepted tissue, and the underlying primary cause. Recurrence and rapid deterioration of affected animals is unfortunately common, since rectal prolapse and intussusception usually reflect an acute complication of a more chronic underlying primary problem.
ESOPHAGEAL DISORDERS Because chinchillas cannot vomit, food items such as raisins, fruit, and nuts as well as bedding material and ingested placentas in postparturient females can become stuck in the oropharynx and upper esophagus.20,38 Clinical signs are a sudden onset of anorexia, drooling, retching, and possible dyspnea. Removal of the foreign material is usually curative and the prognosis is good if the problem is dealt with early.38 Megaesophagus was diagnosed in a 2-year-old chinchilla that presented for recurrent acute episodes of dyspnea and nasal discharge.63 During hospitalization, the animal regurgitated food material episodically, followed by dyspnea, retching, gasping, and nasal discharge.
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B
A
Fig. 24-1 A, Rectal prolapse and intussusception in a 1.5-year-old male chinchilla. The animal presented with a 24-hour history of reduced fecal output and appetite. A firm cylindrical mass, identified as a small intestinal intussusception on necropsy, was palpable in the midabdomen. B, Dissected small intestine of the chinchilla in (A) showing a 4.5-cm section of intussuscepted necrotic jejunum.
Fig. 24-2 Radiograph of a 2-year-old chinchilla 15-minutes after oral administration of barium. The chinchilla presented for recurrent acute episodes of dyspnea and nasal discharge. Megaesophagus was diagnosed based on the dilated and barium-filled thoracic esophagus. Part of the bolus is seen in the stomach. (From Jopp IP, Stengel C, Kraft W. Megaesophagus in a chinchilla (Chinchilla lanigera). A case report. [German] Megaosophagus bei einem Chinchilla (Chinchilla lanigera) Ein Fallbericht. Tierarztl Prax Ausg K Klientiere Heimtiere 2004;32:96-100.)
Upper gastrointestinal contrast radiography aided the diagnosis of megaesophagus in this animal (Fig. 24-2).
DENTAL DISORDERS Dental disease is commonly diagnosed in chinchillas and can affect animals of all ages (see Chapter 32).61 In two studies, abnormalities related to subclinical dental disease were detected in 35%21 and 33%61 of apparently healthy chinchillas presented for routine physical examination. Nutritional (e.g., less abrasive
diet in captivity) and genetic causes have been proposed as the predisposing factors for the development of dental disease in chinchillas.24 Tooth elongation and its secondary complications, affecting the reserve or the clinical crown or both, is the underlying cause for most clinical signs associated with dental disease. Despite sometimes dramatic elongation of reserve and clinical crowns of the cheek teeth, animals often have no difficulty eating and maintain a good body condition until severe changes and complications, such as soft tissue trauma from sharp dental spikes or periodontal infection, have occurred. Common clinical findings associated with dental disease in chinchillas are reduced food intake, changed food preferences toward more easily chewed feed items, weight loss, reduced fecal output with smaller, irregularly shaped fecal pellets, salivastained skin and fur with crusting and alopecia of the perioral area, wetting and crusting of the chin (“slobbers”) and forefeet, epiphora, poor fur condition, and fur chewing.21,61 On clinical examination, palpable irregularities of the ventral borders of the mandible, and overgrown or irregular occlusal surfaces of the incisor teeth may be found.21,61 Facial abscesses of periodontal origin are seen infrequently but can occur.22,61 A limited examination using a pediatric laryngoscope, otoscope, or vaginal speculum can be performed in a conscious animal, but up to 50% of intraoral lesions can be missed.21 Instead, perform a thorough examination of the oral cavity under general anesthesia. Endoscopic-guided intraoral examination (stomatoscopy) provides superior visibility and increases the chance for detection of pathologic lesions (see Chapter 32). Common intraoral findings involving the cheek teeth include coronal elongation, changes of the occlusal surface, formation of sharp spikes buccally and less commonly lingually on the edges of the occlusal surfaces, and widened interproximal coronal spaces that facilitate impaction and promote the development of periodontal disease.22,61 Resorptive and caries-like lesions of the cheek teeth are common in chinchillas; loss of tooth substance or brown discoloration of occlusal and interproximal tooth surfaces is seen.21-23,61 Erosions of the buccal mucosa, gingival hyperplasia, and gingival pocketing are common intraoral findings secondary to dental disease.21,61
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Evaluate radiographs or computed tomography (CT) scans of the skull in any chinchilla with a history or clinical signs suggestive of underlying dental disease. Some clinicians like to use anatomic reference lines for objective interpretation of skull radiographs and staging of dental disease in chinchillas.11 The prognosis for chinchillas diagnosed with dental disease depends on the severity of dental disease, the animal’s general condition, and owner compliance. Repeated intraoral examinations and treatments under general anesthesia at varying intervals, often for the life of the animal, are usually necessary to control complications of dental disease and maintain an acceptable quality of life for the animal. The goals of therapy should include reduction of periodontal infection and soft tissue trauma, which both lead to discomfort and pain, and either recovering or maintaining the animal’s ability to eat unaided. After removing spikes and reducing elongated crowns (see Chapter 32), probe interproximal spaces and gingival pockets and remove impacted debris. Rinse cleaned gingival and periodontal pockets carefully with 2% hydrogen peroxide or diluted chlorhexidine solution.115 Instilling Doxirobe gel (Pfizer Animal Health, New York, NY) in deep (>5 mm) gingival and periodontal pockets may help delay reimpaction with debris and reduce periodontal inflammation.144 If evidence of significant periodontal infection exists, begin systemic antimicrobial therapy with antimicrobials that are effective against anerobic bacteria predominating in periodontal infections (e.g., penicillin G benzathine 50,000 IU/kg SC q5d).133 Limit extraction of cheek teeth to those that are severely diseased and mobile. Removal of cheek teeth may lead to an increased rate of coronal overgrowth of the opposing cheek teeth, owing to the lack of attrition, and can necessitate more frequent coronal crown reductions. Managing pain (e.g., buprenorphine 0.03-0.05 mg/kg SC q8h, meloxicam 0.3-0.5 mg/kg PO, SC q12-24h) and coexisting conditions such as hepatic lipidosis, ketosis, and constipation is critical. With animals in advanced stages of dental disease, offer easy-to-chew food items (soft leafy grass hay, moistened pellets, and “critical-care” formulas offered on a dish). Many animals will require short- and sometimes long-term nutritional support after a dental procedure.
EYE DISORDERS EPIPHORA Epiphora (“wet eye”) is a common condition in chinchillas, usually characterized by unilateral serous discharge, wetting of the periocular fur, and potentially periocular alopecia and dermatitis. Because of secondary bone remodeling, a common underlying cause for epiphora is elongated apical reserve crowns of both the maxillary premolar and first two molar teeth, causing complete or partial compression and consequent obstruction of the nasolacrimal duct.22,25 In contrast to rabbits, obliteration of the nasolacrimal duct at the level of the apical portion of the maxillary incisor is uncommon in chinchillas.16,25,61 Because of the very small size of the lacrimal punctae, visualizing and catheterizing the lacrimal duct for flushing is very difficult and not routinely performed.22 Treat concurrent or underlying infection and inflammation with appropriate topical antibiotics and nonsteroidal anti-inflammatory drugs (see Chapter 37). If underlying dental disease is responsible for the nasolacrimal duct obstruction, it is unlikely that permanent patency of
the nasolacrimal duct will be regained, as there is no effective treatment for apical reserve crown elongation.
CONJUNCTIVITIS Conjunctivitis is common in chinchillas. Irritation from excessive sand bathing, inadequate cage ventilation, or underlying nasolacrimal duct obstruction is often the cause. A variety of bacteria can cause primary bacterial conjunctivitis. Conjunctivitis caused by P. aeruginosa as a localized infection or as part of a systemic infection has been reported.28,138 Depression and anorexia, commonly seen in cases of pseudomonal conjunctivitis, indicates possible underlying systemic infection.28,138 There is controversy as to whether P. aeruginosa is part of the normal conjunctival flora, which consists predominately of gram- positive organisms including Staphylococcus, Streptococcus, and Corynebacterium species.75,85,91 Fluorescein staining is necessary to rule out damage to the corneal surface. Submit a conjunctival swab for bacterial culture and antibiotic susceptibility testing. Treatment includes thoroughly lavaging the conjunctival sac with physiologic saline and applying a topical broad-spectrum antibiotic (e.g., gentamicin) that provides coverage against gram-negative and multiresistant bacteria. Access to a sand bath should be restricted until the chinchilla is fully recovered. Early recognition, systemic antibiotic therapy, and supportive care are critical for chinchillas suffering from systemic Pseudomonas infection.28
CORNEAL DISORDERS Corneal damage and keratitis secondary to trauma are common clinical findings in chinchillas, usually associated with blepharospasm, discharge, and conjunctivitis.35 The large, prominent corneal surface possibly predisposes chinchillas to corneal trauma. Excessive sand bathing, the provision of inappropriate sand for bathing, and inappropriate housing conditions should be considered as possible underlying causes.35,135 Diagnosis is by fluorescein staining of the corneal surface. Avoid direct contact between fluorescein-impregnated test strips and the corneal surface because this can lead to false-positive results.35 Rule out possible nontraumatic underlying causes (e.g., exophthalmos, trichiasis) to avoid reoccurrence. Treatment of acute superficial lesions includes application of antibiotic ophthalmic formulations. In cases of chronic nonhealing ulcers, consider corneal debridement or grid keratomy after controlling any potential bacterial infection.
OTHER EYE DISORDERS Exophthalmos can be caused by a retrobulbar process or increased intraocular pressure. The chinchilla’s orbit is shallow, and manual retraction of the eyelids can cause iatrogenic proptosis of the globe.135 Retrobulbar periapical abscesses of the maxillary cheek teeth—causing exophthalmos—are less common in chinchillas than in guinea pigs or rabbits.21 A rare retrobulbar taenia cyst has been described.56 Primary glaucoma has not been reported in chinchillas. Secondary glaucoma is uncommon but has been reported, and reference intervals for intraocular pressure in chinchillas have been published.85,111 Uveitis and panophthalmitis are uncommon diagnoses in chinchillas and may be caused by trauma or a systemic inflammatory process.135 A rare case of human herpesvirus I infection
CHAPTER 24 Disease Problems of Chinchillas in a chinchilla, causing ulcerative keratitis, retinitis, neuritis and meningoencephalitis, has been reported.140 Lenticular changes such as cataracts, nucleosclerosis, and lens sutures can occur in chinchillas.37,81,85,111 Because diabetogenic cataracts have been reported in chinchillas, diabetes mellitus should be ruled out in animals presenting with uni- or bilateral cataracts.37 Asteroid hyalosis of the vitreous humor, in which lipid- calcium particles are formed as part of a degenerative process, can occur in older chinchillas.111
EAR DISORDERS Chinchillas are used as animal models for human otologic disease research, including hearing loss and otitis media.3,5 Multiple studies on the pathophysiology and antimicrobial treatment of experimentally induced otitis media in chinchillas are available.1,2,4,19,60 Clinical signs caused by otitis externa, media, and interna can range from external ear canal discharge, head shaking, and mild head tilt to facial paresis and severe neurologic deficits.138 Otitis externa is often the result of a perforated tympanic membrane secondary to otitis media; therefore discharge from the ear canal should prompt the clinician to rule out a middle ear infection. Skull radiographs in dorsoventral projection or, preferably, computed tomography (CT) scans are used for the diagnosis of otitis media (Fig. 24-3). Otoscopic examination of the external ear canal and tympanic membrane may reveal purulent exudate and inflammation of the external ear canal. A variety of bacteria may be cultured from the middle ear in chinchillas diagnosed with otitis media. Clinical signs associated with an epizootic outbreak of P. aeruginosa-induced otitis media and interna in a breeding facility were ear discharge, conjunctivitis, neurologic signs, and sudden death due to septicemia.138 Treatment of
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bacterial otitis media and interna remains challenging because of the formation of biofilms in the middle ear that can reduce the efficacy of antimicrobial drugs.3 Bacterial isolation and antimicrobial susceptibility testing are critical. Minimally invasive access to the middle ear through the dorsal roof of the tympanic bulla is a common surgical procedure in chinchillas used for human otologic disease research, and this technique can be used in a clinical setting for sterile sampling of the middle ear.15 Antibiotic selection should be based on susceptibility of the isolated organisms and the potential of the available drugs to eliminate bacteria effectively from the middle ear. Recommendations of fluoroquinolones and chloramphenicol for the treatment of otitis media in chinchillas are based on anecdotal evidence. Azithromycin (30 mg/kg PO q24h) reaches tissue levels high enough to recreate sterile conditions within the middle ear of chinchillas.4,19
RESPIRATORY SYSTEM DISORDERS Primary respiratory disease is uncommon in pet chinchillas. Historically, bacterial pneumonia was an important cause of mortality in ranched chinchillas and is still significant if husbandry is inadequate.8,14,16 Pneumonia is usually bacterial in origin. Important predisposing factors are poor husbandry, such as overcrowding, inadequate ventilation, and poor hygiene. Predominately gram-negative organisms have been isolated from chinchillas diagnosed with pneumonia.7,8,58 Clinical signs that may are tachypnea, dyspnea, and, in severe cases, open-mouth breathing. Presenting animals are often in poor body condition and have a poor hair coat. After ruling out other causes, such as congestive heart failure, initiate systemic antimicrobial therapy. Recommendations for treatment include oxygen therapy, systemic antibiotic therapy, and nebulization with antimicrobials (e.g., gentamicin, tobramycin). Once dyspnea is evident and an animal is in poor body condition, indicative of chronic disease, the prognosis is guarded to poor. Nasal discharge is relatively uncommon in chinchillas but can be associated with underlying dental disease, nasal foreign bodies, rhinitis, or lower respiratory tract disease. Base treatment on the underlying cause. Recurrent pneumonia and mucopurulent nasal discharge were associated with a megaesophagus in a 2-year-old chinchilla (see Fig. 24-2).63
REPRODUCTIVE SYSTEM DISORDERS ENDOMETRITIS AND PYOMETRA
Fig. 24-3 Dorsoventral skull radiograph of a 7-year-old chinchilla that presented for right-sided ear discharge. The changes of increased soft tissue opacity within the thickened bony walls of the right tympanic bulla are consistent with chronic otitis media.
Pet chinchillas may develop endometritis and pyometra. Affected animals can present with a history of acute onset of depression and anorexia or with mild lethargy or behavioral changes.69,136 Clinical signs vary, but an open vulva and vaginal discharge are present in most cases. Vaginal discharge can range from mucoid or mucopurulent to hemorrhagic. Anogenital fur staining may occur. Uterine enlargement or vaginal discharge may be evident on abdominal palpation. Radiographs and abdominal ultrasonography are helpful to evaluate the uterus. Vaginal cytology can be useful to differentiate metritis from physiologic vaginal discharge during estrus.9 Ovariohysterectomy has been recommended as the treatment of choice for endometritis and pyometra.136 In cases of mild endometritis with vaginal discharge, if the animal is in good condition, systemic antimicrobial therapy
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based on culture and susceptibility can be attempted.51 As in other species, prognosis will depend on the underlying and coexisting causes, such as hepatic lipidosis, ketosis, and possible complications in severe cases such as sepsis.
DYSTOCIA Signs of dystocia in chinchillas include restlessness, frequent crying, constant attention to the genital region, and a widened vaginal opening. Allantoic fluid or mucoid-hemorrhagic discharge from the vagina is often seen. Dystocia is usually associated with the presentation of a single oversized fetus or malpresentation of one or more kits.20 Uterine inertia has also been reported as a cause of dystocia.113 Use radiographs to assess the number, size, and position of the fetus(es). In an uncomplicated dystocia, lubrication and gentle traction of the fetus with feline obstetric forceps may correct the condition. Attempt treatment for uterine inertia with oxytocin (0.5-1 IU/kg SQ) and calcium gluconate (25-50 mg/kg SC diluted). Surgical intervention is imperative if the chinchilla is in labor for longer than 4 hours. Pet chinchillas respond well to cesarean section.113 Fetal resorption, mummification, retention, and abortion are common in chinchillas.137 Fetal death may be caused by a variety of infectious and noninfectious causes. Incompletely reabsorbed, retained, or mummified fetuses can remain in the uterus for extended periods and can predispose to sterility and endometritis. The ovarian bursa in chinchillas is underdeveloped and a case of an ectopic pregnancy has been reported.43 Several reports describe pulmonary trophoblastic emboli as incidental findings during postmortem examination in chinchillas; they have no clinical significance.57,132 While neoplasia in chinchillas is rare, several reports describe uterine leiomyosarcomas.
Fig. 24-4 Penile fur ring in an adult single-housed male chinchilla. The fur ring was an incidental finding during a routine physical examination and not associated with any clinical signs or complications.
PENILE DISORDERS Accumulation of hair (“fur ring”), smegma, or both around the base of the glans penis when it is enclosed within the prepuce is common in chinchillas (Fig. 24-4). Fur rings can be found in single-housed as well as breeding males. Male chinchillas that groom excessively, strain to urinate, frequently produce small amounts of urine, or repeatedly clean the penis may have a fur ring.59 The ring of hair can eventually stop the penis from retracting into the prepuce. In severe cases, an engorged penis is seen protruding from the prepuce, resulting in paraphimosis.59 The condition is not only painful but also may cause urethral constriction and acute urinary blockage. Treatment includes carefully removing any fur or debris from the penis. Topical ointments or gels are applied to prevent drying of the exposed penis and everted prepuce. For cases with underlying infection (Fig. 24-5), use systemic antimicrobial treatment. Complete resolution of infection and swelling is possible with topical and systemic treatment, but it may require long-term attention. Always examine the penis of male chinchillas during routine physical examination and remove any accumulated fur or secretions.
URINARY SYSTEM DISORDERS Male chinchillas can develop urinary calculi and urolithiasis.62,125 Quantitative mineral analysis of uroliths retrieved from 73 chinchillas showed that calculi were composed of calcium carbonate in approximately 90% of animals and of other
Fig. 24-5 Preputial abscess in an adult male chinchilla that presented for preputial swelling. The abscess is 2.5 cm in diameter. Despite the abscess, the animal maintained normal micturition.
material in 10%.108 Affected animals present with hematuria, stranguria, or anuria. Abdominal radiographs show radiodense calculi in the bladder and/or urethra. Treatment consists of surgical removal of the calculi if feasible. Submit swab samples of urine or bladder lining taken at surgery for bacterial culture and susceptibility testing to rule out underlying urinary tract infection. Because the underlying cause of urolith formation in chinchillas in unknown, recurrence of uroliths within a few weeks to months after surgical removal is possible. Currently there
CHAPTER 24 Disease Problems of Chinchillas are no specific recommendations for preventing reoccurrence of uroliths in affected chinchillas. Anecdotal recommendations include increased diuresis and a calcium-restricted diet. Spontaneous oxalate nephrosis in six female chinchillas has been reported.46 No access to ethylene glycol (antifreeze) had occurred and the underlying cause for the renal tubular oxalate crystal deposition could not be determined.
NEUROLOGIC DISORDERS SEIZURES Several reports of seizures in chinchillas are described.36,38,51,129,140 Encephalitis, septicemia, toxicosis, dietary deficiencies, hypocalcemia, hypoglycemia, hepatic or renal insufficiency, and heat stroke are described causing either generalized or focal convulsions. Hypersalivation, lateral recumbency, and nonresponsiveness are often seen.36,38 Although uncommon in pet chinchillas, consider infectious causes of encephalitis, such as listeriosis, human herpes simplex virus, or cerebrospinal nematodiasis (see below).119,140 Rule out extracranial causes, such as hypocalcemia, hyperthermia, hypoglycemia, organ failure, and lead poisoning.
HEAT STROKE The ambient temperature range to which chinchillas are adapted is 65°F to 80°F (18.3°C-26.7°C) in a low-humidity environment. Exposure to higher ambient temperatures, especially in the presence of high humidity and poor ventilation, can result in heat stroke. Affected animals are recumbent or ataxic, exhibit rapid breathing, and have bright red mucous membranes, prominent ear vessels, and thick, stringy saliva. Treatment includes cooling the animal and, if the animal is in shock, administering intravenous fluids. Long-term prognosis is guarded to poor and animals often deteriorate after initial improvement.51
LEAD POISONING Lead poisoning is uncommon in chinchillas and incidence varies based on geographic location. Two cases of lead toxicosis in pet chinchillas have been reported, one of which presented for seizures.53,96 Blood lead concentrations of 25 mg/dL or higher are indicative of lead poisoning. Successful treatment with calcium disodium edetate (30 mg/kg SC q12h) has been reported.53
DERMATOLOGIC DISORDERS DERMATOPHYTOSIS Dermatophytosis (ringworm) in chinchillas is most commonly caused by Trichophyton mentagrophytes, although Microsporum canis and Microsporum gypseum have been incriminated in outbreaks of spontaneously occurring dermatophytosis.29 Infected chinchillas may have small, scaly patches of alopecia on the nose, behind the ears, or on the forefeet. Lesions may appear on any part of the body; in advanced cases, a large circumscribed area of inflammation with scab formation is typical. Although most mycologic studies of chinchillas are based on animals with clinical signs, T. mentagrophytes has been
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cultured from 5% of fur-ranched chinchillas with normal skin and 30% of those with fur damage.29 Because T. mentagrophytes does not fluoresce, ultraviolet light is not useful for diagnosis. Diagnosis requires microscopic examination of hair and skin samples and dermatophyte culture. For topical therapy, 2% chlorhexidine/2% miconazole shampoo, or 0.2% enilconazole rinse is effective. Topical treatment removes spores from hair shafts, and systemic treatment acts at hair follicles. Systemic drugs that can be used are terbinafine (20-40 mg/kg PO q24h), itraconazole (5-10 mg/kg PO q24h), or ketoconazole (10-15 mg/kg PO q24h).90 Terbinafine is more effective than itraconazole against Trichophyton species in rodents.90 Continue treatment until two negative dermatophyte cultures have been obtained.
FUR CHEWING Fur chewing can be a common problem in chinchillas; up to 20% of animals in breeding facilities can be affected (see Fig. 22-8). Fur chewing is also commonly seen in chinchillas suffering from dental disease.61,131 Many theories concerning the cause of fur chewing have been proposed, including dietary deficiencies, fungal and endoparasite infections, hormonal dysregulation, environmental stress, boredom, and genetic predisposition.33,131,134 Scant scientific evidence exists for most proposed theories. Currently, the most widely accepted theory suggests that fur chewing is a maladapted displacement behavior triggered by stress and affecting predominately stress-sensitive animals. Adrenocortical hyperplasia and histologic changes of the skin, correlating with increased cortisol secretion, support this theory.131 Clinical signs vary and affected animals may chew their own or their cagemate’s fur, resulting in a coat with a moth-eaten appearance. Fur-chewed areas occur usually along the midspinal area from the lumbar part to the tail. In animals that chew their own fur, the head and distal extremities are usually not affected and the chewed areas are usually covered with short, darker-colored fur.131 Perform a thorough history and physical examination to rule out dietary deficiencies, dental disease, and possible environmental stressors. A variety of dermatologic diagnostic tests can be considered, including dermatophyte culture and skin biopsy, although dermatophytes can be cultured from animals without fur lesions and do not necessarily represent the primary underlying cause of fur damage.33 Obtaining a definitive diagnosis and identifying the primary underlying cause of fur chewing behavior may be difficult. While fur chewing can be annoying for the pet chinchilla owner, it is not a significant threat to the animal’s health. Consider environmental and dietary improvements after ruling out infectious and organic underlying causes. This may include reducing possible stressors, such as frequent handling and light and noise disturbance. Carefully review social dynamics if housing multiple animals together. Avoid overcrowding, and separate affected animals from dominant or aggressive cage mates. Offer multiple sleeping boxes and feeding spots for animals housed in groups. Recommend environmental enrichment such as providing high-quality grass hay and branches for chewing and foraging as well as a structured enclosure and regular exercise. Treatment with antidepressants such as fluoxetine hydrochloride (5 to 10 mg/kg PO q24h) has been suggested for other rodents, but no clinical results have been reported in chinchillas.116
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FUR SLIP Fur slip is a predator avoidance mechanism in chinchillas. When an animal is fighting or roughly handled, it can release a large patch of fur, thus enabling it to escape. A clean, smooth area of skin is left; hair may require several months to regrow.
MATTED FUR Matted fur can develop in chinchillas, especially if they are kept in a warm (>80°F [26.7°C]), humid environment or if they are deprived of dust baths (see Chapter 22). Provide dust baths for approximately 15 minutes per day and reduce humidity levels if necessary.
FOOT DISORDERS In chinchillas, foot disorders predominately affect the hind feet. Lesions can include hyperkeratosis and erythema; less commonly, deep infections or open lesions of the plantar aspect of the feet can occur. Possible predisposing factors are obesity, inappropriate cage flooring or substrate, and poor hygiene. In severe cases, infection can involve the tendons and bones. Treatment depends on the severity of disease. In mild cases, environmental improvements and application of petroleum-based ointment are often sufficient to resolve hyperkeratosis and erythema. Advise owners to reduce body weight gradually in overweight animals. In severe cases, perform surgical debridement followed by open-wound management and bandaging until healing is complete.
MISCELLANEOUS DISEASE PROBLEMS CARDIAC DISEASE Heart murmurs ranging from mild to moderate are often auscultated, particularly in young chinchillas presented for routine examination.55 To date, reports of cardiac disease in chinchillas are scant and the significance of heart murmurs in young, clinically healthy animals remains unknown. Anecdotally, cardiomyopathy, ventricular septal defect, as well as mitral and tricuspid valve insufficiencies have been reported.55,76 Use echocardiography to differentiate innocent from pathologic murmurs. Echocardiographic reference values for chinchillas have been published.76 At present, the clinical management of heart disease in chinchillas is empiric.
HEPATIC LIPIDOSIS AND KETOSIS Hepatic lipidosis because of excessive fat mobilization and, less frequently, toxicosis is one of the most common findings in chinchillas at necropsy.32,112 Chinchillas with hepatic lipidosis often have a history of anorexia and decreased fecal output and, in advanced cases, can be depressed and dehydrated. Hyperglycemia, pronounced ketonuria, and possible glucosuria and acidosis can develop.92 The negative energy balance in hyporectic or anorectic chinchillas leads to increased mobilization of lipids and excessive beta oxidation of fatty acids in hepatocytes, promoting hepatic ketogenesis. Treatment should correct the underlying primary
cause leading to anorexia (e.g., dental disease, gastroenteritis), along with correcting fluid deficits and providing nutritional support. Measure urinary ketones repeatedly as a simple and noninvasive tool to monitor the response to treatment.
DIABETES MELLITUS Diabetes mellitus is rare in chinchillas. Treatment is difficult and the prognosis is poor. Only two cases (suggestive of type 2 diabetes) have been reported.37,81 When presented with an anorexic and hyperglycemic chinchilla, rule out common causes for anorexia (e.g., dental disease, pain). Many chinchillas will become ketoacidotic and hyperglycemic secondary to anorexia. Determine serum insulin levels to confirm a diagnosis of diabetes mellitus caused by endocrine pancreatic insufficiency. Hyperglycemia secondary to peripheral insulin resistance (type 2), as seen in cases of ketoacidosis, should be treated by focusing on the underlying primary cause of anorexia and providing nutritional support. In two cases, a 5-year-old obese female81 and a 1-year-old male37 had blood glucose levels elevated to at least four times the upper limit of the reference range, as well as severe glucosuria and ketonuria. Both animals were treated with insulin (2 IU daily and increasing to 12 IU daily81 and 4 IU daily increasing to 6 IU daily37). After initial improvement, both animals died. Postmortem examination showed atrophy of the pancreas and vacuolation of the islets of Langerhans in the obese female chinchilla and a pancreatic adenoma in the 1-year-old male. Hypoglycemia is always a great risk in treating diabetes with recombinant human insulin or porcine insulin.100,107 Consequently any attempt to treat diabetes with insulin must be conservative and dramatic increases in the insulin dose should be avoided. Ancillary treatment should be aimed at reducing obesity and feeding a diet that is high in protein, low in fat, and high in complex carbohydrates.
FRACTURES Traumatic fractures of the tibia are commonly seen. The tibia is a long straight bone with little soft tissue covering. It is longer than the femur, and the fibula is virtually nonexistent.18 Tibial fractures are usually either transverse or short spiral fractures. Tibial fracture often occurs when a chinchilla is grabbed by its hind limb or a hind limb catches in a cage bar. Like the bones of rabbits, those of chinchillas are thin and fragile; surgical repair can be difficult and complications are common (see Chapter 33). Soft, padded bandages and lateral splints usually do not provide adequate stability for tibial fractures to heal.54 External fixation and intramedullary pins, alone or in combination, have been recommended for surgical stabilization of tibial fractures in chinchillas.38 Restricted exercise in a single-level enclosure, ideally without cage bars, is necessary. The prognosis for tibial fractures is guarded and complications after surgical fixation are common. These include bone-pin loosening and infection, nonunion, necrosis of the distal limb, and automutilation.38 Consider hind-limb amputation if surgical fracture stabilization fails or is not indicated. Chinchillas usually adapt very well after amputation.26,70,130 Fractures
CHAPTER 24 Disease Problems of Chinchillas of the fore limbs distal to the elbow can be managed by external cooptation and splinting; chinchillas usually tolerate such treatment well.
NEOPLASIA Despite the long life span of chinchillas compared with other rodents, references to neoplasia in chinchillas are rare. Postmortem examinations on 1,005 fur-ranched chinchillas before 194913 and another 1,000 fur-ranched chinchillas between 1949 and 195214 ranging in age between less than 6 months and 11 years did not list neoplasia as a cause of death. However, tumors such as neuroblastoma, carcinoma, lipoma, and hemangioma were reported in fur-farmed chinchillas in the annual reports of the San Diego County Livestock Department during the 1950s.101 Individual case reports of neoplasia have included two cases of lymphosarcoma.65,101 Both cases occurred in young male animals (18 months and 4.5 years) and both animals had generalized lymphadenopathy and neoplastic cell infiltration in multiple organs. Four other reports described a nonmetastasizing cholangiohepatic carcinoma in a 3-year-old female chinchilla102; a fibrosarcoma located at the tail base of a 6-year-old male chinchilla38; a nonmetastasizing lumbar osteosarcoma in a 13-year-old female chinchilla123; and an undifferentiated carcinoma of the salivary gland in a 12-year-old female chinchilla.124 A 5-year retrospective from 1991 to 1996 of chinchillas presented to the Animal Medical Center in New York City revealed only one case of a uterine leiomyosarcoma with no associated metastases. A 35-year retrospective evaluation of chinchilla presentations to the University of Tennessee College of Veterinary Medicine showed only two histopathologically documented cases of neoplasia: one was lymphosarcoma and the other adenocarcinoma of the lung.47 A 9-year retrospective between 1990 and 1999 of chinchillas submitted for necropsy at the Institute of Pathology and Forensic Veterinary Medicine in Vienna, Austria, documented a low incidence of neoplasia in chinchillas (2.6%) compared with rabbits, guinea pigs, rats, and mice.39 Three of 115 chinchillas had tumors. The cases included a 2-year-old female with uterine leiomyoma; a 3-year-old male with hemangioma of the subcutis; and a castrated 10-year-old male with an adenoma of the pituitary gland. A second 9-year retrospective between 1994 and 2003 of 325 chinchillas presented to the University of Zurich Veterinary Hospital revealed tumors in only three animals (1%). During the same period, the incidence of neoplasia was higher in rabbits and rodents compared with chinchillas (guinea pigs 7%, rats 34%, and rabbits 6%).72 The low number of reports of neoplasia up to the present may reflect the emphasis on chinchillas as fur producers or research animals, since chinchillas were bred for their fur or kept as laboratory animals before they became popular as pets. Therefore ‘‘geriatric’’ conditions, including neoplasia, may not have been represented in the older reports. Alternatively, the 5- to 35-year surveys of neoplasia in exotic pets presented to veterinary specialty centers suggest that the low incidence of neoplasia in chinchillas may reflect a true low frequency compared with the incidence in other rodents and rabbits. Further clinical surveys are needed to elucidate this question.
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INFECTIOUS DISEASES VIRAL INFECTIONS Diagnoses of viral infections in chinchillas are rare. During several epizootic disease outbreaks in ranched chinchillas, a viral etiology was suspected but diagnosis could not be established.20,38 Chinchillas are susceptible to human herpesvirus type 1 and may play a role as a temporary reservoir for human infections. A spontaneous herpes-like viral infection was reported in a female chinchilla.45 In a 1-year-old male chinchilla with a 2-week history of conjunctivitis and subsequent neurologic signs of seizures, disorientation, recumbency, and apathy, histologic examination showed a nonsuppurative meningitis and polioencephalitis with neuronal necrosis and intranuclear inclusion bodies; these data were confirmed as indicating herpesvirus type 1.140 Both eyes displayed ulcerative keratitis, uveitis, retinitis and retinal degeneration, and optical neuritis. The clinical signs, the distribution of the lesions, and the viral antigen suggested a primary ocular infection with subsequent spread to the central nervous system.
FUNGAL INFECTIONS There are two case reports of Histoplasma capsulatum infection in chinchillas. One case was in a chinchilla imported from the United States to Switzerland.17 The second case was in a female chinchilla that originated from a commercial chinchilla ranch in central Missouri.109 At necropsy, pulmonary lesions included multiple hemorrhagic foci, alveolar consolidation, and bronchopneumonia; the organism was found to be present in numerous giant cells. Multifocal pyogranulomatous splenitis and hepatitis, with H. capsulatum in giant cells, was also noted. Histoplasma capsulatum was subsequently cultured from timothy hay used for food. Cyniclomyces guttulatus is part of the physiologic intestinal flora in chinchillas.145 Animals with soft feces or diarrhea often show increased numbers of this yeast. This finding is indicative of dysbacteriosis, which leads to opportunistic overgrowth of C. guttulatus.38,51 Treatment should address the primary underlying cause of the dysbacteriosis. In severe cases of yeast overgrowth, consider treatment with nystatin (100,000 IU/kg PO q8h for 5 days). Aflatoxicosis is an acute, fatal disease resulting from toxicity from improperly stored feed contaminated with Aspergillus fungi. In one report, the death of 200 chinchillas was attributed to high concentrations of aflatoxin B-1 in the feed.112 The liver is the primary target organ of aflatoxin; in affected animals, it is enlarged, pale yellow, and friable. In the described cases, histopathologic analyses of hepatic parenchyma showed severe, diffuse cytoplasmic vacuolation of hepatocytes.112 Because the histopathologic changes caused by acute aflatoxicosis are nonspecific, the diagnosis of aflatoxicosis is usually made in combination with mycotoxicologic feed analysis.112
PARASITIC INFECTIONS Protozoal Infections Historically, group-housed chinchillas in fur ranches and research colonies had a high prevalence of giardiasis.122 However, the role of Giardia duodenalis (syn. G. lamblia) in
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causing disease in chinchillas is difficult to establish. Giardia are rarely found in fecal samples from wild chinchillas, but Giardia is found in both healthy and sick captive-bred animals.33,40 In pet chinchillas, G. duodenalis is highly prevalent. In 80 pet chinchillas from both pet owners and breeders in Belgium screened for Giardia, 53 (66%) excreted Giardia cysts. Young animals and animals participating in shows were at significantly higher risk of infection.74 A total of 22 isolates based on genetic groups (or assemblages) were characterized. Healthy chinchillas can harbor G. duodenalis organisms in low numbers in the small intestine, and experimental infection of healthy chinchillas with Giardia cysts failed to induce clinical disease.33,38 Predisposing factors, such as stress and poor husbandry, are believed to cause an increase in parasite numbers, resulting in diarrhea and potentially death. Recently weaned juvenile animals seem to be prone to developing clinical signs.38,74 Signs of giardiasis in pet chinchillas can include a cyclic sequence of appetite loss and diarrhea associated with a declining body and fur condition. Zinc sulfate flotation allows identification of cysts, and trophozoites can be identified in fresh fecal smears in acute cases of heavy infection. The use of an antigen enzyme-linked immunosorbent assay (ELISA) (Remel ProSpecT microplate assay; Thermo Fisher Scientific, Lexana, KS) for detecting G. duodenalis antigen in chinchilla feces has recently been reported.110 However, the sensitivity and specificity for this test in chinchillas has not been determined, and because clinically normal chinchillas can also shed G. duodenalis in the feces, the significance of a positive ELISA is of questionable clinical significance.33,40 Quantifying fecal cyst or trophozoite numbers in animals with diarrhea appears to be more clinically relevant to determine if an infection is related to the clinical signs. Assemblage specific polymerase chain reaction (PCR) (e.g., A-G) is clinically and epidemiologically beneficial, as mixed genetic group infections are likely to be missed using conventional PCR approaches.74 Treat chinchillas with giardiasis with metronidazole, albendazole, or fenbendazole. It is unknown if these compounds eradicate Giardia completely or only inhibit cyst production; therefore treated animals may remain a source of chronic cyst shedding. Treat all animals in contact and thoroughly disinfect the environment to prevent reinfection. Replace wooden cage interior parts, such as resting boards. Giardia cysts remain infectious for up to several weeks in a cool, humid environment. Until further research in chinchillas is done, the pathogenicity of G. duodenalis and its association with disease remain controversial. Giardiainfected chinchillas are a potential reservoir for zoonotic transmission.64,74,121 Eimeria chinchillae is strictly host-specific and can occasionally cause enteritis and diarrhea predominately in younger animals. Stress likely leads to clinical manifestation of a previous subclinical infection. This coccidium causes damage of the intestinal mucosa, with subsequent disturbance of the physiologic flora and possible secondary dysbacteriosis. Diagnosis is made by fecal flotation. Treatment with sulfonamide compounds (e.g., sulfadimethoxine) usually resolves clinical signs and cyst shedding. It is unknown if these compounds are able to eradicate E. chinchillae. As for giardiasis, treatment of contact animals and thorough environmental disinfection is mandatory.
Although toxoplasmosis was common in fur-ranched chinchillas in the past, it is now rarely seen.66 At necropsy, lesions include an enlarged spleen and mesenteric lymph nodes as well as hemorrhagic lungs. In two chinchillas with focal necrotic meningoencephalitis caused by Toxoplasma gondii,87 several lobulated Frenkelia cysts up to 0.6 mm in diameter were found in the brains, independent of and remote from the toxoplasmal inflammatory reaction. The authors considered that chinchillas might be susceptible to a Frenkelia species occurring in other free-living species. Tissue cysts of Frenkelia microti were found in the brain of a chinchilla bred in Minnesota and used for biomedical research.30 This was the first report of Frenkelia infection in chinchillas in the United States. A case of acute hepatic sarcocystosis was reported in a pet female chinchilla.114 The owners had housed and fed it with six other chinchillas that remained healthy, and the source of infection was unknown. Gastroenteritis associated with Cryptosporidium species was described in an 8-month-old pet chinchilla that originated from a pet shop.143 Meningitis and optic nerve neuritis due to Cryptococcus species was reported in a 13-year-old chinchilla.10
Helminthic Infections Recent studies report a low prevalence of nematode and cestode infections in pet chinchillas.110 Haemonchus contortus, Trichostrongylus colubriformis, Ostertagia ostertagi, and an unspecified Oxyuris species have been reported in chinchillas.110,127 Disease outbreaks of cerebral nematodiasis caused by the raccoon ascarid Baylisascaris procyonis have been reported in chinchillas from western Canada.119 Affected chinchillas showed ataxia, torticollis, paralysis, incoordination, and tumbling. Outbreaks of fatal central nervous system disease were linked to use of hay contaminated by raccoon feces. Raccoons infected with B. procyonis are more common in temperate regions of North America, especially the midwestern and northeastern United States. Cestodes. Rodentolepis nana (previously Hymenolepis nana) infections are reported in chinchillas.44,105,127 This cestode does not require an intermediate host and infection can occur by direct transmission via fecal-oral route.120 Animals infected by high numbers of R. nana can show anorexia, diarrhea, weight loss, and death,98 but subclinical infections are more common. Rodentolepis nana is zoonotic and can cause severe infections, particularly in immunocompromised humans.31,106 Demonstrate Rodentolepis eggs by fecal flotation. Treat with praziquantel (5 mg/kg PO or SQ q10d). Chinchillas can serve as intermediate hosts for cestodes, including Taenia serialis, Taenia pisiformis, Taenia multiceps, Echinococcus granulosus, and Echinococcus multilocularis.44,56,126 In the 1950s, infections were seen when chinchillas were given feed accidentally contaminated with infected dog feces. A recent report described more than 600 cysts of Taenia crassiceps found in the abdominal cavity of a pet chinchilla imported from the Netherlands into Japan.71
BACTERIAL INFECTIONS Opportunistic bacterial infections in chinchillas can cause disease, which is localized either to one organ or as septicemia. Affected animals are usually immunocompromised by age, underlying disease, nutritional status, or husbandry-related factors (poor hygiene, poor ventilation, contaminated feed).
CHAPTER 24 Disease Problems of Chinchillas Members of the family Enterobacteriaceae and P. aeruginosa have been associated with significant morbidity and mortality in chinchillas.* However, Enterobacteriaceae and P. aeruginosa can also be isolated from clinically healthy animals.12,83,91 Therefore most of these organisms are not considered primary pathogens. Pseudomonas aeruginosa infections and epizootic outbreaks in chinchillas have been reported frequently.49,79,89,138 Initially, infections are often localized to one organ and are associated with conjunctivitis, enteritis, pneumonia, otitis media and interna, metritis, and abortion. As the disease progresses, systemic spread is common. In addition, an acute generalized form with septicemia and often sudden death can occur. P. aeruginosa can be part of the normal intestinal flora in healthy chinchillas. A survey of 67 healthy pet or laboratory chinchillas isolated the organism from 42% of animals.52 Stress, intercurrent disease and/or or contaminated drinking water predispose to infection and clinical disease.28,83,91,138 Conjunctivitis is a common initial sign of Pseudomonas infection in chinchillas. Anorexia, lethargy, and decreased fecal output often follow.27,28,138 In a case of P. aeruginosa infection described in a laboratory chinchilla, the affected animal displayed a variety of clinical signs, including conjunctivitis, scrotal swelling, anorexia, weight loss, and corneal and oral ulcerations.28 Characteristic pathologic lesions are miliary necrosis in the internal parenchymal organs and a necrotizing typhlocolitis. Multidrug-resistant, reduced-antibiotic susceptibility and highly virulent strains of P. aeruginosa are widespread in chinchillas.52 Base antimicrobial drug selection on culture and susceptibility testing. Because affected animals are often in a critically compromised condition, empiric drug selection is necessary. Generally, P. aeruginosa is susceptible to fluoroquinolones, third-generation cephalosporins, and aminoglycosides.117,118 Use topical polymyxin B and gentamicin-containing formulations to treat because of the low prevalence of isolates that are resistant to these drugs. A vaccine against P. aeruginosa has been developed for attempted immunization and is used in fur-ranched chinchillas.79,84 Escherichia coli is not considered part of the physiologic intestinal flora of healthy captive chinchillas but has been isolated from healthy wild-caught chinchillas.12,83 It is a ubiquitous environmental organism and an opportunistic pathogen. The ingestion of large numbers of organisms, a primary underlying disease, or predisposing risk factors are prerequisites to infection and clinical disease. Pathogenicity depends on the serotype, enteropathogenicity, and endotoxin production. Clinical signs associated with E. coli-associated enteritis can very be similar to enteritis caused by P. aeruginosa, such as constipation, diarrhea, variable form and size of fecal pellets, mucoid or hemorrhagic adhesions to fecal pellets, depression, and anorexia. Sudden death due to endotoxemia or septicemia is possible.12 In a study of chinchillas experimentally infected with enteropathogenic E. coli via drinking water, animals became increasingly depressed, exhibited a stretched body posture, initially had diarrhea, and later had reduced fecal output and anorexia.12 In chinchillas that died, E. coli was cultured from all internal organs and the animals
*References 7, 8, 12, 28, 88, 138.
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had suffered from gastroenteritis. Pure growth of E. coli was seen in fecal cultures 4 to 7 days after the start of the experiment. Parenteral treatment with antibiotics caused a significant improvement of clinical signs and disappearance of E. coli from fecal cultures.12 Yersinia pseudotuberculosis (formerly Pasteurella pseudotuberculosis) and Yersinia enterocolitica occur worldwide in areas of moderate and subtropical climate, and outbreaks in chinchillas are commonly described.34,41,42,48,73 The species most frequently isolated from chinchillas is Y. enterocolitica. Yersiniosis is an enteric disease that damages epithelium of the ileum, cecum, and colon, resulting in mucosal hemorrhage and ulceration. Lymphoid infiltration results in hypertrophy of Peyer’s patches and mesenteric lymph nodes and necrotizing granulomas. Systemic spread results in granulomatous lesions in the lungs, spleen, and liver and eventual death. A “chinchillatype” strain of Y. enterocolitica (biovar 3, antigens or serovar 1, 2a, 3) appears to persist enzootically among chinchilla stock worldwide.141 Listeriosis in chinchillas was first reported by MacKay et al. in 1949.80 It was and still is common in fur-ranched chinchillas but not in laboratory or pet chinchillas.67,68,103,139 Listeria monocytogenes is a highly adaptable environmental bacterium that can exist as both an animal pathogen and a plant saprophyte; it is also part of the normal microbial flora in healthy ruminants and is found in environmental sources such as decaying vegetation. Most animal and human cases of listeriosis arise from the ingestion of contaminated food; the disease is common in animals fed on silage.78 Unlike most food-borne pathogens that cause gastrointestinal disease, L. monocytogenes causes several easily recognized invasive syndromes, such as encephalitis, abortion, and septicemia. In chinchillas, listeriosis is a cecal disease with blood-borne dissemination. The main target organ is the liver, where the bacteria multiply inside hepatocytes, followed by cell lysis, bacterial release, septicemia, and, in surviving hosts, the development of lung, brain, spleen, lymph node, and liver abscesses. Enterotoxaemia associated with Clostridium species in chinchillas has been reported.95 Enterotoxemia caused by Clostridium perfringens enterotoxin has been described as well as deaths due to C. perfringens A enterotoxemia.6,104 Salmonellosis characterized by gastroenteritis and abortion has been reported often from ranched chinchillas; it causes significant morbidity and mortality.44,93,99 There are two case reports of Salmonella infection in pet chinchillas. Salmonella arizona septicemia was reported in a chinchilla in the United Kingdom97 and septic infection with Salmonella enteritidis was described in a companion chinchilla in Japan.142
References 1. Alper CM, Doyle WJ, Seroky JT, et al. Efficacy of clarithromycin treatment of acute otitis media caused by infection with penicillin-susceptible, -intermediate, and -resistant Streptococcus pneumoniae in the chinchilla. Antimicrob Agents Chemother. 1996;40:1889-1892. 2. Antonelli PJ, Winterstein AG, Schultz GS. Topical dexamethasone and tympanic membrane perforation healing in otitis media: a short-term study. Otol Neurotol. 2010;31:519-523. 3. Apicella MA. Bacterial otitis media, the chinchilla middle ear, and biofilms. J Infect Dis. 2009;199:774-775.
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4. Babl FE, Pelton SI, Li Z. Experimental acute otitis media due to nontypeable Haemophilus influenzae: comparison of high and low azithromycin doses with placebo. Antimicrob Agents Chemother. 2002;46:2194-2199. 5. Bakaletz LO. Chinchilla as a robust, reproducible and polymicrobial model of otitis media and its prevention. Expert Rev Vaccines. 2009;8:1063-1082. 6. Bartoszce M, Nowakowska M, Roszkowski J, et al. Chinchilla deaths due to Clostridium perfringens A enterotoxin. Vet Rec. 1990;126:341-342. 7. Bartoszcze M, Matras J, Palec S, et al. Klebsiella pneumoniae infection in chinchillas [letter]. Vet Rec. 1990;127:119. 8. Bautista E, Martino P, Manacorda A, et al. Spontaneous Proteus mirabilis and Enterobacter aerogenes infection in chinchilla (Chinchilla lanigera). Scientifur. 2007;31:27-30. 9. Bekyürek T, Liman N, Bayram G. Diagnosis of sexual cycle by means of vaginal smear method in the chinchilla (Chinchilla lanigera). Lab Anim. 2002;36:51-60. 10. Bicknese E, White A, Pessier A, et al. Cryptococcal meningitis and optic nerve neuritis in a chinchilla (Chinchilla lanigera). Proceedings. Annu Conf Assoc Exot Mammal Vet. 2010:Session #165. 11. Boehmer E, Crossley D. Objective interpretation of dental disease in rabbits, guinea pigs and chinchillas. Use of anatomical reference lines. Tierarztl Prax Ausg K Klientiere Heimtiere. 2009;37:250-260. 12. Brem M. Investigations about the diseases of the gastrointestinal tract in the Chinchilla. [German] Untersuchungen über Erkrankungen des Magen-Darmkanals beim Chinchilla. Medizinische Tierklinik. Ludwig-Maximilians Universität: Munich; 1982;125. 13. Brenon HC. Postmortem examinations of chinchillas. J Am Vet Med Assoc. 1953;123:310. 14. Brenon HC. Postmortem examinations of chinchillas. J Am Vet Med Assoc. 1955;126:222-223. 15. Brown C. Middle ear sample collection in the chinchilla. Lab Anim (NY). 2007;36:22-23. 16. Burtscher H. Pathological anatomy of chinchilla diseases. [German] Pathologische Anatomie der Chinchillakrankheiten. Dtsch Tierarztl Wochenschr. 1965;72:376-380. 17. Burtscher H, Otte E. Histoplasma in the chinchilla. [German] Histoplasme beim chinchilla. Dtsch Tierarztl Wochenschr. 1962;69:303-307. 18. Cevik-Demirkan A, Ozdemir V, Turkmenoglu I, et al. Anatomy of the hind limb skeleton of the chinchilla (Chinchilla lanigera). Acta Veterinaria Brno. 2007;76:501-507. 19. Chan KH, Swarts JD, Doyle WJ, et al. Efficacy of a new macrolide (azithromycin). For acute otitis media in the chinchilla model. Arch Otolaryngol Head Neck Surg. 1988;114:1266-1269. 20. Cousens PJ. The chinchilla in veterinary practice. J Small Anim Pract. 1963;4:199-205. 21. Crossley DA. Dental disease in chinchillas in the UK. J Small Anim Pract. 2001;42:12-19. 22. Crossley DA. Dental disease in chinchillas [dissertation]. Manchester, UK: School of Dentistry, University of Manchester; 2003;263. 23. Crossley DA, Dubielzig RR, Benson KG. Caries and odontoclastic resorptive lesions in a chinchilla (Chinchilla lanigera). Vet Rec. 1997;141:337-339. 24. Crossley DA, Miguélez MM. Skull size and cheek-tooth length in wild-caught and captive-bred chinchillas. Arch Oral Biol. 2001;46:919-928. 25. Crossley DA, Roxburg G, Miguélez Vidales MM. Anatomy of the chinchilla (Chinchilla lanigera) lacrimal drainage system and its obstruction in dental disease. Proceedings. 8th Europ Vet Dental Soc. 1999:21-22. 26. Degasperi B, Mosing M, Kunzel F. Limb amputation as salvage procedure in small mammals. [German]. Extremitatenamputation als rettende Massnahme bei kleinen Heimtieren. Wien Tierarztl Monatsschr. 2007;94:93-98.
27. Devos A, Van Impe J, Viaene N, et al. Pseudomonas aeruginosa infection in chinchillas. [Dutch]. Vlaams Diergeneesk Tijdschr. 1966;35:222-232. 28. Doerning BJ, Brammer DW, Rush HG. Pseudomonas aeruginosa infection in a Chinchilla lanigera. Lab Anim. 1993; 27:131-133. 29. Donnelly TM, Rush EM, Lackner PA. Ringworm in small exotic pets. Sem Avian Exot Pet Med. 2000;9:82-93. 30. Dubey JP, Clark TR, Yantis D. Frenkelia microti infection in a chinchilla (Chinchilla laniger) in the United States. J Parasitol. 2000;86:1149-1150. 31. Duclos LM, Richardson DJ. Hymenolepis nana in pet store rodents. Comp Parasitol. 2000;67:197-201. 32. Egri B, Egri J, Hajnovics B. Fatty liver in young male chinchilla (Chinchilla velligera). [German] Uber Fettinfiltration der Leber beim Chinchillabockchen (Chinchilla velligera). Tierarztl Umsch. 1994;49:42, 45-47. 33. Eidmann S. Studies on the etiology and pathogenesis of fur damage in the chinchilla. [German] Untersuchungen zur Atiologie und Pathogenese von Fellscaden beim Chinchilla. Institut fur Pathologie. Hannover, Germany: Tierarztliche Hochschule; 1992;163. 34. Emirsajlow Zalewska W, Furowicz AJ, Aleksic S, et al. Evaluation of pathogenicity determinants of Yersinia pseudotuberculosis strains isolated from chinchillas from the Western Pomerania area. Advances in Agricultural Sciences. 1996;5:19-24. 35. Eule C, Sjoberg JG. Corneal injuries in chinchillas. Exot DVM. 2007;9:7-8. 36. Ewringmann A, Gloeckner B. Neurological deficits. [German] Neurologische Ausfallerscheinungen. In: Ewringmann A, Gloeckner B, eds. Leitsymptome be Meerschweinchen, Chinchilla und Degu. Stuttgart: Enke; 2005:166-185. 37. Ewringmann A, Gobel T. Diabetes mellitus in rabbits, guinea pigs and chinchillas. [German] Diabetes mellitus bei Kaninchen, Meerschweinchen und Chinchilla. Kleintierpraxis. 1998;43:337-348. 38. Fehr M. Chinchilla. In: Fehr M, Sassenburg L, Zwart P, eds. Krankheiten der Heimtiere. 6th ed. Hannover: Schluetersche; 2005:183-213. 39. Ferfschl S. Spontaneous tumors in horses, cattle and small pets: findings of the institute of pathology and forensic veterinary medicine in the years 1990-1999. [German] Spontane Tumoren bei Pferden, Rindern und kleinen Heimtieren: Untersuchungsergebnisse des Instituts für Pathologie und Gerichtliche Veterinärmedizin in den Jahren 1990-1999. Institut für Tierzucht und Genetik. Vienna: Veterinaermedizinische Universitaet Wien; 2004;117. 40. Fialho CG, Oliveira RG, Teixeira MC, et al. Comparison of protozoan infection between chinchillas (Chinchilla lanigera) from a commercial breeding facility in southern Brazil and chinchillas from a natural reserve in Chile. [Portuguese] Comparação da infecção por protozoários em chinchila (Chinchilla lanigera) de uma criação comercial do município de ViamãoRS, Brasil, e de chinchilas no seu habitat natural, Chile. Parasitol Latinoam. 2008;63:85-87. 41. Furowicz AJ, Czernomysy-Furowicz D. Eradication of Yersinia pseudotuberculosis infection from a chinchilla farm. [Polish] Eliminacja jersiniozy (Yersinia pseudotuberculosis) z hodowli szynszyli. Magazyn Weterynaryjny. 1999;8:130-132. 42. Furowicz AJ, Czernomysy-Furowicz D, Misiura M, et al. Yersiniosis in chinchillas caused by the enterotoxigenic strain of Yersinia pseudotuberculosis. [Polish] Enterotoksyczny szczep Yersinia pseudotuberculosis przyczyna jersiniozy szynszyli. Medycyna Weterynaryjna. 1996;52:116-118. 43. Gitlin G, Adler JH. Coexisting intrauterine and abdominal (intraperitoneal) pregnancy with possible superfoetation (superfecundation) and with adhesion of placenta to foetus in a Chinchilla (Chinchilla laniger). Acta Zool Pathol Antverp. 1969;49:65-76.
CHAPTER 24 Disease Problems of Chinchillas 44. Gorham JR, Farrell K. Diseases and parasites of chinchillas. Proceedings. 92nd Ann Meet Am Vet Med Assoc. 1955:228-234. 45. Goudas P, Giltoy JS. Spontaneous herpes-like viral infection in a chinchilla (Chinchilla laniger). J Wildl Dis. 1970;6:175-179. 46. Goudas P, Lusis P. Case report. Oxalate nephrosis in chinchilla (Chinchilla laniger). Can Vet J. 1970;11:256-257. 47. Greenacre CB. Spontaneous tumors of small mammals. Vet Clin North Am Exot Anim Pract. 2004;7:627-651. 48. Gueraud JM. Threat of an epidemic of yersiniosis in chinchillas. [French] Vers une epizootie de yersiniose chez le chinchilla. Bull Acad Vet Fr. 1988;61:95-98. 49. Halen P, Pohl P, Thomas J. Septicaemia caused by Pseudomonas in mink and chinchillas. [French]. Septicémies à pseudomonas chez les visons et chinchillas. Ann Med Vet. 1966;110:397-406. 50. Hallowell GD. Retrospective study assessing efficacy of treatment of large colonic impactions. Equine Vet J. 2008; 40:411-413. 51. Hansen D. Chinchilla. In: Goebel T, Ewringmann A, eds. Heimtierkrankheiten. Stutgart: Verlag Eugen Ulmer; 2005:100-118. 52. Hirakawa Y, Sasaki H, Kawamoto E, et al. Prevalence and analysis of Pseudomonas aeruginosa in chinchillas. BMC Vet Res. 2010;6:52. 53. Hoefer HL. Chinchillas. Vet Clin North Am Small Anim Pract. 1994;24:103-111. 54. Hoefer HL. Chinchillas. Proceedings. North Am Vet Conf. 1995:672-673. 55. Hoefer HL, Crossley DA. Chinchillas. In: Meredith A, Redrobe S, eds. BSAVA Manual of Exotic Pets. 4th ed. Quedgeley, Gloucester: British Small Animal Veterinary Association; 2002:65-75. 56. Holmberg BJ, Hollingsworth SR, Osofsky A, et al. Taenia coenurus in the orbit of a chinchilla. Vet Ophthalmol. 2007;10:53-59. 57. Ilha MRdS, Bezerra Junior PS, Sanches AWD, et al. Trophoblastic pulmonary embolism in chinchillas (Chinchilla laniger). [Portuguese] Embolia pulmonar trofoblastica em chinchilas (Chinchilla laniger). Ciencia Rural. 2000;30:903-904. 58. Ioakimidis I, Tomopoulos D, Vlaikidis N. An outbreak of Bordetella bronchiseptica infection in chinchillas (Chinchilla laniger). Hellenike Kteniatrike. 1970;13:31-33. 59. Ivey ES, Hoefer HL. What’s Your Diagnosis? Pollakiuria in a chinchilla. Lab Anim (NY). 1998;27:21-22. 60. Jauris-Heipke S, Leake ER, Billy JM, et al. The effect of antibiotic treatment on the release of endotoxin during nontypable Haemophilus influenzae-induced otitis media in the chinchilla. Acta Otolaryngol. 1997;117:109-112. 61. Jekl V, Hauptman K, Knotek Z. Quantitative and qualitative assessments of intraoral lesions in 180 small herbivorous mammals. Vet Rec. 2008;162:442-449. 62. Jones RJ, Stephenson R, Fountain D, et al. Urolithiasis in a chinchilla [letter]. Vet Rec. 1995;136:400. 63. Jopp IP, Stengel C, Kraft W. Megaesophagus in a chinchilla (Chinchilla lanigera). A case report. [German] Megaosophagus bei einem Chinchilla (Chinchilla lanigera) Ein Fallbericht. Tierarztl Prax Ausg K Klientiere Heimtiere. 2004;32:96-100. 64. Karanis P, Ey PL. Characterization of axenic isolates of Giardia intestinalis established from humans and animals in Germany. Parasitol Res. 1998;84:442-449. 65. Kast A. Leucosis in chinchillas. [German] Leukose beim Chinchilla. Berl Munch Tierarztl Wochenschr. 1962;75:414-415. 66. Keagy HF. Toxoplasma in the chinchilla. J Am Vet Med Assoc. 1949;94:15. 67. Kimpe A, Decostere A, Hermans K, et al. Isolation of Listeria ivanovii from a septicaemic chinchilla (Chinchilla lanigera). Vet Rec. 2004;154:791-792. 68. Kirinus JK, Krewer C, Zeni D, et al. Outbreak of systemic listeriosis in chinchillas. [Portuguese] Surto de listeriose sistêmica em chinchilas. Ciencia Rural. 2010;40:686-689.
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69. Kottwitz J. Stump pyometra in a chinchilla. Exot DVM. 2006;8:24-28. 70. Kottwitz J, Underwood JG. Midfemoral pelvic limb amputation in a chinchilla. Exot DVM. 2005;7:31-34. 71. Kugi G, Nonaka N, Ganzorig S, et al. Taenia crassiceps larvae in a chinchilla (Chinchilla brevicaudata). [Japanese]. J Vet Med. 1999;52:449-452. 72. Langenecker M. Development of species composition and health problems in exotic pets between 1994 and 2003 [German]. Retrospektive Untersuchung zur Entwicklung der Artenverteilung und den haufigten Krankheitsbildern bei exotischen Heimtieren im Zeitraum von 1994-2003. Division of Zoo Animals Exotic Pets and Wildlife (Departement für Kleintiere, Abteilung für Zoo-, Heim- und Wildtiere). Zurich: Vetsuisse Faculty, University of Zurich (Vetsuisse-Fakultät Universität Zürich); 2006. 73. Lazzari AM, Vargas ACd, Dutra V, et al. Infectious agents isolated from Chinchilla laniger. [Portuguese] Agentes infecciosos isolados de Chinchilla laniger. Ciencia Rural. 2001;31:337-340. 74. Levecke B, Meulemans L, Dalemans T, et al. Mixed Giardia duodenalis assemblage A, B, C and E infections in pet chinchillas (Chinchilla lanigera) in Flanders (Belgium). Vet Parasitol. 2010:[Epub ahead of print]. 75. Lima L, Montiani-Ferreira F, Tramontin M, et al. The chinchilla eye: morphologic observations, echobiometric findings and reference values for selected ophthalmic diagnostic tests. Vet Ophthalmol. 2010;13:14-25. 76. Linde A, Summerfield NJ, Johnston M, et al. Echocardiography in the chinchilla. J Vet Intern Med. 2004;18:772-774. 77. Lopes MAF, White NA, Donaldson L, et al. Effects of enteral and intravenous fluid therapy, magnesium sulfate, and sodium sulfate on colonic contents and feces in horses. Am J Vet Res. 2004;65:695-704. 78. Low JC, Donachie W. A review of Listeria monocytogenes and listeriosis. Vet J. 1997;153:9-29. 79. Lusis PI, Soltys MA. Immunization of mice and chinchillas against Pseudomonas aeruginosa. Can J Comp Med. 1971;35:60-66. 80. MacKay KA, Kennedy AH, Smith DLT, et al. Listeria monocytogenes infection in chinchillas. Annual Report of the Ontario Veterinary College. 1949:137-145. 81. Marlow C. Diabetes in a chinchilla [letter]. Vet Rec. 1995; 136:595-596. 82. Marlow CHB. An outbreak of intussusception in a herd of chinchillas. J S Afr Vet Med Assoc. 1963;34:637-642. 83. Mathieu X, Duran JC, Rivas M. Normal bacterial flora of the wild Chinchilla lanigera Silvestre. [Spanish] Estudio de la flora bacteriana normal de Chinchilla lanigera Silvestre. Rev Latinoam Microbiol. 1982;24:77-82. 84. Matthes S. Vaccination of rabbits and fur animals. [German] Schutzimpfungen bei Kaninchen und Pelztieren. Tierarztl Prax. 1985;13:107-112. 85. Mauler D, Lubke-Becker A, Eule C. Ocular findings in healthy chinchillidae. Abstracts, Annu Meet Europ Col Vet Ophthal/ Europ Soc Vet Ophthal 2009 (Abstract 31). Vet Ophthalmol. 2009;12:387. 86. McGreevy PD, Carn VM. Intestinal torsion in a chinchilla [letter]. Vet Rec. 1988;122:287. 87. Meingassner JG, Burtscher H. Double infection of the brain with Frenkelia species and Toxoplasma gondii in Chinchilla laniger. [German] Doppelinfektion des Gehirns mit Frenkelia species und Toxoplasma gondii bei Chinchilla laniger. Vet Pathol. 1977;14:146-153. 88. Menchaca ES, Martin AM, Moras EV, et al. Infectious diseases of the chinchilla. III. Proteus mirabilis and Proteus vulgaris infection. [Spanish] Enfermedades infecciosas de la chinchilla (Chinchilla lanigera). III. “Proteus mirabilis y Proteus vulgaris”. Gaceta Veterinaria. 1978;40:651-656.
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SECTION III Guinea Pigs and Chinchillas
89. Menchaca ES, Moras EV, Martin AM, et al. Infectious diseases of the chinchilla. IV. Pseudomonas aeruginosa. [Spanish] Enfermedades infecciosas de la chinchilla (Chinchilla laniger). IV. “Pseudomona aeruginosa”. Gaceta Veterinaria. 1980;42:96-102. 90. Mieth H, Leitner I, Meingassner JG. The efficacy of orally applied terbinafine, itraconazole and fluconazole in models of experimental trichophytoses. J Med Vet Mycol. 1994;32:181-188. 91. Miller LG, Finegold SM. Normal bacterial flora of the chinchilla [abstract]. Bacteriological Proceeding. Ann Meet Am Soc Microbiol. 1967;67:66. 92. Mills R, Gilsdorf J. Middle-ear effusions following acute otitis media in the chinchilla animal model. J Laryngol Otol. 1986;100:255-261. 93. Misirlioglu D, Cetin C, Kahraman MM, et al. Salmonella infection in a chinchilla farm. [Turkish] Bir cincila ciftliginde salmonella enfeksiyonu. Turk J Vet Anim Sci. 2002;26:151-155. 94. Misirlioglu D, Ozmen O, Cangul IT, et al. A case of rectum prolapsus and intestinal invagination in a male chinchilla. J Turk Vet Surg. 2000;6:51-53. 95. Moore RW, Greenlee HH. Enterotoxaemia in chinchillas. Lab Anim. 1975;9:153-154. 96. Morgan RV, Pearce LK, Moore FM, et al. Demographic data and treatment of small companion animals with lead poisoning: 347 cases (1977-1986). J Am Vet Med Assoc. 1991;199:98-102. 97. Mountain A. Salmonella arizona in a chinchilla [letter]. Vet Rec. 1989;125:25. 98. Muller M, Haas H, Vogel A, et al. Mass outbreak of Hymenolepis nana in chinchillas. [German] Massenbefall mit dem Zwergbandwurm Hymenolepis nana beim Chinchilla. Tierarztl Umsch. 2010;65:17-20. 99. Naglic T, Seol B, Bedekovic M, et al. Outbreak of Salmonella enteritidis and isolation of Salmonella sofia in chinchillas (Chinchilla laniger). Vet Rec. 2003;152:719-720. 100. Neville RWJ, Weir BJ, Lazarus NR. Hystricomorph insulins. Symposia of the Zoological Society of London. 1974;34:417-433. 101. Newberne PM, Seibold HR. Malignant lymphoma in a chinchilla. Vet Med. 1953;48:428-429. 102. Nobel TA, Neumann F. Carcinoma of the liver in a nutria (Myocaster coypus) and a chinchilla (Chinchilla laniger). Refuah Veterinarith. 1963;20:161-162. 103. Novak S, Ruttkay D, Solar I. Results of screening for bacterial diseases on large-scale chinchilla (Chinchilla laniger) farms. [Slovakian] Vysledky depistaze bakterialnych ochoreni vo vel’kochovoch cincily vlnatej (Chinchilla laniger). Slov Vet Cas. 1994;19:19-21. 104. Nowakowska M, Matras J, Bartoszcze M, et al. Clostridium perfringens enterotoxaemia in chinchillas. [Polish] Zatrucie pokarmowe szynszyli wywolane przez enterotoksyne Clostridium perfringens. Medycyna Weterynaryjna. 1991;47:156-157. 105. Olsen OW. Natural infection of chinchillas with the mouse tapeworm, Hymenolepis nana var. fraterna. Vet Med. 1950; 45:440-442. 106. Olson PD, Yoder K, Fajardo LF, et al. Lethal invasive cestodiasis in immunosuppressed patients. J Infect Dis. 2003; 187:1962-1966. 107. Opazo JC, Soto-Gamboa M, Bozinovic F. Blood glucose concentration in caviomorph rodents. Comp Biochem Physiol A Mol Integr Physiol. 2004;137:57-64. 108. Osborne CA, Albasan H, Lulich JP, et al. Quantitative analysis of 4468 uroliths retrieved from farm animals, exotic species, and wildlife submitted to the Minnesota Urolith Center: 1981 to 2007. Vet Clin North Am Small Anim Pract. 2009;39:65-78. 109. Owens DR, Menges RW, Sprouse RF, et al. Naturally occurring histoplasmosis in the chinchilla (Chinchilla laniger). J Clin Microbiol. 1975;1:486-488.
110. Pantchev N, Globokar-Vrhovec M, Beck W. Endoparasites from indoor kept small mammals and hedgehogs. Laboratory evaluation of fecal, serological, and urinary samples (2002-2004). [German] Endoparasitosen bei Kleinsaugern aus privater Haltung und Igeln Labordiagnostische Befunde der koprologischen, serologischen und Urinuntersuchung (2002-2004). Tierarztl Prax Ausg K Klientiere Heimtiere. 2005;33:296-306. 111. Peiffer RL, Johnson PT. Clinical ocular findings in a colony of chinchillas (Chinchilla laniger). Lab Anim. 1980;14:331-335. 112. Pereyra MLG, Carvalho EC, Tissera JL, et al. An outbreak of acute aflatoxicosis on a chinchilla (Chinchilla lanigera) farm in Argentina. J Vet Diagn Invest. 2008;20:853-856. 113. Prior JE. Caesarian section in the chinchilla. Vet Rec. 1986; 119:408. 114. Rakich PM, Dubey JP, Contarino JK. Acute hepatic sarcocystosis in a chinchilla. J Vet Diagn Invest. 1992;4:484-486. 115. Rams TE, Slots J. Local delivery of antimicrobial agents in the periodontal pocket. Periodontol 2000. 1996;10:139-159. 116. Rossoff IS. Handbook of veterinary drugs and chemicals: a compendium for research and clinical use. 2nd ed. Taylorville, Pharmatox Publishing Company; 1994. 117. Rubin J, Walker RD, Blickenstaff K, et al. Antimicrobial resistance and genetic characterization of fluoroquinolone resistance of Pseudomonas aeruginosa isolated from canine infections. Vet Microbiol. 2008;131:164-172. 118. Saitou K, Furuhata K, Kawakami Y, et al. Isolation of Pseudomonas aeruginosa from cockroaches captured in hospitals in Japan, and their antibiotic susceptibility. Biocontrol Sci. 2009;14:155-159. 119. Sanford SE. Cerebrospinal nematodiasis caused by Baylisascaris procyonis in chinchillas. J Vet Diagn Invest. 1991;3:77-79. 120. Schantz PM. Tapeworms (cestodiasis). Gastroenterol Clin North Am. 1996;25:637-653. 121. Schönball U. Case report: Giardiasis in a chinchilla—possible source of infection for human being? [German] Fallbericht: Gardiaeineninfektion bei einem Chinchilla—mögliche Infektionsquelle für der Menschen? Kleintierpraxis. 1992; 37:785-788. 122. Shelton GC. Giardiasis in the chinchilla. II. Incidence of the disease and results of experimental infections. Am J Vet Res. 1954;15:75-78. 123. Simova-Curd S, Nitzl D, Pospischil A, et al. Lumbar osteosarcoma in a chinchilla (Chinchilla laniger). J Small Anim Pract. 2008;49:483-485. 124. Smith JL, Campbell-Ward M, Else RW, et al. Undifferentiated carcinoma of the salivary gland in a chinchilla (Chinchilla lanigera). J Vet Diagn Invest. 2010;22:152-155. 125. Spence S, Skae K. Urolithiasis in a chinchilla [letter]. Vet Rec. 1995;136:524. 126. Staebler S, Steinmetz H, Keller S, et al. First description of natural Echinococcus multilocularis infections in chinchilla (Chinchilla laniger) and Prevost’s squirrel (Callosciurus prevostii borneoensis). Parasitol Res. 2007;101:1725-1727. 127. Stampa S, Hobson NK. Control of some internal parasites of chinchillas. J Am Vet Med Assoc. 1966;149:929-932. 128. Stoebe W. Rectal prolapse in chinchilla. [German] Zur Behandlung des Darmvorfalls bei der Chinchilla. Tierarztl Umsch. 1965;20:79. 129. Strake GJ, Davis LA, LaRegina M, et al. Chinchillas. In: LaberLaird K, Swindle MM, Flecknell P, eds. Handbook of rodent and rabbit medicine. Tarrytown: Elsevier; 1996:151-171. 130. Thompson L. Amputation of hindlimbs in chinchillas as a salvage procedure. Proceedings. British Vet Zoological Soc Spring Meet: Non-infectious Diseases of Mammals. 2003. Dublin University. 131. Tisljar M, Janic D, Grabarevic Z, et al. Stress-induced Cushing’s syndrome in fur-chewing chinchillas. Acta Vet Hung. 2002;50:133-142.
CHAPTER 24 Disease Problems of Chinchillas 132. Tvedten HW, Langham RF. Trophoblastic emboli in a chinchilla. J Am Vet Med Assoc. 1974;165:828-829. 133. Tyrrell KL, Citron DM, Jenkins JR, et al. Periodontal bacteria in rabbit mandibular and maxillary abscesses. J Clin Microbiol. 2002;40:1044-1047. 134. Vanjonack WJ, Johnson HD. Relationship of thyroid and adrenal function to “fur-chewing” in the chinchilla. Comp Biochem Physiol A. 1973;45:115-120. 135. Wagner F, Fehr M. Eye diseases of chinchilla (Chinchilla lanigera): Anatomical and physiological characteristics and disorders. [German] Augenerkrankungen beim Chinchilla (Chinchilla lanigera)—anatomische, physiologische Besonderheiten, Erkrankungen. Kleintierpraxis. 2008;53:309-318. 136. Ward ML, Morrison LR, Else RW, et al. Endometritis in the chinchilla: 3 Cases (2003-2006). In: Proceedings. British Small Anim Vet Congress Belfast. 2007. 137. Weir BJ. Aspects of reproduction in chinchillas. J Reprod Fertil. 1966;12:410-411. 138. Wideman WL. Pseudomonas aeruginosa otitis media and interna in a chinchilla ranch. Can Vet J. 2006;47:799-800.
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139. Wilkerson MJ, Melendy A, Stauber E. An outbreak of listeriosis in a breeding colony of chinchillas. J Vet Diagn Invest. 1997;9:320-323. 140. Wohlsein P, Thiele A, Fehr M, et al. Spontaneous human herpes virus type 1 infection in a chinchilla (Chinchilla lanigera f. dom.). Acta Neuropathol. 2002;104:674-678. 141. Wuthe HH, Aleksic S. Yersinia enterocolitica serovar 1,2a,3 biovar-3 in chinchillas. Zentralbl Bakteriol. 1992;277:403-405. 142. Yamagishi S, Watanabe Y, Tomura H, et al. Septic infection of a companion chinchilla with Salmonella enteritidis. [Japanese]. Nippon Juishikai Zasshi. 1997;50:345-348. 143. Yamini B, Raju NR. Gastroenteritis associated with a Cryptosporidium sp in a chinchilla. J Am Vet Med Assoc. 1986; 189:1158-1159. 144. Zetner K, Rothmueller G. Treatment of periodontal pockets with doxycycline in beagles. Vet Ther. 2002;3:441-452. 145. Zierdt CH, Detlefson C, Muller J, et al. Cyniclomyces guttulatus (Saccharomycopsis guttulata)—culture, ultrastructure and physiology. Antonie Van Leeuwenhoek. 1988;54:357-366.
CHAPTER
25
Soft Tissue Surgery
R. Avery Bennett, DVM, MS, Diplomate ACVS
Guinea Pigs, Chinchillas, and Degus Ovariectomy Ovariohysterectomy Pyometra Uterine Torsion Dystocia Uterine Prolapse Uterine Tumors Mammary Gland Neoplasia Orchidectomy Penile Prolapse Gastric Trichobezoars Urolithiasis Cutaneous Dermal Masses Cervical Lymphadenitis Thoracotomy Miscellaneous Procedures
Both guinea pigs and chinchillas are hystricomorph members of the order Rodentia and have similar biology, anatomy, physiology, disease susceptibility, and surgical conditions. Degus are also hystricomorph rodents. The natural history of rodents has a significant impact on their ability to survive surgical procedures. Rodents are shy and fearful prey species. With fear, pain, and stress, they lose the will to live and may die for no apparent reason. They may appear to recover from anesthesia and surgery, only to die at home 2 or 3 days later. This seems to be especially true of hystricomorph rodents. It appears that the more human contact the pet is used to receiving, the better it bears the stresses of illness and surgery; those that are frequently held, played with, and coddled are more likely to survive than those that spend all their time in a cage. It is important to bear this fact in mind in discussing options and prognosis with owners. Pet rodents that are stressed may stop eating to the point of starvation. Hystricomorph rodents ferment cellulose in the cecum; if they become anorectic, life-threatening complications 326
can occur. Some patients struggle violently, injuring themselves. Some become so frightened that their levels of circulating catecholamines become very high, which can negatively affect anesthesia, and some die acutely from these high levels of catecholamines. This concern underscores the need for appropriate pre- and postoperative analgesic, antianxiety, and tranquilizing agents. If an animal seems stressed by the hospital environment and surgery is not urgent, it may be best to hospitalize it for a day or more, allowing it to adjust to the new environment before surgery. In some cases it may be best to perform multiple short procedures rather than several procedures at once, which would require a long period of anesthesia. Because these herbivorous rodents eat frequently, a short fast of 1 to 2 hours is generally recommended, only to allow the animal to clear its mouth of food material. Herbivores are physiologically unable to vomit, so the risk of aspiration pneumonia is negligible. Because normal gastrointestinal function is vital to recovery, a long fast is not recommended. It has been shown that small mammals with a negative energy balance are at greater risk for postoperative complications.19 Parenteral fluid administration is recommended for most procedures, as it is in other animals. Vascular access can be difficult to obtain in these small patients; however, obtaining vascular access through an intravenous catheter or an intraosseous cannula allows for the administration of fluids during anesthesia and also provides a means whereby emergency drugs can be administered if necessary. If it is possible to maintain vascular access postoperatively, continue administering fluids at a maintenance level until the patient is eating and drinking normally. The reported total blood volume of small mammals is 57 mL/kg body weight.15,19 With loss of 15% to 20% of the total blood volume, most mammals experience hypovolemic shock and release high levels of catecholamines. Life-threatening consequences usually occur with loss of 20% to 30% of the total blood volume.15,19 This would be equivalent to only 4.5 to 6.8 mL of blood in a 400-g guinea pig. Crystalloid, colloid, whole blood from a conspecific, or a blood substitute can be used in patients experiencing serious blood loss. Guinea pigs, chinchillas, and degus seem to be less likely to bother surgical incisions than are other species of rodents. In selecting suture material, keep in mind the propensity of these Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 25 Soft Tissue Surgery animals to develop a caseous, suppurative response to foreign materials such as sutures. Catgut is degraded by proteolysis and should not be used in rodents because of its reactive nature. Absorbable materials degraded by hydrolysis rather than proteolysis are recommended (e.g., polyglactin 910, polyglycolic acid, polydioxanone, poliglecaprone, glycomer 631, braided lactomer 9-1). Soft, absorbable, braided materials (e.g., poly glycolic acid, polyglactin 910, braided lactomer 9-1) are rapidly absorbed and are less irritating to subcutaneous tissues than stiffer materials such as monofilament absorbable sutures (e.g., polydioxanone, poliglecaprone, glycomer 631). Some surgeons recommend stainless steel for cutaneous sutures; however, this material is very stiff and the cut ends often cause irritation, actually stimulating rather than preventing self-mutilation. Many rodents chew out even steel skin sutures. Skin closure is best accomplished with an intradermal or subcuticular technique. This can be time-consuming if the surgeon is not adept. Skin staples are quickly applied, and most rodents will not bother them because there are no ends to poke and irritate the adjacent skin. Cyanoacrylate tissue adhesive can be used to close small incisions; however, excessive amounts of the adhesive on the skin surface will often attract the attention of the rodent and it will try to groom it off, potentially resulting in dehiscence. If the patient demonstrates a tendency toward self-mutilation, midazolam (0.5-2 mg/kg SC, IM q4-6h) may help modulate the behavior, allowing appropriate recovery and healing. This treatment may be needed for only a day or two until the patient becomes accustomed to having a surgical wound. Most patients that have proper postoperative analgesia do not bother their surgical wound initially, though they may begin to pay attention to it during later stages of healing if it becomes pruritic. Small suture size and use of a small-gauge needle are also important. Suture sizes 4-0 to 7-0 are most commonly used in these species of pet rodents. Hemostatic clips are valuable for controlling hemorrhage, especially because of the small size of these patients and because the vessels are often in locations that are difficult to access. Guinea pig skin is relatively thicker than that of chinchillas. Chinchilla skin is fragile and can easily be damaged during clipping for aseptic surgery, the fur is also very fine and epilates easily. Use a number 50 clipper blade (Oster Professional Products, McMinnville, TN) because the teeth are closer together than those of a No. 40 blade, minimizing the risk of the skin being caught between the teeth and getting nicked. Also, to help minimize the risk of tearing, flatten the skin in front of the clipper blade, move slowly over the skin, and clean and lubricate the clipper blades frequently. Tearing of the skin is not as much of a problem in guinea pigs or degus. For skin preparation, a standard surgical scrub is recommended, alternating with warm sterile saline-soaked gauze (instead of alcohol) to minimize evaporative cooling, because these species are prone to hypothermia. The soaked gauze can be warmed in a microwave before use, being careful not to overheat it. Intraoperatively, monitor the patient’s body temperature closely and provide supplemental heat with a circulating warmwater blanket, thermal pad (Thermally Controlled Surgery Pad, RICA Surgical Products, Schiller Park, IL; Hot Dog Patient Warming, Augustine Biomedical Design, Eden Prairie, MN), a forced warm-air blanket, a radiant heat lamp, or some combination of these.
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GUINEA PIGS, CHINCHILLAS, AND DEGUS OVARIECTOMY Cystic ovaries are common in guinea pigs, with an incidence reported to be as high as 100%.33,34,37 In a study of guinea pigs with reproductive problems, 76% had cystic ovaries.21 In another study of 43 guinea pigs from various owners or breeders, 53% had evidence of cystic ovaries on ultrasound examination and 36% had evidence of bilateral cystic ovaries.28 Of the guinea pigs above 2 years of age, 93% had cystic ovaries, 62% of which were bilateral. There was a direct relationship between age and the size of the ovarian cysts and there was no difference in incidence between breeding and nonbreeding sows.28 There was no difference in reproductive success in sows with or without cysts younger than 15 months of age; however, guinea pigs greater than 15 months of age with cysts had a marked decrease in reproductive success.21 While cystic ovaries are common, most sows show little evidence of clinical disease. In juvenile sows, they are small (5 μm), but they enlarge as the animal matures, reaching up to 7 cm in size.21,33 If the cysts are <5 mm, there are usually multiple cysts; if they are >2 cm, they are usually single cysts.34 The largest are often multilocular. Histologically, ovarian cysts in guinea pigs are similar to those of humans and cats, being cysts of the rete ovarii.21,33,34,37 Rete cells are from the mesonephros; they migrate into the fetal gonads and differentiate into rete testis in males and rete ovarii in females. They form a tubular, blind-ended vestigial structure within the ovary. The function of the rete cells is suspected to be phagocytosis of degenerating oocysts and production of a meiosis-inducing substance. These cells do not produce hormones. The pathogenesis of cyst formation is unknown but is suspected to be the result of a defect in ion pumps, so electrolytes are transported into the tubular structure but not out. Fluid is pulled in as the ion concentration increases, resulting in the formation of cysts. Since these cysts do not produce hormones, patients are usually asymptomatic. Clinical signs include abdominal distention as the cysts enlarge; they may cause anorexia and depression because of the effects of the space-occupying mass. Many sows with ovarian cysts also have uterine disease and may present with a hemorrhagic vaginal discharge or reported hematuria that is actually a result of uterine hemorrhage.9,21 Some guinea pigs have a nonpruritic symmetric alopecia typical of an endocrine alopecia; they become aggressive and begin mounting cage mates9; however, it is considered normal for female guinea pigs to mount cage mates during estrus. These signs are indicative of hormone increases that are difficult to explain, since the cysts do not produce hormones. The diagnosis is confirmed with ultrasound.3 Radiographically, ovarian cysts cannot be differentiated from other abdominal masses because fluid is of the same radiographic density as soft tissues. Ovariohysterectomy is the treatment of choice for guinea pigs with cystic ovaries because uterine disease secondary to these ovarian cysts is common, although a mechanism has not been established. In a study of five guinea pigs with cystic ovaries histologically confirmed to be rete ovarii cysts, all were found to have uterine disease, including endometritis, pyometra, endometrial hyperplasia, and leiomyomas.9,10 If the uterine changes are secondary to the ovarian cysts, ovariectomy at a young age would be expected to prevent both problems.37 Because ovarian rete cysts are so common and become
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SECTION III Guinea Pigs and Chinchillas
larger with age, routine ovariectomy or ovariohysterectomy at a young age should be recommended to owners of female guinea pigs. Drainage of ovarian cysts in guinea pigs provides temporary relief, but fluid quickly reforms. Prior to surgery, drain the cysts as well as possible, using percutaneous centesis to decrease their size. Be aware that there are often adhesions between the diseased ovaries and uterus to the body wall and other viscera. In guinea pigs with alopecia, hair regrowth is generally complete within 3 months after surgery.3,9 Rete ovarii cysts do not produce hormones and should not be affected by the administration of gonadotropin-releasing hormone or its analogs, such as human chorionic gonadotropin.5,13,37 Those hormones are effective in treating follicular cysts, causing them to luteinize. Reports of cysts other than cystic rete ovarii are rare, but a granulosa cell tumor was successfully managed by ovariohysterectomy in a guinea pig.5 Still, administration of 100 IU (1,000 USP units) of human chorionic gonadotropin administered intramuscularly in two doses given 2 weeks apart has been reported to resolve the clinical signs temporarily.5 Gonadotropin-releasing hormone has also been reported to be “very effective” in treating guinea pigs with ovarian cysts.25 Ovarian teratomas have also been reported to occur commonly in sows older than 3 years of age.8,11,13 Some teratomas may be as large as 10 cm in diameter.40 They are usually unilateral and rarely metastasize. Affected sows present for depression, weakness, or collapse due to spontaneous intra-abdominal hemorrhage from the tumor. Acute death from blood loss can occur. Ovariectomy or ovariohysterectomy is the treatment of choice for sows with these tumors. Ovariectomy of young rodents may decrease the incidence of mammary neoplasia; however, this has not been documented in hystricomorph rodents. Ovariectomy can be done through a ventral midline ap‑ proach or a dorsolateral approach.35 The ventral midline approach is as described for other rodents (see Chapter 28). To perform an ovariectomy using the dorsolateral approach, make a 1- to 2-cm incision on each side ventral to the erector spinae muscle and about 1 cm caudal to the last rib. Bluntly penetrate the muscle with a hemostat and enlarge the opening to about 1 cm. With pressure on the abdomen to push the ovary to the incision, reach in with forceps and grasp the ovary. Exteriorize the ovary and use a hemostatic clip or ligature to control any hemorrhage from the ovarian vessels that might occur. Also be sure to remove the entire oviduct surrounding the ovary to prevent cysts from forming. Close the opening in the body wall with 1 or 2 sutures of monofilament absorbable material and appose the skin edges with tissue adhesive or an intradermal suture. No advantage has been documented for performing ovariohysterectomy over ovariectomy unless there is concurrent uterine disease and, in fact, most complications associated with ovariohysterectomy result from removing the uterus.42 The advantage to performing ovariectomy is that the incisions are small and dorsal and the delicate gastrointestinal tract is not disturbed, resulting in less morbidity and a more rapid recovery.
OVARIOHYSTERECTOMY Indications for ovariohysterectomy in hystricomorph rodents include dystocia, uterine prolapse, pyometra, and uterine and/ or ovarian masses. The ovaries are located caudolateral to the kidneys and are approximately 8 mm in length and 5 mm in
a
b
c
Fig. 25-1 The ovary (a) in this guinea pig is located at that caudal pole of the kidney. The uterine horns (b) are long and come together to form the uterine body (c).
width (Fig. 25-1).19 The oviduct lies in close proximity to the dorsal aspect of the ovary, encircling it before joining the uterine horn.31 The uterus is bicornuate and the horns join to form a uterine body, which is divided internally by a well-developed intercornual ligament; however, there is a single cervical os. The mesovarium, mesometrium, and broad ligaments are sites of fat storage in guinea pigs and chinchillas, adding to the difficulty of the procedure. The ovarian artery and vein are branches off the renal vessels that split into an ovarian branch supplying the ovary and a uterine branch to the uterus.31 There is a single artery and vein medial to the ovaries and along the uterus to the uterine body. Once the patient has been anesthetized, clip the abdomen and prepare it for aseptic surgery. Place the patient in dorsal recumbency and drape appropriately. Make a 2- to 3-cm incision centered midway between the umbilicus and pubis. There is usually little subcutaneous tissue and the linea alba is broad, making it easy to identify. Immediately dorsal (deep) to the body wall is the thin-walled cecum, and the bladder (also thinwalled) is just caudal to the cecum. It is vital to avoid iatrogenic injury to these structures, especially the cecum. Leakage of cecal contents can cause life-threatening complications. In addition, because the cecal wall is so thin, it is difficult to achieve typhlotomy closure free of leaks. Because of the potential for damage to these organs, use of a spay hook is not recommended. Locate the uterus between the bladder and the colon. Use a blunt instrument or a finger to move the cecum and bladder to the side on which you are standing, allowing visualization of the uterine horn on the opposite side. Grasp the uterus gently with forceps, and exteriorize it. Trace it cranially to locate the ovary on that side. The ovaries are supported by the mesovarium, which that originates in the area of the caudal pole of the kidney. The mesovarium is short, making the ovaries are
CHAPTER 25 Soft Tissue Surgery more difficult to exteriorize than in carnivores. It may be necessary to extend the incision cranially to avoid accidental tearing of the friable, fat-filled ovarian ligament. The broad ligaments also contain a large amount of fat, which can make identification of the ovarian vessels difficult. A single artery and vein run medial to each ovary and uterine horn.14 Identify the vessels supplying the ovary within the mesovarium and, using gentle blunt dissection, create an opening in the mesovarium to allow placement of two hemostatic clips or two ligatures of an absorbable synthetic suture. Transect the suspensory ligament, mesovarium, and vessels distal to the ligatures. Alternatively, these vessels can be sealed and cut with a tissue-sealing device such as a CO2 laser, a Harmonic device, or a LigaSure (see Chapter 28). It is important to remove the entire oviduct encircling the ovary. Remnants of oviduct can develop into cystic masses within the abdomen.19 Repeat the procedure on the contralateral side and bluntly dissect the broad ligament on each side to the level of the uterine body. Strip the broad ligament on each side caudally to the uterine vessels and uterine body. Ligate the vessels with the uterine body unless they appear particularly large, in which case ligate them separately. The uterus may be ligated with an encircling ligature or with a transfixation ligature. It has been recommended that the uterus be ligated cranial to the cervix to prevent spillage of urine into the abdomen when the uterus is transected19; however, this is of little clinical importance. Place the ligature in the body of the uterus in a convenient location. Remove the ovaries and uterus as a unit. Close the abdomen with 4-0 monofilament absorbable suture in the linea alba in a simple continuous pattern, using 5-0 absorbable suture for the subcutaneous or subcuticular closure. If necessary, use tissue adhesive, 4-0 or 5-0 nonabsorbable suture or skin staples to appose the skin.
PYOMETRA Pyometra is infrequently reported in guinea pigs and chinchillas.4,43 Possible pathogens include Bordetella bronchiseptica, Escherichia coli, Corynebacterium pyogenes, Staphylococcus species, and Streptococcus species. Affected animals are usually presented for vaginal discharge and may be lethargic and anorectic. Some guinea pig owners report polydypsia and decreased appetite. Radiographic and abdominal ultrasound examinations are valuable in obtaining a diagnosis and in ruling out pregnancy, dystocia, and abdominal masses. Vaginal cytology, along with culture and sensitivity testing, confirms the tentative diagnosis. Stabilize the patient and perform a complete blood count and plasma biochemical panel before surgery. Vascular access is required because these patients are usually dehydrated and have other metabolic abnormalities. It also allows intravenous antibiotics to be administered. Administer an appropriate antibiotic intravenously after samples for culture have been obtained intraoperatively. Definitive treatment of pyometra is ovariohysterectomy, which is performed as soon as the patient is stable enough to undergo general anesthesia and surgery. Take care not to spill uterine contents into the abdomen during removal of the uterus. Irrigate the abdomen with warm sterile saline solution before routine closure. Continue fluid therapy and nutritional support postoperatively until the patient is eating and drinking normally.
329
UTERINE TORSION Uterine torsion is uncommon in most domestic pets but has been reported in gravid guinea pigs after 30 days of gestation and in gravid chinchillas.43 Signs are the same as those for dystocia, but usually signs of circulatory shock and acute collapse are also present. The mortality rate is high, and the diagnosis is usually made at necropsy. This is an emergency situation. Establish vascular access and obtain a minimum database before surgery. Stabilize the patient metabolically as well as possible and then perform an emergency ovariohysterectomy.
DYSTOCIA Dystocia is relatively common in guinea pigs and chinchillas because of the relatively large size of the fetuses in these animals.29,43 Degus are smaller and less well developed at birth but are still considered precocious.20 Guinea pigs should be bred before they are 6 months of age, because bony fusion of the pubic symphysis occurs between 6 and 9 months of age. If the pubic symphysis fuses before the first litter is delivered, dystocia can result.29 If a guinea pig delivers a litter before bony fusion of the pubic symphysis has occurred, cartilaginous fusion is preserved for life and future litters are possible without dystocia. Female guinea pigs are sexually mature at 28 to 35 days of age. Weaning typically occurs at 14 to 28 days of age.15 In female chinchillas, fusion of the pubic symphysis is normal and does not cause dystocia. Male and female chinchillas reach sexual maturity at 4 to 12 months of age, much later than guinea pigs.29 Chinchillas are seasonally polyestrus and age at puberty is a function of when they were born. Those born in the late summer do not reach maturity until the next fall breeding season.18 Degus generally wean between 4 and 6 weeks of age and reach sexual maturity at 3 to 4 months of age.20 Gestation is approximately 59 to 72 days (usually 63-68 days) in guinea pigs, 111 days in chinchillas, and 87 to 93 days in degus.15,19,20 Average litter size is 2 to 4 in guinea pigs, 1 to 6 in chinchillas, and 6 to 7 in degus (range, 1-10). In guinea pigs, approximately 10 days before parturition, the pubic symphysis begins to spread. Once the gap is 15 mm, parturition should occur within 48 hours15; at parturition, the symphysis is about 22 mm wide. This gap can be palpated externally; this is a sign of impending parturition.29 If the symphysis is open or the sow has had a previous litter without intervention and the sow has been in unproductive labor for longer than 30 to 60 minutes, give 0.5 to 1 U of oxytocin IM. If no young are delivered after 15 minutes, surgical intervention is likely necessary.30 If guinea pigs become pregnant for the first time after 9 months of age, many will still have normal parturition. If the breeding date is known, palpate the pubic symphysis at about day 60 to determine if it has spread. If so, it is likely that the sow will deliver naturally. If the symphysis does not separate by day 65 or if it has not separated and the sow shows signs of parturition, perform a cesarean section. If the breeding date is unknown, the author has not found a way to determine the due date. Educate owners on the signs of parturition and how to palpate for symphyseal separation. Dystocia in guinea pigs and chinchillas can be surgically treated by either cesarean section or ovariohysterectomy of the intact gravid uterus. Cesarean section is performed to obtain viable fetuses or, if the fetuses are not viable, to preserve the reproductive viability of the sow for future breeding. For either
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SECTION III Guinea Pigs and Chinchillas
procedure, make a routine ventral midline abdominal incision and exteriorize the gravid uterus. For a cesarean section, isolate the uterus with sponges that have been moistened with saline solution and make a longitudinal incision in the dorsal or ventral uterine body, depending on the position of the uterus after it has been exteriorized. Deliver the neonates to an assistant and close the incision with a simple continuous pattern of 4-0 or 5-0 monofilament absorbable suture material. Irrigate the abdomen with warm saline solution before closing. Ovariohysterectomy of the intact gravid uterus can be performed to retrieve the offspring and prevent future pregnancy. The technique is similar to that used for routine ovariohysterectomy. After ligating and dividing the ovarian pedicles, clamp the uterine vessels and uterine body. Transect and remove the gravid uterus, passing it to an assistant before ligating the uterine stump. The assistant opens the uterus and removes and revives the neonates while the surgeon ligates the uterine vessels and uterine stump. It may be necessary to ligate the uterine vessels separately and transfix the uterus, because these structures are enlarged during pregnancy. Clamping of the uterine vessels occludes the blood and therefore also the oxygen supply to the fetuses. Thus it is important to proceed quickly. However, the viability of the neonates is generally not affected by removing them with the uterus en bloc. If the uterus is exceptionally large or engorged with blood and the sow is anemic, the en bloc technique is not recommended. If ovariohysterectomy is indicated, perform a cesarean section first and allow the uterus to involute before removal so that most of the blood in the uterus returns to the sow’s circulation. Give oxytocin IV, IM, or injected directly into a uterine artery (to deliver the hormone directly to the uterus) to speed involution. Guinea pig and chinchilla young are precocious at birth. Their eyes are open, they ambulate well, and they can eat solid foods; however, they should be allowed to nurse as soon as the sow has recovered from anesthesia. Guinea pigs that are orphaned at less than 1 week of age have a high mortality rate, indicating that they do need to have sow’s milk.15 Degus are also precocious at birth but less developed than guinea pigs and chinchillas, having a sparse hair coat and closed eyes.20
UTERINE PROLAPSE Uterine prolapse is usually associated with parturition. In most cases a tissue mass is protruding from the vulva after delivery of young (Fig. 25-2). Most often, the owner discovers live young at the same time the prolapse is noticed. The sow may be stable at presentation or may be debilitated, depending on how long the uterus has been prolapsed. Stabilize the patient medically before administering anesthesia. Consider epidural anesthesia for patients that are not stable enough to undergo general anesthesia. Clean and assess the prolapsed tissue. The prolapsed uterus is usually contaminated by the substrate. If it appears to be viable, clean and reduce the prolapse. A concentrated sugar solution or 50% dextrose applied topically to the prolapsed tissue usually helps reduce edema and the size of the prolapsed tissue, making it easier to replace. If the reproductive viability of the sow is to be preserved, reduce the prolapsed uterine horn into its proper location within the abdomen. If the horn is not returned to its normal location, it is likely to prolapse again. Use an appropriately sized blunt probang to gently push the horn to approximately
Fig. 25-2 This guinea pig was presented for uterine prolapse the morning after farrowing.
midabdomen. After the horn is reduced, monitor the sow closely for reprolapse. If this occurs, perform an emergency ovariohysterectomy. A purse-string suture is not recommended because the prolapsed uterus might be retained in the vagina and cause urinary obstruction, which can be fatal. In most cases, ovariohysterectomy should be recommended. If the exposed uterine tissue is not viable, ovariohysterectomy must be performed after the patient has been adequately stabilized. Prognosis depends on the stability of the patient at presentation. If the patient is alert and active, the prognosis is fair to good.
UTERINE TUMORS Leiomyoma is the most common reproductive tumor of guinea pigs and is usually associated with rete ovarian cysts.10,13,15
MAMMARY GLAND NEOPLASIA Mammary gland neoplasia is uncommon in older guinea pigs and is even rarer in animals younger than 3 years of age.30 There are no published reports of mammary neoplasia in chinchillas, but of 233 specimens submitted to an exotic animal pathology service, 2 were mammary tumors with 1 being a benign adenoma and the other an adenocarcinoma. A third poorly differentiated sarcoma was reportedly collected from a mammary gland, but no glandular tissue was identified. Two specimens were diagnosed as mastitis (D. Reavill, personal communication). No mammary gland tumors were diagnosed in degus. Both male and female guinea pigs have mammary glands located inguinally as a single pair. Chinchillas have two pairs of mammary glands. Neoplasias of the glands, both benign and malignant, have been reported to occur in guinea pigs of both sexes; about 70% are benign fibroadenomas and 30% are mammary adenocarcinomas.7 Adenocarcinomas are locally invasive but rarely metastasize7,8,13,15; however, pulmonary metastasis has been reported.22 Liposarcoma, adenoma, papillary cystadenoma, and carcinosarcoma have also been reported.40 Because of the possibility of malignancy, do a preoperative biopsy or fine-needle aspirate cytology of the mass. If the mass is malignant, it is important to stage the disease before surgery. Staging includes blood work, urinalysis, thoracic radiographs, abdominal ultrasound, and evaluation of regional lymph nodes. Wider excision is recommended for malignant mammary tumors, underscoring the need for a preoperative diagnosis.
CHAPTER 25 Soft Tissue Surgery The left and right inguinal mammary glands of guinea pigs do not have a common blood or lymphatic supply.26 Information regarding the incidence of bilateral disease or of a neoplasm developing in the second gland after one is excised is lacking; however, such cases appear to be uncommon. The benefit of ovariectomy for reducing the risk of the other gland developing a tumor has not been studied. For benign mammary masses, marginal resection is all that is required, and the skin over the tumor can be preserved. The benefit of mastectomy over lumpectomy has not been studied in these rodents, but mastectomy is recommended. Excise all of the mammary tissue, including the areola on the affected side. Usually there are several blood vessels in the subcutaneous tissue supplying the tumor that may have to be ligated. Bluntly dissect around the mass, ligating or clipping vessels as they are encountered. Locate and ligate the caudal superficial epigastric artery and vein where they enter the tissue to be excised. The excision will create dead space, and drains and bandages are not well tolerated by rodents. It is best to tack the skin and subcutaneous tissues to the body wall to eliminate dead space, thus minimizing the risk of seroma formation. Submit the tissue for histologic evaluation. Mastectomy of the affected gland with 0.5 to 1.0 cm of healthy tissue surrounding the mass, including deep to it, is recommended for malignant mammary tumors. In most patients, deep to the mass, remove the body wall to obtain the needed deep margin. Plan carefully to allow for adequate closure, because tissue is not abundant in this area. There is usually adequate body wall to close primarily with simple continuous or interrupted suture. Locate and remove the inguinal lymph node. Close the subcutaneous tissues, tacking to the body wall to eliminate dead space, and close the skin routinely. Submit the tissue for histologic evaluation and request that the pathologist evaluate the margins of the submitted tissue for evidence of tumor. Also ask that the pathologist to quantitate the distance from the tumor to the cut surface if margins are clean. This is vital to determine if you have achieved local tumor control. If bilateral mammary tumors are present (benign or malignant), two unilateral mastectomies are staged 2 to 4 weeks apart, allowing one side to heal and the skin to stretch before the other gland is removed.
ORCHIDECTOMY In hystricomorphs, orchidectomy is primarily used to control reproduction. Seminal plugs have been reported to be a primary cause of urethral obstruction in guinea pigs and can be treated or prevented by orchidectomy. This is easier than performing ovariohysterectomy and is associated with less morbidity and mortality. It may also be indicated to decrease aggression and for medical reasons (e.g., testicular tumor). If a boar guinea pig is to be housed with a female that has a fused pubic symphysis, orchidectomy is recommended to prevent the sow from getting pregnant, which could result in dystocia. The testicles of most rodents are comparatively large and descend during the first week or two of life.14 The inguinal canals remain open, and a functional cremaster muscle allows the testicles to migrate into and out of the abdominal cavity.16 Hystricomorph rodents do not have a well-developed scrotum; instead, the testicles are located lateral to the penis in the inguinal region on each side (Figs. 25-3 and 25-4). Rodents have a large epididymal fat pad within the vaginal tunic that helps prevent intestinal herniation. The large seminal
331
a c
c
b d
Fig. 25-3 The genitalia of a male chinchilla. The penis (a) is directed caudally and almost touches the anus (b), and the testicles (c) are located lateral to the penis. An incision can be made over the tail of the epididymis (d) for castration.
vesicles and coagulating glands also partially occlude the internal inguinal ring, preventing herniation. Because of the anatomy, inguinal hernias are very rare in these rodents, and visceral herniation after orchidectomy has not been reported; however, herniation is frequently discussed as a major concern in performing an orchidectomy in rodents. This concern seems primarily theoretical; because these anatomic mechanisms prevent visceral herniation, it appears unlikely that the inguinal canal would have to be closed. Intratesticular injection of 2% lidocaine at 1 mg/kg per testis has been recommended for intraoperative analgesia. Once the patient is anesthetized, each testicle is injected with lidocaine to minimize the perception of pain when the testicles are manipulated in the anesthetized patient. It must be noted that this does not provide any postoperative analgesic effect but may lower the percentage of inhalant agent needed to maintain anesthesia during the procedure. Orchidectomy can be performed by a closed or open technique. A closed technique requires the least amount of exposure, and viscera cannot herniate into the scrotum because the tunic is tied off near the external inguinal ring. With the patient in dorsal recumbency, clip the fur around the scrotum, penis, and inner thighs and prepare the area for aseptic surgery. Palpate both testes, being careful not to confuse the body of the penis with a testicle. The penis can feel similar to a testicle under the skin, especially if one testicle is retracted into the abdomen. The testes are wider and rounder than the body of the penis, and the penis, which is located on the midline, cannot be pushed back into the abdomen, as the testicles can. If one testicle is in the abdomen, gentle caudoventral pressure results in its return into the scrotum. For the closed technique, holding one testicle between the thumb and forefinger, make a 1.0- to 1.5-cm incision through the scrotum parallel to and 0.5 to 1.0 cm lateral to the penis on each side near the external inguinal ring. The incision should not penetrate the tunica vaginalis. If the
SECTION III Guinea Pigs and Chinchillas
332
b
b
a
A
B Fig. 25-4 A, The genitalia of a male guinea pig; a, penis; b, testicles; anus (arrow). B, The penis has been exteriorized.
b
a
Fig. 25-5 The testicle has been exteriorized for a closed castration. a, Testicle within tunic, b, epididymal fat within tunic. Note: inadvertently, small incisions were made in the tunic (arrow).
incision is too close to the penis, the penis can be detached from the prepuce during dissection. In a degu, such preputial damage was successfully treated by suturing the tip of the penis to the cranial edge of the prepuce with 4-0 polydioxanone suture.32 Two years later, the penis was still adhered to tip of the prepuce. Grasp the tunic and remove the testicle from the scrotum with the tunic intact (Fig. 25-5). The tunic is tightly adhered to subcutaneous tissues. Carefully and gently dissect the tunic from its attachments circumferentially. The tunic is also tightly adhered to the end of the scrotum by the ligament of the tail of the epididymis. Break down this ligament to allow the testicle to be exteriorized. Once the testicle is removed from the scrotum, apply caudal traction to it and strip the fascial attachments, using a dry gauze sponge, until the narrow portion of the cord is exposed. Remember, you need to remove only the testicle; pulling the testicle far out is of no added benefit and may damage the epididymal fat and ipsilateral ureter. Be careful to avoid tearing the vaginal tunic during this dissection. Once the testicle has
been exteriorized adequately, gently push the epididymal fat into the inguinal canal and ligate the cord, using a two-clamp technique. Crushing the tissue with the clamps before placing the ligatures is helpful with the closed technique because the tissue cord is thicker, since the vaginal tunic is incorporated. This ensures a more secure ligature, but it can also cause the tunic to tear, which will defeat the purpose of doing a closed technique (i.e., blocking the inguinal canal). The goal is only to remove the testicle; the epididymal fat must be preserved within the inguinal canal to help prevent hernias. With the closed technique, herniation into the scrotum is not possible unless the ligature fails or the tunic tears. In performing an open castration, make the incisions as described above. Extend the incision through the subcutaneous tissues and the vaginal tunic, exposing the spermatic cord. The testicles are easily exteriorized because they are not attached to the internal (parietal) surface of the vaginal tunic; however, they are attached to the tunic by the ligament of the tail of the epididymis (Fig. 25-6). Traction on the testicles will invaginate the scrotal skin toward the incision. Break down the ligament of the tail of the epididymis, allowing the skin and the vaginal tunic to return to their normal location and leaving the testicle outside the body. With the open technique, the testicle is easily exteriorized and the cord to be ligated is of a smaller diameter than with the closed technique. Be careful to preserve the epididymal fat within the inguinal canal. Double ligate the spermatic cord distal to the epididymal fat and transect it distal to the ligatures. If there is concern about visceral herniation because of trauma or removal of the epididymal fat, identify the external inguinal ring and place a single interrupted suture across it, being careful not to damage or occlude the external pudendal artery and vein, which pass through the canal. The incision in the tunic can be closed or left open. Capello describes a technique for castrating a degu that combines the benefits of an open and a closed castration technique; it is also applicable to chinchillas and guinea pigs because they have similar anatomies.6 Make the incisions as described above. Use a mosquito hemostat to bluntly dissect under and around the spermatic cord within the vaginal tunic without damaging
CHAPTER 25 Soft Tissue Surgery
333
c a b
c a
b
A
B Fig. 25-6 A, Open castration of a chinchilla. B, Open castration of a guinea pig. (a) Testis, (b) epididymis, (c) epididymal fat, vaginal tunic (arrow).
the tunic. Pass a suture ligature around the cord but do not tie it. Make an incision in the tunic and perform an open castration as described above, making sure to keep the epididymal fat cranial to the preplaced ligature. After the testicle is removed, tie the ligature around the tunic as close to the external inguinal ring as possible, thus effectively closing off the inguinal canal. Chinchillas have a particularly large tail of the epididymis. Nelson describes an approach for castration through the skin on the tail of the epididymis.27 Make a small incision over the ventral aspect of the tail of the epididymis, being careful not to cut too deeply into the vaginal tunic. Grasp the tip of the tail of the epididymis with gauze and apply traction while pushing the scrotal skin cranially. The testicle can then be completely exteriorized for either an open or a closed technique. Primary closure of the skin with intradermal suture is recommended, but tissue adhesive and second-intention healing have also been used successfully. Guinea pigs and chinchillas seem particularly prone to the development of scrotal abscesses after orchidectomy. If this occurs, it is more likely that visceral herniation will ensue, because the infection causes tissue necrosis in the area of the inguinal canal, widening the opening and allowing intestine to herniate. It is currently unknown why infection occurs more commonly in hystricomorph rodents. One theory is that the location of the incisions and the way the animal stands contaminate the incisions with feces from the substrate. Owners should be warned of this potential complication before surgery is performed and told that the substrate must be kept clean. Advise owners to use clean paper bedding and change it twice daily for 10 days after surgery. Strictly adhere to aseptic technique. Gentle tissue handling is critical, because traumatic manipulations can cause tissue necrosis and predispose to infection. Application of a layer of cyanoacrylate tissue adhesive over the incision may help to prevent bacterial invasion into the incision. Prophylactic antibiotic therapy should be considered as well.
PENILE PROLAPSE Penile prolapse seems to occur in hystricomorph rodents more commonly than in other small mammals. It is often reported anecdotally after orchidectomy and may be related to nerve trauma during the procedure. Prolapse of the penis may also occur but unassociated with any specific cause.
The penis is protected from the environment within the prepuce. When the penis is prolapsed, it is subject to trauma and contamination. With chronic exposure, the mucosa of the penis becomes thicker and more able to resist such trauma. Most owners are concerned by seeing their pet’s penis outside of its sheath. Powers reported preputial damage and lateral deviation of the penis into the subcutaneous tissues as a complication of orchidectomy in a degu.32 An attempt to maintain the penis within the prepuce by suturing the base of the penis to the base of the prepuce was not successful. Subsequently, the tip of the penis was sutured to the edge of the preputial orifice with four interrupted sutures of polydioxanone, being careful to avoid the urethra. The tissue was not scarified. This technique is simple and easy to perform and maintains the penis in its normal position within the prepuce. In that patient, the penis was still adhered to the prepuce 2 years after surgery. The author recommends this procedure for hystricomorph rodents with penile prolapse.
GASTRIC TRICHOBEZOARS Gastric trichobezoars causing clinical illness have been reported in long-haired Peruvian guinea pigs.2,24,39 The condition is not analogous to what was once called trichobezoars in rabbits. In Peruvian guinea pigs, the hairball is quite large for the animal’s body size (4-5 cm), composed of firmly compacted material consisting primarily of hair. The dense, hard nature of such a bezoar makes it unlikely that it would be broken down and passed simply by medical management; therefore surgery is indicated. In a study looking at the influence of hay on alopecia in guinea pigs, it was determined that when hay was withheld from the diet, a loss of hair density was seen within 4 weeks. This did not resolve when a high-fiber pelleted diet was given.12 The alopecia was observed to result from cage mates eating each other’s hair. Therefore a proposed etiology for trichobezoars in Peruvian guinea pigs is inadequate hay in the diet, resulting in overgrooming of the very long hair combined with inadequate exercise and also possible stressors resulting in the formation of a hard trichobezoar.39 Clinical signs of a gastric trichobezoar can be vague and insidious or acute and severe, depending on whether the bezoar is obstructing gastric outflow. Signs include anorexia,
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SECTION III Guinea Pigs and Chinchillas
depression, weight loss, and decreased production of feces. On physical examination, a large firm mass is palpated in the left cranial abdomen, which does not indent with firm pressure. Radiographs may confirm the presence of a gastric foreign body, but only if there is gas in the stomach to serve as contrast. Orally administered barium confirmed the diagnosis in one patient and ultrasound in another.2,39 The advantage of ultrasound is that other organs can also be evaluated, including the liver, for evidence of hepatic lipidosis, which may complicate the condition and postoperative management. Gastrotomy is recommended as soon as the patient is stable enough for surgery. Pre- and postoperative fluid therapy is vital for a successful outcome. Perform a ventral midline celiotomy from the xiphoid to the pubis to allow the large object to be easily removed. Isolate the stomach with gauze moistened in saline solution and make an incision in a relatively avascular area of the stomach midway between the lesser and greater curvatures. Remove the trichobezoar and irrigate the stomach, being careful to minimize abdominal contamination. Close the stomach in two layers; a simple continuous pattern, over sewn with an inverting pattern such as a Cushing’s with a monofilament absorbable material on an atraumatic needle. Irrigate the abdomen with warm sterile saline solution and close the celiotomy routinely. Postoperative care consists of continuing fluid support, analgesia, nutritional support with a syringe-fed diet, prokinetic medications, and antibiotics if indicated. Keep the animal in the hospital until it is eating and drinking and urinating and defecating normally. Free access to fresh hay, regular grooming of the long hair and potentially cutting it, plenty of exercise, and minimizing stress are considered important to reduce the risk of recurrence. Some have advocated petrolatum-based gastrointestinal lubricants, but their efficacy for preventing trichobezoars and effect on the cecum is unknown.39
UROLITHIASIS Urolithiasis is relatively common in guinea pigs and the primary differential diagnosis for hematuria.14,16 Clinical signs include hematuria, stranguria, dysuria, incontinence, anorexia, and pain manifesting as an animal acting sick.16 Urinary tract obstruction because of calculi can occur at the renal pelvis, ureter, or urethra. In male guinea pigs, urinary tract obstruction occurs more commonly secondary to plugs of sperm and seminal fluid. As a general rule, uroliths should be removed. They cause irritation, which often results in hemorrhage and consequent anemia. Uroliths also cause serious pain and can serve as a nidus for bacterial urinary tract infections. In a patient with stones but without obstruction, the stones may to migrate to a point where they do cause obstruction. Undetected urinary tract obstruction often results in death. Calcium oxalate is the most common type of stone found in guinea pigs, and the etiopathogensis is poorly understood.17 Medical dissolution of stones has not been documented in guinea pigs and recurrence after surgical removal is common (see Chapter 23). Renal mineralization often cannot be distinguished from nephrolithiasis on plain radiographs. Ultrasound will determine the presence of stones rather than mineralization of the renal pelvis. Ureteral calculi can lodge anywhere from the renal pelvis to the entrance to the bladder, but they are most commonly located at the caudal bend of a ureter as it enters the
bladder. The urethra of female guinea pigs is quite large, and large stones often obstruct the urethral papilla because they cannot pass through it (Fig. 25-7). Stones within the bladder do not usually cause signs of obstruction but should be removed because they cause pain and cystitis, predispose to bacterial urinary tract infections, and may migrate into the urethra, causing obstruction. If a nephrolith or ureterolith is causing obstruction, the function of the kidney may be compromised. If the kidney is not functional, the best option is to perform a nephrectomy; however, that will leave the patient with only one functional kidney. It is best to determine whether there is any functional capacity in the kidney prior to surgery. The scintigraphic glomerular filtration rate is very accurate for determining whether a kidney has any remaining function, but this modality is not readily available. An excretory urogram can provide valuable information; however, its accuracy will be compromised if the patient is azotemic. Any blushing of the kidney with contrast indicates that there is still some function remaining and it is worth trying to save it. Since recurrence of urolithiasis is common in guinea pigs, it is best to save the kidney if possible. If the kidney on the affected side is not functional, infected, or severely hydronephrotic, it may be necessary to perform a nephrectomy. In other species, end-stage kidneys have been associated with systemic hypertension.38 Ultrasound is a vital part of working up any patient with urolithiasis. Ultrasound will confirm the location of the stones within the urinary system and determine whether they are causing partial or complete obstruction. Some mineral-dense objects that appear to be within the urinary tract are actually outside it. Nephrotomy and pyelotomy are used to remove nephroliths. For large stones, a pyelotomy is preferred because it causes less damage to the kidney; however, if the stones are small, the pelvis is not dilated and the procedure would be very difficult. Nephrotomy is preferred to remove small stones from the renal pelvis. It is technically easier to perform than a pyelotomy but causes more damage to the renal parenchyma, resulting in at least transient compromise of renal function. For a pyelotomy, perform a standard ventral midline celiotomy and incise the peritoneum lateral but adjacent to the kidney. Bluntly dissect the kidney free from the retroperitoneal fat and reflect it medially, exposing its the dorsal surface. Palpate the stone at the hilum of the renal pelvis and incise the pelvis with a No. 11 blade. Extend the incision to allow the stone to be removed. It is vital that the incision be made on the dorsal aspect of the renal pelvis to avoid the renal artery and vein. Once the stone or stones are removed, flush the renal pelvis with a small Teflon venous catheter and advance the catheter into the ureter to ensure its patency. Close the incision in the renal pelvis with a fine (6-0 to 8-0) monofilament suture on a small atraumatic needle. Replace the kidney to its normal position and place 2 to 4 sutures between the renal capsule and the peritoneum (nephropexy) to hold it in place and prevent renal torsion. Close routinely. To perform a nephrotomy, free the kidney from the retroperitoneal space as described above. Hold it at the hilum between the thumb and first finger. Digitally compress the renal artery and vein or occlude them with atraumatic vascular clamps or Rummel tourniquets while making an incision on the convex surface of the kidney directly through the renal parenchyma all the way down to the renal pelvis. There will be a moderate amount of hemorrhage, so work quickly. Remove the stones
CHAPTER 25 Soft Tissue Surgery
A
C
335
B
D
Fig. 25-7 A and B, A radiodense urethral calculus located at the urethral papilla is noted on abdominal radiographs of this guinea pig (arrow). C, The stone can be seen at the external orifice of the urethra (arrow). D, An incision was made over the urethra to allow the stone to be removed. The incision was allowed to heal by secondary intention. (Images courtesy of Dr. Estella Boehmer.)
and flush the renal pelvis and down the ureter to make sure all stones are removed and the ureter is patent. Once all of the stones have been removed, hold the split kidney together for 5 minutes. This allows a clot to form, and holding the two halves together and should control the hemorrhage. Place a simple continuous suture of fine monofilament material on a small needle in the renal capsule. The renal capsule is very thin and fragile, and sutures easily tear through. Replace the kidney to its normal position, perform a nephropexy, and close routinely. The ureters of hystricomorph rodents are very small, but if there is a ureteral obstruction, the lumen is usually dilated cranial to the obstruction, making surgery more feasible. The blood supply to the ureter is tenuous and must be preserved during ureterotomy. Through a standard ventral midline celiotomy, retract the viscera to the contralateral side, allowing visualization of the kidney and urinary bladder. Palpate between the kidney and bladder just lateral to the hypaxial muscles, where the ureter is located. Attempt to identify the stone by palpation in order to minimize dissection. Some distal ureteral stones can be gently manipulated into the bladder and removed through a cystotomy, which is preferable to performing a ureterotomy; however, often the stones are adhered to the wall of the ureter and cannot be manipulated into the bladder. Once the stone is located, open
the peritoneum over the stone and bluntly dissect the ureter free from the surrounding fat, being very careful not to disturb the small amount of fat directly associated with the ureter, because the ureteral vessels are within that fat. Do not attempt to clear off the fat except in the immediate vicinity of the stone. Isolate the segment with the stone and make an incision in the ureter cranial to the stone in the more dilated segment of ureter and continue the incision onto the stone. Remove the stone through as small an incision as is possible. Irrigate with a fine-gauge venous catheter in both directions. If the catheter will not advance into the caudal, smaller segment, attempt to pass a 4-0 nylon suture to confirm that it is patent caudally. If it is not patent, perform a nephrectomy. In order to minimize the risk of suturing the ureter closed, I prefer to place a stent through the ureter into the urinary bladder while closing the ureterotomy. A cystotomy is done to retrieve the stent after the ureterotomy has been closed. The stent can be a catheter or a suture placed to maintain the lumen’s patency during suture placement. It should pass into the urinary bladder so that it can be removed after the ureterotomy is closed. If the ureterotomy is small, consider closing it transversely to minimize the risk of postoperative stenosis. Once the ureter is closed, monitor the site carefully for any leakage. Urine should be flowing through the ureter and there should be no leakage.
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SECTION III Guinea Pigs and Chinchillas
A simple continuous pattern of a very fine (8-0) monofilament material on a small needle is recommended because a continuous pattern creates a better seal than an interrupted one. Replace the ureter and close the peritoneum and cystotomy prior to routine closure of the celiotomy. Cystotomy for stone removal in rodents is routine. Make a ventral midline celiotomy incision just cranial to the pubis. Exteriorize the bladder, and examine the urinary tract carefully. Isolate the bladder with gauze sponges moistened with saline solution. If possible, place a catheter retrograde into the urethra to prevent stones from migrating into the urethra during surgery. Often, a small (24-26 gauge) Teflon intravenous catheter can be passed into the urethra of males (penis) or females (urethral opening in the clitoris). Magnification is very helpful for this procedure. Make a 2- to 3-cm cystotomy incision on the ventral aspect of the bladder, closer to the apex than the neck to avoid damaging the ureters at the trigone. Trim about 1 mm of bladder wall off one side of the incision and submit the tissue for culture and sensitivity testing. Remove the stones and irrigate the bladder to make sure that all stones are gone. Use a small intravenous catheter passed from the neck of the bladder into the urethra to confirm the patency of the urethra before closure. Submit the calculi for stone analysis and culture and sensitivity testing. Close the cystotomy in a single-layer simple interrupted or simple continuous pattern with a 4-0 or 5-0 monofilament absorbable material on a small atraumatic needle. Irrigate the abdomen with warm saline solution and close routinely. As soon as samples have been collected for culture, administer an appropriate, broad-spectrum antibiotic, which may be extended or changed based on the results of the culture and sensitivity testing. Continue to administer fluids at an appropriate rate (maintenance or higher) for 36 to 48 hours postoperatively. Hematuria may persist for several days. Stones can migrate into the urethra, which seems to be a more common occurrence in females than males, likely because the pelvic urethra in females is wider. Often urethral calculi can be manipulated or retrohydropulsed into the urinary bladder and removed through a cystotomy. In some patients, however, the calculus remains lodged, and adhered to the urethral mucosa, and cannot be moved. Perform a urethrotomy by cutting directly over the stone in the perineum, allowing it to be removed. Once the stone is removed, irrigate the site and allow it to heal by second intention. It is difficult to suture the urethra closed accurately, and it has been shown that there is no difference in healing between those sutured closed and those left to heal by second intention.44 Unsutured urethrotomies do bleed more postoperatively, but it is not enough to be of clinical concern. Not all of the factors that predispose individual rodents to the development of cystic calculi have been determined. It is difficult to cure this condition in guinea pigs, and recurrence is common.1 Cultures of the bladder wall and the calculus should be obtained during the procedure to rule out bacterial cystitis as the cause of the calculus formation. A diet low in calcium along with urine alkalinizers may be indicated; these are discussed further in Chapter 23.
CUTANEOUS DERMAL MASSES Skin and subcutaneous tumors are the second most frequently reported neoplasms in guinea pigs, representing 15% of neoplasms.8,40 Frequently they grow to a large size before the animal
is presented for treatment. Most are trichofolliculomas, which are benign tumors of basal cell origin. They are typically round, firm cutaneous masses usually in the lumbosacral region. Other cutaneous neoplasms reported include trichoepitheliomas, sebaceous adenomas, fibromas, fibrosarcomas, lipomas, fibrolipomas, undifferentiated sarcomas, and adenocarinomas.13,40 It has been suggested that the trichoepitheliomas reported in guinea pigs have actually been trichofolliculomas.13 Trichofolliculomas are typically cystic and may contain sebum, hair, and keratin debris. They can grow quite large without causing clinical problems. Aspiration of sebaceous debris from a cutaneous mass supports a diagnosis of a benign trichofolliculoma. If sebaceous and keratinaceous debris is not present cytologically, submit an aspirate or biopsy to determine the type of tumor, since the surgical and medical management of malignant dermal lesions is different. Marginal excision of a benign mass and histologic confirmation are recommended even if a benign cyst is suspected. Benign cystic masses can become infected. If surgery is delayed and the cyst becomes infected, the guinea pig will likely be ill, complicating anesthesia and surgery. Guinea pigs have a moderate amount of loose skin over the dorsum. After excising the mass, create a single-pedicle advancement flap with the pedicle or base being no less than half the length of the flap. Trim the dog ears away from the pedicle to maximize the blood supply to the flap. Use walking sutures oriented along the long axis of the flap to advance the flap and close dead space. Place simple interrupted monofilament nonabsorbable sutures in the skin. If the mass is malignant, more aggressive surgery is indicated, as described for malignant mammary tumors previously in this chapter.
CERVICAL LYMPHADENITIS Cervical lymphadenitis, a condition known as “lumps” in guinea pigs, is a streptococcal infection of the cervical lymph nodes. It has been suggested to occur secondary to oral mucous membrane trauma and usually results in lymph node abscessation.30 Isolate affected guinea pigs from other guinea pigs until the condition has resolved. The optimal treatment is complete surgical excision of any involved lymph nodes, not just lancing and draining the abscess. Provide supportive care and antibiotic therapy based on the results of culture and sensitivity testing of samples taken at surgery. If the abscesses are small, this approach may result in a cure; however, even with apparent total excision of all grossly infected tissue, the condition can recur in adjacent tissues shortly after surgery. It is important to perform surgery as early as possible, while the abscesses are small. If excision is not possible, it has been recommended that the abscess be lanced, drained, and flushed, leaving the wound open for granulation. Wound irrigation and topical as well as systemic antibiotic therapy may resolve the abscesses; however, recurrence is very common. Persistent, nonhealing abscesses can be cauterized with silver nitrate.26 The silver nitrate is caustic and kills bacteria within the cauterized tissue. Unfortunately it also causes necrosis of healthy tissue, so it must be used judiciously. More recently, antibiotic-impregnated polymethylmethacrylate (AIPMMA) beads have been used to control microscopic disease after excision of these abscesses. Excise the abscesses by removing as much infected tissue as possible so that there is no gross disease remaining. Loosely fill the dead space created by
CHAPTER 25 Soft Tissue Surgery lymph node excision with AIPMMA beads and close the subcutaneous tissues and skin over the beads routinely. The beads release antibiotic into the local tissue for an extended period and in most cases do not have to be removed.
THORACOTOMY The primary indications for thoracotomy in guinea pigs and chinchillas are pulmonary abscesses and neoplasms. Pulmonary tumors are the most common neoplasms observed in guinea pigs.13,40 Most are benign bronchogenic papillary adenomas reported to comprise 30% to 35% of all neoplasms in guinea pigs over 3 years of age,7,8,15,22 but alveolar and bronchogenic carcinomas are also reported. Most pulmonary tumors are slow growing, and clinical signs do not occur until late in the course of the disease. During a 35-year-period, only two chinchillas presented to the University of Tennessee College of Veterinary Medicine had tumors, one of which was a pulmonary adenocarcinoma.13 Thoracotomy is challenging in histricomorph rodents because endotracheal intubation is difficult. Various techniques for intubating guinea pigs have been described.23,41 If thoracotomy is indicated and the patient cannot be intubated per os, a temporary tracheostomy can be performed to establish an airway, allowing for ventilation of the patient during surgery. Make a 1.0- to 1.5-cm skin incision on the ventral cervical midline. Bluntly dissect through the subcutaneous tissues and identify the sternocephalicus muscles. Separate these muscles along the midline, being careful not to damage the thyroid vein. Identify the trachea, and bluntly dissect the peritracheal tissues off the cartilage, preserving the recurrent laryngeal nerves. Place a 3-0 nylon suture around the cartilage ring caudal to the proposed tracheotomy site and create a large loop of suture with long suture tails. Use the suture to pull the trachea to the surface, facilitating tracheostomy tube placement. Make a transverse incision in the trachea approximately one-third of the diameter of the trachea and enlarge it to 50% with hemostats to avoid cutting the recurrent laryngeal nerves. Insert a sterile endotracheal tube (1.5-2.0 mm) into the aborad segment of the trachea. After the procedure is completed and the patient is awake, remove the tracheostomy tube and allow the surgical site to heal by secondary intention. Rats maintained using a tight-fitting face mask for anesthesia and controlled ventilation had thoracotomy for lung lobectomy with very low mortality (see Chapter 28).36 This technique would likely be useful in hystricomorph rodents as well. If the mass is small enough, it can be removed through a lateral thoracotomy at the fourth-to-fifth intercostal space to gain access to the pulmonary hilum. Make a standard intercostal approach and exteriorize the lobe containing the tumor. Magnification is very helpful for this type of procedure. Identify the hilum, the pulmonary artery and vein, and the bronchus. Ideally, the pulmonary artery and vein should be isolated, ligated, and transected individually to minimize the risk of arteriovenous fistula formation. Realistically, it is best to ligate the artery, vein, and bronchus with a single ligature. Transect distal to the ligature and remove the affected lobe. Inspect the stump for hemorrhage or air leakage. Fill the chest with warm saline to observe for bubbles and allow for patient warming, and remove the solution prior to closure. Closure is routine and includes placing a thoracostomy tube to maintain negative intrapleural pressure during recovery. A 5-Fr red rubber catheter serves this
337
purpose well. Once negative pressure has been maintained for a couple of hours, remove the tube. A tube within the thoracic cavity stimulates the production of 1 to 2 mL of effusion per kilogram body weight per day. For large masses and cranial mediastinal masses, a ventral midline thoracotomy is preferred. Place the patient in dorsal recumbency and make a ventral midline incision. The approach is analogous to that used for larger animals with one exception. Because the sternebrae are very narrow, it is not feasible to split them longitudinally. Instead, cut the ribs on one side at their attachment to the sternebrae. After removal of the mass, close with figure-of-eight sutures of monofilament absorbable material encircling the sternebrae at each rib. A second layer, apposing the muscles ventrally, provides additional stability. Close subcutaneous tissues and skin routinely. Place a chest tube to allow control of the pleural space postoperatively. Pulmonary abscesses are also treated by pulmonary lobectomy; however, patients with abscesses are usually more systemically ill. Additionally, during manipulation of the lung lobe, purulent material can flow from the affected lobe into the bronchus and then into other lobes. These factors make the prognosis guarded to poor for guinea pigs, chinchillas, and degus undergoing surgical removal of pulmonary abscesses. Gently lift the affected lung lobe out of the thorax and quickly clamp the hilum to prevent pus from migrating into other lobes. Ligate or clip proximal to the clamp to control the artery, vein, and bronchus, then transect between the clamp and ligature. Check for leaks and close routinely. Long-term antibiotic therapy can be considered as an alternative to pulmonary lobectomy for abscesses; however, because of the caseous nature of the pus, medical management is often not successful in resolving the infection.
MISCELLANEOUS PROCEDURES Exploratory laparotomy for abdominal masses is indicated for hepatic cysts or neoplasms, uterine neoplasms, ovarian cysts or neoplasms, and gastrointestinal obstruction. The etiology and management of pododermatitis are discussed in Chapter 23. These fibrous granulomas of the plantar surface of the feet are usually seen in guinea pigs kept on wire floors. The prognosis for cure is guarded, and many lesions recur after treatment. It is imperative to change the husbandry conditions. Surgery should not be attempted unless the owner is willing and able to house the patient on a soft bedding and can keep the bedding clean by changing it daily. This must be done for the rest of the animal’s life. Aggressive excision of the lesions, followed by bandaging and open-wound management, allowing the surgical wounds to granulate and epithelialize, along with appropriate systemic antibiotic therapy, has met with some success. Alternatively, debridement and placement of AIPMMA beads has resulted in a cure in some cases. Debride the infected areas of purulent material and infected tissue. Place small AIPMMA beads in the defect, and suture the skin together over the beads, across the defect, to hold them in place. Pad and bandage the feet. Change the bandages every day initially and then every 2 to 3 days. The author has left beads in the feet permanently, and they have not appeared to cause lameness. Degloving of the tail, as described in Chapter 28 in gerbils, also occurs frequently in degus.1,20 Tail amputation is recommended.
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SECTION III Guinea Pigs and Chinchillas
References 1. Bennett RA. Rodents: soft tissue surgery. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester: British Small Animal Veterinary Association; 2009:73-85. 2. Bennett RA, Russo EA. What is your diagnosis? Soft tissue density mass in the stomach consistent with a trichobezoar or phytobezoar. J Am Vet Med Assoc. 1985;186:812-814. 3. Beregi A, Zorn S, Felkai F. Ultrasonic diagnosis of ovarian cysts in ten guinea pigs. Vet Radiol Ultrasound. 1999;40:74-76. 4. Bodri MS, Walker LM. What is your diagnosis? Poor intra- and retroperitoneal contrast suggestive of emaciation and alimentary visceral displacement consistent with bladder and uterine mass. J Am Vet Med Assoc. 1993;202:654-655. 5. Burns RP, Paul-Murphy J, Sicard GK. Granulosa cell tumor in a guinea pig. J Am Vet Med Assoc. 2001;218:726-728. 6. Capello V. Prescrotal open technique for neutering a degu. Exot DVM. 2004;6.6:29-31. 7. Collins BR. Common disease and medical management of rodents and lagomorphs. In: Jacobson ER, Kollias GV, eds. Exotic animals. New York: Churchill Livingstone; 1988:261-316. 8. Cooper JE. Tips on tumors. In: Proceedings. North Am Vet Conf. 1994:897-898. 9. Eatwell K. Ovarian and uterine disease in guinea pigs: a review of five cases. Exot DVM. 2003;5.5:37-39. 10. Field KJ, Griffith JW, Lang CM. Spontaneous reproductive tract leiomyomas in aged guinea pigs. J Comp Pathol. 1989; 101:287-294. 11. Frisk CS, Wagner JE, Doyle RE. An ovarian teratoma in a guinea pig. Lab Anim Sci. 1978;28:199-201. 12. Gerold S, Huisinga E, Iglauer F, et al. Influence of feeding hay on the alopecia of breeding guinea pigs. Zentralbl Veterinarmed A. 1997;44:341-348. 13. Greenacre CB. Spontaneous tumors of small mammals. Vet Clin North Am Exot Anim Pract. 2004;7:627-651. 14. Harkness JE. A practitioner’s guide to domestic rodents. Denver: American Animal Hospital Association; 1993. 15. Harkness JE, Turner PV, VandeWoude S, Wheeler C. Harkness and Wagner’s biology and medicine of rabbits and rodents. 5th ed. Ames, IA: Wiley-Blackwell; 2010. 16. Hillyer EV. Common clinical maladies of pet rodents. In: Proceedings. North Am Vet Conf. 1994:909-910. 17. Hoefer HL. Guinea pig urolithiasis. Exot DVM. 2004;6.2:23-25. 18. Hoefer HL. Chinchillas. Vet Clin North Am Small Anim Pract. 1994;24:103-111. 19. Jenkins JR. Surgical sterilization in small mammals. Spay and castration. Vet Clin North Am Exot Anim Pract. 2000;3:617-627. 20. Johnson D. Exotic pet care: degus. Exot DVM. 2002;4.4:39-42. 21. Keller LS, Griffith JW, Lang CM. Reproductive failure associated with cystic rete ovarii in guinea pigs. Vet Pathol. 1987;24:335-339. 22. Kitchen DN, Carlton WW, Bickford AA. A report of fourteen spontaneous tumors of the guinea pig. Lab Anim Sci. 1975;25:92-102. 23. Kramer K, Grimbergen JA, van Iperen DJ, et al. Oral endotracheal intubation of guinea pigs. Lab Anim. 1998;32:162-164.
24. Kuenzel F, Hittmair K. Sonographische diagnosestrellung eines trichobezoars bei einem langhaarmeerschweinchen. Wein Tieraerztl Mschr. 2002;89:66-69. 25. Mayer J. The use of GnRH to treat cystic ovaries in a guinea pig. Exot DVM. 2003;5.5:36. 26. Mullen HS. Nonreproductive surgery in small mammals. Vet Clin North Am Exot Anim Pract. 2000;3:629-645. 27. Nelson WB. Technique for neutering pet chinchillas. Exot DVM. 2004;6.5:27-30. 28. Nielsen TD, Holt S, Rueløkke ML, et al. Ovarian cysts in guinea pigs: influence of age and reproductive status on prevalence and size. J Sm Anim Pract. 2003;44:257-260. 29. Peters LJ. The guinea pig: an overview. Part I. Compend Contin Educ Pract Vet. 1991;4:15-19. 30. Peters LJ. The guinea pig: an overview. Part II. Compend Contin Educ Pract Vet. 1991;5:20-27. 31. Popesko P, Rajtova V, Horak J. Atlas of the anatomy of small laboratory animals, vol. 1. Rabbit and guinea pig. London: Wolfe Publishing; 1992;148–240. 32. Powers MY, Campbell BG, Finch NP. Preputial damage and lateral penile displacement during castration in a degu. J Am Vet Med Assoc. 2008;232:1013-1015. 33. Quattropani SL. Serous cystadenoma formation in guinea pig ovaries. J Submicrosc Cytol. 1981;13:337-345. 34. Quattropani SL. Serous cysts of the aging guinea pig ovary. I. Light microscopy and origin. Anat Rec. 1977;188:351-360. 35. Redrobe S. Soft tissue surgery in rabbits and rodents. Sem Avian Exot Pet Med. 2002;11:231-245. 36. Roman CD, Hanley GA, Beauchamp RD. Operative technique for safe pulmonary lobectomy in Sprague-Dawley rats. Contemp Top Lab Anim Sci. 2002;41:28-30. 37. Rueløkke ML, McEvoy FJ, Nielsen TD, et al. Cystic ovaries in guinea pigs. Exot DVM. 2003;5.5:33-36. 38. Syme HM, Barber PJ, Markwell PJ, et al. Prevalence of systolic hypertension in cats with chronic renal failure at initial evaluation. J Am Vet Med Assoc. 2002;220:1799-1804. 39. Theus M, Bitterli F, Foldenauer U. Successful treatment of a gastric trichobezoar in a Peruvian guinea pig (Cavia aperea porcellus). J Exot Pet Med. 2008;17:148-151. 40. Toft JD. Commonly observed spontaneous neoplasms in rabbits, rats, guinea pigs, hamsters, and gerbils. Sem Avian Exot Pet Med. 1992;1:80-92. 41. Turner MA, Thomas P, Sheridan DJ. An improved method for direct laryngeal intubation in the guinea pig. Lab Anim. 1992;26:25-28. 42. van Goethem B, Schaefers-Okkens A, Kirpensteijn J. Making a rational choice between ovariectomy and ovariohysterectomy in the dog: a discussion of the benefits of either technique. Vet Surg. 2006;35:136-143. 43. Wallach JD, Boever WJ. Diseases of exotic animals: medical and surgical management. Philadelphia: WB Saunders; 1983. 44. Weber WJ, Boothe HW, Brassard JA, et al. Comparison of the healing of prescrotal urethrotomy incisions in the dog: sutured vs nonsutured. Am J Vet Res. 1985;46:1309-1315.
SECTION FOUR
Small Rodents
CHAPTER
26
Basic Anatomy, Physiology, Husbandry, and Clinical Techniques
Angela M. Lennox, DVM, Diplomate ABVP (Avian), and Louise Bauck, DVM, MVSc
General Characteristics Rats Mice Hamsters Gerbils Anatomic and Physiologic Characteristics General Sexing Rats Mice Hamsters Gerbils Husbandry Housing and Equipment Diet and Feeding Zoonosis Clinical Techniques Handling and Restraint Sample Collection Other Diagnostic Testing Procedures Hospitalization Therapeutics
GENERAL CHARACTERISTICS The hundreds of species belonging to the order Rodentia are grouped into three suborders based on the anatomic and functional differences of the masseter muscle. These three groups are the Caviomorpha (guinea pig-like); the Sciuromorpha Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
(squirrel-like); and the Myomorpha (mouse- or rat-like).20 Although body size varies greatly, in general rodents possess a uniform body structure with various adaptations. While the medical approach to the many small rodents species commonly kept as pets is similar, unusual species are sometimes encountered in practice, and more specific information about these species can be found elsewhere.19 The suborder Myomorpha contains many species of small rodents that appear in the pet trade, including rats, mice, hamsters, and gerbils. Species in this group possess elodont incisors and anelodont cheek teeth.6 The testicles and scrotum are usually large in relation to the overall body size, and the inguinal canal is open, allowing the testicles to pass freely from the abdomen to the scrotum.20 Sciruomorph species such as squirrels and chipmunks are less commonly kep.
RATS Rats (Rattus norvegicus) are common pets and are considered one of the better rodent pets because of their larger size and calm nature. Rats are most popular in the pet market in hooded color varieties, in which the coat color is present only over the head and shoulders. Rats are generally hardy as young animals but may suffer from obesity, chronic respiratory disease, and mammary tumors when older. Rats are large enough to be easily grasped by children, and they rarely bite. Some may be excitable and run when removed from their cages; however, rats have been known to return to their cages after “escaping.” Rats are social and can live in mixed sex groups, and males may be present while females are raising litters. Introduction of strangers can be performed successfully on neutral territory.20 Rats are relatively intelligent and seem interested in humans; they can be trained to come when called for a treat. 339
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Giant Gambian pouched rats (Cricetomys gambianus), which weigh up to 1.5 kg, are rare in the pet trade. The U.S. Department of Agriculture (USDA) issued a temporary ban on their importation after monkey poxvirus was identified in a group of Gambian rats shipped to the United States in 2003.23 The ban was lifted in 2008. In 2004, the Gambian pouched rat was discovered to have established a breeding population in the Florida Keys. Efforts to control their populations are under way.28
MICE Standard laboratory mice (Mus musculus) in different color and coat varieties are common as pets. An adult mouse weighs approximately 30 g. Mice make good pets for older children (10 years of age and up) because they rarely bite, but they may move quickly, so younger children may not be able to handle them. Mice are largely nocturnal but are easily roused. Female mice or castrated male mice are recommended over intact males owing to the strong odor of the latter.20 In general, mice are more solitary than other common rodent pets. Female mice usually do well together, but intact males cannot be kept together, as aggression, injury, and death can result. Pet mice are hardy and rarely suffer from infectious disease; however, mite infestations are common and are difficult to treat. Pneumonia and mammary tumors are also frequently seen. Other less common species, including a variety of African spiny mice (Heteromys species) are kept as pets, but their smaller size makes them more difficult for children to handle. Characterized by their dorsal, inflexible spine-like hairs, spiny mice are likely more closely related to the gerbil.9 In the wild, spiny mice are omnivorous.9
HAMSTERS Hamsters include several groups frequently encountered as pets. Hamsters are the least hardy of the small rodents when newly purchased, and stress-related diseases such as proliferative ileitis are common. Hamsters are nocturnal, but if they are scooped up gently with two hands, they usually awaken without attempting to bite. Touching a hamster’s back with a finger in an attempt to rouse it is likely to provoke a startle or threat response. Excited hamsters often jump from hands or tables, so use appropriate caution in handling them. All hamsters are well known for their ability to escape. Chewed or damaged cage parts should be replaced immediately; if a hamster is cared for by a child, an adult should regularly check the pet’s cage for signs of wear. Hamsters may be stressed by hot, humid environments; therefore an effort should be made to keep them in a cool area of the house during the summer months. The most common species is the golden or Syrian hamster (Mesocricetus auratus), which comes in a variety of coats, including the long-haired or Teddy Bear breed.20 Dwarf hamsters include the closely related Campbell’s Russian dwarf (Phodopus campbelli), the Russian dwarf or Djungarian hamster (Phodopus sungorus), and the Roborovskii or desert hamster (Phodopus roborovskii). They are small (average 25-50 g) and have furred short tails, white underparts, and a grayish to tan dorsal surface with or without a dorsal stripe, depending on the species. They are excitable and more difficult to restrain than golden hamsters, and they may bite when restrained. Dwarf
hamsters are more social than golden hamsters and are more likely to live in family groups. Chinese hamsters (Cricetulus griseus) are similar in size to the dwarf hamsters but are not as social and are best kept separately.
GERBILS In nature, gerbils (Meriones unguiculatus) are desert dwellers with efficient kidneys for conserving water.20 Pet gerbils are available in white, black, buff, gray, and spotted varieties. They are extremely active and may be difficult for smaller children to handle; they can slough their tail skin if the tail is caught or grasped too firmly.20 While gerbils are social in nature and live in family groups, they are territorial and often will not tolerate introduction of strangers (cannibalism can result from attempting to keep incompatible pairs or males together). Gerbils are more disease-resistant than hamsters, although older gerbils may develop a variety of neoplastic and degenerative conditions. Epilepsy has been reported in gerbils but is uncommon in many pet strains.
ANATOMIC AND PHYSIOLOGIC CHARACTERISTICS GENERAL The word rodent is derived from the Latin verb rodere, which means “to gnaw.” Small rodents of the Myomorpha suborder possess a common dental formula: 2(I1/1, C 0/0, M 3/3). The four prominent, incisors are elodont, or continuously growing throughout life, while cheek teeth are anelodont, and do not grow after eruption. The enamel of most common rodents is white; however, some species may have enamel that is orange to yellow in color. The crowns of the mandibular incisors are longer than the maxillary incisors and may be mistakenly assumed to be overgrown. In general, the crown/length ratio for the upper to lower incisors is approximately 1:3. Many small rodents have bulging eyes and may appear exophthlamic, especially when scruffed. The harderian gland, which lies behind the eyeball, produces lipid- and porphyrin-containing secretions that aid ocular lubrication and play a role in pheromone-mediated behavior. These secretions impart a red tinge to the tears and fluoresce under ultraviolet light. Normally, the lacrimal secretions are spread over the pelage during daily grooming. However, in stressful situations and in certain disease conditions, there may be an overflow of tears; this can be inaccurately diagnosed as bleeding from the eyes and nose. Rodents are monogastric, with many species having a forestomach that is separated from the glandular stomach by a limiting ridge. They have a relatively large cecum and an elongated colon. Most rodents practice some degree of coprophagy. The ingested fecal pellets presumably provide nutrients, such as B vitamins, produced by the colonic bacteria. Most common rodent species do not vomit, in part because of the limiting ridge in the stomach; but other factors play a role as well, such as the pressure and strength of the esophageal sphincter and crural sling and the innervation of the diaphragm.27,29 For this reason and because these small mammals have such a high metabolic rate, preoperative fasting is not required or recommended. The urinary and reproductive tracts terminate in separate urethral and vaginal orifices in the female. Small rodents are spontaneous ovulators and are polyestrous. Many breed prolifically
CHAPTER 26 Basic Anatomy, Physiology, Husbandry, and Clinical Techniques in captivity. Stages of the estrous cycle can be determined with vaginal cytology. Mammary tissue can be extensive in rodents and ranges from over the shoulders to the perianal region. Mammary tumors can develop anywhere along this tract. Most female rats and hamsters have 6 pairs of nipples, while gerbils have 4 and mice have 5; however, variations in numbers can be seen. Hamsters, rats, and mice possess four front toes and five hind toes, which is opposite in gerbils. All rodents have tails; they are longer than the animal’s body in gerbils, rats, and mice. Golden hamsters have very short tails, while dwarf hamsters have relatively longer tails. Rodents do not pant and have no sweat glands; therefore their ability to withstand high temperatures is limited. Heat dissipation occurs through the ears and tails. Some smaller rodents may salivate in response to warm temperatures.
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Normal physiologic, reproductive, and growth reference values are presented in Tables 26-1 and 26-2.
SEXING Determining the sex of most rodents is easy in mature animals but can be more challenging in very young ones. In general, the distance between the anus and the genital papilla is a reliable method of determining the sex of young animals. The anogenital distance is greater in males than in females, and the genital papilla is usually more prominent and has a round opening in the male. Examining multiple young animals to make a comparison is helpful. The testes of mature males are well developed, especially in rats. Holding the rodent vertically or applying gentle pressure directed caudally on the abdomen allows the testes to pass from the abdomen through the inguinal canal
Table 26-1 Normal Physiologic Reference Values for Gerbils, Hamsters, Mice, and Ratsa Value
Gerbil
Hamster
Mouse
Rat
Average life span (months) Maximum reported life span (months) Average adult weight (g), male Average adult weight (g), female Heart rate (beats per minute) Respiratory rate (breaths per minute) Tidal volume (mL) Minute volume (mL) Rectal temperature (°C) Approximate daily diet consumption of adult (g) Approximate daily water consumption of adult (mL) Approximate daily fecal production (g) Recommended environmental temperature (°C) Recommended environmental relative humidity (%) Total blood volume (mL/kg)
24-39 60 46-131 50-55 260-600 85-160 — — 38.2 5-7 4 1.5-2.5 18-22 45-55 60-85
18-36 36 87-130 95-130 310-471 38-110 0.8 64 37.6 10-15 9-12 2-2.5 21-24 40-60 65-80
12-36 48 20-40 22-63 427-697 91-216 0.15 24 37.1 3-5 5-8 1-1.5 24-25 45-55 70-80
26-40 56 267-500 225-325 313-493 71-146 0.6-1.2 220 37.7 15-20 22-33 9-15 21-24 45-55 50-65
aAverage
reference values from data given in references 3, 12, 13, 15, 17, 18, and 22. Note that the ranges should be considered as guides; values are likely to vary between groups of animals according to such variables as strain, age, sex, fasted, and methodology.
Table 26-2 Normal Reproduction and Growth Reference Values for Gerbils, Hamsters, Mice, and Ratsa Value
Gerbil
Hamster
Mouse
Rat
Estrogen cycle length (days) Estrus (heat) duration (hours) Length of gestation (days) Pups per litter Weight at birth (g) Eyes open (days) Ears open (days) Hair coat starts (days) Start to eat dry food (days) Optimal weaning age (days) Age of maturation of male (weeks) Age of maturation of female (weeks) Recommended minimum breeding age (weeks) Chromosome number (diploid)
4-7 12-18 23-26 3-8 2.5-3.5 16-21 5 6 16 21-28 9-18 9-12 10-14 44
4-5 8-26 15-18 5-10 1.5-3 12-14 4-5 9 7-10 19-21 8 6 8 44
4-5 9-20 19-21 7-11 1-1.5 12-14 10 10 12 18-21 6 6 8 40
4-5 9-20 21-23 6-13 4-6 12-15 2.5-3.5 7-10 14 21 4-5 4-5 9 42
aAverage
reference values from data given in references 1, 3, 4, 12-14, 21, and 22. Note that the reference values may not represent the mean or range for certain populations or strains of animals; as a result, the values should be interpreted as approximations.
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SECTION IV Small Rodents
HAMSTERS
Fig. 26-1 Anatomy of a female hamster, dorsal recumbency. Note the separate urinary (white arrow) and vaginal (yellow arrow) orifices ventral to the anus (green arrow). (Modified from Capello V. Pet hamsters: selected anatomy and physiology. Exot DVM 3.2, Zoological Education Network, 2001.)
into the scrotum, aiding identification. Female rodents have separate vaginal and urethral orifices, the vaginal orifice being between the urethral orifice and the anus (Fig. 26-1). However, it is difficult to identify the vaginal orifice in immature and very small animals. Grossly observable nipples are seen only on the females of these species. Nipples are observable at 10 days of age in female mice and rat pups.
RATS Because of the popularity of rats in the biomedical research community, an extensive amount of information is available on rat anatomy, physiology, behavior, and diseases. In general, rats are typical rodents, many of the pertinent features of which have already been described. Several other points are useful to remember. Albino strain rats, compared with their pigmented peers, have poor eyesight and rely heavily on their vibrissae for spatial orientation. Rats do not have gallbladders. The white hair coat of rats often yellows with age, and the tail becomes more dry and scaly. Aged male rats develop brown, granular sebaceous secretions at the base of their hair shafts, which some owners may mistake for ectoparasitism.
MICE Male mice are typically twice the size of female mice. Like most other male rodents, they have open inguinal canals, an os penis, and a complex urogenital system that contains several prominent accessory glands. Intermale aggression is a common problem, particularly if the males were not raised together or if they are housed in a confined space with mature females. Male mice produce a characteristic musty odor. Pheromones play an important role in mouse behavior and are mediated through tissues such as the vomeronasal (Jacobson’s) organ, which is located in the floor of the nasal cavity. Estrus is suppressed in female mice housed in large groups (the Whitten effect). Recently bred mice that are exposed to a strange male may have impaired implantation (the Bruce effect).
Hamsters are short-tailed, stocky rodents known for their abundance of loose skin. They have large, potentially reversible cheek pouches; these are paired muscular sacs extending as far back as the scapula. The pouches are evaginations of the oral mucosa and are used for transporting food, bedding material, and occasionally young. Hamsters have a distinct forestomach that, like a rumen, has a high pH and contains microorganisms. Golden hamsters have distinctive hip or flank glands that should not be mistaken for skin tumors. These dark brown patches are found bilaterally along the lumbar area. They are poorly developed in the female, but in the mature male they are prominent and become wet and matted during sexual excitement. Dwarf hamsters (Phodopus species) have a midventral sebaceous gland. Secretions from this gland play a role in territorial marking and mating behavior. Female golden hamsters are typically larger than the males and produce a copious vaginal discharge, normally just after ovulation (day 2 of the estrous cycle). These secretions should not be misinterpreted as indicating a bacterial infection of the genital tract. These females also have paired vaginal pouches that collect exfoliated cells and leukocytes. Russian hamster males are larger than females, and the females do not normally produce a vaginal discharge after ovulation. Hamsters are permissive hibernators. Low environmental temperatures stimulate them to gather food. At temperatures of about 41°F (5°C), they may curl up and enter a deep sleep.
GERBILS Gerbils are adapted to a desert environment. They require very little water and produce only a small volume of concentrated urine. In their natural habitat, they can obtain most or all of their water requirements from metabolic processes and from any available fruit or vegetable matter. Despite this remarkable natural ability, pet gerbils should always have access to fresh water. The gerbil’s red blood cell has a life span of approximately 10 days. The rapid turnover of red blood cells is reflected on a stained blood smear as a pronounced basophilic stippling in a high percentage of these cells. Gerbils of both sexes have a distinct orange-tan oval area of alopecia on the midventral region referred to as the ventral marking gland or pad. This structure is composed of large sebaceous glands that are under the control of gonadal hormones. In the pubescent male, the gland starts to enlarge and produces an oily musk-scented secretion. Gerbils can often be seen rubbing their abdomens on objects; this is thought to be a form of territorial marking.
HUSBANDRY HOUSING AND EQUIPMENT Suitable enclosures for rodents should be escape-proof and easy to clean. While colorful and interesting, multilevel cages with tubes, wheels, and hide boxes may be so difficult to disassemble that basic cleaning is neglected. Manufacturers recommend that the entire cage be disassembled and washed thoroughly, which in reality is rarely done. Newer multilevel
CHAPTER 26 Basic Anatomy, Physiology, Husbandry, and Clinical Techniques
Fig. 26-2 Ideal habitat for pet rats. The cage portion can be separated from the base for ease of cleaning. Note the cloth and plastic “dens,” exercise wheel, and cardboard box filled with paper for enrichment.
wire cages sit on a plastic base and can be separated to facilitate cleaning (Fig. 26-2). This type of cage provides the additional benefit of separating living space up and away from urine and feces. Features of optimal enclosures are slide-out or easy-toremove bottoms for ease of cleaning; bottoms with high sides to contain bedding; adequate ventilation (aquariums are not ideal owing to poor ventilation); large doors for easy access to the pet; and a secure locking mechanisms for each cage opening. Frequent cleaning of the cage is critical in the care of pet rodents. Failure to clean the cage results in the buildup of ammonia and contributes to stress and illness. In mice and rats, Mycoplasma pulmonis organisms multiply more rapidly in the presence of ammonia levels of 50 to 100 ppm.31 The frequency of cage cleaning depends on the cage size and number of animals housed. Advise owners to notice the odor of the bedding; anything other than the scent of clean litter indicates that the cage should be cleaned. Provide food in heavy crocks or food dispensers so that the containers will not be tipped over. Bedding choices include recycled paper products, corncob products, shredded paper, and shavings of woods such as pine and various hardwoods. Much debate exists on the use hardwood and aromatic shavings such as cedar; anecdotally their use is linked to skin and respiratory disease. Paper bedding is generally preferable, although these products are more expensive than wood shavings. In a study of the endotoxin, dust, and coliform content of 20 types of rodent bedding, endotoxin and coliform levels were lowest in paper bedding, and these products were recommended to reduce the risk of respiratory disease and immune suppression in laboratory rodents.35 In rats, the rate of sneezing and incidence of lung pathology was higher in animals housed on aspen shavings than in those housed on paper bedding.5 However, results of one study in laboratory mice found no difference
343
in growth, food intake, oxygen consumption, IgE antibody concentrations, or general appearance and behavior in male CD-1 mice kept on CareFRESH (Absorption Corp., Ferndale, WA, USA) original bedding, cedar shavings, or pine shavings over a 4-month period.2 In a study evaluating the dermal toxic effects of cedar and juniper on mice and rabbits, concentrations normally found in wood shavings did not elicit any hypersensitivity reaction; reactions were seen only at much higher concentrations (50% or more).10 The housing of gerbils on shavings is indirectly involved in the development of facial lesions. Sand or dust normally present in the natural environment is absent in cages with shavings, and dust bathing is thought to be part of gerbils’ normal grooming procedure. Sandboxes can be provided for gerbils in much the same way that they are for chinchillas. Enrichment refers to providing mental stimulation and is appropriate for all captive animals. Enrichment for rodents is provided in the form of exercise wheels, hide boxes, materials to shred, treats wrapped in paper or hidden in toys, and time spent outside the enclosure interacting with the owner. Hide boxes can be commercially purchased or constructed from readily available materials such as polyvinyl chloride (PVC) pipe or cardboard boxes. Cardboard provides the additional advantage of a material that can be shredded, destroyed, and discarded when soiled. Smaller cat treat balls have been used successfully with larger rodents. Many small rodents, especially hamsters, enjoy exercise wheels. Plastic exercise wheels that are almost noise-free are available, although some hamsters may chew on them if other materials, such as soft wood blocks, are not available.
DIET AND FEEDING Rodents naturally hoard food items, and exactly how much food is actually being consumed is difficult to gauge by the rate of disappearance from the feeder. Water can be provided in bowls, but good-quality water bottles will prevent bedding from getting into the water and are generally preferred. Water bottles can malfunction over time, resulting in blockage or leakage. Educate owners to change water and test water bottles daily. Most rodents are omnivorous, often eating grasses, seeds, grain, and occasionally invertebrates in the wild.20 Dietary requirements of species in laboratory settings are well established. In the pet environment, needs are best met with a formulated diet supplemented with small amounts of fresh foods and seeds for variety and interest. Although seed mixes are popular choices for rodents, these diets often lead to selective feeding. Animals usually consume high-calorie seeds (sunflower) and ignore formulated pellets, resulting in dietary imbalance. While many rodents have such a short life span that dietary deficiency is rarely recognized, the most common adverse outcome in rats is obesity. Also of interest are various studies demonstrating increased longevity and reduction of certain diseases in rats maintained on a calorie-restricted diet.26 Gerbils have been used extensively in nutritional studies of dietary fat because of their sensitivity to high fat, high cholesterol diets with resultant changes in blood cholesterol levels. Protein requirements for rodents vary from 14% to 17% in hamsters, from 14% to 16% in rats and mice, and up to 22% in gerbils.20 Formulated diets for hamsters, rats, mice, and gerbils should reflect these requirements. Diets for reproduction in rodents should contain higher levels of protein.3
344
SECTION IV Small Rodents
ZOONOSIS Significant public attention has focused on the zoonotic risk of rodents, in particular lymphocytic choriomeningitic (LCM) virus and hantavirus. While the natural reservoirs of these diseases are various species of wild rodents, there are documented cases of human infection with LCM from exposure to pet rodents.8 Pet hamsters, in particular, carry a wide variety of potentially infectious agents. In general, use standard precautions in handling pet rodents, including thorough handwashing. Because many rodent species are popular pets for children, discuss potential anthropozoonoses with clients, especially if pets are kept in a school setting (see Chapter 40).
CLINICAL TECHNIQUES HANDLING AND RESTRAINT The handling and restraint of small rodents can be challenging. The key is to maximize diagnostic information while maintaining safety for both the examiner and the patient. If at any time the animal appears to be in distress, release it immediately and plan an alternative technique. Many owners bring pet rodents to the clinic in their normal enclosures, which is helpful in terms of observing husbandry conditions and examining fecal output. However, attempting to retrieve a rodent from colorful caging tubes is time-consuming and best avoided. Therefore request that owners confine the pet to a separate carrier or a small travel cage within the cage to facilitate capture and restraint. Much information can be gained by examining the pet as it moves about on the examination table, including respiratory rate and effort, overall demeanor, and gait. Take care to prevent the rodent from leaping from the table. Actual
A
restraint technique varies as to species, patient size, temperament, and overall condition. Watching how owners handle their pets will provide information on how the pets will react to restraint. Most small pet rodents do not object to being held loosely in a cupped hand. Exceptions include rodents that have not been sufficiently acclimated to handling and animals in discomfort or distress. As an example, a hamster with enteritis and intestinal gas will appear to be in great discomfort and will roll onto its back, bite, and resist all attempts at handling. The same animal is often calm and will accept handling once the discomfort has subsided. In general, restraint can be accomplished by three methods: (1) examining the animal as it is loose on the table or in the hand; (2) restraining the animal with small towels or cloths; or (3) restraining the animal by some form of scruffing. In some cases, combinations of the above methods are used. For rats, holding firmly at the base of the tail allows the examiner to slow the animal as it walks freely on the table. Do not use this technique for rodents with furred tails, as this presents risk of a degloving injury of the skin (“tail slip”). Scruffing is effective for many species (mice, hamsters, and gerbils) with the exception of rats, which do not tolerate this method. To scruff, grasp generous amounts of loose skin from the lateral aspects of the neck and over the shoulders with thumb and forefinger (Fig. 26-3). Failure to incorporate enough skin allows the animal to rotate and bite the restrainer’s fingers. Calm, large rodents can be restrained by grasping them about the shoulders and thorax (Fig. 26-4). Extremely fractious or distressed rodents benefit from mild sedation for examination or diagnostic testing. Take care, however, with sick or debilitated patients. In general, if the animal is alert enough to defend itself vigorously, it will tolerate sedation. A combination of midazolam (0.25-0.5 mg/kg) with an opioid
B
Fig. 26-3 Restraint methods for rodents. Hamsters, gerbils, and mice can be restrained by scruffing, being careful to include a generous amount of skin to prevent the patient from turning and biting (A). Larger, calm rodents can often be grasped gently around the thorax (B). (Courtesy of Angela Lennox, DVM.)
CHAPTER 26 Basic Anatomy, Physiology, Husbandry, and Clinical Techniques such as butorphanol (0.1-0.4 mg/kg) or hydromorphone (0.1 mg/kg) is generally safe and effective for light sedation (see Chapter 31 and Appendix). Restrain the rodent gently with a cloth and, with a hind limb exposed, inject quickly with a 27-g needle and small syringe (Fig. 26-5) and return the animal to its enclosure. If additional sedation is needed, administer ketamine or consider brief general anesthesia. To induce anesthesia, transfer the small rodent directly into a small to medium-size face mask; afterward, replace the mask with a rodent-sized anesthetic cone or modified syringe case for maintenance (Fig. 26-6). Very small induction chambers can also be used, but avoid using a large induction chamber because of the greater volume of escaped anesthetic gas into the clinic environment and resultant staff exposure.
345
Intubation has been described in large rodents and can been performed with the aid of endoscopy or specialized equipment used in laboratory animals—in particular, a small pediatric laryngoscope and positioning device for the rat.30 Success requires considerable practice. In clinical medicine, rodents are not commonly intubated during anesthesia.
SAMPLE COLLECTION Diagnostic testing can provide the same benefits in rodents as it does in other species. Improved techniques and the ability to acquire meaningful results from small sample sizes make sample collection possible and practical in rodents. Reference intervals for selected species for hematologic and biochemical testing, serum protein electrophoresis, and urinalyses are presented in Tables 26-3 through 26-9. Reference values are often derived from laboratory populations and will vary with animal age, sex, strain, and husbandry as well as with the method of sample collection and laboratory methodology used to derive the values. Many published reference ranges are derived from very small sample numbers and therefore must be interpreted and used with caution (Carolyn Cray, personal communication).
Blood Sample Collection
Fig. 26-4 Most tame rats can be calmed using this technique: the rat is suspended with the rear feet dangling while the head and ears are gently rubbed. (Courtesy of Angela Lennox, DVM.)
Fig. 26-5 Intramuscular injection can be accomplished in most rodents with towel restraint and exposure of the rear leg.
Venipuncture should be attempted only in normovolemic, normothermic patients; therefore blood collection is often delayed until the animal’s condition is stabilized. Before collecting a sample, the clinician must know the sample volume required for the test in question and the maximum blood volume that can be safely acquired from the patient. The blood volume of most rodents is approximately 6% to 7% of total body weight34; removing a maximum of 10% of the blood volume is generally safe (see Table 26-1). For example, a 100-g gerbil is assumed to have approximately 6 to 7 mL total blood volume; of that, no more than 0.7 mL should be collected. Make modifications to these guidelines according to overall patient condition and, in all cases, collect only the blood volume required for testing. If submitting to a reference laboratory, consult the laboratory as to minimum sample requirements and if serum or plasma samples are preferred. Some laboratories require serum for biochemical analysis, whereas others routinely run analyses on plasma samples to maximize sample volume. The volume of blood required for both a biochemical analysis and an automated complete blood count (CBC) may be impractical to collect in small rodent patients. However, the required volume can be reduced if the laboratory is willing to perform a manual count or estimate the CBC from a blood film. In-house analysis of blood samples is convenient for two specific reasons: the speed of results, which is often important for sick rodent patients, and the ability to use very small sample volumes. The Abaxis VetScan (Abaxis, Union City, CA) will perform a preanesthetic or diagnostic biochemical panel on 0.13 mL of whole blood, putting testing within reach for even very small rodent patients. A single drop of blood collected into a hematocrit tube can provide a blood film for an estimated white blood cell (WBC) count, differential, hematocrit, and estimation of total serum solids. Using these techniques, a biochemical panel and modified CBC as described above can be obtained in a 25-g mouse (6% of body weight = 1.5 mL total blood volume; 10% × 1.5 mL = 0.15 mL).
SECTION IV Small Rodents
346
B
A
C Fig. 26-6 Induction of general anesthesia in a sedated rat by using a standard small-animal mask as an induction chamber (A). Once anesthetized, anesthesia is maintained with a modified syringe case (B) or a commercial rodent face mask (C). (Courtesy of Angela Lennox, DVM.)
Table 26-3 Hematologic Data for Micea Value Hematocrit (%) Hemoglobin (g/dL) Red blood cells (x106/μL) Nucleated RBCs (per 100 WBCs) White blood cells (x103/μL) Segmented neutrophils (%) Band neutrophils (%) Lymphocytes (%) Monocytes (%) Eosinophils (%) Basophils (%) Platelets (103/μL)
44 (40-48) 12.6 (9.9-15.3) 7.2 (6.9-7.5) 0 (0-2)
39 (33-34) 13.4 (11.4-15.4) 5.4 (4.5-6.3) 0 (0-2)
34-50 12.8-16.1 7.5-9.7 —
12.9 (5.5-20.3) 27 (10-45) 0 (0-2) 70 (50-85) 2 (0-6) 1 (0-3) 0 (0-2) 1,200 (1,000-1,400) 79 (69-89) 27 (24-30)
8.7 (4.1-13.3) 29 (15-43) 0 (0-2) 66 (49-84) 3 (0-6) 2 (0-4) 0 (0-2) 1,200 (1,000-1,400) 73 (65-81) 25 (22-28)
4.5-9.1 21-57 — 49-82 2-8 0-3 0-3 421-733
34 (32-36)
34 (31-37)
—
Mean corpuscular volume (fL) Mean corpuscular hemoglobin (pg) Mean corpuscular hemoglobin concentration (g/dL) aValues
(n = 20)
Femaleb
Mixed sexc (n = 50)
Maleb
(n = 20)
— —
are given as mean, mean ± 2 SD, or range. River outbred mice, CD-1 (ICR)BR, 32 to 34 weeks, raised under optimal laboratory conditions. (Available at www.criver.com/sitecollectiondocuments/rm_rm_r_hematology_sex_age_outbred_ mice.pdf) cData courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Automated analyzer for cell count, hemoglobin, and hematocrit. Differential based on 100-cell count by using blood smear made at the time of sample acquisition. bCharles
CHAPTER 26 Basic Anatomy, Physiology, Husbandry, and Clinical Techniques
347
Table 26-4 Biochemical Data for Micea Analyte
Male (n = 26)b
Female (n = 26)b
Mixed sex (n = 50)c
Albumin (g/dl) Alkaline phosphatase (IU/L) Alanine aminotransferase (IU/L) Aspartate aminotransferase (IU/L) Bilirubin, total (mg/dl) Blood urea nitrogen (mg/dl) Calcium (mg/dl) Chloride (mEq/L) Cholesterol (mg/dl) Creatinine (mg/dl) Glucose (mg/dl) Gamma-glutamyl transpeptidase (IU/L) Phosphorus (mg/dl) Potassium (mEq/L) Sodium (mEq/L) Total protein (g/dl) Triglycerides (mg/dl)
3.3 ± 0.3 173 ± 45 48 ± 18 101 ± 33 0.22 ± 0.08 16 ± 3 11.3 ± 0.4 106 ± 2 160 ± 20 0.12 ± 0.04 207 ± 18 0.2 ± 0.5
3.6 ± 0.2 191 ± 47 41 ± 19 92 ± 38 0.18 ± 0.04 15 ± 3 10.6 ± 0.5 108 ± 3 133 ± 30 0.12 ± 0.04 225 ± 28 0
2.5-4.8 51-285 29-191 — 0.1-0.9 18-29 8.7-10.1 — — 0.1-0.4 90-193 —
11.9 ± 1.2 10.0 ± 0.2 153.0 ± 1.6 5.1 ± 0.6 100 ± 22
11.2 ± 0.4 8.8 ± 0.9 152.0 ± 2.8 5.2 ± 0.1 225 ± 21
5.4-9.3 — — 4.6-6.9 —
aValues
are given as mean ± SD or range. River, U.S., and Canadian colonies, CD-1(ICR) mice, 56 to 70 days of age, raised under optimal laboratory conditions. (Hitachi 717 Olympus AU 640e). Values rounded. (Available at http:// www.criver.com/sitecollectiondocuments/rm_rm_r_cd1_mouse_biochemistry_jun_dec05.pdf) cSerum samples courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Ortho (Kodak:Ektachem) 700XR. bCharles
The collection technique depends on patient size and temperament and clinician familiarity. The “correct” technique is that which provides consistent diagnostic-quality samples with optimal patient safety. Samples taken after repeated venipuncture attempts and the use of excessive negative pressure often result in hemolysis and poor sample quality, which can have a negative effect on results. Depending on patient size, samples can be collected with small-gauge needles (25-27 g) with attached small syringes (1 mL or smaller). In very small patients or with vessels where negative pressure causes collapse of the vessel, consider puncturing the vessel with a clear-hubbed needle only and collecting blood with a heparinized hematocrit tube as it flows into the needle hub. Most rodents do not tolerate the level of restraint required to safely collect an adequate volume of blood for analysis. Techniques for restraint of laboratory rodents for venipuncture may not be appropriate for pet rodents. Simple sedation as described above, with the addition of brief general anesthesia if needed, greatly facilitates blood collection in many animals. With thoughtful planning, a single sedation procedure can facilitate complete examination, diagnostic testing, and some therapeutic procedures as well. The cranial vena cava is the largest easily accessible vessel in rodent species. In general, it is located just within the thoracic cavity, dorsal to the cranialmost portion of the manubrium; however, species vary in the lateral excursion and depth of the vessel. Accessing this vessel is usually successful and, with few exceptions, is the first choice in rodent species.25 However, venipuncture of the cranial vena cava requires sedation or general anesthesia of the animal. To collect a blood sample from the vena cava, place the animal in dorsal recumbency, and prepare the area over the
manubrium for venipuncture. Using a 27- to 25-g needle with a 1-mL or smaller syringe, approach the vessel from the right or left of the manubrium, angling slightly medially and dorsally (Fig. 26-7). The presence of a well-developed clavicle in some species (gerbils, hamsters) may complicate needle position. As you advance the needle, apply gentle negative pressure. Entry of the cranial vena cava will result in a flash of blood. If blood does not flow freely, the needle may have passed through or oblique to the vessel or the bevel may be contacting the vessel wall. Rotate or redirect slightly. The vessel is just under the manubrium; therefore deep needle penetration is not needed or recommended. Potential complications include vessel rupture and exsanguination, but this appears to occur rarely and is likely associated with excessive patient movement. Other site options for collecting blood samples are the lateral saphenous or lateral tail vein in the rat, gerbil, or mouse; the ventral tail artery of the rat; and the tarsal veins of larger rodents.3,17,34 The lateral tail veins are located on both sides of the tail and are superficial; they can be seen easily in young animals and albino rats. Ideally, warm the tail gently to increase blood flow and occlude the veins by placing a tourniquet around the base of the tail; a rubber band and mosquito hemostat are suitable for this purpose. With a needle of appropriate gauge for the species, enter the skin at a shallow angle at a point approximately one-third down the length of the tail. If the initial attempt at collection is unsuccessful, try again at a site closer to the base of the tail. To avoid collapsing the vessel, use a smallvolume syringe to withdraw the sample or collect the blood into a microhematocrit tube as it flows freely from the needle hub. A modified butterfly catheter (i.e., with all but the proximal 5 mm of tubing removed) may also be used.
348
SECTION IV Small Rodents Table 26-5 Hematologic and Serum Biochemical Data for Ratsa Value/Analyte
Maleb (n > 150)
Femaleb (n > 150)
Mixed sexc (n = 50)
HEMATOLOGIC TESTING Hematocrit (%) Hemoglobin (g/dL) Red blood cells (x106/μL) White blood cells (x103/μL) Segmented neutrophils (%) Lymphocytes (%) Monocytes (%) Eosinophils (%) Basophils (%) Reticulocytes (%) Platelets (x103/μL)
44.2 (38.5-52.0) 15.5 (13.6-17.4) 8.69 (7.62-9.99) 4.28 (1.98-11.06) 22.2 (9.0-49.3) 73.3 (44.7-87.1) 2.0 (1.0-3.6) 1.7 (0.4-4.0) 0.3 (0-0.6) 1.9 (1.4-2.8) 846 (574-1,253)
43.9 (38.5-49.2) 15.4 (13.7-17.2) 8.20 (7.16-9.24) 2.67 (0.96-7.88) 19.3 (8.8-43.8) 75.8 (48.9-88.1) 2.0 (1.0-3.6) 1.9 (0.3-4.7) 0.2 (0-0.7) 2.3 (1.4-3.9) 836 (599-1,144)
33.0-47.0 11.2-15.9 6.4-8.2 4.7-9.4 7.0-32.0 57.0-91.0 2.0-5.0 0-4.0 0-3.0 — 411-626
BIOCHEMICAL ANALYSIS Albumin (g/dL) Alkaline phosphatase (IU/L) Alanine aminotransferase (IU/L) Aspartate aminotransferase (IU/L) Bilirubin, total (mg/dL) Blood urea nitrogen (mg/dL) Calcium (mg/dL) Chloride (meq/L) Cholesterol (mg/dL) Creatinine (mg/dL) Glucose (mg/dL) Phosphorus (mg/dL) Potassium (mEq/L) Protein, total (g/dL) Sodium (mEq/L) Triglycerides (mg/dL)
4.1 (3.6-4.7) 66 (36-131) 30 (19-48)
4.6 (3.7-5.8) 30 (18-62) 30 (14-64)
2.8-5.3 87-381 36-80
96 (63-175)
101 (64-222)
—
0.10 (0.04-0.20) 16 (11-20) 10.3 (9.1-11.9) 103 (98-106) 59 (37-95) 0.4 (0.3-0.5) 141(106-184) 6.2 (3.6-8.4) 4.6 (3.9-6.1) 6.3 (5.6-7.6) 143 (137-147) 62 (27-160)
0.13 (0.07-0.21) 18 (12-25) 10.6 (9.5-12.1) 102 (97-106) 50 (23-97) 0.4 (0.3-0.6) 119 (89-163) 6.7 (4.5-9.5) 4.1 (3.4-5.1) 6.6 (5.7-8.3) 141 (135-146) 42 (16-175)
0.20-0.70 11-23 5.7-12.4 — — 0.3-0.6 50-135 6.5-12.2 — 5.6-7.4 — —
aValues
are given as means and/or reference intervals. River Crl:WI(Han) rats, 17 weeks of age or older, raised under optimal laboratory conditions. Bayer ADVIA 120 analyzer, HitachiP 800 analyzer. (Available at http://www.criver.com/sitecollection documents/rm_rm_r_wistar_han_clin_lab_parameters_08.pdf) cData courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Ortho (Kodak:Ektachem) 700XR; Automated analyzer for cell count, hemoglobin, and hematocrit. Differential based on 100-cell count by using blood smear made at the time of sample acquisition. bCharles
The ventral tail artery in the rat is another option.3 The artery courses along the ventromedial aspect of the tail, although it is not as superficial as the lateral tail veins. Place the sedated or anesthetized animal in dorsal recumbency. Use a 22-gauge needle and a 3-mL syringe from which the plunger has been removed. Alternatively, some practitioners prefer to use a 23-gauge butterfly needle with short tubing connected to a 3-mL syringe. Make the first puncture attempt at a point one-third down the tail’s length. Enter the skin at a 20- to 30-degree angle to the tail, with the bevel of the needle facing upward. A perceptible “pop” usually indicates that the artery has been entered; this is quickly followed by filling of the syringe (or butterfly tubing) with blood. The high blood pressure in this vessel negates the need for the negative pressure produced by withdrawal of the plunger. Indeed, the presence of the plunger within the syringe case impairs recognition of correct penetration of the
vessel. After the required volume of blood has been collected, withdraw the needle and apply pressure to the puncture site. Note that more time is required to stop the bleeding from the tail artery than from the tail vein. The lateral saphenous and tarsal veins are often visible coursing across the surface of the lateral leg or tarsus of larger, lighter-colored rodents. These vessels are very small and are best punctured with a needle only and samples collected directly into heparinized hematocrit tubes. A technique for saphenous venipuncture of laboratory rodents has been described that is minimally invasive, does not require anesthesia, can be performed by a single person when combined with the appropriate restraint, and can be repeated at the same location multiple times.17 The procedure does require skill and is used when relatively small quantities of blood are required. For use in a mouse, place the mouse head first in a 50-mL syringe case that has been modified
CHAPTER 26 Basic Anatomy, Physiology, Husbandry, and Clinical Techniques Table 26-6 Hematologic Data for Hamsters and Gerbilsa Value
Hamsterb
Gerbilc
Hematocrit (%) Hemoglobin (g/dL) Red blood cells (x 106/μL White blood cells (x 103/μL) Neutrophils (%) Lymphocytes (%) Monocytes (%) Eosinophils (%) Basophils (%) Platelets (x 103/μL) Mean corpuscular volume (fL) Mean corpuscular hemoglobin (g/dL) Mean corpuscular hemoglobin concentration (g/dL)
45-52 15.2-17.4 6.5-7.5 6.3-8.9 16-26 65-80 0-4 0-2 0-2 300-570c 68-74 19-24 30-34
41-52 12.1-16.9 7.0-10.0 4.3-21.6 5-34 60-95 0-3 0-4 0-1 400-600 — — —
aValues
are given as mean, mean ± 2 SD, or range. courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Automated analyzer for cell counts, hemoglobin, and hematocrit. Differential WBC count based on 100-cell count using blood smear made at the time of sample acquisition. cAverage values from data given in references 12 and 16. Note that reference values may not represent the range for certain populations or strains of animals; for this reason, the values should be interpreted as approximations. bData
Table 26-7 Biochemical Data for Hamsters and Gerbilsa Analyte Albumin (g/dL) Alanine aminotransferase (IU/L) Alkaline phosphatase (IU/L) Bilirubin, total (mg/dL) Blood urea nitrogen (mg/dL) Calcium (mg/dL) Creatinine (mg/dL) Creatine kinase (CK) Glucose (U/L) Phosphorous (mg/dL) Protein, total (g/dL)
Hamstersb (n = 50)
Gerbilsc (n = 30)
3.5-4.9 22-128 99-186 0.1-0.7 12-26 5.3-12.0 0.4-1.0 — 37-198 3.0-9.9 5.2-7.0
2.5 (2.1-2.9) 91 (56-165) 118 (70-182) — 18 (11-32) — 0.3 (0.2-0.7) 363 (93-752) 91 (24-117) — 5.6 (4.6-6.3)
aValues
are given as reference intervals and means. Samples are mixed plasma and serum. courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Ortho (Kodak Ektachem) 700 XR. cOrtho Vitros 250 Analyzer. bData
Table 26-8 Reference Intervals for Serum Protein Electrophoresis for Selected Rodent Speciesa Analyte
Mouse
Rat
Albumin (g/dL) Alpha-1 globulin (g/dL) Alpha-2 globulin (g/dL) Beta globulin (g/dL) Gamma globulin (g/dL) A/G ratio
2.13-3.35 0.25-0.46 0.75-1.10 1.42-1.73 0.16-0.32 0.7-1.1
3.35-4.09 0.40-0.59 0.20-0.57 1.33-1.69 0.12-0.51 1.1-1.6
Data courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. aAnalytes measured by Beckman Paragon SPEP II gels in adult animals.
349
350
SECTION IV Small Rodents Table 26-9 Urinalysis Reference Values for Gerbils, Hamsters, Mice, and Ratsa Value
Gerbil
Hamster
Mouse
Rat
Urine volume (mL/24 hours) Specific gravity Average pH Protein (mg/dL)
A few drops–4
5.1-8.4
0.5-2.5
13-23
— — —
1.060 8.5 —
1.034 5.0 Males proteinuric
1.022-1.050 5-7 <30
—, Not available. aAverage reference values from data given in references 3, 12, 17, 22. Note that the ranges should be considered as guides; values are likely to vary between groups of animals according to such variables as strain, age, sex, fasting, and methodology.
bleeding; if it is not, apply gentle pressure to the puncture site. This technique may be too stressful for ill or debilitated animals. Collecting blood samples from the orbital plexus or by cardiac puncture is not recommended in pet rodents because of high complication rates and the availability of more suitable options.
Bone Marrow Collection
A
Collecting a bone marrow sample is possible, even in small rodents. The same technique as described below for intraosseous catheterization of the femur or tibia can be used. Indications are the same as for any other species and include nonregenerative anemia of uncertain origin. For optimal quality, prepare samples immediately after collection.
Urine Collection
B Fig. 26-7 Collecting a blood sample from the cranial vena cava in an anesthetized rat and a golden hamster by using an insulin syringe with a 29-g and 25-g needle, respectively. (B) From Capello V. Application of the cranial vena cava venipuncture technique to small exotic mammals. Exot DVM 8.3, Zoological Education Network, 2006.)
Most rodents readily urinate during handling, especially when excited. These samples are useful insofar as free catch samples are useful in any other species (testing with a urinalysis test strip and determining specific gravity); however, urine culture and sensitivity testing requires samples collected sterilely by cystocentesis. For this procedure, deep sedation or full anesthesia is recommended. The bladders of all but the most obese rodents are readily palpable. With the animal in dorsal recumbency, use a very small gauge needle and direct the needle caudally at an oblique angle through the abdominal wall into the bladder. Alternatively, use ultrasound to locate the bladder for cystocentesis. In some cases very small urine samples (adequate for culture) can be obtained by gently expressing the bladder and collecting the sample from the cleansed urinary orifice into a hematocrit tube. Many rodents will void during handling, sedation, and induction of anesthesia; thus it can be difficult to collect an adequate sample volume.34
Fecal Sample Collection by placing one or more small breathing holes in the tip. Extend an exposed hind limb by firmly grasping the skin just in front or just behind the knee, the exact “grip” depending on the leg being bled and the hand being used. Shave the hair over the lateral tarsal area and wet the skin with alcohol to better expose the superficially located vein. Puncture the vessel at a 90-degree angle to the skin with a needle of appropriate size (a 25-gauge needle is usually sufficient in the mouse). A drop of blood immediately appears and can be collected into a standard microcapillary tube or a microtube container. After the required volume has been obtained, release the skin. This usually is sufficient to stop the
Collecting a fecal sample is simple in most rodents because of the large number of fecal pellets produced. Avoid introducing fecal collection loops or devices into the rectum because of the fragile mucosa of the rodent colon.
Miscellaneous Sample Collection Many types of samples can been collected safely in rodents; these include skin scrapings; hair samples; fine-needle aspirates or biopsy samples of external masses, abdominal masses, or cysts; and thoracic fluid samples. For each patient, carefully consider the type of restraint (manual vs. chemical), anatomic
CHAPTER 26 Basic Anatomy, Physiology, Husbandry, and Clinical Techniques
351
variations, and the size of collection equipment (needles and syringes). Use the equipment that is least invasive and as small as possible to collect diagnostic samples.
OTHER DIAGNOSTIC TESTING PROCEDURES Radiology and ultrasound are commonly performed in small pet rodents (see Chapter 35). In brief, diagnostic-quality images require optimal films and settings and an immobilized, wellpositioned patient.7 With few exceptions, this requires sedation or anesthesia.
HOSPITALIZATION Rodents are frequently hospitalized for advanced care or diagnostic testing. The hospital room should be quiet and free of visual or auditory signs of potential predators. Enclosures in the hospital must be escape-proof, even if the patient appears unable to attempt escape. Most rodents can be housed in clear square plastic pet carriers with snap-on tops. These carriers can be placed into most commercial pet incubators for temperature regulation (Fig. 26-8). Ideal bedding is light-colored towels or paper to allow easy identification of urine, feces, and abnormal discharge such as blood. Hospitalized rodents benefit from a hide box, which can be quickly constructed from a small cardboard box or paper. Stock a variety of rodent foods for feeding hospitalized animals or request a supply from the pet’s owner before hospitalization.
Fig. 26-8 Hospitalized rodents are adept at escape. This rat is housed in an escape-proof plastic pet container inside a standard cage.
THERAPEUTICS As in most exotic species, therapeutic agents are not licensed for use in rodents, and such use is considered off label. Some medications may be too concentrated for small rodents and will require dilution or further compounding.32 Fluids can be administered by the subcutaneous, intravenous, intraosseous, and intraperitoneal routes; the method depends on the overall patient condition (see Chapter 38).11 Intravenous or intraosseous administration is most suitable for unstable, severely dehydrated patients. Placement of an intravenous catheter is difficult in most rodent patients; therefore intraosseous catheterization is a better option.24 Studies in multiple species have shown that drug and fluid administration by the intraosseous route results in similar onset of action and peak blood levels as use of the intravenous route.33 Needles manufactured specifically for the purpose of intraosseous catheterization are available; however, in most small rodents, simple injection needles are appropriate (22-27 gauge, depending on patient size). The most commonly used site is the tibia; however, use of the femur has been described as well. Insertion of an intraosseous catheter is uncomfortable for the animal; hence sedation with a local block or full anesthesia is required. To place a catheter into the tibia, infuse the skin, subcutaneous tissues, and periosteum with lidocaine at 1 to 2 mg/kg and wait for at least 10 minutes. Introduce the catheter at the tibial crest at the insertion site of the patellar ligament to avoid penetrating into the joint (Fig. 26-9). The size and shape of the proximal tibia determines the exact angle of placement, and correct placement is best confirmed by radiography in two views.24 Fluid needs for rodent patients that are stable but mildly dehydrated can be met with subcutaneous or oral fluid
Fig. 26-9 Placement of a 25-gauge needle in the bone marrow cavity of the tibia of a young anesthetized rat. The site is prepared aseptically. The needle is taped in placed and can be fitted with a standard catheter injection cap or a small-gauge syringe for intermittent infusion. (Courtesy of Angela Lennox, DVM.)
administration. Administer subcutaneous fluids in the loose skin between the shoulder blades (Fig. 26-10).11 Most parenteral medications are administered subcutaneously in the loose skin above the shoulder blades or intramuscularly in the rear leg or epaxial musculature. Injecting into the lateral leg muscles carries a risk of damage to the sciatic nerve, which can result in lameness of self-mutilation.11 Needle size should be as small as possible to allow passage of medication and penetration of the skin. Intraperitoneal administration of therapeutics is not advised because of the availability of other sites and potential risk of damage to abdominal structures or inadvertent injection into the gastrointestinal tract (especially the cecum).11 Indications for intravenous administration of medications are rare; however, intravenous administration is possible in a larger sedated or anesthetized patient if done carefully. An alternative is injection via an intraosseous catheter.
352
SECTION IV Small Rodents
Fig. 26-10 Administering warmed subcutaneous fluids in a rat by using manual restraint.
Oral administration of medications is generally uncomplicated in most rodents and accomplished by carefully restraining the animal and then introducing a syringe into its mouth. Because the volume of medication is often small, insulin syringes with the needle removed allow accurate measurement and ease of delivery. Compounding medication with sweet sugar bases often increases compliance. The administration of medications in water or food is problematic because of the tendency of rodents to distrust unfamiliar tastes and to cache food as well as the overall inability to control volume and frequency of administration.11 Adding medications to water may reduce water intake. For these reasons, direct administration of medications is preferred. Because of the high metabolic rate of rodents, hand feeding is very important in anorectic animals. Hand feeding is generally easily accomplished with the use of a food supplement that can be fed through a small syringe. A hand-feeding product for herbivores (Critical Care, Oxbow Animal Health, Murdock NE) is nutritionally balanced and convenient for use. Strained vegetable baby foods such as mixed greens or sweet potato can be added to improve palatability if needed.
References 1. Baker HJ, Lindsey JR, Weisbroth SH. The laboratory rat. Volume 1: Biology and diseases. New York: Academic Press; 1979. 2. Becker CE, Mathur CF, Rehnberg BG. The effects of chronic exposure to common bedding materials on the metabolic rate and overall health of male CD-1 mice. J Appl Anim Welf Sci. 2010;13(1):46-55. 3. Bober R. Technical review: drawing blood from the tail artery of a rat. Lab Anim. 1988;17:33-34. 4. Boorman GA, Eutis SL, Elwell MR, eds. Pathology of the Fischer rat: reference and atlas. San Diego: Academic Press; 1990. 5. Burn CC, Peters A, Day MJ, et al. Long term effects of cagecleaning frequency and bedding type on laboratory rat health, welfare and handleability: a cross-laboratory study. Lab Anim. 2006;40(4):353-370. 6. Capello V, Gracis M, Lennox AM, eds. Rabbit and rodent dentistry handbook. Ames: Wiley-Blackwell; 2005. 7. Capello V, Lennox AM. Radiology of exotic companion mammals. Ames: Wiley-Blackwell; 2008.
8. Centers for Disease Control website. Lymphocytic choriomenigitis. Available at http://www.cdc.gov/ncidod/dvrd/spb/ mnpages/dispages/lcmv/qa.htm. Accessed 7/23/10. 9. Chevret P, Denys C, Jaeger J, et al. Molecular evidence that the spiny mouse (Acomys) is more closely related to gerbils (Gerbillinae) than to true mice (Murinae). Proc Nat Acad of Sci of the USA. 1993;90(8):3433-3436. 10. Craig AM, Karchesy JJ, Blythe LL, et al. Toxicity studies on western juniper oil (Jiniperus occidentalis) and Port-Orfordcedar oil (Chamaecyparis lawsoniana) extracts utilizing local lymph node and acute dermal irritation assays. Toxicol Lett. 2004;154(3):217-224. 11. de Matos R. Rodents: therapeutics. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester, UK: British Small Animal Veterinary Association; 2009:52-62. 12. Field KJ, Sibold AL. The laboratory hamster and gerbil. New York: CRC Press; 1999. 13. Foster HL, Small JD, Fox JG, eds. The mouse in biomedical research. Vol. III: Normative biology, immunology, and husbandry. New York: Academic Press; 1983. 14. Fox JG, Cohen BJ, Loew FM, eds. Laboratory animal medicine. Orlando: Academic Press; 1984. 15. Harkness JE, Turner PV, VandeWoude S, et al. Harkness and Wagner’s biology and medicine of rabbits and rodents. 5th ed. Ames: Wiley-Blackwell; 2010. 16. Harkness JE, Wagner JE. The biology and medicine of rabbits and rodents. 4th ed. Baltimore: Williams & Wilkins; 1995. 17. Hem A, Smith AJ, Solberg P. Saphenous vein puncture for blood sampling of the mouse, rat, hamster, gerbil, guinea pig, ferret, and mink. Lab Anim. 1998;32:364-368. 18. Hoffman RA, Robinson PF, Magalhaes H, eds. The golden hamster: its biology and use in medical research. Ames: Iowa State University Press; 1968. 19. Johnson-Delaney C. Rodents: biology, husbandry and clinical techniques in more unusual pet species. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester, UK: British Small Animal Veterinary Association; 2009:96-106. 20. Keeble E. Rodents: biology and husbandry. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester, UK: British Small Animal Veterinary Association; 2009:1-17. 21. Kohn DF, Wixson SK, White WJ, et al, eds. Anesthesia and analgesia in laboratory animals. New York: Academic Press; 1997. 22. Laber-Laird K, Swindle MM, Flecknell P, eds. Handbook of rodent and rabbit medicine. Oxford, UK: Elsevier Science; 1996. 23. Lennox AM. Firsthand encounter with monkeypox. Exot DVM. 2003;5(4):15-17. 24. Lennox AM. Intraosseous catheterization of exotic animals. J Exot Pet Med. 2008;17(4):300-306. 25. Lennox AM. Venipuncture in small exotic companion mammals. NAVC Clinician’s Brief. 2007; October:23-25. Available at http://www.cliniciansbrief.com/topic/146. 26. Masoro EJ. Caloric restriction-induced life extension of rats and mice: a critique of proposed mechanisms. Biochim Biophys Acta. 2009;1790(10):1040-1048. 27. Montedonico S, Godoy J, Mate A, et al. Muscular architecture and manometric image of gastroesophageal barrier in the rat. Dig Dis Sci. 1999;44(12):2449-2455. 28. Perry ND, Hanson B, Hobgood W, et al. New invasive species in southern Florida: Gambian rat (Cricetomys gambianus). J Mammol. 2006;8(2):262-264. 29. Pickering M, Jones JFX. The diaphragm: two physiological muscles in one. J Anat. 2002;201(4):305-312. 30. Rivard A, Simura K, Mohamme S, et al. Rat intubation and ventilation for surgical research. J Invest Surg. 2006;19(4):267-274. 31. Saito M, Nakayama K, Muto T, et al. Effect of gaseous ammonia on Mycoplasma pulmonis infection in mice and rats. Jikken dobutsu. 1982;31(3):203-206.
CHAPTER 26 Basic Anatomy, Physiology, Husbandry, and Clinical Techniques 32. Spenser EL. Compounding, extra label drug use and other pharmaceutical quagmires in avian and exotics practice. J Exot Pet Med. 2004;13(1):16-24. 33. Waren DW, Kissoon N, Mattar A, et al. Pharmacokinetics from multiple intraosseous and peripheral intravenous site injections in normovolemic and hypovolemic pigs. Crit Care Med. 1994;22(5):838-843.
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34. Wesche P. Rodents: clinical pathology. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester, UK: British Small Animal Veterinary Association; 2009:42-51. 35. Whiteside TE, Thigpen JE, Kissling GE, et al. Endotoxin, coliform, and dust levels in various types of rodent bedding. J Am Assoc Lab Anim Sci. 2010;49(2):184-189.
CHAPTER
27
Disease Problems of Small Rodents
Cynthia Brown, DVM, Diplomate ABVP (Avian), and Thomas M. Donnelly, BVSc, Diplomate ACLAM
The Diagnostic Challenge Pet Rodent Etiquette Scheduling an Appointment Reception Area Medical History Clinical Examination Diseases General Comments Clinical Signs and Treatment by Species Medication and Antibiotic Therapy in Pet Rodents Client Education
The treatment of small rodents as pets is still an emerging field, presenting a terrain bristling with complexities for many veterinarians. First among these is the common perception of rodents. Most people do not see these animals as pets, and their wild counterparts are often called vermin. Moreover, almost all research scientists see rodents as experimental tools. Veterinarians are not immune to such prejudices, and some may feel reluctant to examine a fully grown, red-eyed, wheezing rat. Owners of pet rodents often feel the same aversion for unsympathetic veterinarians and therefore travel long distances to see one understanding of their needs. A second area of concern is the clinical veterinarian’s unfamiliarity with rodent biology. Although much information has been accumulated on wild and laboratory rodents, very little of this pertains to pet rodents. Geriatric diseases, the pharmacokinetics of common drugs, and the beneficial and harmful effects of human handling, contact, and care are a few of the phantom areas that array themselves along the frontier of this field. Yet the problems affecting rodents do not differ greatly from those of dogs and cats. The aims of this chapter are to describe the common diseases of pet rodents seen in practice—so that their relative novelty becomes a challenge and not a stumbling block—and to inform clinicians about reasonable methods for accurately diagnosing common diseases of rodents. 354
THE DIAGNOSTIC CHALLENGE PET RODENT ETIQUETTE Establishing and familiarizing the veterinary staff with a few simple rules of so-called pet rodent etiquette can make the physical examination a positive and fruitful experience. This preparation leaves the clinician confident and free of the anxiety that can arise when the he or she is presented with a concerned, overprotective owner. The veterinarian should be ready to apply his or her acumen and therapeutic skills to the treatment of the patient. The establishment of a clear and nonconflictual basis for communication between clinician and owner greatly facilitates the rodent’s treatment and recovery.
SCHEDULING AN APPOINTMENT Healthy rodents are active during the waking part of the normal circadian cycle. Rats, hamsters, and mice are nocturnal, while Mongolian gerbils and degus can be active during both the day and night.80 When a sleep-deprived, drowsy, irritable animal is brought to the clinician, subtle signs of disease may be overlooked. It can be difficult to determine whether the subdued nature of the animal is related to an underlying illness or the circadian cycle. This is especially the case with hamsters. When possible, receptionists should schedule appointments for most rodents for the early evening hours. Appointment times are not as critical for gerbils and degus. By taking the time to explain the reasoning behind appointment scheduling to clients who are unwilling to make evening appointment, one may not only change their minds but also set the veterinarian-client relationship off to a good start. Instruct the client to bring the rodent to the hospital in its own cage if possible. If not, ask the owner for photos or videos of the cage setup. Knowledge of the animal’s husbandry and sanitation is essential to obtaining a good clinical history. Only by seeing the cage, water supply, feed containers, bedding, and food can the clinician understand the environment in which the rodent is lives. Clients should be tactfully instructed not to clean the rodent’s housing in preparation for the appointment, Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 27 Disease Problems of Small Rodents because doing so they may inadvertently destroy information that is important for diagnosis and treatment.
RECEPTION AREA In an ideal world, a separate waiting area for owners with rodents would be quiet and isolated from natural predators. If possible, receptionists should avoid planning appointments for outdoor cats or hunting dogs when a pet rodent is scheduled for an examination. A schedule block can be created for exotic pet/rodent patient appointments only. If none of those options is feasible, aim to escort clients with pet rodents directly into a clean examination room while they wait so that the animal is physically separate from cats, dogs and ferrets. The rodent’s sense of smell is well developed, and its world is rich in olfactory stimuli and pheromonal cues. Rodents exhibit an innate fear-like behavior when they detect chemosignals of predators. Major urinary proteins (MUPs) released by predators are detected by the rodent’s vomeronasal organ— which also detects pheromones involved in sexual behavior— triggering a fear response.74 Lab coats, stethoscopes, clothing, and hands can retain the scent of predators, which can induce defensive behavior in a rodent during the examination process. This is another reason to advise clients not to clean their pets’ cages before an appointment, as the familiar smell of seasoned bedding will afford comfort to the rodent. Alternatively, the opportunity for a habituated rodent to nestle next to the familiar smell of its owner can only be afforded in a quiet and safe waiting area. Rodents are more sensitive to the effects of heat than those of cold. Even though wild hamsters and gerbils are desert-dwelling animals, their main method of thermoregulation involves escaping from the heat by burrowing or seeking cool places. Mice in particular are very sensitive to the effects of heat. Waiting areas as well as hospital cages for rodents should be kept relatively cool. An ideal ambient air temperature range is 68°F to 70°F (20°C-26°C).14 Educating clients and receptionists about ways to make a trip to the hospital less stressful is well worth the time investment required. If proper attention is devoted to education, then the veterinarian is likely to see a rodent that is amenable to examination instead of one ready to fight, flee, or cringe.
MEDICAL HISTORY Facts about the history and nature of a rodent’s problem are generally more useful in reaching a correct diagnosis than is the clinical history of a cat or dog. Skill is required to extract a reliable, unbiased history of a pet’s disease. Some owners are good at noticing changes and can provide important information, whereas others are not. Find out what owners know about rodents. Have they had rodents as pets before? Did they obtain their information on caring for their pet from a book, a pet store, family or friends, websites, or first-hand experience? Books about rodents for owners of all ages are presented in the “Suggested Client Reading” section at the end of this chapter. Websites about rodents often provide incomplete or misleading information; we cannot recommend any of these at present. Veterinarians should be aware of the current popular books and websites, as clients often have questions based on browsing these sources. Knowledge of your clients’ sources of information can further help you in judging
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their ability to provide an accurate history. Furthermore, pet owners report more confidence in information received from veterinarians compared with information from any other accessible source.51 Do not become unsettled if an owner appears to know more than you do. Such a client can be very informative, enthusiastic, and willing to take an active role in treatment. In discussing a pet’s problem with its owner, communicate on a level commensurate with his or her aptitude and background. Parents frequently present a sick rodent that belongs to their child, who is often the one most knowledgeable about the pet’s habits and behavior. In obtaining a medical history in these cases, the young owner’s presence can be invaluable. Over the course of the exchange with the owner, answers to the following specific questions should be obtained: • Where did the pet come from? a pet store? a laboratory? • How long has the owner had the pet? • Are there other pets in the household? If so, are they of the same species or a different species? • What food does the owner give to the pet? Where is the food purchased? • What food does the pet prefer and what does it actually eat? • Where is the food stored and for how long? • Who is responsible for feeding and cleaning? How routinely are these tasks done? • How long have the signs of illness been apparent? Who first noticed them and why? • Has the pet’s condition deteriorated, improved, or remained stable? Pets isolated from other rodents and household animals and those acquired from a private breeder or laboratory are less likely to suffer from infectious disease than are animals obtained from a pet store. Many diseases are the result of poor or inappropriate feeding. When offered mixed-seed, vegetable, and fruit diets, pet rodents often selectively eat only one ingredient (e.g., sunflower seeds). In households with children, a regular feeding routine may not occur, and doting children may feed pets with inappropriate foods. Often owners are ignorant of the availability of specially formulated diets for pet rodents. These diets, which come in the form of pellets, are convenient and nutritionally balanced sources of nourishment. Feed manufacturers such as Oxbow Hay Products (Murdock, NE; www.oxbowhay.com), Kaytee (Chilton, WI; www.kaytee.com), and Mazuri (St. Louis, MO; www.mazuri.com) have developed diets for pet rodents that are available by direct order or from selected retailers. Diets developed for laboratory rodents can also be used. However, these diets are usually available only in 50-pound bags and can be purchased only from wholesale feed distributors. A list of laboratory rodent diet manufacturers can be found in the annual Buyers Guide issue of the journal Lab Animal (New York, NY) or its website www.labanimal.com. For critically ill, anorectic, or convalescing rodents, several products that can be fed by syringe or gavage tube, such as Critical Care (Oxbow) and Emeraid Omnivore (Lafeber, Cornell, IL; www.lafebervet.com) are available.
CLINICAL EXAMINATION Seeing the condition of the rodent’s living quarters provides information that is helpful in reaching a diagnosis or a reasonable prognosis. Information obtained from a physical
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SECTION IV Small Rodents
examination is limited because of a rodent’s size. However, the significance of the rodent’s history and husbandry can be evaluated only after thorough examination of the animal. With appropriate handling and a few specialized but simple pieces of equipment, the major organ systems can be thoroughly evaluated. If the same procedure is followed consistently, it eventually requires less and less time to perform. Observe the pet rodent in its cage for quality of respirations, activity, condition of grooming, and the presence of a head tilt or discharges. If dyspnea or depression is observed, be extremely careful when handling the animal, as it is probably very sick and could die from the stress of a physical examination. At the same time, warn the owner of your guarded prognosis. Pet rodents that have been frequently and gently handled usually require only minimal restraint. Less cooperative patients need to be more firmly restrained, and the use of a towel or even heavy gloves may be required. Although pet rodents do not often bite, their nips can be painful and may elicit in the handler an unfortunate reflex response that causes the pet to be pitched onto the floor or at a wall. In addition to the potential for traumatic injury that this circumstance entails, the rodent may escape and become harmed. In general, the first component of the physical examination is accurate measurement of the animal’s weight. Weight measurement is essential for calculating appropriate doses of medications and provides an opportunity for gauging the rodent’s temperament before the actual physical examination begins. Rodents are easily weighed in metal or plastic containers placed on a small digital scale. The carrier in which the rodent is presented can sometimes also be used as the weigh basket. A transilluminator, binocular loupe, bivalve nasal speculum and otoscope are useful for evaluating physical signs. Start at the head, examining first the ears, eyes, and nose for discharge and the oral cavity for dentition. The otoscope allows careful examination of the mouth and ears in most small rodents except mice. However, general anesthesia is usually required for a thorough dental examination. Lymph nodes and glands of the head can be observed for size and palpated for consistency. Assessment of the head is probably the most time-consuming part of the examination. Palpate the abdomen for consistency and the presence of unusual masses. However, do not squeeze the patient too hard, because overzealous palpation can result in visceral rupture. Keep in mind, in performing abdominal palpation, that some rodents such as degus and prairie dogs have intra-abdominal testes. Examine the anogenital region for discharges and staining of the fur or skin. When a rodent is picked up, it generally urinates and defecates. Have a dipstick ready to perform an immediate urinalysis or a syringe handy to aspirate urine off a clean surface for urinalysis; feces can be caught in a small tube and examined later if required. By this point in the examination, the condition of the fur and the body in general have been assessed. Palpate the limbs for tenderness or fractures and pay special attention to the paws, noting the length of the nails and the state of the footpads. Rodents are fastidious groomers and therefore can groom away evidence of underlying disease/ illness easily. Observe the medial aspect of the front legs for crusts, debris, alopecia, or porphyrin staining, which may occur from excessive grooming associated with nasal discharge. Also, keep in mind that some cage mates are aggressive groomers and can remove any evidence of illness by keeping their mates well groomed.
Respirations and heart rate are difficult to measure in rodents because they are rapid in healthy animals; instead look for signs of dyspnea. A sensitive pediatric stethoscope is useful for auscultation in large rodents. Some respiratory infections, such as mycoplasmosis, are clinically silent. These diseases can be better heard than seen; abnormal sounds called “snuffling” in rats and “chattering” in mice are noticeable without a stethoscope. It may be useful to put the rodent next to your ear and perform a few gentle chest compressions to evaluate audible respiratory excursions. Wheezing or snuffling may not be present at rest, but when chest compressions are performed to create deep respiration and exhalation, they may often become apparent. The value of determining rectal temperature is questionable. Physical examination combined with attempts to measure rectal temperature causes stress, which can increase body temperature of rats by 3.5°F (2°C) above the nonstressed temperature.9 Core body temperature in rats and mice can vary daily from 96.5°F to 100.5°F (36°C-38°C) because of circadian variation, sex, and age.83 Rectal temperatures can be measured safely with the use of small semiflexible temperature probes connected to a digital clinical thermometer. The probes are reusable; they are available in polyvinyl chloride, nylon, and Teflon and range in size from 1 to 3 mm in diameter. They are ideal for monitoring body temperature when surgery on pet rodents is being performed. A list of manufacturers can be found in the annual Buyer’s Guide issue of the journal Lab Animal (New York, NY) or its website at www.labanimal.com. The manufacturers are listed under “Research/Animal Research Equipment/Temperature Probes.” The clinician can obtain a small amount of blood for a smear and microhematocrit from a hind-limb skin stab, nail clip, or nick of the tip of the tail. An excellent website, written by exotic pet veterinarians, from which to obtain information on clinical techniques is Lafebervet.com Small Mammals (www.lafebervet.com/small-mammals/?p=265). Blood sampling in conscious rodents induces increases in blood pressure, heart rate, and body temperature, which may last up to 30 hours; therefore we often sedate or anesthetize patients to obtain samples.29 Low-dose acepromazine (0.5 mg/kg IM) administration results in peripheral vasodilation, making peripheral venipuncture easier. While some authors advise against using acepromazine in gerbils because it may induce seizures, there is no evidence of a proconvulsive effect in gerbils. Furthermore, clinicians reevaluating acepromazine administration and recurrence of seizure activity in epileptic dogs found no correlation.66 Technologic advances have made possible electrocardiography and accurate and sensitive recordings of heart rate, respiratory rate, and blood pressure in research rodents.53 The cost of the equipment for performing these measurements and the invasive procedures that are often necessary for achieving the recordings prohibit routine use of these testing modalities in most veterinary practices. However, advances in high-resolution digital radiography that require relatively low radiographic exposures, developments in ultrasound, and the availability of computed tomography and magnetic resonance imaging have allowed diagnostic imaging to become a useful ancillary examination.84 Two excellent books designed to provide clinicians with normal anatomic and abnormal comparative diagnostic images are Diagnostic Imaging of Exotic Pets: Birds, Small Mammals, Reptiles54 and Radiology of Rodents, Rabbits and Ferrets: An Atlas of Normal Anatomy and Positioning.93
CHAPTER 27 Disease Problems of Small Rodents
DISEASES GENERAL COMMENTS Diseases of Small Rodents Seen in Practice The prevalence and types of small-rodent diseases seen in practice are quite different from what is seen in a research setting. Although this may seem rather obvious, much of the literature describing the maladies of pet rodents has been inferred indiscriminately from conditions seen in laboratory rodents. The diagnosis and treatment of pet rodents involves evaluation and care of an individual animal from a household, not the health management of rodents from a research colony. Derangements likely to be seen in practice include trauma-induced injuries, infectious and parasitic diseases, neoplasia, and problems related to nutrition and aging; genetic disorders are uncommon. Dermatologic conditions make up 25% of the cases in exotic pets presented for small-animal consultations in general practice in the United Kingdom.42 Natural infections that would be considered rare in a laboratory animal colony often are transmitted to pet rodents by other household animals and children; for example, cats and dogs are major reservoirs of dermatophytes,24 and humans are the natural host of Streptococcus pneumoniae and Streptococcus pyogenes, which are now often antibiotic-resistant.2 Rodents used for research are maintained in tightly controlled environments designed to reduce the impact of unwanted variables in animal experiments.14 However, pet rodents are generally exposed to temperature, humidity, and light-cycle changes; a broad range of foods; numerous microorganisms borne by animals and humans; and various types of handling. Rodents obtained from pet stores have had to endure the stress of overcrowding, transport, and on occasion temperature extremes, all of which put them at risk for disease. As a result, pet rodents exhibit a wider range of physiologic and pathologic responses than do rodents used for research. Consequently, the disease presentation of many pet
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rodents is atypical as compared with the classic experimental disease description. Veterinarians must be discerning in their selection of information about rodents. Research-oriented scientific publications are often more obscuring than elucidating for small pet practice. Research articles often treat rodents as part of a herd or as experimental tools, and disease is diagnosed only by necropsy. Successful in vivo disease diagnosis and resolution are not addressed in such articles, and it is in this area that our understanding must be broadened. Exceptions to this observation are becoming more numerous since the first edition of this book was published. Articles with titles such as “What’s Your Diagnosis,” clinical case reports, and articles on clinical and surgical techniques in pet rodents are being published more frequently in laboratory animal medicine and mainstream veterinary journals worldwide. The Association of Exotic Mammal Veterinarians (AEMV) is affiliated with the Journal of Exotic Pet Medicine (www.exoticpetmedicine.com) and has expanded its website at www.aemv.org to include a searchable database of articles, disease descriptions, and current treatments.
Significant Diseases and Life Spans Pet mice, rats, gerbils, hamsters, and degus are subject to a limited number of naturally occurring medical problems. The most common spontaneous outbreaks of disease are caused, or at least stimulated, by shortcomings in husbandry. While caloric restriction has a significant impact on longevity in rodents as well as other species,73 the adverse effects of overfeeding on the early development of many spontaneous tumors and degenerative diseases has also been seen with “diabesity”—diet-induced obesity and type 2 diabetes.49,67 The more common problems seen in general practice, unique to each species, are listed and grouped by the primary organ system affected in Table 27-1. The average life span of each species also is given. Consistent with causes of death in dogs, unpublished surveys we conducted from rodent cases presented to the Animal Center in New York
Table 27-1 Major Disease Problems of Small Rodents Seen in Clinical Practice SPECIES (AVERAGE LIFE SPAN) Organ System
Mice (1.5-2.5 years)
Rats (2-3 years)
Cardiovascular
—
—
Digestive
Endoparasites, incisor overgrowth Endocrine — Integument and Alopecia, bite wounds, mammary ectoparasites gland Ocular — Reproductive — Respiratory
—
Hamsters (1.5-2 years)
Atrial thrombosis, congestive heart failure Incisor overgrowth Enteritis, weight loss — — Mammary neoplasia, Bite wounds, ectoparasites neoplasia Red tears — Chronic respiratory disease, pneumonia
— Vaginal discharge, maternal cannibalism —
Gerbils (2-4 years)
Degus (7-10 years)
—
—
Enteritis
Dental disease
— Diabetes mellitus Nasal dermatitis, Fur chewing tail slip, neoplasia Cataracts Granulosa cell tumor — —
—
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and the Foster Hospital for Small Animals at Tufts-Cummings School of Veterinary Medicine suggest that young (less than 1 year of age) pet rodents die more commonly of gastrointestinal and infectious causes, whereas older rodents (beyond the median life span) die of neoplastic causes.30 Traumatic injuries are frequent in all rodents of all ages. As prey animals, rodents do not show obvious signs of pain or disease until they are near death. Consequently, sick rodents are often presented late in disease progression compared with earlier presentation in cats and dogs. We have found that indicators of death are a form of shock indicated by lethargy, decreased heart and respiratory rates, and a rapid drop in body temperature to below 91°F (33°C). In rodent aging studies, pronounced bradycardia (30% lower than normal) and hypothermia (13% lower than normal) are significant predictors of death 5 to 6 weeks before expiration.101 Treat this type of shock by warming the patient to restore normal temperature, infusing crystalloid fluids, and providing oxygen therapy. Successful treatment does not guarantee resolution of disease but may buy valuable time to establish a diagnosis and treatment plan for the underlying problem.
Prophylaxis for Small Rodents Prevention of disease in rodents is far more successful than treatment. Disease prevention is primarily based on commonsense husbandry practices, such as purchasing healthy, genetically sound animals; supplying balanced fresh food appropriate in protein and caloric content; avoiding obesity; providing clean fresh water; furnishing adequate shelter, including shade from direct sunlight; avoiding drafts and extreme changes in temperature or humidity; keeping cages clean by preventing the accumulation of excess feces and urine; isolating sick animals from a group for treatment; and protecting vulnerable animals from more aggressive members of their group (e.g., young animals from older animals and male hamsters from female hamsters) or from natural predators living in the same household (e.g., mice from cats). Other sound husbandry practices include housing different species separately to prevent interspecies disease transmission (e.g., rats carry Streptobacillus moniliformis, a cause of septicemia in mice, in their nasopharyngeal cavities) and reducing obesity by limiting food intake and providing cage accessories (e.g., exercise wheels, tunnels, and ramps) that allow play and exploration. Companies selling environmental enrichment equipment and accessories for rodents include Otto Environmental (Milwaukee, WI; www.ottoenvironmental.com) and Bio-Serv (Frenchtown, NJ; www.bio-serv.com). Unlike larger companion animals, pet rodents are not vaccinated. The introduction of avermectins (e.g., ivermectin, selamectin, moxidectin), although this agent is not approved for use in any rodent species, has allowed routine systemic treatment of pet rodents for pinworms, mites, and lice. For ecto- and endoparasite treatment recommendations, refer to Table 27-2. Dental problems are commonly seen in pet rodents because of their continually erupting teeth. Overgrown, maloccluded, or malformed incisors are seen as spontaneous background lesions in 3% (females) to 9% (males) of outbred mice and 14.5% (females) to 10.5% (males) of outbred rats in chronic toxicology studies.62 Specially designed tabletop restraint devices, cheek dilators, mouth specula, dental drills, rongeurs, and filing-rasps for treating dental problems are now commonly available. For more information, see Chapter 32.
CLINICAL SIGNS AND TREATMENT BY SPECIES Mice
Integumentary System and Mammary Glands. Nearly all problems seen in pet mice are associated with the skin. A survey from a large diagnostic laboratory housing research animals indicated that skin disease in mice represents 25% of all diagnostic problem-solving cases (for all species) submitted.55 We categorize four groups of skin problems in mice: behavioral disorders, husbandry-related problems, microbiologic and parasitic infections, and idiopathic conditions. Behavioral, husbandry, and infectious causes of skin disease are relatively straightforward to diagnose and treat. However, many skin diseases characterized by chronic or ulcerated skin (often secondarily colonized by bacteria) are diagnosed as idiopathic. This group is commonly unresponsive to topical or systemic treatment, and affected individuals are often euthanatized. Most damage to the skin is done by toenails as the rodent scratches itself. It is difficult to prevent a rodent from scratching. The nails can be trimmed or filed to remove sharp ends, but attempts to fit the rodent with a bandage or protective wrap are often pointless, as these animals are adept at removing all bandages. Restraint collars can be used, but rodents are not able to eat easily with a collar in place and it will not be effective unless the underlying cause is detected and treated simultaneously. Mice exhibit well-studied social and sexual behaviors. Social dominance, a form of behavior relating to the social rank and dominance status of an individual mouse in a group, is manifested as barbering and fighting. Barbering is commonly a condition seen in group-housed mice, where the dominant mouse nibbles off the whiskers and hair around the muzzles and eyes of cage mates. No other lesions are present, and only one mouse (the dominant one) retains all of its fur. Removal of the dominant mouse stops barbering; frequently, however, another mouse then assumes the dominant role. Barbering occurs during acts of mutual grooming in which one member of a mouse pair grasps individual whiskers or hairs with its incisors and plucks them out. Although plucking appears painful, recipients are passive in accepting barbering and even pursue conspecifics for further grooming.85 Barbering may also be seen associated with sexual overgrooming as a form of stress-evoked behavior, and lactating mice may display “maternal” barbering, produced in the process of suckling pups, in which alopecia is seen from tail to chin.47 Although rare in mice, consider infection with Trichophyton mentagrophytes in cases of alopecia that involve face, head, neck, or tail in all or just a few mice.22 Barbering has a genetic-based behavioral background, as aggressive inbred strains of mice do not use barbering in their behavior.47 In pet mice, barbering is often seen in female mice caged together. Male mice except littermates raised together from birth are more likely to fight, often very savagely, and inflict severe bite wounds on one another, especially over the rump, tail, and shoulders. Mechanical abrasion resulting from self-trauma on cage equipment is a form of husbandry-related alopecia. Small patches of alopecia appear on the lateral surfaces of the muzzle, resulting from chaffing on metal feeders, poorly constructed watering device openings, and metal cage tops. Unlike barbering, dermatitis may also be associated with the alopecic area. Treatment consists of replacing the poorly constructed equipment with nonabrading equipment. Individually housed mice can display aberrant stereotypic behaviors such as polydipsia and bar chewing, which result in mechanical abrasion and alopecia. In these cases, environmental enrichment must be
CHAPTER 27 Disease Problems of Small Rodents
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Table 27-2 Drugs for Treating Endo- and Ectoparasites in Pet Rodents Active Substance
Dosage
Application
Chlorpyrifos Chlorpyrifos + Fenoxycarb
6 g per 27 x 48-cm cage —
Mix in bedding Spray
Deltamethrin Dichlorvos
Dependent on body weight 5 g/kg mix/intersperse
Topical Environment
Doramectin Fenthion Fipronil
0.5 mg/kg 10 mg/kg ~7.5 mg/kg topically = 1 to 2 sprays in gloved hand 10 mg/kg 10 + 1 to 2 mg/kg
Subcutaneous Spot on Spray
Moxidectin
10 + 50 mg/kg 0.2 to 0.4 mg/kg 10% solution (propylene glycol) 0.5% solution at 0.5 mg/kg
Spot on Subcutaneous Topical Topical
Permethrin
0.5% solution at 2 mg/kg Dependent on size of animal
PO Topical
Remarks
Ectoparasites
Imidacloprid Imidacloprid + Moxidectin Imidacloprid + Permethrin Ivermectin
Permethrin + Methopren Propoxur Pyrethrins
Spot on Spot on
Spray Dependent on size of animal
Topical Spray Topical
Selamectin
15 mg/kg
Spot on
Doramectin Fenbendazole Metronidazole Niclosamide Praziquantel
0.2 mg/kg 20 mg/kg 10 to 40 mg/kg 100 mg/kg 25 mg/kg
Subcutaneous Oral Oral Oral Oral
Toltrazuril
0.5% solution at 10 to 20 mg/kg
Oral
Used to treat environment against ectoparasites (insecticide + insect growth regulator) Shampoo: wash body completely Mix granules in bedding or hang strip 15 cm above cage for 24 hours, then 2x week for 3 weeks For fur mites, 3 times one week apart Once every 4 weeks Whole body, repeat after 7 to 10 day. For fleas and ticks. 0.1 mL/kg of 10% solution 0.1 mL/kg of 10% solution. Flea species are used as an intermediate host and vector for tapeworms Against fleas and ticks 3x every 7 to 10 days 1 drop behind the ear Repeat in 10 days. For Myocoptes musculinus Repeat in 15 days. For Radfordia affinis Powder: cover body with powder or cotton ball soaked in 5% solution Used to treat environment against ectoparasites (insecticide + insect growth regulator) Flea powder: use to cover with dust Use “kitten-safe” product. Used to treat environment Use 0.05% shampoo every 7 days for four treatments Strictly topical. Can be used for fleas and lice. Repeat in 10 days
Endoparasites
provided. Food items such as Lafeber parrot Nutriberries, Avicakes or Nutriforage (www.lafebervet.com) and toys such as running wheels or hollow tubes are helpful. Most infectious causes of alopecia and dermatitis are associated with fur mites. Ectoparasites are common in mice purchased from pet stores.79 In affected animals, the hair is generally thin, especially on difficult-to-groom areas such as the head and trunk. The coat often has a greasy appearance; in cases of heavy infestation, noticeable pruritus and self-inflicted dermal ulceration may occur. Three mites are commonly seen: Myobia musculi, Myocoptes musculinus, and Radfordia affinis. The
For Syphacia muris For pinworms, twice 5 days apart For protozoa, twice 5 days apart Repeat after 7 days Repeat twice every 10 to 14 days for three treatments. Stings if given SC Give daily for 2 to 3 days, pause 5 days and then repeat
most clinically significant mouse mite is M. musculi. Infestations are usually caused by more than one species. Mites are spread by direct contact with infected mice or infested bedding. Diagnosis is based on the identifying adult mites, nymphs, or eggs on hair shafts with the use of a hand lens or a stereoscopic microscope. Adults and nymphs appear pearly white and elongated (being about twice as long as they are wide); eggs are oval and seen attached to the base of hairs or inside mature females. Treat mite infestations with avermectins (e.g., ivermectin, selamectin) or milbemycins (moxidectin). Ivermectin (0.2 mg/ kg SC or PO) twice at 10-day intervals is effective. Selamectin
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Fig. 27-1 Progressive necrosing dermatitis of the pinna in a mouse. The intense pruritus often causes the mouse to excoriate the area next to the ear pinna and the lesion then extends from the ear to the neck and over the shoulders. The condition is similar to idiopathic ulcerative dermatitis a well-recognized disease with a characteristic distribution on the thorax and head. Both conditions are characterized by severe pruritus, self-mutilation, dermal ulceration, necrosis, and fibrosis. The cause is an underlying vasculitis attributed to immune complex deposition on dermal vessels, although neurogenic abnormalities have been proposed for progressive necrosing dermatitis of the pinna.
(10-12.5 mg/kg topical)35 or moxidectin (0.5 mg/kg topical78 or 2 mg/kg PO77) administered twice at an interval of 10 to 15 days are also effective (see Table 27-2). Sometimes an owner presents a single pet mouse negative for primary ectoparasitic, bacterial, or mycotic infections with severe pruritus characterized by self-mutilation, dermal ulceration, necrosis, and fibrosis. Idiopathic ulcerative dermatitis is a well-recognized disease in black laboratory mice on a C57BL strain background with a characteristic distribution on the thorax and head. The cause is an underlying vasculitis attributed to immune complex deposition on dermal vessels.48 Dietary factors and dysregulated fatty acid metabolism have been implicated in the development of the disease and the severity appears to be modulated by dietary fat and vitamin E content. Gavaging affected mice with 0.1 mL per day of liquid from an essential fatty acid supplement containing omega-3 fatty acids was associated with regressed lesions and resolved pruritus in a small sample of affected mice.60 We have obtained good treatment results using a generic omega-3 fatty acid supplement at 0.1 to 0.2 mL PO q24h. The ulcers may heal with fibrosis and resulting skin contracture or progress to a Staphylococcus xylosus secondary bacterial infection.48,100 Progressive necrosing dermatitis of the pinna in mice, similar to idiopathic ulcerative dermatitis, may also be seen (Fig. 27-1).94 It occurs in outbred mice, and there is no strain background association. Initially a lesion on the dorsum of the pinna, resembling an engorged blood vessel or slight erythema, oozes serum and peripheral necrosis begins. Several days later the necrotic area sloughs and the pinna is left notched. In severe cases, the site becomes secondarily infected, the lesion becomes pruritic, and the mouse self-mutilates from the ear to the neck and over the shoulders. Treat these mice topically twice daily
with of 0.2% cyclosporine in 2% lidocaine gel supplemented with 50 μg/mL gentamicin. Ringworm, caused by Trichophyton mentagrophytes, is uncommon in pet mice. Lesions, when present, are most common on the face, head, neck, and tail. The lesions have a scurfy appearance with patchy areas of alopecia and variable degrees of erythema and crusting. Pruritus is usually minimal to absent and the lesions do not fluoresce under a Wood’s lamp.22 Skin swellings are usually tumors or abscesses. Needle biopsy often reveals the nature of the contents and allows diagnosis. Three opportunistic bacterial pathogens—Staphylococcus aureus, Pasteurella pneumotropica, and S. pyogenes—are often isolated3 and can cause abscesses in other organs (e.g., P. pneumotropica is sometimes associated with conjunctivitis, panophthalmitis, and swollen eye abscesses). Antibiotic therapy with penicillins or cephalosporins, concurrent with drainage and debridement of the abscess, is effective. The most common spontaneous tumors associated with the skin are mammary adenocarcinomas, followed by fibrosarcomas. The incidence of mammary tumors varies according to the mouse strain and the presence or absence of mouse mammary tumor viruses; the incidence is as high as 70% in some strains.98 In wild and outbred mice, the incidence of fibrosarcomas ranges from 1% to 6%.41 Subcutaneous tumors are nearly always malignant and have often ulcerated by the time a diagnosis is made. Tumors can be treated by surgical excision, but the chance of recurrence is high and the prognosis is poor. Attempts to treat skin tumors in pet mice by radiation or chemotherapy have not been reported. Digestive System. Endoparasites are relatively common in mice. However, only two parasites regularly encountered in the digestive tract, the protozoan parasites Spironucleus muris and Giardia muris, are considered pathogenic, even though they are not associated with clinical signs in immunocompetent hosts. Diagnosis is based on demonstrating characteristic trophozoites in wet mounts of fresh intestinal contents or feces. Treatment is metronidazole (two treatments of 10-40 mg/kg PO q5d) (see Table 27-2). Pinworms are ubiquitous, considered nonpathogenic, and found frequently in mice purchased from a pet store.16 Two are commonly encountered in mice: Syphacia obvelata and Aspicularis tetraptera. Often the only indication of pinworm infestation is rectal prolapse due to straining. To establish a diagnosis of S. obvelata infestation, make a clear cellophane tape impression of the perianal skin. Adult S. obvelata females deposit ova around the anus, whereas A. tetraptera does not deposit its ova in this area and fecal smear or flotation is required to confirm a diagnosis. Ivermectin (2.0 mg/kg PO given twice at a 10-day interval) eliminates pinworms from mice. Ivermectin 1% is diluted 1:9 in vegetable oil to establish a concentration of 1.0 mg/mL; affected mice are dosed with a volume of 0.2 mL/100 g PO.31 The recommended package label dose for mice with ectoparasites (0.2 mg/kg given twice at a 10-day interval) does not eliminate pinworms (see Table 27-2). Diarrhea is not usually seen in adult mice. Digestive disease in adult mice usually is caused by a varying combination of pathogenic and opportunistic infectious agents. Fecal flotation and fresh wet mounts of feces usually yield positive results and do not necessarily give a definitive diagnosis. However, these techniques are sometimes helpful in identifying heavy endoparasite infestations. Treatment is generally directed at clinical signs and consists of the judicious use of antimicrobials.
CHAPTER 27 Disease Problems of Small Rodents Respiratory System. Diseases of the upper and lower respiratory tracts are common in pet mice and rats. Animals may be presented with sniffling, sneezing, chattering, and labored breathing. If dyspnea is suspected, do not overhandle the animal during clinical examination, as it may die. Collection of tracheal and nasal secretions is not recommended because swabbing is highly traumatic and the cause of disease is generally a mixed viral, mycoplasmal, and bacterial infection. Antibiotic treatment is helpful but does not eliminate the disease. The two most common causes of clinical respiratory disease in mice are Sendai virus and Mycoplasma pulmonis. Sendai virus is associated with an acute respiratory infection in which mice display chattering and mild respiratory distress. Neonates and weanlings may die. Adults generally recover within 2 months. When the disease’s expression exceeds this pattern, the cause is most likely concurrent mycoplasmal infection. Mycoplasma pulmonis is the cause of chronic pneumonia, suppurative rhinitis, and occasionally otitis media. Chattering and dyspnea are caused by accumulations of purulent exudate in inflamed and thickened nasal passages. Survivors develop chronic bronchopneumonia and bronchiectasis and may develop pulmonary abscesses. Antibiotic therapy may alleviate clinical signs but does not eliminate the infection. Enrofloxacin (10 mg/kg) as an antimicrobial agent in combination with doxycycline hyclate (5 mg/kg) as an immunomodulator (not as an antimicrobial) given every 12 hours PO for 7 days is helpful. Additional treatments such as nebulization therapy, expectorants, and nonsteroidal anti-inflammatory drugs are helpful to ameliorate the disease. Urinary System. Obstruction of the urethra in male mice has been described as resulting from infections of the preputial glands with S. aureus and of the bulbourethral glands with P. pneumotropica. Accessory sex gland secretions and, rarely, urolithiasis have also been implicated. Mice are often presented to the veterinarian because they mutilate their penises as a result. In addition, occasional injury of the penis is seen in young males from aggressive breeding activity and abrasion on the cage. Treatment involves isolating the affected mouse, cleaning and debriding the affected areas, and treating the animal with antibiotics. Theriogenology. Mice experience relatively few complications associated with parturition. Occasionally vaginal and uterine prolapse occurs after parturition. With the animal under anesthesia, clean the prolapsed tissue with isotonic saline, place a lubricated 20-gauge Teflon intravenous catheter (without needle) into the lumen of the uterus and vagina, and manually manipulate the tissue back into the proper anatomic position.12 A purse-string suture with 4-0 polyglycolic acid suture can be placed around the vaginal orifice before removing the catheter. Absorbent cotton or cotton wool is not recommended as bedding material as it may wrap around the legs of suckling mice and cause necrosis and sloughing of limb extremities.76
Rats Integumentary System. Ectoparasitic infestation is more common in rats than in mice. Occasionally the fur mite Radfordia ensifera is seen. Although R. ensifera infestation produces few ill effects, heavy infestation may lead to self-traumatization and ulcerative dermatitis. Other mites, including Demodex species, have been described in rats maintained in laboratories; however, they are seldom seen, and no contemporary reports of infestations in pet rats appear in the literature.107 The tropical rat mite
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Fig. 27-2 The tropical rat mite Ornithonyssus bacoti. Engorged on the host’s blood, the mite appears red. It is an opportunistic ectoparasite often found on pet rats, mice, gerbils, and hamsters. It spends a short time on a host, penetrating the skin only for feeding. Severe infestations can resemble a rodent covered in fine sawdust; the sawdust is thousands of mites. Affected rodents can experience anemia, debilitation, and even death. When an animal host is not available, humans can become the victims of mite infestation. Ornithonyssus bacoti is an opportunistic ectoparasite often found on pet rats, mice, gerbils, and hamsters (Fig. 27-2). It spends a relatively short time on a host and penetrates the skin for feeding only. Severe infestations can cause anemia, debilitation, and death in rodents. When an animal host is not available, humans can become the victims of mite infestation. Locating the resident host of the mite is critical in its successful elimination. Detect adult parasites by macroscopic examination of the host; the mites are orange red. Rats with ectoparasites can be treated with selamectin (15 mg/kg as topical spot on, repeat in 10 days). The environment can be decontaminated with either synthetic pyrethroids or fipronil (see Table 27-2).6 Ringtail is a pathologic condition of the tail of young rats that is typically characterized by dry skin and the formation of annular constrictions. In severe cases, blood vessels distal to the constrictions thrombose, resulting in pain and necrotic tissue. Autoamputation may result. This lesion is highly photogenic and probably for this reason is always described in textbooks and articles on diseases of rats. It occurs primarily during the preweaning period in rats aged 2 to 19 days and occasionally in young mice. Low environmental relative humidity (less than 40%) appears to be the cause, and it is more often seen in rats housed in hanging cages; it is rarely seen in pet rats. If ringtail is diagnosed, treatment options involve adding unsaturated fatty acids to the diet (e.g., 5% corn oil or an essential fatty acid supplement containing omega-3 fatty acids) or topical application of lanolin.32,102 If avascular necrosis has occurred, amputate the tail below the necrotic annular constriction. Ulcerative dermatitis caused by S. aureus infection results from self-traumatization associated with fur mite infestation or, more commonly, from scratching of the skin over an inflamed salivary gland (see sialodacryoadenitis virus under “Digestive System,” below). Rats have a remarkable ability to resist infection
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Fig. 27-3 Mammary fibroadenoma in the inguinal region of a 355-g female rat. The excised tumor weighed 40 g and represented 11% of the rat’s body weight.
with S. aureus.23 Treatment consists of clipping the toenails of the hind paws, cleaning the ulcerated skin, and applying a topical antibiotic. Systemic treatment is rarely necessary. The most common subcutaneous tumor in the rat is fibroadenoma of the mammary glands. The distribution of the mammary tissue is extensive, and the tumors can occur anywhere from the neck to inguinal region (Fig. 27-3). Tumors can reach 8 to 10 cm in diameter and occur in both males and females. Adenocarcinomas represent fewer than 10% of mammary tumors in pet rats. Prevention and treatment of mammary fibroadenomas involves surgery and medical treatment. The surgical technique for mammary tumor removal ranges from straightforward to complicated (for high cervical or inguinal tumors) depending on the location and size of the tumor (see Chapter 28). However, the recurrence of fibroadenomas is common in uninvolved mammary tissue, and often several surgeries are required. The frequency of mammary tumors is significantly lower in ovariectomized versus sexually intact rats.43 Neutering sexually mature females often reduces incidence of tumor recurrence. While tumors do not metastasize, death is often caused by large tumors that ulcerate and become secondarily infected. Digestive System. Sialodacryoadenitis virus, a coronavirus, causes inflammation and edema of the cervical salivary glands. Owners of infected rats often describe their pets as having mumps. Sialodacryoadenitis virus is highly contagious and initially causes rhinitis, followed by epithelial necrosis and inflammatory swelling of the salivary and lacrimal glands. Cervical lymph nodes also become enlarged. There is no treatment for this disease. Glandular healing follows within 7 to 10 days and clinical signs subside within 30 days, with few residual lesions remaining. During acute inflammation, affected rats are at high risk for anesthesia-related mortality because of the decreased diameter of the upper respiratory tract lumen; also, ocular lesions such as conjunctivitis, keratitis, corneal ulcers, synechiae, and hyphema can develop secondary to lacrimal dysfunction. The eye lesions usually resolve but occasionally pro gress to chronic keratitis and megaglobus.
Respiratory System. Respiratory disease caused by infectious agents is the most common health problem in rats. Three major respiratory pathogens cause overt clinical disease: Mycoplasma pulmonis, Streptococcus pneumoniae, and Corynebacterium kutscheri. Other organisms such as Sendai virus (a paramyxovirus), pneumonia virus of mice or PVM (a paramyxovirus), rat respiratory virus (a hantavirus), cilia-associated respiratory (CAR) bacillus, and Haemophilus species are minor respiratory pathogens that rarely cause overt clinical disease by themselves. However, the minor respiratory pathogens interact synergistically as copathogens with the major respiratory pathogens to produce two major clinical syndromes: chronic respiratory disease (CRD) and bacterial pneumonia. The best-understood multifactorial respiratory infection in rats is CRD. The major component of CRD is M. pulmonis, and the disease is also known as murine respiratory mycoplasmosis (MRM). Rats may live 2 to 3 years with CRD. While M. pulmonis is rarely seen in laboratory rats, serologic test results of pet rats are usually positive. A survey of 28 pet ratteries in the United States found that all were positive for M. pulmonis.17 Clinical signs are highly variable: in many cases no signs are present even though significant pulmonary lesions may exist. The prevalence and severity of signs typically increase with the age of the rat and the presence of environmental stresses placed upon the animal. Initial infection commonly occurs without any clinical signs; early signs involve both the upper and the lower respiratory tracts and may include snuffling, nasal discharge, polypnea, weight loss, hunched posture, ruffled coat, head tilt, and red tears.110 The most important aspect of CRD for clinicians is that respiratory mycoplasmosis varies greatly in disease expression because of environmental, host, and organismal factors that influence the host-pathogen relationship. Examples of such factors include intracage ammonia levels, concurrent Sendai virus, coronavirus (sialodacryoadenitis virus), PVM, rat respiratory virus and/or CAR bacillus infection, the genetic susceptibility of the host, the virulence of the Mycoplasma strain, and vitamin A or E deficiency.110 Auscultation is insensitive in determining the severity of respiratory disease, and radiographs are often unremarkable. However, CT scan often reveals significant pulmonary disease. Serology is the preferred choice for diagnostic testing, as Mycoplasma is difficult to grow in a laboratory. Younger rats naturally exposed to M. pulmonis may be seronegative for up to 4 months postexposure.21 The primary lesion of CRD is subacute and chronic bronchitis (SACB), a chronic inflammatory condition resulting in respiratory epithelial dysfunction.50 The underlying airway inflammation and clinical signs result from damage and remodeling of airway epithelium, colonization of the airways with secondary infections, and infiltration and activation of neutrophils, macrophages and lymphocytes. Bronchodilators are the primary treatment for SACB. Inhaled nonselective muscarinic antagonists (e.g., ipratropium bromide) and beta-2 adrenergic agonists (e.g., albuterol, salmeterol) as well as oral theophylline provide significant although modest efficacy. Frequent treatment with broad-spectrum antibiotics is necessary, as microbe colonization is a common finding. Tetracycline antibiotics like doxycycline are efficacious in CRD. Treating with doxycycline (5-10 mg/kg PO q12h) or a long-acting depot such as doxycycline (Vibravenos, Pfizer Animal Health, www.Pfizer.com), 70-100 mg/kg SC or IM q7d) often helps affected rats. Bacterial pneumonia is nearly always caused by S. pneumoniae (Fig. 27-4) but seldom in the absence of some combination
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Fig. 27-5 Gross appearance of the lungs in a rat with pneumonia pneumoniae in a rat.
caused by Mycoplasma pulmonis. Compare the appearance of these lungs with that of the lungs in Fig. 27-4. Rats in the later stages of M. pulmonis chronic respiratory disease may show pulmonary abscesses.
involving M. pulmonis (Fig. 27-5), Sendai virus, or CAR bacillus.110 Infection with C. kutscheri also results in pneumonia but only in conjunction with debilitation or immunosuppression.110 In pet rats, immunosuppression can result from diabetes, neoplasia, or dietary deficiencies. Corynebacterium kutscheri pneumonia is rare in pet rats. Pneumonia caused by S. pneumoniae can be of sudden onset. Young rats are more severely affected than are older ones, and often the only sign they exhibit is sudden death. Mature rats may demonstrate dyspnea, snuffling, and abdominal breathing. A purulent exudate may be seen around the nares and on the front paws (from wiping of the nostrils). A tentative diagnosis is based on the identification of numerous gram-positive diplococci on a Gram stain of the exudate or in a sample submitted for cytologic examination. Severe bacteremia is an important consequence of advanced disease and results in multiorgan abscesses and infarction. Treatment must be aggressive, and the use of beta-lactamase-resistant penicillins such as cloxacillin, oxacillin, and dicloxacillin (all of which can be administered orally) is recommended. We use amoxicillin/clavulanic acid (13.75 mg/kg PO q12h) as our first antibiotic choice. Urinary System. Obtaining a blood sample is important for aging rats as they often suffer from chronic renal disease. Blood urea nitrogen (BUN) concentration can be estimated with the use of a blood dipstick, and proteinuria can be detected on urine dipstick analysis. Chronic progressive nephrosis (CPN) is the best known age-related disease in rats. In CPN, the kidneys are enlarged and pale and have a pitted, mottled surface that often contains pinpoint cysts. Lesions consist of a progressive glomerulosclerosis and myriad tubulointerstitial disease primarily involving the convoluted proximal tubule.36 The most striking change in renal function is proteinuria exceeding 10 mg/ day that increases in severity progressively with age. The features of CPN are qualitatively similar among different strains of laboratory rats, but the onset, incidence, and severity of the disease
vary considerably. The disease occurs earlier and is of greater severity in males than in females: urinary protein excretion averaging 137 mg/day has been documented in 18-month-old male Sprague-Dawley rats, whereas excretion averaging 76 mg/day was reported in female rats of the same age.91 Dietary factors appear to have an important role in the progression of CPN. Caloric restriction, the feeding of low-protein diets (4%-7%), and limiting the source of dietary protein reduce the incidence and severity of CPN. Feeding soybean protein (as opposed to casein) and caloric restriction contribute substantially to reducing the incidence and severity of CPN; low-calorie diets that contain high protein levels do not decrease the incidence or severity. Drugs and exposure to chemicals can also exacerbate CPN. Treatment is supportive and involves feeding a low-protein diet and administering anabolic steroids. Uroliths and renal pelvic calculi are relatively infrequent in rats, especially compared with guinea pigs, rabbits, and chinchillas. Most uroliths that have been described were composed of struvite.71 Musculoskeletal and Peripheral Nervous System. Clinically, old rats show disturbances in motor function, posterior paresis and paralysis, loss of tail control, incontinence, and weight loss. Distinguishing between age-related peripheral or central nervous system changes, neurogenic muscular atrophy, and primary age changes in muscles is very difficult. Posterior paresis is commonly caused by spinal nerve root degeneration (also known as radiculoneuropathy, polyradiculoneuropathy, and degenerative myelopathy).112 Spontaneous tumors of the peripheral nervous system in rats are very rare; only isolated cases are reported in extensive tumor incidence surveys.1 Allowing young rats to stand on their hind legs frequently is a predisposing factor in the development of avascular necrosis of the femoral head (Legg-Calve-Perthes disease).68 Ocular System. The harderian glands of rats are located behind the eyes. These glands secrete various porphyrins
Fig. 27-4 Fibrinopurulent pneumonia caused by Streptococcus
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Fig. 27-6 Chromodacryorrhea, or red tears, in a rat. The color results from porphyrin pigments in the Harderian gland secretions, which are visible around the eyes and occasionally the nares.
that give the tears a reddish color. Harderian gland secretion increases in response to stress and disease, and the tears themselves dry around the eyes and external nares (the nasolacrimal duct drains into the nasal cavity), resembling crusts of blood (Fig. 27-6). Owners commonly report bleeding from the eyes and noses of their pet rats. The porphyrins fluoresce under ultraviolet light and can be readily differentiated from blood with a Wood’s lamp (Fig. 27-7). This condition is known as chromodacryorrhea, or red tears; although it is not pathologic, it is a consequence of an acute onset stress such as that caused by pain, illness, or restraint.19 Red tears are often an indication of a chronic underlying disease, and their presence warrants a thorough evaluation of the affected animal.
Hamsters Although many species of hamsters live in the wild, only a few types are kept as pets. The most common pet hamster is the golden or Syrian hamster (Mesocricetus auratus), which has been kept as a pet since the 1940s. Although two other species of hamsters, the common or European hamster (Cricetus cricetus) and the rat-like Chinese hamster (Cricetulus griseus) are used in research, they do not make good pets because of their aggressive nature. However, dwarf hamsters such as the Djungarian (Phodopus sungorus) and Roborovsky’s (Phodopus roborovskii) are being seen increasingly as pets because they have a docile disposition, do not attempt to bite or run away, and do well in captivity. There is some confusion over the common names of the species of Phodopus. Until about 1980, P. sungorus was considered to have two subspecies—P. sungorus sungorus and P. sungorus campbelli. Phodopus sungorus sungorus was known as the Djungarian hamster. There was no commonly used name for P. sungorus campbelli. Then studies showed that these two subspecies were in fact different species, and they were renamed as P. sungorus and P. campbelli. The name “Siberian hamster” was applied to P. sungorus and “Djungarian hamster” to P. campbelli. Integumentary System. The most common skin problem seen in Syrian hamsters is hair-coat roughness. This is a nonspecific sign of fighting, aging, and a variety of diseases. Female Syrian hamsters are heavier than males and are generally aggressive not only toward other hamsters but also toward their owners; they can inflict severe bite wounds on cage mates. Nonestrous
Fig. 27-7 Chromodacryorrhea in a rat viewed under an ultraviolet light. The porphyrins in the Harderian gland secretions around the eyes and nares readily fluoresce under a ultraviolet light and can be distinguished from blood.
females can be especially aggressive toward young males and may kill them. Length of hair in the long-haired Syrian hamster (“Teddy Bear” hamster) is influenced by testosterone. Longhaired males from the age of sexual maturity have significantly longer hair than females or castrated males, which display fluffy, shorter hair.86 Syrian hamsters possess paired flank organs in the costovertebral area that are androgen-dependent and consist of sebaceous glands, pigmented cells, and terminal hairs. They are larger and heavily pigmented in males and used for territorial marking. Melanomas, not only of the flank organ but also of the skin, are frequently reported in Syrian hamsters. There is a striking 10:1 male:female melanoma ratio.103 Male hamsters have large, pendulous testes, which clients may mistake for tumors. Djungarian hamsters show a high prevalence of neoplastic disease (five times greater than Syrian hamsters), and most tumors are integumental (e.g., mammary tumors, atypical fibromas, and papillomas).52 Bacterial pseudomycetoma has been described in several dwarf hamsters; treatment is excision.26 Digestive System. Hamsters have distensible cheek pouches that may be mistaken for lesions by owners.8 Sometimes the cheek pouches become impacted, and removal of the material from the pouch with fine forceps is necessary. A radiograph of the head often shows the extent of the impaction. Predisposing causes of impaction, such as malocclusion of incisors or molars, should be investigated. The most common problems seen in pet hamsters are enteropathies. Diarrhea may occur in hamsters of any age and is known as “wet-tail,” although this euphemism is frequently used to describe the disease in young hamsters. Proliferative ileitis is the most significant intestinal disease of 3- to 10-week-old hamsters and results in high mortality. It is caused by the intracellular bacterium Lawsonia intracellularis, which is also responsible for proliferative enteropathy in pigs and ferrets.59 Treatment must be aggressive and involves correcting life-threatening electrolyte imbalance, administering antibiotics, and force-feeding. Several antibiotic treatments are recommended, including tetracycline-hydrochloride (400 mg/L of drinking water for 10 days), tetracycline (10 mg/kg PO q12h for 5-7 days), enrofloxacin (10 mg/kg PO or IM q12h for 5-7 days), and trimethoprim-sulfa combination (30 mg/kg PO q12h for 5-7 days). Symptomatic
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Fig. 27-8 Rectal prolapse in a Syrian hamster associated with Lawsonia intracellularis infection, also commonly known as “wet-tail.”
treatment with bismuth subsalicylate may be given if diarrhea persists. Replacement electrolyte and glucose solutions should be given orally, and electrolyte fluid replacement such as saline or LRS should be given at a dose 20 mL/100 g q24h. Diagnosis often depends on necropsy and histologic examination, although fecal polymerase chain reaction (PCR) assays have been developed.75 Sequelae to proliferative ileitis in surviving hamsters may include eventual obstruction, intussusception, or rectal prolapse (Fig. 27-8) (see Chapter 28).15 Diarrhea in adult hamsters is associated with Clostridium difficile enterotoxemia and may occur 3 to 5 days after the administration of antibiotics such as penicillin, lincomycin, or bacitracin.39 Detection of C. difficile by PCR is highly sensitive and can discriminate between toxigenic and nontoxigenic strains of the organism by detecting its toxin-producing genes. Oral administration of bovine antibodies against toxigenic C. difficile has been shown to protect hamsters against experimental antibiotic-associated enterotoxemia.63 Tyzzer disease caused by Clostridium piliforme has been described in hamsters and gerbils obtained from a pet store supplier.69 The supplier’s hamsters and gerbils had a high mortality rate but the rats and mice did not. Affected rodents were depressed, dehydrated, and had scruffy coats and diarrhea; many animals had no clinical signs before death. Clinical outbreaks appear to be precipitated by severe stress, including that caused by overcrowding, high environmental temperature and humidity, heavy internal and external parasite load, and nutritionally inadequate diets despite the prophylactic treatment of drinking water with oxytetracycline. Tyzzer disease is frequently listed as an intestinal disease of rodents and other animals in laboratory animal textbooks. However, the actual prevalence of the infection in contemporary rodents remains unknown. The report concerning the pet store illustrates the opportunistic nature of C. piliforme in immunosuppressed animals. The disease is not seen in healthy immunocompetent animals. Weight loss is seen in older hamsters and is often associated with hepatic and renal amyloidosis. One research report described amyloidosis in 88% of hamsters above 18 months of age.34 Amyloidosis is the principal cause of death in long-term
Fig. 27-9 End-stage hepatic cysts in a Syrian hamster.
research studies, and female hamsters have a higher incidence, increased severity, and earlier age of onset of amyloidosis than do male hamsters.87 In laboratory hamsters, social stress induced by crowding is correlated with amyloidosis.33 The incidence and clinical signs of disease are not described in pet hamsters, because overcrowding is not a problem with pet hamsters, the incidence may be low. However, clinicians would expect edema and ascites caused by hypoproteinemia of hepatic and renal origin. If amyloidosis is diagnosed in a pet hamster, the prognosis is poor and treatment is supportive. Multiple thin-walled cysts of varying sizes (0.25-3.0 cm) and shapes are occasionally found in the livers of hamsters (Fig. 27-9). Initially affected hamsters show no clinical signs, but as the cysts enlarge, affected animals show abdominal enlargement, diffuse alopecia, and wasting.27 The cysts are often found on abdominal palpation, radiographic or ultrasonographic examination, or laparotomy. Grossly they protrude from the liver surface and contain a clear serous fluid. Hepatic parenchyma surrounding the cysts often shows pressure atrophy, necrosis, engorged sinusoids, hemorrhage, mild to extensive fatty or vacuolar degenerative changes, and occasionally proliferation of biliary ducts.96 Such animals are generally over 2 years of age, and cystic proliferations may be found in other abdominal organs. The lesions are caused by developmental defects of the bile duct. Respiratory System. In response to a survey, 6 of 14 laboratories in the United States reported pneumonia as the second most common clinical condition in hamsters after diarrhea.82 An earlier survey conducted in Germany noted respiratory infections in 8% of all clinical conditions in hamsters.61 Histologic evidence of bronchopneumonia resembling bacterial pneumonia and of interstitial pneumonia resembling viral pneumonia has been described, but there are no reports of observed clinical cases. Consequently other authors have stated that respiratory disease is uncommon in hamsters.38 The true prevalence remains to be established.
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Fig. 27-10 Postovulatory discharge drawn out as a thread from a female hamster. The appearance of the discharge marks the end of estrus and start of diestrus.
Fig. 27-11 Atrial thrombosis (arrow) in a Syrian hamster.
Purulent rhinitis associated with pneumonia and gluey eyelids has been described in hamsters and is associated with a poor prognosis.56 Children may inadvertently transmit bacterial pneumonias, especially those caused by Streptococcus species, to pet hamsters. Rapid diagnosis can be made by identifying the characteristic gram-positive diplococci on a Gram’s stain of nasal and ocular discharges. Follow-up culture and treatment with chloramphenicol (chloramphenicol palmitate, 50 mg/kg PO q8h; chloramphenicol succinate, 30 mg/kg IV or IM q8h) are recommended until antibiotic sensitivity results are available. Reproductive System. Female hamsters have a 4-day estrous cycle characterized by a copious postovulatory discharge at the end of the cycle. The discharge is creamy white and has a distinctive odor; it fills the vagina and usually extrudes through the vaginal orifice (female hamsters have three orifices: urinary, genital, and anal). Its stringy nature is distinctive; if touched, it can be drawn out as a thread about 4 to 6 in. long. Owners often describe the discharge as pus and mistakenly believe it to be abnormal (Fig. 27-10). Pyometra has been observed clinically although rarely in pet hamsters. A tentative diagnosis is made by ultrasound examination of the abdomen; ovariohysterectomy is the treatment of choice. Cannibalism of the young accounts for about 95% of all preweaning mortality in group-housed laboratory female hamsters.82 Other factors such as cold ambient temperatures (below 10°C), lean diets, and low body weight (especially during pregnancy) appear to increase cannibalism.89,90 Instruct owners to give the mother ample food and water and to leave her alone in a quiet, warm place for at least 1 week or preferably 2 weeks. Disturbing the mother by handling the young or nest and not providing adequate nesting material, warmth, food, or water often results in desertion of the litter and cannibalism. Cardiovascular System. Atrial thrombosis has been described in aging research hamsters by many authors, and in certain strains it occurs with a high incidence (up to 73%).88 Most thromboses develop in the left atrium secondary to heart failure and lead to a consumptive coagulopathy (Fig. 27-11).
Although the incidence does not differ between the sexes near the end of their respective life spans, atrial thrombosis occurs on average at a younger age in females (13.5 months) than in males (21.5 months).109 Aged pet hamsters present with clinical signs of cardiomyopathy such as hyperpnea, tachycardia, and cyanosis. In untreated hamsters, death usually follows within a week after these signs become evident. The incidence of atrial thrombosis is influenced by the endocrine status of the animal and especially by the amount of circulating androgen. Thus, castrating male hamsters is linked to an increase in the prevalence of atrial thrombosis.92 Cardiomyopathy should be suspected in aged pet hamsters (older than 1.5 years) that present clinically with signs of tachypnea, lethargy, anorexia, and cold extremities.88 Diagnosis of cardiomyopathy in hamsters is based on clinical signs and results of radiography and ultrasound examination of the heart. Treatment of heart disease is symptomatic and involves empiric use of digoxin, diuretics, angiotensin-converting enzyme (ACE) inhibitors, and prophylactic anticoagulants. We base dosages of these drugs on ferret doses and monitor response closely. Verapamil, a calcium antagonist, administered at a dose increasing from 0.25 mg to 0.50 mg given SC q8h over 4 weeks, prevented severe myocardial lesions in untreated 2-month-old inbred female myopathic hamsters.57 Endocrine System. Surveys of spontaneous lesions in laboratory hamsters describe a high incidence of adrenocortical hyperplasia and adenoma.87 However, despite extensive histopathologic study, hyperadrenocorticism or Cushing’s disease has been reported in only four hamsters, with high serum cortisol concentrations documented in only one of the four.5,64 Hamsters with clinical signs resembling those of Cushing’s disease are occasionally seen in practice. Diagnosis is based on identifying classic signs similar to those seen in dogs, such as a history of polydipsia, polyuria, and polyphagia; clinical signs of alopecia and hyperpigmentation; and high concentrations of plasma cortisol and serum alkaline phosphatase. Normal hamster cortisol concentrations are low compared with those of other species and range from 0.5 to 1.0 mcg/dL in normal
CHAPTER 27 Disease Problems of Small Rodents males and females.108 Research has suggested that hamsters may secrete both cortisol and corticosterone.72 Therefore meaningful measurement of plasma cortisol concentrations in hamsters is empiric at present. If hyperadrenocorticism is suspected, the cause—such as hypersecretion by a functional tumor, primary adrenal hyperplasia, or excess adrenocorticotropic hormone production—is often more difficult to determine. Hamsters respond to exogenous adrenocorticotropic hormone stimulation.108 The treatment of hamsters described in the clinical reports indicated that ketoconazole (5 mg/kg q12h for 1 month) did not work, o,p’-DDD, now known as mitotane (5 mg PO q24h for 1 month) did not work, and metyrapone (8 mg PO q24h for 1 month) worked in one of two hamsters. Further research needs to be done on this syndrome. Ocular System. Exophthalmos is commonly seen in hamsters and usually occurs because of ocular infection or trauma to the periorbital area or the application of excess pressure in holding the animal during restraint. Hamsters with sialodacryoadenitis may develop keratoconjunctivitis sicca and exophthalmos and subsequent proptosis. Occasionally, a hamster’s eye is displaced forward if the caregiver restrains the animal too tightly by holding the skin at the back of the neck. If the hamster is treated soon after the exophthalmos occurs, the prognosis for saving the eye is good. Cleanse the ocular area gently with an ophthalmic wash and lubricate the eye with sterile ophthalmic lubricant. Gently retract the lid margins around the globe until the eye returns to its normal position. Treat the eye with an antibiotic ophthalmic ointment for a minimum of 7 to 10 days. Occasionally, tarsorrhaphy will be needed to prevent recurrence. Enucleation may be necessary if the eye cannot be replaced or if significant trauma to the proptosed eye has occurred. Lymphoma. Lymphoma is the most common neoplasm in hamsters. Clinicians see three variations. In older hamsters, lymphoma is the most frequently observed neoplasm of the hematopoietic system.99 These tumors are often multicentric, involving the thymus, thoracic lymph nodes, mesenteric lymph nodes, superficial lymph nodes, spleen, liver, and other sites. Cytology of the tumors is variable. A second variation, cutaneous lymphoma, is seen in adult hamsters and resembles mycosis fungoides, an epidermotropic T-cell lymphoma in humans.40 Clinicians have described lethargy, anorexia, weight loss, patchy alopecia, and exfoliative erythroderma in affected animals. Pathologists have observed dense infiltrates of neoplastic lymphocytes in the dermis with extension into the epidermis. The third variation is an epizootic lymphoma in young hamsters caused by hamster polyomavirus (HaPV).4 When HaPV is first introduced into a naive population of breeding hamsters, it can result in epizootics of lymphoma, with an incidence as high as 80% among animals. Once enzootic in a hamster population, the occurrence of lymphoma declines to a much lower level. Enzootically infected hamsters develop HaPV skin tumors rather than lymphoma. Hamsters with HaPV lymphoma appear thin, often with palpable masses in their abdomens. Tumors associated with HaPV often arise in the mesentery but can also arise in the axillary and cervical lymph nodes. The tumors are often lymphoid, but erythroblastic, reticulosarcomatous, and myeloid types occur.
Gerbils Integumentary System. Facial eczema, sore nose, and nasal dermatitis all describe a common skin condition seen in gerbils. Clinical lesions adjacent to the external nares appear
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Fig. 27-12 Sore nose (facial eczema, nasal dermatitis) in a gerbil. This condition may result from an increase in Harderian gland secretion complicated by infection with Staphylococcus species.
erythematous initially; these lesions progress to localized alopecia and then to an extensive moist dermatitis (Fig. 27-12). The cause is believed to be an increase in the secretion of porphyrins by the Harderian gland (as in chromodacryorrhea in rats), which acts as a primary skin irritant.20 Various staphylococcal species (S. aureus and S. xylosus) may act synergistically to produce the dermatitis.95 Stress may cause excessive harderian gland secretion. Two examples of stress are overcrowding and exposure to an environmental humidity of greater than 50%, which causes the fur coat to stand out and appear matted. Gerbils require sand baths to keep their coats from becoming oily. Keeping the gerbil in a dry environment, cleaning its face, and providing soft clay or sand bedding instead of abrasive wood chip bedding will usually alleviate the problem. Use topical or parenteral antibiotics (except streptomycin) in gerbils with severe dermatitis. The tail of the gerbil is covered by thin skin. Unlike rats or mice, if a gerbil is picked up by the tip of its tail, the skin often slips off, leaving a raw, exposed tail that eventually becomes necrotic and sheds (Fig. 27-13).18 If the tail skin is lost, surgically amputate the bare tail where the skin ends. The tail usually sloughs if it is left untreated. In picking up a gerbil, take care to avoid grasping the tail unless it is gently held at the base. The best holding technique involves placing the palm of the hand over the gerbil’s back and encircling the body with thumb and fingers. Gerbils will bite if they are not handled securely, despite the claim in many reviews that they rarely bite human handlers regardless of provocation. Gerbils have large ventral abdominal marking glands that are androgen-dependent. Owners may mistake these normal glands for tumors. In aged animals, the gland may become infected or neoplastic. Local debridement and topical antibiotics are indicated for the treatment of infected glands. Do a wide excisional biopsy if you suspect a tumor such as adenocarcinoma.44 Digestive System. Tyzzer disease, due to Bacillus piliformis, is the most frequently described fatal infectious disease of gerbils.111 Common findings are sudden death or death after a short period of illness and the presence of multiple foci of hepatic necrosis. Diarrhea and gross and microscopic lesions in the intestinal tract are variably present. Experimentally induced Tyzzer disease in gerbils has confirmed that these animals are
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Degus
Fig. 27-13 A normal gerbil tail and a degloved gerbil tail caused by improper restraint of the gerbil by its tail.
extremely susceptible to infection.106 The probable route of infection is oral, explaining why gerbils exposed to infected bedding contract the disease. Gerbils will develop spontaneous, insidious periodontal disease if fed on standard rat or mouse diets for more than 6 months.104 On the same diets, about 10% of the animals become obese, and some may even develop diabetes. When feeding pelleted diet to gerbils, use diets labeled for gerbils (e.g., Mazuri [St. Louis, MO] makes a hamster/gerbil diet). Central Nervous System. Approximately 20% to 40% of gerbils develop reflex stereotypic epileptiform (clonic-tonic) seizures from around 2 months of age. The susceptibility is inherited, seen in selectively bred lines, and caused by a deficiency in cerebral glutamine synthetase.58 There is no treatment. Most animals outgrow the behavior with time. The seizures generally pass in a few minutes; they may be mild or severe and have no lasting effects. Reproductive System. Cystic ovaries are reported to occur frequently in laboratory Mongolian gerbils.70 Removal of the affected ovary is recommended. Ovarian granulosa cell tumors are the most common tumors in gerbils. In young gerbils (less than 2 years of age) tumors are often incipient and not macroscopically visible.37 However, in older gerbils, local invasion is seen in the ovarian hilum, periovarian fimbriae, and large ligaments. Metastases occur in the abdomen, with the omentum being the most affected organ. Metastases are not found in the thorax. The incidence of granulosa cell tumor is higher in virgin females than breeding females; therefore ovariectomy is recommended in pet gerbils that are not destined for breeding. Tumors and Aging. After 2 to 3 years of age, approximately 25% to 40% of gerbils develop neoplasia.65 After granulosa cell tumor, tumors of the skin are the next most frequent. Squamous cell carcinoma of the sebaceous ventral marking gland in males and melanoma, usually of the ear, foot, or base of the tail, are seen.81 Besides neoplasia, older gerbils have a high incidence of chronic interstitial nephritis.7 Aged gerbils have a remarkable propensity for the development of aural cholesteatoma, a nonneoplastic keratinizing epithelial mass that occurs in the middle ear and mastoid region; it erodes bone and invades the labyrinth and cranial cavity.13
Degus, or trumpet-tail rats, come from Chile. Taxonomically, they are in the same diverse order as guinea pigs and chinchillas. Degus have been used as laboratory animals for 20 years and in recent years have become popular as pets in the United States and Europe. They are highly social, demonstrating a broad array of communication methods that make them appealing as pets. Research studies with degus have produced a wealth of information that should make care of this species in captivity easy. However a study of 300 pet degus indicated that most of their diseases are caused by improper diet, selfmutilation, and improper handling45; consequently client education is critical. Degus are herbivorous rodents adapted anatomically and behaviorally to use a fibrous diet with moderate to low levels of nonstructural carbohydrate. Captive degus should consume foods containing nutrients comparable to those consumed by free-ranging animals.25 Like chinchillas, captive degus must be provided with dust baths twice a week.25 Although degus do not drink much water, owners should change their water bottles regularly to prevent bacterial overgrowth. The testicles of male degus are intra-abdominal and the method of castration is usually by laparotomy,28 although a prescrotal open technique has been described.11 Integumentary System and Behavior. Alopecia due to fur chewing is common, especially in younger degus (less than 2 years of age). In one report, alopecia was the second most frequent disorder seen in pet degus.45 Degus live in nature in groups of up to 10 animals and, if kept alone, especially in a cage without environmental enrichment, will develop stereotypical behavior and self-mutilation.105 Self-barbering of the medial aspect of the hind legs and of the forepaws is common. Alopecia may also be seen around the nose and muzzle from a degu rubbing itself continually against cage bars. Other stereotypical behaviors include constant gnawing of the cage bars, rubbing on the bars, continuous grooming, or sitting immobile for hours and stopping all regular activities such as play or grooming. Treatment involves more interaction with the owner, environmental enrichment, and addition of a companion degu. In group-housed degus, barbering, as a form of dominant behavior, or fight wounds and abscesses from bite wounds are also seen. Ectoparasites are infrequent in degus. Do not hold degus by the tail because they will spin like a top and deglove the tail skin. Degus that are familiar with their owners do not show this behavior, and tail shedding is uncommon. Dental Disease. Dental diseases are the most common problem seen in degus, being recorded in 60% of these pet animals. It is seen more frequently in older degus (75% in animals more than 2 years of age). Incisor malocclusion usually results secondary to coronal elongation of cheek teeth; traumatic injury is a rare cause. Clinical crowns of maxillary cheek teeth elongate toward the cheek and mandibular crowns elongate toward the tongue. Apical elongation of mandibular cheek teeth can be palpated on the ventral mandibular surface. The likely cause is the refusal of pet degus to eat an optimal amount of hay, instead consuming commercial feeds containing alfalfa, cornflakes, maize, grain mixture, nuts, raisins, and other sweets. Such diets have less abrasive properties, resulting in reduced chewing duration owing to high dietary energy content and lower coarse fiber
CHAPTER 27 Disease Problems of Small Rodents content. 113 The lack of tooth wear likely results in continuous eruption and abnormal coronal and apical elongation of the incisor and cheek teeth. In early dental disease, the mandibular cheek teeth elongate apically with palpable prominences on ventral mandibular surface. In severe dental disease, maxillary cheek teeth may extend into the nasal cavity, narrowing the nasal passages. Degus appear to be prone to elodontoma (odontoma) formation.46 This results from germinal tooth tissue damage from stereotypical wire-cage chewing. In one survey of 300 degus, 20 cases (7%) were seen.45 Elodontomas partially obstruct the nasal cavity, leading to respiratory problems; secondary bacterial rhinitis is a common consequence. Elodontomas are easily seen on skull radiographs. Endocrine and Ocular System. Degus develop spontaneous diabetes mellitus; the lesion is amyloidosis of the Langerhans islets. Cytomegalovirus-induced insulitis, alpha-cell crystals with a herpes-type viral presence, and foods such as guinea pig chow or fresh fruit (which increase blood sugar levels) are associated with the development of diabetes.97 Owners should give degus a commercial rodent diet supplemented with vegetables. Like prairie dogs, it is easy to overfeed degus and obesity is likely to occur. No treatments have been described for diabetic degus. Diabetic degus can develop cataracts within 4 weeks.10 Cataracts were the third most frequent lesion seen in pet degus.45 Check for diabetes in degus with cataracts, since a congenital cataract unrelated to diabetes has also been described.114
MEDICATION AND ANTIBIOTIC THERAPY IN PET RODENTS Because of the small size of pet rodents, even pediatric-strength medications must often be diluted for use in these species. Knowing the precise body weight of the animal, diluting medications, and administering medications with a tuberculin or insulin syringe will permit greater accuracy of dosing. Medication is often given by mixing it into feed or water. However, rats do not drink if they find the taste of their water objectionable. Ball-ended dosing needles are ideal for gavage, but always carefully calculate the volume of the dose and depth of penetration when using the dosing needle to prevent gastric rupture. Intravenous injections are difficult to administer, and the substitution of intravenous administration with intraperitoneal injection (for anesthetics) and intramuscular or subcutaneous injections is common. Exercise caution in administering antibiotic therapy to rodents. Streptomycin and procaine are toxic in mice; nitrofurantoin causes neuropathologic lesions in rats; and gerbils cannot tolerate dihydrostreptomycin and streptomycin. Hamsters are similar to guinea pigs in their susceptibility to the development of clostridial enterotoxicity when they are given penicillins, erythromycin, or lincomycin. Antibiotics that are apparently safe to use in rodents (especially guinea pigs and hamsters) include enrofloxacin, ciprofloxacin, trimethoprim-sulfa combinations, and chloramphenicol. Avoid ampicillin and amoxicillin; other sulfonamides, tetracycline, and piperacillin should be used sparingly in hamsters. Many compounding pharmacies now prepare medications in flavored syrups or treats that are palatable to rodents. (See also Chapter 26 for a discussion of antibiotic therapy in rodents.)
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CLIENT EDUCATION Rodents kept as pets, especially unusual species that may have been imported or animals that are subjected to stress from crowded conditions during transportation or sale, have the potential of transmitting zoonotic diseases (see Chapter 40). The risk of zoonotic disease should be discussed with clients, particularly if the pet is in a household with children or immunosuppressed persons. Occasionally a client will present a wild rodent that has been caught and that he or she wants to keep as a pet; this should always be discouraged because of the risk that these animals may carry bacteria, viruses, or parasites that might be infectious to persons with whom the animal is in contact. Most clients purchase books on pet rodents in pet stores or look at information on the Internet. They often rely on the recommendations of the pet store owner before asking for advice from a veterinarian. Unfortunately many of the available owner’s manuals are not familiar to veterinarians. Having some knowledge about pet rodents from these handbooks, clients often raise questions about what they have read, and clinicians may not appear well informed from the client’s perspective if they are unfamiliar with such references. The rodent owner then often return to the pet store owner for guidance; unless their animals are very sick, such owners may not return to the veterinarian for advice on husbandry and diseases. At this point, the prognosis for very ill pets is poor. Familiarizing oneself with the pet hobbyist literature breaks the cycle of mistrust and ignorance. Many hobby books on pet rodents are highly entertaining and informative about the husbandry and biology of the animals; some are not. In any case, the veterinarian should carefully review the medical information in these books. Purchasing and recommending some of these books to pet rodent owners is an effective method not only of educating clients but also of establishing good rapport.
Suggested Client Reading Barron’s Complete Pet Owner’s Manuals (Chinchillas, Degus, Dwarf Hamsters, Gerbils, Guinea Pigs, Hamsters, Mice, Rats, Fancy Rats, Rabbits, Dwarf Rabbits, and others): paperback, 64-104 pages, $9.00. Since 1999, all publications in the Complete Pet Owner’s Manuals are written by experienced veterinarians. Many of the small-rodent books are authored by Sharon Vanderlip, a clinical veterinarian. These books have good husbandry and basic diseases sections and are the best value for their price.
References 1. Abbott DP. Malignant schwannoma of the dorsal spinal nerve root in a laboratory rat. Lab Anim. 1982;16:265-266. 2. Albrich WC, Monnet DL, Harbarth S. Antibiotic selection pressure and resistance in Streptococcus pneumoniae and Streptococcus pyogenes. Emerg Infect Dis. 2004;10:514-517. 3. Baker DG. Natural pathogens of laboratory mice, rats, and rabbits and their effects on research. Clin Microbiol Rev. 1998;11:231-266. 4. Barthold SW, Bhatt PN, Johnson EA. Further evidence for papovavirus as the probable etiology of transmissible lymphoma of Syrian hamsters. Lab Anim Sci. 1987;37:283-288. 5. Bauck L, Orr JP, Lawrence KH. Hyperadrenocorticism in three teddy bear hamsters. Can Vet J. 1984;25:247-250.
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6. Beck W. Common endo- and ectoparasitic diseases in small mammals—clinical feature, diagnosis and treatment. A review of the literature and own experiences. [German] Haufige Endo- und Ektoparasitosen bei kleinen Heimsaugern - Klinik, Diagnostik und Therapie Literaturubersicht und eigene Erfahrungen. Tierarztl Prax Ausg K Klientiere Heimtiere. 2004;32:311-321. 7. Bingel SA. Pathologic findings in an aging Mongolian gerbil (Meriones unguiculatus) colony. Lab Anim Sci. 1995;45: 597-600. 8. Bivin WS, Olsen GH, Murray KA. Morphophysiology. In: Van Hoosier GL, McPherson CW, eds. Laboratory hamsters. Orlando, FL: Academic Press Inc; 1987:9-41. 9. Briese E. Normal body temperature of rats: the setpoint controversy. Neurosci Biobehav Rev. 1998;22:427-436. 10. Brown C, Donnelly TM. Cataracts and reduced fertility in degus (Octodon degus). Cataracts secondary to spontaneous diabetes mellitus. Lab Anim (NY). 2001;30:25-26. 11. Capello V. Prescrotal open technique for neutering a degu. Exot DVM. 2005;6:29-31. 12. Castro PA, Sohn CS, Roman L. Nonsurgical correction for vaginal/uterine prolapse in mice [P27]. Proceedings. 61st Annu Meet Am Assoc Lab Anim Sci. 2010;150. 13. Chole RA, Henry KR, McGinn MD. Cholesteatoma: spontaneous occurrence in the Mongolian gerbil. Meriones unguiculatis. Am J Otol. 1981;2:204-210. 14. Committee for the Update of the Guide for the Care and Use of Laboratory Animals. National Research Council. Guide for the care and use of laboratory animals. 8th ed. Washington, D.C. National Academies Press; 2010. 15. Cunnane SC, Bloom SR. Intussusception in the Syrian golden hamster. Br J Nutr. 1990;63:231-237. 16. Dammann P, Hilken G, Hueber B, et al. Infectious microorganisms in mice (Mus musculus) purchased from commercial pet shops in Germany. Lab Anim. 2011;0. 17. Deeb B. Respiratory disease in pet rats. Exot DVM. 2005;7:31-33. 18. Donnelly TM. Tail-slip in gerbils—Improper handling. Lab Anim (NY). 1997;26:15-16. 19. Donnelly TM. What’s your diagnosis? Blood-caked staining around the eyes in Sprague-Dawley rats. Lab Anim (NY). 1997;26:17-18. 20. Donnelly TM. What’s your diagnosis? Nasal dermatitis (‘‘facial eczema’’ or ‘‘sore nose’’). Lab Anim (NY). 1997;26:17-18. 21. Donnelly TM. Application of laboratory animal immunoassays to exotic pet practice. Exot DVM. 2006;8:19-26. 22. Donnelly TM, Rush EM, Lackner PA. Ringworm in small exotic pets. Sem Avian Exot Pet Med. 2000;9:82-93. 23. Donnelly TM, Stark DM. Susceptibility of laboratory rats, hamsters, and mice to wound infection with Staphylococcus aureus. Am J Vet Res. 1985;46:2634-2638. 24. Drouot S, Mignon B, Fratti M, et al. Pets as the main source of two zoonotic species of the Trichophyton mentagrophytes complex in Switzerland, Arthroderma vanbreuseghemii and Arthroderma benhamiae. Vet Dermatol. 2009;20:13-18. 25. Edwards MS. Nutrition and behavior of degus (Octodon degus). Vet Clin North Am Exot Anim Pract. 2009;12:237-253. 26. Eshar D, Mayer J, Keating JH. Dermatitis in a Siberian hamster (Phodopus sungorus). Bacterial pseudomycetoma. Lab Anim (NY). 2010;39:71-73. 27. Fehr M, Luerssen D, Kaup FJ. Hepatic cysts in golden hamsters. [German] Leberzysten beim Goldhamster (Mesocricetus auratus). Kleintierpraxis. 1987;32(283):287-288. 28. Fehr M, Schanen H, Grof D, et al. Anatomical basics and description of a method of castration in the degus. [German] Anatomische Grundlagen und Beschreibung einer Kastrationsmethode beim Degu (Octodon degus Molina). Kleintierpraxis. 1994;39:837-840.
29. Fitzner Toft M, Petersen MH, Dragsted N, et al. The impact of different blood sampling methods on laboratory rats under different types of anaesthesia. Lab Anim. 2006;40:261-274. 30. Fleming JM, Creevy KE, Promislow DE. Mortality in North American dogs from 1984 to 2004: An investigation into age-, size-, and breed-related causes of death. J Vet Intern Med. 2011;25:187-198. 31. Flynn BM, Brown PA, Eckstein JM, et al. Treatment of Syphacia obvelata in mice using ivermectin. Lab Anim Sci. 1989;39:461-463. 32. Flynn RJ. Notes on ringtail in rats. In: Conalty ML, ed. Husbandry of laboratory animals. London & New York: Academic Press; 1967:285-288. 33. Germann PG, Kohler M, Ernst H, et al. The relation of amyloidosis to social stress induced by crowding in the Syrian hamster (Mesocricetus auratus). Z Versuchstierkd. 1990;33:271-275. 34. Gleiser CA, Van Hoosier GL, Sheldon WG, et al. Amyloidosis and renal paramyloid in a closed hamster colony. Lab Anim Sci. 1971;21:197-202. 35. Gonenc B, Sarimehmetoglu HO, Ica A, et al. Efficacy of selamectin against mites (Myobia musculi, Mycoptes musculinus and Radfordia ensifera) and nematodes (Aspiculuris tetraptera and Syphacia obvelata) in mice. Lab Anim. 2006;40:210-213. 36. Gray JE. Chronic progressive nephrosis, rat. In: Jones TC, Mohr U, Hunt RD, eds. Urinary System. Berlin: Springer-Verlag; 1986:174-178. 37. Guzman-Silva MA, Costa-Neves M. Incipient spontaneous granulosa cell tumour in the gerbil, Meriones unguiculatus. Lab Anim. 2006;40:96-101. 38. Harkness JE. A practitioner’s guide to domestic rodents. Denver: American Animal Hospital Association; 1993. 39. Hart M, O’Connor E, Davis M. Multiple peracute deaths in a colony of Syrian hamsters (Mesocricetus auratus). Lab Anim (NY). 2010;39:99-102. 40. Harvey RG, Whitbread TJ, Ferrer L, et al. Epidermotropic cutaneous T-cell lymphoma (mycosis fungoides) in Syrian hamsters (Mesocricetus auratus). A report of six cases and the demonstration of T-cell specificity. Vet Dermatol. 1992;3: 13-19. 41. Heider K, Eustis SL. Tumors of the soft tissues. In: Turusov VS, Mohr U, eds. Tumours of the mouse. 2nd ed. Lyon: International Agency for Research on Cancer; 1994:611-631. 42. Hill PB, Lo A, Eden CA, et al. Survey of the prevalence, diagnosis and treatment of dermatological conditions in small animals in general practice. Vet Rec. 2006;158:533-539. 43. Hotchkiss CE. Effect of surgical removal of subcutaneous tumors on survival of rats. J Am Vet Med Assoc. 1995;206:1575-1579. 44. Jackson TA, Heath LA, Hulin MS, et al. Squamous cell carcinoma of the midventral abdominal pad in three gerbils. J Am Vet Med Assoc. 1996;209:789-791. 45. Jekl V, Hauptman K, Knotek Z. Diseases in pet degus: a retrospective study in 300 animals. J Small Anim Pract. 2011;52:107-112. 46. Jekl V, Hauptman K, Skoric M, et al. Elodontoma in a degu (Octodon degus). J Exot Pet Med. 2008;17:216-220. 47. Kalueff AV, Minasyan A, Keisala T, et al. Hair barbering in mice: implications for neurobehavioural research. Behav Processes. 2006;71:8-15. 48. Kastenmayer RJ, Fain MA, Perdue KA. A retrospective study of idiopathic ulcerative dermatitis in mice with a C57BL/6 background. J Am Assoc Lab Anim Sci. 2006;45:8-12. 49. Keenan KP, Hoe CM, Mixson L, et al. Diabesity: a polygenic model of dietary-induced obesity from ad libitum overfeeding of Sprague-Dawley rats and its modulation by moderate and marked dietary restriction. Toxicol Pathol. 2005;33:650-674. 50. Kerstjens HA, Postma DS, Ten Hacken N. COPD. Clin Evid (Online). 2008; pii: 1502.
CHAPTER 27 Disease Problems of Small Rodents 51. Kogan LR, Goldwaser G, Stewart SM, et al. Sources and frequency of use of pet health information and level of confidence in information accuracy, as reported by owners visiting small animal veterinary practices. J Am Vet Med Assoc. 2008;232:1536-1542. 52. Kondo H, Onuma M, Shibuya H, et al. Spontaneous tumors in domestic hamsters. Vet Pathol. 2008;45:674-680. 53. Kramer K, Kinter LB. Evaluation and applications of radiotelemetry in small laboratory animals. Physiol Genomics. 2003;13:197-205. 54. Krautwald-Junghanns ME, Pees M, Reese S, et al. Diagnostic imaging of exotic pets: birds, small mammals, reptiles. Hannover: Schlutersche; 2010. 55. Krogstad AP, Franklin CL, Besch-Williford CL. An epidemiological and diagnostic approach to murine skin lesions [meeting abstract]. Contemp Top Lab Anim Sci. 2001;40:82-83. 56. Kuntze A. Diseases of guinea pigs and golden hamsters important in practice. [German] Praxisrelevante Erkrankungen bei Meerschweinchen und Goldhamster. Monatsh Veterinarmed. 1992;47:143-147. 57. Kuo TH, Ho KL, Wiener J. The role of alkaline protease in the development of cardiac lesions in myopathic hamsters: effect of verapamil treatment. Biochem Med. 1984;32:207-215. 58. Laming PR, Cosby SL, O’Neill JK. Seizures in the Mongolian gerbil are related to a deficiency in cerebral glutamine synthetase. Comp Biochem Physiol C. 1989;94:399-404. 59. Lawson GHK, Gebhart CJ. Proliferative enteropathy. J Comp Pathol. 2000;122:77-100. 60. Lawson GW, Sato A, Fairbanks LA, et al. Vitamin E as a treatment for ulcerative dermatitis in C57BL/6 mice and strains with a C57BL/6 background. Contemp Top Lab Anim Sci. 2005;44:18-21. 61. Lindt VS. About the diseases of the Syrian golden hamster (Mesocricetus auratus). [German] Uber Krankheiten des Syrischen Goldhamsters (Mesocricetus auratus). Schweiz Arch Tierheilkd. 1958;100:86-97. 62. Losco PE. Dental dysplasia in rats and mice. Toxicol Pathol. 1995;23:677-688. 63. Lyerly DM, Bostwick EF, Binion SB, et al. Passive immunization of hamsters against disease caused by Clostridium difficile by use of bovine immunoglobulin G concentrate. Infect Immun. 1991;59:2215-2218. 64. Martinho F. Suspected case of hyperadrenocorticism in a golden hamster (Mesocricetus auratus). Vet Clin North Am Exot Anim Pract. 2006;9:717-721. 65. Matsuoka K, Suzuki J. Spontaneous tumors in the Mongolian gerbil (Meriones unguiculatus). [Japanese; abstract in English] Exp Anim. 1995;43:755-760. 66. McConnell J, Kirby R, Rudloff E. Administration of acepromazine maleate to 31 dogs with a history of seizures. J Vet Emerg Crit Care. 2007;17:262-267. 67. Mihaiu M, Rotaru O, Dan SD, et al. Experimental studies regarding obesity prevention in pets. Cluj Vet J. 2010;18:30-35. 68. Mihara K, Hirano T. Standing is a causative factor in osteonecrosis of the femoral head in growing rats. J Pediatr Orthop. 1998;18:665-669. 69. Motzel SL, Gibson SV. Tyzzer disease in hamsters and gerbils from a pet store supplier. J Am Vet Med Assoc. 1990;197:1176-1178. 70. Norris ML, Adams CE. Incidence of cystic ovaries and reproductive performance in the Mongolian gerbil, Meriones unguiculatus. Lab Anim. 1972;6:337-342. 71. Osborne CA, Albasan H, Lulich JP, et al. Quantitative analysis of 4468 uroliths retrieved from farm animals, exotic species, and wildlife submitted to the Minnesota Urolith Center: 1981 to 2007. Vet Clin North Am Small Anim Pract. 2009;39:65-78. 72. Ottenweller JE, Tapp WN, Burke JM, et al. Plasma cortisol and corticosterone concentrations in the golden hamster, (Mesocricetus auratus). Life Sci. 1985;37:1551-1558.
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73. Page MM, Robb EL, Salway KD, et al. Mitochondrial redox metabolism: Aging, longevity and dietary effects. Mech Ageing Dev. 2010;131:242-252. 74. Papes F, Logan DW, Stowers L. The vomeronasal organ mediates interspecies defensive behaviors through detection of protein pheromone homologs. Cell. 2010;141:692-703. 75. Pedersen KS, Holyoake P, Stege H, et al. Diagnostic performance of different fecal Lawsonia intracellularis-specific polymerase chain reaction assays as diagnostic tests for proliferative enteropathy in pigs: a review. J Vet Diagn Invest. 2010;22:487-494. 76. Percy DH, Greenwood JD, Blake B, et al. Diagnostic exercise: sloughing of limb extremities in immunocompromised suckling mice. Contemp Top Lab Anim Sci. 1994;33:66-67. 77. Pollicino P, Rossi L, Rambozzi L, et al. Oral administration of moxidectin for treatment of murine acariosis due to Radfordia affinis. Vet Parasitol. 2008;151:355-357. 78. Pullium JK, Brooks WJ, Langley AD, et al. A single dose of topical moxidectin as an effective treatment for murine acariasis due to Myocoptes musculinus. Contemp Top Lab Anim Sci. 2005;44:26-28. 79. Reeves WK, Cobb KD. Ectoparasites of house mice (Mus musculus) from pet stores in South Carolina, U.S.A. Comp Parasitol. 2005;72:193-195. 80. Refinetti R. Variability of diurnality in laboratory rodents. J Comp Physiol A Neuroethol Sens Neural Behav Physiol. 2006;192:701-714. 81. Rembert MS, Johnson AJ. What’s your diagnosis? Pigmented mass in an experimental gerbil. Spontaneous malignant melanoma. Lab Anim (NY). 2001;30:22-25. 82. Renshaw HW, Van Hoosier Jr GL, Amend NK. A survey of naturally occurring diseases of the Syrian hamster. Lab Anim. 1975;9:179-191. 83. Sanchez-Alavez M, Alboni S, Conti B. Sex- and age-specific differences in core body temperature of C57Bl/6 mice. Age (Dordr). 2011;33:89-99. 84. Sandhu GS, Solorio L, Broome AM, et al. Whole animal imaging. Wiley Interdiscip Rev Syst Biol Med. 2010;2:398-421. 85. Sarna JR, Dyck RH, Whishaw IQ. The Dalila effect: C57BL6 mice barber whiskers by plucking. Behav Brain Res. 2000;108:39-45. 86. Schimke DJ, Nixon CW, Connelly ME. Long-hair growth influenced by sex in the Syrian hamster. J Hered. 1974;65:57-58. 87. Schmidt RE, Eason RL, Hubbard GB. Pathology of aging Syrian hamsters. Boca Raton: CRC Press; 1983. 88. Schmidt RE, Reavill DR. Cardiovascular disease in hamsters: review and retrospective study. J Exot Pet Med. 2007;16:49-51. 89. Schneider JE, Wade GN. Effects of maternal diet, body weight and body composition on infanticide in Syrian hamsters. Physiol Behav. 1989;46:815-821. 90. Schneider JE, Wade GN. Effect of ambient temperature and body fat content on maternal litter reduction in Syrian hamsters. Physiol Behav. 1991;47:135-139. 91. Short BG, Goldstein RS. Nonneoplastic lesions in the kidney. In: Mohr U, Dungworth DL, Capen CC, eds. Pathobiology of the aging rat. Washington: International Life Sciences Institute Press; 1992:211-225. 92. Sichuk G, Bettigole RE, Der BK, et al. Influence of sex hormones on thrombosis of the left atrium in Syrian (golden) hamsters. Am J Physiol. 1965;208:465-470. 93. Silverman S, Tell L. Radiology of rodents, rabbits and ferrets: an atlas of normal anatomy and positioning. Philadelphia: WB Saunders; 2005. 94. Slattum MM, Stein S, Singleton WL, et al. Progressive necrosing dermatitis of the pinna in outbred mice: an institutional survey. Lab Anim Sci. 1998;48:95-98. 95. Solomon HF, Dixon DM, Pouch W. A survey of staphylococci isolated from the laboratory gerbil. Lab Anim Sci. 1990;40:316-318.
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96. Somvanshi R, Iyer PK, Biswas JC, et al. Polycystic liver disease in golden hamsters. J Comp Pathol. 1987;97:615-618. 97. Spear GS, Caple MV, Sutherland LR. The pancreas in the degu. Exp Mol Pathol. 1984;40:295-310. 98. Squartini F, Pingitore R. Tumors of the mammary gland. In: Turusov VS, Mohr U, eds. Tumours of the mouse. 2nd ed. Lyon: International Agency for Research on Cancer; 1994:47-100. 99. Strauli P, Mettler J. Tumours of the haematopoietic system. In: Turusov VS, ed. Tumours of the hamster. Lyon: International Agency for Research on Cancer; 1982:343-369. 100. Sundberg JP, Brown KS, McMahon WM. Chronic ulcerative dermatitis in black mice. In: Sundberg JP, ed. Handbook of mouse mutations with skin and hair abnormalities. Boca Raton: CRC Press; 1994:485-492. 101. Tankersley CG, Irizarry R, Flanders SE, et al. Unstable heart rate and temperature regulation predict mortality in AKR/J mice. Am J Physiol Regul Integr Comp Physiol. 2003;284:R742-R750. 102. Taylor DK, Rogers MM, Hankenson FC. Lanolin as a treatment option for ringtail in transgenic rats. J Am Assoc Lab Anim Sci. 2006;45:83-87. 103. Van Hoosier Jr GL, Trentin JJ. Naturally occurring tumors of the Syrian hamster. Prog Exp Tumor Res. 1979;23:1-12. 104. Vincent AL, Rodrick GE, Sodeman Jr WA. The pathology of the Mongolian gerbil (Meriones unguiculatus): a review. Lab Anim Sci. 1979;29:645-651. 105. Visticot M- E. A new companion animal: the degu, Octodon degus. [French] Un nouvel animal de compagnie: l’octodon, Octodon degus. Thesis for Doctorat Veterinarire. Alfort: École nationale vétérinaire d’Alfort; 2002.
106. Waggie KS, Ganaway JR, Wagner JE, et al. Experimentally induced Tyzzer’s disease in Mongolian gerbils (Meriones unguiculatus). Lab Anim Sci. 1984;34:53-57. 107. Walberg JA, Stark DM, Desch C, et al. Demodicidosis in laboratory rats (Rattus norvegicus). Lab Anim Sci. 1981;31:60-62. 108. Wardrop KJ, Van Hoosier GL. The hamster. In: Loeb WF, Quimby FW, eds. The clinical chemistry of laboratory animals. New York: Pergamon Press; 1989:31-39. 109. Wechsler SJ, Jones J. Diagnostic exercise. A case of atrial thrombosis and consumptive coagulopathy in a hamster. Lab Anim Sci. 1984;34:137-139. 110. Weisbroth SH, Kohn DF, Boot R. Bacterial, mycoplasmal and mycotic infections. In: Mark AS, Steven HW, Craig LF, eds. The laboratory rat. 2nd ed. Burlington: Academic Press; 2006:339-421. 111. White DJ, Waldron MM. Naturally-occurring Tyzzer’s disease in the gerbil. Vet Rec. 1969;85:111-114. 112. Witt CJ, Johnson LK. Diagnostic exercise: rear limb ataxia in a rat [radiculoneuropathy]. Lab Anim Sci. 1990;40:528-529. 113. Wolf P, Kamphues J. A critical assessment of commercial supplementary feedstuffs for rabbits, guinea pigs, chinchilla as companion pets. [German] Kritische Einschatzung kommerzieller Erganzungspraparate fur Kaninchen, Meerschweinchen und Chinchilla. Praktische Tierarzt. 2003;84:674-678. 114. Worgul BV, Rothstein H. Congenital cataracts associated with disorganized meridional rows in a new laboratory animal: the degu (Octodon degus). Biomedicine. 1975;23:1-4.
CHAPTER
28
Soft Tissue Surgery
R. Avery Bennett, DVM, MS, Diplomate ACVS
Presurgical Considerations Patient Support Instrumentation Magnification Focal Light Hemostatic Aids Patient Preparation Sutures, Needles, and Closure Orchidectomy Ovariectomy and Ovariohysterectomy Ovarian Disease Uterine Disease Ovariectomy Ovariohysterectomy Cesarean Section Mammary Gland Neoplasia Other Cutaneous Masses Miscellaneous Abdominal Procedures Cystotomy Abdominal Masses Pulmonary Lobectomy Abscesses Subcutaneous Abscesses Dental Abscesses in Hamsters Enucleation and Exenteration Cheek Pouch Eversion in Hamsters Prolapsed Bowel in Hamsters Tail Amputation Recovery Analgesia
Small rodents such as hamsters, gerbils, rats, and mice are popular pets. Despite their small size and relatively low purchase price, their owners are frequently as emotionally attached to them as they would be to a dog or a cat. This attachment often leads owners to request surgical and medical treatments that have costs much greater than the replacement value of the pet. Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
As members of the Myomorph group of rodents, hamsters, gerbils, rats, and mice share similar anatomy. Rats and mice are murid rodents, whereas gerbils and hamsters are cricetid rodents. All are monogastric, have a bicornuate uterus and uterine body, and have testicles that are large relative to their body size and usually descended into a large, well-developed scrotum, which, however, can easily be retracted into the abdomen through relatively large inguinal canals. The spermatic cords can be palpated cranial to the penis on each side as they exit the inguinal canal. Caudally, within the scrotum, the testicles are separated by the median raphe. The female reproductive tract is more like that of dogs and cats than rabbits and hystricomorph rodents. The mesovarium is short, making it difficult to exteriorize the ovaries for removal. The uterine body is relatively short, with a single cervical os. The oviducts encircle the ovaries, as they do in other rodents. This group of rodents is sexually mature at 40 to 60 days of age.23 Rats and mice have a long, hairless tail; gerbils have a long, haired tail; and hamsters have a short, lightly haired tail. Hamsters have large cheek pouches in which they store food.
PRESURGICAL CONSIDERATIONS PATIENT SUPPORT Most surgeries in small rodents are analogous to those performed in dogs and cats. The small size of these patients makes surgery more challenging for several reasons. As surgical patients, rodents are especially prone to complications such as hypovolemia from blood loss, hypothermia, and renal and respiratory compromise. Because they also serve as a food source for predator species, fear, pain, and stress have profound effects on their survival during and after surgery. It is believed that the release of large amounts of catecholamines, which can have effects on anesthesia as well, is responsible for such stress-related death. Efforts should be made to minimize pain, fear, and stress during the hospitalization period. The provision of preemptive analgesic and antianxiety medications is a vital component of a successful outcome. A preoperative database should include a complete blood count (CBC), plasma biochemical analysis, urinalysis, and whole-body radiographs, especially to evaluate the lungs, 373
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because they are especially prone to asymptomatic respiratory disease as well as to renal disease. Rats and mice frequently have subclinical mycoplasmal pneumonia. Although renal disease is common in older rodents, it can be difficult to obtain urine in a small rodent. Because they urinate frequently, however, one can put the rodent in a clean, dry, plastic or glass container, and a few drops of urine can then be collected from the container within a few minutes. This is all that is required to evaluate the urine by dipstick and determine the specific gravity. Proteinuria and dilute urine suggest underlying renal disease. At the very least, a preoperative database should include a packed cell volume, total protein, urine specific gravity, and a blood urea nitrogen (BUN) estimate by an azotemia test strip. These tests require only a few drops of blood and one drop of urine. Rodents are physiologically unable to vomit, so a prolonged fast is not necessary.35 They also have relatively low hepatic glycogen stores and are prone to developing hypoglycemia during a prolonged fast.16,35 A short fast of 1 hour will allow the animal to clear food material from its mouth for oral procedures or to minimize the risk of an endotracheal tube carrying food into the trachea if intubation is anticipated. Administration of fluids containing dextrose (SC, IV, or IO) will help combat loss of energy stores. A prolonged fast has not been shown to decrease the volume of abdominal contents in these hindgut cellulosefermenting species. Rodents have a large body surface area/volume ratio predisposing to the development of hypothermia during long anesthetic and surgical procedures. A temperature monitoring probe is ideal for continuous evaluation of core body temperature. The body temperature of a rat can drop 18°F (10°C) after 20 minutes of anesthesia.17 Hypothermia decreases the metabolism and excretion of many anesthetic drugs. A short operative time—accomplished by having all the necessary equipment ready and accessible—will help to minimize the development of hypothermia. Alcohol should be avoided for skin preparation because the evaporative cooling will potentiate the development of hypothermia. Warm, sterile saline is as effective as alcohol in patient preparation. Alcohol is also flammable and can ignite if electrosurgery or laser is used. A circulating warmwater blanket under the patient should be set at 104°F (40°C), because these patients have a higher resting body temperature than dogs and cats. Forced warm air blankets (Bair Hugger, Arizant, Eden Prairie, MN) are also useful in combating hypothermia. The Thermally Controlled Surgery Pad (RICA Surgical Products, Inc., Schiller Park, IL) is a safe plastic pad placed under the patient that does not overheat and turns on automatically when a patient is placed on it and turns off automatically when not in use. Also, the Hot Dog Warming System (Augustine Biomedical & Design, Eden Prairie, MN) is a safe and effective conductive-fabric warming unit. Other sources of supplemental heat, including hot water bottles or examination gloves filled with warm water and radiant heat lamps, can cause thermal burns and must be used with caution. Placing a towel between the heat source and the patient does not ensure safety. Closely monitor all heat supplement devices to avoid accidentally burning the patient. The patient should be draped as quickly as possible because the drape will help hold in heat. Seemingly small amounts of hemorrhage can be disastrous in such small patients. Although it may not be practical, ideally it is best to have blood from a conspecific available for transfusion when significant blood loss is anticipated. Strict attention to hemostasis is vital. The average cotton-tipped applicator holds
Fig. 28-1 The Thermally Controlled Surgery Pad (RICA Surgical Products, Inc., Schiller Park, IL) provides safe, supplemental heat and serves as a patient restraint and positioning board. The pad, with the patient secured to it, can be tilted and moved during the surgery as needed.
approximately 0.1 mL of blood when completely soaked. Loss of 10% to 15% of the total blood volume (approximately 1% of body weight) is usually safe. Loss of more than five cottontipped applicators full of blood (0.5 mL, or 10 drops) in a 50-g mouse is equivalent to more than 20% of the blood volume and is potentially dangerous.16 Approximately 1.2, 1.4, and 4.0 mL of blood is equivalent to more than 20% of the blood volume (a potentially dangerous amount to lose) in the average-sized gerbil, hamster, and rat, respectively.16 Intravascular fluid volume may also be supported with the aid of crystalloid fluids such as lactated Ringer’s solution with or without dextrose. Intraosseous cannulas such as spinal needles are commonly inserted in a normograde fashion into the proximal femur or proximal tibia of pet rodents in a manner analogous to normograde insertion of an intramedullary pin for fracture management. This provides access to the vascular system for fluid support and emergency drug therapy if needed. Fluids may also be administered subcutaneously or intraperitoneally with slower absorption. These routes are not acceptable for treating animals that are seriously ill, severely dehydrated, or in shock. The standard recommendation for IV fluid support during anesthesia and surgery is 10 mL/ kg per hour; however, a single dose of 10 mL/kg SC of 4% dextrose has been recommended for short procedures in rodents.35 The thoracic cavity of many small rodents is small relative to their body size. Placing the patient in dorsal recumbency can compromise respiration because of the pressure of the abdominal viscera on the diaphragm. Tilting the patient so that the viscera are displaced caudally may be beneficial for the respiratory system of the anesthetized patient.35 Securing the patient to a restraint board such as the Thermally Controlled Surgery Pad (Fig. 28-1) rather than the surgical table allows the patient to be tipped in any direction and to be easily repositioned intraoperatively.
INSTRUMENTATION In general, ophthalmic instruments are not suited to surgery in small mammals (Fig. 28-2). The short length of the instruments makes them difficult to control and manipulate the tissues within the body cavity. Microsurgical instruments are constructed so
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A Fig. 28-3 Jeweler’s forceps (bottom) have a very small point. Ring-tip forceps (top) have a fine tip that is broader and can be used to hold tissue as well as suture for knot tying.
B Fig. 28-2 A, Ophthalmic instruments (right) are short, with flat finger grips, compared with microsurgical instruments, which are of a standard length and have rounded grips (left). B, They should be held like a pen. Microsurgical instruments are long enough to extend beyond the fingers into a body cavity and balance in the hand, requiring very little muscle action and thus reducing fatigue and tremor.
that only the tips are miniaturized, which is different from ophthalmic instruments, where the whole instrument is miniaturized.4 The handles should be of normal length (5.5 to 7 in.) to help provide stability to the tips, thus diminishing the effects of tremors. The tips of the instruments should extend beyond the fingers far enough to use within the patient’s body cavity, while ophthalmic instruments may be used for surface work. The handles should be round to facilitate the required rolling action between the thumb and first two fingers. Round handles are most important for the needle holders, where a curved needle must be rolled through the tissue. Many prefer needle holders without a clasp or box lock because the motion that occurs when the lock is set and released may cause the needle to tear tissues. Hold a microsurgical instrument as if you were holding a pen. An across-the-palm grip is inappropriate and limits the function of these precision instruments (see Fig. 28-2). A microsurgical pack should consist minimally of a microsurgical needle holder (Micro Surgery Needle Holder, straight, T/C without lock, 8 in.; Micro Surgery Needle Holder, curved, without lock, 7.125 in., pencil-style grip), microsurgical scissors (Tew-Barraquer Scissors, curved, 7 in.), and microsurgical thumb forceps (Micro Surgical Forceps, curved, 7 in., pencilstyle grip) (all from RICA Surgical Products, Inc., Schiller Park, IL). Many microsurgeons use jeweler’s forceps as microsurgical forceps (Long Jeweler’s Forceps No. 3, RICA Surgical Products, Inc.). These are significantly less expensive and serve their purpose adequately, but they are not ideal. They have sharp tips, are short, and do not have round handles. I prefer curved ringtipped microsurgical forceps (Ring Tip Forceps, Cat. No. 843025,
Fig. 28-4 The Lone Star Retractor (Veterinary Specialty Products, Inc., Mission, KS) is autoclavable. The tissue hooks on Silastic bands are used to gently retract tissues during surgery. The hooks are placed into the tissue and the bands, under gentle tension, are wedged into the notches on the plastic frame.
RICA Surgical Products, Inc.) and Counterweight Round Handle Ring Tip Tissue Forceps (Cat. No. 2200-260, Sontec Instruments, Centennial, CO) because they are of a standard length, counterbalanced, and delicate, but the tips have rings that distribute the pressure over a wider surface, making them less traumatic for tissue handling while one can still hold the fine strand of suture for knot tying (Fig. 28-3). Eyelid retractors do not work well in small rodents because the tension cannot be adjusted to fit the individual patient. Heiss and Alm Self Retaining Retractors (RICA Surgical Products, Inc.), analogous to Bennett and Doolan retractors (Sontec Instruments), respectively, work well as abdominal and tissue retractors in small rodents. The Lone Star Retractor (Veterinary Specialty Products, Inc., Mission, KS) consists of Silastic bands with tissue hooks on the ends and a plastic frame (Fig. 28-4). All components are fully autoclavable and quite durable. The stay hooks are placed in the tissue and the Silastic bands inserted into the notches of the frame to maintain tissue retraction. Use the least amount of tension necessary to maintain exposure. Hemostatic clips (Hemoclips and Samuels Hemoclip Applier, 6.25 in., RICA Surgical Products, Inc.) are also very useful and available in five sizes, with small and
SECTION IV Small Rodents
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B
A
Fig. 28-5 A, Surgitel (General Scientific Corp., Ann Arbor, MI) loupes have a focal range rather than a set focal distance and are available in various styles (glasses or mounted on a head band) with options for a focal light source. B, They are ergonomically designed for the surgeon to maintain an appropriate posture while providing magnification. The surgeon diverts the view down to use the magnification and over the lenses when magnification is not needed. The head and neck remain upright, reducing neck fatigue. medium being most applicable for pet rodent surgery. A microsurgical pack can be put together for $1,500 to $2,000. After experience has been gained, other microsurgical instruments such as micro mosquito hemostats may be added to the pack. Small gauze pads (2 × 2 in.) can be cut from the standard 4 × 4-in. sponges. Sterile cotton-tipped applicators should also be available. These are useful for absorption of fluids as well as gentle tissue dissection and manipulation. Absorbable gelatin sponge (Gelfoam, Pfizer, New York, NY; Vetspon, Novartis Animal Health US, Inc., Greensboro, NC) and other absorbable hemostatic agents are valuable for controlling hemorrhage. Surgicel (Ethicon, Inc., Sommerville, NJ) is oxidized regenerated cellulose resembling cloth. It is a hemostatic aid that adheres nicely to moist tissues but is not capable of absorbing much fluid. Topical thrombin (Thrombin-JMI, Gentrac, Inc. Middleton, WI) is available in various-sized vials. Thrombin is a liquid that stimulates fibrin clot formation using the patient’s fibrinogen. Either gelatin sponge or oxidized regenerated cellulose can be soaked in topical thrombin before being placed on the source of hemorrhage to aid in hemostasis. Ophthalmic bulb syringes work well for tissue irrigation in small rodents, but a syringe with a needle attached is also applicable. Fine Barron suction tips have a release hole providing two degrees of suction. For rodent patients, the tip is used mainly with the hole open, providing suction that will not traumatize tissues. If there is a larger volume of fluid, occlude the hole with the tip of your index finger to increase the force of the suction.
MAGNIFICATION Some form of magnification is recommended for surgery in rodent patients. The dexterity and manipulation required of fingers and hands are far greater than can be achieved with unaided vision. Small amounts of hemorrhage appear more significant under magnification and individual vessels are much more easily identified for coagulation, minimizing the degree of hemorrhage associated with a procedure. Binocular loupes are available in various styles, with the hobby loupe being the least expensive and simplest. Hobby loupes have several disadvantages, including a set focal distance, no focal light source, and the inability to look around the lenses when magnification is not required.
A modification of the hobby loupe is marketed with interchangeable lenses and an LED focal light source (MDS, Inc., Brandon, FL). These allow the surgeon to change the degree of magnification as appropriate for the size of the patient and to illuminate body cavities. This type of loupe still has a set focal length and the surgeon is committed to looking through the lenses. The higher the power, the shorter the focal distance, and the closer the surgeon must be to the working area for objects to be in focus. These factors all contribute to dizziness in the wearer. These loupes also require the surgeon to bend his or her neck to be able to see through the lenses. This results in fatigue and neck pain. SurgiTel (General Scientific Corp., Ann Arbor, MI) eliminates most of the problems associated with the hobby loupes (Fig. 28-5). It is ergonomically engineered to be used with a proper neck posture, the lenses do not cover the entire visual field so the surgeon is not committed to looking through the lenses, it has a focal LED light source, and the lenses also have a focal range, rather than a set focal distance, which generally allows the entire patient to be in focus during surgery.
FOCAL LIGHT Overhead lights are not adequate for surgery in small rodents. A focal light provides illumination of a smaller area, allowing better visualization of the tissues. In working within a body cavity, a focal light is essential to illuminate the relevant structures. Various types of lights with or without magnifying loupes are available through various companies. LED lights are commonly used because they provide high-quality cool light with high intensity and durability. An inexpensive headband-mounted LED light, the LED Headlight, is available (MDS, Inc., Brandon, FL). SurgiTel loupes are available with different light options, including fiberoptic lights that adapt to endoscope light sources and halogen or LED lights that use a battery or wall outlet. These mount over the lenses and are adjusted to illuminate the surgeon’s field of view. General Scientific Corp. also offers SurgiCam, a digital video camera with 3× magnification that mounts over the lenses. This system allows the procedure to be viewed on a monitor. It makes still photos or video recordings that can be edited—a useful client education tool.
CHAPTER 28 Soft Tissue Surgery
Fig. 28-6 The Surgitron Dual Frequency 120 radiosurgical unit (Ellman International, Inc., Hewlett, NY) is small and has both monopolar and bipolar capabilities. It operates with radiofrequency electromagnetic waves, with the ground plate acting as an antenna.
HEMOSTATIC AIDS Electrocautery refers to the use of electricity to heat metal until it is red hot to heat-coagulate tissues. Electrosurgery uses highfrequency alternating current to generate energy. There are two electrodes (an active electrode and an indifferent electrode) with concentration of current density at the tip of the smaller (active) electrode. Burns can result if the ground plate contacts only a small area. The Surgitron (Ellman International, Inc., Hewlett, NY) uses radiofrequency current that is received, with the indifferent electrode acting as an antenna. The area of contact between the patient and the indifferent electrode is irrelevant, which is especially advantageous in working with small mammal patients. The bipolar forceps is most useful for hemostasis within body cavities because the current passes between the tips of the electrodes, coagulating only the tissue held in the tips of the forceps. Ellman International offers a switch that allows the surgeon to change from monopolar (used for cutting the skin) to bipolar (used for coagulating vessels within a body cavity) and back without having to unplug electrodes. The Surgitron Dual Frequency 120 (Ellman International, Inc., Hewlett, NY) radiowave energy is emitted at 4.0 MHz, providing control of absorption depth in tissue and resulting in minimal cellular damage and heat conduction (Fig. 28-6). This unit features a unique isolated circuitry system that maintains a constant frequency of 4.0 MHz. With this constant frequency, there is only 15 μm of lateral heat damage compared with 90 μm from the older Surgitron units and more than 100 μm with a carbon dioxide laser. Standard electrosurgical units generally produce more than 750 μm of lateral heat damage. The finger switch has a button for cutting, another for coagulating, and a third for a blend of cutting and coagulating. A technician is not required to switch from bipolar to monopolar, as is required with the other electrosurgical units. The carbon dioxide (CO2) laser (Luxar and Aesculight, LuxarCare, Woodinville, WA) is popular in surgery involving small exotic animals. It produces a beam of light energy at a wavelength of 10,600 nm. This wavelength of energy is highly absorbed by water molecules, making it ideal for cutting with a focal beam or vaporizing tissue with a diffused beam. The CO2 laser seals vessels less than 0.6 mm in diameter, so many of the incisions are bloodless, resulting in less postoperative swelling because the lymphatic vessels are also sealed. Incisions made with the CO2 laser are reportedly less painful than those made
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with a blade or electrosurgical or radiosurgical units because the laser beam seals nerve endings as well. When used appropriately, the amount of lateral heat damage is said to be less with a laser than with some other modalities;4 however, when used incorrectly, lasers can cause significant lateral thermal damage. Lasers can cause injury to the operator or assistant as well as to the other tissues of the patient. Proper safety training for all personnel, therefore, is important in using surgical lasers. In the past several years, Harmonic Systems (Ethicon Endosurgery, Inc. Cincinnati, OH) has gained popularity for providing hemostasis during minimally invasive and open procedures. This is an ultrasonically activated tissue cutting device whereby electrical energy is converted to mechanical energy. Ultrasonic vibrations at the tip generate heat that results in tissue coagulation. After the tissue is coagulated and hemostasis is achieved, the device is reactivated to cut the tissue within the tips. It is approved by the U.S. Food and Drug Administration (FDA) to seal vessels up to 5 mm in diameter with less lateral heat than electrosurgery or laser. No electrical current passes through the patient. Electricity does not pass through the tissues, so muscles do not spasm when cut. The tissue is grasped between the Teflon anvil (cool) and the cutting blade (hot); it is then sealed at the low setting and cut at the high setting. The hand pieces are intended for single use but can be gas-sterilized and reused 10 to 15 times, making them affordable for veterinary practice. LigaSure (Covidien, Mansfield, MA) is a tissue sealing system that has gained popularity in veterinary surgery. The newest model combines the tissue sealing technology with monopolar and bipolar electrosurgery (Force Triad). While the specific technology is proprietary, the LigaSure is a type of bipolar cautery. The generator senses the electrical impedance of the tissues within the tips and delivers the amount of current needed to melt the elastin and collagen; it then allows the elastin and collagen to re-form, creating a permanent seal in a single application. It is FDA-approved to seal vessels up to 7 mm in diameter and generates less than 1 mm of lateral heat. A translucent “seal zone” is created and the tissue is transected through this zone.
PATIENT PREPARATION Standard aseptic technique is essential with pet rodent patients. Rodents are susceptible to infections and their cages often require them to be in close proximity to their urine and feces. Be careful to avoid damaging the skin during preparation for surgery. The area clipped should be minimized to help control heat loss. A No. 50 clipper blade (Oster Professional Products, McMinnville, TN) is ideal for fine, thin hair in patients with thin skin, such as small rodents. The teeth are closer together than those of the No. 40 blade, making it less likely that the patient’s skin will get caught between the teeth and be cut accidentally. Clear plastic drapes allow respiration to be monitored during the procedure. A plastic drape with a 3.5- × 5-in. adhesive center and an overall size of 40 × 54 in. is commercially available (Veterinary Transparent Surgical Drape, Veterinary Specialty Products, Inc.). This allows the surgeon to maintain a sterile field over the entire table and still monitor the patient. A smaller drape (24 × 24 in.) is also available for minor procedures but is not large enough to create a sterile field on a surgical table. These drapes may also hold heat better than cloth or paper drapes. Place the four quarter drapes around the patient rather than the proposed incision, then place the plastic drape over the patient to cover the entire table and create a sterile field.
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SECTION IV Small Rodents
SUTURES, NEEDLES, AND CLOSURE In general, the sutures and needles in the average veterinary practice are too large for use in small rodents. It is best to use the smallest size suture and needle to accomplish the desired task. For these small patients, sizes 5-0 and smaller with short, fine, swaged-on needles are applicable. Cutting needles are mainly used for skin. Atraumatic taper-point needles are recommended for soft tissues. Monofilament rapidly absorbable suture is most appropriate for rodents (polyglecaprone, glycomer, polyglactin 910, and polyglycolic acid). Suture that is present for many months, such as polydioxanone and polyglyconate, is not usually needed in rodents. The subcutis in small rodents is relatively thick and holds subcuticular sutures well. Rodents are fastidious groomers and tend to remove skin sutures. They also seem prone to self-mutilation of surgical wounds. Gentle tissue handling, strict attention to aseptic technique, and closing with no external sutures will help minimize the risks. Pinching the skin with forceps during suture placement causes bruising and irritation, predisposing to selfmutilation. In placing sutures, use forceps to provide counterpressure and guide the tissues without pinching. Chromic catgut should be avoided because of its reactive nature, which might stimulate self-mutilation and can delay healing. I prefer 5-0 to 7-0 rapidly absorbable suture in a subcuticular or intradermal closure to avoid the use of skin sutures. If necessary, tissue glue may also be used to facilitate skin apposition. Avoid using large amounts of tissue glue, because the patient will notice it and work to get it off, potentially resulting in self-trauma. The placement of intradermal sutures is time-consuming. If time is of the essence, small incisions can be apposed with only tissue adhesive or skin staples. Rodents tend to chew at skin sutures and can chew out wire sutures as well as nylon ones. Steel skin staples appear to be more resistant to being chewed out.
reported in gerbils.41 Orchidectomy is indicated to prevent or treat these conditions. The procedure for orchidectomy in rats, mice, hamsters, and gerbils is similar. The testicles are located caudoventrally in the inguinal area and are readily retracted into the abdomen (Fig. 28-7). They can easily be pushed back into the scrotum with gentle rolling pressure on the caudal abdomen in a caudal direction just cranial to the pubis. The testicles are relatively large and contained in a well-defined scrotal sac caudal to the penis. The epididymal (testicular) fat is responsible for preventing intestinal herniation.23 This fat extends into the abdominal cavity near the kidneys bilaterally (Fig. 28-8). The fat, along with the testicular artery and vein, passes through the inguinal canal into the scrotum. It can be challenging to remove the testicle without removing or damaging the epididymal fat at the proximal end of the testicle and attached to the head of the epididymis. In performing a closed orchidectomy, removal of the entire testicle but leaving as much of the epididymal fat as possible avoids the risk of herniation or evisceration; however, the risk of this has not been clearly documented. Alternatively, the vaginal tunic can be ligated closed proximally after an open orchidectomy is performed with minimal trauma to the epididymal fat. Preservation of this fat appears to be important for preventing visceral herniation following open or closed orchidectomy in small rodents. In small rodents there is little hair on the scrotum itself; however, adjacent to the penis on each side and cranially into the inguinal region, clip the fur with a No. 50 blade and perform a standard surgical preparation of the scrotum. Isolate the penis and scrotum and make a single transverse 0.5- to 1.0-cm incision at the distal (caudal) tip of the scrotum. Alternatively, two incisions oriented dorsal to ventral can be made, one in each side of the scrotum at the tail of the epididymis. Place the incision as far dorsally as possible to minimize postoperative incisional contamination by the cage substrate (Fig. 28-9).
ORCHIDECTOMY The primary indication for orchidectomy in rodents is to prevent breeding, as it is easier to castrate males than to spay females. Urethral plugs have been described in male rats, gerbils, golden hamsters, mice, and guinea pigs as a normal finding, although they have caused urethral obstruction in rats and mice.27 They are present in all healthy adult male rats, but their size decreases by 99% following orchidectomy. Urethral plugs are composed of proteins I-V from the seminal vesicles mixed with vesiculase, from the coagulating glands; these congeal to form the plug.6 Because orchidectomy virtually eliminates the risk of a urethral plug causing obstruction in rodents, some recommend routine orchidectomy at a young age for rats. Whether it has an effect to suppress aggressive behavior is a subject of debate and there is no scientific evidence that orchidectomy will ameliorate aggression. In most species, orchidectomy is recommended prior to puberty because often, once sexual behaviors manifest, they become learned behaviors not necessarily affected by the loss of hormones. Testicular tumors, such as Leydig cell tumors, occur in rodents, especially rats, and orchidectomy is indicated for treatment. The incidence of testicular tumors varies with strain of rat and increases with age. Although testicular tumors are usually bilateral and multilobulated, unilateral tumors are generally very large, with the unaffected testis becoming atrophied. Leydig cell tumors are benign but can get quite large.41 Prostatic and testicular tumors are also
a
b
c
Fig. 28-7 The external genitalia of a male rat showing the preputial orifice (a), the scrotum (b), and the tail of the epididymis (c).
CHAPTER 28 Soft Tissue Surgery Do not incise the vaginal tunic in performing a closed orchidectomy. The tunic is thin, fragile, and easily torn. Exteriorize the testicles only far enough to allow the entire testicle to be removed without removing the epididymal fat blocking the inguinal canal. If the tunic is inadvertently incised or torn,
a b
A
a b
B Fig. 28-8 A, The epididymal fat is attached to the head of the epididymis and extends cranially through the inguinal canal and into the abdomen to the caudal pole of the kidney, obstructing the inguinal canal and preventing inguinal hernia formation. The body wall, inguinal canal, scrotum, and vaginal tunic have been incised to show the normal location of the epididymal fat. B, The fat is being retracted cranially within the abdomen, pulling the testicle into the abdomen. In this image about half of the testicle is within the abdomen.
A
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convert the procedure to an open technique. Gently pry the scrotum off the vaginal tunic with gauze, breaking the ligament of the tail of the epididymis and isolating the spermatic cord. Do not clamp the cord because it will likely tear the tunic and testicular vessels. Rather, place a single encircling ligature of a 4-0 rapidly absorbable suture just distal to the epididymal fat (between the fat and the head of the epididymis) and transect the cord distal to the ligature. For the open technique, extend the incision through the tunic on each testicle to allow the testicles to be exteriorized through the tunic. The ligament of the tail of the epididymis attaches the tail of the epididymus to the tunic and the scrotal skin. Manually break the ligament of the tail of the epididymis, freeing the testicle from the tunic and scrotum. Replace the everted scrotum to its normal position. Apply caudal retraction to the testicle to expose the spermatic cord (testicular artery and vein and ductus deferens). Ligate or clip the cord as close to the testicle as possible with a single ligature (between the head of the epididymis and the fat), leaving the epididymal fat intact, and transect it distal to the ligature (Fig. 28-10). Replace the fat into the inguinal canal. The tunic may be left open or ligated closed as cranial as possible, thereby blocking the inguinal canal. The skin incision may be left open to heal by second intention, or it may be closed with a tissue adhesive. The scrotal skin in this area is too thin to allow placement of intradermal sutures. Another technique involves making bilateral skin incisions at the cranial extent of the spermatic cord near the inguinal canal, cranial to the penis.9 The skin is thicker here, and this approach allows access to the external inguinal ring, which can then be sutured closed as part of the procedure. Make a 1-cm incision in the skin approximately 0.5 to 1.0 cm lateral to the penis on each side (Fig. 28-11). Do not make the incisions too close to the prepuce because the support for the penis within can be damaged.33 Identify the spermatic cord within the vaginal tunic. An open or modified closed orchidectomy can be performed through this inguinal approach. Gently grasp the spermatic cord within the tunic and carefully free the tunic from its attachments circumferentially to pass a ligature. For a modified closed technique,
B Fig. 28-9 A, An incision for castration of this mouse was made in the skin at the dorsal aspect of the tail of the epididymis. B, For a closed castration, the testicle is exteriorized with the vaginal tunic; an encircling ligature is placed just proximal to the head of the epididymis to preserve the epididymal fat, preventing an inguinal hernia. (Surgical drape removed for photographic purposes.)
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A
B Fig. 28-10 A, During an open castration in a mouse, the vaginal tunic is incised, exposing the testicle and the spermatic cord. B, An encircling ligature is placed between the head of the epididymis and the epididymal fat; the spermatic cord is then transected proximal to the ligature. The stump is replaced in the inguinal canal to prevent an inguinal hernia. (Surgical drape removed for photographic purposes.)
Fig. 28-11 An inguinal incision can be used to castrate rodent pets. The dotted line in this image shows the location of the incision in a rat.
the tunic is opened to facilitate removal of the testicles, but it is ligated closed proximal to the opening in the tunic. Preplace a ligature (4-0 to 6-0) around the cord and incise the tunic distal to the ligature. Identify the epididymal fat and push it proximal to the ligature, then tie the ligature incorporating the spermatic cord within the tunic at the junction of the epididymal fat with the head of the epididymis. The head of the epididymis can easily be detached from the testis and left within the tunic, helping to block the inguinal canal. The goal is to block the inguinal canal with the epididymal fat. Exteriorize the testicle through the incision in the tunic, detach the scrotal skin from the ligament of the tail of the epididymis, replace the scrotum with the attached tunic into its normal position, and then ligate the exposed spermatic cord distal to the first ligature and transect
between the two, allowing the testicle to be removed. A closed technique can be performed using this inguinal approach; however, the scrotal skin is adhered to the vaginal tunic, making it difficult to free the tunic-enclosed testicle through the more cranial incision. The open technique is performed in a similar manner but a ligature is not placed around the vaginal tunic containing the spermatic cord. Incise the tunic and exteriorize that testicle.9 Do not pull it far from the body, as this will pull the epididymal fat out of the inguinal canal. Detach the ligament of the tail of the epididymis from the scrotum and replace the scrotal skin with the attached vaginal tunic to its normal position. Ligate the open spermatic cord, making sure that the fat is replaced into the inguinal canal, and close the skin. Regardless of the technique used, close the incisions with an intradermal suture or cyanoacrylate tissue adhesive. The theoretical advantages of this cranial inguinal approach result from its being close to the external inguinal ring, allowing it to be occluded either by ligating the cord within the tunic or placing a suture across the external inguinal ring. The inguinal skin is also thicker, making it is easier to place intradermal sutures there. A disadvantage of this approach is that the incisions are in contact with the substrate during normal ambulation, which could predispose to infection at the surgery site. After surgery, the substrate must be kept clean and changed at least twice daily for 1 week. Generally sexual activity should cease in 1 to 2 weeks after orchidectomy; however, in rodents that have had sexual experience before orchidectomy, mounting and intromission usually persist for several weeks.16 Viable sperm remain in the ductus deferens. Therefore, to prevent unwanted pregnancy, orchidectomized males should not be put in contact with females for 6 to 8 weeks after surgery. Complications associated with orchidectomy include hematoma formation, self-trauma, excess activity, and infection.35 Visceral herniation can occur if the epididymal fat has been disrupted. The inguinal canal in rodents is large enough for the testicles to move freely into the abdominal cavity even in adults. Some surgeons prefer an abdominal approach for orchidectomy in rodents.17,21 The technique is analogous to that described in Chapter 25.
CHAPTER 28 Soft Tissue Surgery
OVARIECTOMY AND OVARIOHYSTERECTOMY Indications for ovariectomy in rodents include control of reproduction, preventing and treating cystic ovaries, reducing the risk of mammary and pituitary tumors, and suppressing anxiety.12 In most species, ovariectomy is effective at preventing uterine disease; in one review, virtually all complications associated with performing ovariohysterectomy were due to removal of the uterus.42 Therefore, unless there is uterine pathology at the time of surgery, ovariectomy is preferred over ovariohysterectomy. Ovariectomy of young female rodents has been shown to significantly decrease the potential for the development of mammary neoplasia.20 Ovariectomized female rats also had a significantly lower incidence of pituitary tumors and a higher rate of survival to day 630. Ovariectomy, however, also predisposes rats to osteopenia.20 Rats ovariectomized at a young age (3 months), at adulthood (6 months), and at an advanced age (18 months) had significantly suppressed anxiety behavior compared with intact rats, indicating that ovariectomy performed at any age may have a behavioral benefit.12 Depressive behaviors were shown to worsen with age regardless of the presence or absence of ovaries.
OVARIAN DISEASE Cystic ovaries have been reported in all the small rodents, but most often in hamsters and gerbils. Cystic ovaries are reported to occur commonly in gerbils over 2 years of age and affect one or both ovaries.28 Granulosa cell, thecal cell, and lutein cell tumors are also common in gerbils.5,15 In one report, there was a 12.5% incidence of ovarian tumors in gerbils.3 Of female reproductive tract tumors in gerbils, 29 of 37 were ovarian in origin and the other 8 were uterine.41 The incidence of thecomas in hamsters is approximately 2%.41 These cystic masses may become very large without causing significant signs of discomfort; however, they can become large enough to compress abdominal organs and compromise function. They can also make it difficult for the hamster to take in adequate amounts of nutrition. Animals typically present with abdominal distention on one or both sides but may present for dyspnea from the cystic mass compressing the chest and lungs. Ultrasound is a useful tool in determining the origin (ovary or other structure) and character (fluid-filled or solid tissue) of the mass. It is used to obtain a sample for cytology with fineneedle aspiration. If cystic ovaries are documented, drain the fluid contents to improve ventilation and make ovariectomy easier to perform. While malignant cystic ovarian diseases occur, there are no reports of tumor seeding subsequent to needle aspiration of the contents. Once removed, the fluid rapidly reaccumulates, making ovariectomy or ovariohysterectomy the treatment of choice. No reports for treatment with human chorionic gonadotropin (HCG) in hamsters and gerbils, as is done in guinea pigs, were found. Because of the high incidence of ovarian disease in these companion rodents, routine juvenile ovariectomy should be considered. The procedure is easier to perform with less morbidity in young, healthy animals with normal ovaries.
UTERINE DISEASE Diseases of the uterus include pyometra, mucometra, metritis and endometritis, and tumors. Tumors of the uterus have been reported15,41 and tend to be benign in rats and malignant in
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hamsters and gerbils. Benign endometrial stromal tumors occur in up to 66% of some strains of rats more than 21 months old; these are more common in virgin rats than in breeding females. Uterine adenocarcinomas are common in Syrian and Chinese hamsters, with vaginal hemorrhage being the most common presenting complaint.15,30 These tumors have been documented to metastasize and to be spread by implantation of malignant cells during surgery.41 Endometrial polyps, leiomyomas, leiomyosarcomas, cervical carcinomas, and squamous papillomas of the vagina have been reported in hamsters.15 Pyometra is rare in small rodents but can occur in any species. Female hamsters have an odorous, mucoid vaginal discharge at estrus, which should not be mistaken for pyometra.
OVARIECTOMY Ovariectomy is indicated for the treatment of ovarian disease and the prevention of ovarian and uterine disease. The ovaries are located at the caudal pole of the kidneys. The oviduct wraps around the ovary, encircling it cranially. The ovaries are deep to the viscera, which—through a ventral midline celiotomy—must be retracted to gain exposure. In some patients, the ovaries are difficult to exteriorize through a small ventral midline incision because of the short suspensory ligament. A single artery and vein runs medial to each ovary and the ipsilateral uterine horn. Ovariectomy can be performed from either a ventral midline or a dorsal approach. The advantages of the dorsal approach are that the gastrointestinal tract does not have to be manipulated; the incisions are dorsal, so not subjected to the weight of the viscera, making evisceration unlikely; and there is less morbidity, since the incisions are smaller. For a ventral midline approach, place the animal in dorsal recumbency and prepare it for aseptic surgery.9 Make a 2- to 3-cm ventral midline celiotomy incision midway between the umbilical scar and the pubis. In rodents, the incision must be relatively large in order to provide access to the ovarian vessels. Because gerbils of both sexes have a sebaceous gland located midventrally on the abdomen near the umbilicus, make the skin incision paramedially to avoid the gland and undermine the skin to expose the linea alba. Identify the uterine horns dorsal to the apex of the bladder. Grasp one horn and trace it cranially to locate the ovary. Carefully retract the cecum to the opposite side. Gentle tissue handling and minimal manipulation of the gastrointestinal tract are important because rodents, especially hamsters, are prone to the development of adhesions.35 Identify the vessels within the mesovarium supplying the ovary and create an opening in the mesovarium to allow placement of hemostatic clips or a 4-0 to 6-0 absorbable synthetic suture ligature. Place a second clip or ligature at the proper ligament of the ovary (between the ovary and uterus), thus isolating the ovary from its blood supply and connection to the uterus. Transect the suspensory ligament, mesovarium, and vessels distal to the ligature on the suspensory ligament and proximal to the ligature on the proper ligament of the ovary, allowing the ovary to be removed. Repeat the procedure on the contralateral side. Make sure there is no hemorrhage before closing the abdominal wall with 5-0 or 6-0 monofilament synthetic absorbable material in a simple continuous pattern. Close the skin with an intradermal suture or skin staples. The substrate should be changed at least twice daily to minimize incisional contamination. In laboratory rodents, ovariectomy is performed routinely using a dorsal approach.23,35 A single dorsal skin incision can
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be made longitudinally or transversely. The skin incision can be moved from one side to the other, allowing removal of both ovaries through one incision. Alternatively, separate incisions can be made to approach each ovary. Approximately 1 cm ventral to the dorsal spinous processes of the third lumbar vertebra, immediately caudal to the last rib, bluntly dissect through the body wall with a mosquito hemostat. Pressure on the abdomen will cause the ovary to extrude through the incision, allowing its removal. If the ovary does not exteriorize, insert a forceps or hemostat and retrieve it located just caudal to the kidney on that side. In some laboratory animal facilities, the ovaries are removed by blunt dissection, often with no ligation, because hemorrhage in young rodents is generally minimal. In a clinical setting, place a ligature or hemoclip on the ovarian vessels to control hemorrhage. Transect the pedicle distal to the ligature and remove the ovary. The oviduct, which encircles the ovary, is also removed. If it is incompletely removed, the lining produces fluid and a cyst can develop.23 Close the opening in the muscle with 4-0 to 6-0 synthetic monofilament absorbable suture. Because of the dorsal placement of the incision and the blunt dissection used to enter the abdominal cavity, a suture is recommended but probably not essential. If a single skin incision was made, move the incision to the other side and repeat the procedure. Close the skin with a subcuticular suture or tissue adhesive. With increasing age and amount of perirenal fat, it is more difficult to locate the ovaries using this approach; however, with practice the procedure is easily accomplished even in older female rodents.
OVARIOHYSTERECTOMY Ovariohysterectomy is indicated for the treatment of rodents with uterine disease and with or without ovarian disease. In treating uterine disease, ovariectomy is generally indicated, along with hysterectomy, because nearly all uterine diseases occur as a result of ovarian hormones.42 A ventral midline celiotomy is performed as described above. Identify the vessels within the mesovarium supplying the ovary and create an opening in the mesovarium to allow placement of hemostatic clips or an absorbable synthetic suture ligature. Monofilament absorbable synthetic suture of size 4-0 or 6-0 is suitable. Transect the suspensory ligament, mesovarium, and vessels distal to the ligature on the suspensory ligament and bluntly dissect the broad ligament on each side to the level of the uterine body. The uterine body of small rodents is short, and if the ovaries are removed, it is not usually critical where the uterus is ligated. Some recommend that the uterus be ligated cranial to the cervix to prevent urine from spilling into the abdomen when the uterus is transected. This appears to be of little concern clinically. Care must be taken to avoid damaging the urinary bladder, which lies immediately dorsal to the uterus. Ligate the uterine body at a convenient location and transect it distal to the ligature. Inspect the wound to make sure that there is no hemorrhage and close it routinely. Ovariohysterectomy can also be performed from a dorsolateral approach similar to that described above for ovariectomy.24 The dorsolateral abdomen is opened on only one side. Exteriorize the ovary and double ligate or clip the ovarian vessels, then transect between the ligatures. Retract the ovary and associated uterine horn out the opening in the dorsal body wall. Continue until you identify the uterine body and the contralateral uterine horn. Trace the contralateral uterine horn to the contralateral
ovary, double ligate the ovarian vessels, and transect between ligatures. Exteriorize both ovaries and uterine horns, allowing access to the uterine body. Ligate the uterus and transect it distal to the ligature. It is technically more difficult to remove the uterus from this approach, especially in older, obese rats or if there is uterine pathology.
CESAREAN SECTION Cesarean section has been reported in a gerbil.31 In rodent species, the patient is tilted head up to allow the viscera and gravid uterus to fall away from the thoracic cavity, making ventilation easier. Avoid the enlarged mammary glands and associated blood vessels in making the skin incision. The uterus is more vascular when gravid. Make the incision in the uterus carefully in a relatively avascular area. Close the uterus in a simple continuous pattern with a 5-0 to 6-0 monofilament synthetic absorbable suture on a small atraumatic needle. Oxytocin can be administered after removal of the fetuses to aid with uterine involution. Ovariohysterectomy is an alternative to cesarean section for managing dystocia in small rodents, especially if the fetuses are not viable and future reproduction is not desired.
MAMMARY GLAND NEOPLASIA Mammary gland neoplasia is the most common spontaneous neoplasm of mice and rats11,41 but appears to be rare in gerbils and hamsters. Most mammary tumors in rats and hamsters are benign, whereas in mice and gerbils they are usually malignant.11,41 In mice, they are rapidly metastatic, invasive, and much more difficult to remove.10 Mouse mammary tumor virus predisposes to the development of mammary adenocarcinoma and is passed transplacentally and through milk. In rats, mammary gland tumors are usually benign mammary fibroadenomas (50%-90%, depending on the strain)26; they are uncommon in rats younger than 1 year of age.20 Fewer than 10% of mammary tumors are malignant.15 Mammary tumors are usually single, large, firm, not attached to deeper structures, and well tolerated by the animal. They do not metastasize to distant locations and clinical problems are generally associated with their large size, sometimes being larger than the animal (Fig. 28-12). Rat mammary gland tumors are known to be sensitive to hormone stimuli,41 and Sprague-Dawley rats ovariectomized at 90 days of age had a significantly lower incidence of mammary tumors (4%) than intact rats (47%).20 These ovariectomized rats also had a significantly lower incidence of pituitary chromophobe adenomas (4% compared with 66%). Estrogen does not seem to be the major factor in tumor development because exogenous estrogen did not increase the incidence of tumor development.20 It is theorized that high levels of prolactin from pituitary tumors actually cause the development of mammary tumors. However, estrogen may contribute to the development of pituitary tumors, which in turn release large quantities of prolactin, resulting in mammary tumor development. Ovariectomy also improved survival to 630 days of age (89% compared with 59% of intact rats). Mammary tumor removal has been reported to prolong survival to 630 days14; however, a study by Hotchkiss20 comparing data from other studies with those regarding rats in which tumors were not removed failed to confirm this. Clinically it appears that ovariectomy in conjunction with mammary tumor removal helps to prevent tumor recurrence and new tumor development.
CHAPTER 28 Soft Tissue Surgery
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A
Fig. 28-13 Mammary tissue is extensive in rats and tumors occur
B
C
Fig. 28-12 A, Rat mammary fibroadenomas can grow quite large. This tumor weighed more that the rat did after tumor removal. B, Because speed is important in removing large mammary tumors, a single-layer closure with skin staples is usually adequate. C, Mammary tumors occur in 16% of older male rats.
Prophylactic ovariectomy at 90 days of age is recommended for rats. In treating rats with mammary tumors, remove the mammary tumor with minimal margins and, if the animal is stable, perform ovariectomy as well. If the animal’s status dictates a shorter anesthesia and surgery time, remove the mammary tumor, and then, at a later date, remove the ovaries. Estrogen antagonists are potentially hepatotoxic in rats and have not been shown to be effective in treating rat mammary tumors.15 It has been reported that 16% of aged male rats develop mammary fibroadenomas.20 If the theory is correct and estrogen contributes to the development of pituitary tumors that secrete prolactin resulting in mammary tumor formation, castration of male rats should not affect the incidence of tumors; however, no scientific studies have been conducted to determine any effect castration might have on the development of mammary masses in males. Mammary tumors in mice generally carry a poor prognosis because they are more commonly malignant and associated with mouse mammary tumor virus.10,11,15 The incidence in older female mice ranges from 30% to 70%, with tumors being invasive and vascular. They are difficult to remove surgically, and surgery has not been shown to prolong survival, likely because they are virus-induced. Similarly, ovariectomy does not have a protective effect. Fine-needle aspirate cytology is recommended
in any location. 1-4, Mammary glands; 1, cervical gland; 2, thoracic gland; 3, abdominal gland; 4, inguinal gland; 5, mammae papillae; 6, superficial cranial epigastric vein (subcutaneous abdominal vein); 7, preputial gland; 8, prepuce; 9, opening of vagina; 10, anus. (From Popesko P, Rajtová V, Horák J. Atlas of the anatomy of small laboratory animals. Vol. I. Rabbit and guinea pig; Vol. 2: Rat, mouse and golden hamster. Bratislava: Príroda Publishing, 1990 [Czechoslovak edition]).
to determine whether the mass is benign prior to recommending treatment. Restriction of energy content in the diet regardless of the nutrient restricted resulted in a significant reduction in the incidence of mammary tumors in mice. Mammary tumors have been reported in hamsters, with Russian hamsters being more commonly affected. In one report,15 it was the most common type of tumor diagnosed at a veterinary teaching hospital in a 35-year period (n = 6). Mammary adenocarcinomas have been reported in two gerbils.41 Mammary gland tissue is extensive in rats and mice, extending from the cervical to the inguinal region ventrally and as high as the shoulders and flanks laterally (Fig. 28-13). Mammary gland tumors can be found anywhere in these areas and grow rapidly to large sizes. Mammary tissue is confined to the ventral thorax and abdomen in hamsters and gerbils. Treatment of mammary tumors in rodents consists of excising the tumor and associated mammary gland, in conjunction with ovariectomy in some cases. Prior to surgery, it is important to determine whether the mass is benign or malignant. Fineneedle aspirate cytology is inexpensive and easy to perform, can potentially differentiate benign from malignant mammary tumors, can be helpful in planning the best surgical treatment, and can provide vital prognostic information for the owner. If the mass is ulcerated or known to be malignant, the skin overlying the tumor should be removed; however, the skin over benign tumors can be preserved, making closure easier. Remove the mass, saving as much skin over the mass as possible. After
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the mass has been removed, trim excess skin but preserve enough to allow tension-free closure. Generally there are several blood vessels in the subcutaneous tissue supplying the tumor that need to be ligated. Bluntly dissect around the mass, ligating or clipping vessels as they are encountered. With large tumors, ligate arteries before veins to allow some blood to drain from the tumor into the patient’s circulation. Speed and hemostasis are vital for a successful outcome. Electrosurgical or laser dissection and tissue-sealing devices that coagulate or seal vessels are useful for mammary tumor removal. Close the surgical wound in one or two layers. If possible, close the subcutaneous tissue separately from the skin. Drains are difficult to maintain in rodents, so consider tacking the subcutaneous tissue to deeper structures to minimize dead space. Be aware that tacking can cause irritation and stimulate the patient to chew at the incision postoperatively. If time is critical, close only the skin using staples, which are rapidly applied.
OTHER CUTANEOUS MASSES A variety of cutaneous neoplasms have been reported in rodents.15,41 These include fibromas, fibrosarcomas, lipomas, undifferentiated sarcomas, squamous cell carcinomas, and hemangiosarcomas. Fibromas tend to be small, firm, and well circumscribed, whereas fibrosarcomas are usually large, locally invasive, and can metastasize. Hamsters and gerbils have scentmarking glands (Fig. 28-14). It is important to know the normal location and appearance of these glands so as not to confuse them with a pathologic process. Hamsters have a bald spot near each hip, which marks the location of the scent gland, and Djungarian hamsters have a midventral sebaceous gland. Gerbils have a similar single gland on the ventral abdomen. Cutaneous neoplasms are uncommon in mice and rats.15 In mice, squamous cell carcinoma is most common. Dermal fibromas and fibrosarcomas are also reported, and there are rare reports of hemangiomas and myxolipomas. Subcutaneous tumors (fibromas, fibrosarcomas, lipomas, and undifferentiated sarcomas) are more common than cutaneous tumors and usually occur on the face, tail, and feet. Rats have a specialized sebaceous gland around the external ear canal called Zymbal’s gland. Malignancy of this gland pre sents as a large, ulcerated swelling or mass within or ventral to the ear canal. Squamous cell carcinoma is the most common
histologic diagnosis. These tumors are locally aggressive but late to metastasize to the lymph nodes and lungs. Surgical removal can be curative if adequate surgical margins can be obtained. This can be difficult with large masses and usually requires total ear canal ablation and reconstructive surgery. Rats of both sexes also have bilateral preputial glands (also called clitoral glands in females) (Fig. 28-13). Although inflammation of the glands is most commonly reported, malignancy can occur. These are modified sebaceous glands and one or both can be affected. Carcinomas of these glands are invasive and can metastasize. Early detection and removal with adequate surgical margins can affect a cure. Because of the aggressive nature of this tumor type, it is often diagnosed late in the course of the disease; therefore it can be difficult to obtain clean surgical margins. It is important to protect the urethral papilla during resection. Use a 24- to 26-gauge intravenous catheter as a urethral catheter to assure the urethra is not damaged during the resection. Skin and subcutaneous tumors are uncommon in hamsters. Melanoma is the most commonly reported tumor and often involves the pigmented scent glands. These are larger in males and the incidence of melanoma of the gland is also higher in male hamsters.17,30 Mast cell tumors were diagnosed in 8 of 86 skin biopsies and usually occur around the head and neck.15,32 Various other cutaneous and subcutaneous tumors have been reported in hamsters.15 The skin and subcutis are reported to be the second most common locations for tumors in gerbils, with adenomas and adenocarcinomas of the scent gland being the most common.10,15 A review of the literature in one report documented that 8 of 34 tumors reported in gerbils were associated with the scent gland, including three squamous cell carcinomas, a sebaceous adenoma, three sebaceous adenocarcinomas, and a papilloma.41 Wide excision of malignant tumors is recommended because they tend to be locally invasive.22 Of the 34 cutaneous tumors reported in gerbils, there were 12 squamous cell carcinomas, 8 melanomas, 5 sebaceous adenomas, 3 fibrosarcomas, and 2 mammary adenocarcinomas. Melanomas occur most commonly on the ear, foot, or base of the tail.10,15,36 Auricular cholesteatomas occur in gerbils and gerbils are the only species other than humans reported to develop this type of tumor.5,18
MISCELLANEOUS ABDOMINAL PROCEDURES Exploratory laparotomy is indicated in pet rodents for a variety of reasons. Most procedures performed in other species can be performed in small rodents with the aid of magnification, a focal light, microsurgical instruments, and appropriate hemostasis. Some rodents, especially hamsters, are prone to developing adhesions, which can cause clinical problems postoperatively.35 Gentle tissue handling, minimizing manipulation of viscera, and avoiding the use of reactive suture material, such as chromic catgut, are important in preventing adhesion formation. Verapamil (200 μg/kg PO q8h for 3 days), a calcium channel blocker, has been shown to reduce the formation of adhesions in hamsters after laparotomy.13
CYSTOTOMY Fig. 28-14 The normal scent gland of this hamster should not be confused with a cutaneous lesion.
Cystotomy is generally performed for removal of cystic calculi, which are relatively uncommon but do occur in small rodents.2,29 Clinical signs include hematuria, stranguria,
CHAPTER 28 Soft Tissue Surgery dysuria, incontinence, and anorexia. All the factors predisposing individual rodents to the development of cystic calculi have not been determined. It is difficult to cure this condition in rodents and recurrence is common.17 Calculi are usually calcium-based and may respond to dietary management; however, reduction of dietary calcium has not been shown to prevent their formation or recurrence. Rodent urine pH is alkaline. If the urine is acidic, depending on the type of calculi, urine alkalizing agents, such as potassium citrate, might help prevent recurrence. Obtain cultures of the bladder wall and the calculus during the procedure to direct postoperative antibiotic therapy. Cystotomy is performed as for other species except that the suture material and needle must be small and delicate tissue handling is required. Make a midline skin and body wall incision just cranial to the pubis and exteriorize the bladder. Isolate the bladder with moist 2- × 2-in. gauze sponges to minimize urine spilling into the abdomen. Make an incision on the ventral surface of the bladder, thus avoiding the trigone. Remove the calculi and collect samples for culture by excising a thin strip of bladder mucosa from the edge of the cystotomy incision. Irrigate the bladder and antegrade flush the urethra with an intravenous catheter before closing the cystotomy. The bladder wall of rodents is relatively thin. Close with 6-0 to 8-0 monofilament synthetic absorbable suture material on a small atraumatic swaged-on needle in a simple continuous pattern. In general, a continuous pattern creates a better seal than an interrupted pattern, minimizing the risk of urine leakage. Irrigate the abdominal cavity with warm saline before closing the celiotomy. If the patient is not urinating postoperatively, catheterize the urethra with an appropriate-sized intravenous catheter (24-26 gauge) and, if possible, leave it in place for 2 to 3 days to allow tissue swelling to subside.35 Nonsteroidal anti-inflammatory drugs (NSAIDs) can be used to reduce inflammation but should be avoided in patients with renal compromise.
ABDOMINAL MASSES Abdominal masses can arise from any organ. In general, hamsters are resistant to developing spontaneous neoplasms, but they are used in cancer research because it is easy to induce tumor formation in these animals. Hamsters are prone to polycystic disease. The cysts can become quite large and are most often found in the liver and kidneys.8 Hamsters are also prone to lymphosarcoma. In addition to the peripheral lymph nodes, masses may be found in the liver, kidney, spleen, and bowel.41 For a solitary mass or to relieve an intestinal obstruction due to lymphosarcoma, excision is indicated; however, for diffuse disease, chemotherapy is recommended. Gerbils have a high incidence of spontaneously occurring neoplasia, especially if they are older than 2 years. Tumors of the reproductive tract, liver, pancreas, and spleen have all been described.41 Renal tumors have been reported in rats. Nephroblastomas are seen most often in young rats; older rats with renal neoplasia usually have renal tubular adenomas or adenocarcinomas. These tumors are often bilateral.11,15,41 Hepatocellular carcinomas are common in mice and have been linked to Helicobacter species, causing chronic active hepatitis progressing to hepatocellular carcinoma.43 The incidence of hepatocellular carcinoma is also high in inbred strains of
385
mice with spontaneous hepatic lipidosis suggesting that fatty liver may predispose to the development of hepatocellular carcinomas.39 Adrenal tumors occur commonly in older rodents but are usually found incidentally at necropsy. There are at least 15 reports of malignant and benign adrenal tumors in gerbils.38,41 Adrenal tumors also occur commonly in older rats and are typically adrenocortical in origin. They may be malignant or benign, and pheochromocytomas have also been reported. The incidence varies with the strain of rat, and metastasis has been reported.41 In hamsters, adrenal adenomas are commonly reported benign tumors, occurring more often in males.41 Clinical signs include alopecia, skin hyperpigmentation, and changes in behavior. In a study that evaluated 4,575 Syrian hamsters, the incidence of adrenal masses in hamsters less than 1 year of age was only 1.7%, but it was 35.5% at 2 years and 40% in hamsters over 2 years of age.40 Adrenocortical adenocarcinoma has been reported in conjunction with pituitary chromophobe adenomas in two hamsters.1 The location of the adrenal glands is similar across mammalian species, with the left being easier to remove than the right because of the intimate association of the caudal vena cava and the right adrenal gland. Reports of successful surgical removal of adrenal tumors in rodents are lacking.
PULMONARY LOBECTOMY Many sources consider pulmonary tumors to be the most common tumor of mice, with an incidence of 28% in one study.34 Primary lung tumors appear to be uncommon in other rodent species. Aside from pulmonary masses, the other main indication for pulmonary lobectomy is for the removal of lung abscesses. A technique that does not require endotracheal intubation and is applicable to other rodents when intubation is not feasible has been described for pulmonary lobectomy in laboratory rats.37 The rat is maintained under anesthesia with a face mask that fits tightly around the muzzle, allowing the patient to be ventilated. During thoracotomy, ventilation is supported at 20 breaths per minute and the degree of lung expansion is directly visualized to evaluate for sufficient expansion and not overinflation. Make an intercostal approach with an incision over the lateral thoracic wall at the appropriate (generally at the fourth) intercostal space. Divide the muscles, paying attention to hemostasis, and then enter the thoracic cavity using a hemostat to penetrate the intercostal space. Spread the hemostat to open the intercostal space, avoiding vessels. Place a Bennett or Heiss retractor to maintain exposure while the lung is exteriorized. For lobectomy, place hemostatic clips or ligatures on the hilum to occlude the pulmonary artery and vein as well as the primary bronchus and transect distal to the clip. Place two sutures of 4-0 or 5-0 monofilament slowly absorbable suture around the ribs adjacent to the thoracotomy to oppose the ribs. Tighten the suture to leave a gap between ribs (intercostal space); the ribs should not overlap. Before tying the sutures, place a catheter into the chest to allow air and fluid to be evacuated after surgery. Close the muscle layers routinely along with the subcutis and skin. The chest tube can be maintained if needed based on the amount of fluid and air that accumulates or removed postoperatively after the chest cavity has been evacuated. This procedure takes less than 10 minutes to perform and the chest is generally open for less than
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3 minutes.37 Of the 54 rats on which this procedure was performed, 51 survived. The fatalities occurred early in the study and were related to hemorrhage.
ABSCESSES SUBCUTANEOUS ABSCESSES Abscesses in small rodents, particularly hamsters, are usually a result of bite wounds from cage mates or other trauma. Some affected animals are asymptomatic except for having an abnormal swelling. Others are systemically ill and are given supportive care as necessary. Radiographs are indicated when abscesses are on the face or feet to evaluate for the presence of bone or tooth involvement. Abscesses are easily diagnosed by finding pus on cytologic examination of a fine-needle aspirate. Because rodent pus is thick and the abscess wall is often thick, excision of the abscess without rupturing the capsule is more likely to result in a cure than lancing, draining, and irrigating the wound and leaving it open to heal by second intention. Antibiotic-impregnated polymethylmethacrylate beads are useful if there is tissue contamination or if the abscess cannot be excised because of its location. If it is not feasible to excise the entire abscess, remove as much of the abscess as possible, including the fibrous capsule. Submit a section of the wall and some pus for culture and sensitivity testing as well as histopathologic evaluation. Fill the resulting defect loosely with beads containing an antibiotic to which you suspect the organisms may be susceptible. For example, if no preoperative culture results are available, place some beads containing cefazolin and some beads containing amikacin for broad spectrum coverage. Close the subcutis and skin over the beads routinely. Administer a systemic broad-spectrum antibiotic based on culture and sensitivity results or empirically pending results. While systemic absorption from the beads should be minimal, some of the antibiotic does reach the circulation. There are few studies looking at systemic absorption from the beads in different species. Until more is known, it is safer not to use the same antibiotic systemically as the one that is in the beads because of the potential risk of having an additive effect when using antibiotics that have a narrow margin of safety.
DENTAL ABSCESSES IN HAMSTERS Hamsters are prone to periodontal disease and dental caries. Their incisors grow continuously but their cheek teeth do not. Therefore, once a cheek tooth is removed, no new tooth will grow to replace it. As in most rodents, the roots of the incisor teeth are long and deeply seated in the skull. A facial abscess located ventral or rostral to the eye may be caused by a tooth root abscess. These facial abscesses must be irrigated and drained; however, they will recur if the underlying cause, the abscessed tooth root, is not addressed. By the time a facial abscess has formed, the tooth root is usually so loose that extraction is not difficult. Remove the tooth by elevating the periodontal membrane while alternately and steadily retracting the exposed tooth. Close the defect in the gingiva with a single absorbable suture if possible or allow the defect to heal by secondary intention. Generally, once the devitalized tooth is removed the infection rapidly resolves. Clean the skin wound several times daily until it has granulated closed.
ENUCLEATION AND EXENTERATION Rodents have a large orbital venous sinus that surrounds the muscles and Harderian gland caudal to the globe. This is the site used for blood collection with a capillary tube in a laboratory setting. If this is damaged during enucleation, significant hemorrhage occurs. Indications for enucleation are those that cause end-stage ocular disease. Murid rodents do not have a third eyelid. Proptosis is somewhat common in small rodents, especially hamsters. If left untreated even a few hours, it may not be possible to save the eye. Two techniques are described for enucleation of rodents, the transconjunctival and the transpalpebral approaches.4,35 Both techniques involve excision of the globe, all of the conjunctiva, and the third eyelid in species that have one. The difference is based on whether the eyelid margin and palpebral conjunctiva are removed at the beginning of the procedure along with the globe and bulbar conjunctiva (transpalpebral) or if they are removed at the end of the procedure, after the globe and its conjunctiva have been removed (transconjunctival).4 For a transconjunctival enucleation, perform a lateral canthotomy first. Make an incision about 1 to 2 mm from the limbus circumferentially in the bulbar conjunctiva.19 Grasp the edge of the 2 mm of conjunctiva attached to the limbus securely while the globe is excised. Bluntly dissect deep to the conjunctiva through Tenon’s fascia to the sclera. Identify and transect the extraocular muscles as close as possible to their insertion on the globe to minimize muscle hemorrhage. Use care in applying traction to the globe, since this pulls on the optic chiasm and can damage the ocular nerve to the contralateral eye. Once all the muscles are transected, the globe will rotate freely within the orbit. Gently lift the globe and ligate, clip, or clamp the optic nerve along with the associated blood vessels. Incise distal to the clip, allowing the globe to be removed, and inspect the optic nerve for hemorrhage. If a clamp was used, a ligature can be more easily placed once the globe is removed. If the sinus has been damaged and hemorrhage is more serious, pack a piece of cellulose sponge into the defect and apply gentle pressure for 5 minutes. After the globe is removed, the conjunctiva must be removed. If these structures are not removed, cysts can form.19 Using Metzenbaum scissors, excise the upper and lower eyelid margins about 2 mm from the edges to join at the medial canthus. The medial angular vein is superficial and medial to the orbital rim and it is best to avoid this, using meticulous dissection. Inspect for any remaining conjunctiva and remove the Harderian gland. Be prepared for hemorrhage. One author suggests suturing the eyelids partially closed prior to removing the gland of the third eyelid and Harderian gland so that the rest of the lid margins can be closed quickly to provide pressure for hemostasis.19 Alternatively, excise the tissue quickly and control the hemorrhage with cellulose sponge and pressure. Once hemostasis is achieved, suture the eyelids closed. It is acceptable to place skin sutures in the eyelids since rodents are not able to chew them out from this location. Because rodent eyes are prominent, a transconjunctival approach is most commonly used. The transpalpebral approach is used if there is superficial ocular disease, such as bacterial conjunctivitis, to prevent the periocular structures from being contaminated during the procedure. Suture the eyelids together with fine nylon suture. Make an incision about 2 mm from the eyelid margins to the palpebral conjunctiva but not through it. Dissect down to the sclera rostral to the extraocular muscles without penetrating
CHAPTER 28 Soft Tissue Surgery the conjunctiva. Transect the extraocular muscles at their insertion and continue the dissection as described above. With the transpalpebral approach, all of the conjunctiva and glands are removed with the globe and any surface infection is contained within the sutured eyelids. Hemorrhage is controlled and lids closed as described for the transconjunctival approach. Exenteration involves removing all of the structures within the bony orbit including the globe, the ocular muscles, the glands, and the nictitans if present. Exenteration is indicated if there is periocular disease such as a retrobulbar abscess or neoplasia. Perform a lateral canthotomy, remove the eyelid margins, and excise all of the tissue within the orbit, staying as close as possible to the bone to minimize the risk of damaging the sinus. Control hemorrhage with hemostatic clips and cellulose sponge as needed. Close the eyelids as previously described. Regardless of the technique used, a polymethylmethacrylate bead with (if infection is present) or without an antibiotic can be placed as a prosthetic globe to help control infection and prevent the sunken appearance. Cellulose sponge is absorbed by fibrous tissue ingrowth which will also help prevent the concavity. Suture the eyelids closed over the prosthesis.
CHEEK POUCH EVERSION IN HAMSTERS Hamsters have well-developed cheek pouches bilaterally, are lined by a thin epithelial membrane. These pouches are used to store extra food, and they extend very far caudally (Fig. 28-15). Hamsters use their forelimbs to massage the food out of the pouch, then to be ingested. Cheek pouch impaction, eversion or prolapse, abscessation, and neoplasia have been reported.8 Cheek pouch impaction occurs when excessive amounts of inappropriate types of food are fed. For example, very large or very small seeds in the diet can impact the pouch; bedding materials such as cotton or paper stored in the pouches can also cause impaction. These materials desiccate and adhere to the epithelium, so that the hamster is unable to remove them. If materials stay in the pouch, the pouch may be predisposed to abscess formation. If the hamster works hard enough to get the material out and the epithelium is adhered, the pouch will prolapse or evert. One or both pouches can evert. If the prolapse is acute, sedate the hamster and remove the material adhered to the mucosa. If the mucosa is relatively healthy, replace the pouch to its normal position. Insert a 1-mL syringe case into the pouch and place a single full-thickness percutaneous mattress suture into the cheek pouch with 4-0 or 5-0 monofilament nonabsorbable suture material on a small needle (Fig. 28-16). If a cotton-tipped applicator is used, be careful not to suture the cotton tip into the pouch. A stent is not necessary because the suture is not tightened enough to cause skin necrosis. The hamster should be able to eat right away. Remove the suture in 14 days. If the mucosa is severely damaged or infected, it should be excised (Fig. 28-15).8 Place a hemostat at the base of the prolapsed pouch and transect the tissue distal to the clamp. Remove the clamp and place a single layer of fine monofilament, rapidly absorbed suture material in a simple continuous pattern. Postoperatively, it is best to withhold the normal diet because the animal will attempt to pack it into the pouch, resulting in incisional dehiscence. Syringe-feed a formula diet (e.g., Critical Care for Herbivores, Oxbow Animal Health, Murdock, NE) for 3 to 5 days and remove any bedding materials that the patient might attempt to pack into the pouch.
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Cheek pouch neoplasia often goes undetected until the mass prolapses with the pouch. Amputation as described above; and histopathologic evaluation of the tissue is recommended. If the cheek pouch is not everted, the skin of the cheek will be adherent and must be dissected from the cheek pouch. Make an incision in the epithelium of the pouch but not through the skin and undermine to free the pouch from the skin (see Fig. 28-15). Resect the mass and associated cheek pouch and suture the mucosal edges together with a fine suture.
PROLAPSED BOWEL IN HAMSTERS Hamsters are prone to develop proliferative ileitis (“wet tail” or regional enteritis), a disease characterized by excessive glandular proliferation in the epithelium of the ileum. Signs seen with this disease include diarrhea (sometimes containing blood) and tenesmus. Intestinal neoplasia of the colon and small intestine can cause similar signs and must be differentiated from proliferative ileitis. Rectal prolapse or intussusception of the small intestine or colon can result from the excessive straining seen with these conditions. Other conditions causing tenesmus, such as parasitism and diet-induced diarrhea, can also lead to prolapse or intussusception. Hamsters with rectal prolapse or intestinal intussusception are usually critically ill and are treated with fluids, dextrose, and antibiotics before they undergo emergency surgery. It is important to ascertain what segment of bowel is exposed before surgery. If there is an orifice at the end of the tissue, it is a section of bowel. If the tissue is solid, it may be a rectal mass or polyp that is protruding. Once you have determined that it is intestine, gently pass a small, blunt-ended probe (e.g., a tomcat catheter or cotton-tipped miniapplicator) alongside the exposed tissue. If the probe passes into the pelvic canal, it is an intestinal intussusception. If it does not advance, it is a rectal prolapse. A nonnecrotic rectal prolapse can be replaced and maintained in reduction with a purse-string suture or transanal sutures; however, a necrotic rectal prolapse must be amputated and sutured circumferentially. A small intestinal or colonic intussusception requires abdominal exploration to replace the intestine to its normal location and to assess the viability of the tissue and its blood supply. All these conditions are considered surgical emergencies. Lubricate a simple, nonnecrotic rectal prolapse and gently replace it with a cotton-tipped miniapplicator or a soft urinary catheter. Push the tissue cranially into its normal intrapelvic location cranial to the anal sphincter, then place a purse-string suture in the anus with 5-0 or 6-0 nonabsorbable suture material, snugly but not tightly, over a 5-Fr catheter. Alternatively, place two transanal sutures from cranial to caudal allowing fecal material to pass but holding the rectum inside. Remove the sutures in 3 to 5 days; meanwhile, treat the medical condition causing the prolapse. Amputate a necrotic rectal prolapse by making a fullthickness incision through the prolapsed tissue where it is healthy around half of the prolapse. Suture healthy rectum to healthy anus with 6-0 monofilament synthetic absorbable suture in a simple interrupted pattern. Amputate the remaining prolapsed tissue and close the second half of the anastomosis in the same manner. The terminal colon will thus be sutured to the anal mucocutaneous junction with suture knots in the lumen. A purse-string suture is not needed. Hamsters with an intussuscepted bowel have a poor prognosis for survival because they usually present in a debilitated
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A
B
C
D
E Fig. 28-15 A, This hamster developed a cheek pouch fistula at the caudal extent of the pouch. B, Food can be seen passing through the fistula. C, To remove the pouch, a skin incision has been made without incising the cheek pouch mucosa. D, The pouch is carefully dissected from the surrounding tissue from the caudal extent to the oral cavity; the oral mucosa is then closed with simple continuous or interrupted sutures. E, After copious irrigation of the remaining soft tissues, the skin is closed with a simple continuous pattern of polyglactin 910.
condition from the primary disease. Perform a routine ventral midline abdominal exploratory celiotomy and identify the intussuscepted segment. Gently reduce it, if possible, using a combination of traction on the orad segment and pushing the intussusceptum within the aborad segment. Resect any necrotic portion of the intussusceptum, preserving the blood supply to
the viable bowel, and perform an end-to-end anastomosis. If it is not possible to reduce the intussusception, transect the intestine orad to the intussusception. The aborad segment contains the intussusceptum. Incise the outer layer of intestine, which will allow the intussusceptum to be pulled out the anus by an assistant. Perform an end-to-end anastomosis of the healthy
CHAPTER 28 Soft Tissue Surgery
A
389
B Fig. 28-16 A, An everted cheek pouch in a dwarf hamster. B, A cotton-tipped applicator swab is used to hold the cheek pouch in position while a full-thickness percutaneous suture is placed.
intestine with 6-8 simple interrupted sutures of 6-0 or 7-0 monofilament absorbable material. Close the defect in the mesentery with 6-0 absorbable suture material. Intestinal plication is no longer recommended because it takes a lot of time and is associated with more postoperative complications than not plicating; however, an enteropexy is quickly performed and minimizes the risk of recurrence. Make a 1- to 2-cm incision in the peritoneum and tack the intestine at the site of the anastomosis to the incised peritoneum with 6-0 monofilament absorbable suture. Follow proper technique for enteric surgery, which includes changing gloves and instruments after completing the anastomosis, followed by irrigating the abdomen with warm saline. Closure is routine.
TAIL AMPUTATION Degloving injury to the tail is common in gerbils because the skin on the tail is more loosely attached than in other rodents; however, it does occur in other rodents. The skin over the tail is thin and easily pulled off during restraint or if it becomes caught. The tail can be mutilated by cage mates if the animals do not get along. The exposed, skinless tail will eventually desiccate and slough off, but it may be painful or, if it is denervated, the patient may chew on it. It is unlikely that sepsis would occur from this type of injury, but local infection is a risk. For these reasons, amputation is recommended. Sensation to the tail stump can be blocked by injecting 0.01 mL of bupivicaine or lidocaine proximal to the site. Place the animal in ventral recumbency and suspend the tail with tape while the site is prepared for aseptic surgery. A tourniquet can be used to control hemorrhage. Incise the healthy skin 1 to 2 mm proximal to the edge of the degloving to create fresh skin edges for suturing. Retract the skin proximally as far as possible and disarticulate the tail between the two most cranial coccygeal vertebrae that are exposed. This will allow adequate soft tissue coverage over the vertebral bone. Bleeding from the coccygeal vessels is generally easy to control
with cautery or a ligature. Close the subcutis with 5-0 or 6-0 absorbable suture over the exposed ends of the vertebrae and close the skin with an intradermal suture or tissue adhesive. Self-trauma to the amputation site is uncommon if gentle tissue handling is used.
RECOVERY During the recovery period, every attempt should be made to prevent heat loss. Hypothermia decreases the patient’s metabolic rate and the rate of excretion of drugs, prolonging the recovery period. During the recovery period, the patient should be placed in a well-ventilated, oxygen-enriched, warm, quiet environment. Incubators are ideal for this purpose. They allow for proper temperature control, administration of oxygen, and adequate humidity. If an incubator is not available, circulating-warm-water heating blankets, heat lamps, and warm-water bottles may be used for thermal support. Use caution with heat lamps and warm-water bottles, which can cause thermal burns. Forced-warm-air blankets and the Thermally Controlled Surgery Pads are safe and effective. Turn the animal every 30 to 60 minutes if it is recumbent to minimize the risk of hypostatic pulmonary congestion. Maintain it in a cage lined with clean paper towels to keep the incision clean as the scab forms; keep it isolated so that other animals do not traumatize the surgical wound. Also, other animals, including conspecifics, often unexpectedly attack an animal they perceive to be injured. Continue providing fluid therapy until the animal is eating and drinking well. Provide nutritional support with a syringe-feeding diet until no longer indicated. The animal should be eating and drinking well before it is discharged to the owner’s care. Try to avoid using bandages, because they are not well tolerated by rodents. A paste made from metronidazole tablets applied to the wound will often discourage self-trauma because of its bitter taste. Analgesic and antianxiety medications are very helpful at preventing self-trauma. A yoke can be fashioned to help prevent the animal from chewing at its wound.
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The technique for construction and application of such a yoke has been previously described.21 After applying the yoke, and before discharge, observe the animal carefully to make sure that it is eating and drinking normally. Evaluate the incision in 10 to 14 days and, if it is adequately healed, remove any sutures or staples that remain present.
ANALGESIA Providing preemptive, intraoperative, and postoperative analgesia is important in small rodents.35 Signs of pain in rodents include an increased heart rate, increased respiratory rate, depression, decreased appetite, trembling, and bruxism (see Chapter 39). Many agents have been given with good success and appear to be effective in providing the recovering patient with pain relief.7 Butorphanol (0.1-0.5 mg/kg SC, IM q4-6h for mice, and 2 mg/kg SC, IM q4-6h for rats) can be given pre- and postoperatively. Other analgesics include buprenorphine (0.050.1 mg/kg SC q8-12h in mice, and 0.01-0.05 mg/kg SC q8-12h in rats), which has a longer analgesic effect than butorphanol. Morphine (2-5 mg/kg SC q2-4h) is recommended in mice and rats. The veterinarian should consider the species variation in the dosage being used and the lasting analgesic effect. Morphine is not suitable as an analgesic agent in hamsters because this species shows high resistance to its effects. NSAIDs can have detrimental effects on kidney function, especially during periods of hypotension. Unless blood pressure can be monitored during surgery, it is safer to administer NSAIDs as postoperative rather than preemptive analgesics. Carprofen (4 mg/kg PO, SC q12h for rats and mice) has been recommended. Meloxicam (0.2-0.3 mg/kg PO q12h) has shown great efficacy as an analgesic in a clinical setting. For more information on analgesics used in rodents, see Chapter 31.
References 1. Bauck L, Orr JP, Lawrence KH. Hyperadrenocorticism in three teddy bear hamsters. Can Vet J. 1984;25:247-250. 2. Bauck LA, Hagan RJ. Cystotomy for treatment of urolithiasis in a hamster. J Am Vet Med Assoc. 1984;184:99-100. 3. Benitz KF, Kramer AW. Spontaneous tumors in the Mongolian gerbil. Lab Anim Care. 1965;15:281-294. 4. Bennett RA. Rodents: soft tissue surgery. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester: British Small Animal Veterinary Association; 2009:73-85. 5. Bingel SA. Pathologic findings in an aging Mongolian gerbil (Meriones unguiculatus) colony. Lab Anim Sci. 1995;45:597-600. 6. Bradshaw BS, Wolfe HG. Coagulation proteins in the seminal vesicle and coagulating gland of the mouse. Biol Reprod. 1977;16:292-297. 7. Cantwell SL. Ferret, rabbit, and rodent anesthesia. Vet Clin North Am Exot Anim Pract. 2001;4:169-191. 8. Capello V. Surgical techniques in pet hamsters. Exot DVM. 2003;5.3:32-37. 9. Capello V. Techniques for neutering pet hamsters. Exot DVM. 2003;5.4:21-26. 10. Collins BR. Common diseases and medical management of rodents and lagomorphs. In: Jacobson ER, Kollias GV, eds. Exotic animals. New York: Churchill Livingstone; 1988:261-316. 11. Cooper JE. Tips on tumors. Proceedings. North Am Vet Conf. 1994:897-898. 12. de Chaves G, Moretti M, Castro AA, et al. Effects of long-term ovariectomy on anxiety and behavioral despair in rats. Physiol Behav. 2009;97:420-425.
13. Dunn RC, Steinleitner AJ, Lambert H. Synergistic effect of intraperitoneally administered calcium channel blockade and recombinant tissue plasminogen activator to prevent adhesion formation in an animal model. Am J Obstet Gynecol. 1991;164:1327-1330. 14. Goya RG, Lu JK, Meites J. Gonadal function in aging rats and its relation to pituitary and mammary pathology. Mech Ageing Dev. 1990;56:77-88. 15. Greenacre CB. Spontaneous tumors of small mammals. Vet Clin North Am Exot Anim Pract. 2004;7:627-651. 16. Harkness JE. Anesthesia, surgery. In: Harkness JE, ed. A practition er’s guide to domestic rodents. Denver: American Animal Hospital Association; 1993:37-50. 17. Harkness JE, Turner PV, VandeWoude S, Wheeler C. Harkness and Wagner’s biology and medicine of rabbits and rodents. 5th ed. Ames, IA, Wiley-Blackwell; 2010. 18. Henry KR, Chole RA, McGinn MD. Age-related increase of spontaneous aural cholesteatoma in the Mongolian gerbil. Arch Otolaryngol. 1983;109:19-21. 19. Holmberg BJ. Enucleation of exotic pets. J Exot Pet Med. 2007;16:88-94. 20. Hotchkiss CE. Effect of surgical removal of subcutaneous tumors on survival of rats. J Am Vet Med Assoc. 1995;206:1575-1579. 21. Hoyt Jr RF. Abdominal surgery of pet rabbits. In: Bojrab MJ, ed. Current techniques in small animal surgery. 4th ed. Baltimore: Williams & Wilkins; 1998:777-790. 22. Jackson TA, Heath LA, Hulin MS, et al. Squamous cell carcinoma of the midventral abdominal pad in three gerbils. J Am Vet Med Assoc. 1996;209:789-791. 23. Jenkins JR. Surgical sterilization in small mammals. Spay and castration. Vet Clin North Am Exot Anim Pract. 2000;3:617-627. 24. Johnson-Delaney C. Ovariohysterectomy in a rat. Exot DVM. 2002;4.4:17-21. 25. Krohn B, Erben RG, Weiser H, et al. Osteopenia caused by ovariectomy in young female rats and prophylactic effects of 1,25-dihydroxyvitamin D3. Zentralbl Veterinarmed A. 1991; 38:54-60. 26. Krohn DE, Barthold SW. Biology and diseases of rats. In: Fox JG, Cohen BJ, Loew FM, eds. Laboratory animal medicine. Orlando: Academic Press; 1984:116-122. 27. Lejnieks DV. Urethral plug in a rat (Rattus norvegicus). J Exot Pet Med. 2007;16:183-185. 28. Lewis W. Cystic ovaries in gerbils. Exot DVM. 2003;5.1:12-13. 29. Lidderdale JA, St Pierre SJ. Cystotomy for treatment of urolithiasis in a hamster. Vet Rec. 1990;127:364. 30. Lipman NS, Foltz C. Hamsters. In: Laber-Laid K, Swindle MM, Flecknell P, eds. Handbook for rodent and rabbit medicine. New York: Pergamon; 1995:65-82. 31. Mighell JS, Baker AE. Caesarean section in a gerbil. Vet Rec. 1990;126:441. 32. Nishizumi K, Fujiwara K, Hasegawa A. Cutaneous mastocytomas in Djungarian hamsters. Exp Anim. 2000;49:127-130. 33. Powers MY, Campbell BG, Finch NP. Preputial damage and lateral penile displacement during castration in a degu. J Am Vet Med Assoc. 2008;232:1013-1015. 34. Prejean JD, Peckham JC, Casey AE, et al. Spontaneous tumors in Sprague-Dawley rats and Swiss mice. Cancer Res. 1973;33:2768-2773. 35. Redrobe S. Soft tissue surgery of rabbits and rodents. Sem Avian Exot Pet Med. 2002;11:231-245. 36. Rembert MS, Johnson AJ. What’s your diagnosis? Pigmented mass in an experimental gerbil. Spontaneous malignant melanoma. Lab Anim. 2001;30:22-25. 37. Roman CD, Hanley GA, Beauchamp RD. Operative technique for safe pulmonary lobectomy in Sprague-Dawley rats. Contemp Top Lab Anim Sci. 2002;41:28-30.
CHAPTER 28 Soft Tissue Surgery 38. Shumaker RC, Paik SK, Houser WD. Tumors in Gerbillinae: a literature review and report of a case. Lab Anim Sci. 1974;24:688-690. 39. Soga M, Kishimoto Y, Kawamura Y, et al. Spontaneous development of hepatocelluar carcinomas in the FLS mice with hereditary fatty liver. Cancer Lett. 2003;196:43-48. 40. Tanaka A, Hisanaga A, Ishinishi N. The frequency of spontaneously-occurring neoplasms in the male Syrian golden hamster. Vet Hum Toxicol. 1991;33:318-321. 41. Toft JD. Commonly observed spontaneous neoplasms in rabbits, rats, guinea pigs, hamsters, and gerbils. Sem Avian Exot Pet Med. 1992;1:80-92.
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42. van Goethem B, Schaefers-Okkens A, Kirpensteijn J. Making a rational choice between ovariectomy and ovariohysterectomy in the dog: a discussion of the benefits of either technique. Vet Surg. 2006;35:136-143. 43. Ward JM, Fox JG, Anver MR, et al. Chronic active hepatitis and associated liver tumors in mice caused by a persistent bacterial infection with a novel Helicobacter species. J Natl Cancer Inst. 1994;86:1222-1227.
SECTION FIVE
Other Small Mammals
CHAPTER
29
Sugar Gliders
Robert D. Ness, DVM, and Cathy A. Johnson-Delaney, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal)
Biology Natural History Anatomy and Physiology Behavior Husbandry Clinical Techniques Handling and Restraint Blood Collection Treatment Techniques Radiography Drug Dosages Diseases and Syndromes Gastrointestinal Disease Respiratory Disease Urogenital Disease Reproductive Disorders Reproductive Tract Infection Dermatologic Disorders Ophthalmic Disorders Retrobulbar Abscess Musculoskeletal Disease Neurologic Disease Infectious Disease Neoplasia Surgery and Anesthesia Anesthesia Soft Tissue Surgery
Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
BIOLOGY NATURAL HISTORY Sugar gliders (Petaurus breviceps) are small nocturnal arboreal marsupials native to New Guinea and the eastern coast of Australia.1,5,20 They inhabit open areas in tropical or coastal forests and dry inland sclerophyll tropical forests. They are social animals, with colonies of 6 to 10 animals occupying a territory of up to 1 hectare. Dominant males mark territory and other group members with scent gland secretions. Animals in a group nest communally in leaf-lined tree holes. During periods of extreme cold or food scarcity, sugar gliders conserve energy by going into torpor for as many as 16 hours per day.5
ANATOMY AND PHYSIOLOGY Marsupials: General Information Marsupials are best known for possessing a pouch, in which the female raises its young. The degree of pouch enclosure is dependent on the species. The pouch is absent in males and in female South American short-tailed opossums, which are considered to be more primitive marsupials. The female sugar glider has a pouch containing four teats, in which she raises one or two young. Epipubic bones (ossa marsupialia or eupubic bones) are unique to certain marsupials, but they are diminished or absent in gliders. These small bones are thought to provide an attachment for muscles that support the pouch. Their absence may be an adaptation to gliding, which reduces skeletal weight. The metabolism of marsupials is approximately two-thirds that of placental (eutherian) mammals. The normal heart rate 393
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SECTION V Other Small Mammals
Fig. 29-1 The patagium, or gliding membrane, of sugar gliders stretches between the front and hind legs.
Fig. 29-3 The male sugar glider has a frontal scent gland on its head. Secretions from this gland are used to mark territory and group members.
Fig. 29-2 The hind foot of the sugar glider has five toes, with syndactyly of the second and third digits and an opposable first digit.
Fig. 29-4 The male glider also has a gular scent gland, located on the ventral aspect of the throat.
of a sugar glider is 200 to 300 beats per minute; the respiratory rate is 16 to 40 breaths per minute.1 The cloaca is a common terminal opening of the rectum, urinary ducts, and genital ducts. Cloacal temperature is lower than the actual body temperature; the average cloacal temperature being 89.6°F (32°C).1,4,8,10 True rectal temperature in marsupials can be measured by directing the thermometer dorsally into the rectum from within the cloaca. The rectal temperature is usually 97.3°F (36.3°C).4 Measurement of the tympanic temperature is another means of determining core body temperature.
Anatomic Characteristics Wild sugar gliders have soft, velvety fur that is gray, with a central black stripe dorsally; it is cream-colored ventrally. Like American flying squirrels (Glaucomys volaris, Glaucomys sabrinus), sugar gliders have a patagium (gliding membrane) that stretches between their front and hind legs (Fig. 29-1). There are at least seven recognized subspecies of sugar gliders. Although average body weight varies among the subspecies, typical adult males weigh between 100 and 160 g and females between 80 and 130 g. Body length ranges from 16 to 21 cm and tail length from 16.5
to 21 cm. Gliding distances are reported to be as long as 50 m.26 Being a nocturnal prey species, sugar gliders have large, protruding, widely spaced eyes. They have five toes on their hind feet, with an opposable first digit and syndactyly of the second and third digits (Fig. 29-2). Dominant males mark territory and group members with secretions from androgen-sensitive frontal (forehead) (Fig. 29-3), gular (throat) (Fig. 29-4), and paracloacal scent glands (Fig. 29-5).
Color Variations and Genetics Sugar gliders have become available in a variety of colors through selective genetic breeding in recent years. Although there has not been thorough genetic testing on these variations, some genetic trends have been realized by applying Mendel’s laws of heredity to well-documented breeding programs. Several color variations have been described and accepted as “standard” mutations. These variations of the natural gray line include the white face, white tip, leucistic, and mosaic. There is also a redseries classification that includes red cinnamon, lion, butter cream, and chocolate.
CHAPTER 29 Sugar Gliders
395
Fig. 29-6 The leucistic sugar glider is also known as the blackFig. 29-5 Paracloacal glands are most developed in males and
eyed white, a rare color variation.
may become infected and impacted, as seen here.
The white face is not as much a separate coloration as it is a mutation of the stripes around the face. These gliders are missing the black bar below the ears and usually have lighter or broken eye rings. They are the natural gray-and-black coloration otherwise. The white tip coloration is also just a slight variation of other color varieties. As the name implies, these gliders have white on the ends of their tails. This may indicate the presence of the subspecies Petaurus breviceps ariel in the lineage, but the precise genetic connection is not known. It is believed that the white tip is recessive to normal, but not as a single recessive gene. The leucistic sugar glider is all white with black eyes; therefore it is also referred to as black-eyed white (Fig. 29-6). Some leucistic gliders may have faint yellowish markings where the black would be on the standard gray coloration. The leucistic gene is recessive to the normal gray, following the principles of Mendelian genetics. The breeding of a heterozygous leucistic to a leucistic yields 50% leucistic and 50% heterozygous. Breeding two heterozygous leucistics can produce heterozygous, standard, or leucistic offspring in the expected ratios. This is a relatively rare and valuable coloration, whether the glider is white or just carrying the gene as a heterozygous leucisitc. The albino sugar glider differs from the leucistic in that it is all white with red eyes, as seen in other albino animals. As in other species, albinism is the genetic inheritance of recessive alleles manifesting as a lack of melanin pigmentation in the eyes, skin, and hair. These sugar gliders are more rare and valuable. A mosaic sugar glider has characteristic white patches on the body, feet, or tail. If the mosaic coloration involves the tail, the sugar glider is referred to as a ring-tail mosaic. The mosaic trait is codominant to the standard gray coloration; therefore gliders cannot be heterozygous for the mosaic gene. Certain lines of mosaics may produce sterile males, although the females are fertile. Breeding of mosaics over several generations, however, can breed out the sterile male condition. Mosaic sugar gliders are a relatively rare and expensive color variation. The red-series gliders are subdivided into four basic colors; chocolate, butter cream, lion, and red cinnamon. These may indeed just be shade variations of the reddish coloration, since these variations are not clearly defined or well established. The
chocolate is described as having a brown coat with either brown or black markings. The butter cream has a brown coat with cream to yellow highlights and red or brown markings. The lion is considered to be a standard variation and has a golden brown coat with red markings. The red cinnamon has a reddish coat and red or brown markings. The color of these strains becomes more faded when bred to the standard gray coloration, and the color can be bred out of a line. These colorations are not as valuable as the lighter-colored leucistic or mosaic varieties. Other color varieties have been described, with more being identified and developed by genetic breeding programs. The variations described above are those more commonly accepted, but they are not fully standardized or genetically defined. As fanciers continue to breed for more unusual mutations and unique colorations, inadvertent selection for genetic disorders and instability is bound to develop as well.
Reproduction Sugar gliders are seasonally polyestrous, with the natural breeding season in Australia occurring between June and November. They are polygamous, with a dominant male that breeds with the mature females in the colony.5 Young (joeys) are typically born in the spring, when insects are plentiful. Litter size is usually two (81%) or, less commonly, one (19%).5 Two litters in a single breeding season are common. The estrous cycle is 29 days in length, and gestation is 15 to 17 days.1,14,15,27 Young weigh only 0.2 g at birth, when they migrate to the pouch. They remain in the pouch for 70 to 74 days (Fig. 29-7). After they outgrow the pouch, the young are left in the nest until they are weaned at 110 to 120 days of age. Table 29-1 lists the expected growth rate and specific developmental milestones as the joey ages.1,10 The young remain with the colony until they are forcibly dispersed, at 7 to 10 months of age.1,5 Sexual maturity is reached at 8 to 12 months of age in females and 12 to 15 months in males.1,14,15 The reproductive anatomy of sugar gliders is unique compared with that of other mammals routinely treated in practice. Female sugar gliders have two uteri and two long, thin, lateral vaginas that open into a single cul-de-sac divided by a septum. Both sexes have paracloacal glands, which are more developed in the male.25 Males have a large prostate with a constriction at the anterior third. They also have two pairs of Cowper’s glands in addition to three paracloacal glands. The testes are located in a prepenile pendulous scrotum, and the penis is bifid
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SECTION V Other Small Mammals
(Fig. 29-8). Males do not urinate from the forked end of the penis but rather from the proximal portion. Therefore the distal penis can be amputated in cases of penile trauma or paraphimosis. A captive male sugar glider was presented to one of the authors (RDN) with the unusual development of a pouch as well as having the typical bifid penis. This individual lacked a scrotum but may have cryptorchid testes, since he also developed the classic alopecia on the forehead and upper chest present with male scent glands. It is not known if this is a true hermaphrodite, since an abdominal evaluation was not performed to identify a female reproductive tract. It is anecdotally reported that similar gliders have produced offspring and raised them in their pouches.
Males develop a scent gland on the forehead and may rub this on the female’s chest. Males also have scent glands on the chest and paracloacal glands. Both sexes mark territory with secretions from scent glands. In addition, the female uses urine to mark territory. The female’s scent glands are within her pouch, and she will secrete and increase marking to indicate breeding readiness to the male.
Sugar gliders are quite vocal, with a whole series of alarm yaps and screams. They are highly social animals and should not be kept as solitary pets; they become clinically depressed when housed singly. Self-mutilation is not uncommon in solitary gliders. Those without proper socialization or exercise and territorial space may become aggressive. Males will fight if there is not enough distance between nest boxes. Both sexes may fight and become irritable at the beginning of breeding. Males without adequate stimulation may become aggressive toward humans. Neutering may help, but they will continue to scentmark and will not be “human socialized” like a domesticated animal. Sugar gliders make acceptable pets with the appropriate caretakers if they are given sufficient socialization and ample space. When hand-reared, they adapt well to captivity and develop a strong bond with their human companions. As pets, their care is time-consuming, with recommended socialization periods of at least 2 hours per day. Basic handling for socialization and companionship is best achieved at night, when the animals are more interactive and playful. If awakened and handled during the day, sugar gliders become agitated and irritable. Sugar gliders are most responsive to and trusting of individuals they know, but they respond well if approached with
Fig. 29-7 The female sugar glider raises one or two joeys in its
Fig. 29-8 In male sugar gliders, the penis is bifid. Male gliders
pouch.
do not urinate from the tip of the forked penis but from its base.
BEHAVIOR
Table 29-1 Growth and Development of Young Sugar Gliders1,10 Age (days)
Weight (g)
Milestones
Total Daily Intake (mL) — 0.2 0.6 2.5
1 20 40 60
0.2 0.8 3.2 12
70 80-90 100 120-130
20 35-44 54 78
Birth Ears free from head; papillae of whiskers visible Ears pigmented; whiskers erupted; eye slits present Intermittently detached from teat; dorsal stripe, fur, and gliding membrane developing Left in nest; eyes open; fully furred Fur lengthens Emerging from nest, starting to eat solids Weaning
200
100
Subadult
4 6-8 8-10 Feed up to 20% of body weight in pureed solids Feed up to 20% of body weight in solids
CHAPTER 29 Sugar Gliders patience and gentle handling. They enjoy cuddling and curling up in shirt pockets or pouches, where they feel safe. They should not be allowed to crawl into tight-fitting clothes, such as pants pockets, because of the risk of injury. Advantages of these pets include their small size, playfulness, and intelligence. The life span of sugar gliders is longer than that of other comparably sized pets; they can live from 10 to 12 years in captivity.1 Disadvantages include their nocturnal nature, housing requirements, specific dietary needs, and musky odor.
HUSBANDRY Caging Because of their active nature, sugar gliders should have cages as large as possible. These animals need space to climb, run, and jump. Minimum cage size is 36 × 24 × 36 in. (91 × 61 × 91 cm), but larger is better. Cages should be made of wire for good ventilation, with wire spacing no more than 1.0 × 0.5 in. (2.5 × 1.3 cm) wide. Sugar gliders tolerate temperatures between 65°F and 90°F (18°C and 32°C), with an ideal range of 75°F to 80°F (24°C-27°C).4,22 The cage must have designated areas for food, water, shelter, and exercise. Several food and water dishes should be placed in various locations throughout the cage. A nest box or sleeping pouch positioned high in the cage gives the sugar glider a place to sleep during the day. Branches, perches, and shelves can be placed at various levels of the cage to satisfy the natural behavior of climbing. Sugar gliders enjoy playing with bird toys, such as swings and chew toys. Plastic wheels without open rungs, such as a hamster wheel, are used for exercise. A variety of objects may be placed throughout the cage to stimulate and entertain these animals. Sugar gliders need permanent access to a nest box for sleeping and hiding, with minimal daytime disturbances. Nest box size depends on colony size, but boxes should be at least 6 × 6 in. (15 × 15 cm) with a hinged lid and a circular opening on the front with a sliding closure.1,5,16,22 Bird nest boxes or small hollow logs are suitable. Nesting material can consist of hardwood shavings, recycled paper products, shredded bark, dried leaves, coconut fibers, sea grass, or equivalent materials. Artificial wool, cloth strips, and similar materials are not suitable because fibers have been known to entrap an animal’s limbs. Nest boxes must be cleaned and bedding material changed regularly, at least every 1 to 2 weeks. If nest box closures are secure, the nest box can be relocated from an outdoor aviary into the house for interaction each evening and then returned to the aviary to allow for nocturnal activity. These animals need large areas for activity and exercise, ideally the freedom of a whole room equipped with vertical “trees.” This may be impractical in a typical household, and many owners are reluctant to allow such freedom because gliders scentmark and are active at night.
Nutrition and Feeding Natural Diet. Sugar gliders are omnivorous. The diet of a wild sugar glider can include sap and gums (carbohydrate-rich) from eucalyptus and acacia trees, nectar and pollen, manna and honeydew, and a wide variety of insects and arachnids.5,13 Sugar gliders have specialized lower incisors for chewing and gouging into the bark of trees. The lengthened fourth digit on the manus aids in extracting insects from crevices. Sugar gliders also have an enlarged cecum, which functions principally in microbial fermentation of complex polysaccharides in gum.
397
The diet varies with the season. During the spring and summer months, these animals are primarily insectivorous. During the winter months, sugar gliders feed on gum from the eucalyptus and acacia trees as well as on sap and sugar from the trees and sap-sucking insects. Small insects trapped in wattle or acacia gum are consumed. Favorite trees include the Australian “bloodwood,” the red sap of which crystallizes, mixes with the decaying pulp of the trees, and attracts insect activity. Other trees produce “manna,” a deposit of white encrusting sugars from wounds made by sap-sucking insects, birds, other gliders, or possums. The sugar glider has been observed to eat honeydew, secreted by sap-sucking insects. Field energetics studies have demonstrated that wild sugar gliders consume 182 to 229 kilojoules (kJ)/day.19 This is equivalent to approximately 17% of body weight in wet weight of food.19 Captive gliders expend less energy in exercise and consume more easily assimilated foods than do those in the wild; therefore the total energy offered to gliders in captivity should be less. Captive Diet. Captive diets must satisfy the numerous specializations of these insectivorous omnivores. The captive diet should include nectar, insects, and other protein sources as well as very limited amounts of fruits and vegetables.3,13,14 Portion size for one glider is roughly a tablespoon of insects or insectivore diet, a tablespoon of nectar, and one-half teaspoon of fruits, other types of insects, etc. The diet should be offered in fresh portions in the evening. Protein is a critical nutrient in the diet of sugar gliders. Various protein sources include insects (mealworms and crickets), eggs, newborn mice, lean meat, and commercial protein sources (pelleted insectivore diets or monkey chow). Commercially available adult insects should be fed a calcium-rich insect diet for several days before being offered to the gliders. Larval forms should be kept to a minimum. Approximately 50% of the diet should consist of sources of fruit sugars, preferably in the form of a sap or nectar. Sources include fresh nectar, maple syrup, honey, and artificial nectar products.3,13,14,22 Examples of commercial products include prepared lory diets and Gliderade (Avico, Fallbrook, CA). Gum Arabic (acacia) can be purchased as a powder, mixed into a thick paste, and used to simulate native gums: it can be used in holes in branches and on surfaces, with insects or bits of fruit stuck to it for enrichment and foraging. Various commercial diets for sugar gliders and insectivores are available and may be included as a part of the diet. Leafy green vegetables provide a source of fiber and some vitamins. Sugar gliders accept a wide variety of other foods, including fruits, vegetables, nuts, and seeds (sunflower and pumpkin), but these should be offered in very limited quantities. Fruit juices and strained baby food can be offered if they are free of preservatives, but they are not as appropriate as the nectar-based formulas. Because these foods are not a significant component of the natural diet, they should constitute less than 10% of the captive diet. Sprinkle a broad-spectrum vitamin and mineral supplement with a good calcium supply on the food daily. Contrary to the nutritional needs observed in the wild, much of the information found in lay publications lists fruits and vegetables as a major portion of the captive diet. Fruit-based diets are harmful to captive sugar gliders because they provide inadequate protein and calcium and predispose animals to osteoporosis and periodontal disease.3,22 Although sugar gliders readily accept fruits, nuts, and grains, these are not a substantial part of their natural diet.
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SECTION V Other Small Mammals
Leadbeater’s diet has been recommended as a base mixture for many sugar glider diets. This is an artificial nectar mix originally formulated for Leadbeater’s possums (Gymnobelideus leadbeateri).3,13,16 Leadbeater’s recipe consists of 150 mL water, 150 mL honey, 1 shelled hard-boiled egg, 25-g high-protein baby cereal, and 1 teaspoon vitamin and mineral powder. This mixture is kept refrigerated until served, with unused refrigerated portions discarded after 2 to 3 days. The mixture can be kept longer if frozen. The original Leadbeater’s recipe is often modified, with adjustments usually being made for palatability rather than nutritional content. Several modified versions are found on the Internet, such as Boubon’s Modified Leadbeater’s Diet and the High Protein Wombaroo Diet (Wombaroo, Adelaide, SA, Australia). These diets should be scrutinized closely, because none of them have undergone thorough nutritional dietary trials and analysis. Several diets recommended by the authors are listed in Table 29-2. Choose one of these recipes and use it in its entirety. Chop the pieces together so that the gliders cannot pick out only their favorite items. No one commercial diet seems adequate, but long-term nutritional studies are still pending.3
Hand-Rearing Orphaned marsupial-pouch young must be maintained at a stable temperature close to the body temperature of the mother, which may range from 86°F to 93°F (30°C-34°C). The young depend less on artificial heat as their fur begins to grow. A snug artificial pouch can be made from a child’s cotton sock to provide security and warmth. Place the pouch in a temperaturecontrolled box. By the time the pelage is thick and complete, the orphan has reached a homeothermic state and no longer requires an artificial heat source. If the age of the orphaned young is not known, estimate the age from the body weight and stage of development (see Table 29-1). Sugar glider pouch life is 70 to 74 days, and weaning occurs at 120 days.1,10,12 From day 75 to day 100, the young are left in an insulated nest while the mother is foraging. When an orphan has reached this stage, a soft toy the size of an adult sugar glider is useful to provide security from the time of pouch exit to weaning. The artificial nest can be a fleecy cotton bag that is big enough to accommodate the animal and the toy; it should be placed in a box maintained at 77°F (25°C). Feed unfurred young every 1 to 2 hours (including throughout the night) and feed just-furred young every 4 hours. Gradually reduce the frequency to twice daily, then once daily, until the young are weaned. Feed a low-lactose milk formula (e.g., puppy Esbilac [PetAg, Inc., Hampshire, IL]). Marsupial milk increases in energy at the time of pouch exit to provide for the young’s increased energy demands of locomotion and thermoregulation. This change can be simulated by adding canola oil (rapeseed oil) to the milk at the rate of 1 mL of oil per 20 mL of milk. Juvenile sugar gliders usually lap readily from the tip of a syringe, or they can be taught to lap from a small plastic lid. At each feeding, measure and record milk intake. Measure body weight daily until the weight stabilizes, then weekly. Frequency of feeding and quantity of food can be adjusted to achieve a satisfactory growth rate. Start offering solids at about 100 days, at which time the young glider should weigh approximately 54 g, and wean at 130 days, when body weight is approximately 78 g. Pureed baby food with meat and vegetables or blended adult diet is a suitable starter food.
CLINICAL TECHNIQUES HANDLING AND RESTRAINT Sugar gliders are rather easy to handle but can be difficult to restrain. As mentioned previously, they are very energetic and hyperactive during the late evening hours, which makes them difficult to handle or restrain. Therefore schedule clinical examinations early in the day when the animals are normally less active. Nest boxes make useful transport boxes if they are constructed with hinged lids and lockable slides over the circular entrance. If the hinged lid is partially opened, the glider can be captured with a hand protected by a small cotton bag, which is then inverted over the glider. Bag seams should be double-sewn so that no frayed threads can become tangled around the sugar glider’s appendages. Palpate the head through the bag, then grasp the glider firmly around the back of the head, behind the ears, as you cup its body in the rest of your hand. Peel the bag back to expose the face for examination or to apply a face mask to induce anesthesia.20 Other methods of restraint include grasping the glider at the base of its tail or cupping the animal in the palm of your hand. The ventrum and hindquarters can be examined by lifting the animal by the base of its tail while allowing the glider to grasp a surface with its front feet. This method also permits palpation of the abdomen and examination of the pouch. To examine the head and chest, hold the sugar glider in a small towel with its head exposed. These animals typically do not tolerate being scruffed.20 Full clinical examination or diagnostic sampling is best performed with the animal under sedation or isoflurane anesthesia. Do a complete head-to-toe examination, assessing the function of all major systems. Measure the temperature, pulse, and respiratory rate and weigh the animal before it recovers from anesthesia. During recovery, place the sugar glider in a cotton bag or pouch and administer oxygen by face mask. Recovery is usually rapid. Assess locomotion before anesthesia or after full recovery, when the sugar glider is contained in a small cage with simple horizontal perches.
BLOOD COLLECTION Collecting a blood sample from a sugar glider can be difficult. Sedation or full anesthesia is required for diagnostic sampling. Use a 0.5-mL insulin syringe or a 1-mL tuberculin syringe with a 25- to 27-gauge needle to collect blood samples. Because of the small size of these animals, only small volumes of blood (up to 1% of body weight in grams) can be safely drawn. Reliable venipuncture sites in sugar gliders are the jugular vein and the cranial vena cava (Fig. 29-9).11,21,22 These sites yield the greatest blood volume (up to 1 mL) from an adult sugar glider. For jugular venipuncture, insert the needle midway between the point of the shoulder and the ramus of the mandible on the ventral aspect of the neck, lateral to the trachea and esophagus. The cranial vena cava can be sampled at the thoracic inlet. The needle is directed caudally at a 30-degree angle off midline toward the opposite hind leg.11,21 Small blood samples can be obtained from various peripheral veins. The medial tibial artery can be accessed to collect up to 0.5 mL of blood.1,21 Although this vessel is readily visible, it is quite mobile. For venipuncture of the cephalic, lateral
CHAPTER 29 Sugar Gliders Table 29-2 Suggested Sugar Glider Diets A. Dr. Cathy Johnson-Delaney’s Sugar Glider Diet Feed in the evening. 50% Leadbeater’s Mix, 50% insectivore/carnivore diet. Leadbeater’s Mix: 1 glider portion is approximately 2 tablespoons: Warm water 150 mL. Honey 150 mL. 1 shelled boiled egg. 1 tablespoon powdered/flaked baby cereal. 1 teaspoon vitamin/mineral supplement such as Prime or Vionate; additional calcium carbonate can be added (100 mg). Mix warm water, honey. In separate container, blend egg until homogenized, gradually add water/honey, then vitamin powder, then baby cereal, blending after each addition until smooth. Keep refrigerated until served. This can also be frozen in ice cube trays; one well is approximately one meal’s worth. Insectivore/carnivore diet: Reliable Protein Products Insectivore Diet (760-321-7533, fax 760-321-0395, www.zoofood.com;
[email protected]) or commercial sugar glider pellet. Treat foods: should not be more than 1 to 2 teaspoons/glider/day: Fruit, various, chopped; may add bee pollen, vitamin/mineral supplement. Live insects, calcium-gut loaded for several days, adult insects preferred. Can also dust insects prior to giving them to the gliders. Blooming eucalyptus branches when available. Commercial lorikeet nectar can also be offered several times a week. Gum arabic can have additional calcium added, mix with fruit juice or lorikeet nectar, and use it smeared on branches, as enrichment treat.
B. Chicago Zoological Park Diet Recipe for One Sugar Glider 1 teaspoon-sized piece each, chopped: apple, carrot, sweet potato, banana. 1 teaspoon leaf lettuce. 1 hard-cooked egg yolk. 1 tablespoon Nebraska Feline Diet, ZuPreem, or Mazuri (or other zoo-quality feline diet). 1 dozen mealworms.
C. Taronga Zoo Sugar Glider Diet Recipe Feeds Two Sugar Gliders (Taronga Zoo, Sydney, Australia) Leadbeater’s mix (see above): 2 teaspoons. Apple, 3 g; banana/corn, 3 g; dog kibble, 1.5 g; fly pupae, 1 teaspoon. Grapes/kiwi fruit, 3 g; orange with skin, 4 g; pear, 2 g; Rock melon/melon/pawpaw (papaya), 2 g; sweet potato, 3 g. Weekly: feed day-old chick; when available, large insects or mealworms.
D. Dr. Debra McDonald’sa Sugar Glider Diet Water: ad lib. Daily diet per animal: 1 piece of Eukanuba dog chow. 6 g fruit, chopped (approx 1 tablespoon). 1 teaspoon nectar mix (commercial lorikeet nectar). 1 g fly pupae (1⁄4 teaspoon). 5 g corn (1⁄4 thin slice). 2 g sprouted seed. 2 mealworms. Supplement 5 pollen grains weekly. 3 sultanas, 3-4 times weekly. 2 sunflower seeds weekly. 1 g Pet Health Food (Australian product) (small cube) weekly. 1 almond weekly. Insects, 3-4 times weekly. Acacia, eucalypts, other blossoms as available (gliders will lick/eat nectar). Continued
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SECTION V Other Small Mammals Table 29-2 Suggested Sugar Glider Diets—cont’d E. Dr. Rosemary Booth’s Sugar Glider Diet Offer a total of 15% to 20% of body weight daily. Select 1 diet from each group each day. Animals will benefit from a major effort to provide a regular supply of vitamin/mineral-enriched insects. Group 1: Insects: 75% moths, crickets, beetles. 25% fly pupae, mealworms. Meat mix: commercial small carnivore or insectivore diet. Group 2: Nectar mix: 1.5 cups fructose, 1.5 cups sucrose (brown sugar), 0.5 cup glucose made up to 2 L with warm water. (Commercially available mixes have the advantage of some vitamin/mineral additives.) Dry lorikeet mix: 4 cups rolled oats, 1 cup wheat germ, 1 cup brown sugar, 0.5 cup glucose, 0.5 cup raisins or sultanas. Group 3: Fruit and vegetables: select from diced apple, nectarine, melons, grapes, raisins, sultana, figs, tomato, sweet corn kernels, sweet potato, beans, shredded carrot, butternut pumpkin. Greens: mixed sprouts, lettuce, broccoli, parsley, with a vitamin/mineral supplement.b Equal portions of all three groups; do not skimp on ingredients. It is necessary to have a variety of insects. aFormer
nutritionist at Healesville Sanctuary, Victoria, Australia. Vionate (Gimborn Pet Specialties LLC, Atlanta, GA, 1-800-755-7056) is a good vitamin/mineral supplement. 1⁄8 teaspoon per glider per group 3 portion. Nursing/breeding: add in additional calcium carbonate per group 3 portion.
bNote:
TREATMENT TECHNIQUES
Fig. 29-9 The cranial vena cava is a common venipuncture site in sugar gliders, made with the use of a 1-mL tuberculin syringe with a 26-gauge needle.
saphenous, femoral, or ventral coccygeal veins, use a 0.5-mL insulin syringe with a 27-gauge needle. These veins are superficial and easily collapse if too much negative pressure is applied. Up to 0.25 mL of blood can be drawn from these veins. Hematologic and plasma biochemical values obtained from captive sugar gliders in clinical practice are listed in Table 29-3.21 Data from the International Species Information System on reported hematology and biochemistry in sugar gliders have been included for comparison.9 However, these values should be used only as guides to normal values, as the sample sizes used in both collections were too small for these to be considered as definitive reference ranges.
Fluid therapy is administered by routes commonly used in other small animals and includes the oral, subcutaneous, and intraosseous routes. Administer subcutaneous fluids along the dorsal midline of the thorax. Take care not to administer the fluids laterally into the patagium because fluids are more slowly absorbed from this area and can cause discomfort and distress. Administer intraosseous fluids in the proximal femur by using the same technique to place an intraosseous catheter as is used in other small mammals. Calculate the volume of fluids to be administered by estimating the percent dehydration in addition to the daily fluid requirements (comparable to methods used in similarly sized mammals). The choice of fluids depends on the animal’s condition and diagnosis. Injections can be given by a variety of routes and at various sites. Give intramuscular injections in the epaxial muscles of the neck and upper thorax or in the muscle mass in the anterior thigh (biceps femoris). Subcutaneous injections are safely administered along the dorsal midline of the thorax. Intravenous injections are difficult and must be administered in the cephalic or lateral saphenous veins with the animal under sedation or general anesthesia.
RADIOGRAPHY General anesthesia is required to position a sugar glider for diagnostic radiographs. If young are present in the pouch but detached from the teat, remove them before radiography. A marsupial, however, should never be pulled off a teat. Reflect the patagium away from the body wall to minimize artifact and improve visualization on lateral views. In very small animals, digital radiography or mammography yields superb definition of both soft tissues and skeletal
CHAPTER 29 Sugar Gliders
401
Table 29-3 Hematologic and Plasma Biochemical Reference Values for Captive Sugar Gliders CLINICAL STUDY21 ISIS Measurement
n
Values9
(95% CI)a
Hemoglobin (g/dL) Hematocrit (%) Red blood cell count (x106/μL) Mean corpuscular hemoglobin concentration (g/dL) Mean cell volume (μm3) Mean corpuscular hemoglobin (pg) Platelets (x103/μL) White blood cell count, total (x103/μL) Neutrophils (x103/μL) Bands (x103/μL) Lymphocytes (x103/μL) Monocytes (x103/μL) Eosinophils (x103/μL) Basophils (x103/μL) Sodium (mmol/L) Potassium (mmol/L) Chloride (mmol/L) Blood urea nitrogen (mg/dL) Creatinine (mg/dL) Bilirubin, total (mg/dL) Alkaline phosphatase (AP) (IU/L) Alanine aminotransferase (ALT) (IU/L) Aspartate aminotransferase (AST) (IU/L) Lactase dehydrogenase (LDH) (IU/L) Creatine phosphokinase (IU/L) Protein, total (g/dL) Albumin (g/dL) Globulin (g/dL) Glucose (mg/dL) Cholesterol (mg/dL) Calcium (mg/dL) Phosphorus (mg/dL)
2 8 9 3
13-15b 45-53 5.1-17.8 31-33b
15.4±1.6 43.9±4.0 7.8±0.9 35.1±2.0
3 3 7 10 10 10 10 9 10 10 9 9
58-62b 18-21b 105-220 5.0-12.2 1.5-3.0 0 2.8-9.2 0.1-0.2 0-0.1 0 135-145 3.3-5.9 — 18-24 0.3-0.5 0.4-0.8 — 50-106 46-179 — 210-589 5.1-6.1 3.5-4.3 — 130-183 — 6.9-8.4 3.8-4.4
56.8±5.4 19.9±1.3 728±176 6.7±4.9 1.2±1.0 0.12±0.05 5.2±4.4 0.18±0.17 0.18±0.25 0.04 143±4 3.5±0.7 106±1 17±7 0.8±0.3 0.3±0.2 196±35 70±40 72±67 246±33 639±477 6.1±0.6 4.0±0.7 2.3±0.8 139±78 161±2 7.5±3.5 7.0±2.2
9 11 11 10 9 7 11 10 11 9 5
aConfidence
interval. Values shown are the 95% confidence intervals after outliers were removed. Statistically, 90% of the population should have values within these limits. Because the sample size is small, these values should be used as guidelines. Data were analyzed using SPS version 10.0. bSample size too small to calculate meaningful confidence intervals (actual range quoted).
elements (Figs. 29-10 and 29-11). When digital radiography is not an option, special cassettes with slightly slower screens are recommended, but otherwise standard radiographic equipment can be used. Because of the greater detail provided by this film and screen combination, a greater exposure time is required (preferably higher milliamperage). Some experimentation will be required to arrive at the best settings for each individual radiographic machine. A suitable starting point for a sugar glider is 60 kilovolts (kV) at 30 milliamperes (mA) for 0.2 to 0.3 seconds (see also Chapter 35).
DRUG DOSAGES Various antimicrobial agents have been used successfully in captive sugar gliders, despite the absence of pharmacologic studies in this species. In using metabolic scaling, it should be
Fig. 29-10 Digital radiograph of a male sugar glider, lateral view.
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SECTION V Other Small Mammals
Fig. 29-11 Digital radiograph of a male sugar glider, ventrodorsal (VD) view.
remembered that the small body size is offset to some extent by the low metabolic rate of marsupials. Approximate drug dosages can be extrapolated from the low-end ranges for cats, ferrets, and hedgehogs.17 Dosages provided in Table 29-4 have been extrapolated from dosages used in other species and have been used successfully in sugar gliders.12,17,23,24
DISEASES AND SYNDROMES GASTROINTESTINAL DISEASE Malnutrition Because of the misinformation regarding the dietary requirements, sugar gliders in captivity commonly suffer from malnutrition. Common diet-related conditions in captive sugar gliders include hypoproteinemia, hypocalcemia, hypoglycemia, and anemia. Hypocalcemia is primarily caused by an imbalance of dietary calcium, phosphorus, and vitamin D. Lack of dietary protein is a cause of anemia and hypoproteinemia. With chronic malnutrition, hepatic and renal biochemical values are abnormal, as these organ systems become affected. Malnourished sugar gliders are weak, lethargic, and debilitated upon presentation. These animals are usually thin and dehydrated; if they are severely hypocalcemic, seizures and pathologic fractures can develop. Pale mucous membranes, edema, and bruising may be present in anemic and hypoproteinemic patients, and secondary infections are common in debilitated animals. Treatment involves general supportive care and correction of the underlying dietary problems.
Dental Disease Sugar gliders have specialized lower incisors for chewing and gouging into the bark of trees (Fig. 29-12). These animals are not rodents, therefore the incisors do not overgrow because of
malocclusion and should never be trimmed. If the teeth are trimmed or chipped and the pulp is exposed, they become infected and painful, potentially requiring extraction. Periodontal disease and tartar buildup are common in sugar gliders fed soft, carbohydrate-rich diets.1 Tartar can be scaled while the patient is anesthetized. Treat associated gingivitis with systemic antibiotics. Including insects with hard exoskeletons in the diet helps deter tartar buildup. Necrotic teeth should be extracted with care not to damage adjacent tooth roots. Advanced tooth decay or traumatic incisor fracture can lead to exposure of root canals. The root canal is too small for filling. Because of the anatomy of the mandibular incisors, extraction leaves little mandibular bone. The surgical procedure should begin with local anesthetic blocking using 0.02 to 0.05 mL of 2% lidocaine at the level of the exit of the nerves from the mandibular fossa on the lateral side of the mandible approximately at the level of the premolars. The mandibular fossa is palpable. The lidocaine dosage can also be injected at the base of the tooth itself, particularly if it is fractured or abscessed. Gingivectomy using laser, radiosurgery, or a No. 15 scalpel blade is used to reflect the gingiva, incise the periodontal ligaments, and elevate the tooth from the alveolar bone. A number 18to 20-gauge needle can also be used to incise the attachments of the tooth root and elevate the tooth. The alveoli should be debrided and flushed with a 2% chlorhexidine solution, particularly in cases of abscess. To restructure the mandible, synthetic bone matrix (Consil, Nutramax Laboratories, Inc., Edgewood, MD) is applied and the gingiva sutured over the deficit using absorbable monofilament suture with a taper-point swaged-on needle. If both mandibular incisors are removed, the mandible may be shortened, as in the procedure for hemimandibulectomy. Gingivectomy with similar tooth elevation is preferred for any extractions. Care after dental extraction includes analgesia, nonsteroidal anti-inflammatory drugs (NSAIDs), and antibiotics. Some gliders will permit their owners to swab the extraction site daily using a children’s fruit-flavored antiseptic mouthwash on a cotton swab. Soft foods are appropriate for the immediate postoperative period. Proper diet is the best preventive measure for dental disease.
Enteritis and Enteropathy Diarrhea is a common symptom of digestive disease in the sugar glider as in many other species. Such disease can be due to a bacterial enteritis (caused by Escherichia coli, Clostridium species, or other bacteria), intestinal parasites (such as various nematodes, cestodes, and protozoa including Giardia species), metabolic disorder (i.e., hepatic or renal disease), malnutrition, or stress. Standard diagnostics including fecal analysis, bacterial cultures, fecal Gram’s stains, complete blood count (CBC)/ biochemistry profiles, and radiographs are useful in discerning the cause of the digestive disorder. Specific therapy is directed at the underlying cause, along with nutritional support and fluid therapy.
Rectal Prolapse Rectal or cloacal prolapse can occur secondarily to tenesmus and is more common in gliders in generally poor nutritional health. Reduction of the prolapsed tissue is similar to the procedure done in birds because of the structure and function of the cloaca. The prolapsed cloaca should first be irrigated with a hypertonic solution (50% dextrose) and inspected closely for
CHAPTER 29 Sugar Gliders Table 29-4 Drug Dosages for Sugar Gliders12,14,17,24 Agent
Dosage and Administration
Comments
Carbaryl 5% powder
Use topically or in nest box
Fenbendazole
20-50 mg/kg PO q24h x 3 days
Ivermectin Levamisole Metronidazole Oxfendazole
0.2 mg/kg PO/SC, repeat at 14 days 10 mg/kg PO 25 mg/kg PO q12h 5 mg/kg PO once
Use sparingly for ectoparasites Roundworms, hookworms, whipworms, and tapeworms Roundworms, hookworms, whipworms, and mites
Piperazine Pyrethrin powder
100 mg/kg PO Use topically
Antiparasitics
Intestinal protozoa Roundworms and adult tapeworms Ectoparasites
Antimicrobials Amikacin sulfate Amoxicillin Cephalexin Clavamox Enrofloxacin
10 mg/kg IM q12h x 5 days 30 mg/kg PO, IM q24ha 30 mg/kg SC, PO q24h 12.5 mg/kg SC, PO q24h 2.5-5 mg/kg IM, PO q12-24h
Gentamicin Lincomycin Metronidazole Penicillin Sulfadimethoxine Trimethoprim/sulfa
1.5-2.5 mg/kg q12h SC, IM, IV 30 mg/kg IM, PO q24ha 25 mg/kg PO q24ha 22,000-25,000 IU/kg q12-24h 5-10 mg/kg q12-24h 15 mg/kg PO q12h 10-20 mg/kg PO q12-24h
Gram-negative pneumonia Dermatitis (general) Injectable formb Injectable formb Possible tissue necrosis when administered parenterally Nephrotoxicity Also use probiotics Monitor hydration
Antifungals Griseofulvin Itraconazole Nystatin
20 mg/kg q24h PO x 30-60d 5-10 mg/kg PO q12h 5,000 IU/kg q8h x 3 days
Antidermatophyte Antifungal Candida
Acepromazine/ butorphanol
1.7 mg/kg (A)/1.7 mg/kg (B) PO
Acepromazine/ketamine
1 mg/kg (A)/10 mg/kg (K) SC
Buprenorphine Butorphanol Flunixin meglumine
0.01-0.03 mg/kg PO, SC q12h 0.1-0.5 mg/kg SC, IM q6-8h 0.1 mg/kg IM q12h
Meloxicam
0.1-0.2 mg/kg PO, SC q24h
Postoperative analgesia and sedation to prevent selftrauma to incision Postoperative analgesia and sedation to prevent selftrauma to incision Analgesia Analgesia Analgesia and anti-inflammatory NSAID
Analgesics
Preanesthetics Atropine
0.02-0.04 mg/kg IM, SC, IV
Glycopyrrolate
0.01-0.02 mg/kg IM, SC, IV
Control salivation with anesthesia Control salivation with anesthesia
Anesthetics Diazepam Enflurane Isoflurane Ketamine Ketamine/acepromazine
0.5-2.0 mg/kg PO, IM, IV To effect 1%-5% 30-50 mg/kg IM 30 mg/kg (K)/2 mg/kg (A)
Sedative; calming to seizures Anesthetic agent Anesthetic agent of choice Follow with isoflurane Immobilization Continued
403
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SECTION V Other Small Mammals Table 29-4 Drug Dosages for Sugar Gliders12,14,17,24—cont’d Agent
Dosage and Administration
Comments Immobilization
Ketamine/xylazine Midazolam Sevoflurane Tiletamine/zolazepam
2-3 mg/kg (K)/0.05-0.10 mg/kg (M) 10-20 mg/kg (K)/0.35-0.50 mg/kg (M) 10-25 mg/kg (K)/5 mg/kg (X) 0.25-0.50 mg/kg IM, IV 1%-5% 8-12.4 mg/kg IM
Yohimbine
0.2 mg/kg IV
Ketamine given 5-10 min after midazolam Immobilization Sedation, anti-anxiety Anesthetic agent Use with caution: reports of toxicity Reverse xylazine
0.22-0.44 mg/kg q24h PO 1-4 mg/kg q6-8h SC, IM 1-5 mg/kg q12h PO 100 mg/kg q12h PO
Cardiac disease Diuretic Diuretic Cardiac disease
Anesthetics—cont’d Ketamine/medetomidine Ketamine/midazolam
Cardiac Medications Enalapril Furosemide l-Carnitine
Gastrointestinal Medications Cisapride Metoclopramide
0.25 mg/kg q8-24h 0.05-0.10 mg/kg q6-12h
Enhance GI motility Enhance GI motility
50-100 IU/kg SC, IM 0.2 mg/kg q12-24h SC, IM, IV 0.5-2.0 mg/kg IV, IM 0.1-0.2 mg/kg q24h PO, SC, IM
Calcium mobilization Anti-inflammatory Shock Corticosteroid
150 mg/kg PO q24h 7 mg/kg IM 500-5,000 IU/kg IM 0.01-0.02 mL/kg 25-100 IU per glider/day 2 mg/kg q24-72h SC
Calcium deficiency Calcium deficiency Skin disorders Stings on injection; dilute
Steroid/Hormone Therapy Calcitonin Dexamethasone Prednisolone
Vitamin/Mineral Therapy Calcium glubionate Calcium glycerophosphate Vitamin A Vitamin B complex Vitamin E Vitamin K
Adjunctive, cardiac and liver disease
aDaily bNot
dose can be divided. available in the United States.
Fig. 29-12 The lower incisors are adapted for gouging into tree bark. These teeth normally angle forward; thus they are not maloccluded and should never be trimmed.
necrotic tissue as well as to identify the anatomic structures. A sedative (midazolam, 0.35 mg/kg IM) along with an analgesic (butorphanol, 0.4 mg/kg IM) prior to the administration of isoflurane anesthesia should be administered by mask and the prolapsed tissue irrigated with diluted 2% lidocaine (0.05 mL diluted with sterile water to 0.1 mL). Any necrotic tissue should be removed and resected if the necrosis extends beyond the superficial mucosa. The tissue is gently replaced using moistened cotton swabs. After replacement, the abdominal/rectal area should be gently massaged to make sure that the tissue has been replaced into normal anatomic position. One or two vertical sutures using monofilament suture can be placed near the lateral canthi of the vent. Care must be taken not to obstruct the urogenital opening. An antibiotic cream can be placed into the cloaca after the sutures have been placed. Postoperative care includes addressing the etiology for the prolapse as well as continued antibiotics and NSAIDs (meloxicam, 0.2 mg/kg PO q24h; Metacam, Boehringer Ingleheim Vetmedica, Inc., St. Joseph, MO).
CHAPTER 29 Sugar Gliders
Impaction of the Paracloacal Gland The paracloacal glands are similar to anal glands in placental mammals. As such, these glands can similarly become infected or impacted. Expression of the impacted glands produces a thick mucoid to caseated discharge, depending on the degree and type of infection. Bacterial culture and sensitivity may reveal an assortment of bacterial species and determine the best course of antimicrobial therapy. The glands can be surgically removed if they become repeatedly or chronically impacted. The procedure is described under “Surgery and Anesthesia,” below.
RESPIRATORY DISEASE The differential diagnoses in sugar gliders presented with signs of tachypnea or dyspnea include trauma, bacterial pneumonia, cardiac disease, heat stress, and abdominal distention due to various conditions. Radiography under sedation will assist with diagnosis. In addition, standard blood work and bacterial culture of any discharge will help to establish the diagnosis and guide therapy. Treatments may include antibiotics, diuretics, antihistamines, bronchodilators, nebulization, and specific therapy for underlying conditions.
UROGENITAL DISEASE Self-Mutilation of the Penis and Scrotum Self-mutilation is commonly seen in single or stressed males (see later discussion). In animals with severe injuries, amputation of the penis may be necessary. Because males urinate from the base of the penis, amputation of the forked end of the penis will not interfere with urination. The urethral opening must be closely monitored for patency after amputation of the penis, as postoperative swelling may occlude the orifice. Because sexual frustration may be a factor, castration is recommended for a pubescent male that mutilates its penis or scrotum.10,14 A necrotic penis may be associated with systemic sepsis and bacteremia. Parenteral antibiotics and supportive care should be instituted. Blood chemistries and hematology are indicated to assess the extent of the infection.
Cystitis, Crystalluria, and Urolithiasis Hematuria, stranguria, and dysuria are common signs of these urinary disorders in sugar gliders as in other species. Complicating factors that can lead to this presentation include poor nutrition, subclinical dehydration, inactivity, and improper environmental conditions that disrupt normal urine marking behavior. Urinalysis and urine culture and sensitivity help establish the specific urinary disorder and guide the course of therapy. Standard therapy as performed in other species is indicated in these cases, including medical or surgical approaches in addition to correction of underlying factors.
Urinary Tract Obstruction A potential complication to the urinary disorders mentioned above is obstruction of the urinary tract, especially in males. Radiographs or ultrasound can assist in locating the obstruction. It is not anatomically possible to catheterize the sugar glider’s bladder through the urethra; therefore a urethrostomy may be indicated if attempts at relieving the pressure by cystocentesis and flushing of the distal urethra are inadequate. NSAIDs,
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analgesics, antimicrobials, and fluid therapy are also medically indicated.
Nephritis and Renal Failure Renal disease can be evaluated in sugar gliders as in other mammals by biochemical analysis, specifically blood urea nitrogen (BUN), creatinine, phosphorus, and electrolyte levels. Clinical signs include polyuria and polydypsia in addition to nonspecific signs of illness. Prognosis is guarded to poor in many cases, with therapy being limited as in other species.
REPRODUCTIVE DISORDERS Infertility Infertility is common in certain lines of the mosaic color mutations. The male is more likely to be infertile than the female in these mutations. The cause of the infertility is not known but likely genetically linked to one of the genes associated with the color mutation. Failure to breed in female sugar gliders can be related to obesity or inappropriate social situations, as well as underlying medical conditions. In the closely related Leadbeater’s possum, abscesses in both lateral vaginas were observed in an aged female. This animal presented with lethargy and abdominal distention, and the diagnosis was made by radiography, ultrasonography, and exploratory laparotomy.
Failure to Thrive in a Joey Baby sugar gliders are born in a fetal stage and migrate to the dam’s pouch, where they develop further. This is a very delicate stage for the joey, where detachment from the nipple or dislodgement from the pouch is often fatal. Trauma to the pouch or infection in it may contribute to developmental problems. The diet of the dam is critically important for the well-being of her joeys, as is the stress level of her environment.
Pouch Infection and Mastitis Bacteria and yeast have been isolated from infected pouches. These can be infections of the pouch itself or a true mastitis of the dam’s mammary gland. Culture and sensitivity of the exudates from the pouch provides information on effective antimicrobials. Improvement in environmental and cage-mate hygiene can be crucial in the prevention of these infections.1,15
REPRODUCTIVE TRACT INFECTION Because the cloaca is the common opening for the reproductive tract as well as digestive and urinary systems, infection of the reproductive organs can be an extension of disease in these other systems. Ascending infections into the lateral vaginas and uterus from the cloaca may also be a significant cause of poor fertility. Systemic therapy with antimicrobials and general improvements in hygiene and nutrition are necessary to treat reproductive tract infections.
DERMATOLOGIC DISORDERS Stress-Related Disorders Stress in sugar gliders can manifest in numerous ways. Selfmutilation, as described below, is a common manifestation of stress in the sugar glider. Stressed sugar gliders can also pre sent with coprophagy, hyperphagia, polydipsia, and pacing.12
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Fig. 29-13 Self-mutilation of the tail is common in solitary, stressed sugar gliders.
Fig. 29-15 Endocrine alopecia is a recognized but poorly understood syndrome in the sugar glider.
Traumatic Injuries Trauma is a common presentation in captive sugar gliders. Bite wounds from pet dogs, cats, and ferrets are potentially fatal to these little animals, not only because of the physical trauma but also because of the potential infection acquired from the wound. Sugar gliders can also be injured by household accidents, such as falling into toilet bowls, chewing on electric cords, being shut behind a door or window, or being stepped on by family members. Common injuries include cuts, punctures, and fractures.
Ear Margin Canker
Fig. 29-14 A modified E-collar can be used to prevent chewing of a surgical site or deter self-mutilation.
Irregular ear margins with variable proliferation and crusting is commonly encountered in sugar gliders with ear-mite infestations. Differential diagnoses include cage-mate aggression, frostbite, or ergot poisoning. Ear mites can be diagnosed by cytology of swabs from the ear or skin scrapings from the pinna. These can become secondarily infected, leading to otitis externa; therefore bacterial and fungal cultures may be indicated. Appropriate treatment with ivermectin and topical antimicrobials is indicated.
Endocrine Alopecia Some patients may present with alopecia, presumably from increased adrenal activity caused by stress. The provision of proper nutrition and hygiene, normal social grouping, appropriate nesting areas and cage accessories, and protection from potential predatory species can help reduce stress in captive sugar gliders.
Self-Mutilation Self-mutilation of the tail (Fig. 29-13), limbs, scrotum, and penis is common. Common underlying causes of the stress leading to self-mutilation include lack of proper socialization with other gliders, disregard to their nocturnal nature, poor husbandry, and inappropriate nutrition. The therapeutic protocol for these patients includes correction of the underlying causes as well as treatment of the inflicted wounds. Appropriate use of analgesics and antimicrobials is usually indicated. Antidepressants and sedatives may be necessary. Modified Elizabethan collars can assist in preventing further trauma while underlying causes are treated and the lesion heals (Fig. 29-14).
Alopecia in a bilateral, symmetric pattern has been reported primarily in older female gliders (Fig. 29-15). A similar condition is frequently reported in older female short-tailed opossums (Monodelphis domestica) and has been linked to prolactin-secreting pituitary adenomas.7,14 Etiology of the condition in gliders has not been documented.
OPHTHALMIC DISORDERS Ocular Injury Ophthalmic traumas occur frequently in sugar gliders because of their slightly protruding eyes. Corneal scratches with ulceration and conjunctivitis are most common. These may be inflicted by cage mates or by rubbing against cage accessories.
Cataracts Nutritional imbalances and congenital predisposition are both potential causes of cataracts in sugar gliders. Hypovitaminosis A and hyperglycemia can both contribute to cataract formation.
CHAPTER 29 Sugar Gliders Correction of the causative imbalance can halt progression but may not restore vision, depending on the severity of the opacity. A hereditary predisposition is also suspected in captive sugar gliders.14
RETROBULBAR ABSCESS Bite wounds to the eye from competing males can lead to retrobulbar abscesses. The glider will be presented with a large semifluctuant mass extending from the eye to the ear. Radiographs are useful in determining the extent of the abscess and whether there is underlying osteomyelitis from penetration of the orbit or skull. Treatment involves surgical removal of the abscess material, debridement, and in some instances placement of a Penrose drain for irrigation. Systemic treatment involves antibiotics as well as use of an NSAIDs for the pain and inflammation. Separation of males or incompatible gliders and reorganization of the social structure may not eliminate future fights, however, because some gliders seem to have repeat fight wounds and abscesses.
MUSCULOSKELETAL DISEASE Nutritional Osteodystrophy Also known as nutritional secondary hyperparathyroidism or metabolic bone disease, this is a common cause of hind-limb paresis and paralysis in pet sugar gliders. The condition resembles the syndrome seen in calcium-deficient captive reptiles. The importance of full-spectrum light, particularly ultraviolet light, is well established in reptiles; however, sugar gliders are nocturnal and rely more on gut absorption of vitamin D3 than on the conversion of vitamin D2 in the skin from exposure to ultraviolet light.10 The clinical presentation of nutritional osteodystrophy is a sudden onset of hind-limb paresis; spinal trauma and general malnutrition are differential diagnoses. Radiographs may reveal long bone, pelvic, and vertebral osteoporosis. Hypocalcemia and hypoproteinemia are seen on plasma biochemical profiles. The clinical history often indicates a calcium-deficient diet comprised of mostly fruit and minimal protein.1 Patients identified in the early stages may respond to cage rest, parenteral calcium, and vitamin D3 with dietary correction. Calcitonin (50 to 100 IU/kg (Miacalcin, Sandoz Ltd, Hanover, NJ) can be used in more severely affected animals.15 However, calcitonin should be given only when plasma calcium concentrations are within reference ranges. Calcitonin decreases calcium resorption from bone because it inhibits osteoclastic activity; however, because serum calcium and phosphorus concentrations also decrease, calcitonin must be used in conjunction with a calcium supplement. Until more is known about the specific nutritional requirements of sugar gliders, their diets should contain about 1% calcium, 0.5% phosphorus, and 1,500 IU/kg vitamin D3 on a dry weight basis.1,10 Insects should be gut-loaded with calcium before being fed to sugar gliders.
Obesity Captive sugar gliders easily become overweight when fed a diet that is too high in fat or protein. Lack of exercise also contributes to the problem. Obesity may contribute to cardiovascular or hepatic disease, as seen in other species.1 Lipid deposits can form in the eyes of juvenile sugar gliders if the mother is fed a diet too high in fat. These deposits appear as small white spots
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within the eyes and can affect the animal’s sight. Treatment of obesity includes modifying the diet and increasing exercise.
Fractures Because of their small size and propensity to chew injured extremities, severe fractures of the limbs may be most successfully treated with amputation, although internal fixation and external splinting can be performed, as in rodents of equivalent size. The patagium often interferes with placement of splints and bandages. Three-limbed gliders can often locomote well, particularly if cage furnishing adjustments are made. Attention to analgesia and NSAIDs complements any orthopedic procedure.
NEUROLOGIC DISEASE The most common neurologic condition of pet sugar gliders is paresis or paralysis secondary to nutritional spinal osteopathy. If a sugar glider presents with neurologic signs, the differential diagnosis should include trauma and, less commonly, middle ear infection, bacterial meningitis, toxoplasmosis, Baylisascaris procyonis, vitamin E nutritional encephalomalacia, and cryptococcosis. A full diagnostic workup including hematology and serum chemistries and radiographs should be performed. Reflex testing is often difficult even in tame gliders, as they tend to resist limb and joint manipulation in preference to “clinging” to the handler. Lack of proprioception or clutching along with prick sensitivity and toe/tail pinches, balance, and righting response may help the clinician determine if the problem is central or peripheral in nature.
Encephalomalacia and Encephalitis Encephalomalacia has been diagnosed in sugar gliders postmortem. The brain tissue exhibits vacuolation and changes consistent with hypovitaminosis E, although controlled dietary studies have not been performed to prove that this is the complete etiology. Encephalitis due to parasitic infection has been diagnosed postmortem histologically, with identification of the parasite. Baylisascaris procyonis aberrant migration has been found in gliders housed outdoors, which began showing seizures and collapse shortly before death. Mortality was highest in juvenile gliders. Raccoons had been known to climb on the wire mesh roof of the outdoor glider habitat. Toxoplasmosis causes a fatal encephalitis similar to that seen in other species. Antemortem diagnosis may be aided by serology, although specific serologic tests are not validated commercially for sugar gliders. Treatment may be tried in suspected cases using trimethoprim sulfa (15 mg/kg PO q12h) and supportive care. Bacterial and fungal organisms have been implicated in neurologic disease, which is often confirmed postmortem. Cerebrospinal fluid for culture and analysis can be obtained from the cisterna magna, although the volume is usually less than 0.05 mL. The procedure is similar to that done in rodents of comparable size and should be practiced first on a cadaver.
Tremors and Seizures Seizure activity to any degree constitutes an emergency regardless of etiology. The glider should immediately be warmed and given a benzodiazepine anticonvulsant (i.e., diazepam, initially administered IM and subsequently given IV or IO). Isotonic fluids with additional calcium gluconate should be administered, preferably intravascularly or intraosseously, but immediately intraperitoneal or subcutaneous administration may suffice.
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Intrarectal anticonvulsants can be administered at the parenteral dosage if the seizures cannot be controlled and IV or IO administration has not been established. The glider can also be warmed by irrigating the cloaca with warmed saline. Full anesthesia may have to be administered to control the seizure while diagnosis is pursued. Intubation is recommended, as many gliders may vomit in the course of the seizure or anesthesia.
INFECTIOUS DISEASE Bacterial Diseases Infectious diseases are not well documented in captive sugar gliders, but these animals are presumed to be susceptible to the same pathogens as are other marsupials in their family. Sugar gliders have died of infection with Pasteurella multocida, contracted from rabbits kept in the same area.10,14,16 The disease is characterized by generalized abscessation in various organs and subcutaneous areas as well as sudden death. Clostridium piliforme infection has been diagnosed in captive sugar gliders.10,14 Significant pathogens in related possum species include Yersinia pseudotuberculosis, Salmonella species, Mycobacterium species, Cryptococcus neoformans, and Leptospira species.10
Parasitic Diseases Diseases caused by parasites have not been specifically reported in captive sugar gliders. Parasites are most often found in wild sugar gliders or in captive gliders kept outdoors. Various nematodes, trematodes, tapeworms, mites, lice, and fleas are all potential parasites of sugar gliders.1,9,10 Specific parasites identified in sugar gliders include trematodes (Athesmia species) in the liver and nematodes (Parastrongyloides, Paraustrostrongylus, and Paraustroxyuris species) in the gut.1 Giardiasis and cryptosporidiosis have also been diagnosed in captive sugar gliders.10,14 Ectoparasitic mites recovered from sugar gliders include trombiculid (Guntheria kowanyam), astigmatid (Petauralges rackae), and atopomelid mites.1,10 Marsupials are susceptible to Toxoplasma gondii, which causes neurologic signs and sudden death.10 Various anthelmintics have been used in gliders without apparent side effects, but pharmacologic studies have not been conducted. Dosages for several anthelmintics are provided in Table 29-4. Pyrethrin or carbaryl powder can be applied topically and in nest boxes to control mite infestation.1,12,24
NEOPLASIA Neoplasia, in particular lymphoid neoplasia, is relatively common in captive gliders.1 Cutaneous lymphosarcoma has been reported in a sugar glider.7
SURGERY AND ANESTHESIA ANESTHESIA A balanced anesthesia approach—including the use of preemptive analgesics, anxiolytic tranquilizers, and local anesthetics— followed by gas anesthesia results in fewer fluctuations in heart rate and blood pressure as well as a decreased concentration of inhalant gas anesthetic and a smooth, rapid recovery. Maintenance of preoperative body temperature prevents prolonged recovery and the trigger of torpor and decreased overall metabolism associated with the drop in body temperature. Body temperature can be effectively maintained using a forced-air heating
system such as a Bair Hugger (Augustine Medical, Inc, Eden Prairie, MN). Because insertion of an IV catheter for intraoperative fluid and venous access may be difficult, an intraosseous catheter in the femur or tibia can be used instead. For surgeries predicted to last less than 1 hour and/or that do not enter the abdominal cavity, subcutaneous fluids administered as part of the preanesthetic medication regimen is recommended. A successful preanesthetic protocol may include atropine or glycopyrrolate, midazolam, and either butorphanol or buprenorphine, along with the administration of intraosseous or subcutaneous fluid. Table 29-4 lists analgesics, anesthetics, and suggested protocols. Local anesthesia using a 50:50 mixture of lidocaine 2% and bupivicaine 5%—of a volume necessary to infiltrate the incision site, operative area, or achieve a dental nerve block— has the further advantage of providing analgesia to the postoperative site, thus decreasing the possibility that the glider will chew at the incision site. For castration, 0.03 mL of each of the above local anesthetics diluted with sterile water to a volume of 0.1 mL, infiltrated at the base of the stalk on the abdominal wall is done following the first cleansing of the skin with a surgical scrub. Castration can then be performed using a very low concentration of inhalant anesthetic. For example, atropine (0.08 mL SC), midazolam (0.3 mg/kg IM), and buprenorphine (0.03 mg/kg IM) can be administered 30 minutes prior to surgery. Concurrently, 10 to 14 mL of an isotonic crystalloid (such as Normosol) with 1 mL of 50% dextrose warmed to 95°F (i.e., body temperature) is administered subcutaneously. This combination of fluids is slightly hypertonic but provides dextrose to prevent hypoglycemia induced by anesthesia and the resultant drop in metabolism. The glider is then induced with isoflurane via small mask at 5% for 2 to 3 minutes while an electrocardiogram (ERM 3010, Vetronics, Devon, UK) is attached. Isoflurane is then reduced to 2.5% while the skin is shaved and a surgical scrub with a chlorhexidine scrub 3 to 5 times is performed. For castration, the glider is usually not intubated. Alcohol is not used because of its cooling effect. A local block is done after the first skin wipe. The heart rate is maintained in the range of 175 to 200 beats per minute. Isoflurane is then reduced to 1.5% to 2% during the incision of the stalk and ligature of the two cords. Isoflurane is reduced to 1% after the testicles and scrotal skin are removed. The ligated stalks are incorporated into a suture in the abdominal fascia to prevent herniation. Orchiectomy and scrotal ablation may also be performed using a carbon dioxide laser.18 Isoflurane and sevoflurane are the anesthetics of choice for sugar gliders. The use of either of these agents facilitates physical examination, venipuncture, and radiographic examination as well as anesthesia for general surgery. Anesthesia can be induced with the use of a large face mask as an induction chamber to deliver 5% isoflurane. Transient apnea may occur during induction; therefore monitor the heart rate closely. If the heart rate is stable, gentle pressure on the thorax usually stimulates breathing. Once anesthesia is induced, a small face mask or a 1-mm Cook endotracheal tube (Global Veterinary Products, New Buffalo, MI; www.globalvetproducts.com) can be used to deliver isoflurane at 2% to 3% for maintenance. Intubation of gliders is difficult, but it can be aided by using a stylet and finebladed laryngoscope.1 While extending the glider’s head to lift the soft palate maximally, extend its tongue and use the laryngoscope to see the larynx. A mixture of tiletamine hydrochloride and zolazepam hydrochloride (Telazol, Fort Dodge Animal Health, Fort Dodge, IA)
CHAPTER 29 Sugar Gliders
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reportedly caused neurologic signs and death in apparently healthy sugar gliders when administered at 10 mg/kg.6 Other reports suggest that Telazol can be used in sugar gliders at 8.4 to 12.8 mg/kg without problems.2 Therefore use this drug combination with caution in sugar gliders. However, there is seldom an indication for the use of any injectable anesthetics in this species.
SOFT TISSUE SURGERY General Surgical Considerations Sugar gliders should be fasted for at least 4 hours before surgery.23 For gastrointestinal surgery, a longer fasting period may be indicated. Administer subcutaneous fluids before surgery or place an intraosseous catheter to administer fluids during the operation. Maintain the patient’s body temperature during surgery with the use of a circulating hot-water pad or forced-warmair unit. Magnification with a binocular head loupe during surgery aids in identifying small structures and vessels. Minimize blood loss during surgery by using a radiosurgery unit or carbon dioxide laser and ligate small vessels with fine nonreactive monofilament suture, such as 5-0 polydioxanone (PDS). Sugar gliders commonly chew at skin sutures; therefore, close the skin with subcuticular sutures and apply tissue glue. Sugar gliders typically recover smoothly from anesthesia. Administer analgesics after surgery to minimize self-trauma to the incision and provide postoperative pain relief. Butorphanol (0.5 mg/kg IM) can be administered up to three times daily or buprenorphine at 0.01-0.03 mg/kg IM q12h for postoperative pain.14,23,24 Recommended dosages for other analgesic choices are listed in Table 29-4.12,14,24 To maintain body temperature, allow the patient to recover from anesthesia in an incubator. Restricted activity and usual postoperative procedures are recommended as in other species.
Castration and Scrotal Ablation Following preanesthesia medication—administration of atropine, midazolam, and either butorphanol or buprenorphine— induce the glider via isoflurane or sevoflurane using a mask. For castration, the glider is usually not intubated. Clip the skin and surgically prepare around the base of the scrotal sac and stalk. Do not use alcohol because of its cooling effect. A local block is done after the first skin wipe. Maintain the heart rate in the range of 175 to 200 beats per minute. Isoflurane or sevoflurane can then be reduced to 1.5% to 2% during the incision of the stalk and the blunt dissection of the two cords (vas deferens, blood vessels). Several methods have been described for exposure of the cords, including a longitudinal incision by scalpel, radiosurgery electrode, or surgical laser made close to the body wall in the stalk skin. Clamp the cords with hemostats and ligate, using 5-0 PDS suture or surgical laser, or cut and cauterize using radiosurgery. Then remove the scrotal sac containing the testicles and the distal portion of the stalk. The ligated stalks can be incorporated with a suture in the abdominal fascia to prevent herniation (Fig. 29-16).15,18 Close the skin incision using tissue glue.
Ovariohysterectomy Induce and maintain general anesthesia as described previously. Clip and surgically prepare the area around the pouch on the ventral abdomen. Make a 1- to 2-cm skin incision paramedial to the pouch, leaving enough skin along the edge of the pouch for closure.15,23 Identify the linea alba by blunt dissection, then
Fig. 29-16 Castration is performed by ligating the spermatic cord and vessels with complete scrotal ablation.
incise into the abdominal cavity. The bladder is visible as the abdominal cavity is entered in this area. Carefully exteriorize the bladder to bring the reproductive tract into view. The ovaries appear as small red granular structures. Identify and ligate the ovarian branch of the ovarian artery. Ligate and remove the uterus above the lateral vaginal canals. Close the linea alba; then close the skin routinely with subcuticular sutures and tissue glue.
Repair of the Patagium Small wounds (less than 5 mm in diameter) to the patagium may be left to heal by secondary intention, but they should be thoroughly clipped, debrided, and cleaned. Take particular care to remove hair from the wound edges with a scalpel blade. Larger wounds may require suturing, and care must be taken to align the two skin layers correctly. Magnification will assist in this process. Fine (4-0 or 5-0) absorbable suture material with a swaged-on suture is recommended. If the gliding membrane is extensively damaged, it may have to be reshaped to remove any skin tags or flaps that could become snagged on a wire enclosure. To discourage self-trauma, sutures must not be tight. Sugar gliders can reach almost all parts of their bodies, even with a fitted Elizabethan collar, so comfortable sutures are important. Some patagial repairs will result in contraction of the gliding membrane and mobility may be reduced.10
Removal of the Paracloacal Glands Surgical removal of the paracloacal glands is similar to removal of the anal gland in the ferret. A single skin incision over the center of each firm swollen gland will reveal the gland and allow the skin to be peeled away from the thin connective tissue and glandular wall (Fig. 29-17). Care should be taken to prevent rupturing the glands, especially if they are infected and impacted. The bulk of the procedure is performed with blunt dissection of the gland from the surrounding connective tissue around the cloaca. A single ligation of 5-0 or 6-0 PDS, or using radiosurgery or carbon dioxide laser at the base, may be necessary in closing the small blood vessel to the gland. Close the incision with a single subcuticular 5-0 PDS suture and a small drop of tissue glue on the surface (Fig. 29-18). Inject a small amount of diluted lidocaine at the surgery site to reduce the risk of postoperative self-trauma.
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Fig. 29-17 Dissection of the impacted paracloacal gland from the surrounding connective tissue reveals a small stalk-like attachment for ligation and excision.
Fig. 29-18 Closure of the incision after paracloacal gland removal surgery is similar to that following anal gland removal in the ferret.
References 1. Booth RJ. General husbandry and medical care of sugar gliders. In: Bonagura JD, ed. Kirk’s current veterinary therapy XIII. Philadelphia: WB Saunders; 2000:1157-1163. 2. Bush MJ, Graves AM, O’Brien SJ, et al. Dissociative anesthesia in free-ranging male koalas and selected marsupials in captivity. Aust Vet J. 1990;67:449-451. 3. Dierenfeld ES. Feeding behavior and nutrition of the sugar glider (Petaurus breviceps). Vet Clin North Am Exot Anim Pract. 2009;12:209-215. 4. Fleming MR. Thermoregulation and torpor in the sugar glider Petaurus breviceps (Marsupilia: Petauridae). Aust J Zool. 1980;28:521. 5. Henry SR, Suckling GC. A review of the ecology of the sugar glider. In: Smith PA, Hume ID, eds. Possums and gliders. Sydney: Australian Mammal Society; 1984:355-358. 6. Holz P. Immobilization of marsupials with tiletamine and zolazepam. J Zoo Wildl Med. 1992;23:426-428. 7. Hough I, Reuter RE, Rahaley RS, et al. Cutaneous lymphosarcoma in a sugar glider. Aust Vet J. 1992;69:93-94. 8. Hubbard GB, Mahaney MC, Gleiser CA, et al. Spontaneous pathology of the gray short-tailed opossum (Monodelphis domestica). Lab Anim Sci. 1997;47:19-26. 9. International Species Information System (ISIS), 12101 Johnny Cake Road, Apple Valley, MN, 2002. 10. Johnson R, Hemsley S. Gliders and possums. In: Vogelnest L, Woods R, eds. Medicine of Australian mammals. Collingwood Vic: CSIRO Publishing; 2008:395-438. 11. Johnson-Delaney CA. Common procedures in hedgehogs, prairie dogs, exotic rodents, and companion marsupials. Vet Clin North Am Exot Anim Pract. 2006;9:415-435. 12. Johnson-Delaney CA. Exotic companion medicine handbook for veterinarians (Supplement). Lake Worth: Wingers Publishing. 1997. 13. Johnson-Delaney CA. Feeding sugar gliders. Exot DVM. 1998;1:4.
14. Johnson-Delaney CA. Marsupials. In: Meredith A, JohnsonDelaney C, eds. BSAVA manual of exotic pets. 5th ed. Quedegley: BSAVA; 2010:103-126. 15. Johnson-Delaney CA. Reproductive medicine of companion marsupials. Vet Clin North Am Exot Anim Pract. 2002;5:537-553. 16. Johnson-Delaney CA. The marsupial pet: sugar gliders, exotic possums, and wallabies. Proceedings. Assoc Avian Vets. 1998:329-339. 17. Johnson-Delaney CA. Therapeutics of companion marsupials. Vet Clin North Am Exot Anim Pract. 2000;3:173-181. 18. Morges MA, Grant KR, MacPhail CM, et al. A novel technique for orchiectomy and scrotal ablation in the sugar glider (Petaurus breviceps). J Zoo Wildl Med. 2009;40:204-206. 19. Nagy KA, Suckling GC. Field energetics and water balance of sugar gliders, Petaurus breviceps (Marsupialia: Petauridae). Aust J Zool. 1985;33:683. 20. Ness RD. Introduction to sugar gliders. Proceedings. North Am Vet Conf. 1998:864-865. 21. Ness RD. Clinical pathology and sample collection of exotic small mammals. Vet Clin North Am Exot Anim Pract. 1999;2:591-619. 22. Ness RD. Sugar glider (Petaurus breviceps): general husbandry and medicine. Proceedings. Exot Small Mam Med Suppl, Assoc Avian Vet. 2000:99-107. 23. Pye GW, Carpenter JW. A guide to medicine and surgery in sugar gliders. Vet Med. 1999;94:891-905. 24. Pye GW. Sugar gliders. In: Carpenter JW, ed. Exotic animal formulary. St. Louis: Elsevier Saunders; 2005:347-358. 25. Smith MJ. The reproductive system and paracloacal glands of Petaurus breviceps and Gymnobelideus leadbeateri (Marsupialia: Petauridae). In: Smith PA, Hume ID, eds. Possums and gliders. Sydney: Australian Mammal Society; 1984:321-330. 26. Suckling GC. Sugar glider. In: Strahan R, ed. The mammals of Australia. Chatswood: Reed Books; 1995:229-231. 27. Tyndale-Biscoe H, Renfree M. Reproductive physiology of marsupials. Cambridge: Cambridge University Press; 1987; 18, 19, 22, 59, 123.
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African Hedgehogs
Evelyn Ivey, DVM, Diplomate ABVP (Avian), and James W. Carpenter, MS, DVM, Diplomate ACZM
Biology and Anatomy Taxonomy and Natural History Anatomy and Physiology Husbandry Housing Diet Breeding and Neonatal Care Basic Procedures and Preventative Medicine Restraint and Examination Clinical Techniques Preventative Medicine Common Diseases Ocular Otic Oral and Dental Respiratory Cardiovascular and Hematologic Gastrointestinal and Hepatic Urinary Reproductive Musculoskeletal Neurologic Integumentary Neoplastic Nutritional The Lethargic, Weak, Anorectic Hedgehog Anesthetic and Surgical Considerations Zoonoses and Suitability as Pets Because of their small size, spiny coat, and relatively recent appearance in the pet trade, African hedgehogs can be challenging patients. Hedgehogs are illegal in some states and municipalities; in other states, a permit is required. Additionally, a permit from the U.S. Department of Agriculture (USDA) is required for persons who breed, transport, sell, exhibit, or use hedgehogs for research or teaching purposes. Even breeders who give away their progeny must be registered with the USDA, although breeders with three or fewer intact females are now Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
excluded from this requirement. In 1991, it became illegal to import hedgehogs from Africa into the United States because of the potential transmission of foot and mouth disease to cattle.59 Hedgehog owners are advised to check with appropriate regulatory agencies, including state fish and game or wildlife departments and their regional USDA office for specific regulations.
BIOLOGY AND ANATOMY TAXONOMY AND NATURAL HISTORY Hedgehogs (Fig. 30-1) are members of the family Erinaceidae within the order Insectivora. The two most familiar hedgehog species are the central African hedgehog (Atelerix albiventris) and the European hedgehog (Erinaceus europaeus). The central African hedgehog, also known as the white-bellied, four-toed, or “African pygmy” hedgehog, is native to the savannah and steppe regions of central and eastern Africa.58 Some pet African pygmy hedgehogs are descended from hybrids of the central African and similar Algerian hedgehog (Atelerix algirus). In the pet trade, Atelerix species are generally referred to as African pygmy hedgehogs. Unless otherwise noted, the information in this chapter refers to pet African pygmy hedgehogs. Because of the ban on importation, pet hedgehogs in the United States are captive-bred; several color varieties exist and at least one show standard has emerged. Wild African hedgehogs are found in diverse dry open habitats including grassland, scrub, and suburban yards.45 They are nocturnal and spend their daylight hours hidden in burrows or other cavities. At night, they are very active invertebrate predators, jogging several miles in search of insects, earthworms, slugs, and snails.58 Males and females are territorial and solitary except during courtship and when females raise young. Although all hedgehog species appear to be capable of entering a torpid state under cold conditions, the central African hedgehog probably does not normally experience cold temperatures in the wild, and torpor is considered to be undesirable in captives.35,58 Excessively high temperatures can also induce a torpid state.58
ANATOMY AND PHYSIOLOGY Radiographic anatomy of the normal hedgehog is presented in Figures 30-2 and 30-3.9 411
412
SECTION V Other Small Mammals
Musculoskeletal
Alimentary
Hedgehogs are adept at climbing, digging, swimming, and constant jogging. The tibia and fibula are fused distally. The stance is plantigrade. The manus has five digits, the pes has four. The normal hedgehog gait is a slow and steady waddle, but they are capable of bursts of speed.
Hedgehogs have brachydont (closed-rooted) teeth. The first incisor in each quadrant is large and projects forward. The mandibular first incisors occlude into a space between the maxillary first incisors, an arrangement that is suited to spearing insects. The molar teeth are relatively flattened with multiple cusps and are suited for crushing food items. The stomach is simple, and a vomiting reflex is present. There is no cecum, and the colon is smooth. Feces are normally soft but formed.
Integument The crown and dorsum (collectively called the mantel) are covered in a dense coat of several thousand smooth spines. Hair and sebaceous glands are absent in the spiny skin.58 The epidermis in this area is thin, and there is a thick fibrous dermal layer that contains much fat and few blood vessels. Hedgehog spines are composed of keratin and have a complex internal structure that confers lightness, strength, and elasticity. Each spine has a round basal bulb that firmly attaches it within the follicle, while a more narrowed portion at the skin surface allows each spine to bend when force is applied. Healthy spines are difficult to pull from the follicle without breaking at this narrowed portion. Spines may last up to 18 months and are replaced
Fig. 30-1 Captive African hedgehog with young.
A Scapula Spiny coat Ilium Caudal vertebrae Axis (C2) Atlas (C1) C3-C7 T1-T13 L1-L7 Sacrum
Skull Trachea Maxilla Humerus Mandible Radius Carpal bones Manus
Ribs
Colon
Lungs Heart
Femus
B Ulna
Olecranon
Sternebrae
Pubis Ischium Fibula Tibia Calcaneus Metatarsal bones Phalanges
Stomach Patella Liver Small intestine
Fig. 30-2 Normal, whole body radiograph (lateral view) of a female African hedgehog; A, unlabeled; B, labeled anatomic structures.9 (Reprinted with permission of John Wiley & Sons, Inc.)
CHAPTER 30 African Hedgehogs individually. Spines are absent from the midline of the crown, and hair on the feet and muzzle is sparse to absent. The haired skin and the soles of the feet are rich in sweat and sebaceous glands. The toenails are round in cross section and are highly curved. A wary hedgehog will raise the spines and crouch. If an alert hedgehog is frightened by touch or noise, contraction of the panniculus muscle pulls the loose spiny skin over the entire body (Fig. 30-4). Several muscles assist in pulling the panniculus over the body and tucking in the legs and rump. The panniculus is thickened at the rim to form the orbicularis muscle, a purse-string-like muscle that closes the loose skin over the animal. A hedgehog may remain rolled up for hours with relatively little muscular effort.
Reproductive An unrolled hedgehog’s gender is easily determined; the male has a conspicuous prepuce that opens on the midabdomen, whereas the female’s urogenital opening is a few millimeters cranial to the anus. The penis has a prominent glans that protrudes laterally into horn-like structures.4 There is no scrotal sac; the testes are located in a para-anal recess,4 are surrounded by fat, and can be palpated in reproductively active males. Male accessory glands include paired prostate glands, seminal vesicles, bulbourethral glands, and Cowper’s-like glands.58 The uterus is bicornate with no uterine body but rather a continuous lumen over the cervix. There is a single cervix and a long vagina that is always patent. A fan-shaped gland, homologous to the Cowper’s-like gland of the male, lies on each side of the
413
vagina. The urethral opening is located in the distal vagina, several millimeters from the vulva. The mesosalpinx and ovarian bursa are fat-laden, and the fallopian tubes are relatively short and emerge from the side of each uterine horn.4 Females have three pairs of mammary glands.45 African hedgehogs are polyestrous and breed throughout the year in captivity. The estrus cycle for A. albiventris empirically seems to be similar to that of other hedgehog species, for which the estrus cycle has been reported to be 3 to 17 days of estrus followed by 1 to 5 days of diestrus.58 There is some indirect evidence to suggest that ovulation is induced.4,58 Sterile matings with pseudopregnancy may occur. Although the gestation period is 34 to 37 days, it is possible that delayed implantation may occur and extend the apparent gestation period to 40 days.58 Hedgehogs are born hairless, with closed eyes and ears. At birth, the spines are buried within a layer of skin; within the first few hours, the spines are exposed by an inflating and subsequent deflating of the skin with fluid and removal of the covering by the mother. These first spines are white, but they are replaced by darker spines over the first 3 days of life. Colostral transfer is believed to occur early in lactation.
Neurology and Behavior Hedgehogs have a very sensitive olfactory system and well- developed olfactory lobes. The vomeronasal organ is also prominent, and hedgehogs may occasionally exhibit a flehmen response.29 It is believed that olfaction plays a key role in a wild hedgehog’s ability to navigate within its home range, detect food, avoid predators, and communicate with conspecifics.29 Mandible
Maxilla
Skull
Atlas (C1)
Clavicle
Axis (C2)
Scapula Heart
C3-C7 Ribs
Humerus Manus
Lungs
Radius Ulna
T1-T13 Liver
Diaphragm
Small intestine
Olecranon Stomach Spiny coat Colon
L1-L7
Ilium
Sacrum
Patella
Caudal vertebrae
Femur Fibula Tibia Tarsal bones Metatarsal bones
A
B
Phalanges Ischium
Pubis
Fig. 30-3 Normal whole-body radiographs (ventrodorsal view) of a female African hedgehog. A, unlabeled. B, Labeled and showing the ischiopubic symphysis in this nonpregnant animal with no history of parturition.9 (Reprinted with permission of John Wiley & Sons, Inc.)
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SECTION V Other Small Mammals
Fig. 30-4 If the animal is frightened, contraction of the panniculus muscle pulls the loose spiny skin over the entire body. The panniculus is thickened at the rim to form the orbicularis muscle, a purse-string-like muscle that closes the loose skin over the animal (i.e., permits it to roll up into a ball).
Hedgehogs also have sensitive hearing, especially in the ultrasonic range. Their sense of vision is not as well developed and is essentially monochromatic.58 Foraging hedgehogs normally emit a variety of snuffling sounds. Frightened or agitated hedgehogs make a distinctive hissing sound that may be punctuated with various puffing and cough-like sounds. With intense distress, and occasionally impatience for food, a scream may be emitted.29 Other vocalizations include grunts, squeals, clucking, snuffling, and high-pitched sounds, but these are uncommon except during courtship and between mothers and offspring. The cerebrum of the hedgehog is relatively small, and their learning abilities seem to be less than those of rodents or carnivores.58 They do demonstrate the ability to recognize their owners and with patient training may learn simple commands. With patient, gentle handling, most hedgehogs will learn to accept and even enjoy human contact. Both genders of wild and captive hedgehogs demonstrate a unique behavior called self-anointing, or anting (Fig. 30-5). This behavior may be elicited by a variety of substances, particularly those with a strong or unusual odor, such as fish, wool, and various plants and vegetables. The hedgehog takes the material or object into its mouth, mixes it with frothy saliva, and applies the mixture to its spines with its tongue. Many speculations have been offered as to the purpose of this behavior, the most plausible of which seems to be one of imparting an individual odor to the animal and its home range.58
HUSBANDRY HOUSING Hedgehogs are solitary in the wild; in captivity they are usually maintained in individual cages. Some fanciers successfully keep groups of females with or without a single male or even groups of males, but this can lead to disproportionate feeding and injuries from fighting. Although young animals raised together may tolerate each other as adults, hedgehogs typically become aggressive toward cage mates when they reach sexual maturity. Should multiple animals be housed together, at least one hide box should be provided for each animal. Like-colored individuals may be
Fig. 30-5 Self-anointing or anting behavior in a hedgehog. This behavior is elicited by a variety of substances; the hedgehog takes the material or object into the mouth, mixes it with frothy saliva, and applies the mixture to its spines with its tongue. (Courtesy of Lyndsey Smith, Manhattan, KS.)
identified with either a microchip implanted in the subcutaneous fat pad or nontoxic marker or fabric paint applied to the dorsal spines. These methods of identification have the advantage of being readable even when the animal is curled in a tight ball.24 Healthy hedgehogs are very active, and as large a cage as possible should be provided; 2 × 3 feet (0.6 × 0.9 m) are minimal floor dimensions. Hedgehogs are able to climb and escape through small holes, so the cage must be secure and lidded; smooth walls are ideal. Glass tanks are suitable but are heavy when they are sufficiently large. Hedgehog droppings can be soft and messy, making wood cages difficult to keep clean. Plasticbottomed cages with wire walls are suitable provided that the wire spacing is appropriate (1-in. square is acceptable). Widely spaced wires can lead to limb entrapment or death if the hedgehog puts its head between the wires and becomes ensnared by its spines. A hiding place—which may be a cardboard or wooden box, flowerpot, or polyvinyl chloride (PVC) tube—is an essential furnishing. The cage substrate should be soft and absorbent and will require frequent changing. Recycled newspaper bedding is a good choice; aspen shavings, alfalfa pellets, and hay are other options. Wire, cedar, corncob, and dusty or scented substrates are not recommended. Pine bedding remains a somewhat controversial choice; it is used by many fanciers without apparent ill effects but is discouraged by many practitioners as being too aromatic. The substrate should be 3 to 4 in. deep to allow for digging; hiding mealworms or other treat items in the bedding is a good way to stimulate foraging behavior. Cloth in the cage poses a risk, as hedgehog digits and limbs are easily entrapped by loose fibers. Hedgehogs should be maintained at ambient temperatures between 72°F and 90°F (22°C and 32°C); 75°F to 85°F (24°C to 29°C) is optimal. African hedgehogs may go into torpor if they are too cool or too warm. A heating pad placed under part of the enclosure or a ceramic reptile heater may be used. Low humidity (less than 40%) is preferred. Hedgehogs avoid bright light; however, a day cycle of 10 to 14 hours of mild light should be provided. Although some owners may attempt to convert their pets to a diurnal schedule, most hedgehogs will retain the nocturnal lifestyle of their wild counterparts. Strategies include reverse lighting schedules and feeding during the day.
CHAPTER 30 African Hedgehogs Hedgehog droppings are relatively soft and, depending on diet, can be quite messy. Although some hedgehogs use a litter tray, others deposit their droppings at random. Placing all of the droppings in the litter tray on a daily basis may facilitate litter training. Other litter training tips include providing a cardboard enclosure over the tray, placing the animal in the tray after feeding, placing another hedgehog’s feces in the tray, and placing the tray where the animal seems most inclined to eliminate. Natural plant litters used for cats make the best litter substrate. Clay, clumping-type litter, or sand may stick to the animal and should not be used. Because many hedgehogs defecate in their hide boxes and exercise wheels and subsequently walk in their feces, daily spot cleaning of the cage is often necessary to prevent dermatitis and possible dissemination of Salmonella species. Exercise wheels are highly recommended. The wheel must have a solid or fine plastic mesh for hedgehogs to run on because their legs tend to become entrapped by traditional wire rodent wheels. Hedgehogs should also be let out into a large area on a daily basis for exercise. Cardboard tubes, straw, safe climbing structures, swimming tubs, and other toys provide enrichment. Dirty hedgehogs may be bathed with a mild pet shampoo and a soft-bristle vegetable brush.
DIET Wild African hedgehogs feed on a diversity of invertebrate prey as well as plant materials and occasional eggs, vertebrate prey, and carrion.58 Although many of the natural food items are known, the nutritional content of invertebrates varies tremendously; this makes it very difficult to deduce nutrient requirements based on the wild diet. Insectivorous mammals are traditionally fed diets that are 30% to 50% protein and 10% to 20% fat (dry-matter basis).1 Hedgehogs seem to require a higher level of dietary fiber than carnivores; this may be related to the large quantity of insect exoskeletons in their natural diet. The base of the captive diet should consist of a commercially prepared hedgehog food. Scientific studies regarding hedgehog nutritional needs are lacking; however, commercial diets appear to be the most balanced staple that a pet owner can offer. If commercially prepared hedgehog food is not used, premium food for less active cats or dogs should form the basis of the diet. Ferret food is high in fat and is not recommended. With many individuals, food must be rationed to prevent obesity. Depending on the animal’s weight and activity, 3 to 4 teaspoons of the main diet is typically fed daily. Growing animals and reproductively active females may be fed the usual diet ad libitum, and calcium-rich foods are recommended. In addition to the main diet, approximately 1 to 2 teaspoons of varied moist foods and/or invertebrate prey (e.g., canned cat or dog food, cooked meat or egg, or low-fat cottage cheese; mealworms, earthworms, waxworms, gut-loaded crickets) and about 1 teaspoon of vegetable/fruit mix (e.g., beans, cooked carrots, squash, peas, tomatoes, leafy greens; bananas, grapes, apples, pears, berries) should also be provided daily. Invertebrate prey and other dry food items may be hidden in the bedding to promote foraging activity. Hedgehogs should not be fed raw meat or eggs, which may harbor Salmonella species. Milk, although relished by many hedgehogs, can cause diarrhea. Nuts, seeds, and large items or hard foods such as raw carrots can become lodged in the roof of the mouth and should be avoided. The need for vitamin or mineral supplementation, if any, is not known, but supplementation does not appear to be necessary
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for animals fed a commercial diet. Moist or perishable foods should be offered in the evenings. Hedgehogs may be slow to accept novel foods, and any dietary changes must be made with care. Fresh water should be available at all times. Most hedgehogs can learn to drink from water bottles.
BREEDING AND NEONATAL CARE Although pet hedgehogs may become sexually mature at 2 months, females should be at least 6 months of age before breeding. Pregnancy is most easily determined by weighing the female every few days; a gain of 50 g or more within 2 to 3 weeks of being placed with a male is suggestive of pregnancy.63 At 30 days, a general swelling of the abdomen or mammary enlargement may be detected. Infanticide, usually followed by cannibalism of the young, can occur. Novice hedgehog breeders should give the female strict privacy from other hedgehogs and humans from about 5 days before delivery through 5 to 14 days after delivery. Females conditioned to frequent handling are less likely to desert or kill their young in response to human contact.63 Providing a large cage and an additional hiding place for the female as a refuge from her litter may reduce stress.29 Male hedgehogs must not be allowed near the neonates because cannibalism (by either parent) often results. Normal pups (or “hoglets”) stay close to their dam and littermates when resting. In cases of lactation failure or abandonment by the female, fostering of the pups to another dam with similarly aged pups is usually successful. Weaning generally occurs at 5 to 6 weeks, and the young may be moved to separate cages at 8 weeks.29 Daily handling starting at 3 weeks of age will produce hedgehogs that remain tame. If a surrogate dam is unavailable, a milk replacer may be fed through a dropper, feeding tube, or narrow-tipped syringe.64 Based on the composition of European hedgehog milk, a canine milk replacer with added lactase (Lactaid, McNeil-PPC, Ft. Washington, PA) seems to be the most logical formula.34,58,64 Hand-rearing of hedgehogs is often associated with high mortality. Neonates should be fed as much as they will consume every 2 to 4 hours for about 3 weeks. The ambient temperature should be maintained at 90°F to 95°F (32°C to 35°C) for the first few weeks.63 Neonates should gain approximately 1 to 2 g/day during the first week, 3 to 4 g/day during the second week, 4 to 5 g/day during the third and fourth weeks, and 7 to 9 g/day until they are 60 days old.58,63 Neonates should be stimulated to eliminate after each feeding by massaging the ventrum and perineal area with a cloth or swab moistened in warm water. At 4 to 6 weeks, parent- or hand-raised young should be weaned by offering canned dog or cat food, minced beef, or freshly molted mealworms. A slight weight loss may occur during weaning.63
BASIC PROCEDURES AND PREVENTATIVE MEDICINE RESTRAINT AND EXAMINATION Even very tame hedgehogs often roll up when an examination or other procedure is attempted. Hedgehogs do bite, but infrequently. Patience and a quiet room with subdued light may help calm wary hedgehogs. High-pitched sounds, such as the jingling of instruments, should be avoided. A small towel and light gloves may facilitate handing. Some hedgehogs may be induced to uncurl voluntarily if supported in normal standing position
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SECTION V Other Small Mammals Table 30-1 Biologic and Physiologic Data for African Pygmy Hedgehogs10,34,43,58,63,64 Average body weight
Fig. 30-6 Holding a rolled-up hedgehog face-downward over a table may induce extension of the legs as the animal seeks to reach the surface to avoid falling.
and gently rocked up and down in a see-saw fashion. When the snout pokes out, place a thumb firmly on the back of the neck to prevent the head from tucking in. Press the thumb of the other hand into the back so that it also uncurls.58 Other techniques include holding the animal face-downward over a table (Fig. 30-6), placing it on its back, or pushing it toward the edge of a table; these maneuvers may induce extension of the legs as the animal seeks, respectively, to reach the surface, right itself, or avoid falling. The hind legs are then grasped, and the hedgehog is held in a face-down position. Some hedgehogs uncurl if their rump spines are stroked slowly in a circular or backward motion for several minutes; the animal is then pinned against the table by grasping the spiny dorsal skin. Once uncurled, hedgehogs may be restrained by holding the spined skin as one would hold the scruff of other species. A superficial examination may be performed as the animal moves about within a transparent container, but a thorough examination usually requires chemical restraint. Healthy hedgehogs should be active and inquisitive or curled up in a tight ball. Hydration may be assessed by eyelid turgor. Body temperature is lower than that of most mammals (Table 30-1). The eyes should be clear, and the pinnal margins should be free of crusting or ragged edges. The teeth should be white and the gingiva a uniform pink. Inspect the oral cavity and tongue for ulcers, foreign material, and masses. The nose is normally moist and active. Normal lymph nodes are difficult to palpate, but lymph nodes may become enlarged in cases of neoplasia or infection.26,55 The heart should have a regular rhythm and no murmurs, and a femoral pulse should be palpable. Respiration is normally silent except in the defensive or aggressive animal, in which forceful expulsion of air through the nose creates a loud hissing sound. The abdominal contour as the animal rests in the hand of the clinician should be flat. Palpate the abdomen for organomegaly, masses, and fluid. Check the prepuce or vulva for inflammation, discharge, or adherent debris. Testicles may be palpable in the para-anal area. Normal stools are very dark brown, and the consistency varies from very soft to pellet-like. Healthy, untroubled hedgehogs walk with the ventrum raised clear off the table, but weak or wary hedgehogs tend to crouch. Inspect the toes for encircling fibers and overgrown nails. The skin in the spiny areas may have a mildly dry or flaky appearance, but excessive flaking, quill loss, erythema, and crusting are abnormal.
Captive: male 400-600 g, female 300-400 g Life span Average 4-6 years,a may live to 8 years Body temperature 95.7°F-98.6°F (35.4°C-37.0°C) Adult dental formula 2 (I3/2:C1/1:P3/2:M3/3) = 36; variations have been noted Gastrointestinal transit time 12-16 hours Heart rate 180-280 beats per minutea Respiratory rate 25-50 breaths per minutea Age at sexual maturity 2-3 months Reproductive life span Female, 2-3 years; male, throughout life Gestation 34-37 days Milk composition Protein, 16 g/100 g; carbohydrate, trace; fat, 25.5 g/100 g Litter size 3-4 (range, 1-9) Birth weight 10-18 g Eyes open 14-18 days Deciduous teeth eruption Begins on day 18; all deciduous teeth erupted by 9 weeks Permanent teeth eruption Begins at 7-9 weeks Age at weaning 5-6 weeks (start eating solids at 3 weeks) aPersonal
observation (EI).
CLINICAL TECHNIQUES The jugular vein is usually used to collect blood samples. Although visualization of the vein may be difficult, its anatomic location is similar to that in other small mammals. Alternatively, the cranial vena cava may be used. The technique for caval venipuncture is similar to that used in ferrets, but there is a greater risk of cardiac puncture because of the relatively cranial position of the hedgehog heart. The femoral, lateral saphenous (Fig. 30-7), or cephalic veins may be used for injections or to collect small samples (up to 0.5 mL). Reference ranges for hematologic and serum biochemical values are presented in Table 30-2. A urine sample may be collected by cystocentesis or catheterization; however, reference values are not available.5,64 Subcutaneous injections can be given in the spiny or furred areas; the furred area is more elastic and vascular, but it is less accessible in a balled hedgehog. The dermis under the spiny skin is poorly vascularized, so drugs or fluid given in this location may not be absorbed for several hours. For subcutaneous fluid administration, the junction of furred and spined skin midbody provides an accessible and reliable site.31 Intramuscular injections may be given in the triceps, quadriceps, gluteal, or obicularis muscles.31 Maintaining an intravenous catheter in all but extremely weak hedgehogs is difficult, as the catheter tends to become dislodged if the animal curls. A femoral or tibial intraosseous catheter may be placed as a substitute for vascular access.5,35 A 22- or 25-gauge needle or 1-in. spinal needle is used and secured with tape; a tibial crest catheter remains accessible even when the animal is in a curled position.36 For long-term administration of subcutaneous fluids or medication, a 5-Fr polyvinylchloride (PVC) feeding tube may
CHAPTER 30 African Hedgehogs
417
Table 30-2 Hematologic and Serum Biochemical Reference Values of African Hedgehogs10 Measurement
Reference Rangea
Hematologic Values
Fig. 30-7 Venipuncture of the hedgehog can be challenging. The lateral saphenous vein is a site that may be used for collecting small samples.
be implanted under the mantel.49 The tube is inserted through an incision in the spineless area in the dorsal cervical region, into a 5- to 6-cm tunnel made with a periosteal elevator caudal to the incision. The tube is fenestrated along the distal 3 cm prior to insertion and secured with nylon suture using a fingertrap pattern. A Luer-lock injection plug provides the injection port and remains accessible even when the patient curls into a ball. Oral medication may be difficult to impossible to administer to some animals. Some animals accept pleasant-tasting medications via a syringe; grape juice and other fruit flavors have been recommended.24 Many hedgehogs will consume mealworms that have been injected with medication; for those that do not, medications may be mixed with a favorite food. Applying topical medications is complicated by the presence of spines and self-grooming, although bandages can be used to protect the extremities.39 Some odors may initiate anting behavior. Radiographic detail is greatly diminished by the presence of the spines. Anesthesia usually is required for proper positioning unless the patient is too weak to roll up. Extend the legs and body for both the lateral and ventrodorsal views. For the lateral view, the elasticity of the dorsal skin makes it possible to pull the spines away from most of the chest and abdomen; a plastic kitchen clip (as is used to seal potato chip bags) is useful. Other imaging techniques—including CT (Fig. 30-8), MRI, and echocardiography—may also be useful diagnostic procedures in the hedgehog. When weak or debilitated hedgehogs are hospitalized, cage temperatures of 80°F to 85°F (27°C to 29°C) should be provided. Hedgehogs may shun unfamiliar food items, and having the owner bring food the animal is accustomed to may enhance voluntary feeding in the hospital. Providing live invertebrates may also stimulate appetite. Anorectic animals should be fed a protein-rich, high-calorie canine or feline diet via syringe or tube.64 For ongoing assisted feeding, a pharyngostomy or esophagostomy tube may be placed, although this can be difficult in obese patients.36,61
PREVENTATIVE MEDICINE Because hedgehogs have a somewhat compressed life span and often hide signs of illness, semiannual examinations with blood testing are recommended. Hedgehogs are prone to a number of
Hematocrit (%) Red blood cell count (x106/μL) Hemoglobin (g/dL) Mean corpuscular volume (fL) Mean corpuscular hemoglobin (pg) Mean corpuscular hemoglobin concentration (g/dL) Platelets (x103/μL) White blood cell count (x103/μL) Neutrophils (x103/μL) Lymphocytes (x103/μL) Monocytes (x103/μL) Eosinophils (x103/μL) Basophils (x103/μL)
36 ± 7 (22-64) 6 ± 2 (3-16) 12.0 ± 2.8 (7.0-21.1) 67 ± 9 (41-94) 22 ± 4 (11-31) 34 ± 5 (17-48) 226 ± 108 (60-347) 11 ± 6 (3-43) 5.1 ± 5.2 (0.6-37.4) 4.0 ± 2.2 (0.9-13.1) 0.3 ± 0.3 (0-1.6) 1.2 ± 0.9 (0-5.1) 0.4 ± 0.3 (0-1.5)
Biochemical Values Alanine aminotransferase (IU/L) Alkaline phosphatase (IU/L) Amylase (IU/L) Aspartate aminotransferase (IU/L) Bilirubin, total (mg/dL) Blood urea nitrogen (mg/dL) Calcium (mg/dL) Chloride (mEq/L) Cholesterol (mg/dL) Creatine kinase (IU/L) Creatinine (mg/dL) Gamma-glutamyl transferase (IU/L) Glucose (mg/dL) Lactate dehydrogenase (IU/L) Phosphorus (mg/dL) Potassium (mEq/L) Protein, total (g/dL) Albumin (g/dL) Globulin (g/dL) Sodium (mEq/L) Triglycerides (mg/dL) aMean
53 ± 24 (16-134) 51 ± 21 (8-92) 510 ± 170 (244-858) 34 ± 22 (8-137) 0.3 ± 0.3 (0-1.3) 27 ± 9 (13-54) 8.8 ± 1.4 (5.2-11.3) 109 ± 10 (92-128) 131 ± 25 (86-189) 863 ± 413 (333-1964) 0.4 ± 0.2 (0-0.8) 4 ± 1 (0-12) 89 ± 30 441 ± 258 (57-820) 5.3 ± 1.9 (2.4-12.0) 4.9 ± 1.0 (3.2-7.2) 5.8 ± 0.7 (4.0-7.7) 2.9 ± 0.4 (1.8-4.2) 2.7 ± 0.5 (1.6-3.9) 141 ± 9 (120-165) 38 ± 22 (10-96)
± SD (range in parentheses).
dental disorders, including periodontal disease. A palatable pet enzymatic dentifrice may be placed on a crunchy treat daily to help prevent this. Tartar-control treats made for dogs and cats may also help prevent the accumulation of calculus. Some aged individuals may demonstrate excessive wearing of the teeth; these animals should be fed soft foods. An accurate scale to monitor weight allows early detection of obesity or weight loss. Owners should be advised of the risks associated with cold temperatures, which include metabolic derangement and immunocompromise. Nails must be trimmed to prevent ingrowth; trimming the nails as the animal climbs a small screen is one way to attempt nail trimming without anesthesia.61 Currently, there are no vaccines for pet hedgehogs. Because hedgehogs are not typically maintained in mixed-gender groups, elective castration or ovariohysterectomy is not usually needed to prevent unwanted offspring.
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SECTION V Other Small Mammals
COMMON DISEASES OCULAR Hedgehogs are prone to corneal ulcers and other ocular injuries. Diagnosis and treatment is as for other species, although treatment may be difficult if owners are unable to administer topical medication. Blind hedgehogs seem to navigate their captive environments with minimal detriment to their quality of life.
Ocular proptosis was a relatively common presenting complaint in one report.70 The ocular sequelae to proptosis were severe and resulted in enucleation or euthanasia in all eight cases. Moderate to marked orbital inflammation was present in each case; sinusitis, neoplasia, and fungal elements were not observed. Two of the animals had concurrent neurologic disease, which may have resulted in ocular trauma; no known trauma occurred in the other cases. Hedgehogs have a shallow orbit that may predispose them to proptosis, especially if excessive fat accumulation or orbital inflammation is present. In hedgehogs with a unilateral proptosis, tarsorrhaphy may be indicated as a prophylactic measure for the remaining eye.70
OTIC Pinnal dermatitis is common in pet hedgehogs; skin crusts, accumulated secretions, and a ragged pinnal margin may be observed. Dermatophytes and acariasis are important causes; other possibilities include nutritional deficiencies, dry skin, nonspecific seborrhea with hyperkeratosis, and extension of ear canal disease.26 Ear mites (Notoedres cati) are occasionally seen.58 Signs include accumulation of waxy otic debris and pruritus. Diagnosis is confirmed by visualizing the mites in the ear or in swabbed material and they may be treated as for cats. Bacterial or yeast otitis externa also occurs; these infections are often secondary to acariasis or another cause of chronic inflammation. Signs include purulent discharge, odor, and sensitivity of the face and ear. Otic cytology, skin scrapings, cleansing, and topical antimicrobial/anti-inflammatory therapy are employed as for dogs. Otitis media/interna has also been described.7
ORAL AND DENTAL Fig. 30-8 Computed tomography (CT) imaging may occasionally be of diagnostic value in hedgehogs.
A
Oral neoplasms, particularly squamous cell carcinomas (Fig. 30-9), are relatively common. Dental disease is also common; calculus, gingivitis, and periodontitis may occur. On
B Fig. 30-9 A, Oral mass in a hedgehog (arrow). B, Open-mouth rostrocaudal skull radiograph of the hedgehog in (A) showing severe osteolysis of the left mandible associated with soft tissue swelling. Histopathologic findings revealed the mass to be a squamous cell carcinoma.
CHAPTER 30 African Hedgehogs examination, reddened and swollen gingiva, calculus, gingival recession, and loose teeth may be present. Signs include decreased appetite, ptyalism, halitosis, and pawing at the mouth. Dental radiographs, extractions, cleaning, and antibiotic administration should be performed as in other species. In cases in which advanced periodontal disease requires extraction of all the teeth, hedgehogs can be maintained on soft food.64 Tooth fractures and dental abscesses also occur. A mandibular abscess with extensive osteomyelitis and cellulitis due to Actinomyces species infection has been reported.40 It is not known if this abscess was secondary to periodontal disease or some other cause, nor is it known how prevalent Actinomyces infection is in hedgehogs. Based on this report and the prevalence of Actinomyces in the dental abscesses of other species, anaerobic culture and treatment should be considered for dental abscesses in hedgehogs. Excessive tooth wear may occur with advancing age, and animals with this condition should be fed a predominantly moist diet. Hedgehog teeth are not continuously growing and should not be trimmed or filed. Hedgehogs are susceptible to wedging of hard items (e.g., peanuts) against the palate; signs are similar to those seen with dental disease. Stomatitis may occur in males that bite their mates during copulation; treatment is with soft food and antibiotics.35
RESPIRATORY A case of fatal corynebacterial bronchopneumonia has been reported in a pet African hedgehog.57 Other bacteria such as Bordetella bronchiseptica and Pasteurella multocida can cause respiratory infections in European hedgehogs and are possibly important in Atelerix as well.64 Predisposing factors for upper and lower respiratory tract infection include suboptimal environmental temperature; aromatic, dusty or unsanitary bedding; malnutrition; concurrent disease; and other causes of immunocompromise. Aspiration of infected material from an oral infection is another potential cause.40 Signs include nasal discharge, increased respiratory noise, dyspnea, lethargy, inappetence, and sudden death. As in other species, diagnostic testing includes radiographs, hematologic testing, and culture of tracheal or lung lobe aspirates. Treatment includes antibiotics, nebulization, supportive care, and correction of husbandry problems. Additional differential diagnoses for dyspnea include pulmonary neoplasia and cardiac disease. Lungworms can also cause pneumonia, but this is unlikely in indoor pets.26,64 The existence of cytomegalovirus in African hedgehogs has been questioned and is in any case highly unlikely in domestically raised pets.6
CARDIOVASCULAR AND HEMATOLOGIC Dilated cardiomyopathy is common in pet hedgehogs, and necropsy findings from several cases have been described.52 Affected hedgehogs are typically 3 years of age or older, although the disease may occur in animals as young as 1 year of age. Signs include dyspnea, decreased activity, weight loss, heart murmur, ascites, and acute death. Radiographs typically demonstrate varying degrees of cardiac enlargement, pulmonary edema, pleural effusion, hepatic congestion, and abdominal fluid. Reported histologic lesions of the myocardium have included fibrosis, edema, degeneration, and necrosis, with myofibrillar atrophy, hypertrophy, and disarray.52 Histologic lesions may be confined to either ventricle (more commonly the left), but in
419
some cases both ventricles are affected. Pulmonary and renal infarcts may occur in some affected animals. The etiology of cardiomyopathy in hedgehogs is not known, but there may be a genetic or dietary component. If cardiac disease is suspected, obtain full-body radiographs and an echocardiogram. Normal echocardiographic measurements have not been published, but a subjective evaluation of wall motion and chamber size is often sufficient to confirm a diagnosis of cardiomyopathy. In our experience with a small number of healthy hedgehogs, fractional shortening should be at least 25%, and wall thickness should be at least 1.5 mm. A complete blood count and biochemical profile are useful to screen for concurrent problems and serve as a reference for monitoring the effects of therapeutic agents. Therapy with digoxin, furosemide, and enalapril may be helpful initially, but the long-term prognosis for hedgehogs with congestive heart failure is poor. Myocardial mineralization or calcinosis has been described as a necropsy finding in pet hedgehogs with concurrent diseases.2,7 The clinical significance of this lesion is not known. Splenic extramedullary hematopoiesis, the clinical significance of which is not known, has been noted at necropsy in several hedgehogs with diverse diseases.52,55 Saddle thrombus and pulmonary fibrin thromboemboli was described in a hedgehog with a very large uterine polyp.2 Congenital erythropoietic porphyria was reported in a 6-month-old male inbred pet African hedgehog.71 The animal exhibited pink-colored urine and teeth, hepatomegaly, and, under ultraviolet light, fluorescence of the urine, teeth, feet, and spines. The animal behaved normally despite high levels of porphyrins in the urine and feces; it is not known whether the animal was anemic or how long it lived.
GASTROINTESTINAL AND HEPATIC Enteritis may be caused by Salmonella species or other bacteria. Salmonellosis in hedgehogs may be subclinical or cause diarrhea, weight loss, decreased appetite, dehydration, lethargy, and death. It must be emphasized that Salmonella species have been identified in pet African hedgehogs as well as European hedgehogs.11,12 Diagnosis should be confirmed with fecal culture, using a Salmonella-enriching medium. Although treatment is indicated in animals with clinical signs of disease, owners should be advised of the zoonotic potential and the risks of creating antibiotic resistance. Alimentary candidiasis (Candida albicans) was suspected based on results of fecal cytology and culture in a hedgehog that presented with weight loss, depression, and blood in the stool.8 Fatal cryptosporidiosis of the ileum, jejunum, and colon was reported in a juvenile hedgehog.20 Although numerous species of nematodes, cestodes, and protozoa have been identified in wild hedgehogs, their significance in pets appears to be minimal. Nevertheless, fecal examination by ultracentrifugation and wet mount has been recommended in newly acquired animals and in those with signs of enteritis.5 Pyloric and intestinal obstructions can occur and are most often caused by rubber, hair, or carpet fibers. Signs include acute anorexia, lethargy, and collapse. Vomiting may be present, but very often is not. Presurgical diagnosis of acute gastrointestinal obstruction may be complicated by the fact that marked gaseous dilation of the gastrointestinal tract can be a nonspecific finding in ill hedgehogs. A fatal 720-degree intestinal mesenteric torsion has also been reported.14 Alimentary inflammation—including
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SECTION V Other Small Mammals
esophagitis, gastritis, enteritis, colitis, and gastric ulceration with perforation—has also been seen in African hedgehogs.7,15 Most of these animals had nonspecific signs such as decreased appetite and weight loss; gastrointestinal signs such as vomiting and diarrhea were not observed. Diarrhea can also be associated with some commercial diets or inappropriate foods such as milk. Changing the brand of pellets and eliminating inappropriate food items may resolve the condition. Gastrointestinal neoplasia, particularly lymphosarcoma, is relatively common.53,54 Other differential diagnoses for gastrointestinal signs include dietary change, toxins, hepatic disease, and malnutrition. Hedgehogs do not rely on bacterial fermentation for digestion, and there is no evidence of antibiotic sensitivity, as is seen in herbivorous mammals.26 Hepatic lipidosis is relatively common in hedgehogs. In one survey of common necropsy findings in 14 animals, hepatic lipidosis was found in 50% of the animals.55 Hepatic lipidosis may be a sequela of cardiomyopthy, neoplasia, nutritional imbalances, starvation, obesity, toxicosis, pregnancy, and infectious disease. Signs include lethargy, inappetence, and icterus, particularly in the axillary region. Diarrhea or signs of hepatic encephalopathy may also be present. Treatment for hepatic lipidosis in hedgehogs is similar to that in other species. Other important causes of liver failure include primary and metastatic hepatic neoplasia.38 Multifocal hepatic necrosis caused by human herpes simplex virus 1 was reported in a hedgehog that had been treated with dexamethasone for intervertebral disk disease.3 This animal was found dead after a single day of anorexia; the dexamethasone had been administered 2 weeks before death (the dose and duration of treatment were not specified). Transmission from the hedgehog’s owners was suspected in this case.
URINARY Cystitis and urolithiasis have been seen in pet hedgehogs.28,30 Signs may include changes in urine color, stranguria, pollakiuria, inappetence, and lethargy. Urinalysis with culture and radiographs should be obtained. Renal disease is also common and in many cases may be secondary to systemic disease. Nephritis, tubular necrosis, nephrocalcinosis, glomerulosclerosis, infarcts, polycystic kidneys, neoplasia, and various glomerulonephropathies have been identified histologically.7,14,18,55 As in other species, signs associated with renal diseases tend to be nonspecific, although polyuria or polydipsia may be noted. Treatment consists of correcting the underlying cause if possible, fluid therapy, and supportive care.
REPRODUCTIVE Posthitis caused by substrate entrapment in the prepuce is common. Hemorrhagic vulvar discharge is often caused by uterine neoplasia35 or endometrial polyps.2,48 Pyometra and metritis have been reported.14 Dystocia also occurs and is treated as in other small mammals.63 One report describes a case of a perianal fetal hernia in a primigravid hedgehog; three viable offspring were delivered via cesarean section, and a nonviable herniated fetus was removed from an incision over the perianal area.68 Premature births occasionally occur, and the prognosis for young without a sucking reflex is poor.63 Agalactia may be suspected if neonates lose condition within 72 hours after birth. Diagnosis may be confirmed by attempting to express the
mammary glands63; however, this procedure usually requires anesthesia and may cause the dam to abandon or cannibalize her young. Possible causes of agalactia include malnutrition of the dam, stress, lack of oxytocin, inadequate mammary development in young females, and mastitis. Supportive care for weak neonates includes warming to normal body temperature over 1 to 3 hours, fluid support, and caloric support once normothermia has been achieved.63
MUSCULOSKELETAL Myositis secondary to cellulitis has been reported.14 Osteoarthritis and intervertebral disk disease have also been observed.3,14 Fractures most frequently occur when a limb becomes entrapped in a rodent wheel or wire cage. Splinting may be performed for distal limb fractures and requires anesthesia.62 Surgical correction may also be performed, but any fixation device must be able to withstand the strong rolling-up mechanism. Lameness may be caused by ingrown toenails, arthritis, nutritional deficiencies, pododermatitis, constriction of a foot or digit by fibrous foreign material, neurologic disease, or neoplasia.
NEUROLOGIC Common causes of neurologic disease with ataxia include torpor, demyelination (“wobbly hedgehog syndrome,” or WHS), intervertebral disk disease, trauma, toxins, infarcts, malnutrition, and neoplasia. Hedgehogs experiencing cold or sometimes excessively high temperatures may enter a state of torpor or dormancy. In this state, the animal remains sensitive to touch, but the response is greatly diminished. Heart and respiratory rates are decreased, and these animals may have increased susceptibility to infection. Dormancy can last for several weeks, during which the animal may have periods of activity with ataxia.29 As in other species, head tilt or circling may be caused by otitis media/interna or primary neurologic disease. Hypocalcemia may occur in cases of postpartum eclampsia, with malnutrition, or for unknown reasons and usually responds to calcium supplementation.39 Neurologic signs resulting from rabies, Baylisascaris species, and polioencephalomyelitis are also possible.39 Intervertebral disk disease (IVD) has been reported.3,14,55,56 In one case series of four hedgehogs, affected animals were 4 years of age and older and both genders were affected.56 Signs included chronic progressive ataxia, lameness, proprioceptive deficits, and bladder atony with urine retention. Radiographic findings included spondylosis, disk-space narrowing, and disk mineralization. In all four animals, multiple disks were affected; necropsy findings in all cases included degeneration of the nucleus pulposus and annulus fibrosus, dorsal extrusion of disk material, and mineralization of the nucleus pulposus. Cervical lesions were present in 3 of 4 cases, while the fourth case demonstrated lesions in the lumbar vertebrae. One case had evidence of fibrocartilaginous embolism; in another case, spinal nerve roots were affected, resulting in forelimb lameness. Temporary improvement with corticosteroids has been described in two cases of hedgehog IVD.3,56 Since the mid-1990s, an increasing number of cases of WHS (also known as demyelinating paralysis) have been reported in captive African hedgehogs, a condition that had been previously described in European hedgehogs.15,21,22,35,39 This progressive paralysis occurs in approximately 10% of hedgehogs in North America.22 One of the earliest indications of WHS is the
CHAPTER 30 African Hedgehogs
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Fig. 30-10 Hedgehog in the final stage of wobbly hedgehog syndrome (WHS). This animal suffered from the progressive paralysis associated with WHS over several months and demonstrated clinical signs typical of the final stages of the disease, including tetraplegia, muscle atrophy, and emaciation.22 (Reprinted with permission of Elsevier.)
inability to “close the hood” (ball up). Hedgehogs often present with mild ataxia and lack of coordination, becoming off balance and stumbling, tripping, or wobbling. In the early stages of WHS, clinical signs are usually relapsing and remitting.22 Over several months, the signs become progressively more severe and may include falling to one side, tremors, exophthalmos, scoliosis, seizures, muscle atrophy, and self-mutilation (Fig. 30-10).22 The paralysis ascends from hind limbs to forelimbs in about 70% of the cases. The disease ultimately leads to complete paralysis. The onset of WHS commonly occurs in animals under 2 years of age but can occur at any age. In contrast, IVD disease typically occurs in older animals. In 60% of cases in one study, the hedgehog was immobilized within 9 months after the onset of ataxia.22 In 90% of cases, immobility occurred by 15 months after the onset of ataxia. The progression of WHS is generally accompanied by severe weight loss. However, in most cases there is no apparent loss of appetite until the terminal stages of the disease, when most hedgehogs become dysphagic.22 Death usually occurs within 18 to 25 months after the onset of signs. The diagnosis of WHS can be determined only postmortem. Gross lesions are not evident, but histopathologic lesions reveal a vacuolization of the white matter of the cerebrum, cerebellum, brainstem, and spinal cord as well as and associated neurogenic muscle atrophy.22 Other histological lesions include axonal swelling, degeneration of spinal cord ventral tracts, and the degeneration of axons and myelin in brain white matter.21 Peripheral nerves may also be involved. Inflammation of the central nervous system is not associated with WHS. To date there has been no histologic evidence of a viral or autoimmune cause. The etiology of this syndrome is unknown, but pedigree analyses from numerous animals suggests a familial tendency to develop the disease. Although no evidence of an infectious agent has been observed, the familial tendency of WHS does not rule out a transmissible component.22 Numerous treatments for this idiopathic neurodegenerative disease have been attempted, with little or no success; none have stopped the progression of the paralysis. Although supportive care and handfeeding can initially be performed, euthanasia is recommended when the hedgehog becomes significantly immobilized and its quality of life is compromised. Differential diagnosis is important, however, because brain tumors, intervertebral disk disease,
Fig. 30-11 Severe pruritus, erythema, and dermatitis in a hedgehog with acariasis.
and hepatic encephalopathy are other causes of progressive paralysis.22
INTEGUMENTARY Acariasis is very common. Caparinia tripilis is probably the most common mite infesting pet hedgehogs, whereas Caparinia erinacei may be more common in wild African hedgehogs.58,66 Chorioptes species have also been implicated in pet hedgehog acariasis, although it is probable that mites in some of these cases were misidentified as Caparinia, which they closely resemble. Notoedres mites may also infest pet hedgehogs.30,58 Some hedgehogs have subclinical infestations, which may account for the high prevalence of mites in the pet population. Infested bedding or fomites from pet stores may be another source. Signs include hyperkeratosis, seborrhea, quill loss, and white or brownish crusts (mite droppings) at the base of the quills and around the eyes. Hedgehogs can scratch themselves (Fig. 30-11) with their hind limbs or rub against stationary objects, but many individuals do not demonstrate obvious signs of pruritus. Acariasis also causes nonspecific signs such as lethargy and decreased appetite. Diagnosis is confirmed by a skin scraping. Treatment consists of ivermectin (0.3-0.4 mg/kg PO, SC q10-14d × 3-5 treatments). In some cases, however, ivermectin alone may not be effective in treating mite infestations. Therefore some have recommended a combination of ivermectin with amitraz (0.3% topically q7d × 2-3 treatments).67 All bedding must be removed and cage furnishings disinfected or discarded. During the treatment period, the cage is lined with paper that is changed daily. All hedgehogs in the home should be treated concurrently. Ear mites (Notoedres cati) are occasionally seen and can cause crusting lesions around the head and ears.41,58 Signs include accumulation of waxy otic debris and otic pruritus. Diagnosis is confirmed by identifying mites in the ear canal or in swabbed material. Ear mites in hedgehogs may be treated as in cats. Pet hedgehogs may be infested with fleas; shampoo and powder products that are safe for kittens appear to be safe for hedgehogs. An infestation of tropical rat mites (Ornithonyssus bacoti) was reported on a pet African hedgehog with flaky skin and loss of spines.37 These mites are black and 1 to 2 mm long; they resemble the snake mite (Ophionyssus natricus). Treatment with fipronil spray (one spray on the dorsum, repeated in 10 days)
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SECTION V Other Small Mammals Pinnal dermatitis occurs when skin secretions accumulate along the margins of the pinnae, causing a ragged-edged or crusty appearance. Dermatophytes are an important cause; other possibilities include localized acariasis, nutritional deficiencies, dry skin, and nonspecific seborrhea with hyperkeratosis.26 Cutaneous and subcutaneous nodules may be seen. Differential diagnoses include papillomas, abscesses, mycobacteriosis, Cuterebra larvae, and neoplasia.26,55,62 Pruritus may occur with the development of new spines, as is seen in young hedgehogs.
NEOPLASTIC
Fig. 30-12 Pruritus and dermatitis resulting from a yeast infection, suspected to be a result of an allergy, in a hedgehog.
was an effective treatment, while selamectin dosed topically at 16 mg/kg was ineffective. Ticks and fly larvae are uncommon in indoor hedgehogs, and are removed individually. Fungal organisms most often identified as infecting hedgehog skin include Trichophyton erinacei, Trichophyton mentagrophytes, and Microsporum species. These dermatophytes can cause a crusting, usually nonpruritic dermatitis, especially around the face and pinnae. Yeast, probably a secondary problem, may also be identified in cases of dermatitis (Fig. 30-12). Some infections are secondary to other dermatopathies, such as acariasis or trauma.64 Diagnosis is confirmed by culturing spines from the affected areas in dermatophyte testing media. Treatment consists of topical antifungal agents, with systemic griseofulvin (30-40 PO mg/kg q24h × 30 days)44 or ketoconazole if needed. Lyme sulfur dips may also be used. Other hedgehogs in the home may be subclinically infected, and treatment of all animals may be indicated.5 Several cases of Trichophyton infections in humans have originated from contact with infected hedgehogs. Contact dermatitis may result from unsanitary bedding.5 Cellulitis has been linked to secondary myositis and sepsis; the primary cause in most of these cases was found to be trauma.14 Skin neoplasia is common in middle-aged and older hedgehogs.16 Papilloma, squamous cell carcinoma, lymphosarcoma, and sebaceous gland carcinoma have been described. Osteosarcoma has also been reported in the skin of a hedgehog47 (Fig. 30-13). Papillomas of suspected viral etiology have been reported, although histologic findings were not described; recurrence at other sites after excision is common.5,62 Suspected allergic dermatitis has been anecdotally described. Cases of facial, axillary, and groin dermatitis with pruritus and histologic evidence of allergy of unknown source have been reported.16,39 Restricted-antigen diets, antihistamines, and glucocorticoids may be helpful in such cases.16 A single case of pemphigus foliaceus has been reported.69 Signs included generalized loss of spines; dry, flaking skin; moist erythema on the legs, anus, ears, and chin; and epidermal collarettes on the ventral abdomen and limbs. Dexamethasone injections were successful in resolving the condition; initially injections were given twice daily, but over 16 months this frequency was gradually reduced to once every 10 days.
Neoplasms in African hedgehogs are common, and a wide variety of tumors and disseminated neoplastic processes affecting virtually every body system have been reported. Several reviews are available detailing the specifics of numerous reported cases;19,23-25 typical signalment, clinical signs, and tumor behavior, and appropriate diagnostics and treatment options are outlined. In one retrospective study of 74 necropsies, neoplasms were diagnosed in 24 animals (32%); tumors were present in 40% of animals between 1 and 36 months and in 69% of animals over 3 years of age.15 In a survey of 97 hedgehogs submitted for histopathology (including ante- and postmortem samples), 50 animals (52%) had neoplastic disease.54 The body systems in which the tumors were found were integumentary (18 tumors), hemolymphatic (12 tumors), alimentary (9 tumors), endocrine (8 tumors), genital (6 tumors), musculoskeletal (2 tumors), and nervous system (1 tumor). The most commonly diagnosed tumors in this survey were mammary gland tumor (9), lymphosarcoma (8), and oral squamous cell carcinoma (7). Four (8%) of the 50 animals had more than one type of tumor. The occurrence of tumors was unrelated to gender, and the median age at time of tumor diagnosis was 3.5 years (range 2-5.5 years). More than 80% of the tumors in this survey were malignant, and they usually carried a poor prognosis. In another report, 28 proliferative uterine lesions were diagnosed in fifteen 3- to 5-year-old hedgehogs.42 Lesions were associated with vaginal bleeding in all cases, hematuria in 11 of 13 cases, and weight loss in 7 of 12 cases. Lesions were multiple in 8 cases. Lesions identified were 13 adenosarcomas, 7 endometrial stromal sarcomas, 6 endometrial polyps, 1 adenoleiomyosarcoma, and 1 adenoleiomyoma.42 Mean survival time was 303 days (n = 10). Ovariohysterectomy allows prolonged survival of hedgehogs with uterine tumors.42 In retrospective studies of hedgehogs at necropsy, the prevalence of neoplasia has ranged from 29% to 52%.25 Numerous neoplasms have been reported in the hedgehog; they include squamous cell carcinomas (especially of the skin, maxillary, and oronasal areas); cutaneous mast cell tumors; mammary gland tumors51; cutaneous hemangiosarcoma; an acinic cell carcinoma; alimentary lymphosarcomas and plasmacytoma; a pancreatic exocrine carcinoma; endocrine tumors (including a pancreatic islet cell tumor, thyroid tumors, adrenocortical carcinomas,33 a pituitary adenoma, and a parathyroid tumor); gastric adenocarcinoma; myelogenous leukemia; osteosarcomas (including a spinal osteosarcoma, and a rib osteoblastic osteosarcoma); fibrosarcomas; cutaneous histiocytic sarcoma; malignant fibrous histiocytoma; nerve cell tumors (peripheral nerve sheath tumor, astrocytoma, neurofibroma, neurofibrosarcoma, and schwannoma); and a wide variety of tumors of the reproductive system (including uterine leiomyoma, leiomyosarcoma,
CHAPTER 30 African Hedgehogs
A
B
C
D
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Fig. 30-13 A, Anesthetizing a hedgehog with isoflurane administered via a nose cone for evaluating a mass located on its posterior dorsum. B, Performing a fine-needle aspirate of the mass. C, Patient is intubated and, with clippers, site is prepared for surgical excision of the tumor (histologically classified as an osteosarcoma). D, Postsurgical appearance of site.47 (Reprinted with permission of Elsevier.) uterine spindle cell tumor, and adenocarcinoma; and a mammary adenocarcinoma [Fig. 30-14] and a granulosa cell tumor). Certain types of sarcomas in hedgehogs have been associated with retroviral infection.46 In addition to the presence of masses, clinical signs of neoplasia may include weight loss, anorexia, lethargy, diarrhea, dyspnea, ascites, and neurologic signs. Cases of oral squamous cell carcinoma usually present as swelling of the maxillary or mandibular gingiva. Hedgehogs with oral squamous cell carcinomas generally present with loose teeth, swollen gingivae, and/or gingivitis. This tumor generally appears to be locally infiltrative. Diagnosis of neoplasia is based on biopsy (needle, incisional, or excisional) or necropsy and histopathology. Radiographs for evidence of invasion or metastasis, a CBC and serum chemistry panel, and abdominal ultrasound may be useful in determining long-term prognosis. Treatment generally includes surgical excision and supportive care, although different treatment
modalities may be helpful. Prognosis and management depend on type and stage of the neoplastic process. Because African pygmy hedgehogs appear particularly prone to neoplasia, semiannual examinations of animals more than 2 years old should be performed and should include an appropriate workup for neoplasia.24 Despite the high incidence of neoplasia in this species, it should be emphasized that not every mass in pet hedgehogs is neoplastic. For example, dental abscesses, bone cysts, papillomas, and uterine polyps have been described.7,40,48
NUTRITIONAL Obesity in captive hedgehogs is common. Healthy hedgehogs should be able to roll up completely, without any fat deposits protruding. Obese hedgehogs may have large axillary fat deposits or a decreased ability to roll up (Fig. 30-15). Treatment
SECTION V Other Small Mammals
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A
B Fig. 30-14 A, Lateral radiographic view of an adult female with a mammary adenocarcinoma. B, Excised mammary tumor. Gross postexcisional view of the mammary adenocarcinoma.25
without respiratory signs. Hedgehogs with advanced neoplasia, hepatic disease, or renal failure may have nonspecific signs at presentation. Multisystemic diseases should also be considered as causes of lethargy, weakness, and inappetence. For example, disseminated histoplasmosis was reported in a pet hedgehog; fungal organisms were recovered from the spleen, liver, kidneys, intestines, and lungs.65 Other causes include suboptimal environmental temperature, inanition due to inappropriate or novel foods, dental problems, and cage-mate competition. The frequent presentation of hedgehogs emphasizes the importance of diagnostic testing, even if anesthesia is required.
ANESTHETIC AND SURGICAL CONSIDERATIONS Fig. 30-15 Obesity in a hedgehog. This hedgehog is so obese that it cannot roll up completely.
includes eliminating high-fat foods, rationing the main diet, and increasing exercise. Weight reduction should be gradual to prevent hepatic lipidosis, and owners should monitor their pet’s weight with an accurate scale. Hedgehogs, particularly those whose diet consists mainly of cultured invertebrates, may have health problems associated with calcium deficiency.1 Also, vitamin excess or deficiency may occur with unbalanced diets.64 Other nutritional diseases are uncommon.
THE LETHARGIC, WEAK, ANORECTIC HEDGEHOG Nonspecific signs of lethargy, weakness, and anorexia frequently are the only historical and physical findings, even in hedgehogs with severe system-specific disease. Examples include advanced meningeal lymphoma without neurologic signs, debilitating acariasis without evident pruritus, severe gastric ulceration without gastrointestinal signs, and extensive pneumonia
Anesthesia is often required for thorough physical examination, restraint for imaging and other diagnostic testing, as well as for surgical procedures. A 4- to 6-hour fast is recommended prior to anesthetic procedures lasting longer than 20 minutes.24 As with other species, thermal and fluid support should be maintained during anesthesia. Following a period of preoxygenation, isoflurane alone is commonly used for the induction and maintenance of anesthesia. The animal may be placed within an induction chamber (for smaller animals, a large cone for dogs may serve this purpose); alternatively, a small cone may be placed near the animal’s nose until it unfolds. Hypersalivation may occur, and premedication with atropine is often advised if anesthesia is to be maintained with a mask.38 Tracheal intubation may be indicated for longer or oral procedures and is accomplished with a 1.0- to 1.5-mm endotracheal tube, Teflon IV catheter (stylet removed), or feeding tube.5,64 When intubation is to be performed, atropine may be contraindicated, as thickened secretions may obstruct the breathing tube. Injectable agents, including ketamine, diazepam, midazolam, xylazine, toletamine/zolazepam, and medetomidine have also been used—but may prolong recovery.50,64 Combinations include ketamine with midazolam and ketamine with
CHAPTER 30 African Hedgehogs medetomidine.10 Although medetomidine is no longer commercially available and has been replaced by dexmedetomidine, medetomidine can still be obtained through select compounding services. Analgesics include butorphanol, buprenorphine, tramadol, and meloxicam.10,32 When removal of spines is necessary, they may be clipped at the skin surface or removed with steady traction at the base.24 Lacerations and other wounds are occasionally seen. Hedgehogs often bite cage mates below the spined skin on the hind legs. Hedgehogs may self-mutilate traumatic or surgical wounds; therefore prompt primary closure with subcuticular sutures should be implemented whenever possible.24 Bandages and dilute chlorhexidine baths are well tolerated. If the cutaneous muscle is damaged, it must be closed in a separate layer. Contraction of the “rolling-up” musculature can cause wound dehiscence. Elizabethan collars are not practical in this species.24 Because hedgehogs are generally housed individually, castrations and ovariohysterectomies are generally performed to remove diseased organs rather than for population control. Surgical alteration as a means of preventing neoplasia of the reproductive organs has not been reported. Castration is performed through a para-anal skin incision over each testicle; the closed technique is preferred.24 Ovariohysterectomy is similar to this procedure in other mammals, although the practitioner should be prepared to encounter substantial fat surrounding the ovaries and mesosalpinx.24
ZOONOSES AND SUITABILITY AS PETS Several strains of Salmonella occur commonly in hedgehogs, particularly S. tilene, S. typhimurium, and S. enterititis; in many cases hedgehogs are asymptomatic carriers.11,12,59 Many cases of transmission from pet African hedgehogs to humans have been documented, particularly in young children living in homes with hedgehogs. In many cases the children had no direct contact with the hedgehogs.59 Hedgehogs’ messy droppings and propensity to walk in their feces may facilitate dissemination of Salmonella in the household. As with reptiles, it should be assumed that all pet hedgehogs can carry and transmit this pathogen, and animals should be handled accordingly (e.g., wash hands after handling, do not allow animals or fomites to contact food or food preparation areas, keep cages clean, etc.). Subclinically infected animals may shed intermittently, so cultures should not be used to rule out the carrier state. Treatment aimed at eliminating the carrier state is unlikely to be successful and should not be attempted because resistance may result. Several cases of human dermatophytosis transmitted from pet hedgehogs have been documented.59,60 In addition, some people are extremely sensitive to contact with African hedgehog spines and develop a transient but markedly pruritic urticaria within minutes after handling a hedgehog.17 One person reportedly developed a more persistent papular eruption after the urticaria. Human herpes simplex virus 1 was recovered from the liver of a pet African hedgehog that died acutely following glucocorticoid treatment for intervertebral disk disease.3 The owner of the animal reported that several family members suffered from cold sores, and it seems likely that the animal contracted the infection from a human in the household. A single case of Chagas’ disease (Trypanosoma cruzi) was reported in a captive African hedgehog that was housed outdoors in Texas.13 Monkeypox virus DNA was recovered from an African hedgehog
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housed with many other exotic species at a pet distributor facility in Illinois; the report does not specify whether this hedgehog was wild-caught or captive-bred.27 Other potentially zoonotic organisms recovered from pet hedgehogs include Cryptosporidium and Candida albicans.59 Tickborne diseases—including tularemia, Q fever, and CrimeanCongo hemorrhagic fever—have been reported in wild African hedgehogs,58,59 and Tahyna virus, Bhanja virus, and tick-borne encephalitis have been recovered from wild European hedgehogs.59 However, these diseases have not been reported in captive-bred hedgehogs. Wild-caught African hedgehogs can be infected with foot-and-mouth disease; signs include vesicular lesions on the tongue, muzzle, and paws. This virus can be transmitted to humans but presents a far greater threat to hoofed stock. To prevent introduction of this disease to the United States, importation of African hedgehogs was banned by the USDA in 1991.59 A single report of rabies in a hedgehog exists; the virus was recovered from a wild European hedgehog in Hungary.59 Rabies has not been reported in captive African hedgehogs, but the profuse salivation that occurs during selfanointing is occasionally mistaken as a sign of rabies. Additionally, Yersinia pseudotuberculosis and Mycobacterium marinum have been reported in captive European hedgehogs and Chlamydophila psittaci and Toxoplasma gondii have been recovered from wild European hedgehogs.59 Because hedgehogs are not particularly personable and have a relatively short life span, their suitability as pets has been questioned. In our experience, most hedgehogs are tame with their owners if handled consistently from an early age. As pets, hedgehogs have the advantages of being small, quiet, and solitary, and they have minimal odor. Unlike many other exotic species, their care is relatively straightforward and husbandry-related problems (other than obesity) are infrequent. Hedgehogs do not appear to suffer from boredom or stress in appropriate captive environments, and behavioral problems are rare. From a conservation perspective, the current pet hedgehog trade in the United States does not affect wild populations and has not led to the establishment of feral populations. Although some adult hedgehogs do resent handling, they are nevertheless very active and interesting animals.
References 1. Allen ME. The nutrition of insectivorous mammals. Proceedings of Specialty Program. Joint Conf Am Assoc Zoo Vet/Am Assoc Wildl Vet. 1992:113-115. 2. Allison N. A hyperplastic endometrial polyp and vascular thrombosis in a hedgehog. Vet Med. 2003;98:298-303. 3. Allison N, Chang TC, Steele KE, et al. Fatal herpes simplex infection in a pygmy African hedgehog (Atelerix albiventris). J Comp Pathol. 2002;126:76-78. 4. Bedford JM, Mock OB, Nagdas SK, et al. Reproductive characteristics of the African pygmy hedgehog. Atelerix albiventris. J Reprod Fertil. 2000;120:143-150. 5. Bennett RA. Husbandry and medicine of hedgehogs. Proceedings of Specialty Program. Annu Conf Assoc Avian Vet. 2000: 109-114. 6. Brunnert SR, Hensley GT, Citino SB, et al. Salivary gland oncocytes in African hedgehogs (Atelerix albiventris) mimicking cytomegalic inclusion disease. J Comp Pathol. 1991;105:83-91. 7. Buergelt CD. Histopathologic findings in pet hedgehogs with nonneoplastic conditions. Vet Med. 2002;97:660-665. 8. Campbell T. Intestinal candidiasis in an African hedgehog (Atelerix albiventris). Exot Pet Pract. 1997;2:79.
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9. Capello V, Lennox AM. African pygmy hedgehog. In: Capello V, Lennox AM, eds. Clinical radiology of exotic companion mammals. Ames: Wiley-Blackwell; 2008:481-487. 10. Carpenter JW. Hedgehogs. In: Carpenter JW, ed. Exotic animal formulary. 3rd ed. St. Louis: Elsevier Saunders; 2005:361-373. 11. Centers for Disease Control and Prevention (CDC). African pygmy hedgehog-associated salmonellosis—Washington, 1994. MMWR Morb Mortal Wkly Rep. 1995;44:462-463. 12. Craig C, Styliadis S, Woodward D, et al. African pygmy hedgehog–associated Salmonella tilene in Canada. Can Commun Dis Rep. 1997;23:129-131:discussion, 131-132. 13. deMaar TW, Kassell NL, Blumer ES. Chagas’ disease in an African hedgehog. Proceedings. Joint Conf Assoc Rept Amph Vet/Am Assoc Zoo Vet. 1994:151-153. 14. Done LB. What you don’t know about hedgehog diseases. Proceedings. North Am Vet Conf. 1999:824-825. 15. Done LB, Dietze M, Cranfield M, et al. Necropsy lesions by body systems in African hedgehogs (Atelerix albiventris): clues to clinical diagnosis. Proceedings. Joint Conf Am Assoc Zoo Vet/Am Assoc Wildl Vet. 1992:110-112. 16. Ellis C, Mori M. Skin diseases of rodents and small exotic mammals. Vet Clin North Am Exot Anim Pract. 2001;4:493-542. 17. Fairley JA, Suchniak J, Paller AS. Hedgehog hives. Arch Dermatol. 1999;135:561-563. 18. Fisher PG. Exotic mammal renal disease: causes and clinical presentation. Vet Clin North Am Exot Anim Pract. 2006;9:33-67. 19. Garner MM, Kiupel M, Munoz JF. Brain tumors in African hedgehogs (Atelerix albiventris). Proceedings of Specialty Program. Annu Conf Assoc Avian Vet/Assoc Exot Mam Vet. 2010:65. 20. Graczyk TK, Cranfield MR, Dunning C, et al. Fatal cryptosporidiosis in a juvenile captive African hedgehog (Atelerix albiventris). J Parasitol. 1998;84:178-180. 21. Graesser D, Dressen P, Spraker T. Wobbly hedgehog syndrome. Proceedings. Annu Conf Am Assoc Vet Lab Diagn. 2001:82. 22. Graesser D, Spraker TR, Dressen P, et al. Wobbly hedgehog syndrome in African pygmy hedgehogs (Atelerix spp.). J Exot Pet Med. 2006;15:59-65. 23. Greenacre CB. Spontaneous tumors of small mammals. Vet Clin North Am Exot Anim Pract. 2004;7:627-651. 24. Heatley JJ. Hedgehogs. In: Mitchell MA, Tully Jr TN, eds. Manual of exotic pet practice. St. Louis: Saunders Elsevier; 2009:433-455. 25. Heatley JJ, Mauldin GE, Cho DY. A review of neoplasia in the captive African hedgehog (Atelerix albiventris). Sem Avian Exot Pet Med. 2005;14:182-192. 26. Hoefer HL. Clinical approach to the African hedgehog. Proceedings. North Am Vet Conf. 1999:836-838. 27. Hutson CL, Lee KN, Abel J, et al. Monkeypox zoonotic associations: insights from laboratory evaluation of animals associated with the multi-state US outbreak. Am J Trop Med Hyg. 2007;76:757-768. 28. Johnson DH. Hedgehog with suspected bilateral renal calculi. Exot DVM. 2001;3:5. 29. Johnson DH. Miscellaneous small mammal behavior. In: Bays TB, Lightfoot T, Mayer J, eds. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: Saunders Elsevier; 2006:263-344. 30. Johnson-Delaney CA. Other small mammals. In: Meredith A, Redrobe S, eds. BSAVA manual of exotic pets. 4th ed. Gloucester, UK: British Small Animal Veterinary Association; 2002:102-115. 31. Johnson-Delaney CA. Common procedures in hedgehogs, prairie dogs, exotic rodents, and companion marsupials. Vet Clin North Am Exot Anim Pract. 2006;9:415-435. 32. Johnson-Delaney CA. What veterinarians need to know about hedgehogs. Exot DVM. 2007;9:38-44. 33. Juan-Sallés C, Raymond JT, Garner MM, et al. Adrenocortical carcinoma in three captive African hedgehogs (Atelerix albiventris). J Exot Pet Med. 2006;15:278-280. 34. Landes E, Zentek J, Wolf P, et al. Investigations on composition of milk and development of sucklings in hedgehogs. Kleintierpraxis. 1997;42:647-658.
35. Larsen RS, Carpenter JW. Husbandry and medical management of African hedgehogs. Vet Med. 1999;94:877-890. 36. Lennox AM. Emergency and critical care procedures in sugar gliders (Petaurus breviceps), African hedgehogs (Atelerix albiventris), and prairie dogs (Cynomys spp). Vet Clin North Am Exot Anim Pract. 2007;10:533-555. 37. Leonatti SR. Ornithonyssus bacoti mite infestation in an African pygmy hedgehog. Exot DVM. 2007;9:3-4. 38. Lightfoot TL. Clinical examination of chinchillas, hedgehogs, prairie dogs, and sugar gliders. Vet Clin North Am Exot Anim Pract. 1999;2:447-469. 39. Lightfoot TL. Therapeutics of African pygmy hedgehogs and prairie dogs. Vet Clin North Am Exot Anim Pract. 2000;3:155-172. 40. Martínez LS, Juan Sallés C, Cucchi-Stefanoni K, et al. Actinomyces naeslundii infection in an African hedgehog (Atelerix albiventris) with mandibular osteomyelitis and cellulitis. Vet Rec. 2005;157:450-451. 41. Meredith A. Skin diseases and treatment of hedgehogs. In: Patterson S, ed. Skin diseases of exotic pets. Ames: Blackwell Science; 2006:264-274. 42. Mikaelian I, Reavill DR, Practice A. Spontaneous proliferative lesions and tumors of the uterus of captive African hedgehogs (Atelerix albiventris). J Zoo Wildl Med. 2004;35:216-220. 43. Morgan KR, Berg BM. Body temperature and energy metabolism in pygmy hedgehogs. Am Zool. 1997;5:150A. 44. Ness RD. Clinical pathology and sample collection of exotic small mammals. Vet Clin North Am Exot Anim Pract. 1999;2:591-620. 45. Nowak RM. Walker’s mammals of the world. Vol I. 6th ed. Baltimore: Johns Hopkins University Press; 1999. 46. Peauroi JR, Lowenstine LJ, Munn RJ, et al. Multicentric skeletal sarcomas associated with probable retrovirus particles in two African hedgehogs (Atelerix albiventris). Vet Pathol. 1994;31:481-484. 47. Phair K, Carpenter JW. Management of extraskeletal osteosarcoma in an African hedgehog (Atelerix albiventris). J Exot Pet Med. 2011. In press. 48. Phillips ID, Taylor JJ, Allen AL. Endometrial polyps in 2 African pygmy hedgehogs. Can Vet J. 2005;46:524-527. 49. Powers LV. Subcutaneous implantable catheter for fluid administration in an African pygmy hedgehog. Exot DVM. 2002;4:16-17. 50. Pye GW. Marsupial, insectivore, and chiropteran anesthesia. Vet Clin North Am Exot Anim Pract. 2001;4:211-237. 51. Raymond JT, Garner M. Mammary gland tumors in captive African hedgehogs. J Wildl Dis. 2000;36:405-408. 52. Raymond JT, Garner MM. Cardiomyopathy in captive African hedgehogs (Atelerix albiventris). J Vet Diagn Invest. 2000;12:468-472. 53. Raymond JT, Garner MM. Spontaneous tumors in captive African hedgehogs (Atelerix albiventris): a retrospective study. J Comp Pathol. 2001;124:128-133. 54. Raymond JT, Garner MM. Spontaneous tumors in hedgehogs: a retrospective study of fifty cases. Proceedings. Joint Conf Am Assoc Zoo Vet/Am Assoc Wildl Vet/Assoc Rept Amph Vet/Nat Assoc Zoo Wildl Vet. 2001:326-327. 55. Raymond JT, White MR. Necropsy and histopathologic findings in 14 African hedgehogs (Atelerix albiventris): a retrospective study. J Zoo Wildl Med. 1999;30:273-277. 56. Raymond JT, Aguilar R, Dunker F, et al. Intervertebral disc disease in African hedgehogs (Atelerix albiventris): four cases. J Exot Pet Med. 2009;18:220-223. 57. Raymond JT, Williams C, Wu CC. Corynebacterial pneumonia in an African hedgehog. J Wildl Dis. 1998;34:397-399. 58. Reeve N. Hedgehogs. London: T & AD Poyser Ltd; 1994. 59. Riley PY, Chomel BB. Hedgehog zoonoses. Emerg Infect Dis. 2005;11:1-5. 60. Rosen T. Hazardous hedgehogs. South Med J. 2000;93:936-938.
CHAPTER 30 African Hedgehogs 61. Simone-Freilicher EA, Hoefer HL. Hedgehog care and husbandry. Vet Clin North Am Exot Anim Pract. 2004;7:257-267. 62. Smith AJ. Husbandry and medicine of African hedgehogs (Atelerix albiventris). J Small Exot Anim Med. 1992;2:21-28. 63. Smith AJ. Neonatology of the hedgehog (Atelerix albiventris). J Small Exot Anim Med. 1995;3:15-18. 64. Smith AJ. General husbandry and medical care of hedgehogs. In: Bonagura JD, ed. Kirk’s current veterinary therapy XIII small animal practice. Philadelphia: WB Saunders; 2000:1128-1133. 65. Snider TA, Joyner PH, Clinkenbeard KD. Disseminated histoplasmosis in an African pygmy hedgehog. J Am Vet Med Assoc. 2008;232:74-76. 66. Staley EC, Staley EE, Behr MJ. Use of permethrin as a miticide in the African hedgehog (Atelerix albiventris). Vet Hum Toxicol. 1994;36:138.
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67. Tully Jr TN, Mitchell MA. A technician’s guide to exotic animal care. Lakewood, CO: AAHA Press; 2001;179-186. 68. Vuolo S, Whittington JK. Dystocia secondary to a perianal fetal hernia in an African hedgehog. Exot DVM. 2008;10:10-12. 69. Wack R. Pemphigus foliaceus in an African hedgehog. Proceedings. North Am Vet Conf. 2000:1023. 70. Wheler CL, Grahn BH, Pocknell AM. Unilateral proptosis and orbital cellulitis in eight African hedgehogs (Atelerix albiventris). J Zoo Wildl Med. 2001;32:236-241. 71. Wolff CF, Corradini PR, Cortés G. Congenital erythropoietic porphyria in an African hedgehog (Atelerix albiventris). J Zoo Wildl Med. 2005;36:323-325.
SECTION SIX
General Topics
CHAPTER
31
Anesthesia, Analgesia, and Sedation of Small Mammals
Michelle G. Hawkins, VMD, Diplomate ABVP (Avian), and Peter J. Pascoe, BVSc, Diplomate ACVA, DVA, Diplomate ECVAA
General Principles and Challenges in Anesthetizing Small Exotic Mammals Balanced Anesthesia/Analgesia Preanesthetic Considerations Patient Evaluation Nutritional Status and Fasting Equipment Breathing Circuits Ventilators Clinical Techniques Vascular Access Intubation Preanesthetic Medications Injectable Medications Sedatives and Tranquilizers Induction and Maintenance of Anesthesia Injectable Anesthetics Inhalant Anesthesia Local and Regional Anesthesia Epidural Anesthesia/Analgesia Anesthetic Monitoring and Supportive Care Emergencies Small Mammal Analgesia Opioids Tramadol Hydrochloride Nonsteroidal Anti-Inflammatory Drugs Acupuncture Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
GENERAL PRINCIPLES AND CHALLENGES IN ANESTHETIZING SMALL EXOTIC MAMMALS There are many challenges to be faced when one is anesthetizing very small patients. It is more difficult to obtain preoperative blood work, secure an airway or intravenous access, monitor anesthesia, and prevent hypothermia once the animal is anesthetized. Small mammals have higher metabolic rates and smaller glycogen reserves, predisposing them to hypoglycemia. Metabolism and excretion of parenterally administered drugs are faster; therefore they have shorter durations of action. Small mammals have higher oxygen consumption and may thus suffer irreversible central nervous system (CNS) injury more rapidly during respiratory arrest. Time becomes critical during anesthesia; complications occur faster, and the time for intervention is shorter. Much dosing information has been established in terminal studies of young, healthy laboratory animals, so critically evaluate the information prior to using it in a sick or debilitated pet animal. A recent large-scale prospective study reported an overall perianesthetic mortality rate of healthy pet rabbits and guinea pigs of approximately 1.39% and 3.80%, respectively, values 5 to 10 times higher than those found in dogs and cats.9 There is substantial room to decrease mortality during anesthesia in these patients.
BALANCED ANESTHESIA/ANALGESIA Isoflurane has been the most common anesthetic used in small exotic mammals for the last two decades and is often administered as the sole agent. Inhalant anesthetics have the advantage of allowing rapid changes in anesthetic depth and rapid 429
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recoveries owing to their low solubility in tissues and elimination via the respiratory tract. But inhalants also induce dosedependent cardiopulmonary depression. Clinically, 1.3 to 1.5 times the minimum alveolar concentration (MAC) of an inhalant is required for immobility during surgery, but inhalants do not block activation of nociceptive pathways by noxious stimuli. High inhalant doses (1.5-3 times MAC) are required to block undesirable responses to pain, such as changes in heart rate, blood pressure, and respiratory rate, but the margin of safety at these doses is low and circulatory collapse can occur.121 While inhalants are still the main agents used for many procedures, there are many ways to minimize their use and thus decrease their adverse effects. The concept of “balanced anesthesia” originated to provide the elements of anesthesia (unconsciousness/amnesia, immobility, muscle relaxation, reduced autonomic response to noxious stimuli) by using multiple drugs at lower overall doses so as to minimize the adverse effects of each drug while still providing appropriate anesthesia and analgesia for the procedure. This approach reduces the magnitude of cardiovascular depression under anesthesia and promotes hemodynamic stability. Even with balanced anesthesia protocols, morbidity and mortality increases with increased duration of anesthesia. Minimize anesthesia time by preparing the patient as well as all drugs and equipment in advance and provide close patient monitoring so that adjustments in anesthesia are identified and corrected immediately. Morbidity and mortality also occurs during recovery, and it is important to continue to monitor animals until they are fully recovered.
PREANESTHETIC CONSIDERATIONS PATIENT EVALUATION A complete history should be obtained and a physical examination performed to assess preexisting clinical problems and record baseline values for comparison. Age should be considered, because the margin of safety is wider in younger animals. Obtain an accurate body weight for fluid and drug dose calculations. Evaluate hydration status, and correct dehydration prior to anesthesia. Compensatory mechanisms are blunted under anesthesia, exacerbating underlying hypotension and poor peripheral perfusion. Whenever possible, preanesthetic noninvasive blood pressure should be assessed to provide a more objective assessment of perfusion. Ideally, a complete blood count and biochemical profile should be performed prior to anesthesia. Severe anemia should be corrected, because the patient may not be able to compensate for decreased oxygen delivery. The packed cell volume (PCV) may decrease 3% to 5% during anesthesia because of vascular hemodynamic changes. In ferrets, isoflurane and sevoflurane are associated with significant reductions in PCV.77-79 Glucose concentration should be monitored and supplemented as necessary. The type of surgery to be performed and potential for blood loss should be assessed for fluid plan preparation.
NUTRITIONAL STATUS AND FASTING Prolonged anorexia leads to negative energy balance and hypoglycemia, requiring nutritional support in the perianesthetic period. Nutritional support for the debilitated patient is discussed in Chapter 38. Overnight fasting prior to anesthesia is not performed in small mammals because of their rapid
gastrointestinal (GI) transit times and potential for GI stasis. Fasting has been advocated to reduce stomach and cecal volumes in large and obese rabbits, which can theoretically compress their small thoracic cavities, but this generally does not change GI volumes enough to be clinically important. Fasting exhausts glycogen stores, resulting in hypoglycemia, and may significantly contribute to perianesthetic ileus and potentially exacerbate pancreatic tumors in ferrets. Fasting reduces regurgitation in ferrets, but vomiting/regurgitation is uncommon in most other small mammal species. Rabbits and rats cannot vomit because of their well-developed cardiac sphincter and limiting ridge of the stomach, respectively. In general, recommended fasting times are 0 to 4 hours, with prudence used to weigh the risks and benefits in the specific patient. Schedule procedures early in the day to reduce fasting times. Many small rodents store food in their oral cavities; thus the mouth is cleaned out by gently syringing water into the oral cavity to remove food materials in tolerant awake patients or with wet cotton tips immediately after induction. Elevate the head during anesthesia if there is any concern of aspiration.
EQUIPMENT BREATHING CIRCUITS Nonrebreathing circuits (Ayre’s T-piece, Mapleson systems, Magill, Jackson-Rees, Norman elbow, and the Bain circuit) are most commonly used for anesthetizing small mammals. The nonrebreathing system depends upon high rates of fresh gas flow to remove carbon dioxide (CO2) and provides rapid changes in inhalant concentration when the vaporizer setting is changed. Circle systems are heavier and bulkier, which predisposes to accidental extubation and difficulty working around the head region. These circuits also have a greater resistance to breathing. For these reasons, the nonrebreathing circuit is used most commonly in small mammals. In general, the flow rate used for a T-piece or Norman elbow should be two to three times the minute ventilation. But in the Bain circuit, a flow rate of 150 to 200 mL/kg per minute appears effective. Removal of waste gases from the circuit occurs during the pause between expiration and inspiration. Thus if the respiratory rate is high (shorter pause and therefore more mixing of inspired and expired gases), a higher flow rate should be used (e.g., if RR >40, increase the flow rate to 300 mL/kg/min).
VENTILATORS Ventilators appropriate for small mammals must cope with the range of tidal volumes in the different species. In very small patients, this may be less than 1 mL; but some giant-breed rabbits might need to be as high as 500 mL. In general the three things that must be controlled by a ventilator are tidal volume, the rate at which the tidal volume is delivered, and the number of breaths per minute. The way in which each of these is controlled varies from ventilator to ventilator.45
CLINICAL TECHNIQUES VASCULAR ACCESS Vascular access is necessary for replacement fluids and to deliver anesthesia and emergency medications. Sites for intravenous catheterization include the cephalic, lateral saphenous, and
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals
A
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B Fig. 31-1 Catheterization of the cephalic vein is the most common intravenous site used (A), but the lateral saphenous, jugular, or femoral veins and the superficial lateral tail veins in the rat can also be used. Catheters can be taped and bandaged for security; additional stability can be provided with tongue depressors for very small limbs. In cases where intravenous catheterization is not possible, an intraosseous catheter may be placed in the tibia (B) or femur for short-term use.
femoral veins in larger mammals and the superficial lateral tail veins in the rat (Fig. 31-1). Sedation may be necessary for intravenous catheter placement. A surgical cut-down procedure under anesthesia is generally necessary for jugular catheterization. Small-bore over-the-needle catheters (≤24 gauge) are used. The catheter site should be aseptically prepared; catheters are then secured with tape and/or sutured in place and bandaged for extra security. Jugular catheters, if left indwelling, require 24-hour monitoring, as fatal hemorrhage can occur if the patient pulls or chews on the catheter and damages the vessel. Many small rodents, even when severely compromised, are intolerant of bandaging material and indwelling catheters and will attempt to remove them. Intraosseous (IO) cannulation can be useful in smaller patients or during cardiovascular collapse (see Fig. 31-1).89 Products that can be used as IO cannulas include 18- to 24-gauge 1- to 1½-in. spinal needles or 18- to 25-gauge 1-in. hypodermic needles, depending upon the animal’s size. The cannula should be long enough to extend through one-third to one-half the length of the medullary cavity. A wire stylet reduces the potential for a bone core. Prepare several hypodermic needles (25- to 18-gauge) with wire stylets (stainless steel suture) and sterilize them for IO catheterization. Common sites for IO cannula placement include the trochanteric fossa of the femur and the tibial crest (see Fig. 31-1, B). Placement is similar to that of a normograde intramedullary pin and requires strict aseptic technique during placement and maintenance. Once the cortex is penetrated, the cannula should advance easily, with little resistance. Flush the cannula with heparinized saline immediately to prevent clotting. Cover the insertion site with an antibiotic ointment and secure the cannula with tape, suture, and a bandage. IO cannulas can remain patent for 72 hours without flushing but should be flushed with heparinized saline twice daily if fluid therapy is not continuous. Complications associated with IO catheterization include penetration of both cortices, failure to properly enter the medullary cavity, and extravasation of fluids with associated pain.89 IO catheters are used primarily for short-term vascular volume expansion until an IV catheter site can be obtained. Many animals become uncomfortable on limbs supporting IO catheters even after short-term placement. IO catheterization is contraindicated in septic patients and those with metabolic bone disease. Osteomyelitis may occur owing to duration or placement of the
IO catheter; administration of alkaline or hypertonic solutions can contribute to osteomyelitis and cause pain.89 Dilute these solutions before administration and flush the catheter with heparinized saline after any drug injection. Arterial catheterization for the evaluation of blood pressure, blood gas tension, and pH can be performed in many species. The most common sites include the pedal arteries in most species, the central auricular artery in the rabbit, and the ventral tail artery in the rat and ferret. The catheters are usually placed percutaneously, but a cut-down procedure is sometimes used.
INTUBATION Endotracheal intubation provides a patent airway, reduces dead space, and facilitates positive-pressure ventilation. Disadvantages include tracheal mucosal trauma, increased airway resistance, and airway occlusion due to mechanical forces or secretions. Increased resistance is of greater importance in very small patients because it is inversely related to the fourth power of the tube radius. For example, decreasing the tube radius from 5 to 3 mm increases the resistance 7-fold, whereas a decrease from 3 to 1 mm increases it more than 80-fold. Increased resistance can be overcome by positive pressure ventilation. Specialized endotracheal tubes and light sources are available to aid intubation of small mammals. The smallest commercial uncuffed tubes have an internal diameter (ID) of 1 mm. However, tubes of less than 2-mm ID are often highly flexible and kink easily. The smallest-diameter cuffed tube is 3-mm ID. Uncuffed tubes do not provide a sealed airway, so clean the oral cavity prior to intubation, elevate the head, and monitor during the procedure. If you are using a cuffed tube, the cuff is carefully inflated with just enough air to prevent leakage when 10 to 15-cm H2O pressure is applied. Very small mammals may be intubated with Teflon IV, red rubber, or urinary catheters (Fig. 31-2, A). Care is taken to ensure that no sharp edges are present at the end. This is achieved by using a small piece of silicone tubing over the end of the catheter (see Fig. 31-2, B,C). Endotracheal tube obstruction is detected by monitoring for a prolonged expiratory phase. Anticholinergics reduce the production of mucus and mucous plug formation but also increase mucous viscosity, making it harder to clear secretions. The use of an endotracheal tube with a Murphy eye decreases the
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A
B
C Fig. 31-2 A, Commercial endotracheal tubes are useful for larger mammals, but Teflon intravenous, red rubber, and urinary catheters are often necessary to intubate very small mammals. A small piece of silicone tubing can be glued over the end of the intravenous catheter to minimize sharp edges (B, C [inset]).
likelihood of mucous occlusion. Humidification of the gases reduces mucous plug formation. Commercial endotracheal tube humidifiers are available (Humidi-vent Mini Agibeck Product, Hudson RCI, Temecula, CA). Disadvantages of their use include increased dead space and plugging of the filter with secretions. Care must be taken to minimize head and neck movement in intubated patients, as movement of the tube and changes in positioning may induce mucosal trauma. Sublaryngeal tracheal injury, ulceration, and postintubation tracheal strictures have been described in rabbits from several institutions that were intubated with both cuffed and uncuffed endotracheal tubes.38,92 Other factors that predispose to mucosal injury include ventilation technique and endotracheal tube disinfection protocols. Ferret intubation is straightforward and should be performed routinely as in the cat. All pet rabbits over 1-kg body weight can be intubated, but this is more difficult. Rabbits have large incisors, long narrow oral cavities, and thick tongues, making laryngeal visualization more difficult, and laryngospasm is easily induced. There are a number of tracheal intubation techniques that have been advocated for use in rabbits: direct visualization of the larynx with a laryngoscope and intubation, blind intubation with the neck in extension, endoscope-guided intubation, and nasotracheal intubation. Apply topical local anesthetic on the larynx 60 seconds before intubation to decrease laryngospasm. The authors use the blind technique routinely. Guide the tube to the larynx and listen for louder breath sounds, place one drop of 2% lidocaine via tomcat catheter through the endotracheal tube directly on to the larynx, then use the same approach
for intubation (Fig. 31-3). Placement of the tube is confirmed by visualizing the movement of water vapor in the tube, by the rabbit coughing, by auscultation of breath sounds in the lungs, and definitively by attaching a capnograph and seeing a characteristic capnographic trace. Rabbits are obligate nasal breathers and the patency of both nares must be carefully assessed postextubation, especially if the animal has been in dorsal recumbency. Reported complications include postextubation obstruction, respiratory arrest, and tracheal mucosal injury.38,92 Intranasal intubation or catheterization can be used in the rabbit if endotracheal intubation cannot be performed or for complicated oral procedures, such as extensive dental procedures. The ventral meatus of the rabbit’s nasal passage is surprisingly large. A blind intubation technique with a 2.5- to 3-mm ID endotracheal tube is used, but these are often stiff for smaller rabbits; standard IV tubing has a thinner wall and so is more flexible. IV tubing can be cut to the appropriate length and a bevel fashioned on the distal end; the proximal end can be fitted with a 2- to 2.5-mm endotracheal tube adapter. Administer anticholinergics with nasal intubation, as vagally mediated bradycardia can be associated with this procedure. The tube is passed through the ventral meatus with the head held in a normal or slightly extended position. Because of the difficulties of intubating rabbits, newer techniques such as laryngeal mask airways are being evaluated.7,66,111 Rodents also have a long narrow oral cavity, but equipment and techniques for intubating small rodents have been developed.87,117 In guinea pigs and chinchillas, orotracheal intubation is complicated by the fusion of the soft palate to the base of the tongue, creating the palatal ostium, which is highly vascular and easily traumatized (Fig. 31-4, A). Endotracheal tubes of 1.0- to 2.5-mm ID are most often needed for small rodents. Very small rodents may be intubated with catheters or IV tubing as in the rabbit, but occlusion with mucous plugs, due to the small internal diameter of these tubes, occurs frequently. An otoscopic cone, modified pediatric blade, commercially available rodent work stands and intubation packs, or endoscopy can help facilitate intubation. Otoscopic cones that have been modified by removing a section laterally can facilitate visualization of the epiglottis and direct placement of the endotracheal tube. A stylet placed first may help facilitate endotracheal tube placement (see Fig. 31-4, B,C). Endoscopy provides the best visualization of the epiglottis and minimizes trauma during tube placement. Nasal intubation/catheterization has also been performed for guinea pigs and other small rodents with similar issues arising as for rabbits.
PREANESTHETIC MEDICATIONS Parasympatholytics are most commonly used to minimize salivary and bronchial secretions and vagally induced bradyarrhythmias, but they can increase secretion viscosity. Many rabbits have high circulating concentrations of atropine esterases, thus reducing the efficacy of atropine and prompting the use of much higher atropine doses, with redosing as often as every 10 to 15 minutes during bradycardia.44 Large doses of parasympatholytics can alter GI motility in hind-gut fermenters; therefore the lowest doses necessary are used.48 Glycopyrrolate has a longer duration of activity than atropine (Table 31-1). Glycopyrrolate increased heart rates in rats for 240 minutes, versus 30 minutes with atropine.86 The same study demonstrated that 0.1 mg/kg glycopyrrolate administered to rabbits increased heart rate for an equal duration to the rats, but atropine doses ≤2 mg/ kg did not increase heart rate for any duration.86
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals
A
C
B
Fig. 31-3 Intubation of the rabbit using the blind technique. Provide fresh oxygen and anesthetic gas through a nasal mask during the procedure. Guide the tube to the larynx, listen for louder breaths and/or visualize water vapor in the tube (A), place one drop of 2% lidocaine via tomcat catheter through the endotracheal tube directly onto the larynx (B), wait 60 seconds, then use the same approach for intubation (C). Placement of the tube is confirmed by visualizing movement of water vapor in the tube, by the rabbit coughing, by auscultation of breath sounds in the lungs, and definitively by attaching a capnograph and seeing a characteristic capnographic trace.
A
B
C Fig. 31-4 In guinea pigs and chinchillas, orotracheal intubation is complicated by the fusion of the soft palate to the base of the tongue, creating the palatal ostium, which is highly vascular and easily traumatized (A). Intubation can be accomplished with the use of an otoscope or endoscope and placement of a stylet (B), then threading the endotracheal tube over the stylet (C).
433
434
Table 31-1 Injectable Preanesthetic, Sedative, and Tranquilizer Drugs Used in Small Exotic Mammalsa
Glycopyrrolate Sedatives/ tranquilizers Acepromazine
Ferret
Guinea Pig
Chinchilla
Rat
0.2-1.0 SC, IM, IV 0.01-0.02 SC, IM, IV
0.05 SC, IM, IV 0.01-0.02 SC, IM, IV
0.05 SC, IM, 0.05 SC, IM, 0.05 SC, IM, IV IV IV, IP 0.01-0.02 0.01-0.02 SC, 0.02-0.5 SC, SC, IM, IV IM, IV IM, IV25
0.25-1.0 SC, IM
0.1-0.2 SC, IM25 ≥ 1 SC, IM, IV
0.5-1.0 SC, IM25 ≥ 1 SC, IM, IV
Hamster
Gerbil
Mouse
0.04 SC25
0.04 SC25
0.5 IM25
−
0.05 SC, Higher doses may be IM, IP necessary in rabbits 0.02-0.5 SC, − IM25
0.5-1.0 SC, IM ≥ 1 SC, IM, IV54
0.5-2.5 SC, IM, IP25 0.1-1.0 SC, IM, IV, IP25
−
3 IM25
−
−
Atipamezole
≥ 1 SC, IM, IV68,88
Dexmedetomidine
0.05-0.125 IM 0.05-1.0 μg/kg IM, IV
0.04-0.1 IM 0.05 SC 0.05-1.0 μg/kg IM, IV
0.05 SC
0.015-0.5 SC, IP
−
−
Diazepam
0.5-2.0 IV
0.5-2.0 IV
0.5-3.0 IV
0.5-3.0 IV
2.5-5.0 IV, IP
5 IM, IP25
5 IM, IP25
Flumazenil
0.05-0.1 SC, IM, IV 0.1-0.25 SC, IM 1-2 μg/kg IM, IV
0.05-0.1 SC, IM, IV 0.01-0.2 SC, IM25,70 1-2 μg/kg IM, IV
0.05-0.1 SC, 0.05-0.1 SC, IM, IV IM, IV54 0.1 SC 0.1 SC
−
−
−
0.03-0.1 SC, IP25 −
0.1-0.2 IP25
−
0.1-1.0 IM, IV, IP25 0.03-0.1 SC, IP25 −
−
0.5-2.0 SC, IM, IV 2-10 IM, IV
1-2 SC, IM, IV 1.0-2.5 SC, IM, IV; 5 IP25 2-10 IM, IV 1-5 IM, IV, IP
5 IM, IP25
1-5 IM, IP25
2 IM25
Yohimbine
0.2-0.4 IV
0.25-0.5 SC, IM, IV 0.1-0.5 SC, IM25 0.2-1.0 IM, IV
5 IP25
Xylazine
0.25-2.0 SC, IM, IV 2-5 IM, IV25
0.5-1.0 IM, IV
0.5-1.0 IM, IV 0.2 IV; 0.5 IM25
−
−
Medetomidine
Midazolam
Comments
2-5 SC, IM, IP25 ≥ 1 SC, IM, IV, IP
May induce seizures in gerbils 1:1 volume reversal of medetomidine or dexmedetomidine 0.015-0.5 Half the dose of SC, IP medetomidine; Microdoses combined with benzodiazepines and opioids for sedation, induction 3-5 PO, IP25 Significant irritation if given IM − Reversal of benzodiazipines 0.03-0.1 SC, Light sedation, rarely IP25 used alone; variable − effects in guinea pigs Microdoses combined with benzodiazepine and opioid for induction 1-5 SC, Lower doses for IM, IV25 premedication 5-10 IP25 Seldom used 0.5-1.0 IM, IV
Reversal for xylazine
aThese are suggested doses based on published information and the authors’ clinical experience. Species and individual variation in response to a given drug can be uncertain so adjust the dose depending on the clinical response of the animal. All doses are in milligrams per kilogram unless specified otherwise.
SECTION VI General Topics
Preanesthetics Atropine
Rabbit
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals
INJECTABLE MEDICATIONS A considerable increase in the use of injectable anesthetic combinations in small exotic mammals has recently been documented.* Certain breeds or even strains of small mammals have differing responses to injectable anesthetics.5 There is little published work evaluating different injectable combinations for common surgical procedures in pet animals of differing age, sex, breed, and health status in the clinical setting,40,88 and so use caution when extrapolating injectable doses from laboratory animal studies. Using reversible injectables provides better control of anesthetic depth, less hypothermia, less cardiopulmonary depression, and shorter recovery times.54-56,98 Muscle necrosis and cutaneous reactions can occur in any patient administered IM injections, depending on the drug formulation or volume delivered. Drugs are often not uniformly absorbed when administered SC, resulting in erratic and unpredictable anesthesia. The intraperitoneal (IP) route of injection is commonly used in laboratory medicine, but errors during administration (intravisceral, SC, or intra-adipose tissue) can result in organ damage or delayed onset of action. Injections are usually performed into the lower left quadrant of the abdomen with the rodent restrained in dorsal recumbency and the cranial end of the rodent directed downward. Supplemental oxygen and assisted ventilation should always be available when injectable anesthetics are used, regardless of their route of administration.
SEDATIVES AND TRANQUILIZERS Acepromazine In ferrets, acepromazine provides adequate sedation for simple procedures such as ear cleaning,71 however, one author discourages its use in ferrets owing to its vasodilatory effects (see Table 31-1).64 Acepromazine has been used extensively in rabbits and rodents and reported doses vary widely.4,48,120,122 The peak effect in rabbits is often not seen for 30 to 40 minutes, even when given IV. It is used in laboratory rabbits specifically for blood collection because of the drug’s vasodilatory effects. Vasodilation occurs at very low doses of acepromazine, therefore low doses will not avoid this effect. Animals that are hypovolemic, anemic, or hypotensive before anesthesia should not receive acepromazine. Acepromazine has also been shown to decrease tear production in rabbits.35 Acepromazine is not recommended for use in gerbils because it may lower the seizure threshold.49
Benzodiazepines Diazepam can be administered PO or IV, but if used IV, the patient must be monitored for hypotension caused by the propylene glycol found in many diazepam formulations (see Table 31-1). IM and SC absorption is erratic because of the solution’s high osmolality, which is due to an increased pH with this carrier. Because parenteral formulations of diazepam are available, large volumes must often be given. Transient lameness after IM injection has been reported in ferrets.72,74 Diazepam can be used alone in rabbits to provide preoperative anxiolysis and sedation,15 but it is most commonly used in combination with other drugs to enhance their activity or decrease muscle rigidity. Midazolam is a short-acting benzodiazepine that is watersoluble, so it can be given IM (see Table 31-1). Midazolam has
*References 4, 40, 53-55, 72, 74, 88, 120, 122.
435
been used as a sole sedative for minor, nonpainful procedures in rabbits and rodents. Lower doses are most often used for preanesthetic sedation. Flumazenil will reverse midazolam, but because its half31-life is shorter than that of midazolam, resedation may occur. Titrate the flumazenil dose to avoid the reversal of the beneficial anxiolysis, sedation, and muscle relaxation. Benzodiazepines coupled with opioid medications provide sedation and preoperative analgesia.
Alpha-2 Agonists Xylazine, medetomidine, and now dexmedetomidine are the most commonly used alpha-2 adrenergic receptor agonists in small exotic mammals (see Table 31-1). Their major advantages are that they provide good muscle relaxation and can be reversed with yohimbine, atipamezole, or tolazoline (see Table 31-1). If all analgesic components are reversed, postoperative analgesia should be provided before reversal of anesthesia. These drugs can have significant cardiopulmonary effects, including respiratory depression, second-degree heart block, bradyarrhythmias, and increased sensitivity to catecholamine-induced cardiac arrhythmias (xylazine).53,54 Because of the many adverse effects reported for this class of drugs, their use is not recommended in small exotic mammal patients with evidence of cardiopulmonary compromise. Supplemental oxygen and assisted ventilation should always be available when these drugs are being used. Xylazine/ketamine given over multiple anesthetic episodes and detomidine alone or in combination with ketamine or diazepam have been associated with myocardial necrosis and fibrosis in rabbits.61,80 Significant breed and strain differences in response to medetomidine sedation doses have been reported for rabbits and rats.5,6,48 In rabbits, ketamine/medetomidine allowed for more rapid intubation, a greater isoflurane-sparing effect, and less esophageal temperature loss than with ketamine/ midazolam, but the rabbits thus treated were more prone to laryngospasm.40 Medetomidine/ketamine provided better quality and duration of surgical anesthesia (38.7 ± 30.0 minutes) in healthy rabbits than medetomidine/midazolam/fentanyl (MMF). Arterial pH and PaO2 were significantly decreased in both groups and apnea occurred postintubation with MMF.53 Atipamezole (10%-20% of calculated dose) may reduce these adverse effects without reversing their anesthetic and analgesic effects, but supplemental oxygen must be available. The effects of alpha-2 agonists with ketamine seem to be less uniform in guinea pigs, and they may not become sufficiently anesthetized.96 Chinchillas administered MMF IM had wellcontrolled anesthesia for 1.5 hours, but the heart rate (HR) and respiratory rate (RR) were decreased and recoveries were prolonged without reversal.54 Chinchillas receiving medetomidine/ ketamine had greater HR and RR depression, and recovery was longer than with MMF combinations (Table 31-2).54 Alpha-2 agonists are combined with ketamine for IP injection for surgical anesthesia with good muscle relaxation in rats and mice.4,96
INDUCTION AND MAINTENANCE OF ANESTHESIA Preoxygenation should be performed routinely, and always when there is potential for hypoxemia. Preoxygenation can produce an oxygen reservoir and the benefits are achieved in ≤1 minute of high inspired oxygen concentration in the healthy patient but may take ≥5 minutes in a patient with compromised respiratory function. Preoxygenation is accomplished with an
436
Rabbit
Ferret
Guinea Pig
Chinchilla
Rat
Hamster
Gerbil
Mouse
Comments
Injectable Anesthetics/Analgesics 1-2 IP20
Etomidate
1-2 IV
1-2 IV
Ketamine
5-50 SC, IM, IV25
5-50 SC, IM, IV70
5-40 SC, IM, IV
5-40 IM, IV
10-40 IM, IV; 50-100 IP25
50-100 IP25
100-200 IM, IV25
Ketamine (K)/ diazepam (D)
10-25 K/1-5 D IV25,48
10-25 K/1-3 D IV25,70
20-30 K/1-2 D IV
40-100 K/3-5 D IP33
70 K/2 D IP25
50 K/5 D25 100 K/5 D IP25,33
Ketamine (K)/ diazepam (D)/ butorphanol (B) Ketamine (K)/ medetomidine (M) Ketamine (K)/ medetomidine (M)/butorphanol (B) Ketamine (K)/ midazolam (Mi) Induction Anesthesia
−
15 K/3D/0.2 B IM, IV70
100 K/5 D IM25; 20-30 K/1-2 D IV −
−
−
−
−
−
4-8 K/0.085-35 K/0.25-0.5 M 0.1 M IM17,39,40,50,53,68,88 IM25,70 5 K/0.08 5-15 K/0.1-0.5 M/0.1-0.2 M/0.4-0.5 B SC, B IM70 IM50
5-40 K/0.050.5 M IM25
5 K/0.06 M IM54
100 K/0.25 M IP25
75 K/0.5 M IV25
−
−
45-75 K/0.30.5 M IM, IP42,51 −
−
−
50-75 K/0.11.0 M IP25,42 −
5 K/0.25-1.0 Mi IM, IV
5 K/0.1-0.3 Mi IM, IV
5 K/0.5 Mi IM, IV
5 K/0.5 Mi IM, IV
−
−
−
−
10-40 K/1-3 Mi IM, IV39,40,48
5-10 K/0.25- 20-50 K/0.50.5 Mi IM, 5.0 Mi IM, IV70 IV 25 K/1-2 X 20-40 K/1-2 IM25 X IM
20-50 K/0.55.0 Mi IM, IV 20-40 K/1-2 X IM
40-150 K/3.0- − 5.0 Mi IV, IP25 40-100 K/5-10 200 K/10 X X IM, IP25,33 IP25
−
−
0.05 M/0.02 − F/1 Mi IM54
40-150 K/3.0- Lighter anesthesia at 5.0 Mi IP25 lower dosages of ketamine 35-100 K/2.5- − 15.0 X IP25,42 − Must be prepared to intubate because apnea, respiratory depression common; complete reversal with atipamezole, flumazenil, naloxone
Ketamine (K)/ xylaxine (X)
10-50 K/3-5 X IM17,53
Medetomidine (M)/ 0.25 M/0.02 F/1 Mi fentanyl (F)/ IM53 midazolam (Mi)
−
−
50 K/2 X IP25 −
100-200 IM, IV25
Administer 15-20 minutes after benzodiazepine Lower doses for induction; can cause muscle necrosis in guinea pigs Poor analgesia Provided 20-25 minutes anesthesia in ferrets − −
−
SECTION VI General Topics
Table 31-2 Injectable Anesthetic Drugs and Drug Combinations Used in Small Exotic Mammalsa
3-10 IV10
−
−
3-25 IV25
3-10 IM, IV70 −
−
40-80 IV, IP25,96,104
50-80 IP25
−
40-80 IP25,96
Bupivacaine (local, <2 regional blocks)
<2
<2
<2
<2
−
−
<2
Lidocaine (local, regional blocks)
<4
<4
<4
<4
−
−
<4
3-6 IV
3-6 IV
Tiletamine/ zolazepam
5-15 IM, IV18
3-6 IV
Doses for routine induction Avoid higher doses in rabbits due to renal injury18
Local Anesthetics
<4
Short duration of 20-30 min in rats Dosages for local blocks; can extend to 2 mg/kg as maximum dose Short duration of 20-30 min in rats
aThese are suggested dosages based on published information and the authors’ clinical experience. Species and individual variation in response to a drug or combination of drugs can be uncertain, so the dosage should be adjusted depending upon the clinical response of the animal. All doses are in milligrams per kilogram unless specified otherwise.
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals
3-6 IV
Propofol
437
438
SECTION VI General Topics
oxygen cage or an induction chamber in the unrestrained animal or with a face mask in the physically restrained patient. Levels rise more slowly in an unprimed oxygen cage or induction chamber. It is expected that PaO2 will decrease rapidly if oxygen delivery is interrupted and the animal begins to breathe room air again. Supplemental O2 should be supplied throughout the anesthesia, and intubation should be performed whenever possible.
INJECTABLE ANESTHETICS Ketamine Ketamine is a noncompetitive n-methyl-d-aspartate (NMDA) receptor antagonist that can prevent central sensitization, provide analgesia, and cause dissociative anesthesia (see Table 31-2). Ketamine is generally well tolerated in the stable patient but it has sympathomimetic properties that can cause increases in HR, myocardial contractility, and peripheral vascular resistance. It is a negative inotrope; therefore patients that are highly stressed (especially rabbits and chinchillas) may become hypotensive following ketamine injection because they have maximized their sympathetic output, thus “unmasking” the negative inotropic effect of the drug. Patients with hypertrophic cardiomyopathy may be at an increased risk of cardiac failure, as they may not be able to cope with the increased myocardial demand caused by sympathetic stimulation. Renal impairment can markedly prolong recovery. The authors do not recommend using ketamine for patients with cardiac or renal disease or in severely stressed small mammal patients. Salivation is increased and secretions can obstruct the airway. In small rodents, higher IM doses have been associated with muscle necrosis at the injection site. Ketamine has a very short duration of clinical effect in rabbits and varies within strains of rabbits, partly owing to redistribution and renal elimination.5 Another disadvantage is the potential for the development of apneustic ventilation, a pattern characterized by a prolonged pause after inspiration. Despite these disadvantages, ketamine combined with one or more drugs is still considered a first-choice injectable anesthetic.97 Ketamine is commonly combined with alpha-2 agonists or benzodiazepines as induction agents to improve relaxation and anesthetic depth (see Table 31-2).* Ketamine/alpha-2 agonist combinations, particularly at higher doses, produce mild-tosevere dose-dependent hypotension and bradyarrhythmias; ketamine/benzodiazepine combinations produce less cardiopulmonary depression and analgesia but provide good muscle relaxation.40 Low-dose ketamine/midazolam IV is used by the authors to induce anesthesia in rabbits, ferrets, and small rodents; it is given IP in very small rodents. Opioids added to ketamine combinations further reduce the dose and minimize the adverse effects of each drug. Reversal of other drugs in ketamine combinations allows for its specific undesirable effects to become apparent.
Tiletamine-Zolazepam In mice, tiletamine-zolazepam (Telazol, Fort Dodge, IA) alone appears to produce poor analgesia, but in combination with xylazine (7.5 mg/kg tiletamine-zolazepam and 45 mg/kg xylazine) this improved, with an anesthetic time of 30 to 90 minutes (see Table 31-2). In rats, doses of 30 to 60 mg/kg were tested
*References 17, 40, 52-54, 69, 88, 107, 120.
and found to have a dose-dependent effect on the duration of anesthesia (59-124 minutes), but not much change in cardiorespiratory effects was seen over this high dose range.104 Much smaller doses of tiletamine-zolazepam (1.5-3.0 mg/kg) are used in ferrets with xylazine (1.5-3.0 mg/kg) or with xylazine (1.5 mg/kg)/butorphanol (0.2 mg/kg) with a dose-dependent duration of effect.70 Tiletamine-zolazepam has been reported to cause nephrotoxicity, related to the tiletamine, in New Zealand white rabbits, and this appears to be dose-dependent.18 The advantage of this combination over ketamine is that much smaller volumes of drug are needed, so it is less likely to cause muscle damage if given IM.
Propofol The need for IV access and ventilatory support limits the use of propofol. Rapid, excitement-free induction and recovery occur at clinically relevant doses in most mammals (see Table 31-2). Early studies evaluating propofol anesthetic maintenance as the sole anesthetic agent in rabbits demonstrated a very narrow therapeutic window,2 but balanced anesthesia may produce propofol-sparing effects, thus reducing its respiratory disadvantages.73,81 The authors suggest a calculated propofol induction dose be given in one-quarter increments, each administered over 30 to 60 seconds, to allow more accurate dosing and minimize apnea and hypotension during induction. The effects of propofol given IP in small rodents are unpredictable; very little information is available on using this drug via this route in small exotic mammals.83 New formulations of propofol have been developed that may change some of the uses in these species. A clear solution of nanoparticulate propofol in an aqueous base eliminates the addition of other solvents and gives this agent a much longer shelf life after the container is opened.95
Etomidate Etomidate is a potent short-acting hypnotic induction and maintenance agent (see Table 31-2). It has a rapid onset, a short recovery period, and minimal cardiopulmonary effects.82 As an IV induction agent, etomidate has minimal depressive effects on the sympathetic nervous system but may stimulate sympathetic outflow at low doses.3 Etomidate is sold as a hyperosmolar solution with propylene glycol and can cause hemolysis, thrombophlebitis, and pain on injection. An aqueous formulation has been developed, which may be safer than the current commercial product.82 It is usually administered IV, but in rats and mice it has been shown to be readily absorbed IP.20 In order to reduce the impact of the hyperosmolarity, IV crystalloid fluids are given simultaneously. The larynx is often reactive with this drug, so topical laryngeal lidocaine is helpful.
Constant-Rate Infusions Intravenous constant-rate infusions (CRIs) of anesthetic and analgesic drugs are being evaluated in small mammal balanced anesthesia/analgesia protocols (Table 31-3).14,37,85 Analgesic medications administered IV CRI can be titrated to effect, potentially reducing other drug doses within the anesthetic protocol. Disadvantages to the use of CRIs in small mammals are the need for patient IV access and for a syringe pump to accurately deliver low IV dosages. Plasma drug concentrations increase slowly when used as a CRI, so a loading dose of the drug is commonly given prior to initiation of the CRI. Microdose ketamine via IV CRI can be an effective analgesic (see Table 31-3). It is very useful in patients that cannot be
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals
439
Table 31-3 Injectable Drugs Administered as Constant-Rate Infusions (CRIs) Used for Perioperative and Postoperative Analgesia in Small Exotic Mammalsa Injectable Analgesics
Rabbit
Ferret
Guinea Pig
Chinchilla
Rat
Mouse
Butorphanol
Loading dose, Loading dose, Loading dose, Loading dose, Loading dose, − 0.2-0.4 0.05-0.2 0.2-0.4 mg/kg; 0.2-0.4 0.2-0.4 mg/kg; mg/kg; maintenance, mg/kg; mg/kg; maintenance, maintenance, 0.2-0.4 mg/ maintenance, maintenance, 0.2-0.4 mg/ 0.1-0.4 mg/ kg/h 0.2-0.4 mg/ 0.2-0.4 mg/ kg/h kg/h kg/h kg/h Fentanyl citrate Loading dose, Loading dose, Loading dose, Loading dose, − 1.25 µg/ kg/h Perioperative 5-10 μg/ 5-10 μg/kg IV; 5-10 μg/kg IV; 5-10 μg/ IV33 CRI kg IV; maintenance, maintenance, kg IV; maintenance, 10-30 μg/ 10-30 μg/kg/h maintenance, 10-30 μg/ kg/h IV IV 10-30 μg/ kg/h IV kg/h IV
Postoperative 1.25-5.0 μg/ analgesia kg/h
1.25-5.0 μg/ kg/h
1.25-5.0 μg/kg/h 1.25-5.0 μg/ kg/h
−
−
Ketamine Loading dose, Loading dose, Loading dose, Loading dose, − Perioperative 2-5 IV mg/kg; 2-5 IV mg/kg; 2-5 IV mg/kg; 2-5 IV mg/kg; CRI maintenance, maintenance, maintenance, maintenance, 0.3-1.2 mg/ 0.3-1.2 mg/ 0.3-1.2 mg/ 0.3-1.2 mg/ kg/h IV kg/h IV kg/h IV kg/h IV
−
Postoperative 0.1-0.4 mg/ analgesia kg/h
Oxymorphone − Perioperative CRI
0.1-0.4 mg/kg/h 0.1-0.4 mg/kg/h 0.1-0.4 mg/ kg/h
0.1-0.4 mg/kg/h −
−
0.03 mg/kg/h IV36
−
−
−
Comments Less respiratory depression than with fentanyl The authors have used up to 40 μg/kg/h in rabbits; apnea common, expect to ventilate patient Combine with ketamine CRI to reduce overall doses More useful when intubation is not possible because there is less respiratory depression than opioids given CRI Can be combined with fentanyl to reduce overall doses of both drugs −
aThese
are suggested dosages based on published information and the authors’ clinical experience. If using for postoperative analgesia only, a loading dose should be used. Gradually wean from postoperative CRI over 12-24 hours. Species and individual variation in response to a drug or combination of drugs can be uncertain, so the dosage should be adjusted depending on the clinical response of the animal.
intubated, as it has a low potential for respiratory depression at these doses. Opioid medications can also be used for CRI. All mu (μ) receptor agonists can cause respiratory depression; thus they should be used only when an airway is secure and ventilation is available.14 These have also historically been avoided in small hind-gut fermenters for fear of inducing GI stasis, but when used as low-dose IV CRIs, these drugs can be part of balanced anesthesia and analgesia protocols.14 Fentanyl citrate is a μ receptor agonist that is used as a single-agent CRI (see Table 31-3),14 or it can be combined with ketamine in the clinical setting; to date, however, there are no published studies evaluating
these drugs via this route in small mammals. Remifentanil is an ultra-short-acting μ agonist opioid, making it very suitable for CRI.14,37 It is metabolized by esterase hydrolysis in the blood and tissues, not via the liver or kidneys, and accumulation is prevented even when given at high doses over prolonged periods. Because of its very short half31-life, undesirable respiratory and cardiovascular effects are not expected to last for more than 10 to 15 minutes after discontinuation. Acute tolerance can develop with remifentanil,37,47 which may require increasing doses to maintain intraoperative analgesia. Butorphanol has also been used successfully as a CRI in the clinical setting, but
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SECTION VI General Topics
there are no published data for small exotic mammals. Anesthesia has been maintained with doses of propofol at 0.4 to 0.6 mg/kg/min, but been deaths have reported rabbits following propofol infusions.2,67,103
INHALANT ANESTHESIA Inhalant anesthetics offer rapid induction and recovery and the ability to rapidly change anesthetic depth; moreover, their use does not require an accurate determination of body weight. Very little is metabolized, reducing the impact on hepatic and renal function, and recovery is independent of either. Small mammals require approximately the same concentration of inhaled anesthetic for an equivalent amount of stimulation as other mammals. But inhalant anesthetics can induce cardiovascular and respiratory depression, which may be life-threatening. All inhalants currently in use are potent negative inotropes; therefore a dose-related decrease in cardiac output is expected, which is reflected as a decrease in blood pressure. Thus the primary function of the anesthetist using inhalants is to maintain the lightest plane of anesthesia possible, allowing for minimal cardiopulmonary depression and completion of the procedure. In some cases these goals may not be achieved by simply reducing the inhalant; balanced anesthetic/analgesic protocols or the addition of dopamine or dobutamine (only in the volumeloaded patient) may be necessary to offset inhalant-induced hypotension. Isoflurane, sevoflurane, and desflurane are the currently available inhalants. Their speed of onset and offset is largely determined by their solubility and isoflurane > sevoflurane > desflurane. Despite these physiochemical differences, the clinical differences overall may be fairly small (e.g., induction and recovery times, heart rate, and indirect blood pressure measurements were not different between sevoflurane and isoflurane in ferrets).77 All three agents appear to have many of the same dose-dependent cardiovascular effects. Sevoflurane has a less pungent odor than other inhalants and therefore is generally better tolerated during mask induction. Many practitioners induce using sevoflurane and then maintain anesthesia with the more cost-effective isoflurane.
Mask or Chamber Induction Induction with inhalants is performed with either a mask or an induction chamber. Commercially available masks for dogs and cats can be used, but appropriate-sized masks and nose cones are also commercially available or can be fashioned from syringe cases or other materials (Fig. 31-5, A). The mask should have minimal dead space and a diaphragm, so that a seal can be achieved on the animal’s nose or neck and a piece of suture can be placed around the upper incisors and through the breathing circuit for security (see Fig. 31-5, B,C). Commercially available induction chambers sized for mice to small dogs can be used, especially for very stressed patients that may resist face-mask induction. Advantages of appropriately sized chambers are less waste gas and shorter induction times. In stressed patients, the chamber is darkened by covering it with a towel. Alternatively, for induction of large or stressed patients, an animal carrier or the front of a hospital cage can be sealed with a plastic bag and inhalant anesthetic administered into the enclosure. Disadvantages of induction chambers include inability to assess the patient, gas pollution when the chamber opened, and trauma during the excitement phase of anesthesia. Pad the chamber and
minimize its size to reduce movement and trauma. Gas flow rates are adjusted to chamber size to provide an optimum rate of rise of the anesthetic concentration. The flow rate can be calculated using the formula: f = V/t ln (S/(S-C), where f = the O2 flow rate, V = chamber volume, t = time to reach the desired concentration, S = concentration being put out by the vaporizer, and C = desired concentration.58 The patient is promptly removed after the righting reflex is lost. The final phase of induction is accomplished using a mask to facilitate monitoring.
LOCAL AND REGIONAL ANESTHESIA Local anesthetics are ideal for use in preemptive analgesic combinations. Although local anesthetics provide sufficient analgesia for some procedures without the addition of another anesthetic, the stress of handling and restraint generally precludes their sole use. Local or regional nerve blocks are commonly used, as are epidural or spinal administration (see “Epidural Anesthesia/Analgesia,” below).19,27,60,84 Regional infiltration of incision lines and specific peripheral nerve blocks (i.e., brachial plexus, sciatic) are used; dental nerve blocks are very useful in performing dental extractions. Intratesticular blocks can be used for castration; the authors use 1 mg/kg of 2% lidocaine per testicle. The toxic dose of local anesthetics in small exotic mammals clinically appears to be similar to that in cats and dogs. Toxic signs may include depression, drowsiness, muscle tremors, vomiting, hypotension, and arrhythmias as well as CNS signs such as ataxia and nystagmus. Toxicity is prevented by using appropriate concentrations and volumes. For example, the commercial lidocaine (2%) solution often must be diluted to achieve a suitable injection volume while not exceeding a total dose of 4 mg/kg. Dilution can decrease duration of action and drug effectiveness, but clinical studies are lacking on dilution limits in small mammals. Bupivacaine is used conservatively because of concerns that its toxic effects may take longer to resolve. New long-acting local anesthetic formulations have been shown to produce nerve blocks for 5 days without toxic effects in rats.119 Eutectic mixture of local anesthetics (EMLA, AstraZeneca, Wilmington, DE) is a mixture of 2.5% lidocaine and 2.5% prilocaine used topically to desensitize skin for catheter placement or superficial biopsies. Reported optimal contact time requires application and occlusion with a bandage for 30 to 60 minutes. EMLA toxicity is associated with application to large or traumatized areas and prolonged contact time. Systemic uptake may occur in smaller patients if the skin is damaged during shaving. Blanching of the skin and local erythema have also been reported, in addition to the typical toxic effects of local anesthetics. Prilocaine can induce methemoglobinemia.
EPIDURAL ANESTHESIA/ANALGESIA Epidural anesthesia/analgesia can be an extremely useful adjunct to a balanced anesthesia/analgesia protocol and can significantly reduce the concentration of inhalant anesthetics for surgical procedures (Table 31-4). Anesthetic/analgesic effects are achieved with little to no systemic drug effects, thus further reducing the use of other cardiopulmonary depressant anesthetics in the protocol. Epidural anesthesia is more commonly used in the larger exotic mammals,19,60 but it can be performed routinely in smaller species as well.22,84 The lumbosacral junction site is most commonly used, and the techniques
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B
A
C Fig. 31-5 A, Appropriate-sized masks and nose cones are commercially available or can be fashioned from syringe cases or other materials for induction and to maintain anesthesia. The mask should have minimal dead space and a diaphragm so that a seal can be achieved on the animal’s nose or neck (B). A piece of suture can be placed around the upper incisors and secured between the mask and breathing circuit (B,C).
Table 31-4 Injectable Drugs Used for Epidural Anesthesia/Analgesia in Small Exotic Mammalsa Injectable Epidural Drugs Buprenorphine Bupivacaine (0.125%)
Morphine (preservative-free) aThese
Rabbit
Ferret
Guinea Pig
Chinchilla
Rat
Mouse Comments
12 μg/kg 1
12 μg/kg 1
12 μg/kg 1
12 μg/kg 1
− 1
− 1
0.1
0.1
0.1
0.1
0.1
0.1
− Concentrations of 0.125% or less minimize motor block Little evidence of systemic uptake
are suggested dosages based on published information and the authors’ clinical experience. Species and individual variation in response to a drug or combination of drugs may be uncertain, and doses should be adjusted for the needs of each patient. All doses are in milligrams per kilogram unless specified otherwise. All drugs should be diluted with preservative-free saline only. Total volumes for epidural administration should not exceed 0.33 mL/kg regardless of drug or drug combination. Opioid and local anesthetics can be administered in combination to reduce the dosage needed of each drug.
SECTION VI General Topics
442
B
A
C
D Fig. 31-6 Landmarks for epidural analgesia are the wings of the ileum bilaterally and the depression between the last lumbar and first sacral vertebrae (A). Sterile technique is used throughout. Appropriate-sized spinal or hypodermic needles are introduced with a firm rotating motion until a “pop” is felt (B). Placement is confirmed by lack of resistance to air introduced with a glass syringe (C) before analgesics are delivered (D). Only half the calculated dose is delivered if spinal fluid is identified in the needle hub.
for administration are similar to those in dogs and cats (Fig. 31-6). Morphine is the most commonly used opioid because it has a high potency and long duration of analgesic action (1824 hours), but oxymorphone and buprenorphine have similar effects and duration (see Table 31-4). Bupivacaine at concentrations ≤0.125% appear to have the least motor effect while producing good sensory block, which is important for minimizing recovery stress and potential trauma (see Table 31-4). Epidural bupivacaine in rats provided potent antinociceptive effects for only 20 to 30 minutes.84 There appears to be synergism in the epidural space between local analgesics and opioids; drug combinations reduce doses and minimize potential adverse effects of each drug. In general, the total epidural administration volume for all drugs combined should be ≤0.33 mL/kg.
ANESTHETIC MONITORING AND SUPPORTIVE CARE Effective monitoring requires minute-to-minute patient assessment. Regardless of the anesthetic used, the eyes should be well lubricated often to prevent corneal dessication and ulceration. Covering the eyes with damp gauze sponges helps maintain the moisture provided by the eye lubrication.
Withdrawal reflexes that are used in small mammals include toe pinch, stimulation of the interdigital tissue, tail pinch, and rectal pinch. The toe pinch does not always reliably measure anesthetic depth in guinea pigs, as involuntary leg movements sometimes occur under anesthesia. Slight corneal and palpebral reflexes are commonly maintained at a surgical plane of anesthesia; absence of these reflexes indicates an excessively deep plane of anesthesia. Heart rates vary widely between species. Base knowledge of expected heart rates for the species is important, but absolute trends in rate during anesthesia are of the utmost importance. The anesthetic concentration is decreased with any sudden HR reductions and supportive care is provided. Auscult the HR and record the rate and any arrhythmias. Because of the rapid HR of many small rodents, ECG complexes can be difficult to assess at the standard sweep speed of 25 mm/sec. Sweep speeds of 100 and 200 mm/sec are available on some ECG machines and can allow for more accurate assessment. Needle ECG leads are ideal for small rodents, as they provide excellent conduction without the use of gels. Alternatively, metal alligator clips attached to a hypodermic needle or a saline-soaked cotton ball or adhesive pad can be used (Fig. 31-7). Peripheral pulse quality can be subjectively assessed by changes in pulse loudness gauged with
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B
A
Fig. 31-7 Monitoring and supportive care equipment for surgery including Doppler and noninvasive blood pressure equipment, capnography, electrocardiography (ECG), esophageal/rectal temperature probes, intravenous catheters, and temperature support with pads and forced-warm-air blankets can be used even on small patients (A). ECG pads can be cut and taped to the footpads of many rodents (B).
a Doppler ultrasonic probe placed over an artery. This is quite insensitive, but a change in Doppler loudness should alert the anesthetist to reevaluate the patient. Noninvasive blood pressure monitoring can be performed using a Doppler probe, a pressure cuff, and a sphygmomanometer (see Fig. 31-7). Oscillometric devices can be unreliable in the small hypotensive or hypothermic patient with a high heart rate. In rabbits, oscillometric readings from the forelimb correlated well with arterial blood pressures at low and normal pressures, but hind-limb readings did not.125 The size of the blood pressure cuff should approximate 40% of the limb circumference. Cuffs that are too large can give falsely decreased pressures. The cuff is placed over the radius/ulna, humerus, tibia, or femur, or a tail cuff can be placed over the base of the tail. The Doppler probe is placed between the carpal/tarsal pad and pads of the feet, above the carpus on the ventromedial aspect, or on the ventral surface of the tail over the ventral tail artery; shaving of these areas is sometimes necessary. The pressures determined with a Doppler and sphygmomanometer are taken as systolic pressures, whereas oscillometric devices can provide systolic, diastolic, and mean arterial pressures. Normal systolic blood pressure measurements obtained with the Doppler range from 90 to 120 mm Hg. Perfusion is assessed by evaluating the color and capillary refill time of the oral, rectal, or vaginal mucous membranes, femoral pulse quality, heart rate, and blood pressure. Fluid types, uses, and volume calculations are provided in Chapter 38. Respiratory rate and character should be monitored closely. Visual monitoring of the respiratory rate can be complicated by surgical drapes, especially in small rodents. Ventilation is monitored by watching the frequency and degree of movement of the chest or movement of the reservoir bag on the anesthesia machine. Small reservoir bags, such as balloons, can be used to visualize small movements. High respiratory frequency can be misleading and does not mean that the patient is necessarily light and hyperventilating. High respiratory rates can be associated with small tidal volumes, resulting in more dead-space ventilation than pulmonary ventilation. Lighten the plane of anesthesia and institute manual ventilation for any period of apnea greater than 10 to 20 seconds. Respiratory monitors that are triggered by a thermistor in the airway function well in patients weighing more than 500 g, but they may not respond to small respiratory movements and lower tidal volumes in animals weighing less than 200 g. In general, normal ventilation
produces little to no noise. Whistling, wheezing, or harsh sounds may indicate obstruction. Both injectable and inhalant anesthetics cause dose-dependent respiratory depression; this and the increased resistance to breathing imposed by the endotracheal tube provide compelling reasons to ventilate these animals. Sick or debilitated patients may not be able to accommodate these physiologic changes and thus require intermittent positive-pressure ventilation (IPPV). The authors recommend initial inspiratory pressures of 5 to 10 cm H2O and rates of 6 to 10 breaths per minute. Thoracic excursions are assessed and the tidal volume is adjusted to achieve appropriate thoracoabdominal expansion. In general, the average tidal volume for many species is 10 to 15 mL/kg; however there is variation. Inspiration should take 1 to 1.5 seconds. If a mechanical ventilator is not available, hand ventilation delivered from the reservoir bag provides assisted ventilation. End-tidal carbon dioxide (PETco2) monitoring using a capnograph is useful to assess ventilation (see Fig. 31-7). Mainstream capnographs have a sampling device that reads directly from the gas in the airway, whereas sidestream capnographs pull gas along a tube to the measuring device in the machine. Mainstream capnographs may add an unacceptably large dead space in small patients (<500 g), and the weight of the detector on the end of the endotracheal tube may predispose to inadvertent extubation. With sidestream sampling, the port should be located inside the endotracheal tube; but it is important that this not take up excessive volume and lead to an increased resistance to breathing. Sidestream capnographs draw samples at a rate of 50 to 200 mL/min. Instruments with a sample rate of 50 mL/min provide more accurate measurements in small patients.96 Evaluation of the capnograph tracing can provide important information regarding trends in PETco2. Pulse oximetry (SpO2) is used in many species to evaluate arterial hemoglobin oxygen saturation, but the high normal heart rates in these species may exceed the upper limits of standard monitors. Pulse oximeters that can detect high heart rates and low signal strengths are now available. SpO2 probes are placed on the tongue, ear, toe, tail, or vaginal/rectal mucous membranes. Smaller probes are available through laboratory animal supply companies. An angled probe placed into the mouth has proven reliable.25 Since many anesthetized animals are breathing high concentrations of oxygen, the usefulness of
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SpO2 is very low unless the procedure is expected to cause desaturation (e.g., thoracotomy). To reduce potential anesthetic-associated morbidity and mortality, it is mandatory to monitor core body temperature and to provide thermal support (see Fig. 31-7). Most anesthetics depress thermoregulatory function, and this is exacerbated in small patients prone to heat loss because of their high body surface-to-volume ratios. Hypothermia certainly depresses cardiopulmonary function, and arrhythmias and disturbances of coagulation can occur with deep hypothermia in mammals (<86°F; <30°C). Hypothermia decreases the anesthetic requirement and metabolism and prolongs recovery. Supplemental heat sources should be used regardless of the length of the procedure. Core body temperature is most accurately monitored with an esophageal temperature probe, but rectal temperatures can also be obtained if necessary. Most heat is lost via radiation, so the most effective technique to prevent heat loss is to minimize the temperature gradient between the patient and the room. This can be done by increasing room temperature; insulating the patient with clear plastic drapes, bubble wrap, or foil; and wrapping nonsurgical fields. Surgical time and the time the body cavities are open are minimized, as are the time for hair clipping and alcohol/water use for skin preparation. Latex gloves, empty fluid bags, or plastic bottles can be filled with water, warmed in a hot-water bath, and placed next to the patient. Water bottles are wrapped in a towel to prevent burns. Circulating-warm-water blankets insignificantly diminish the rate of heat loss in small patients. Radiant heat lamps are effective at maintaining core body temperature, but the optimal distance between the heat source and the patient differs with patient size, heat lamp strength, and the heat setting. Forced-air warmers are more effective at minimizing hypothermia during anesthesia than other methods and are particularly effective when set up to have warm air rising around the patient either by wrapping the patient in the blanket or by using a perforated table (see Fig. 31-7). Lubricate the eyes well to prevent corneal ulceration due to drying from the warm air. Warm the replacement fluids prior to administration. Heated, humidified anesthetic gases also help to reduce or prevent the development of hypothermia. One of the most important ways in which to reduce the potential for complications during anesthesia and recovery is
to minimize anesthetic time. Postprocedural recovery should have the patient connected to the circuit or on a face mask with 100% O2 for as long as possible in a quiet area. Animals wet from the procedure are gently dried with a loose towel or with warm air. Always monitor the character of respiration during towel restraint to ensure that chest movement is not impaired. Once recovery is considered to be of minimum potential for self-trauma, place the patient in a warm, quiet, darkened, welloxygenated environment for continued recovery. Respiratory depression can continue into the postanesthetic period, so supplemental oxygen should be continued. Use thermal support in the recovery areas, but do so with caution, because many small mammals are susceptible to heat stress. Chinchillas are particularly prone to be stressed by temperatures greater than 75°F (24°C) and should be monitored closely. Food and water are provided as soon as the patient has recovered to a point of minimal to no ataxia, and the patient is encouraged to eat. The GI transit times are very rapid for virtually all small mammals; therefore carefully consider the patient’s nutritional plane in advance. To minimize the potential for GI stasis after the anesthetic period, administer nutritional support to small mammals exhibiting anorexia of concerning duration. Replacement fluid therapy plays a role in nutritional support, as the GI tract must be hydrated to facilitate motility and function. If IV fluids were not provided during surgery, administer warm SC fluids at this time.
EMERGENCIES Emergencies during anesthesia should be anticipated and planned for in advance. Most are averted by careful monitoring of HR, RR, blood pressure, and body temperature. Endotracheal tubes, oxygen, IV catheters and materials for securing them, ventilatory support, and emergency drugs should be close by and ready for use. Emergency drug doses should be calculated and predrawn before induction (Table 31-5). Treat respiratory depression and arrest by turning off the anesthetic machine and controlling ventilation. The standard approach of maintaining airway, breathing, and circulation (ABC) should be followed. Cardiac massage can be performed and in many of these small species, in which the heart is palpable through the thoracic wall.
Table 31-5 Emergency and Cardiopulmonary Resuscitative/Supportive Drug Dosages Commonly Used in Small Exotic Mammalsa Drug
Dosea
Route of Administration
Indications
Atropine
0.04-0.1
SC, IM, IV, IO, IT
Diazepam Dobutamine Dopamine Doxapram Epinephrine (1:1000) Glycopyrrolate
To effect 5 μg/kg/min 5-20 μg/kg/min 1-10 0.01-0.1 0.01-0.05
IV IV CRI IV CRI IV, sublingual IM, IV, IO, IT IM, IV
Lidocaine Midazolam
1-2 0.1-2.0
IV IM, IV
In all small animals except the rabbit where much higher doses are required (0.2-1.0); bradycardia, CPR Seizures; slow delivery due to propylene glycol carrier Nonhypovolemic hypotension during surgery Nonhypovolemic hypotension during surgery Ineffective during cardiac arrest or severe hypoxia Cardiac arrest; dilute to 1:10,000 before IV use Bradycardia; slower onset than atropine but may provide longer benefit Antiarrhythmic Seizures, sedation
CPR, cardiopulmonary resuscitation. aNote that these are suggested doses based on the authors’ clinical experience. Species and individual variation in response to a given drug can be uncertain, so adjust the dose based on clinical response. All doses are in milligrams per kilogram unless specified otherwise.
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals If this is not immediately successful, then administration of epinephrine IV, transpulmonary, or intracardiac is recommended (in that order of preference). Treat severe bradycardia, assuming that it is due to increased parasympathetic tone, with atropine.
SMALL MAMMAL ANALGESIA The recognition and alleviation of pain in small mammals is an essential component of every animal’s evaluation. Recent surveys evaluating veterinarians’ use of analgesics in companion exotics have shown that many small mammals are still undertreated and are less likely to receive analgesics than are dogs or cats.76,118 For example, postoperative analgesia was given to 50% to 70% of dogs and cats but to only 21% of rabbits and rodents after the same procedures.76 Recognition of pain (see Chapter 39) is critical for providing timely selection of analgesia and pain relief. Human pain is mostly assessed by verbal statements, whereas in animals pain is assessed by observed behaviors. Pain assessment is complex because it requires consideration of differences in age, gender, species, breed, strain, individual behaviors, and environment. It must also take into consideration the different types and sources of pain, such as acute versus chronic pain and visceral versus somatic pain. To assess pain in the small mammal, become familiar with the normal and abnormal behaviors of each species as well as those of the individual. Many clinical signs have been associated with pain, including change in temperament (either aggressive or passive), restlessness, reduction in mobility or a reluctance to stand, lethargy, hunched appearance, constipation/GI stasis, increased respiratory rate, and lameness. A reduction in food consumption may occur with animals in pain. Monitoring body weight in exotic animals fed ad libitum is extremely important because of their small body size, and in some, such as the rabbit and guinea pig, it may help to recognize disturbances in GI motility and function before these become critical. Rabbits, ferrets, and some rodents will often grind their teeth with abdominal pain. As with other species, exotic animals with abdominal pain may exhibit abdominal tensing with a “tucked up” or “hunched” appearance to the abdomen. Animals may avoid a painful area of the body, or they may bite or chew at the site. Decreased grooming in small mammals may be exhibited by a dull or greasy appearance or, conversely, the animal may overgroom a painful site. Porphyrin staining is a nonspecific clinical sign of stress in rodents, but it can alert the clinician that the animal may be in pain. Some species respond to strong pain stimuli with tonic immobility. Because there are few reliable and consistent indicators of pain in exotic species, there is tremendous room for error in our assessment of their pain. To provide some standards for determining whether to provide analgesics, a set of universal questions is used to determine whether a patient is in pain: 1. Would the lesion or procedure be painful to any species? 2. Is the lesion or procedure damaging to tissues in any species? 3. Does the patient display any abnormal behavioral responses? If the answer to any of these questions is yes, we must assume that the patient is in pain and that an analgesic plan must be developed. Ideally the use of drugs in any species is based on knowledge of the drug’s pharmacokinetics and pharmacodynamics. Pharmacokinetic predictions are useful if the plasma concentration is reflective of the effect site activity, but this is often not the case (e.g., buprenorphine and most of the nonsteroidal anti-inflammatory drugs [NSAIDs]). Pharmacodynamic
445
studies provide more clinically relevant information about dose intervals and route of administration but are often carried out in the laboratory setting with healthy animals. There is a paucity of information for many of the drugs used clinically in small mammals; thus doses and dose intervals are often extrapolated from experience in other species. Therefore published doses and dosing frequencies should be critically evaluated for each patient prior to clinical use. Balanced analgesia should be considered for every patient, as combinations of drugs acting at different points in the nociceptive system may be more effective and less toxic than one drug given alone. For example, synergy has been demonstrated in laboratory animals between opioids, which act mainly in the CNS, and NSAIDs, which act mainly peripherally to decrease inflammation and peripheral sensitization. “Preemptive analgesia” with opiates, NSAIDs, and/or local anesthetics administered before a painful procedure can block noxious sensory stimuli from onward transmission to the CNS, thus reducing the overall potential for pain and inflammation.
OPIOIDS Opioids are most commonly used for moderate to severe pain, such as fractures and traumatic or surgical pain. The most common adverse effects reported with opioids are respiratory or cardiac depression and constipation/GI dysmotility. Changes in GI motility are concerning but can be overcome by providing excellent hydration and assisted feeding during drug administration. Most opioids can be reversed with antagonists, but this will also terminate analgesia if the antagonist crosses the bloodbrain barrier. Opioid antagonists that do not cross the bloodbrain barrier can be used to minimize the effects on the GI tract without reversing analgesia. Methylnaltrexone has been investigated in horses and humans and has been shown to minimize GI effects associated with μ agonists.8 Pure antagonists such as naloxone and naltrexone at high doses are most effective. These antagonists compete for the receptor but have minimal effects at the receptor themselves. They reverse both μ and kappa (κ) receptor agonists but will reduce or eliminate any analgesic effect of the agonist. Nalbuphine is a κ agonist; it can be used to reverse μ agonists and has some residual analgesic effect via the κ receptors (Table 31-6). Butorphanol can be used in the same way, but it is a much weaker antagonist and may increase sedation. Buprenorphine has a very strong affinity for μ receptors and is difficult to reverse. Antagonists should be given carefully, preferably titrating the dose slowly IV to the desired effect. The effects of opioids can be variable between species and within species at the same dose. Most opioids are used parenterally because of their poor oral bioavailability associated with the first-pass effect. Buprenorphine at 8 to 10 times the parenteral dose has been incorporated into gelatin and administered orally to rats.27 Most parenteral routes of administration are effective and the lipid solubility of some opioids leads to routes not normally effective with other drugs. Oral transmucosal buprenorphine has been used successfully in cats, and anecdotal use via this route in ferrets has been reported.65 Transdermal fentanyl and buprenorphine are available in some countries, but dosing studies in small mammals are limited or have not been performed. Morphine, hydromorphone, and oxymorphone are μ receptor agonists with similar durations of action (see Table 31-6). Both hydromorphone and oxymorphone have been used by the
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Table 31-6 Analgesic Drug Dosages Used in Small Exotic Mammalsa Drug
Rabbit
Ferret
Guinea Pig
Chinchilla
Rat
Mouse
Comments
NSAIDs Carprofen
1.5-5.0 PO, 2-5 PO, SC 2-5 PO, SC SC q12-24h q12-24h q12-24h 1-3 IM 1-3 SC 1-2 SC, IM q12-24h26 or IM q12-24h q12-24h26 0.1-0.3 0.1-0.3 PO, 0.1-0.3 PO, PO, SC SC q24h SC q24h q24h114,115
2-5 PO, SC 2-5 PO, SC 2-5 PO, SC q12-24h q12-24h30,99,102 q12-24h 1-2 SC, IM 2-5 SC, IM 2-5 SC, IM q12-24h q12-24h30,99 q12-24h
−
0.1-0.3 PO, SC q24h
−
Buprenorphine
0.01-0.05 SC, IM, IV q6-12h29,62
0.01-0.03 0.01-0.05 SC, IM, IV SC, IM, IV q6-12h q6-12h26
Butorphanol
0.2-2.0 SC, IM, IV q2-4 h50,94,115
0.05-0.4 SC, IM, IV q2-4h
0.2-0.5 IM, IV q4h
Fentanyl citrate − Hydromorphone 0.05-0.2 SC, IM q6-8h
− 0.1-0.2 SC, IM q6-8h
− 0.2-0.5 SC, IM q6-8h
Morphine
0.5-2.0 SC, IM q4h
0.2-2.0 SC, IM q2-4h
2-5 SC, IM q2-4h28
Nalbuphine HCl Naloxone
1-2 IM, IV q2-4h 0.01-0.1 SC, IM, IV
1-2 IM q2-4h26
Oxymorphone
0.05-0.2 SC, IM q6-8h
0.5-1.5 IM, IV q2-4h 0.01-0.1 SC, IM, IV 0.05-0.2 SC, IM q6-8h
0.01-0.05 0.05-0.1 SC, IM 0.05-0.1 SC − SC, q6-12h26 q6-12h62,102,113 IM, IV q6-12h 1-2 SC, IP Lower doses 0.2-2.0 IM, 1-2 SC, IM, IV q2-4h26 used in preIV q2-4h q2-4h26 medication combinations − − 0.025-0.223 0.2-0.5 0.2-0.5 SC q6-8h 0.2-4.0 SC SC, IM q6-8h q6-8h 2-5 SC, IM 2-5 SC, IM q4h32 2-10 SC, IM Most commonly q4h q4h32 used as a single dose pre-operatively − 1-2 SC, IM 2-4 SC, IM − q2-4h26 q2-4h26 0.01-0.1 SC, 0.01-0.1 0.01-0.1 Opioid IM, IV25 SC, IM, SC, IM, antagonist IV, IP IV54 − 0.2-0.5 1.2-1.5 SC 0.2-4.0 SC, IM SC13 q6-8h
Ketoprofen Meloxicam
0.5-2.0 PO, SC q24h101,106
1-5 PO, SC q24h105 1-5 IP105
−
Opioids
0.01-0.1 SC, IM, IV 0.2-0.5 SC, IM q6-8h
Other medications Tramadol
2-5 PO q4-8h
−
−
−
5-20 PO, SC, IV,16 5-40 SC, IP26,41 IP24,26
Little is known about this drug to date in small mammals; higher dosing frequency necessary to maintain therapeutic concentrations of other species
aThese
are suggested dosages based upon published information and the authors’ clinical experience. Species and individual variation in response to a given drug can be uncertain, so the dosage should be adjusted depending upon the clinical response of the animal. Although there is no good data available for hamsters and gerbils, extrapolation from rat dosages clinically appears to be appropriate. Many published dosages are based upon laboratory animal data; differences in pet breeds of these animals are unknown. Doses are in milligram per kilogram unless specified otherwise.
authors in ferrets and anecdotally by others in rabbits as primary analgesics for the treatment of moderate to severe pain, but they are also useful for preemptive analgesia and postoperative pain. In ferrets, these drugs cause profound sedation, making assessment of analgesia difficult.65 Fentanyl citrate has
an effect for only approximately 30 minutes after a single IV injection and is thus most commonly used as a CRI during the perianesthetic and postoperative periods (see “Constant-Rate Infusions,” above, and Table 31-3).14 As with all μ receptor agonists, fentanyl can cause respiratory depression; thus a secure
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals airway and ability to ventilate are required when using it as a CRI. Systemic uptake of transdermal fentanyl in rabbits with the 25-mcg/h patch was highest, with the longest duration of activity (3 days) when the hair was clipped at the patch site. But the rabbits were more sedated and had shorter plasma concentrations when the hair was chemically depilated, and no systemic concentrations were identified when hair was present at the patch site.31 Although extrapolated effective therapeutic plasma concentrations were obtained, loss of body weight occurred in this study. If this method of analgesia is to be used, appetite and fecal/urine output must be monitored very closely. Remifentanil is an ultra-short-acting μ receptor agonist, making it very suitable for CRI (see “Constant-Rate Infusions,” above). Buprenorphine is a slow-onset, long-acting partial μ opioid (see Table 31-6).1 It is the preferred opioid in small mammals for postoperative analgesia because of its longer duration of effect.100 It can antagonize the effects of other pure μ agonists and is also described as a κ-receptor agonist and antagonist.1 Buprenorphine may exhibit a plateau or “ceiling” analgesic effect; administration of additional doses produces either detrimental effects,62 no additional analgesia, or prolonged analgesia.29,62 Analgesic effects at the same dose can also be variable among different strains of rodents.62 GI effects are the most commonly reported adverse effect with buprenorphine. “Pica behavior” may occur when rats are housed on certain types of bedding, especially sawdust or wood chips. Buprenorphine is most commonly administered SC, IM, or IV; however other routes have also been evaluated. Buprenorphine is considered safe and effective when administered at 0.01 to 0.05 mg/kg parenterally, but higher doses are sometimes necessary in smaller mammals. For example, 0.5 mg/kg provided 6 to 8 hours of analgesia in rats, while 2 mg/kg provided 3 to 5 hours of analgesia in mice.32 Buprenorphine administered PO in gelatin to rats was effective at 8 to 10 times the parenteral dose.27,91 Oral transmucosal buprenorphine is used in cats and anecdotally in ferrets. Epidural administration in rats shows consistent mild to moderate antinociception for at least 2 hours.84 Buprenorphine administered as a preanesthetic may not provide timely analgesia for surgical procedures and may prolong recoveries, so it is most commonly administered for postoperative pain.110 Butorphanol is a mixed agonist/antagonist with low intrinsic activity at the μ receptor and strong agonist activity at the κ receptor (see Table 31-6). Butorphanol is commonly used in small mammals to circumvent the potential for GI disturbances and generally does not produce dose-related respiratory depression. The half31-life of butorphanol in rabbits administered 0.5 mg/kg IV was shown to be 1.6 hours and 3.2 hours when given SC, consistent with results from other pharmacokinetic studies of this drug.94 In rabbits administered 0.4 mg/kg butorphanol SC, ketamine/medetomidine, anesthesia was prolonged, but only minor anesthetic-sparing effects were seen.50 Butorphanol (0.4 mg/kg IV) significantly reduced the MAC of isoflurane in rabbits.115 Butorphanol has also been used as a CRI in the clinical setting, although to date there are no published studies evaluating this in small exotic mammals.
TRAMADOL HYDROCHLORIDE Tramadol hydrochloride is an analgesic that has become popular recently despite minimal evidence of its efficacy (see Table 31-6). It is active at opiate, alpha-adrenergic, and serotonergic receptors.109 Tramadol is a very weak μ agonist, but
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the O-desmethyl metabolite (M1) is a much more potent agonist. The conversion to the M1 metabolite is variable among species, but it is known that it is produced in rats, mice, and rabbits.90,112,123 In the United States, only the PO formulation is available. In humans, less respiratory depression and constipation are seen with tramadol than with μ-agonist opioids, but tramadol caused a significant decrease in guinea pig intestinal motility in vitro.57 The pharmacokinetics of tramadol have been evaluated in rats34,90,127 and rabbits,112 but analgesic plasma concentrations have not yet been established. Clinically insignificant isoflurane-sparing effects have been shown in both rats and rabbits administered 10 and 4.4 mg/kg tramadol PO, respectively.16,21 Significant decreases in HR and transient decreases in systolic arterial pressure were identified in rabbits after a single 4.4-mg/kg IV dose.21 Results of several studies in rats have shown that tramadol can be an effective analgesic for acute pain.12,41,43 In rats, tramadol provided analgesia for osteoarthritis,12 but its efficacy decreased with increased duration of pain, and its antinociceptive mechanism changed over time, which may partially explain its inconsistent efficacy in patients with chronic pain.43 Anecdotally, doses of 2 to 5 mg/kg PO have been well tolerated in rabbits and rats. In one study, there was evidence of tolerance in rats with chronic use, so dosing may need to be adjusted based on individual needs.116 While this analgesic holds great promise for use in small mammals, much work is still needed to evaluate its appropriate dosing, efficacy, and safety in different species.
NONSTEROIDAL ANTI-INFLAMMATORY DRUGS Nonsteroidal anti-inflammatory drugs (NSAIDs) are the most commonly prescribed analgesics in veterinary medicine. Reported adverse effects include GI ulceration and renal damage.108 Cyclo-oxygenase-2 (COX-2) has been shown to be constitutively expressed in the kidney in all mammalian species studied and is highly regulated in response to alterations in intravascular volume. In conditions of relative intravascular volume depletion and/or renal hypoperfusion—such as dehydration, hemorrhage, hemodynamic compromise, heart failure, and renal disease—interference with COX-2 activity can have deleterious effects on maintenance of renal blood flow and glomerular filtration rate. NSAIDs are generally well absorbed after PO, SC, and IM administration. All NSAIDs are highly proteinbound, some greater than 99%, which explains their extended therapeutic activity in inflamed tissues. Plasma half-life therefore may not be the most important factor in determining NSAID dosing intervals. Ketoprofen is a potent nonselective COX inhibitor (see Table 31-6). Ketoprofen is used parenterally in exotic patients because of limited oral pharmacokinetic data and difficulty in accurately dosing oral formulations. Ketoprofen administered at 5 mg/kg SC preoperatively in rats undergoing exploratory laparotomy showed an effect for at least 5 hours, but this dose was not effective PO, suggesting that higher doses may be needed for PO administration.30 Ketoprofen is available as a 2.5% topical gel in some countries, but systemic uptake may occur if it is ingested. Carprofen is available in both injectable and PO forms (see Table 31-6). It is a weak COX inhibitor and may achieve its therapeutic effects, at least partially, through other pathways. The halflife of carprofen varies considerably among mammalian species, which may affect dosing intervals. Very little work has been done
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to date evaluating this drug in exotic animal species.30,46,101 Carprofen administered at 5 mg/kg SC provided postlaparotomy analgesia for at least 5 hours,30 but an equivalent effect was found using only 2.5 mg/kg in another study.101 Postoperative food and water consumption were improved when rats were administered 5 mg/kg SC but not PO, suggesting that higher doses may be necessary with PO dosing.30 Unlike most species studied, the S (+) carprofen enantiomer predominates in rabbit plasma, which may have significant therapeutic and safety implications.46 The authors have used 1 to 4 mg/kg PO, SC, and IM q12-24h shortterm (<7 days) in many small mammal species. Meloxicam is a COX-2 selective oxicam NSAID (see Table 31-6). In recent years, meloxicam has become the most widely used anti-inflammatory medication in pet exotic animal practice. It is currently available as an oral suspension and an injectable form in the United States. Meloxicam at a dose of 1 mg/kg SC reduced acute postlaparotomy pain behaviors in rats, but a dose of 0.5 mg/kg SC was not effective.101 The pharmacokinetics of single or repeated PO doses (daily for 5 days) of either 0.3 or 1.5 mg/kg meloxicam in 3-month-old rabbits showed that after single oral administration of either dose, maximal plasma concentrations were achieved at 6 to 8 hours and were nearly undetectable by 24 hours. No drug accumulation in plasma was reported at either dose after 5 days, and meloxicam was rapidly eliminated after drug discontinuation.114 A separate study of the pharmacokinetics of single or repeated PO doses (daily for 10 days) of 0.3 mg/ kg meloxicam in 8-month-old rabbits identified a slight increase in plasma concentrations over time.11 It is difficult to interpret the importance of these data because of the differences in age and diet between the two studies.11 Meloxicam administered 0.5 mg/kg PO produced significant analgesia in visceral pain tests in guinea pigs.63 These studies suggest that some rodents and rabbits may need higher meloxicam doses, but clinical efficacy and safety studies are necessary to determine appropriate meloxicam analgesic doses and dosing frequency in small mammal patients. Single- and multiple-dose pharmacokinetics of tepoxalin, a dual inhibitor of cyclooxygenase and 5-lipoxygenase, have recently been published in rabbits.93 Clinical efficacy studies to evaluate safety and appropriate dosing are necessary before this drug can be recommended.
ACUPUNCTURE Acupuncture has become more popular and accepted in veterinary medicine because of a recent increase in scientifically based studies investigating its mechanism of action and documenting results. The main mechanism of action for analgesia appears to be an increased release of endogenous opioids, and the analgesic effects of acupuncture are reversible with naloxone. Many of these studies have been performed in rodents.59,75,124,126
References 1. ——Buprenorphine hydrochloride. American Hospital Formulary Service Drug Information. Bethesda, MD: Board of Directors of the American Society of Hospital Pharmacists; 2003;2061-2069. 2. Aeschbacher G, Webb AI. Propofol in rabbits. 2. Long-term anesthesia. Lab Anim Sci. 1993;43:328-335. 3. Aono H, Hirakawa M, Unruh GK, et al. Anesthetic induction agents, sympathetic nerve activity and baroreflex sensitivity: a study in rabbits comparing thiopental, propofol and etomidate. Acta Med Okayama. 2001;55:197-203.
4. Arras M, Autenried P, Rettich A, et al. Optimization of intraperitoneal injection anesthesia in mice: drugs, dosages, adverse effects, and anesthesia depth. Comp Med. 2001;51: 443-456. 5. Avsaroglu H, Versluis A, Hellebrekers LJ, et al. Strain differences in response to propofol, ketamine and medetomidine in rabbits. Vet Rec. 2003;152:300. 6. Avsaroglu H, van der Sar AS, van Lith HA, et al. Differences in response to anaesthetics and analgesics between inbred rat strains. Lab Anim. 2007;41:337-344. 7. Bateman L, Ludders JW, Gleed RD, et al. Comparison between facemask and laryngeal mask airway in rabbits during isoflurane anesthesia. Vet Anaesth Analg. 2005;32:280-288. 8. Boscan P, Van Hoogmoed LM, Farver TB, et al. Evaluation of the effects of the opioid agonist morphine on gastrointestinal tract function in horses. Am J Vet Res. 2006;67:992-997. 9. Brodbelt DC, Blissitt KJ, Hammond RA, et al. The risk of death: the confidential enquiry into perioperative small animal fatalities. Vet Anaesth Analg. 2008;35:365-373. 10. Cantwell SL. Ferret, rabbit and rodent anesthesia. Vet Clin North Am Exot Anim Pract. 2001;4:169-191. 11. Carpenter JW, Pollock CG, Koch DE, et al. Single and multiple-dose pharmacokinetics of meloxicam after oral administration to the rabbit (Oryctolagus cuniculus). J Zoo Wildl Med. 2009;40:601-606. 12. Chandran P, Pai M, Blomme EA, et al. Pharmacological modulation of movement-evoked pain in a rat model of osteoarthritis. Eur J Pharmacol. 2009;613:39-45. 13. Clark MD, Krugner-Higby L, Smith LJ, et al. Evaluation of liposome-encapsulated oxymorphone hydrochloride in mice after splenectomy. Comp Med. 2004;54:558-563. 14. Criado AB, Gomez de Segura IA. Reduction of isoflurane MAC by fentanyl or remifentanil in rats. Vet Anaesth Analg. 2003;30:250-256. 15. Czabak-Garbacz R, Cygan B, Chomicki M, et al. The influence of diazepam on the behaviour of rabbits in spontaneous conditions. Ann Univ Mariae Curie Sklodowska Med. 2002;57:264-270. 16. de Wolff MH, Leather HA, Wouters PF. Effects of tramadol on minimum alveolar concentration (MAC) of isoflurane in rats. Br J Anaesth. 1999;83:780-783. 17. Difilippo SM, Norberg PJ, Suson UD, et al. A comparison of xylazine and medetomidine in an anesthetic combination in New Zealand white rabbits. Contemp Top Lab Anim Sci. 2004;43:32-34. 18. Doerning BJ, Brammer DW, Chrisp CE, et al. Nephrotoxicity of tiletamine in New Zealand white rabbits. Lab Anim Sci. 1992;42:267-269. 19. Dollo G, Malinovsky JM, Peron A, et al. Prolongation of epidural bupivacaine effects with hyaluronic acid in rabbits. Int J Pharm. 2004;272:109-119. 20. Duysens J, Inoue M, Van Luijtelaar EL, et al. Facilitation of spike-wave activity by the hypnotic etomidate in a rat model for absence epilepsy. Int J Neurosci. 1991;57:213-217. 21. Egger CM, Souza MJ, Greenacre CB, et al. Effect of intravenous administration of tramadol hydrochloride on the minimum alveolar concentration of isoflurane in rabbits. Am J Vet Res. 2009;70:945-949. 22. Eisele PH, Kaaekuahiwi MA, Canfield DR, et al. Epidural catheter placement for testing of obstetrical analgesics in female guinea pigs. Lab Anim Sci. 1994;44:486-490. 23. El Mouedden M, Meert TF. Evaluation of pain-related behavior, bone destruction and effectiveness of fentanyl, sufentanil, and morphine in a murine model of cancer pain. Pharmacol Biochem Behav. 2005;82:109-119. 24. Erhan E, Onal A, Kocabas S, et al. Ondansetron does not block tramadol-induced analgesia in mice. Methods Find Exp Clin Pharmacol. 2005;27:629-632.
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals 25. Flecknell PA. Anaesthesia of common laboratory species: special considerations. In: Flecknell PA, ed. Laboratory animal anaesthesia. 3rd ed. London: Elsevier; 2009:181-241. 26. Flecknell PA. Analgesia and post-operative care. In: Flecknell PA, ed. Laboratory animal anaesthesia. 3rd ed. London: Academic Press, Elsevier; 2009:139-179. 27. Flecknell PA. Analgesia of small mammals. Vet Clin North Am Exot Anim Pract. 2001;4:47-56. 28. Flecknell PA. Guinea pigs. In: Meredith A, Redrobe S, eds. BSAVA manual of exotic pets. 4th ed. Gloucester: BSAVA; 2002:52-64. 29. Flecknell PA, Liles JH. Assessment of the analgesic action of opioid agonist-antagonists in the rabbit. J Assoc Vet Anaesth Gr Brit Ireland. 1990;17:24-29. 30. Flecknell PA, Orr HE, Roughan JV, et al. Comparison of the effects of oral or subcutaneous carprofen or ketoprofen in rats undergoing laparotomy. Vet Rec. 1999;144:65-67. 31. Foley PL, Henderson AL, Bissonette EA, et al. Evaluation of fentanyl transdermal patches in rabbits: blood concentrations and physiologic response. Comp Med. 2001;51:239-244. 32. Gades NM, Danneman PJ, Wixson SK, et al. The magnitude and duration of the analgesic effect of morphine, butorphanol, and buprenorphine in rats and mice. Contemp Top Lab Anim Sci. 2000;39:8-13. 33. Gaertner DJ, Hallman TM, Hankenson FC, et al. Anesthesia and analgesia for laboratory rodents. In: Fish RE, Brown MJ, Danneman PJ, et al, eds. Anesthesia and analgesia in laboratory animals. 2nd ed. San Diego: Elsevier; 2008:239-297. 34. Garrido MJ, Sayar O, Segura C, et al. Pharmacokinetic/pharmacodynamic modeling of the antinociceptive effects of (+)-tramadol in the rat: role of cytochrome P450 2D activity. J Pharmacol Exp Ther. 2003;305:710-718. 35. Ghaffari MS, Moghaddassi AP, Bokaie S. Effects of intramuscular acepromazine and diazepam on tear production in rabbits. Vet Rec. 2009;164:147-148. 36. Gillingham MB, Clark MD, Dahly EM, et al. A comparison of two opioid analgesics for relief of visceral pain induced by intestinal resection in rats. Contemp Top Lab Anim Sci. 2001;40:21-26. 37. Gomez de Segura IA, de la Vibora JB, Aguado D. Opioid tolerance blunts the reduction in the sevoflurane minimum alveolar concentration produced by remifentanil in the rat. Anesthesiology. 2009;110:1133-1138. 38. Grint NJ, Sayers IR, Cecchi R, et al. Postanaesthetic tracheal strictures in three rabbits. Lab Anim. 2006;40:301-308. 39. Grint NJ, Murison PJ. Peri-operative body temperatures in isoflurane-anaesthetized rabbits following ketaminemidazolam or ketamine-medetomidine. Vet Anaesth Analg. 2007;34:181-189. 40. Grint NJ, Murison PJ. A comparison of ketamine-midazolam and ketamine-medetomidine combinations for induction of anaesthesia in rabbits. Vet Anaesth Analg. 2008;35:113-121. 41. Guneli E, Karabay Yavasoglu NU, Apaydin S, et al. Analysis of the antinociceptive effect of systemic administration of tramadol and dexmedetomidine combination on rat models of acute and neuropathic pain. Pharmacol Biochem Behav. 2007;88:9-17. 42. Hahn N, Eisen RJ, Eisen L, et al. Ketamine-medetomidine anesthesia with atipamezole reversal: practical anesthesia for rodents under field conditions. Lab Anim (NY). 2005;34:48-51. 43. Hama A, Sagen J. Altered antinociceptive efficacy of tramadol over time in rats with painful peripheral neuropathy. Eur J Pharmacol. 2007;559:32-37. 44. Harrison PK, Tattersall JE, Gosden E. The presence of atropinesterase activity in animal plasma. Naunyn Schmiedebergs Arch Pharmacol. 2006;373:230-236. 45. Hawkins MG, Pascoe PJ. Cagebirds. In: West G, Heard D, Caulkett N, eds. Zoo animal & wildlife immobilization and anesthesia. Ames: Blackwell Publishing; 2007:269-297.
449
46. Hawkins MG, Taylor IT, Craigmill AL, et al. Enantioselective pharmacokinetics of racemic carprofen in New Zealand white rabbits. J Vet Pharmacol Ther. 2008;31:423-430. 47. Hayashida M, Fukunaga A, Hanaoka K. Detection of acute tolerance to the analgesic and nonanalgesic effects of remifentanil infusion in a rabbit model. Anesth Analg. 2003;97:1347-1352. 48. Heard DJ. Lagomorphs (rabbits, hares, and pikas). In: West G, Heard D, Caulkett N, eds. Zoo animal & wildlife immobilization and anesthesia. Ames: Blackwell Publishing; 2007:647-654. 49. Heard DJ. Rodents. In: West G, Heard D, Caulkett N, eds. Zoo animal & wildlife immobilization and anesthesia. Ames: Blackwell Publishing; 2007:655-664. 50. Hedenqvist P, Orr HE, Roughan JV, et al. Anaesthesia with ketamine/medetomidine in the rabbit: influence of route of administration and the effect of combination with butorphanol. Vet Anaesth Analg. 2002;29:14-19. 51. Hedenqvist P, Roughan JV, Flecknell PA. Effects of repeated anaesthesia with ketamine/medetomidine and of pre-anaesthetic administration of buprenorphine in rats. Lab Anim. 2000;34:207-211. 52. Hellebrekers LJ, de Boer EJ, van Zuylen MA, et al. A comparison between medetomidine-ketamine and medetomidinepropofol anaesthesia in rabbits. Lab Anim. 1997;31:58-69. 53. Henke J, Astner S, Brill T, et al. Comparative study of three intramuscular anaesthetic combinations (medetomidine/ ketamine, medetomidine/fentanyl/midazolam and xylazine/ ketamine) in rabbits. Vet Anaesth Analg. 2005;32:261-270. 54. Henke J, Baumgartner C, Roltgen I, et al. Anaesthesia with midazolam/medetomidine/fentanyl in chinchillas (Chinchilla lanigera) compared to anaesthesia with xylazine/ketamine and medetomidine/ketamine. J Vet Med A Physiol Pathol Clin Med. 2004;51:259-264. 55. Henke J, Roberts U, Otto K, et al. Clinical investigations of an i.m. combination anesthesia with fentanyl/climazolam/xylazine and postoperative i.v. antagonism with naloxone/sarmazenil/yohimbine in guinea pigs. Tierarztl Prax. 1996;24:85-87. 56. Henke J, Schneider E, Erhardt W. Medetomidine combination anaesthesia with and without antagonisation - influence on vital parameters in mongolian gerbils (Mesocricetus unguiculatus). Proceedings. Switzerland: 7th World Congress Vet Anesth, Berne. 2000:99-100. 57. Herbert MK, Weis R, Holzer P. The enantiomers of tramadol and its major metabolite inhibit peristalsis in the guinea pig small intestine via differential mechanisms. BMC Pharmacol. 2007;7:5. 58. Hodgson DS. Anesthetic concentrations in enclosed chambers using an innovative delivery device. Vet Anaesth Analg. 2007;34:99-106. 59. Huang C, Wang Y, Han JS, et al. Characteristics of electroacupuncture-induced analgesia in mice: variation with strain, frequency, intensity and opioid involvement. Brain Res. 2002;945:20-25. 60. Hughes PJ, Doherty MM, Charman WN. A rabbit model for the evaluation of epidurally administered local anaesthetic agents. Anaesth Intensive Care. 1993;21:298-303. 61. Hurley RJ, Marini RP, Avison DL, et al. Evaluation of detomidine anesthetic combinations in the rabbit. Lab Anim Sci. 1994;44:472-478. 62. Jablonski P, Howden BO. Oral buprenorphine and aspirin analgesia in rats undergoing liver transplantation. Lab Anim. 2002;36:134-143. 63. Jain NK, Kulkarni SK, Singh A. Modulation of NSAID-induced antinociceptive and anti-inflammatory effects by alpha-2 adrenoceptor agonists with gastroprotective effects. Life Sci. 2002;70:2857-2869. 64. Johnson-Delaney CA. Ferrets: anaesthesia and analgesia. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Quedgeley, Gloucester: BSAVA; 2009:245-253.
450
SECTION VI General Topics
65. Johnston MS. Clinical approaches to analgesia in ferrets and rabbits. Sem Avian Exot Pet Med. 2005;14:229-235. 66. Kazakos GM, Anagnostou T, Savvas I, et al. Use of the laryngeal mask airway in rabbits: placement and efficacy. Lab Anim. 2007;36:29-34. 67. Kemmochi M, Ichinohe T, Kaneko Y. Remifentanil decreases mandibular bone marrow blood flow during propofol or sevoflurane anesthesia in rabbits. J Oral Maxillofac Surg. 2009;67:1245-1250. 68. Kim MS, Jeong SM, Park JH, et al. Reversal of medetomidineketamine combination anesthesia in rabbits by atipamezole. Exp Anim. 2004;53:423-428. 69. Ko JC, Heaton-Jones TG, Nicklin CF. Evaluation of the sedative and cardiorespiratory effects of medetomidine, medetomidine-butorphanol, medetomidine-ketamine, and medetomidine-butorphanol-ketamine in ferrets. J Am Anim Hosp Assoc. 1997;33:438-448. 70. Ko J, Marini RP. Anesthesia and analgesia in ferrets. In: Fish RE, Brown MJ, Danneman PJ, et al, eds. Anesthesia and analgesia in laboratory animals. San Diego: Elsevier; 2008:443-456. 71. Ko JC, Nicklin CF, Heaton-Jones TG, et al. Comparison of sedative and cardiorespiratory effects of diazepam, acepromazine, and xylazine in ferrets. J Am Anim Hosp Assoc. 1998;34:234-241. 72. Ko JC, Smith TA, Kuo WC, et al. Comparison of anesthetic and cardiorespiratory effects of diazepam-butorphanol- ketamine, acepromazine-butorphanol-ketamine, and xylazine-butorphanol-ketamine in ferrets. J Am Anim Hosp Assoc. 1998;34:407-416. 73. Ko JC, Thurmon JC, Tranquilli WJ, et al. A comparison of medetomidine-propofol and medetomidine-midazolam-propofol anesthesia in rabbits. Lab Anim Sci. 1992;42:503-507. 74. Ko JC, Villarreal A, Kuo WC, et al. Evaluation of sedative and cardiorespiratory effects of diazepam-butorphanol, acepromazine-butorphanol, and xylazine-butorphanol in ferrets. J Am Anim Hosp Assoc. 1998;34:242-250. 75. Koo ST, Park YI, Lim KS, et al. Acupuncture analgesia in a new rat model of ankle sprain pain. Pain. 2002;99:423-431. 76. Lascelles BDX, Capner CA, Waterman-Pearson AB. Current British veterinary attitudes to perioperative analgesia for cats and small mammals. Vet Rec. 2000;145:601-604. 77. Lawson AK, Lichtenberger M, Day T, et al. Comparison of sevoflurane and isoflurane in domestic ferrets (Mustela putorius furo). Vet Ther. 2006;7:207-212. 78. Marini RP, Callahan RJ, Jackson LR, et al. Distribution of technetium 99m-labeled red blood cells during isoflurane anesthesia in ferrets. Am J Vet Res. 1997;58:781-785. 79. Marini RP, Jackson LR, Esteves MI, et al. Effect of isoflurane on hematologic variables in ferrets. Am J Vet Res. 1994;55:1479-1483. 80. Marini RP, Li X, Harpster NK, et al. Cardiovascular pathology possibly associated with ketamine/xylazine anesthesia in Dutch belted rabbits. Lab Anim Sci. 1999;49:153-160. 81. Martin-Cancho MF, Lima JR, Luis L, et al. Relationship of bispectral index values, haemodynamic changes and recovery times during sevoflurane or propofol anaesthesia in rabbits. Lab Anim. 2006;40:28-42. 82. McIntosh MP, Narita H, Kameyama Y, et al. Evaluation of mean arterial blood pressure, heart rate, and sympathetic nerve activity in rabbits after administration of two formulations of etomidate. Vet Anaesth Analg. 2007;34:149-156. 83. McKune CM, Brosnan RJ, Dark MJ, et al. Safety and efficacy of intramuscular propofol administration in rats. Vet Anaesth Analg. 2008;35:495-500. 84. Morimoto K, Nishimura R, Matsunaga S, et al. Epidural analgesia with a combination of bupivacaine and buprenorphine in rats. J Vet Med A Physiol Pathol Clin Med. 2001; 48:303-312.
85. Mustola ST, Rorarius MG, Baer GA, et al. Potency of propofol, thiopentone and ketamine at various endpoints in New Zealand white rabbits. Lab Anim. 2000;34:36-45. 86. Olson ME, Vizzutti D, Morck DW, et al. The parasympatholytic effects of atropine sulfate and glycopyrrolate in rats and rabbits. Can J Vet Res. 1994;58:254-258. 87. Ordodi VL, Mic FA, Mic AA, et al. A simple device for intubation of rats. Lab Anim. 2005;34:37-39. 88. Orr HE, Roughan JV, Flecknell PA. Assessment of ketamine and medetomidine anaesthesia in the domestic rabbit. Vet Anaesth Analg. 2005;32:271-279. 89. Otto C, Crowe D. Intraosseous resuscitation techniques and applications. In: Kirk R, Bonagura J, eds. Kirk’s current veterinary therapy XI: small animal practice. Philadelphia: WB Saunders; 1992:107-112. 90. Parasrampuria R, Vuppugalla R, Elliott K, et al. Routedependent stereoselective pharmacokinetics of tramadol and its active O-demethylated metabolite in rats. Chirality. 2007;19:190-196. 91. Pekow C. Buprenorphine Jell-O recipe for rodent analgesia. Synapse. 1992;25:35-36. 92. Phaneuf LR, Barker S, Groleau MA, et al. Tracheal injury after endotracheal intubation and anesthesia in rabbits. J Am Assoc Lab Anim Sci. 2006;45:67-72. 93. Pollock CG, Carpenter JW, Koch DE, et al. Single and multiple-dose pharmacokinetics of tepoxalin and its active metabolite after oral administration to rabbits (Oryctolagus cuniculus). J Vet Pharmacol Ther. 2008;31:171-174. 94. Portnoy LG, Hustead DR. Pharmacokinetics of butorphanol tartrate in rabbits. Am J Vet Res. 1992;53:541-543. 95. Ravenelle F, Vachon P, Rigby-Jones AE, et al. Anaesthetic effects of propofol polymeric micelle: a novel water soluble propofol formulation. Br J Anaesth. 2008;101:186-193. 96. Richardson C, Flecknell PA. Rodents: anaesthesia and analgesia. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Quedgeley, Gloucester: BSAVA; 2009:63-72. 97. Richardson CA, Flecknell PA. Anaesthesia and post-operative analgesia following experimental surgery in laboratory rodents: are we making progress? Altern Lab Anim. 2005;33:119-127. 98. Roberts U, Henke J, Brill R, et al. Fully antagonizable anaesthesia of the guinea pig. Part I: experimental investigations. In: Schmidt-Oechtering G, Alef M, eds. Neue aspekte der veterinaranasthesie und intensivtherapie. Berlin: Verlag Paul Parey; 1993:295-296. 99. Roughan JV, Flecknell PA. Behavioural effects of laparotomy and analgesic effects of ketoprofen and carprofen in rats. Pain. 2001;90:65-74. 100. Roughan JV, Flecknell PA. Buprenorphine: a reappraisal of its antinociceptive effects and therapeutic use in alleviating postoperative pain in animals. Lab Anim. 2002;36:322-343. 101. Roughan JV, Flecknell PA. Evaluation of a short duration behaviour-based post-operative pain scoring system in rats. Eur J Pain. 2003;7:397-406. 102. Roughan JV, Flecknell PA. Behaviour-based assessment of the duration of laparotomy-induced abdominal pain and the analgesic effects of carprofen and buprenorphine in rats. Behav Pharmacol. 2004;15:461-472. 103. Rozanska D. Evaluation of medetomidine-midazolam-atropine (MeMiA) anesthesia maintained with propofol infusion in New Zealand white rabbits. Pol J Vet Sci. 2009;12:209-216. 104. Saha DC, Saha AC, Malik G, et al. Comparison of cardiovascular effects of tiletamine-zolazepam, pentobarbital, and ketamine-xylazine in male rats. J Am Assoc Lab Anim Sci. 2007;46:74-80. 105. Santos AR, Vedana EM, De Freitas GA. Antinociceptive effect of meloxicam, in neurogenic and inflammatory nociceptive models in mice. Inflamm Res. 1998;47:302-307.
CHAPTER 31 Anesthesia, Analgesia, and Sedation of Small Mammals 106. Santos M, Kunkar V, Garcia-Iturralde P, et al. Meloxicam, a specific COX-2 inhibitor, does not enhance the isoflurane minimum alveolar concentration reduction produced by morphine in the rat. Anesth Analg. 2004;98:359-363. 107. Schernthaner A, Lendl C, Busch R, et al. [Clinical evaluation of three medetomidine–midazolam–ketamine combinations for neutering of ferrets (Mustela putorius furo)]. Berl Munch Tierarztl Wochenschr. 2008;121:1-10. 108. Schmassmann A, Zoidl G, Peskar BM, et al. Role of the different isoforms of cyclooxygenase and nitric oxide synthase during gastric ulcer healing in cyclooxygenase-1 and -2 knockout mice. Am J Physiol Gastrointest Liver Physiol. 2006;290:G747-G756. 109. Scott LJ, Perry CM. Tramadol: a review of its use in perioperative pain. Drugs. 2000;60:139-176. 110. Sharp J, Zammit T, Azar T, et al. Recovery of male rats from major abdominal surgery after treatment with various analgesics. Contemp Top Lab Anim Sci. 2003;42:22-27. 111. Smith JC, Robertson LD, Auhll A, et al. Endotracheal tubes versus laryngeal mask airways in rabbit inhalation anesthesia: ease of use and waste gas emissions. Contemp Top Lab Anim Sci. 2004;43:22-25. 112. Souza MJ, Greenacre CB, Cox SK. Pharmacokinetics of orally administered tramadol in domestic rabbits (Oryctolagus cuniculus). Am J Vet Res. 2008;69:979-982. 113. Stewart ASL, Martin WJ. Evaluation of postoperative analgesia in a rat model of incisional pain. Contemp Top Lab Anim Sci. 2003;42:28-34. 114. Turner PV, Chen HC, Taylor WM. Pharmacokinetics of meloxicam in rabbits after single and repeat oral dosing. Comp Med. 2006;56:63-67. 115. Turner PV, Kerr CL, Healy AJ, et al. Effect of meloxicam and butorphanol on minimum alveolar concentration of isoflurane in rabbits. Am J Vet Res. 2006;67:770-774. 116. Valle M, Garrido MJ, Pavon JM, et al. Pharmacokinetic-pharmacodynamic modeling of the antinociceptive effects of main active metabolites of tramadol, (+)-O-desmethyltramadol and (−)-O-desmethyltramadol, in rats. J Pharmacol Exp Ther. 2000;293:646-653.
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117. Vergari A, Gunnella B, Rodola F, et al. A new method of orotracheal intubation in mice. Eur Rev Med Pharmacol Sci. 2004;8:103-106. 118. Wagner AE, Hellyer PW. Survey of anesthesia techniques and concerns in private veterinary practice. J Am Vet Med Assoc. 2000;217:1652-1657. 119. Wang CF, Djalali AG, Gandhi A, et al. An absorbable local anesthetic matrix provides several days of functional sciatic nerve blockade. Anesth Analg. 2009;108:1027-1033. 120. Welberg LA, Kinkead B, Thrivikraman K, et al. Ketamine-xylazine-acepromazine anesthesia and postoperative recovery in rats. J Am Assoc Lab Anim Sci. 2006;45:13-20. 121. Wolfson B, Hetrick WD, Lake CL, et al. Anesthetic indices– further data. Anesthesiology. 1978;48:187-190. 122. Wood AK, Klide AM, Pickup S, et al. Prolonged general anesthesia in MR studies of rats. Acad Radiol. 2001;8:1136-1140. 123. Wu WN, McKown LA, Codd EE, et al. Metabolism of two analgesic agents, tramadol-n-oxide and tramadol, in specific pathogen-free and axenic mice. Xenobiotica. 2006;36:551-565. 124. Yang J, Liu WY, Song CY, et al. Through central arginine vasopressin, not oxytocin and endogenous opiate peptides, glutamate sodium induces hypothalamic paraventricular nucleus enhancing acupuncture analgesia in the rat. Neurosci Res. 2006;54:49-56. 125. Ypsilantis P, Didilis VN, Politou M, et al. A comparative study ofinvasiveandoscillometricmethodsofarterialbloodpressuremeasurement in the anesthetized rabbit. Res Vet Sci. 2005;78:269-275. 126. Zhang RX, Lao L, Wang X, et al. Electroacupuncture combined with indomethacin enhances antihyperalgesia in inflammatory rats. Pharmacol Biochem Behav. 2004;78:793-797. 127. Zhao Y, Tao T, Wu J, et al. Pharmacokinetics of tramadol in rat plasma and cerebrospinal fluid after intranasal administration. J Pharm Pharmacol. 2008;60:1149-1154.
CHAPTER
32
Small Mammal Dentistry
Vittorio Capello, DVM, Diplomate ABVP (Exotic Companion Mammal), Diplomate ECZM (Small Mammal), and Angela M. Lennox, DVM, Diplomate ABVP (Avian)
Equipment Diagnostic Testing The Clinical Examination Imaging Other Diagnostic Testing Rabbits Anatomy and Physiology of the Skull and Teeth Pathophysiology of Dental Disease Dental Disease Treatment of Dental Disease Treatment of Periapical Infections and Abscesses Facial Surgery and Surgical Treatment of Specific Conditions Rodents Anatomy and Physiology of the Skull and Teeth Pathophysiology of Dental Disease Clinical Presentation by Species Dental Disease Treatment of Dental Disease Treatment of Periapical Infections and Abscesses Ferrets Anatomy and Physiology of the Skull and Teeth Dental Disease Treatment and Prevention Hedgehogs and Sugar Gliders Dentistry of exotic companion mammals has received considerable attention during the past few years, focusing in particular on rabbits and rodents. Dental disease is very common in these species; it has been underdiagnosed or misdiagnosed because of factors related to their anatomic and physiologic differences compared with traditional pet species, the unusual clinical presentations, improper understanding of the pathophysiology, and limited diagnostic tools and dedicated instruments. 452
Outcomes were often unrewarding because of the lack of accurate diagnosis and prognosis. Today, many resources and references are available about this critical topic of medicine and surgery in exotic companion mammals—a field that can benefit from advanced diagnostic imaging like endoscopy, high-quality standard and digital radiology, computed tomography (CT), and magnetic resonance (MR) imaging. Advances in general and local anesthesia as well as in postoperative analgesia greatly help in exploring more accurate and advanced surgical options, especially in cases that involve frequent complications, such as periapical infections, abscesses, and focal and extensive osteomyelitis. The proper clinical approach comes from understanding that dental disease is a syndrome (i.e., a complex of clinical signs and symptoms) with associated primary and secondary diseases or complications.
EQUIPMENT The dental equipment required for pet rabbits and other small mammal species falls into two groups: instruments for diagnosis and instruments for treatment.13,14 Because of the small, long, narrow oral cavity of rabbits and herbivorous rodent species, mouth gags are essential instruments for oral examination under general anesthesia. Special mouth gags designed for rabbits and rodents fit with the clinical crowns of incisors, but these gags should be used with extreme care to prevent excessive stretching of the masticatory muscles and the ligaments of the temporomandibular joint. Be aware that wide opening of the oral cavity is not a natural movement for herbivorous elodont species. The “rabbit and rodent table retractor/restrainer” (also named: “tabletop mouth gag”) is a special platform acting as a combined mouth gag and patient positioner. The second essential instrument to access the oral cavity of rabbits and rodent species is the cheek dilator. Different sizes and shapes are available. Longer blades fit better into the rabbit mouth and are more effective in dilating cheeks. A small, modified “open blade” (rather than flat) cheek dilator is especially designed for rodents, but it also works well for dwarf rabbits weighing less than 1 kg. It is particularly useful in Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 32 Small Mammal Dentistry guinea pigs because of the double-folded cheek opening in this species. Other dental diagnostic instruments used in small mammals are the periodontal probe and the dental explorer, which are commonly used in dog and cat dentistry. A small dental mirror, especially when oral endoscopy is not an option, can be used in small mammals as well. Proper lighting is critical, and magnification is useful in rabbits and mandatory for intraoral inspection of rodent species. A penlight or other handheld focused light source can be used but requires the help of an assistant to allow the operator use both hands. Effective light spots can be connected to magnifying loupes when endoscopy is not an option. Miscellaneous diagnostic instruments include a 21-gauge blunt needle or 22to 24-gauge Jelco catheter to assess the patency of the nasolacrimal duct and culturettes to collect microbiologic samples from any lesion where a bacterial or fungal pathogen is suspected. Special instruments for intraoral dental treatment are needed. Elevators for rabbit incisor and cheek teeth have been specially designed.32 Small extraction forceps can be used to extract incisors; needle holders and small hemostats can be used as well, depending on the patient’s size. For rodents, appropriately contoured needles are used for extracting incisors and are shaped to match the size and curvature of the tooth. A dental unit is recommended for dental treatment of small mammal species. While not necessary for ferrets and other carnivorous or omnivorous species, a straight dental handpiece is needed for intraoral treatment of rabbits and herbivorous rodent species. Many different types, shapes, and sizes of burrs are available. Diamond rasps and cutters for rabbit cheek teeth are available, but their use must be discouraged because forces applied by these instruments can cause fractures of cheek teeth and severe damage to adjacent soft tissues. Facial surgery for treating dental-related abscesses and osteomyelitis requires a basic surgical set of small instruments. In addition, small bone curettes and burrs are needed to debride necrotic bone or to perform selected osteotomy/ostectomy procedures. Transparent or adhesive surgical drapes and small, delicate retractors are especially useful for such small surgical fields.
DIAGNOSTIC TESTING THE CLINICAL EXAMINATION Dental disease is a syndrome and can produce a wide range of clinical signs and symptoms.13 These may be related specifically to the primary dental problem (reduced food intake, anorexia, dysorexia, dysphagia, changes in fecal quantity and size, weight loss) or to complications associated with dental disease (excessive grooming, excessive salivation and drooling, facial abscesses, epiphora, exophthalmos, nasal discharge, dyspnea). Other signs and symptoms are indicative of diseases or conditions secondary to dental disease (poor general condition, gastrointestinal problems, poor coat and skin diseases, ocular diseases, death).13,58 Before performing a physical examination, obtain a thorough history and review the diet and feeding habits with the owner. Considering the capability of prey species to mask or hide symptoms, the absence of a clear clinical history does not rule out the possibility of dental disease. Reduced food intake and abnormal feces in rabbits and rodents are common early symptoms frequently missed by owners. A thick hair coat can
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hide evident signs like weight loss or the presence of facial swellings. Use careful and proper restraint during a dental examination.13,32 The first step in rabbits and larger rodents is to palpate the external maxillary and mandibular profiles (including the ventral aspect of the mandible and the temporomandibular joint). The goal is to detect bony irregularities or swellings indicative of apical elongation of cheek teeth, periapical deformities, or abscesses. Inspect the incisors from both the frontal and lateral aspects. Evaluate the lateral mobility of the mandible to indirectly assess the cheek teeth arcades. Directly palpating the lingual or palatal profile of cheek teeth, using a finger, may be feasible in some rabbits and can help to detect sharp spurs. In unsedated rabbits and some large rodents, the oral cavity can be inspected with an otoscope. A plastic otoscope cone is preferable to a metal instrument because the latter might cause iatrogenic dental fracture if the animal chews on it. However, if used carefully, a nasal speculum with an attached light source is helpful in viewing spurs, elongated crowns, or buccal or lingual ulcerations. Include examination of the eye and periocular structures in the dental examination. Evaluate patency of the nasolacrimal duct in rabbits by flushing with saline or with fluorescein dye (see Chapter 37). When dental disease is a presumptive or a differential diagnosis, complete the oral examination with the animal sedated or under general anesthesia. Because of the natural behavior of most pet rodents, safe restraint and effective oral examination is much more difficult in unanesthetized rodents than in rabbits. Oral inspection other than that of the incisors is not feasible without sedation. Complete inspection and proper diagnosis of dental disease in rodent species must always be performed under general anesthesia.13 Pet ferrets may be presented for chewing problems, but apart from advanced dental disease, they rarely show clear symptoms of dental disease. For this reason, routinely perform a thorough dental examination on ferrets brought in for physical examination. Oral examination can usually be done with manual restraint. Pay special attention to the most caudal cheek teeth, which may be covered by the commissure of the lips.
IMAGING Radiography In rabbits with suspected dental disease, a full radiographic examination of the skull and teeth is an essential diagnostic tool.13,20 Deep sedation or general anesthesia is necessary for optimal positioning for radiographs. Although rabbits are routinely intubated for anesthetic procedures, the endotracheal tube may interfere with the radiographic image.13 Good-quality skull radiographs can be obtained with the use of standard or digital radiographic equipment (see Chapter 35). For standard radiography, ultra-slow-speed films, dedicated intensifying screens, and high-resolution mammography x-ray films are particularly advantageous.13,19 Multiple views are necessary to fully evaluate the dental anatomy. 13,20,70 Never base diagnosis and prognosis on any single radiographic image. In rabbits, include five standard projections in the radiographic series: lateral, left-to-right oblique, right-toleft oblique, ventrodorsal or dorsoventral, rostrocaudal.13,19,20,39 The lateral view is useful to evaluate the occlusion of the maxillary and mandibular incisors, the occlusal plane of the cheek teeth, and the overgrowth of clinical crowns (Fig. 32-1). The normal appearance of the occlusal plane of the cheek teeth is
SECTION VI General Topics
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B Fig. 32-1 Gross (A) and radiographic (B) anatomy of the skull and teeth of the European rabbit (Oryctolagus cuniculus). Lateral view. 1, Nasal bone. 2, Incisive bone. 3, First maxillary incisor tooth. 4, Second maxillary incisor tooth. 5, Mandibular incisor tooth. 6a, Maxillary bone. 6b, Palatine process of the maxillary bone. 7, Perforated surface of the maxillary bone. 8, Maxillary diastema. 9, Mandibular diastema. 10, Alveolar bulla. 11, Zygomatic arch. 12, Maxillary cheek teeth. 13, Mandibular cheek teeth. 14, Incisive part of the mandible. 15, Body of the mandible. 16, Masseteric fossa. 17, Coronoid process of the mandible. 18, Orbital fossa. 19, Temporomandibular joint. 20, Tympanic bulla. (Modified from: Capello V, Gracis M: Rabbit and rodent dentistry handbook. Zoological Education Network (2005), Wiley-Blackwell Publishing (2007), with permission.)
a zigzag line. An ideal oblique projection has minimal (15-20 degrees) rotation, and the contralateral oblique view should always be obtained for comparison. Each projection is especially useful to evaluate reserve crowns and apices of mandibular cheek teeth on one side of the head and the same dental structures of maxillary cheek teeth on the other. The ventrodorsal or dorsoventral view is useful to evaluate overgrowth of cheek teeth and deformities of the mandible. Superimposition of mandibular and maxillary structures impairs evaluation; shifting the tip of the mandible laterally permits visibility of an entire quadrant of maxillary cheek teeth. Ideal positioning in the rostrocaudal projection allows evaluation of the occlusal plane of the cheek teeth and the temporomandibular joints. Excellent symmetrical positioning is mandatory for both the ventrodorsal and the rostrocaudal projections.13,19,39 One or more intraoral dental views using small dental films may be useful in addition to the radiographic study of the skull and teeth. Either the standard radiographic equipment or a dental radiographic unit can be used.3,13,19,39 In rodents, either very high definition intensifying screens and films such as those used for mammography are critical for adequate detail.13 In guinea pigs, the normal bending of cheek teeth (see “Anatomy,” below) does not allow visualization of their occlusal plane on the lateral projection; the rostrocaudal projection is the only one allowing a proper evaluation.13,20,39 In ferrets, radiographs of the skull and teeth are very important when the oral examination suggests the possible presence of retained roots, periodontal infection or abscess, and osteomyelitis.21,55 Open-mouth projections and intraoral projections using small films can be added to standard projections.19 Mouth gags used in ferrets (Jorgensen Laboratories, Loveland, CO; www.jorvet.com) are miniature replicas of dog and cat mouth gags14 and are applied to the tip of canine teeth to open the mouth. When a radiotransparent device is needed
for open-mouth radiographic projections, a presized piece of syringe can be used.
Oral Endoscopy Even during examination of the anesthetized rabbit, experienced observers can still miss several lesions. Oral endoscopy allows thorough inspection of the oral cavity and greatly facilitates detection of subtle lesions because it provides magnified perspective of dental structures.10,13,48,49,74 Instruments for oral endoscopy are described in Chapter 34; normal and pathologic endoscopic patterns have been reported (see “Dental Disease,” below).8,10,13,49,60 While endoscopy is very useful in rabbits, it is an integral part of the examination in rodent species. Other magnification devices are helpful but not always sufficient, and many lesions can be missed without endoscopy.
Computed Tomography Computed tomography (CT) has received great attention in exotic animal medicine during the past few years (see Chapter 35). With the advent of newer spiral CT scanners and the availability of postcapture image manipulation software, this diagnostic imaging modality has become feasible, practical, and extremely useful as an adjunct to standard radiography and provides excellent detail even in small mammals.5,16,19,20,22,28 CT is particularly useful over traditional radiography (visualization of the internal anatomy without superimposition of adjacent and overlying structures) in evaluating complications of dental disease such as facial abscesses and osteomyelitis.15,20,21,72 Optimal traditional radiographs (both digital and analog) still provide higher resolution, but CT allows visualization of important details impossible to see on radiographs because of the superimposition of different anatomic structures on a single plane. In selected cases, CT provides detailed diagnostic information that may be critical for proper prognosis and surgical planning of
CHAPTER 32 Small Mammal Dentistry difficult dental cases.15,21 In particular, three-dimensional volume and surface renderings of the skull can provide excellent information for the diagnosis of dental disease or dental-related problems, such as abnormalities of the temporomandibular joint in guinea pigs.20
OTHER DIAGNOSTIC TESTING A complete blood count (CBC) and biochemical panel are routinely obtained in rabbits with dental disease.10 However, even in cases of periapical infection and abscessation, markedly abnormal findings in the CBC results are uncommon in rabbits.38 Unlike other species, where either leukocytosis and neutrophilia occur in response to bacterial infection, total leukocyte counts in rabbits rarely exceed two or three times the normal value.38,43 For these reasons, the CBC does not provide useful diagnostic or prognostic indications in cases of chronic bacterial infections and abscesses in rabbits. Laboratory findings are instead related to secondary diseases, when present, that may be associated with this syndrome.10 Bacterial culture and sensitivity testing is important in cases of periapical infections and abscesses.13,75,76 Anaerobic gramnegative and aerobic gram-positive pathogenic bacteria have been identified as causative agents, including Fusobacterium nucleatum, Prevotella species, Pseudomonas species, Streptococcus species, and Actinomyces israelii.1,75,76,81 Common rabbit pathogens such as Pasteurella multocida have not been isolated.76 Always request screening for both aerobic and anaerobic organisms when specimens are submitted for culture. Purulent material from the core of the abscess may be sterile and culture results of this are often unrewarding. A biopsy sample of the capsule wall obtained during surgical excision is more appropriate to submit for culture and sensitivity testing.13 Histopathology can be useful in selected cases, particularly when bone neoplasia is suspected.71
RABBITS ANATOMY AND PHYSIOLOGY OF THE SKULL AND TEETH Rabbits are highly specialized herbivorous placental mammals, and the evolutionary process has adapted the anatomy and physiology of the skull and teeth to accommodate their unique diet.13,26,33 Skull morphology is typical of true herbivores, similar to larger species like horses, cattle, sheep, and goats (see Fig. 32-1).13,32,64 The oral cavity is long and narrow, with a narrow opening. The temporomandibular joint is dorsal to the dental occlusal plane. Rabbits are anisognathic, with the mandible narrower than the maxilla.13,26,32,66 The incisors of lagomorphs are similar anatomically and partially functionally to those of rodents.13,26,32,33 For this reason, the order Lagomorpha was previously classified as a rodent suborder (Rodentia). However, lagomorphs differ from rodents because they have two pairs of maxillary incisors (duplicidentata), while rodents have only one pair (simplicidentata). Also, lagomorphs are diphyodont, with two sets of teeth (deciduous and permanent); while rodent species are monophyodont.13,25,26,33,43 However, this physiologic trait does not have practical relevance because the first set is shed before or immediately after birth. The labial maxillary incisors (also named: “first,” or “primary”), are bigger than the palatal incisors (also named: “second,” “secondary,” or “peg”). The
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mandibular and the maxillary first incisors are typically chiselshaped. Rabbits lack canine teeth; incisors are separated from premolar and molar teeth by an edentulous gap (diastema). Premolar and molar teeth are anatomically similar and are therefore simply called “cheek teeth.” They are set closely together, without any significant interproximal space. As maxillary and mandibular cheek teeth differ in number, each mandibular tooth occludes with two maxillary cheek teeth, with the exception of the smaller first and sixth maxillary cheek teeth. The occlusal surface is not an even plane. Transverse ridges form a zigzag occlusal surface that can be seen as a zigzag line from a lateral view both on the skull and on a radiograph (see Fig. 32-1).13,26,32,33,43 All rabbit teeth are hypsodont (long-crowned), and elodont (continuously growing and erupting) teeth. They are defined as aradicular teeth because they do not develop anatomic roots. The anatomic parts of rabbit elodont teeth are the portion above the gingival margin (clinical crown), the portion below the gingival margin and inside the alveolous (reserve crown), and the open root (apex).13,26,33 The diastema separates the dental arcade into two functional units: the incisors and the cheek teeth. The function of the incisors of lagomorphs is to cut grass and other plants. Food is then chewed with the rough occlusal surface of the cheek teeth. Mandibular movements occur in three directions. Rostrocaudal movements are limited in lagomorphs.* Dorsoventral chewing movements similar to those of carnivores are very limited as well. Anisognathism explains the importance of laterolateral movements of the mandible. Because the mandibular dental arcades are closer to each other than the maxillary arcades, mandibular and maxillary cheek teeth are not in occlusion at rest. They come into occlusion alternatively, one arcade at a time, during chewing. Transverse ridges on the occlusal surface interlock with the opposite teeth during chewing, providing an efficient rough surface for grinding and crushing fiber.13,26,33,66 During normal chewing of fibrous foods, the clinical crowns of rabbit elodont teeth are constantly worn and continuously replaced by the reserve crown. The lateral movements allow wearing of the occlusal plane of primary incisors.13,26,43,66
PATHOPHYSIOLOGY OF DENTAL DISEASE The pathophysiology of dental disease is mostly related to the continuous growth of teeth. Any process interfering with normal eruption or wear will result in some aspect of dental disease.13,32,33,66 Four different primary causes have been reported in the rabbit: congenital and developmental abnormalities, traumatic injuries, abnormal wear, and metabolic bone disease.† Congenital jaw mismatch may be caused by prognathism of the mandible or brachygnathism of the maxilla.13,32 This anatomic condition is common in purebred dwarf rabbits weighing less than 1 kg and in lop rabbits. Traumatic injuries often cause fracture of the incisors13; if the apex has been damaged, it may result in malocclusion. Fractures of the incisors that expose the pulp can lead to pulpitis and abscesses. The most important and most frequent cause of dental disease is related to improper nutrition and abnormal wear.13,32,33 The natural diet of rabbits is rich in crude fiber and abrasive silicates. Even pet rabbits properly fed with hay and vegetables,
*References 13, 26, 32, 33, 63, 82. †References 13, 32, 33, 40, 41, 43, 45, 66.
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including those kept in the garden eating the same kind of grass, do not receive the same types and variety of food as free-ranging lagomorphs. Therefore, all pet rabbits can potentially develop at least some subclinical degree of acquired dental disease during their lifetime.32,33 Moreover, despite the increased understanding of proper nutritional requirements, many pet rabbits are fed improper diets including seeds, bread, fruit, or treats and deficient in hay and vegetables. The pathophysiologic sequela follows a typical pattern, with few exceptions.13,32,33 The lack of fiber causes insufficient occlusal wear, and because teeth are elodont, the clinical crowns elongate. This elongation is partially counteracted by the strength of the masticatory muscles, and the increased pressure on cheek teeth (both at rest and during chewing movements) reduces elongation of the clinical crowns. This initiates two subsequent concurrent steps: bending of the axis of cheek teeth and elongating of the reserve crowns. A cascade of abnormal changes then follows. Bending of the cheek teeth follows a typical pattern: the mandibular cheek teeth begin to bend medially (showing lateral convexity), while maxillary cheek teeth begin to bend laterally, with medial convexity. This further reduces the occlusal wear of cheek teeth arcades because the lateral chewing movements are not enough to fill the gap in occlusal surfaces. When occlusal planes begin to malocclude, the rabbit will reject hay, looking for food easier to crush (if available) and thus further reducing the amount of fiber and the lateral chewing movements. The occlusal planes do not wear on the lateral (buccal) margin of the maxillary cheek teeth and on the medial (lingual) margin of the mandibular cheek teeth, causing spikes and spurs to form (Fig. 32-2). These can be very sharp, creating discomfort and leading the rabbit to further reduce the degree of lateral chewing movement. Injuries and ulcerations of the tongue (from the mandibular cheek teeth) and the buccal mucosal surface (from the maxillary cheek teeth), often severe, represent secondary complications. On the apical ends of the teeth, elongation of reserve crowns leads to apical deformity and stretching and perforation of
the cortical bone. Vascular support to the apex is reduced or impeded and diseased teeth partially loose their capability to grow, becoming more fragile and prone to fractures. Increased interproximal space and widened periodontal space, caused by bending of the cheek teeth, lead to food impaction and increase the risk of periapical infection and focal osteomyelitis. Perforation of the cortical bone and periapical infection lead to the most common complication of acquired dental disease: infection of soft tissues and facial abscesses. Excessively elongated cheek teeth, particularly the mandibular first premolars, may also lead to functional prognathism of the mandible.13,32 Once the crowns have elongated, the jaw is held more open and the mandible slides rostrally into a functional prognathic position, leading to malocclusion of incisors (Fig. 32-3). This condition is often incorrectly diagnosed as congenital, even in aging rabbits. Metabolic bone disease as a cause of acquired dental disease has been extensively studied and reported.40,41,43,45 In a study in 81 pet rabbits, parathyroid hormone concentrations were abnormally high and calcium concentrations were low, suggesting metabolic bone disease as a predisposing condition for acquired dental disease.41 Most affected rabbits demonstrate demineralized skull bones and radiographic findings consistent with osteodystrophic disease of both the teeth and the skull bones. Demineralization of the bone matrix and weakness of the alveolar bone lead to loosening of the teeth, bending and rotation of the cheek teeth within the alveolus, and widened interproximal spaces. These changes lead to improper occlusion of cheek teeth and resulting abnormal wear of the clinical crowns. The ventral cortical bone becomes deformed, and the periosteum of the mandible and the maxilla eventually becomes perforated, leading to secondary complications such as abscesses. Different pathophysiologic patterns (in particular abnormal wear and underlying metabolic bone disease) may be present in the same patient.
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Fig. 32-3 Acquired malocclusion of incisor teeth secondary to
Fig. 32-2 Pathophysiologic pattern of acquired dental disease following improper nutrition and insufficient wear. (Modified from Crossley DA: Oral biology and disorders of Lagomorphs. Vet Clin No Am Exot Anim Pract 2003, 6: 629-659.)
prognathism of the mandible in a rabbit. The rostral view (A) shows an abnormal oblique occlusal plane, abnormal curvature and wider interproximal space of maxillary teeth, and abnormal enamel of the right mandibular incisor tooth. The lateral view (B) shows the most common pattern of malocclusion of incisor teeth: the mandibular incisors tend to elongate rostrally, while the maxillary first incisors tend to elongate caudally. (Used by permission from Vittorio Capello, DVM.)
CHAPTER 32 Small Mammal Dentistry
DENTAL DISEASE The three most common clinical presentations of dental disease relate to overgrowth of the incisors, reduced food intake, anorexia or dysphagia, and presence of a facial swelling.13 Two or three of these conditions can be present at the same time. While dental disease affecting the cheek teeth is more frequent, diseases affecting the incisors are more apparent to the owners. Typically, the mandibular incisors tend to elongate labially, while the maxillary incisors tend to elongate palatally.13
Fig. 32-4 Early stage of acquired dental disease of cheek teeth in a rabbit following improper nutrition and insufficient wear. Radiographic signs are elongated clinical and reserve crowns, widened interproximal spaces of cheek teeth, abnormal curvature of first premolars, and deformity of the ventral mandibular profile caused by apical overgrowth. Malocclusion of incisor teeth and clinical symptoms are usually not present at this stage. (Reprinted from: Capello V, Gracis M: Rabbit and rodent dentistry handbook. Zoological Education Network (2005), Wiley-Blackwell Publishing (2007), with permission.)
For this reason, overgrown mandibular incisors usually do not produce secondary lesions because they are anterior to the upper lip and nose, while overgrown maxillary incisors curve dorsally and may damage the lips or the palate. Very elongated incisors may fracture. With end-stage dental disease, growth may be slowed or arrested. Dental disease of the incisors also affects the reserve crown. Abnormal elongation and apical deformity leads to partial or complete obstruction of the nasolacrimal duct. Epiphora, dacryocystitis, subsequent ocular lesions, or facial dermatitis are common possible sequelae (see Chapter 37).7,13,44,77 Rabbits with acquired dental disease of the cheek teeth may present at different stages. The earliest stage is elongated crowns. Radiographically the normal occlusal plane and its zigzag pattern are lost; the first premolars are curved, interproximal spaces widen, and the ventral profile of the mandibular cortical bone deforms slightly due to increased pressure (Fig. 32-4). Coronal elongation and the abnormal occlusal plane appear as “wave mouth” or “step mouth” on oral examination (Fig. 32-5).13,20 Clinical signs and symptoms are usually not present at this stage. In later stages, these abnormalities become more severe. Sharp spur and spikes may develop on the cheek teeth, leading to lesions of the tongue or the buccal mucosa (see Fig. 32-5). A common sequela of excessive coronal elongation of cheek teeth is fracture, especially longitudinal fracture of mandibular premolars.13 End-stage dental disease of the cheek teeth is frequently associated with resorption of bony and dental tissue. The most common complications of acquired dental disease are periapical infections, osteomyelitis, and facial abscesses.13,18,43 The latter appear as large masses usually located at the ventrolateral aspect of the mandible or the lateral aspect of the maxilla. Some rabbits may show clear unilateral exophthalmos. Abscesses are typically firm, cool, and nonpainful. Early small masses are usually missed by the owners because of their location and the presence of fur, especially in long-haired rabbits.
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Fig. 32-5 Oral endoscopy in the rabbit. A, Normal appearance of cheek teeth, right mandibular arcade. B, Malocclusion of the right mandibular arcade. Abnormal elongation (or shortening) of clinical crowns, uneven occlusal plane (referred as “step mouth”) and initial lingual bending of the teeth. Clinical symptoms are usually absent at this stage. C, Sharp lingual spike of the second molar tooth caused by improper wearing, and related lesion of the lingual mucosa (arrow). P, premolar; M, molar; T, tongue. (Reprinted from: Capello V, Gracis M: Rabbit and rodent dentistry handbook. Zoological Education Network (2005), Wiley-Blackwell Publishing (2007), with permission. B and C used by permission from Vittorio Capello, DVM.)
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A
B Fig. 32-6 A, Coronal elongation and malocclusion of the incisor teeth following congenital brachygnathism of the maxilla in a rabbit. The maxillary diastema is shorter than normal, and its length is similiar to that of the mandibular diastema (compare with Fig. 32-1, B). Cheek teeth are normal, including the zig-zag line at the occlusal plane. B, Postoperative radiograph after extraction of incisor teeth. (Used by permission from Vittorio Capello, DVM.)
The prognosis of dental disease is based on an accurate diagnosis, which results from a thorough workup. Discuss every clinical and practical aspect of treatment and management with owners, including severity of disease, general patient condition, continued required care and subsequent dental procedures, compliance, and expected costs. Many rabbits can live in good condition for years by eating soft foods, ground pellets, and chopped hay, even with severe to end-stage dental disease. In other cases, aggressive facial surgery is needed to stop or control the disease.
TREATMENT OF DENTAL DISEASE In general, the goals of dental procedures are to reduce abnormal tooth length, to restore the occlusal plane to as near normal as possible, and to extract diseased teeth.59 Complications and secondary diseases like periapical infections, osteomyelitis, and abscesses are treated with combined dental procedures and extraoral surgery. Surgical treatment of diffuse osteomyelitis may require more invasive facial surgery. Anesthesia of small mammals is discussed in Chapter 31. Many different anesthetic protocols are suitable for diagnosis and treatment of dental disease in rabbits and other small mammal species.57 Local anesthesia is an important adjunct for extraction of both incisor and cheek teeth.57,59
Medical Treatment Medical therapy alone is generally inadequate for the treatment of dental disease but is an important adjunct to dental and surgical procedures. Medical therapy is aimed to address several concerns such as antibiotic, analgesic, and supportive treatment.13 In choosing an antibiotic, base the selection on both aerobic and anaerobic culture and sensitivity testing when possible, and consider any species-specific contraindications, such as oral administration of penicillins in rabbits and herbivorous rodents.13,43,68 Analgesia is critical to prevent pain-related
anorexia. Supportive treatment includes hand-feeding products for convalescing small mammals, such as Oxbow Critical Care (Oxbow Pet Products, Murdock, NE, www.oxbowhay.com), especially for herbivorous species. Fluid therapy may also be necessary in some patients.13
Dental Procedures Treatment of disease of the incisors includes coronal reduction and extraction.13,43,58 Coronal reduction is limited to cases in which the malocclusion is not severe and has been diagnosed early and proper occlusion of both incisor and cheek teeth can be restored. Reduction and reshaping of the incisors must be performed on anesthetized animals with low or high-speed dental equipment and appropriate burrs.23,32,33 Do not use trimmers, clippers, or similar instruments to cut overgrown or maloccluded incisors. These instruments do not allow the restoration of a normal incisal edge, and they frequently lead to complications such as fractures, damage of the apical germinative tissue, pulp exposure, endodontic infection, and periapical abscessation.12,13,30 The only definitive and completely effective treatment for severe malocclusion and acquired dental disease of incisors is extraction (Fig. 32-6).6,11,13,33 Rabbits adapt easily to lack of incisors, much better than when teeth are short or maloccluded, and they are able to eat normally using the lips and tongue to prehend food. The technique for extraction of incisors is described in Box 32-1. Extracting the entire set of six incisors is usually necessary. In selected cases of fracture or infection of a single mandibular incisor tooth where malocclusion is not present, extracting that incisor tooth may be indicated. The contralateral incisor may be able to maintain a normal occlusal plane because of lateral chewing movements.13 Postoperative skull radiographs help in detecting possible remaining tooth fragments. Partially removing a tooth or failing to destroy pulp tissues can result in partial or complete regrowth, which can be addressed at a later time.13,73
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Box 32-1 Step-by-Step Procedure for Extracting Elodont Incisor Teeth 1. Take diagnostic radiographs in five standard projections before the dental procedures. 2. Perform intraoral dental procedures as needed before extracting the incisor teeth. 3. Place the patient under general anesthesia in dorsal or lateral recumbency. 4. Scrub the gingiva with 0.1% chlorhexidine solution. 5. For optional adjunct analgesia, perform local blocks for the mental nerve and the rostral infraorbital nerve for extraction of mandibular and maxillary incisor teeth, respectively.
Extraction of the Mandibular Incisor Teeth 6. Incise the gingival attachment circumferentially around the whole tooth with the tip of a No. 1 or 15 scalpel blade. 7. Insert a luxator in the periodontal space on the medial side of the tooth until resistance is felt. Hold the luxator in position for a few seconds to stretch and disrupt the periodontal ligament. Gradually move the tip of the luxator toward the apex of the tooth. Use your free hand to hold and stabilize the mandible at all times. Perform the same procedure on the lateral aspect of the tooth. 8. Following the same technique, use a contoured, 18-gauge hypodermic needle to break down the periodontal ligament on the lingual and labial sides of the tooth. The tooth should be loose and very mobile at this point. 9. Grasp the tooth with the extraction forceps as close to the gingival margin as possible and extract it using steady, slowly increasing force, following the curvature of the root. Once loosened, gently intrude the tooth in the socket to destroy the germinal tissues at the bottom of the alveolus. If the periodontal ligament has been completely and correctly severed and the tooth is not severely deformed, it can be extracted without the use of significant force. Bleeding is usually minimal. 10. Once it is extracted, examine the tooth to ensure that the entire tooth and its pulp tissues have been removed. 11. After extraction, insert a needle into the alveolus to curette the alveolar walls and damage any germinal tissue to prevent tooth regrowth. 12. Flush the alveolar cavity with abundant saline solution to remove any debris and dental or bony fragments. If periapical infection is present, use dilute 2% povidone iodine or 0.05% to 0.1% chlorhexidine solution to flush.
Dental procedures on cheek teeth in rabbits include reducing clinical crowns and extracting diseased teeth.* Coronal reduction should be aimed at restoring the normal height and angle of the occlusal plane as near to normal as possible. Even if it is impossible to restore the normal zigzag-shaped occlusal plane or to reverse the abnormal changes affecting the reserve crowns and the apices of cheek teeth, this procedure is still very useful for many reasons. Sharp spurs are burred, allowing mucosal lesions to heal and reducing discomfort. Clinical crowns are shortened, reducing pressure and allowing easier lateral movements during chewing. Diminished pressure counteracts some of the pathophysiologic steps described above, like bending of
*References 12, 23, 32, 33, 43, 58.
13. Close the alveolus by suturing the gingiva with simple interrupted sutures or a purse-string suture pattern using 3-0 or smaller absorbable suture material. The pursuestring suture must have at least four points of fixation before being tightened. Do not place postextraction sutures when infection is present.
Extraction of the Maxillary Incisor Teeth 14. Use a contoured needle to sever the periodontal ligament on the labial and palatal sides of the maxillary incisor teeth, similar to the procedure described for mandibular incisors. Lift the upper lip with your free hand, which is used to hold and stabilize the patient’s head without obstructing the nares (if the patient is not intubated). 15. Sever the periodontal ligament on the medial and lateral aspects of the teeth with Crossley’s luxator. The periodontal ligament is particularly tight on the medial aspect of the tooth. 16. Once it is loose, gently grasp the tooth with suitable extraction forceps or a pair of needle holders. To avoid dental and bony fractures, gently extract the tooth following the natural shape of the tooth, applying slight distal rotation. 17. Completely extract the tooth and examine it for the presence of pulp tissues. Repeat the procedure on the contralateral incisor tooth. Control bleeding with sterile cotton swabs. 18. Use a thin (22-gauge) hypodermic needle to loosen the second incisor teeth. When the tooth is completely luxated, extract it with small extraction forceps or thin hemostats, taking care to avoid crushing the tooth. 19. After extraction, curette the alveoli to remove any remaining pulp tissues and rinse with saline or 0.1% chlorhexidine solution. 20. To promote gingival healing, close the extraction site with simple interrupted sutures or a purse-string suture pattern with 3-0 or 4-0 absorbable suture material. Do not suture the extraction site when infection is present. In using a purse-string pattern, fix the suture material at a minimum of six points before tightening it. 21. Obtain a lateral view postoperative radiograph to confirm complete extraction of all six incisor teeth.
the teeth, apical deformity, and cortical bone deformity. Therefore reduction of crown length is especially useful in the early stages of acquired dental disease, and in rabbits with concurrent metabolic bone disease, when the surrounding bone is weaker than normal. In many cases, overgrowth of gingival tissue may mask elongated crowns, making accurate analysis of preoperative radiographs very important. Postoperative radiographs are also necessary to determine whether crowns are adequately reduced and the occlusal plane is restored. Because a contra-angle hand piece is impossible to use in the narrow oral cavity of rabbits, use a straight hand piece for coronal reduction. In using the drill, protect soft tissues with a flat or concave dental spatula and remove accumulated tooth dust with moistened cotton swabs. A burr protector is available that fits over the end of the burr tip and protects soft tissues
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from damage during drilling. Prevent thermal damage to teeth by cooling the teeth with saline. The indications for extracting cheek teeth are fracture or luxation associated with periapical infection. Marked deformities of individual teeth not easily corrected by burring may represent an indication for extraction. As many cheek teeth as possible should be retained to allow proper crushing of food. Even if rabbits adapt surprisingly well to progressive dental disease and chronic lack of cheek teeth, extracting more than two premolars or molars from each quadrant can cause discomfort during chewing, and should be avoided when possible. The opposing cheek teeth may or may not have to be burred at regular intervals, depending on the stage of dental disease and their ability to grow. The extraction of cheek teeth can be challenging, depending on the position and condition of the affected teeth. Even after proper luxation and with the use of proper extraction forceps, teeth are easily fractured during extraction. Complete intraoral extraction is also impossible when fracture of the reserve crown, severe apical deformity, retained root tips, and dental ankylosis are present. In these cases, combined intraoral and extraoral extraction of fragments is necessary. When periapical infection is not present, the socket must be protected after extraction to prevent food impaction and infection.34 When feasible, the gingiva may be sutured over it, or the socket can be filled with Doxirobe gel (Pfizer Animal Health, Kalamazoo, MI). In case of periapical infection, osteomyelitis, and concurrent abscessation, the socket may be left open and treated postoperatively as described later.
TREATMENT OF PERIAPICAL INFECTIONS AND ABSCESSES Because periapical infections, osteomyelitis, and subsequent abscessation of soft tissues are true bacterial infections, antibiotic treatment would appear to be the treatment of choice. Nevertheless, with the exception of some anectodal reports, medical treatment alone is ineffective in treating this condition in rabbits.13,18 To obtain long-term therapeutic success and prevent frequent recurrence, three pathologic conditions of this syndrome must be addressed: the presence of the abscess capsule, osteomyelitis, and necrotic tissue (soft, dental or bony) acting as a sequestrum.18 The aim of most treatment protocols, except wound-packing as described below, is to remove the entire abscess including the capsule, extract the tooth fragment(s) involved, and debride the osteomyelitic bone.1,12,13,18,43 Different surgical options have been reported beyond the simple (and usually ineffective) incision of the abscess and flushing of the purulent material. Excision of the entire abscess—followed by either primary closure or after packing of the surgical site with antibiotic-impregnated polymethylmethacrylate (AIPMMA) beads* or marsupialization and packing of the surgical site with calcium hydroxide,67 honey42,62 or sugar61 solution, or bioactive ceramics1—has been reported. A technique in which the capsule is not removed has been reported in rabbits with dental abscesses.75 The procedure involves lancing the abscess, minimally debriding and cleaning the abscess cavity, and packing the cavity with strips of 3to 5-mm-diameter sterile gauze impregnated with antibiotics,
*References 2, 35, 36, 43, 47, 84.
most commonly ampicillin. Rabbits are concurrently treated with systemic antibiotics. This technique may be an option in rabbits with minimal osteomyelits; however, efficacy in rabbits with extensive osteomyelitis or bony sequestra is not known. Marsupialization of the soft tissues around the area of the affected bone is our treatment of choice, followed by daily postoperative flushing and further gentle debridement of the surgical site, applying antiseptics or other products to promote healing, and constant direct monitoring of healing process.12,13,29 Slow healing by secondary intention is also critical in cases of simultaneous extraction of one or two cheek teeth, where suturing the gingiva may be very difficult or impossible. In these cases, the alveolus may become impacted with food. Marsupializing the tissues actually allows flushing of the intraoral fistula through the cutaneous opening until the new bone apposes and the gingiva heals, effectively closing the fistula. Despite exposure of part of the deep bone, marsupialization is well tolerated by rabbits and most of them do not need an Elizabethan collar or hand-feeding. In our experience, this procedure is associated with a higher percentage of successful outcomes and long-term postoperative follow-up, particularly in cases of severe osteomyelitis.13 A longer postoperative period, temporarily unattractive cosmetic appearance, and significant owner commitment must be discussed before surgery. The technique for excising odontogenic abscesses, debriding the osteomyelitic site, and marsupialization of the soft tissues is described in Box 32-2 and shown in Figure 32-7.
FACIAL SURGERY AND SURGICAL TREATMENT OF SPECIFIC CONDITIONS In cases of severe complications of acquired dental disease in rabbits, the severity of the bone infection is not directly related to the severity of the clinical signs and symptoms. This is why accurate diagnosis and proper prognosis are the keys to successful surgical treatment. Before planning surgical debridement of an abscess and the related osteomyelitic bone, determine which tooth or teeth are involved and the extent of the osteomyelitis. Even if exploratory surgery remains the most important step, a complete presurgical evaluation allows poor surgical candidates to be ruled out and assessment of the degree of difficulty of the surgical procedure. Beside conventional radiographs, the use of CT is extremely helpful for accurate diagnosis, proper prognosis, and surgical planning. When treatment of periapical infections and osteomyelitis fails, extensive osteomyelitis of the body of the mandible is a common sequela.15,18,21 Thorough and deep surgical debridement cannot be performed, and bone healing by secondary intention may be impossible when the entire body of the mandible is affected. In severe cases, rostral partial unilateral mandibulectomy is an option; it was successful in the treatment of two pet rabbits18,21 (Fig. 32-8). In rabbits, the alveolar bulla includes the reserve crowns and the apices of the four caudal maxillary cheek teeth.13,64 Because of the anatomic relationship with the orbital fossa, retrobulbar abscesses are a relatively common sequelae of periapical infection of these teeth. The alveolar bulla can fill with food debris, tooth fragments, and pus and the infection can then spread to the retrobulbar space. The typical presentation is unilateral exophthalmos without a history of trauma, possibly complicated by panophthalmitis. Advanced cases necessitate enucleation of the globe. Diagnosis of impaction and periapical
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Box 32-2 Step-by-Step Procedure for Treatment of Surgical Abscesses and Osteomyelitis 1. Perform diagnostic radiographs in five standard projections before the dental procedures. 2. For optional adjunct analgesia, perform local nerve blocks of the mental and inferior alveolar nerves for extracting mandibular cheek teeth and surgical debridement of the mandible. Perform a local nerve block of the rostral infraorbital nerve for extracting the maxillary cheek teeth and surgical debridement of the maxilla. 3. Place the patient under general anesthesia in dorsal or lateral recumbency, depending on necessary intraoral procedures and the site of infection. 4. Perform intraoral dental procedures as needed; in particular, extract the tooth/teeth affected by periapical infection. 5. Shave and aseptically prepare the surgical site. Place an adhesive transparent drape on the surgical field, facilitating the view of the orientation of the head. 6. Make a 1- to 2-cm skin incision over the mass, taking care not to enter the underlying abscess. 7. Gently dissect subcutaneous tissue and muscle layers to free as much of the abscess capsule as possible, taking care not to disrupt connection to the cortical bone. 8. Incise the junction between the capsule and mandibular bone with the tip of a No. 11 scalpel blade. Portions of the wall of the abscess are typically composed of thick connective tissue and/or thin cortical bone, which usually
prevents removal of the entire capsule in one piece. Carefully dissect the wall of the abscess free from the bone and remove it. 9. Remove the purulent exudate and flush the bone cavity. Collect samples for culture and sensitivity from the capsule wall, as the purulent material itself is often sterile. Remove remaining debris or purulent material with a bone curette. 10. Debride infected or necrotic cortical bone to the point of bleeding with a bone curette. 11. After thorough debridement, the fragment of the diseased tooth/teeth, if present, can be seen. Use a hypodermic needle to gently free the attachment of the fragment to the bone. Extract the tooth and fragment of necrotic alveolar bone. 12. Debride the bone cavity again and thoroughly flush with saline and dilute povidone iodine or 0.1% chlorhexidine solution. 13. Marsupialize the surgical site with 3-0 or smaller nonabsorbable suture material. 14. Obtain postoperative radiographs with lateral and oblique views to confirm proper coronal reduction of the cheek teeth, bone debridement, and extraction of tooth fragments. 15. Show and discuss with the owner detailed instructions for flushing. Schedule frequent rechecks.
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Fig. 32-7 Surgical treatment of periapical infection of a mandibular cheek tooth in a rabbit. A, Mandibular abscess, after shaving. B, Exposure of the abscess. After complete exposure, the junction between the capsule and mandibular bone is incised with blunt scissors. C, Appearance of the osteomyelitic bone site after the abscess was excised and the adjacent soft tissues were debrided. The diseased fragment of the involved tooth is extracted by the extraoral approach. D, Marsupialization of the surgical site. E,F, Follow-up 10 days and 20 days after surgical treatment. (Used by permission from Vittorio Capello, DVM.)
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Fig. 32-8 Computed tomography image of the skull of a rabbit with end-stage dental disease and diffuse osteomyelitis of the right mandible, before (A,B,C) and 6 months after (D,E,F) rostral partial mandibulectomy. A,D, Scout views adapted from the patient’s lateral radiographs, demonstrating the scanning angle perpendicular to the palatine process of the maxillary bone. The yellow lines represent the 0.5-mm slices shown in (B) and (E), respectively. B,E, Axial views. Scanning parameters: scan speed: 1 second; mA: 125; kVp: 120; slice thickness: 1.0 mm; reconstruction index: 0.5 mm; WL: 345; WW: 1,500; image size (matrix) 512 × 512 pixels. C,F, Three-dimensional surface rendering (shaded surface display [SSD]) of the skull. Viewer software for DICOM images enables virtual three-dimensional renderings of the two-dimensional scans, providing a detailed, realistic anatomic view of both the surface and the internal bone structure and their relationship with soft tissues. B, Axial view at the level of mandibular first premolars showing the normal left mandible on the left side, and the osteomyelitic right mandible. C, Diffuse osteomyelitis and severe periosteal reaction of the body and of the incisive portion of the right mandible. The masseteric fossa including the cranial margin is still normal. E, Axial view at the level of the caudal portion of left mandibular CT3, showing the normal left mandible and the missing right mandible. The cranial margin of the osteotomy site is partially included in this slice (arrow). F, Three-dimensional surface rendering shows a cortical fragment attached to the mandibular symphysis (I, left mandibular incisor tooth), and the missing body of the right mandible. The osteotomy site is also visible. The masseteric fossa and the branch of the right mandible are still normal. (Used by permission from Vittorio Capello, DVM.)
infection or abscessation of the alveolar bulla is ideally made before a retrobulbar abscess develops and the eye is compromised. The extraoral lateral surgical approach for abscesses of the two rostral premolars (zygomatic abscesses) is feasible because they are located cranial to the zygomatic arch. However, because of the presence of the zygomatic arch, the same approach cannot be applied to the alveolar bulla. Access to the alveolar bulla through the intraoral fistula after extracting the diseased maxillary cheek tooth or teeth is feasible, but proper debridement is very difficult because the fistula is much smaller than the bulla itself.61 A lateral extraoral surgical approach to the alveolar bulla by partial ostectomy of the zygomatic arch was done in a Flemish giant rabbit, allowing thorough debridement of the alveolar bulla and introduction of small AIPMMA beads.15,21 Periapical infection and osteomyelitis of the mandibular cheek teeth 4 and 5 are less common but carry a difficult prognosis because the abscess usually spreads beneath the masseter muscle, making both surgical access and marsupialization more
challenging. Also, debriding the osteomyelitic bone is not feasible because the cranial portion of the masseteric fossa is much thinner than the body of the mandible, carrying much higher risk of iatrogenic fracture.15,21 Unilateral rostral mandibulectomy has been reported for surgical treatment of bone tumors: a cementoma and a chondrosarcoma of the mandible.85 Osteogenic sarcomas (one involving the maxilla50; three involving the mandible80,83,85) have also been reported.
RODENTS ANATOMY AND PHYSIOLOGY OF THE SKULL AND TEETH All rodent species have one pair of well-developed maxillary and mandibular incisors, representing the best-known anatomic peculiarity of this order. Unlike rabbits, rodents have just one pair of maxillary incisors (simpicidentata).13,25,56,64,66 Incisors
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Table 32-1 Dental Formulas of Most Common Species of Exotic Companion Mammals INCISOR TEETH
CANINE TEETH
CHEEK TEETH Premolars
Molars
Total # of Cheek Teeth
Total # of Teeth
Maxillary/ Mandibular
Maxillary/ Mandibular
Maxillary/ Mandibular
Maxillary/ Mandibular
Maxillary/ Mandibular
Rabbit
2 x 2/1 Elodont/ Hypsodont
0/0
2 x 3/2 Elodont/ Hypsodont
2 x 3/3 Elodont/ Hypsodont
2 x 6/5 = 22 Elodont/ Hypsodont
28
Porcupine-like (hystricomorph) rodents: Guinea pig, chinchilla, degu
2 x 1/1 Elodont/ Hypsodont
0/0
2 x 1/1 Elodont/ Hypsodont
2 x 3/3 Elodont/ Hypsodont
2 x 4/4 = 16 Elodont/ Hypsodont
20
Rat-like (myomorph) rodents: Rat, mouse, hamster, gerbil
2 x 1/1 Elodont/ Hypsodont
0/0
0/0
2 x 3/3 Anelodont/ Brachyodont
2 x 3/3 = 12 Anelodont/ Brachyodont
16
Squirrel-like (sciuromorph) rodents: Prairie dog, other squirrels
2 x 1/1 Elodont/ Hypsodont
0/0
2 x 1-2/1 Anelodont/ Brachyodont
2 x 3/3 Anelodont/ Brachyodont
2 x 4 – 5/4 = 16-18 Anelodont/ Brachyodont
20-22
Ferret
2 x 3/3 Anelodont/ Brachyodont
2 x 1/1 Anelodont/ Brachyodont
2 x 3/3 Anelodont/ Brachyodont
2 x 1/2 Anelodont/ Brachyodont
2 x 4/5 = 18 Anelodont/ Brachyodont
34
Hedgehog
2 x 2/2 Anelodont/ Brachyodont
2 x 1/1 Anelodont/ Brachyodont
2 x 3/2 Anelodont/ Brachyodont
2 x 3/3 Anelodont/ Brachyodont
2 x 6/5 = 22 Anelodont/ Brachyodont
34
Sugar glider
2 x 3/2 Anelodont/ Brachyodont
2 x 1/0 Anelodont/ Brachyodont
2 x 3/3 Anelodont/ Brachyodont
2 x 4/4 Anelodont/ Brachyodont
2 x 7/7 = 28 Anelodont/ Brachyodont
40
vary in shape, color, and thickness among species.13,25,66 Incisors of rodents are covered by enamel only over the labial surface. The enamel can be white (guinea pigs, Russian hamsters), yellow, or orange-pigmented (chinchillas, golden hamsters, prairie dogs). The length of the clinical crown of incisors differs from species to species. For example, in guinea pigs, the length of the mandibular incisors is normally triple the length of the maxillary incisors. Differences in normal anatomy should be kept in mind to avoid misinterpreting abnormal patterns. In all rodent species, both maxillary and mandibular incisors pre sent a chisel-shaped occlusal surface and are elodont teeth. As in rabbits, all rodents lack canine teeth and a diastema is pre sent between the incisor and the first premolar (or molar) tooth. Premolar and molar teeth vary among rodent species. In most species, they are anatomically indistinguishable and are simply called “cheek teeth.”13 The hundreds of species belonging to the Rodentia are grouped into three suborders based on anatomic and functional differences of the masseter muscle. These are the Hystricomorpha (“porcupine-like”) or Caviomorpha (“guinea pig-like”), Myomorpha (“mouse-like” or “rat-like”), and Sciuromorpha (“squirrellike”).13,34 The suborders have different dental formulas (Table 32-1), but from the standpoint of the dental anatomy and physiology, they can be simply reduced to two groups.13,56 The
porcupine-like rodent species have elodont incisor and cheek teeth. Despite differing dental formulas, guinea pigs, chinchillas, and degus have dentition physiologically similar to rabbits. Rat-like and squirrel-like rodent species have elodont incisors but anelodont (rooted, not growing throughout life) cheek teeth. This distinction between species with elodont and anelodont cheek teeth is clinically very important. Malocclusion and acquired dental disease of the cheek teeth is not common in myomorph or sciuromorph species, as it is in rabbits and hystricomoph species, and coronal reduction of cheek teeth must never be performed. A minor distinction between myomorph and sciuromoph species is that the first group lacks premolar teeth. Rodent species are monophyodont with a single set of teeth.13,25,56,66 The occlusal surface of cheek teeth of hystricomoph rodents, as true herbivores, is rough and uneven owing to enamel crests and dentinal grooves.13,25,66 Unlike those of rabbits, these occlusal surfaces are flat and do not present a zigzag pattern. In guinea pigs, the cheek teeth are curved; the mandibular teeth have a buccal (lateral) convexity while the maxillary teeth have a palatal (medial) convexity.13,25,66 This results in a 30-degree oblique occlusal plane that slopes from dorsal to ventral, lateral to medial (Fig. 32-9). The clinical crowns are much shorter than those of rabbits.
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SECTION VI General Topics
Fig. 32-9 Rostrocaudal projection of the skull of a guinea pig with
Fig. 32-10 Pseudo-odontoma of incisor teeth in a prairie dog.
normal dentition. Green superimposition indicates the shape and size of the left premolars; the orange line indicates the oblique occlusal plane. (Modified from Capello V, Gracis M: Rabbit and rodent dentistry handbook. Zoological Education Network (2005), Wiley-Blackwell Publishing (2007), with permission.)
Typical radiographic abnormalities are represented by the abnormal occlusal plane; enlarged and deformed apices (especially those of maxillary incisor teeth); irregular dorsal profile of the maxillary incisors, showing the typical folds of newformed dentine; and irregular ventral profile of the mandibular incisors. Abnormalities of maxillary incisor teeth reduce the size of both the nasal cavity and its opening (arrow), resulting in severe dyspnea. Excessive wearing of mandibular cheek teeth is also present. (Used by permission from Vittorio Capello, DVM.)
Cheek teeth of prairie dogs have multiple roots. Although these animals are herbivorous ground squirrels, their dental structure is similar to that of primates, with cusps and ridges that create a rough occlusal surface.13 The temporomandibular joint and masticatory muscles of rodent species (especially those with anelodont cheek teeth) allow much greater rostrocaudal movements than in rabbits.13,25,33 Like those of rabbits, elodont cheek teeth of hystricomorph rodents are worn during normal chewing activity.
PATHOPHYSIOLOGY OF DENTAL DISEASE Congenital malocclusion is rare because prognathism of the mandible (or brachygnathism of the maxilla) is not recognized in rodents. True dwarfism as recognized in pet rabbits has not been documented because rodent species have not been selectively bred for size variation.17 The primary cause of dental disease in hystricomorph rodents (like guinea pigs) is insufficient or improper wearing of cheek teeth due to inappropriate diet, in particular lack of fiber.9,13,28,33,56 This syndrome does not occur in rodent species with anelodont cheek teeth, such as rats, hamsters, and prairie dogs. However, these patients can develop dental problems of the cheek teeth due to excessive wearing.13,17 Metabolic bone disease as an underlying cause of dental disease has not been investigated and reported in rodent species. Primary congenital deviation and malocclusion of the incisors in growing rats, hamsters, and squirrels has been described.56 This may be difficult to distinguish from malocclusion due to incisor fractures in adult animals, as owners often are not aware that an injury has occurred. Acquired malocclusion and severe deviation of incisors occurs most often after repeated trauma and fractures. Odontomas have been reported in many rodent species, both companion and laboratory animals: rats, mice, guinea pigs, prairie dogs, degus, and tree squirrels. I have encountered radiographic abnormalities consistent with this lesion also in a
pet rabbit, a chinchilla, a chipmunk, and a citellus (Spermophilus citellus). In all rodent cases, the incisors were affected. Terminology about this dental disease has not been clear in the past. The term “odontoma” was meant including both odontogenic tumors (invasive and locally destructive) and the odontogenic dysplasia of incisors, very frequent in prairie dogs and other squirrels (mostly locally compressive on the cranial portion of nasal cavities). Two different kinds of odontomas are classified.46 The nonneoplastic, dysplastic malformation, occurring when normal tooth eruption is impaired or arrested, has been properly named pseudo-odontoma.33 It is a very frequent condition in prairie dogs.* Apical growth continues, causing primary deformation of the apex and of the reserve crown and secondary abnormalities of the surrounding structures, such as the incisive bone. The outcome is severe apical deformity and folds of new-formed tooth substance, especially on the labial surface of the reserve crown, acting as a space-occupying mass and leading to progressive obstruction of the nasal openings (Fig. 32-10).17,33 The term elodontoma has been suggested because those dental changes develop in elodont incisors.4,51 The pathogenesis of odontomas is not clear. Different hypotheses have been proposed, including osteopetrosis (an autosomal recessive disease) in rats and mice. Nevertheless, I have diagnosed and treated dental dysplasia of a mandibular incisor tooth in a citellus with concurrent metabolic bone disease. In prairie dogs and other squirrels, repeated trauma, fractures, and subsequent acquired dental disease are the most important causes of dysplastic changes (pseudo-odontomas). Trauma can be the result of constant chewing of cage bars, fractures, or improper trimming of maloccluded incisors.
*Reference 8, 13, 17, 20, 33, 51, 56, 78, 79.
CHAPTER 32 Small Mammal Dentistry
CLINICAL PRESENTATION BY SPECIES Guinea pigs are commonly presented with symptoms directly related to and suggesting dental disease, such as labored chewing, dysphagia, reduced food intake, or anorexia. When many cavies are housed together, owners may be unaware of decreased appetite in one individual, and the only presenting sign may be weight loss. Clinical symptoms tend to be more specific than in rabbits, because guinea pigs are much less capable of masking or hiding symptoms and they tolerate dental abnormalities for much shorter periods of time.13,17 Because of the peculiar anatomy of cheek teeth, a slight alteration of the sloped occlusal plane is enough to hamper chewing, and a slight overgrowth of clinical crowns is enough to interfere with movements of the tongue and swallowing.17 The history of guinea pigs with dental disease often includes improper feeding, most particularly lack of fiber. Malocclusion of incisors often presents as excessive elongation or lateral deviation of the clinical crown of mandibular incisors. As primary incisor malocclusion is rare, it is often secondary to dental disease of cheek teeth.13,17,56 Chinchillas tend to demonstrate clinical signs and symptoms only when dental disease is advanced.13,28 The most common presenting signs and symptoms are reduced activity, food intake, and fecal production. Because dental disease often leads to ptyalism and pawing of the mouth, other presenting signs are wet fur over the mouth, chin, and fore limbs. Weight loss and emaciation are common in chinchillas with dental disease but are frequently missed by the owner because of the heavy fur coat.13,17 As in guinea pigs and rabbits, incisor malocclusion is rarely the only cause of dental disease and is usually associated with acquired dental disease of cheek teeth. Two signs of early dental disease that are often missed during routine physical examination in healthy chinchillas are epiphora and cortical bone deformities of the ventral mandible.13,17,20,29 Both are related to elongated reserve crowns and deformed apices of maxillary and mandibular cheek teeth, respectively. These abnormalities represent more advanced dental disease, as these changes are permanent and cannot be corrected. Therefore preventive diagnosis in chinchillas is based on a thorough dental examination starting at 2 years of age, before the onset of any clinical signs and symptoms. In prairie dogs, the most common presenting signs are respiratory symptoms. Reduced activity, food intake, and stool production as well as weight loss or emaciation are often present as well. True dyspnea is common, but milder symptoms are sneezing or a unique snoring sound sometimes referred to as “reverse sneezing.” Respiratory symptoms are related to dystrophic changes and apical deformaties of maxillary incisors (elodontoma), leading to reduced nasal air passage.* Like rabbits and porcupine-like rodent species, prairie dogs are obligate nasal breathers. Another common presentation, especially in prairie dogs older than 5 years, is reduced food intake and emaciation without respiratory signs and symptoms. This may be associated with other underlying medical conditions, but the primary cause is end-stage dental disease of cheek teeth with flattened crowns due to excessive wearing and advanced cavitation.13,17 Fracture and malocclusion of incisors also are common in prairie dogs but are often missed by owners.
*References 13, 17, 20, 33, 78, 79.
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Rats, hamsters, and other rat-like species are frequently presented for evident malocclusion of incisors.13 However, dental disease of incisors may missed by the owner, and patients are presented for reduced activity and food intake, emaciation, and in some cases ptyalism. Myomorph rodents generally do not display symptoms related to diseases of the cheek teeth. More common signs include facial swellings related to periapical infection and abscessation, which occasionally affect the ocular and periocular structures.13,17,56
DENTAL DISEASE Typical incisor malocclusion, where mandibular incisors elongate labially, is rarely seen in rodents. This may be because of their physiologic ability to compensate for any moderate primary incisor abnormalities through rostrocaudal movements of the jaw, which permit normal wearing of incisors.13,17,25 In guinea pigs, the most frequent cause of malocclusion of incisors is elongated crowns that develop because of maloccluded cheek teeth.9,13,17 In guinea pigs, maxillary incisors are not prone to curve palatally, while this pattern is more common in chinchillas. Both species can fracture the incisors. As in rabbits, excessive crown elongation and malocclusion of the cheek teeth is very common in guinea pigs, but with different traits. Because of the orientation of the cheek teeth, mandibular crowns always elongate lingually and maxillary crowns elongate laterally. Overgrowth of the crown is not as evident and spike or spur formation is much less frequent than in rabbits. Mandibular cheek teeth tend to bend lingually over the tongue, and a typical bridge-like malocclusion develops when the first mandibular premolars occlude or even cross each other (Fig. 32-11). The sharp border of the entire maxillary arcade can cause severe discomfort to the buccal mucosa. Even very early malocclusion and subsequent alteration of the sloped occlusal planes is enough to elicit reduced food intake or anorexia.* In chinchillas, elongated crowns of mandibular cheek teeth are not as evident as in guinea pigs. The occlusal planes can develop “step mouth” and “wave mouth” patterns in a rostrocaudal direction. Excessive elongation of the reserve crowns leads to apical deformities of both the maxillary and mandibular cheek teeth (Fig. 32-12). Typical firm swelling can be palpated on the ventrolateral aspect of the mandible. Apical deformities can also result in perforated cortical bone, with resulting exposure of the apices. On oral examination, this marked abnormal curvature appears as widened interproximal spaces on the mandibular arcades, and buccal sharp edges on the maxillary arcades. Also, elongated crowns of maxillary cheek teeth are frequently accompanied by increased height of both the alveolar crest and the gingival margin.† Caries, resorptive lesions, and fractures may also be present.27,31,56 In myomorph rodents, every pattern of malocclusion of incisors can be seen, from mild to severe. The most common is slightly elongated or fractured mandibular incisors and maxillary incisors curved toward the palate. Secondary lesions of the lips, tongue, and palate can be present, with perforation of the hard palate and resulting oronasal fistulas.9,13,20 Sciuromorph rodents (especially prairie dogs and ground squirrels) often present with fractured incisors. Fractures can occur along the clinical crown, resulting in a visibly shortened
*References 9, 13, 17, 20, 33, 56. †References 9, 13, 17, 20, 33, 56.
SECTION VI General Topics
466
S M3 M2 T
M1 T P1
A
B
T
C
Fig. 32-11 Oral endoscopy in the guinea pig. A, Normal appearance of cheek teeth, left mandibular arcade. Note the enamel crests and dentinal grooves. B, Severe coronal elongation and uneven occlusal plane of cheek teeth. Right and left mandibular CT1 cross each other in a “bridgelike” malocclusion over the tongue. C, Sharp buccal spike of the first maxillary premolar tooth following improper wearing, causing ulceration of the buccal mucosa (arrow). Spikes must be carefully detected during inspection with a curved spatula. P, premolar; M, molar; T, tongue; S, spatula. (Used by permission from Vittorio Capello, DVM.)
M3 M2 M1
T
T
P1
A
B
C
T
Fig. 32-12 Oral endoscopy in the chinchilla. A, Normal appearance of cheek teeth, right mandibular arcade. Note the enamel crests and dentinal grooves. Clinical crowns are much shorter than in rabbits and guinea pigs, and the occlusal plane is not oblique, as in guinea pigs. B, Acquired dental disease of cheek teeth. Coronal elongation, “wave mouth,” widened interproximal spaces, and a laterally bent occlusal plane are present. C, Severe coronal elongation and sharp buccal spikes of the first maxillary premolar teeth caused by improper wearing. Increase of both the alveolar crest and the gingival margin of the maxillary cheek teeth is typical in chinchillas with advanced dental disease. P, premolar; M, molar; T, tongue. (Used by permission from Vittorio Capello, DVM.)
tooth, or along the reserve crown under the gum line, resulting in a tooth that appears loose because it is attached only to the gingiva. Pseudo-odontoma is a dysplastic disease affecting the apices and the reserve crown of incisors, particularly the maxillary incisors.* A typical firm swelling of the hard palate is visible intraorally, caused by the deformed apices. Fractures and carious lesions of the cheek teeth are frequently reported in older animals. The prognosis for dental disease in rodent species is usually more guarded than in rabbits.17 However, because pathologic patterns vary among the numerous species, prognosis must be formulated for each individual case.
*References 13, 17, 20, 33, 78, 79.
Unless the animal is presented in poor general condition, prognosis for treatment of dental malocclusion of cheek teeth in guinea pigs is fair to good. Malocclusion of incisors is usually addressed as a consequence of elongated cheek teeth, and extraction is rarely needed. Herbivorous rodent species can adapt well to extraction of incisors, although not as readily as rabbits, since rodents tend to use the incisors for chewing to a greater degree than do rabbits. Unlike rabbits, guinea pigs may not improve immediately after a dental procedure because of stretching of the masticatory muscles and associated pain and inflammation. Subluxation of the temporomandibular joint may also be present. Guinea pigs that do not begin eating immediately may not wear teeth down to the degree required to prevent repeated overgrowth. These patients may require additional dental treatment
CHAPTER 32 Small Mammal Dentistry
Fig. 32-13 Three-dimensional surface rendering (shaded surface display [SSD]) of the skull of a chinchilla affected by advanced dental disease. Severe radiologic signs typical of this condition in this species—such as apical elongation, deformity, and perforation of the cortical bone—are visible. Deformed apices of right maxillary CT1 and CT2 cause subocclusion of the nasolacrimal duct and subsequent epiphora. (Used by permission from Vittorio Capello, DVM.)
until the soft tissues heal and the animal is able to eat enough high-fiber food to allow normal wearing.56 In chinchillas, prognosis is related to the stage of dental disease when diagnosis is made, because severe to end-stage dental disease is frequently diagnosed at first presentation (Fig. 32-13). In most cases, repeated dental treatments are only palliative, as restoring dental anatomy to normal is not possible. Gingival proliferation is associated with increased discomfort and more guarded prognosis. Conversely, chinchillas tolerate advanced dental disease remarkably well. Incisors can be extracted in chinchillas, but this is rarely indicated. Prognosis for elodontomas in prairie dogs is related to the stage of disease at the time of diagnosis, whether disease is unilateral or bilateral, the impact on the respiratory tract, and overall patient condition. Prognosis for end-stage dental disease of cheek teeth is usually poor. Dental disease of smaller myomorph or sciuromorph rodents carries a fair prognosis in cases of uncomplicated malocclusion of incisors. Prognosis for diseases related to cheek teeth is guarded to poor because of patient size and the difficulties related to any surgical approach.
TREATMENT OF DENTAL DISEASE Always consider the established or potential toxicity of many common antibiotics in rodent species during medical treatment.13,17,68 In hystricomorph rodents, reducing the crowns of the incisors is usually performed in conjunction with treating the cheek teeth. Always evaluate the cheek teeth thoroughly before shortening the incisors.17 Extraction of the incisors is a rarely indicated in guinea pigs and chinchillas. The technique is similar to that described in rabbits, and 21-gauge-contoured needles are used as dental elevators. To adjust the occlusal surface of the cheek teeth in guinea pigs, use the tip of a delicate, slightly abrasive burr. The goals
467
are to shorten the elongated clinical crowns and restore the proper oblique occlusal plane.9,13,17,56 In chinchillas, reducing the crowns of maxillary cheek teeth to normal length may be difficult to impossible because of concurrent elongation of the gingival margin. Crowns can be reduced to the level of the gingival margin, but additional reduction may require gingivectomy. Because of the normal curvature of cheek teeth in guinea pigs, extraction is virtually impossible unless the tooth is loose secondary to periodontal infection. Unfortunately diseased teeth fracture easily, making complete extraction extremely difficult. The same is true of mandibular cheek teeth of chinchillas, despite the fact their teeth are less curved than those of the guinea pig. Successful extraction of maxillary cheek teeth is more likely. The extraoral approach for extracting cheek teeth as described for rabbits is possible but much more difficult because of the animal’s small size. Different surgical techniques have been reported for treating psuedo-odontomas in prairie dogs. The goal of primary treatment is extraction of the affected maxillary incisor tooth or teeth.8,13,17 Mandibular incisors can be extracted during the same surgery, or a second procedure can be delayed to shorten anesthesia. Extraction is similar to that described for rabbits but is extremely challenging in this species because apical deformities and dental ankylosis are always present to some degree. The most common complication is fracture of the tooth, which represents treatment failure. In these cases, an intraoral transpalatal approach to the apices can be considered.33 An alternative approach is the dorsal approach to the apical mass by rhinotomy.33 Depending on the case, the apical mass can be removed entirely or simply debulked. A lateral approach (uni- or bilateral) to the apical mass, burring through the lateral surface of the incisive bone, has also been described and seems to be a promising surgical option.33 Palliative treatment by dorsal rhinostomy and positioning of a tubular stent can be considered when radiographs or CT scans reveal that the maxillary incisors are fractured or when the preoperative plan is to minimize risks related to anesthesia and surgical treatment.78,79 However, this surgical option does not stop the dysplastic process. For this procedure, incise the skin with a linear incision or with a biopsy punch, as the rhinostomy site and the stent opening will be round. Use a 2.5- or 3-mm intramedullary pin for rhinostomy of the nasal bones to expose the nasal cavity. Cut a small plastic catheter to the proper length and insert it into the nasal cavity just dorsal to the mass if the goal is to keep the rhinostomy site open or place it beyond the pseudo-odontoma to maintain a larger airway passage. A few weeks after surgery, remove the stent or exchange it for a smaller one to keep the skin from closing over the bony rhinostomy opening. Postoperative management of the rhinostomy site includes removing mucus and debris. Extracting the incisors in myomorph rodents is done by carefully dissecting the periodontal ligament with a small (25-gauge) contoured needle. The most frequent complications are fractures of the maxillary incisors or diastasis of the mandibular symphysis during extraction of mandibular incisors.17,23
TREATMENT OF PERIAPICAL INFECTIONS AND ABSCESSES Despite similar anatomy and physiology of the teeth and frequent perforation of the cortical bone by deformed apices in chinchillas, guinea pigs and chinchillas are much less prone than rabbits to periapical infections and osteomyelitis. One case of retrobulbar abscess secondary to molar overgrowth has
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SECTION VI General Topics
been reported in a guinea pig.69 The general surgical guidelines (excision of the capsule, debridement, extraction of the tooth involved, flushing, and marsupialization) are the same as those described for rabbits.13,17 Unlike the case in rabbits, the cheek teeth of guinea pigs always lie beneath the masseter muscle, making both surgical access and marsupialization more challenging.64 The same anatomic consideration is true for smaller species. Hamsters and rats are sometimes presented for huge abscesses involving the masseter muscle subsequent to fractures and cavities of the anelodont (rooted) cheek teeth. Periapical infections subsequent to malocclusion and fractures of incisors are also relatively frequent. Unfortunately, early diagnosis is virtually impossible in these species because clinical signs and symptoms are rarely seen before severe involvement of the bone and adjacent tissues occurs. Surgical prognosis is usually guarded to poor because of the small size of the animal and extensive involvement.
FERRETS ANATOMY AND PHYSIOLOGY OF THE SKULL AND TEETH Ferrets are strict carnivores; therefore the morphology of the skull and teeth is similar to that of larger carnivores like cats and structured to accommodate the diet.33,37,55 The oral cavity is short and wide, with a wide opening. Ferrets are anisognathic, with the mandible narrower than the maxilla, allowing shearing of food during chewing. The temporomandibular joint has a transverse articular fossa and a well-developed articular process that prevents dislocation of the mandible if the mouth is opened wide for a strong bite.55 Ferrets have 28 to 30 deciduous teeth that shed between 7 and 11 weeks of age. The dental formula of the 34 permanent teeth is reported in Table 32-1, and dentition is shown in Figure 32-14. Permanent teeth are anelodont (rooted) and brachyodont (short-crowned). Mandibular canine teeth occlude rostrally to maxillary canine teeth. Unlike other carnivorous species having 4 premolar teeth, only 3 premolars are present in ferrets because the first has been lost during development. The fourth maxillary premolar (CT3) is the most developed, has three roots, and is called the carnassial tooth. There is a single maxillary molar and two mandibular molars. The first mandibular molar (CT4) is the most developed and is the mandibular, carnassial37,55 tooth (see Fig. 32-14).
DENTAL DISEASE A grading system for dental disease in ferrets has been reported.54,55 Stage 1 is gingivitis, mostly due to plaque; stage 2 is early periodontitis, usually with gingival swelling, retraction, and pockets, but the teeth are not loose and there are no radiographic abnormalities; stage 3 corresponds to moderate periodontitis, with possible root exposure and food impaction of gingival pockets; and stage 4 is advanced periodontitis with evident gingival and bone recession. At stage 4, teeth may be loose, with more than 50% loss of attachment; pulp or enamel lesions and complications such as periodontal abscess and osteomyelitis may also be present. Diseased teeth usually appear dark. Calculus is the outcome of mineralized plaque on the tooth surface. The accumulation of plaque depends on the content and consistency of the diet, the presence of food debris, bacterial
I1 I2
I3
I1 I2 I3 C
C
P2
P2 P3
P3
P4 P4 M1
M1
M2
A
B
Fig. 32-14 Dentition of the ferret. A, Maxillary arcades, ventral view. B, Mandibular arcades, dorsal view. I, incisor; C, canine; P, premolar; M, molar. This specimen is from an old ferret and has some abnormalities, like blunting of the maxillary canine teeth and malalignment of mandibular incisor teeth. (Used by permission from Vittorio Capello, DVM.)
by-products, and the pH of saliva. The powerful chewing movements typical of carnivores seem to be a predisposing factor because it allows pocketing of food material beneath the gingival sulcus. Plaque and calculus accumulate and extend, creating gingival pockets, gingival retraction, and gingivitis. Later stages of periodontal disease are represented by periodontal infection, involvement and destruction of periodontal ligaments, and root infection. Further complications may include periapical abscessess and osteomyelitis.23,24,33,54,55 Ferrets are frequently presented for fracture of the tips of the canine teeth.55 Iatrogenic fracture was routinely performed in some countries where hunting with ferrets was popular. Pet ferrets frequently fracture the canine teeth from biting cage bars, playing with toys, or because of falls. Fractures of the tips of the maxillary canine teeth are more common than those of the mandibular teeth. Usually, the pulp is exposed after fracture of the tip, and infection is a common sequela. Teeth may be lost with advanced periodontal or dental disease or from severe trauma. Tooth loss may involve only the crown, but retained roots can be present. Periapical infections and abscesses represent a complications of either periodontal or endodontal diseases mentioned above. Local osteomyelitis is not an obligate sequela, as in in rabbits and other hypsodont species, but may occur. Congenital dental abnormalities are seen in ferrets. Supranumerary incisors are relatively common.23 Lack of the small second mandibular molar tooth has been reported.23
TREATMENT AND PREVENTION For both preventive and therapeutic dental procedures, anesthetize the ferret with general anesthesia. Removal of calculus and polishing are routinely performed with a dental scaler and a low-speed hand piece. Advise the owner to schedule this procedure regularly well before the ferret shows clinical symptoms of dental disease.55 Endodontic procedures like partial pulpectomy and pulp capping in case of pulp exposure of fractured canine teeth are difficult because of the small size of the teeth.54 These procedures require a high-speed dentistry unit and a restorative
CHAPTER 32 Small Mammal Dentistry material and are done as in other species. Refracture may occur because the biting behavior is difficult to control or prevent.53,55 For extraction of teeth or retained roots, a local block with lidocaine is a useful adjunctive analgesia. Partial gingivectomy and removal of a small section of the alveolar bone may be necessary. An 18- or 20-gauge needle can be used as an elevator. Pack the cavity of an open alveolous with bone matrix material or antibiotic gel and suture the gingiva to prevent food impaction. Administer antibiotics, nonsteroidal anti-inflammatory drugs (NSAIDs), and analgesic drugs routinely.55 Marsupialization of the infected site is not necessary. Encourage owners to institute a dental prophylactic program. Routine tooth cleaning and brushing can be performed with cotton swabs and enzymatic toothpaste. Tooth brushing may or may not be feasible depending on the ferret’s behavior.55 The impact of diet on the dentition and dental disease of pet ferrets as compared with wild ferrets and polecats is under investigation and somewhat controversial.24,54 The consumption of kibble reduces dental attrition because teeth, in particular carnassial teeth, fracture the dry food. In consuming the whole prey, carnassial teeth act like scissors, increasing and prolonging the contact. Those movements likely keep the occlusal plane sharper and prevent the formation of plaque and calculus.24 However, many other factors beyond the type of food can pose a risk for dental disease: dental bacterial flora, pH of saliva, systemic disease, higher inbreeding compared with wild counterparts, differences in skull development, and reduced muscle mass after early spay and neutering.54
HEDGEHOGS AND SUGAR GLIDERS Hedgehogs belong to the order Insectivora, but they are not strictly insectivorous. Besides invertebrates, nutrition in the wild includes eggs and fruits, making dentition more similar to that of carnivores and omnivores.33,52 All teeth are anelodont brachyodont. The dental formula is reported in Table 32-1.33 Abnormalities in tooth number have frequently been recorded in hedgehogs, including supranumerary incisors or the absence of the second pair of mandibular incisors.65 For this reason, their dental formula has also been reported with 2 x 2/2 incisors.33 The absence of the second maxillary premolar has been reported in the African hedgehog (Atelerix albiventris). The first maxillary incisors are long and widely spaced, projecting slightly forward33; for this reason, they are sometimes misinterpreted as canine. Canine teeth are small and similar to the second (or third) pair of incisors and the first premolars. Premolars show prominent cusps, more similar to those of carnivores. As in ferrets, the fourth maxillary premolar is actually carnassial tooth 3 (CT3) because of a reduction in tooth number. It has three cusps and shears against the first mandibular molar, resembling the carnassial tooth form of carnivores. The first two maxillary teeth have four cusps, resembling the molars of primates. Dental disease similar to that described for ferrets is common in pet hedgehogs. Calculus, gingivitis, gingival recession, tooth fractures, and periodontal abscesses may be present. Clinical symptoms are ptyalism, pawing at the mouth, decreased food intake, and bad breath. Diagnosis and treatment should be performed as in other species with anelodont teeth. Sugar gliders are marsupials; in the wild, their nutrition comprises plant exudates (from eucalyptus and acacia trees) and
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insects. They are also partially frugivores. Being adapted to this diet, the teeth are small with the exception of the mandibular first incisors. They are prominent and sharp and are specialized for gouging.33 Although they resemble the incisors of hamsters, the teeth are not elodont.
References 1. Aiken SA. Small mammal dentistry (Part II). In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits and rodents: clinical medicine and surgery. 2nd ed. St. Louis: Elsevier Saunders; 2004:379-382. 2. Bennet RA. Management of abscesses of the head in rabbits. Proceedings. North Am Vet Conf. 1999:822-823. 3. Böhmer E. Intraoral radiographic technique in lagomorphs and rodents. Exot DVM. 2007;9:21-27. 4. Boy SC, Steenkamp G. Odontoma-like tumours of squirrel elodont incisors-elodontomas. J Comp Path. 2006;135:56-61. 5. Brenner SZG, Hawkins MG, Tell LA, et al. Clinical anatomy, radiography, and computed tomography of the chinchilla skull. Comp Cont Ed Pract Vet. 2005;27:933-944. 6. Brown SA. Surgical removal of incisors in the rabbit. J Small Exot Anim Med. 1992;1:150-153. 7. Burling K, Murphy CJ, da Silva Curiel, et al. Anatomy of the rabbit nasolacrimal duct and its clinical implications. Prog Vet Comp Ophthalmol. 1991;1:33-40. 8. Capello V. Incisor extraction to resolve clinical signs of odontoma in a prairie dog. Exot DVM. 2002;4:9. 9. Capello V. Dental diseases and surgical treatment in pet rodents. Exot DVM. 2003;5:21-27. 10. Capello V. Endoscopic assessment and treatment of cheek teeth malocclusion in pet rabbits. Exot DVM. 2004;6:37-40. 11. Capello V. Extraction of incisor teeth in pet rabbits. Exot DVM. 2004;6:23-30. 12. Capello V. Extraction of cheek teeth and surgical treatment of periodontal abscessations in pet rabbits with acquired dental disease. Exot DVM. 2004;6:31-38. 13. Capello V, Gracis M, Lennox A. Rabbit and rodent dentistry handbook. Ames: Wiley-Blackwell (formerly Zoological Education Network); 2005. 14. Capello V. The dental suite: equipment needed for handling small exotic mammals. J Exot Pet Med. 2006;15:106-115. 15. Capello V. Management of difficult periapical infections in rabbits. Proceedings. Annu Conf Assoc Exot Mam Vet. 2007:91-97. 16. Capello V, Cauduro A. Application of computed tomography for diagnosis of dental disease in the rabbit, guinea pig and chinchilla. J Exot Pet Med. 2008;17:93-101. 17. Capello V. Diagnosis and treatment of dental diseases in pet rodents. J Exot Pet Med. 2008;17:114-123. 18. Capello V. Clinical technique: Treatment of periapical infections in pet rabbits and rodents. J Exot Pet Med. 2008;17:124-131. 19. Capello V, Lennox A, Widmer WR. The basics of radiology. In: Capello V, Lennox AM, eds. Clinical radiology of exotic companion mammals. Ames: Wiley-Blackwell; 2008:2-51. 20. Capello V, Lennox AM. Clinical radiology of exotic companion mammals. Ames: Wiley-Blackwell; 2008. 21. Capello V. Extreme periapical abscesses in rabbits. Proceedings. North Am Vet Conf. 2009:1835-1837. 22. Chesney CJ. CT scanning in chinchillas. J Small Anim Pract. 1998;39:550. 23. Church B. Ferret dentition and pathology. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. 2nd ed. Edinburgh: Saunders Elsevier; 2007:467-485. 24. Church RR. The impact of diet on the dentition of the domesticated ferret. Exot DVM. 2007;9:30-39. 25. Crossley DA. Clinical aspects of rodent dental anatomy. J Vet Dent. 1995;12:131-135.
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26. Crossley DA. Clinical aspects of lagomorph dental anatomy: The rabbit (Oryctolagus cuniculus). J Vet Dent. 1995;12:137-140. 27. Crossley DA, Dubielzig RR, Benson KG. Caries and odontoclastic resorptive lesions in a chinchilla (Chinchilla lanigera). Vet Rec. 1997;141:337-339. 28. Crossley DA, Jackson A, Yates J, Boydell IP. Use of computed tomography to investigate cheek tooth abnormalities in chinchillas (Chinchilla laniger). J Small Anim Pract. 1998;39: 385-389. 29. Crossley DA, Roxburgh G. The site of obstruction of the lacrimal drainage system in chinchillas (Chinchilla lanigera) with “wet eye.” Proceedings. Brit Small Anim Vet Assoc Congress. 1999. 30. Crossley DA. The risk of pulp exposure when trimming rabbit incisor teeth. Proceedings. 10th Ann Cong Europ Vet Dent. 2001:21-22. 31. Crossley DA. Dental disease in chinchillas in the UK. J Small Anim Pract. 2001;42:12-19. 32. Crossley DA. Oral biology and disorders of lagomorphs. Vet Clin North Am Exot Anim Pract. 2003;6:629-659. 33. Crossley DA. Small mammal dentistry (Part I). In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: Elsevier Saunders; 2004:370-379. 34. Delany MJ, et al. Rodents. In: Cogger HG, Gould E, Forshaw J, eds. Encyclopedia of animals. Barnes & Noble Books; 2002:214-218. 35. Divers SJ. Mandibular abscess treatment using antibioticimpregnated beads. Exot DVM. 2000;2:15-18. 36. Ethell MT, Bennet RA, Brown MP, et al. In vitro elution of gentamicin, amikacin, and ceftiofur from polymethylmethacrylate and hydroxyapatite cement. Vet Surg. 2000;29:375-382. 37. Evans HE, An NQ. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:19-69. 38. Fudge AM. Rabbit hematology. In: Fudge AM, ed. Laboratory medicine. Avian and exotic pets. Philadelphia: WB Saunders; 2000:273-275. 39. Gracis M. Normal dental radiography of rabbits, guinea pigs and chinchillas. J Exot Pet Med. 2008;17:78-86. 40. Harcourt-Brown FM. Calcium deficiency, diet and dental disease in pet rabbits. Vet Rec. 1996;139:567-571. 41. Harcourt-Brown FM, Baker SJ. Parathyroid hormone, haematological and biochemical parameters in relation to dental disease and husbandry in rabbits. J Small Anim Pract. 2001;42:130-136. 42. Harcourt-Brown FM. Honey to treat rabbit abscesses. Exot DVM. 2002;3:13-14. 43. Harcourt-Brown F. Textbook of rabbit medicine. Oxford: Butterworth Heinemann; 2002. 44. Harcourt-Brown FM. Dacryocystitis in rabbits. Exot DVM. 2002;4:47-49. 45. Harcourt-Brown FM. Metabolic bone disease as a possible cause of acquired dental disease in pet rabbits. Fellowship Thesis. The Royal College of Veterinary Surgeons. 2006. 46. Head KW, Cullen JM, Dubielzig RR, et al. Histological classification of tumors of the alimentary system of domestic animals. Second series. Washington DC: Armed Forces Institute of Pathology; 2003;46-57. 47. Hernandez-Divers SJ. Molar disease and abscesses in rabbits. Exot DVM. 2001;3:65-69. 48. Hernandez-Divers SJ, Murray M. Small mammal endoscopy. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: Elsevier Saunders; 2004:392-394. 49. Hernandez-Divers SJ. Clinical technique: dental endoscopy of rabbits and rodents. J Exot Pet Med. 2008;17:87-92. 50. Hoover JP, Paulsen DB, Qualls CW, et al. Osteogenic sarcoma with subcutaneous involvement in a rabbit. J Am Vet Med Assoc. 1986;189:1156-1158.
51. Jekl V, Hauptman K, Skoric M, et al. Elodontoma in a degus (Octodon degus). J Exot Pet Med. 2008;17:216-220. 52. Johnson-Delaney CA. Hedgehogs. In: Johnson-Delaney CA, ed. Exotic companion medicine handbook. West Palm Beach: Zoological Education Network; 1999. 53. Johnson-Delaney CA. A rapid procedure for filling fractured canine teeth of ferrets. J Small Exot Anim Med. 1992;3:100-102. 54. Johnson-Delaney CA. Ferret dental disorders: pictorial of common clinical presentations. Exot DVM. 2007;9:40-43. 55. Johnson-Delaney CA. Diagnosis and treatment of dental disease in ferrets. J Exot Pet Med. 2008;17:132-137. 56. Legendre LFJ. Oral disorders of exotic rodents. Vet Clin North Am Exot Anim Pract. 2003;6:601-628. 57. Lennox AM. Small exotic mammal dentistry-anesthetic considerations. J Exot Pet Med. 2008;17:102-106. 58. Lennox AM. Diagnosis and treatment of dental disease in pet rabbits. J Exot Pet Med. 2008;17:107-113. 59. Lichtenberger M, Ko J. Anesthesia and analgesia for small mammals and birds. Vet Clin North Am Exot Anim Pract. 2007;10:293-315. 60. Martinez-Jimenez D, Hernandez-Divers SJ, Dietrich U, et al. Endosurgical treatment of a retrobulbar abscess in a rabbit. J Am Vet Med Assoc. 2007;230:868-872. 61. Mathews KA, Binnington AG. Wound management using sugar. Comp Cont Ed. 2002;24:41-50. 62. Mathews KA, Binnington AG. Wound management using honey. Comp Cont Ed. 2002;24:53-60. 63. Morimoto T, Inoue T, Nakamura T, et al. Characteristics of rhythmic jaw movements of the rabbit. Arch Oral Biol. 1985;30: 673-677. 64. Popesko P, Rjtovà V, Horàk J. A colour atlas of anatomy of small laboratory animals. Vol. I: Rabbit, guinea pig. Vol. II: Rat, mouse, hamster. London: Wolfe Publishing Ltd; 1992. 65. Reeve N. Hedgehogs. London: T& A D Poyser, Ltd; 1994. 66. Reiter AM. Pathophysiology of dental disease in the rabbit, guinea pig and chinchilla. J Exot Pet Med. 2008;17:70-77. 67. Remeeus PG, Verbeek M. The use of calcium hydroxyde in the treatment of abscesses in the cheek of the rabbit resulting from a dental periapical disorder. J Vet Dent. 1995;12:19-22. 68. Rosenthal KL. Therapeutic contraindications in exotic pets. Sem Avian Exot Pet Med. 2004;13:44-48. 69. Ruelokke ML, Arnbjerg J. Retrobulbar abscess secondary to molar overgrowth in a guinea pig. Exot DVM. 2003;5:10-16. 70. Silverman S, Tell LA, eds. Radiology of rodents, rabbits, and ferrets. An atlas of normal anatomy and positioning. St. Louis: Elsevier Saunders; 2005. 71. Slayter MV, Boosinger TR, Pool RR, et al. Histological classification of bone and joint tumors of domestic animals. Second series. Vol. 1. Washington DC: Armed Forces Institute of Pathology; 1994. 72. Souza MJ, Greenacre CB, Avenell JS, et al. Diagnosing a tooth root abscess in a guinea pig (Cavia porcellus) using micro computed tomography imaging. J Exot Pet Med. 2008;15:274-277. 73. Steenkamp G, Crossley DA. Incisor tooth regrowth in a rabbit following complete extraction. Vet Rec. 1999;145:585-586. 74. Taylor M. Endoscopy as an aid to examination and treatment of the oropharyngeal disease of small herbivorous mammals. Sem Avian Exot Pet Med. 1999;8:139-141. 75. Taylor WM, Beaufrère H, Mans C, et al. Long-term outcome of treatment of dental abscesses with a wound packing technique in pet rabbits: 13 cases (1998-2007). J Am Vet Med Assoc. 2010;237:1444-1449. 76. Tyrrel KL, Citron DM, Jenkins JR, et al. Periodontal bacteria in rabbit mandibular and maxillary abscesses. J Clin Microbiol. 2002;40:1044-1047. 77. Van der Woerdt A. Ophthalmologic diseases in small pet mammals. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits and rodents: clinical medicine and surgery. 2nd ed. St. Louis: Elsevier Saunders; 2004:421-428.
CHAPTER 32 Small Mammal Dentistry 78. Wagner RA, Garman RH, Collins BM. Diagnosing odontomas in prairie dogs. Exot DVM. 1999;1:7-10. 79. Wagner R, Johnson D. Rhinotomy for treatment of odontoma in prairie dogs. Exot DVM. 2001;3:29-34. 80. Walberg JA. Osteogenic sarcoma with metastasis in a rabbit (Oryctolagus cuniculus). Lab Anim Sci. 1981;31:407-408. 81. Ward GS, Crumrine MH, Mattloch JR. Inflammatory exostosis and abscessation associated with Fusobacterium nucleatum in a rabbit. Lab Anim Sci. 1981;31:280-281. 82. Weijs WA, Brugman P, Grimbergen CA. Jaw movements and muscle activity during mastication in growing rabbits. Anat Rec. 1989;224:407-416.
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83. Weisbroth SH, Hurvitz A. Spontaneous osteogenic sarcoma in Oryctolagus cuniculus with elevated serum alkaline phosphatase. Lab Anim Care. 1969;19:263-265. 84. Weisman DL, Olmstead ML, Kowalski JJ. In vitro evaluation of antibiotic elution from polymethylmethacrylate (PMMA) and mechanical assessment of antibiotic-PMMA composites. Vet Surg. 2000;29:245-251. 85. Yasutsugu M. Mandibulectomy for treatment of oral tumors (cementoma and chondrosarcoma) in two rabbits. Exot DVM. 2006;8:18-22.
CHAPTER
33
Orthopedics in Small Mammals
Ashley Zehnder, DVM, Diplomate ABVP (Avian), and Amy S. Kapatkin, DVM, MS, Diplomate ACVS
Initial Fracture Management/First Aid Amputations Fracture Fixation Methods External Coaptation Intramedullary Pinning Bone Plating External Skeletal Fixation Repair of Specific Fracture Types Pectoral Limb Pelvic Limb Spinal Fractures and Luxations Skull Fractures Joint Luxations Fracture Healing and Postoperative Management Complications Posttraumatic Osteomyelitis Fracture Incidence
Small-mammal veterinary care has progressed considerably since the first edition of this text. The primary reasons for this are the training of specialists that care for small mammals and the increasing number of veterinarians that treat these types of pets. Many of the veterinarians treating these species are adept at both medical and surgical care for their patients. Much of what we know in small mammal orthopedics is based on our experiences rather than well-controlled clinical or scientific studies designed specifically for small mammals. Although the rabbit is commonly used as a fracture model for humans in the scientific literature, these situations do not necessarily translate into what we see clinically. Whether we are orthopedic surgeons or small-mammal specialists, our treatment of orthopedic trauma in small mammals is modeled after what we do in dogs and cats. However, we know from experience 472
that some of the orthopedic approaches and treatments used in small mammals are different from those used in dogs and cats, much as orthopedics differs between dogs, cats, and horses. Evidence or outcome-based surgery currently does not exist in veterinary medicine, yet it is a major goal of the surgical college in the near future.2,23 Even the human orthopedic college struggles with the fact that many treatments used for common orthopedic conditions are those derived from case experience and not from good clinical trials. Much of the information in this chapter is therefore drawn from both personal experiences with small mammals and established orthopedic principles. Hopefully, a team approach between small-mammal specialists and small-animal orthopedic surgeons will help to guide and encourage well-designed research on these topics in the future.
INITIAL FRACTURE MANAGEMENT/FIRST AID With the exception of ferrets, the small-mammal species commonly kept as companions are prey species. This means that they react to stress and pain differently than do dogs and cats. They may exhibit a state of tonic immobility and not show evidence of pain, even in reaction to severe injury.32 It is important to assess these animals for indicators of shock (see Chapter 38) and neurologic function and to treat them for pain and stress with antianxiolytics (e.g., midazolam), as well as opioids (e.g., butorphanol or buprenorphine). These treatments may make it possible to perform a thorough physical examination as well as to provide additional supportive care (e.g., catheter placement, bandaging). In handling all open fractures, wear sterile gloves and use aseptic technique, even if the wounds are grossly contaminated. Clip the area around the wound carefully to avoid tearing the delicate skin. A surgical clipper is the best tool for removing fur. Be patient and take care not to further complicate the wound. Carefully debride the wound and lavage the area copiously with warm sterile saline. We recommend the Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
CHAPTER 33 Orthopedics in Small Mammals use of a 35- or 60-mL syringe with an 18-gauge indwelling catheter attached to a three-way stopcock and a saline drip set for lavage and debridement, as this combination provides a pressure of 7 to 8 psi to remove debris without damaging tissues.10,49,52 However, this may not be appropriate for very small patients (<200 g); for these patients, gentle lavage may be indicated. Take care not to chill the animal, as small patients lose body heat very quickly. Remove underpads as they become saturated and provide warmth in the form of recirculating water blankets or gel packs. After lavage and debridement, appropriate wound care for the condition of the wound must be instituted.17 Ideally, take swab samples from the fracture site for aerobic and anaerobic bacterial cultures and then start broadspectrum antibiotic therapy.49 Parenteral formulations are preferred if it is unknown how well the gastrointestinal tract is functioning. External coaptation in the form of temporary splints or padded bandages may be indicated for some fractures. Manufactured veterinary splints are often too large for use with small mammals; therefore use cuttable, moldable splints or bandages for temporary immobilization.
AMPUTATIONS In some patients, orthopedic trauma is so severe that it is unmanageable and in these cases, amputation is indicated. These cases include open, infected fractures with exposed bone and compromised vasculature; fractures affecting a joint that cannot be properly aligned; or a limb with fractures involving multiple bones with a high risk of implant failure or future complications (i.e., severe degenerative joint disease, limb deformity). Limbs with pathologic fractures should be amputated, as for dogs and cats. Financial constraints of an owner may also dictate the need for amputation. Fortunately, most small mammals seen in practice will tolerate amputations. The level of amputation depends on the lesion site. It is very important to cover the amputation site with sufficient soft tissue to prevent trauma to the limb and self-mutilation. After surgery, provide extra cage padding and monitor the animal for foot lesions caused by uneven weight bearing. Because postoperative pain may induce mutilation of the amputation site, consider analgesia in planning these procedures. Use epidural analgesia and local nerve blocks where indicated, either preemptively or intraoperatively (see Chapter 31). For the thoracic limb, removing the scapula is easier and more cosmetically satisfactory than amputating at the scapular-humeral joint. Some surgeons prefer to leave the scapula because it serves as a protective barrier to the chest wall. The surgical approach to the scapula in most small mammals is similar to that in dogs and cats; however, rabbits have a prominent suprahamate process on the caudal border of the scapula. The scapula can be abducted from the body after transecting the omotransversarius, the rhomboideus, the thoracic part of the trapezius muscles, and the insertions of the latissimus dorsi, teres major, and cutaneous trunci muscles. This allows for easier isolation and transection of the thoracodorsal nerve, the axillary and the external thoracic arteries, and the brachial and axillary veins. Additionally, transect the median, ulnar, radial, axillary, subscapular,and suprascapular nerves. The scapula may then be removed and closure is routine.
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The pelvic limb can be amputated at the midfemur or coxofemoral joint. There is no evidence to suggest that mid-femoral amputation helps with balance or ambulation, and it may make the patient more prone to traumatizing the remaining limb or infection. The techniques for pelvic limb amputation are the same as those used for dogs and cats. The bone has a tendency to shatter, so use caution when the bone is cut if a mid-femoral amputation is performed. Also, transect the femur proximal to the site where the muscle is transected to allow for sufficient coverage of the remaining femur.
FRACTURE FIXATION METHODS The approach to fracture fixation in small mammals requires a basic understanding of the different types of bone healing (direct and indirect). Additionally, the surgeon must determine whether a biological approach versus an open anatomic approach is best for each fracture. How this is applied to an individual fracture and patient is determined by many factors; generally, a biological approach is used in high-energy fractures that cannot be reconstructed; anatomic reconstruction is used in low-energy, simple fractures. Other considerations are the age of the animal (which affects how rapidly bone heals and the patient’s activity level), whether the fracture is open or closed (open fractures involve a loss of blood supply to bone), if the patient has other orthopedic trauma (multitrauma cases need rigid fixation), ease of managing certain types of fixation, and the finances of the owner. Rabbits have a lower skeletal mass (8%) than cats (13%),11 yet rabbits have a haversian bone structure, like that of dogs and cats.34 Therefore, although the fracture healing process is similar, cortical bone is thinner in rabbits than in dogs or cats. Small rodents (mice and rats) have a primitive bone system; therefore, bone healing is somewhat different in them than in rabbits.34 Because guinea pigs, ferrets, chinchillas, and other rodent species discussed in this book are not used as animal models of bone healing, there is little to no information on the degree of haversian system development in these species. Many of the surgical implants that veterinarians are accustomed to using are too large for these small species. Even the smallest bone plates available from Synthes Vet (West Chester, PA; http//www.synthesvet.com) can be too large. Although several other companies make bone plates for animals, the smallest screw size, a 1.5-mm cortical screw, is often greater than 40% of the diameter of the bone and therefore too large for use.25 Because they should not exceed 25% of the bone diameter, the positive profile pins that are used with external skeletal clamp systems are too large for use in small mammals.43 Available implants are discussed under the individual headings. Regardless of the fixation method used, high-quality radiographs are essential to evaluate conditions associated with the trauma. All fractures require two views of the joint above and below the trauma site, both before stabilizing the fracture and after the fixation is applied. Additionally, obtain two-view radiographs of the whole body to determine if other injuries are present that were not observed on physical examination.
EXTERNAL COAPTATION External coaptation in small mammals can be used effectively as a definitive orthopedic treatment for closed fractures. These methods work best for simple fractures affected by bending and
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rotational forces because they often interdigitate with manipulation and remain reduced. Fractures affected by compressive or shear force, such as short oblique fractures, are poorly immobilized with external coaptation.13 Animals will have to be anesthetized or heavily sedated to allow for proper alignment and prevent further injury. In trying to reduce a fracture in these small animals, take care not to damage soft tissues and the blood supply. External coaptation does not achieve rigid fixation; therefore maintaining the blood supply is paramount to ensure healing. For a good result, it is important to achieve good alignment of the joints and at least 50% cortical contact visible from two radiographic views. Apply standard splinting principles: maintain the affected limb in normal position, immobilize the joint above and below the fracture, and use a bandage and splint that can maintain adequate reduction until bony union (Fig. 33-1). At each splint change, radiograph the limb after placing the splint to ensure adequate reduction. Once the fracture has developed a soft callus with inherent stability, the limb will not have to be radiographed after each change. Standard materials used for splints and casts in dogs and cats can be applied in managing fractures of rabbits, ferrets, and other small mammals. Several products work well for splinting small mammals, such as Orthoplast (Johnson & Johnson, New Brunswick, NJ), and Vet-lite (Runlite SA, Micheroux, Belgium; http://www.runlite.com/vet-lite/). These are materials that are very lightweight, thin, easy to use, and biomechanically strong. They are malleable when heated in boiling water and conform to any shape when molded and allowed to cool. These products work best when used in a half cast with a light modified Robert Jones bandage, avoiding excessive pressure on the limb. Although these materials cost more than routine casting materials, the final cost is reasonable because only a small piece of material is used. Vet-lite can be layered so that splint thickness and stiffness can be increased if desired. These products are very resistant to chewing by the animal and will not cause oral trauma or break apart and cause gastrointestinal obstruction. Encircling fiberglass casts are difficult to place without causing pressure sores and irritation, but they can be used with care. Fiberglass casting material can be used in strips to make a splint, or they can be bivalved; however, the plastic materials, like Orthoplast, are easier to use and maintain. Monitor animals with splints and casts weekly to prevent complications. Common complications include chewing at the
A
splint; soiling, which leads to the formation of pressure sores or local skin infections; and swelling, which results from excessive activity or from the splint being too tight. Joint laxity and stiffness from the coaptation are also seen.50
INTRAMEDULLARY PINNING Intramedullary (IM) pinning can be an effective method of internal fixation for small mammals. The small diameter K wires (pin diameters of 0.9-1.6 mm) will fit many of these species. Ideally, the intramedullary pin should fill about 70% of the diameter of the medullary cavity. In one rabbit study, an IM pin that was 75% of the diameter of the bone caused a fracture when being inserted.2a Using a pin that is less than 70% of the bone diameter is unlikely to efficiently control bending unless an external skeletal fixator (ESF) is added to the construct. Ideal pin sizes for small mammal bone are not known. Intramedullary pins primarily resist bending forces, and the recommended pin diameter should provide sufficient stiffness and strength for the fracture.42,43 Because intramedullary pins do not counteract other forces on fractures—such as rotation, tension, and compression— many of the diaphyseal fractures in small mammals will need additional support. External skeletal fixators, separately or in a tie-in configuration, or external coaptation may be combined with IM pins (Fig. 33-2). Because these are small bones, combining apparatuses may not work as well in small mammals as they do in larger domestic species. In rabbits, fractures occurred from pin insertion of both the IM and ESF pins.2a Therefore choose an intramedullary pin as the fixation method when bending is the major force acting on the bone—for example, in spiral or long oblique fractures. Cerclage wires can be used for spiral and long oblique fractures to help counteract shear forces. One or more K wires can be used alone for fractures at the physis or metaphysis. If the fracture fragment has tension forces on it from ligaments or tendons, wire used as a tension band must be added. The technique for pins and tension-band wires requires two pins, yet only one may fit. However, many of these small mammals will do well with only one pin despite the fact that rotational forces may not be counteracted. Biodegradable implants are now available that provide adequate mechanical properties yet do not need to be removed and may stress-shield bone less than stainless steel. After implantation,
B
C
Fig. 33-1 Splinting of a fractured tibia/fibula in a chinchilla. A, Tape stirrups placed on the fractured limb. B, Padding complete. C, Completed splint.
CHAPTER 33 Orthopedics in Small Mammals
A
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B
Fig. 33-2 Comminuted middiaphyseal femoral fracture in a rabbit. A, Preoperative view. B, After application of an intramedullary pin and external skeletal fixator.
biodegradable intramedullary pins are designed to dissolve into water and carbon dioxide, like polylactides, after a period of time.3,37 These would be excellent features for an implant in small mammals. Some research on these products has been done in a rabbit osteotomy model.44 Although results showed adequate stiffness for healing in this model, clinical fractures in small mammals have both biological and biomechanical features that may affect the outcome. Whether these implants would be stiff enough for clinical use in small mammals is unknown. Interlocking nails are not made small enough for small mammals at this time and are not discussed in this chapter.
BONE PLATING In certain situations, the use of a plate is appropriate for fracture repair in a rabbit or ferret. Because of the conformation of these species, a fracture of the humerus or femur can be difficult to manage with an external fixator. The soft tissue coverage over the femur and humerus is considerable, and external skeletal fixation can be uncomfortable when it is used in this location. If plating is chosen, the implants commonly used are 1.5- and 2.0-mm cuttable plates (Synthes, Ltd., Wayne, PA or Veterinary Instrumentation, Sheffield, UK). These offer the surgeon flexibility in choosing the appropriate length, and the plates can be stacked for added stiffness. The limited-contact dynamic compression plates (LCDCP) or the dynamic compression plates (DCP) that would be long enough to bridge the fracture are too large and stiff for use in these small species. Although DCP and LCDCP plates are made in 2.0-mm size, they are short and therefore may not be the appropriate length for a weightbearing bone. A recent biomechanical study in rabbit bone had similar results; screw insertion caused cortex fractures. The 6-hole, 2.0-mm DCP in rabbit femurs failed at low loads, usually at the plate or the plate-bone interface. The authors of the study acknowledged that the short plate contributed to the low
compressive load in this transverse fracture model.2a The principles of applying AO bone plates are covered in other texts and are beyond the scope of this chapter.25,42 Bone plating in these species presents some specific problems. Clinically, the bones of rabbits have extremely thin cortices, which make screw placement difficult without stripping the cortex. One of the present authors (AK) has experienced complete collapse of the bone while trying to place a screw. In a clinical setting, rabbit fractures are usually comminuted. The blood supply to the bone is already compromised, predisposing the rabbit to osteomyelitis and nonunion. Placement of a bone plate with removal of the remaining periosteum and soft tissues only worsens this problem. Plating in these species often overprotects a fracture, preventing load sharing and causing subsequent delayed healing or nonunion.26,47 If a plate is used, the animal may require staged plate removal, which is costly and unpopular with owners. Another disadvantage is the high cost of equipment and implants. The new locking compression plate (LCP) (Synthes Vet) uses the concept of external fixation internally near the bone. The stability is from the screw locking into the plate instead of the screw compressing the plate to the bone. Therefore LCPs can be placed without contouring, over the periosteum, and minimally invasively so that rigid fixation is combined with preservation of blood supply to the bone. However, the smallest LCP is 2.0 mm, which may still be too large, depending on the animal. Locking screws have excellent holding power even when placed unicortically, and this may eliminate the complication that was observed in the one recent study on rabbit bone.2a,24,25,30,46 The LCP functions differently than conventional plates and may prove useful in small mammals. To date, there are no reports of the use of LCPs in small mammals. Like biodegradable pins, biodegradable plates and screws were evaluated in the 1980s in rabbit osteotomy models; however, the perfect balance of a biologically sparing and
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mechanically stiff implant does not yet exist.37 Biodegradable implants are used clinically only on non-weight-bearing bones.44,45 It is unknown whether these implants will be useful in small mammals.
EXTERNAL SKELETAL FIXATION External skeletal fixation is probably the most common method of repairing fractures in small mammals. External fixators provide rigid stability, can be adapted for use in these small species, and cause minimal soft tissue damage. Basic principles for applying external fixators are similar in all species. The pins should be inserted with low rotational speed (150-300 rpm). Manual insertion causes wobble and leads to premature loosening of the pins. High-speed drilling can cause bone necrosis, which also leads to the premature loosening of pins.14,15 Smooth pins have poor holding power and tend to loosen prematurely. Negative-profile threaded pins tend to break at the thread-nonthread interface and are rarely used now that positive-profile pins are available in small sizes.28 Predrilling is the correct method for inserting threaded (positive-profile) pins.28 Imex (IMEX Veterinary Inc., Longview, TX; www.imexvet.com) produces a miniclamp and bar system that can hold positive profile K-wires as small as 0.035 in. (0.9 mm). Many small mammals are too tiny for the connecting bars that are 1⁄8 in. (3.2 mm) in size, but a smaller acrylic or polymethylmethacrylate (PMMA) connecting bar can be used with the Imex miniature INTERFACE pins (IMEX Veterinary, Inc.). The miniature Imex pins are designed as positive-profile pins with a central area that is roughened only to allow acrylic bonding to it. The roughened area is not designed for placement in the bone. With the design of external fixation clamps that are biomechanically better and the use of positive-profile pins, type II (bilateral-uniplanar) and type III frames (bilateral-biplanar) have become almost obsolete. In large species or specific fracture locations, frames other than a type Ia (unilateral-uniplanar) or Ib (unilateral-biplanar) are sometimes indicated, but this is rare in small mammals. The use of an IM-pin tie-in frame is useful in some of the larger small mammals, especially when the bending forces are large and repair with an ESF alone may not counteract the forces successfully. In small species, the use of a tie-in frame is constrained by the limited space to place the ESF pins past the IM pin. Before repairing a fracture with an ESF, decide whether a limited open approach or closed placement of the pins is best. In species with limited soft tissue covering the bone, it is advantageous not to destroy the tissues by opening the fracture. Therefore, in simple fractures, place the external fixator with a closed technique. With highly comminuted or open fractures that need debridement, use a limited open technique to reduce the fracture and place the pins. Mechanical studies have tested pin placements and configurations for dogs, and some may apply to small mammals.4 Place the fixator pins approximately 1 cm away from the fracture line. Four pins per bone segment are ideal for maximum stiffness: additional pins increase stiffness insignificantly. However, in these small species, four pins provide too much stiffness and probably will not fit in the fracture segments; therefore we rarely use more than three pins per segment. The fixator pin’s diameter should not exceed 25% to 30% of the bone diameter. Position the connecting bar approximately
0.5 to 1 cm away from the skin. Placement of the bar too far away from the skin decreases the stiffness and strength of the external fixators. Remember to allow for swelling in placing the bars. Spread the pins evenly across the fracture segment to achieve maximum strength. Also remember to place the pins through a separate stab skin incision and not through the fracture opening or original incision.4,15,28 Results of one biomechanical study in rabbits demonstrated that an IM pin-ESF construct can cause fractures on pin insertion and that this construct had less than ½ the compressive and bending strength of normal rabbit bone.2a Bone healing in rabbits with fractures stabilized by external fixation has been well studied as a research model. The current recommendation for external fixation in dogs and other research models is to stage the removal of the apparatus so that bone is allowed to take over load bearing at an early stage of fixation. This process has been termed dynamization, or staged disassembly.28 In studies of rabbits with tibial osteotomies, the ideal time to remove external fixators was 6 weeks.48 The strength and stiffness of the bone were greater if the fixator was removed at 6 weeks than if it was left in place for 12 weeks.48 Clinically, many factors affect optimal removal time of a fixator, such as the age of the animal, the type of fracture, and the degree of disruption of vascular structures. Therefore the key is to examine these patients every 2 to 3 weeks after surgery and to stage removal when there are indications that the fracture has regained normal stiffness, even though strength may not be normal. Experimentally, removing the fixator at 4 weeks causes some loss of reduction and is probably not advisable, even in perfectly reduced fractures.48
REPAIR OF SPECIFIC FRACTURE TYPES Generally, because the anatomy of small mammals resembles that of dogs and cats, small-animal orthopedic surgery texts may be used as guides for surgical approaches as well as for additional information on fixation techniques for fracture types not discussed here. Where relevant, important differences are discussed below, and anatomic texts are available for some species.9,21,39
PECTORAL LIMB Scapula Scapular fractures are uncommon but can occur, often secondary to significant trauma involving the chest wall. Concurrent broken ribs or pulmonary complications are common. Treatment recommendations will vary by the fracture location (body and spine, acromion, neck, glenoid or supraglenoid tuberosity). Fractures not involving the articular surface and that are minimally displaced heal acceptably with exercise restriction. Non-weight-bearing slings (Velpeau) may not be tolerated by all patients. Fractures involving the neck or articular surface will lead to osteoarthrosis if they are not properly aligned. For these fractures, rigid internal fixation or amputation is recommended. Anatomic Considerations. In rabbits, a prominent suprahamate process is present on the caudal aspect of the acromion in addition to the hamate process. The axillary artery should be avoided medially as well as the subscapular, suprascapular, ulnar, and radial nerves.
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477
Table 33-1 Summary of Types of Orthopedic Trauma (Fractures/Luxations) in Small Mammals Treated Between January 1998 and July 2011 at the College of Veterinary Medicine, University of California at Davisa SPECIES RABBIT Anatomic Location
Fracture Site
Total #
GUINEA PIG %
Total #
Appendicular Scapular 1 1.9 0 Humeral 4 7.7 0 Radius/ulna 5 9.6 1 Pelvic 1 1.9 0 Femoral 14 26.9 2 Tibial/fibular 9 17.3 3 Multiple 0 0 bones Spinal Cervical 1 1.9 0 Thoracic 1 1.9 0 Lumbar 4 7.7 1 Skull Multiple 0 0 Mandibular 4 7.7 0 Maxillary 0 0 Luxation Spinal 2 3.8 1 Elbow 1 1.9 0 Tarsal 3 5.8 0 Coxofemoral 2 3.8 0 52 (67.5%) 8 (10.4%) Total Fractures/ Luxations (%)b Total patients/ 1998-2011 1293 326 species Cumulative 1998-2011 3.9% 2.5% incidence
CHINCHILLA
HAMSTER/RAT
%
Total #
%
Total #
%
12.5 25 37.5
0 1 1 2 1 3 0
10 10 20 10 30
0 0 0 0 0 2 1
40 20
12.5 12.5 -
0 1 0 0 1 0 0 0 0 0 10 (13%)
10 10 -
FERRET
TOTAL
Total # % Total # 0 0 0 0 0 0 0
0 0 0 1 0 0 2 40 1 0 0 0 0 0 0 0 0 0 0 0 0 5 (6.5%) 2 (2.6%)
169
542
166
5.9%
0.9%
1.2%
%
-
1 5 7 3 17 17 1
1.3 6.5 9.1 3.9 22.1 22.1 1.3
50 50 -
1 3 5 3 5 0 3 1 3 2 77
1.3 3.9 6.5 3.9 6.5 1.3 3.9 2.6 (100%)
aNote
that ferrets were only seen after 2005 and these numbers may not be representative of practices that see a larger proportion of ferret cases. bOne animal had multiple fractures.
Types of Fixation. Fixation methods are K-wires in a crosspin fashion, small cuttable plates, or tension banding or lag screws for the repair of articular fractures.
pins with or without external fixators (generally type Ia); external fixators on the lateral aspect of the limb; and lag screws for distal humeral articular fractures.
Humerus
Radius/Ulna
Fractures of the humerus most commonly occur in the diaphysis in dogs and cats; this is consistent with the fractures seen in rabbits at this institution (4/4, 100% at the diaphysis).51 These fractures result in a non-weight-bearing lameness with the limb dropped at the elbow. Temporary or permanent radial nerve damage, in our experience, is uncommon in these species. If there is persistent loss of withdrawal or lack of pain sensation in the limb, amputation is recommended. External coaptation is not recommended because it is difficult to immobilize the scapulohumeral joint. Anatomic Considerations. The brachial artery and vein and the median, musculocutaneous, and ulnar nerves are located medially. The radial nerve runs laterally along the distal third of the humerus to lie superficial to the brachialis muscle and underneath the lateral head of the triceps muscle. Types of Fixation. Acceptable fixation methods are bone plating on the lateral, cranial, or medial surface; intramedullary
Fractures of the radius and ulna are common (Table 33-1) and usually occur together. These fractures are more likely to be open because the soft tissue coverage in this area is poor. This should be considered in the choice for fixation. Fractures in young animals that affect the growth of either the radius or ulna can cause angular limb deformities. If they occur, early surgical intervention to prevent asynchronous growth is indicated. Anatomic Considerations. The median, ulnar, and radial arteries are present on the caudal aspect of the limb as well as the ulnar nerve. Types of Fixation. External coaptation can be used for minimally displaced fractures or those with either the radius or ulna intact (Fig. 33-3). Extending the splint from the distal limb to the midhumerus is critically important in order to provide appropriate support. Because an IM pin can compromise either the proximal or distal joint during placement, it can be used for
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A
B
Fig. 33-3 Middiaphyseal radial/ulnar fracture in a rabbit. A, External coaptation with poor alignment of the fracture ends. Note that the end of the temporary splint is distal to the proximal aspect of the ulna. B, The fracture was realigned with the animal under heavy sedation. Note the improved alignment of the fracture ends and that the splint extends from the toes to the midhumerus. The splint is slightly long and was trimmed to allow the toes to be visualized.
ulnar fractures but is never indicated for radial fractures. External fixators of types Ia, Ib, or II can be used; type Ia is usually sufficient and is placed on the medial, cranial, or lateral surface, depending on where the fracture is located.
Metacarpals These fractures are uncommonly seen in small mammals but may be managed with external coaptation. Although malunion is common, it is rarely clinically significant. The bones are too small for definitive fixation.
PELVIC LIMB Pelvis Pelvic fractures (sacroiliac, ilial wing, ilial body, acetabular, ischial, pubic, or pelvic symphysis) usually involve multiple bones because of the box configuration of the pelvis. The animal should be carefully evaluated for spinal trauma, nerve damage affecting the pelvic limbs, as well as damage to the urinary tract because of the significant trauma associated with these injuries. Complications may involve altered urination or defecation if caudal nerve roots are affected. Always consider surgical fixation for an animal with a severely compromised pelvic canal, acetabular fractures, or ilial fractures that compromise the sciatic nerve. Conservative management is indicated for minimally displaced ilial, ischial, or pubic fractures or sacroiliac luxations. Complications of malunion may include colonic stricture if the pelvic canal is compromised. Healing time for animals managed conservatively is much like that for other mammals, and significant healing should occur by 6 to 8 weeks. Monitor rabbits, guinea pigs, and chinchillas closely for signs of gastrointestinal stasis if exercise is restricted for long periods.
Anatomic Considerations. The surgical approach to the hip depends largely on the bones involved and the surgical plan. Take care to prevent damage to the ventral nerve roots, the sciatic nerve, and the internal iliac artery and its branches during repair of the pelvis. Types of Fixation. Bone plates can be used for larger rabbits and ferrets. Other methods are interfragmentary wiring and pin-and-wire techniques. The type of fixation required will vary greatly by the type of fracture. Consult an orthopedic text for more detailed descriptions of repairs.
Femur Femoral fractures are common and often significantly displaced because of the strong musculature in this area. Because Spica-type splints are not tolerated, external coaptation is not acceptable in these animals. Additionally, significant malunion will occur with limb angulation or shortening that limits ambulation. However, some fractures will heal with prolonged strict cage rest (Fig. 33-4). With these fractures, evaluate the sciatic and femoral nerves by testing deep pain perception in the lateral and medial aspects of the digits, respectively. Anatomic Considerations. The lateral approach to the femur is most common for diaphyseal fractures. Use extreme care to avoid the sciatic nerve, which is located caudal to the femoral diaphysis and deep to the biceps femoris muscle. Additionally, the popliteal and caudal femoral arteries are present caudal to the stifle. Types of Fixation. For fixation, IM pins can be placed either normograde or retrograde. Normograde insertion at the trochanteric fossa allows more lateral pin placement and easier avoidance of the sciatic nerve. Type Ia external fixators are generally placed on the lateral aspect of the limb. These may be
CHAPTER 33 Orthopedics in Small Mammals
A
479
B
Fig. 33-4 A, Radiographs of a middiaphyseal femoral fracture in a rabbit. Note the congenital malformation of the stifle joints bilaterally. B, Repeat radiographs 6 months after strict cage rest. A callous is noted at the fracture site and there is remodeling of the fracture ends.
placed alone or in combination with an IM pin (see Fig. 33-2) and can be tied in. Bone plates may be used as described above for rabbits and ferrets.
Tibia/Fibula In our experience, tibial fractures are common in small mammals, particularly in guinea pigs and chinchillas (see Table 33-1). Depending on the location and degree of displacement, these may be amenable to external coaptation (see Fig. 33-1). The area of the proximal tibia in small mammals, particularly rodents, is characterized by an abrupt change in the diameter of the limb distal to the stifle. This makes it difficult to properly apply splints to the lower limb. Additional padding is needed for the bandage to be cylindrical. If the splint is not placed far enough proximally (at least to the distal third of the femur), the splint may slip distally and act as a fulcrum at the fracture site. A lateral-cranial splint or a combination of caudolateral splint may be applied. Anatomic Considerations. Avoid the saphenous artery, vein, and nerve on the medial aspect. Types of Fixation. For fixation, IM pins can be inserted normograde from the cranial aspect of the tibia, on the tibial crest. Type Ia external fixators may be applied medially and may be used in conjunction with an IM pin (Fig. 33-5). Generally, type II fixators are not required for stabilization. Plates are difficult to use because of the thinness of the bone in most small mammals as well as the lack of good soft tissue coverage.
Metatarsals These are relatively uncommon fractures in small mammals and may be treated as described above for metacarpal fractures.
SPINAL FRACTURES AND LUXATIONS Spinal fractures can occur from trauma in any small mammal. Falls, cage trauma, or dog bites are common causes in companion small mammals. In rabbits, improper handling is another cause of vertebral fractures or luxations (see Chapter 19).35 There are no large case series describing surgical treatment results after spinal fracture or luxation in small mammals. In eight cases of spinal trauma in rabbits presented to UC Davis Veterinary Teaching Hospital over the past 10 years, all but one were euthanized because of the severity of the neurologic dysfunction and therefore poor prognosis for recovery. Three rabbits had cervical trauma: one had a fracture of the C7-T1 junction and two had cervical luxations. One rabbit had a T5-6 fracture, two rabbits had lumbar fractures at L3 and L5, respectively and two had L7 fractures. Four of the eight rabbits were nondomestic and had injuries from traps or other trauma. The other four rabbits had cage-related injuries. Of the four other small mammals with spinal fractures in the database, three (a ferret, a guinea pig, and a chinchilla) were euthanized because of their injuries. The ferret had a T11 fracture from a dog bite. The guinea pig was dropped and had an L1-2 luxation. The chinchilla fell off a countertop and had a T13 fracture and complete paralysis. The surviving animal, a guinea pig, had a chronic L6 fracture, found on a routine health check that included whole-body radiographs. Although this guinea pig had spinal pain, the fracture had healed and the animal did well. The cause of the fracture in this animal was unknown. Even though surgical intervention in small mammals that have some neurologic function intact is feasible, results and prognosis on success of surgery and recovery is unknown. Because of the size of small mammals, use of the small positive-profile K-wires and an acrylic connecting bar either internally or externally is suggested.
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A
B
Fig. 33-5 A, Postoperative radiographs of a midtibial fracture in a rabbit with an intramedullary pin tie-in. B, Repeat radiographs approximately 6 weeks postfixation.
SKULL FRACTURES There is very little published information describing treatment of skull fractures in small mammals. These generally occur secondary to severe trauma, and the animal should be carefully assessed for other lesions. A perfectly positioned skull series or computed tomography (CT) scan done while the animal is under anesthesia is recommended to fully assess the injuries (see Chapter 35). For animals such as guinea pigs, chinchillas, and rabbits with aradicular hypsodont dentition (long crown without closed root) and continuously erupting teeth, the longterm complications of malocclusion secondary to skull trauma should be clearly explained to the owner. These animals may require dental adjustments (under anesthesia) every 6 to 12 weeks for life. For rats and mice, this applies to the incisors only. Fractures that do not affect occlusion or significantly reduce the opening of the nasal passage may heal well with conservative management (rest and analgesics). Animals that are reluctant to eat will require placement of an esophageal feeding tube. Nasogastric tubes are inappropriate for long-term management of hind-gut fermenters because it is difficult to provide the fiber necessary for normal digestion and the tubes are not well tolerated. Esophageal feeding tubes are placed as for dogs and cats. Types of Fixation. Many of the therapies used in dogs and cats for skull fractures apply to small mammals. These include interdental appliances and wire fixation. We strongly recommend consulting a veterinary dentist or orthopedic surgeon before fixation of a skull fracture that affects dental occlusion.
JOINT LUXATIONS Elbow luxations occur occasionally in ferrets and rabbits.41 These cases present with a non-weight-bearing lameness and a very painful, swollen elbow joint. Many elbow luxations need internal fixation to heal. For internal fixation, place a transarticular pin through the joint and then place a light Orthoplast or Vetlite splint. The transarticular pin keeps the joint reduced while
the splint prevents full weight-bearing by the animal. The pin must be removed in approximately 3 weeks. The splint is usually needed for support for another 2 to 3 weeks after pin removal. Elbow luxation repair is usually successful with this technique. Another option is to use a transarticular ESF with positive-profile pins and acrylic connecting bar extending from the humerus to the distal radius. We prefer the transarticular pin technique because it poses less risk of iatrogenic fracture of the bones than the transarticular ESF technique. Because of the size of the collateral ligaments and the thinness of the joint capsule, primary repairs of these structures alone rarely keep the elbow reduced. Luxations of some of the other joints can be managed in the same way as the elbow luxation. Shoulder luxations are rare and can be managed with a transarticular pin technique. Coxofemoral luxations are also rare and, although many techniques are successful in keeping the femoral head reduced in larger domestic pets, many of the techniques require implants that are too large for this species.20 Coxofemoral luxations can be treated with femoral head and neck excisions; many small mammals do well with this procedure. Tarsal and carpal luxations occur and often present severely displaced. Primary repairs of the ligaments pose the same difficulty as is seen in repair of the elbow. External coaptation, allowing ankylosis of the joints, can be successful (Fig. 33-6). The patient will develop significant degenerative joint disease; yet clinically, most small mammals do well. Displaced carpal and tarsal joints can be arthrodesed with plates or transarticular external fixation, but the apparatuses used will fit only the larger small mammals. Small K-wires can be used for arthrodesis in conjunction with splints. See orthopedic texts for arthrodesis techniques. External coaptation for luxated joints requires reducing the joint and placing a splint made from Orthoplas, Vet-lite, or an aluminum rod (Fig. 33-7). For the elbow, keep the limb in extension while the soft tissues fibrose to keep the elbow in place. Place carpal and tarsal splints with the joints in a functional position.
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A
481
B
Fig. 33-6 A, Lateral radiograph of an intertarsal luxation (arrow) in a rabbit. B, Repeat lateral radiograph after 8 weeks of external coaptation. Note the extensive periarticular bony proliferation (arrow).
Fig. 33-7 A rabbit with a splinted tarsal luxation.
FRACTURE HEALING AND POSTOPERATIVE MANAGEMENT Bone healing in rabbits, ferrets, and small rodents is the same as in other mammals. There are two major categories of bone healing: direct and indirect. Direct bone healing is osteonal reconstruction and occurs when there is anatomic reduction and rigid stability of the fracture. Direct healing takes place by either contact healing or gap healing. With contact healing, bone union and haversian remodeling occur together. Gap healing takes place when a gap smaller than 800 mm is present. Layers of bone are laid down on two surfaces of the fracture. This transverse lamellar bone is then remodeled longitudinally through osteons and haversian remodeling. Indirect bone healing involves the stages of inflammation, soft callus, hard callus, and then remodeling. For further details on bone healing, refer to other texts.22 Postoperative management of small mammals involves confinement, good husbandry, and frequent monitoring and
maintenance of the fixation apparatus. Animals with closed fractures need only perioperative antibiotics (a dose given intravenously every 2 hours during surgery); but open fractures may require extended antibiotic therapy. Ideally, the antibiotic is chosen according to the sensitivity pattern of the bacterial culture taken initially. Some of the techniques used in managing open fractures in dogs and cats are not feasible in small mammals. The placement of drainage systems or treatment of open wounds daily with sterile flushes and bandages is extremely difficult. Animals chew excessively at bandages or will reach the wound unless a hard splint, made of one of the discussed casting materials, has been placed on the limb. In some species, daily flushes can be managed only with heavy tranquilization or anesthesia. This is expensive and stressful, especially for rodent species. Therefore wound management should be definitive at the time of surgery if at all possible. Whenever a splint is used, review bandage care with the owner. Have the owner check the toes for swelling or discharge. These species often chew at splints even if they appear comfortable. Extreme discomfort in these animals often manifests as quietness and anorexia. Recheck the splints every 1 to 2 weeks, with the first recheck scheduled within a few days of splint placement. For fractures repaired with one of the above fixation methods, evaluate the fracture radiographically in 4 to 6 weeks. In these species, this usually requires heavy sedation or anesthesia. Because radiographic findings often lag behind clinical healing, good palpation of the fracture is also important. Depending on the type of fixation, changes can be made in the apparatus as healing becomes evident on radiographs.
COMPLICATIONS The ultimate goal of fracture repair is bony union. Unfortunately this does not always occur despite our use of sophisticated techniques, resulting in delayed union, nonunion, or malunion. Please see orthopedic textbooks for specifics of each condition.31 Because of their small size, light body weight, and
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SECTION VI General Topics
locomotive stance, these animals seem to tolerate severe deformities with minimal clinical significance.
POSTTRAUMATIC OSTEOMYELITIS Posttraumatic osteomyelitis is another potential complication of fractures in rabbits, ferrets, and small mammals. This posttraumatic or postoperative infection of the bone is caused by wound contamination and bone avascularity. An infected wound needs more than 1 × 105 organisms per gram of tissue to induce posttraumatic osteomyelitis.5,56 Therefore good surgical debridement and copious lavage are very important in managing open fractures. Adherence to strict sterile surgical technique, careful wound handling, hemostasis, and anatomic closure helps prevent posttraumatic osteomyelitis.5,6,56 Radiographically, rabbits with posttraumatic osteomyelitis show periosteal reactions and osteolysis.55 Rabbits develop caseous, nondraining abscesses; thus typical drainage techniques are not useful.34 Treatment of osteomyelitis in rabbits must be aggressive. Collect samples for aerobic and anaerobic cultures, debride the wound, and administer an appropriate antibiotic. Additionally, polymethylmethacrylate (PMMA) antibiotic-impregnated beads are recommended to avoid antibiotic inactivation by the loss of blood supply to the infected, traumatized area.12,16,29,33,54 Many in vitro studies show varied results depending on the antibiotic used, the form used (powder vs. liquid), the antibiotic concentration, the carrier medium (PMMA, hydroxyapatite cement, calcium sulfate, and biodegradable polymers), and the carrier medium’s surface area.* The use of multiple antibiotics within a single bead has been shown to significantly increase the rate of elution.38,40 Certain antibiotics can delay the polymerization of PMMA.1,40 Local tissue conditions—including seroma formation, fibrous tissue, and blood supply—also affect elution.5-7 Many recent studies have shown efficacy in vivo and in vitro with a variety of antibiotics (cephalosporins and aminoglycosides). These studies demonstrate a bimodal elution, with significantly higher elution occurring within the first 48 hours and then continued elution from 2 weeks to months.† Antibiotic-impregnated beads are being used clinically in rabbits for osteomyelitis, with perceived success in some cases.8 Because antibiotic beads may continue to elute over an extended period, it may be advantageous to leave them permanently at the infection site.16,19 Also, many owners do not want a second surgery to remove the beads. However, beads can potentially act as foreign bodies and harbor bacteria once the antibiotic concentration decreases.7 In using impregnated beads, use an appropriate antibiotic and dose. Higher doses elute more antibiotic1,16,40,54; however, systemic levels of drugs are often undetectable or present only initially.1,7 Use caution if beads contain high levels of antibiotics known to elicit enteritis in rabbits and other hind-gut fermenters.36 Beads can be made with special bead molds (usually too large for small mammals) or injected into syringes and then cut to an appropriate size or placed on the suture. Some compounding pharmacies will prepare impregnated beads to order. All uses must be performed under sterile conditions. A similar type of abscess can develop in small rodents, but the incidence of posttraumatic osteomyelitis appears to be low.
*References 12, 16, 18, 27, 29, 33, 53-55. †References 1, 12, 16, 19, 29, 33, 38, 40.
Ferrets with posttraumatic osteomyelitis have manifestations similar to those of dogs and cats. Therefore treatment and the use of drainage systems can be similar. In severe cases, antibiotic-impregnated beads can be used as well. When treatment for posttraumatic osteomyelitis is unsuccessful, which is common in rabbits, amputation of the limb is the only option. This is a salvage procedure that is reserved for only those animals for which prognosis for successful treatment is very poor.
FRACTURE INCIDENCE Because of the lack of published reports of fracture incidence in small mammals, we performed a retrospective analysis of small mammal orthopedic cases seen between January 1998 and July 2011 at this institution (UC Davis) (see Table 33-1). Only two ferrets were seen in this period for orthopedic conditions because they were only seen at this institution after 2005 and are illegal to own in the state of California. Seventy-six animals presented during this time period with orthopedic injuries, the majority being rabbits. However, as a percentage of total cases seen, chinchillas had the highest prevalence of fractures at 5.9%. Consistent with other reports, tibia/fibular fractures were most common (30% of cases). Interestingly, in rabbits, of the 52 fractures presented, surgical fixation was pursued in only 6 cases, splinting or bandaging in 6 cases, amputation in 9 cases, and euthanasia in 16 (including all of the spinal fractures except 1 managed conservatively). The remainder were managed conservatively with supportive care. It was not possible to make conclusions regarding the outcomes of various treatments with such small numbers, but it does appear that very few animals are being treated with primary surgical fixation compared with more conservative measures. This trend is similar in the other small mammal species. When guinea pigs, chinchillas, hamsters and rats are combined, only 2 had surgical fixation, 2 were managed with external coaptation, 2 with amputation, and 4 were euthanized. Additional, larger, multi-institutional retrospective analyses need to be performed with particular attention paid to outcomes of various treatment for orthopedic injuries to obtain a better understanding of what therapies are most appropriate for small mammals.
References 1. Anguita-Alonso P, Rouse MS, Piper KE, et al. Comparative study of antimicrobial release kinetics from polymethylmethacrylate. Clin Orthop Relat Res. 2006;445:239-244. 2. Aragon CL, Budsberg SC. Applications of evidence-based medicine: cranial cruciate ligament injury repair in the dog. Vet Surg. 2005;34(2):93-98. 2a. Barron HW, McBride M, Martinez-Jimenez D, et al. Comparison of two methods of long bone fracture repair in rabbits. J Exot Pet Med. 2010;19(2):183-188. 3. Böstman O, Pihlajamäki H. Clinical biocompatibility of biodegradable orthopaedic implants for internal fixation: a review. Biomaterials. 2000;21(24):2615-2621. 4. Bouvy BM, Markel MD, Chelikani S, et al. Ex vivo biomechanics of Kirschner-Ehmer external skeletal fixation applied to canine tibiae. Vet Surg. 1993;22(3):194-207. 5. Braden T. Posttraumatic osteomyelitis. Vet Clin North Am Small Anim Pract. 1991;21(4):781-811. 6. Bubenik L, Smith MM. Orthopedic infections. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:1862-1875.
CHAPTER 33 Orthopedics in Small Mammals 7. Calhoun JH, Mader JT. Antibiotic beads in the management of surgical infections. Am J Surg. 1989;157(4):443-449. 8. Capello V, Gracis M. Surgical treatment of periapical abscesses. In: Capello V, ed. Rabbit and rodent dentistry handbook. Lake Worth: Zoological Education Network; 2005:249-273. 9. Cevik-Demirkan A, Ozdemir V, Turkmenoglu I, et al. Anatomy of the hind-limb skeleton of the chinchilla (Chinchilla lanigera). Acta Vet Brno. 2007;76(4):501-507. 10. Crowley DJ, Kanakaris NK, Giannoudis PV. Irrigation of the wounds in open fractures. J Bone Joint Surg Br. 2007;89B(5):580-585. 11. Cruise L, Brewer NR. Anatomy. In: Manning P, Ringler DH, Newcomer CE, (ed). The biology of the laboratory rabbit. San Diego: Academic Press, 199:47-62. 12. Cui QJ, Mihalko WM, Shields JS, et al. Antibiotic-impregnated cement spacers for the treatment of infection associated with total hip or knee arthroplasty. J Bone Joint Surg Am. 2007;89A(4):871-882. 13. DeCamp C. External coaptation. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:1835-1848. 14. Egger EL, Histand MB, Blass CE, et al. Effect of fixation pin insertion on the bone-pin interface. Vet Surg. 1986;15(3):246-252:1986. 15. Egger EL. Complications of external fixation. A problemoriented approach. Vet Clin North Am Small Anim Pract. 1991;21(4):705-733. 16. Ethell MT, Bennett RA, Brown MP, et al. In-vitro elution of gentamicin, amikacin, and ceftiofur from polymethyl methacrylate and hydroxyapatite cement. Vet Surg. 2000; 29(5):375-382. 17. Graham JE. Rabbit wound management. Vet Clin North Am Exot Anim Pract. 2004;7(1):37-55. 18. Heard GS, Oloff LM, Wolfe DA, et al. PMMA bead versus parenteral treatment of Staphylococcus aureus osteomyelitis. J Am Podiatr Med Assoc. 1997;87(4):153-164. 19. Holcombe SJ, Schneider RK, Bramlage LR, et al. Use of antibiotic-impregnated polymethyl methacrylate in horses with open or infected fractures or joints: 19 cases (1987-1995). J Am Vet Med Assoc. 1997;211(7):889-893. 20. Holsworth I, DeCamp CE. Coxofemoral luxation. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:2002-2008. 21. Howard E, Nguyen QA. Anatomy. In: Fox J, ed. Biology and diseases of the ferret. Baltimore: Williams & Wilkins; 1998:19-70. 22. Hulse D, Hyman B. Fracture biology and biomechanics. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:1785-1792. 23. Kapatkin AS. Outcome-based medicine and its application in clinical surgical practice. Vet Surg. 2007;36(6):515-518. 24. Keller MA, Voss K, Montavon PM. The ComPact UniLock 2.0/2.4 system and its clinical application in small animal orthopedics. Vet Comp Orthop Traumatol. 2005;18(2):83-93. 25. Koch D. Screws and plates. In: Johnson A, Houlton JE, Vannini R, eds. AO principles of fracture management in the dog and cat. Switzerland: AO Publishing, Davos; 2005:26. 26. Låftman P, Nilsson OS, Brosjö O, et al. Stress shielding by rigid fixation studied in osteotomized rabbit tibiae. Acta Orthop Scand. 1989;60(6):718-722. 27. Mader JT, Calhoun J, Cobos J. In-vitro evaluation of antibiotic diffusion from antibiotic-impregnated biodegradable beads and polymethylmethacrylate beads. Antimicrob Agents Chemother. 1997;41(2):415-418. 28. Marcellin-Little. External skeletal fixation. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:1818-1834.
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29. Miclau T, Dahners LE, Lindsey RW. In-vitro pharmacokinetics of antibiotic release from locally implantable materials. J Orthop Res. 1993;11(5):627-632. 30. Miller DL, Goswmi T. A review of locking compression plate biomechanics and their advantages as internal fixators in fracture healing. Clin Biomech. 2007;22(10):1049-1062. 31. Millis D, Jackson AM. Delayed unions, nonunions, and malunions. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:1849-1861. 32. Monassi CR, Hoffmann A, Menescal-de-Oliveira L. Involvement of the cholinergic system and periaqueductal gray matter in the modulation of tonic immobility in the guinea pig. Physiol Behav. 1997;62(1):53-59. 33. Nijhof M, Stallmann HP, Vogely C, et al. Prevention of infection with tobramycin-containing bone cement or systemic cefazolin in an animal model. J Biomed Mater Res. 2000;52(4):709-715. 34. Nunamaker DM. Experimental models of fracture repair. Clin Orthop Relat Res Suppl. 1998:S56-S65. 35. Oglesbee B. Vertebral fracture or luxation. The 5-minute veterinary consult: ferret and rabbit. Ames: Blackwell Publishing; 2006;381. 36. Percy D, Barthold SW. Rabbit: Bacterial infections. In: Pathology of laboratory rodents and rabbits. Ames: Blackwell Publishing; 2007:264-286. 37. Perren S, Mathys R. Pohler O. Appendix—Implants and materials in fracture fixation. In: Johnson A, Houlton JE, Vannini R, eds. AO principles of fracture management in the dog and cat. Switzerland: AO Publishing, Davos; 2005:476. 38. Phillips H, Boothe DM, Shofer F, et al. In-vitro elution studies of amikacin and cefazolin from polymethylmethacrylate. Vet Surg. 2007;36(3):272-278. 39. Popesko P, Rajtová V, Horák J. A colour atlas of the anatomy of small laboratory animals. Vols 1 & 2. Bratislava: Wolfe Publishing Ltd; 1990. 40. Ramos JR, Howard RD, Pleasant RS, et al. Elution of metronidazole and gentamicin from polymethylmethacrylate beads. Vet Surg. 2003;32(3):251-260. 41. Ritzman TK, Knapp, D. Ferret orthopedics. Vet Clin North Am Exot Anim Pract. 2002;5(1):129-155. 42. Roe S. Internal fracture fixation. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:1798-1818. 43. Roe S. External fixators, pins, nails, and wires. p. 52. In: Johnson A, Houlton JE, Vannini R, eds. AO Principles of fracture management in the dog and cat. Switzerland: AO Publishing, Davos; 2005:52-70. 44. Saikku-Bäckström A, Tulamo RM, Raiha JE, et al. Intramedullary fixation of cortical bone osteotomies with absorbable selfreinforced fibrillated Poly-96L/4D-lactide (SR-PLA96) rods in rabbits. Biomaterials. 2001;22(1):33-43. 45. Saikku-Bäckström A, Tulamo RM, Raiha JE, et al. Intramedullary fixation of femoral cortical osteotomies with interlocked biodegradable self-reinforced Poly-96L/4D-lactide (SR-PLA96) nails. Biomaterials. 2004;25(13):2669-2677. 46. Snow M, Thompson G, Turner PG. A mechanical comparison of the locking compression plate (LCP) and the low contactdynamic compression plate (DCP) in an osteoporotic bone model. J Orthop Trauma. 2008;22(2):121-125. 47. Terjesen T. Bone healing after metal plate fixation and external fixation of the osteotomized rabbit tibia. Acta Orthop Scand. 1984;55(1):69-77. 48. Terjesen T. Healing of rabbit tibial fractures using external fixation—effects of removal of the fixation device. Acta Orthop Scand. 1984;55(2):192-196. 49. Tillson M. Open fracture management. Vet Clin North Am Small Anim Pract. 1995;25(5):1093-1110. 50. Tomlinson JL. Complications of fractures repaired with casts and splints. Vet Clin North Am Small Anim Pract. 1991;21(4):735-744.
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51. Tomlinson JL. Fractures of the humerus. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:1905-1918. 52. Waldron D, Zimmerman-Pope N. Superficial skin wounds. In: Slatter D, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders; 2003:259-273. 53. Wang GH, Liu SJ, Ueng SW, et al. The release of cefazolin and gentamicin from biodegradable PLA/PGA Beads. Int J Pharm. 2004;273(1-2):203-212. 54. Weisman DL, Olmstead ML, Kowalski JJ. In-vitro evaluation of antibiotic elution from polymethylmethacrylate (PMMA) and mechanical assessment of antibiotic-PMMA composites. Vet Surg. 2000;29(3):245-251.
55. Wenke JC, Owens BD, Svoboda SJ, et al. Effectiveness of commercially-available antibiotic-impregnated implants. J Bone Joint Surg Br. 2006;88(8):1102-1104. 56. Worlock P, Slack R, Harvey L, et al. An experimental model of post-traumatic osteomyelitis in rabbits. Br J Exp Pathol. 1988;69(2):235-244.
CHAPTER
34
Exotic Mammal Diagnostic and Surgical Endoscopy
Stephen J. Divers, BSc (Hons), BVetMed, Diplomate ACZM, Diplomate ECZM (Herpetology), Diplomate ZooMed, FRCVS
Patient Selection Patient Evaluation Anesthesia Instrumentation Procedures Otoscopy Stomatoscopy Endotracheal Intubation Tracheobronchoscopy Rhinoscopy Vaginoscopy/Cystoscopy Gastroscopy/Colonoscopy Laparoscopy Thoracoscopy Complications Postoperative Care Outcome Acknowledgments
With over 10 million pet rabbits (Oryctolagus cuniculus), ferrets (Mustela putorius furo), and rodents (order Rodentia) in the United States, these exotic mammals represent the third largest group of companion mammals (behind dogs and cats).1 This group represents an expanding component of small-animal practice, with many clients expecting the same level of medicine for them as for our more traditional clientele. The advent of the certifying examination in zoological companion species (American College of Zoological Medicine) and exotic companion mammals (American Board of Veterinary Practitioners) has further galvanized the need for veterinarians to provide the expected high-quality care being demanded for these species. The majority of rabbits, ferrets, and rodents presented to practitioners weigh less than 2 kg, and, given their small size, are ideal candidates for minimally invasive diagnostic and surgical endoscopy. Indeed, in some situations, the development of Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
endoscopy has enabled many procedures to be performed for the first time or with significantly reduced morbidity and mortality compared with traditional surgery. Considerable advances in exotic animal endoscopy have been made over the past 5 years, and further development and refinement seems assured.8
PATIENT SELECTION Unlike ferrets, which are carnivores, rabbits and rodents are prey animals; as such, they will generally mask symptoms until disease is advanced. Consequently, many rabbits and rodents may require preanesthetic nursing support (particularly analgesia, fluid therapy, and nutrition) prior to anesthesia and endoscopy. The most common endoscopic procedures performed in rabbits and rodents are stomatoscopy (examination of the oral cavity), rhinoscopy, otoscopy, cystoscopy, laparoscopy, and endotracheal intubation. In ferrets, laparoscopy and gastroscopy are probably most common. In addition to anesthetic contraindications, obesity can hamper laparoscopy; in most cases, however, it is small patient size that presents the greatest challenge to the exotic mammal endoscopist.
PATIENT EVALUATION Detailed husbandry and medical anamneses are essential, as many ailments are associated with poor captive management. Ideally, hematology, plasma biochemistry, and urinalysis should precede anesthesia; in many cases, however, short-term anesthesia may be required for the collection of such samples, while in small rodents the required volume may prove difficult to obtain. In addition, regional survey radiographs are advised prior to endoscopy.
ANESTHESIA The diversity within the exotic mammal group necessitates generalities rather than specifics, and reference should be made to the extensive literature on rabbit, ferret, and rodent anesthesia for precise drug dosages.3-5 Ferrets are typically fasted for 6 to 485
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8 hours. Rabbits and rodents seldom vomit or regurgitate and are fasted (which includes removal of food and all bedding materials) for only 1 to 2 hours to reduce material within the oral cavity. An opioid-benzodiazepine premedicant is generally effective and, following preoxygenation by mask, induction can be accomplished using intravenous or intramuscular ketamine alone or in combination with an additional benzodiazepine (rabbits and rodents), or intravenous propofol (ferrets). Unlike ferrets, in which it is relatively easy to place an endotracheal tube, rabbits and rodents are more challenging, and ketamine provides sufficient time for visualization, application of a local anesthetic, and careful visual intubation. Small rodents that cannot be intubated are best induced and maintained using isoflurane or sevoflurane with oxygen via a close-fitting rodent mask and dedicated nonrebreathing rodent circuit. Maintain animals on isoflurane or sevoflurane in oxygen adjusted to individual patient requirements. Ventilation is often required, especially in rabbits and rodents in dorsal recumbency, because of gastrointestinal pressure on their diaphragms, reducing ventilation. However, ventilation is essential for all laparoscopy, thoracoscopy, and procedures requiring neuromuscular blockade (e.g., rhinoscopy). Hypothermia is difficult to prevent unless warm-water- and air-circulating blankets and drapes or covers are used. Warming air filters attached to the endotracheal tube can also be useful for maintaining temperature, but these often require ventilator support. In addition to reflexes and muscle tone, anesthesia monitoring should include temperature, pulse, respiratory rate, end-tidal capnography, pulse oximetry, and direct or indirect blood pressure. Many of these aids become increasingly problematic as animal size decreases. During rhinoscopy of rabbits and larger rodents, intranasal lidocaine and short-term neuromuscular blockade using atracuronium further enhance immobilization; otherwise a much deeper plane of general anesthesia would be required. Intraoperative fluid support is important and can be delivered through intravenous (aural, cephalic, saphenous, or jugular) or intraosseous (femoral, humeral) catheters using syringe pumps. In cases where catheterization is not possible, administer fluids subcutaneously in the dorsal region immediately following induction and again on recovery. After extubation, provide small mammals with oxygen via a loose mask until they are fully conscious; then return them to a warm incubator until they are normothermic. Postoperative fluids, nutrition, and analgesia are essential for rapid recovery and return to normal function.
INSTRUMENTATION Given the variation in size and the nature of the procedures that may be performed, a variety of different scopes and instruments may be used (Table 34-1). For most practices the 2.7-mm system offers the greatest versatility, which can be built upon as individual practice caseload dictates. This system offers several advantages, including single-entry procedures, ports for air or saline infusion, and an operating channel for the introduction of 5-Fr instruments (Fig. 34-1). In addition, the 1.9-mm telescope with integrated sheath and the 1-mm semirigid mini scope are extremely useful for smaller mammals (Fig. 34-2). For multiple-entry endoscopy, the recent application of human pediatric 2- and 3-mm instruments to exotic animal endoscopy has enabled laparoscopy and thoracoscopy to become a reality (Figs. 34-3 and 34-4).6
PROCEDURES In general, the approach to exotic mammal endoscopy is similar to that in domesticated dogs and cats, and much can be learned and applied from the domestic animal and human literature.10,12 However, in addition to the anatomic peculiarities of these creatures, the exotic mammal endoscopist must be more precise, given the confines within which it is often necessary to work. It is therefore particularly important to use the fingers and thumb of the inferior hand to support the tip of the telescope in order to ensure accurate control at all times.
OTOSCOPY With the anesthetized animal in sternal or lateral recumbency, a detailed evaluation of the ears can be undertaken. In cases of severe disease, gently remove superficial exudates and debris before employing sterile saline infusion and working within a fluid environment. Abnormalities can be sampled for histopathology and microbiology using the biopsy forceps, which tends to provide more precise results than introducing a culturette down the vertical canal. Depending upon the size of the animal, diameter of the scope, and nature of the aural disease, it is often possible to examine the vertical and horizontal canals down to the tympanum (Fig. 34-5). It is especially important to examine the tympanum in rabbits and rodents as head tilt due to otitis media or interna is common. In ferrets, ear mites and aural neoplasia appear to be more common than bacterial infection.
STOMATOSCOPY Dental disease is undoubtedly one of the most common presentations for rabbits and rodents, and stomatoscopy under general anesthesia ensures a far more detailed examination than can be achieved in the conscious animal in the examination room.7,8,13 The limited access to the oral cavity may preclude the use of endotracheal tubes; however, anesthetic gas and oxygen can be supplied via nasal intubation or by placing a small face mask over the nostrils. It is important to consider using an active scavenging device in the area to avoid exposing the staff to anesthetic gas. Alternatively, injectable anesthetic agents may be used, and although this does not negate the need for supplying oxygen via nasal line or mask, it does reduce staff exposure to inhalant agents. Position the animal in sternal recumbency with the head supported and mouth held open using a rodent/rabbit table retractor restrainer (Sontec Instruments Inc., 7248 South Tuscon Way, Centennial, CO) and cheek spreaders (Sontec Instruments Inc.). The 1.9- or 2.7-mm telescopes are preferred because the 30-degree angle provides a better view of the occlusal surfaces; with the light-guide cable down, the view is angled toward the maxillary arcades. With the light-guide cable up, the mandibular teeth are preferentially seen (Fig. 34-6). Evaluate the lingual, buccal, and occlusal aspects of every tooth using appropriately sized curved dental probes. Note any tooth laxity, exudates, and gingival changes. In the vast majority of the small herbivores, the most commonly encountered malocclusions involve overgrowth to the lingual aspect of the lower arcades and buccal aspect of the upper arcades (Figs. 34-7 and 34-8). Once identified, trim the malocclusion with ronguers, a dental rasp, or preferably a motorized dental handpiece, which is less likely to result in dental fracture. During dental trimming, it is vital that the telescope be protected by using a guard or that it be temporarily removed
CHAPTER 34 Exotic Mammal Diagnostic and Surgical Endoscopy
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Table 34-1 Endoscopic Instrumentation for Ferrets, Rabbits, and Rodents Equipment Description
Primary Indications
Visualization & Documentation Endovideo camera and monitor Xenon light source and light guide cable Digital capture device (e.g., AIDA-Vet)
Required for all endoscopy procedures
Endoscopes 1-mm x 20-cm semirigid miniscope, 0° 1.9-mm x 18.5-cm telescope, 30° oblique, with integrated 3.3-mm operating sheath 2.7-mm x 18-cm telescope, 30° oblique 4.8-mm operating sheath 5-mm x 8.5-cm otoendoscope, 0°, with integrated operating sheath
Stomatoscopy, otoscopy, rhinoscopy in animals <500 g Endotracheal intubation in animals <1 kg Stomatoscopy, otoscopy, rhinoscopy in animals 500 g-10 kg Endotracheal intubation in animals 1-8 kg Laparoscopy in animals <1 kg Stomatoscopy, otoscopy, rhinoscopy in animals 500 g-10 kg Endotracheal intubation in animals 1-8 kg Laparoscopy and thoracoscopy in animals 1-8 kg Stomatoscopy and otoscopy in animals 3-40 kg
Flexible Instruments for Use with Operating Sheaths 1-mm biopsy forceps 1-mm grasping forceps 1.7-mm x 34-cm biopsy forceps 1.7-mm x 34-cm single-action scissors 1.7-mm remote injection needle 1.7-mm x 34-cm grasping/retrieval forceps 1.7-mm x 60-cm wire basket retrieval 1.7-mm needle end radiosurgery electrode
For use with 1.9-mm telescope and integrated sheath For use with 2.7-telescope and 4.8-mm operating sheath, and 5-mm otoendoscope
Insufflation CO2 insufflator with silicone tubing Sterile saline suspended above endoscopy table with intravenous drip line to a port on the operating sheath
Used for insufflation during laparoscopy Used for sterile saline infusion for otoscopy, rhinoscopy, cystoscopy, vaginoscopy, and colonoscopy
Rigid Instruments and Cannulas for Multiple-entry Laparoscopy and Thoracoscopy 2.5-mm graphite and plastic cannula Used with the 1.9-mm telescope for laparoscopy and 2-mm Reddick-Olsen dissecting forceps, plastic handle without racket thoracoscopy in animals <1 kg 2-mm Metzenbaum scissors, plastic handle without racket 2-mm Babcock forceps, plastic handle with racket 3.9-mm graphite and plastic cannula (accommodates 2.7 mm Used with the 2.7-mm telescope for laparoscopy and telescope and 3.5-mm protection sheath) thoracoscopy in animals 1-8 kg 3.5-mm graphite and plastic cannula (accommodates 3-mm instruments) 3-mm fenestrated grasping forceps 3-mm Reddick-Olsen dissecting and grasping forceps 3-mm short curved Kelly dissecting and grasping forceps 3-mm atraumatic dissecting and grasping forceps 3-mm Babcock forceps 3-mm Blakesley dissecting and biopsy forceps 3-mm scissors with serrated curved double action jaws 3-mm micro-hook scissors, single action jaws 3-mm Mahnes bipolar coagulation forceps 3-mm irrigation and suction cannula 3-mm palpation probe with centimeter markings 3-mm distendable palpation probe 3-mm ultramicro needle holder 3-mm knot-tier for extracorporeal suturing 2 plastic handles without rackets 1 plastic handle with Mahnes-style racket 1 plastic handle with hemostat-style racket
Radiosurgery Equipment 3.8- or 4.0-MHz dual radiofrequency unit with foot pedal Monopolar lead to connect to plastic instrument handles Bipolar lead to connect to 3-mm Mahnes bipolar coagulation forceps
Enables endoscopic instruments to be used as monopolar devices and facilitates bipolar coagulation
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Fig. 34-1 2.7-mm telescope system. A, 2.7-mm telescope housed within a 4.8 operating sheath, connected to a light cable and an endovideo camera. B, 1.7-mm biopsy forceps inserted down the instrument channel and emerging directly in front of the telescope. C, A variety of 1.7-mm instruments can be used through the operating channel, including retrieval forceps (1), biopsy forceps (2), remote injection/aspiration needle (3), and single-action scissors (4).
A Fig. 34-3 Multiple-entry 3-mm endoscopy system. A, Plastic, nonracket handle (1) attached to radiosurgery (2), and a 3-mm instrument (3), which is inserted through a 3.5-mm cannula (4). B, Instruments (1) are interchangeable by pressing the button (arrow), and the radiosurgery connector (2) on the handle enables the instrument to be used as a monopolar device. glottis to aid direct intubation. The author favors inserting the endoscope into the endotracheal tube, passing the endoscope through the glottis and then advancing the tube off the endoscope and into the trachea (Fig. 34-10).
A
B Fig. 34-2 A, 1.9-mm telescope with integrated operating sheath. B, 1-mm semirigid miniscope with a 2-mm endotracheal tube placed over the scope in preparation for exotic mammal intubation. and then be reinserted to evaluate the teeth following reduction of the malocclusion (Fig. 34-7). The telescope can also be used periodically to evaluate the intraoperative progress of premolar or molar extractions or to examine the cavity that is left following extraction (Fig. 34-8). Indeed, the telescope has also been used to target flushing and antimicrobials into dental cavities via the oral cavity (Fig. 34-9). These techniques have been used to successfully treat retrobulbar abscesses in rabbits via the oral cavity, thereby avoiding enucleation.9 Although rare, soft tissue masses may be biopsied using the 5-Fr biopsy forceps, while foreign bodies may be removed using retrieval forceps.
ENDOTRACHEAL INTUBATION Intubation is more difficult in rodents and rabbits, and the endoscope can serve as a useful aid for intubation prior to any prolonged diagnostic or surgical procedure. The endoscope can be used as a laryngoscope to provide visualization of the
TRACHEOBRONCHOSCOPY Small flexible bronchoscopes can be used to examine the trachea and bronchi of larger rabbits and rodents; however, small telescopes can also be used to gain access to the level of the tracheal bifurcation and beyond (Fig. 34-11). It is vital that on entering the glottis, the head and neck be kept straight and extended to avoid mucosal damage as the telescope is advanced. Unless oxygenation can be maintained, tracheobronchoscopy evaluations must necessarily be brief.
RHINOSCOPY With the animal intubated and in sternal recumbency in a 10to 20-degree head-down position, pack the oropharynx with moistened gauze. Flush the nasal cavities using warm sterile saline to remove any debris and excess mucus (Fig. 34-12, A). Placing towels under the animal’s head helps prevent flooding of the table and floor. Use the 2.7-mm telescope for animals weighing more than 2 kg; for smaller animals, the 1.9-mm sheathed telescope is preferred (Fig. 34-12, B). The use of a sheath enables intraoperative flushing to maintain visualization; however, in small animals, the naked telescope can be used with care along with intermittent syringe flushing through the nostrils. The ventral and middle nasal meati can be exploited to examine the ventral and middle conchae. In larger animals, the endoturbinates and opening to the nasopharynx can also be seen. Care is required to avoid damaging the delicate nasal turbinates, which are prone to hemorrhage (Fig. 34-13, A). Keep the telescope as medial as possible and always keep it within the meati. Even so, hemorrhage can rarely be completely avoided. Exudates, abscesses, masses, and foreign bodies can be appreciated and biopsied or removed (Fig. 34-13, B-F).
CHAPTER 34 Exotic Mammal Diagnostic and Surgical Endoscopy
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Fig. 34-4 Multiple-entry 3-mm endoscopy system. A, Trocar (1) and 3.5-mm plastic/graphite cannula (2) with side insufflation port (3). B, Forceps: Babcock forceps (1), atraumatic dissecting and grasping forceps with single-action jaws (2), Reddick-Olsen dissecting forceps (3), fenestrated grasping forceps (4), long curved Kelly dissecting and grasping forceps (5), short curved Kelly dissecting and grasping forceps (6). C, Scissors and biopsy instruments: micro-hook scissors with single-action jaws (1), Blakesley dissecting and biopsy forceps (2), scissors with long, sharp, curved double-action jaws (3), scissors with serrated curved double-action jaws (4). D, Probes: distendable palpation probe (1), palpation probe with centimeter markings (2), irrigation and suction cannula (3).
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Fig. 34-5 Rabbit otoscopy. A, Normal horizontal canal and tympanum. B, Inflamed horizontal canal with caseous debris visible behind an intact tympanum. C, Ruptured tympanum without evidence of inflammation or infection. D, Otitis media with ruptured tympanum and caseous debris visible.
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Fig. 34-6 Rabbit stomatoscopy. A, Positioning of an anesthetized rabbit for intraoral endoscopy using a dental rack, cheek dilators, and nasal oxygen. This system allows positional versatility for a variety of small exotic mammals. B, Positioning of the telescope and light-guide cable to capitalize on the 30-degree viewing angle for examination of the mandibular teeth. Rotating the telescope 180 degrees around its longitudinal axis (such that the light-guide cable faces down) facilitates examination of the maxillary teeth. In this case the rabbit’s anesthesia is being maintained via a small nasal cone.
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Fig. 34-7 Intraoral endoscopy in a normal rabbit and guinea pig. A, General overview of the oral cavity of a normal rabbit. The hard palate (1) and tongue (2) are labeled for orientation; note that the maxillary crowns (black arrows) are naturally shorter than the mandibular crowns (white arrows). B, Normal left maxillary arcade in a rabbit demonstrating the third premolar (PM3) and the first, second, and third molar teeth (M1-3). C, General overview of the oral cavity of a normal guinea pig. The hard palate (1) and tongue (2) are labeled for orientation; note the angular slope of the dental arcades as compared with the rabbit. D, Close-up of a maxillary molar in a guinea pig demonstrating the enamel-dentine interface (arrows), which produces the cusps that aid forage mastication.
CHAPTER 34 Exotic Mammal Diagnostic and Surgical Endoscopy
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Fig. 34-8 Endoscopic dental pathology in rabbits. A, Abnormally long right maxillary premolars 1 and 2 due to inadequate dietary forage. B, Close-up of more severe elongation of the right maxillary premolars 1 and 2. In this case the teeth have caused severe ulceration in the buccal mucosa (arrow). C, Mild elongation of right maxillary premolar 3, molar 1, and molar 2; molar 3 appears normal. Note that molar 1 also has a small sharp spur associated with its buccal border (arrow). This spur would be buried in the buccal mucosa and would be impossible to visualize without endoscopy and lateral retraction of the mucosa. D, Gross elongation of left mandibular premolar 2 with associated lingual spur (arrow) and fracture or severe wear of premolar 1 to the level of the gingiva. E, Caseous exudate emanating from around the left maxillary molar 2 following the application of pressure with a dental probe. F, End-stage dental disease. In this view of the left maxillary arcade, only premolar 1 and molar 3 are clearly visible, premolar 2 and molar 2 are missing, while part of molar 1 (arrow) can be seen emerging from swollen gingival, which may well indicate an underlying dental abscess.
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Fig. 34-9 Endoscopic dental treatment in rabbits. A, Preparation for trimming elongated maxillary premolars and molars using a dental burr (1). Note the metal probe (2) being used to retract and protect the buccal mucosa from the burr. B, High-speed dental burring requires great care to protect the soft tissues and the telescope. Metal probes and guards should always be used. C, Immediate postoperative view of the maxillary arcade following burring down to the level of the gingiva. D, Immediate postoperative view following intraoral extraction of molars 2 and 3, causing a retrobulbar abscess. E, Close-up of the extraction site demonstrating the ability of the 30-degree telescope to look into the abscess cavity and visualize the maxillary bone (arrow). F, Placement of an intravenous catheter (arrow) into the abscess cavity to instill antibiotic-impregnated synthetic bioactive ceramic material.
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Fig. 34-10 Rabbit intubation. A, Normal resting position of the epiglottis engaged over the caudal edge of the soft palate in an obligate nasal-breathing rabbit. B, Following induction and with the head and neck extended, the endoscope is used to displace the soft palate dorsad to free the epiglottis. C, With the epiglottis now lying ventrad in the oropharynx, the glottis can be clearly seen. D, Following the application of local anesthetic, the endotracheal tube and stylet are inserted into the trachea. The recent advent of 2- and 3-mm rigid instruments also permits biopsy and debridement within the nasal or paranasal sinuses via limited surgical access.6 There are occasions when dental disease requires an extraoral approach, either alone or in conjunction with intraoral surgery, and the telescope can serve as a useful surgical aid. The extension of hypsodont roots into the nasal cavity may warrant rhinoscopy via the nostrils, as detailed above, or surgical rhinotomy. Dental abscesses affecting the maxilla may enter the paranasal sinuses; in such a case, the telescope can provide evaluation via a small (4-5 mm) osteotomy. Even when extensive surgical osteotomy or rhinotomy is performed, surgical access is often very limited in small herbivores. However, the telescope enables detailed evaluation, including areas cranial and caudal to the surgical site (Fig. 34-14).
VAGINOSCOPY/CYSTOSCOPY Hematuria is not uncommon in rabbits and rodents; it can be related to diseases affecting the urinary or reproductive systems. With the animal in dorsal recumbency and the perineum close to the table edge, insert a small 30-degree sheathed telescope
through the vulva and into the vagina. Using a sterile saline infusion, it is possible to evaluate the vagina, urethra, bladder, and surface of the cervices of animals as small as 500 g (Fig. 34-15).
GASTROSCOPY/COLONOSCOPY Gastrointestinal disease is especially common in ferrets; however, there has been a noticeable absence of endoscopy in the pursuit of definitive diagnoses. This is unfortunate because ferrets and other small mammals weighing more than 1 kg can often accommodate the smaller flexible gastroscopes, while the stomach can often be reached using telescopes in animals weighing less than 1 kg (Fig. 34-16). The ability to confirm the presence of gastric ulceration and collect mucosal biopsies for cultures and histology should be considered routine in the investigation of gastric diseases in ferrets (Fig. 34-17, A,B). Unfortunately the stomachs of small herbivores are never empty, which can make endoscopic evaluation nearly impossible. Rodent colonoscopy recently became important because researchers were looking for a means of following the
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Fig. 34-11 Rabbit tracheobronchoscopy. Telescopic views of the normal anterior trachea (A), bifurcation (B), bronchus and secondary bronchi (C). D, Flexible endoscopic view of the bifurcation with a foreign body lodged in a primary bronchus (arrow).
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Fig. 34-12 Rabbit rhinoscopy. A, Flushing the nasal cavities using warm sterile saline delivered via a 60-mL syringe and cut-down 8-Fr red rubber catheter. Packing the oral cavity with gauze and use of a cuffed endotracheal tube are essential. B, Performing rhinoscopy using a 2.7-mm telescope and 4.8-mm operating sheath. A bag of sterile saline suspended above the table is connected to one of the sheath ports by an intravenous administration set, facilitating intraoperative flushing. A second administration set is connected to the second sheath port, providing egress to a collection bucket under the table.
CHAPTER 34 Exotic Mammal Diagnostic and Surgical Endoscopy
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Fig. 34-13 Rabbit rhinoscopy. A, Normal nasal turbinates viewed from within the ventral nasal meatus. B, Abscess in the caudal aspect of the ventral nasal meatus in a rabbit unsuccessfully treated for chronic rhinitis. C, Same abscess following endoscopic debridement. It was resolved with the help of postoperative antimicrobials based on biopsy culture and sensitivity. D, Granulomatous rhinitis with destruction of the nasal septum due to Mycobacterium species. E, Hay foreign body (arrow) within the cranial aspect of the ventral nasal meatus, just caudal to the alar fold. F, Ventral conchal atrophy and chronic rhinitis.
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Fig. 34-14 Surgical rhinotomy and rhinoscopy in a rabbit. A, Intraoperative view illustrating the use of the telescope to examine the nasal cavity following rhinotomy. Insert, closeup of the introduction of the telescope through the rhinotomy. B, Endoscopic view of caseous exudate (arrow) within the ventral nasal meatus following the removal of chronically infected turbinates. The nasal septum (1) is labeled for orientation. C, Exposure of an overgrown maxillary tooth (arrow) that has extended into the nasal cavity and caused the infection. The nasal septum (1) is labeled for orientation; the suction probe (2) is also visible. D, Intraoperative view following tooth extraction and flushing of the site, revealing a now clear and unhindered ventral nasal meatus leading to the nasopharynx (arrow).
progression of human colon cancer in a rodent model. The 1.9-mm sheathed telescope has been used to examine the rectum and descending colon of mice weighing as little as 20 g. The ability to infuse methylene blue and other chemical markers via the operating sheath has further improved the ability to identify colonic cells undergoing early neoplastic transformation through the use of so-called chouromoendoscopy (Fig. 34-17 C,D).2 The same system and technique can be employed in companion rodents, with the animal in sternal or lateral recumbency and the perineum close to the table edge.
LAPAROSCOPY For a detailed discussion of methodology, the reader is referred to the dedicated laparoscopy literature, as only small-mammal specifics are highlighted here.10-12 Laparoscopy has been shown to offer significant advantages over traditional surgical options,
both in human and in veterinary medicine. In particular, laparoscopy is, with practice, faster and less traumatic, resulting in less postoperative pain and a faster return to normal function. Until the advent of 2- and 3-mm human pediatric equipment, laparoscopy was limited to a single-entry system using the sheathed telescope. However, multiple-entry techniques are now possible and practical for animals weighing more than 500 to 1,000 g. Indeed, laparoscopic ovariectomy is now the author’s sterilization method of choice for female rodents and rabbits because it involves less tissue manipulation and results in less postoperative discomfort and a faster return to normal feeding and behavior. Single-entry laparoscopy has been used most extensively for the collection of visceral biopsies from rodents and rabbits. In general, make a 3- to 4-mm surgical approach through the umbilicus or at some other convenient point along the linea alba (Fig. 34-18). Following insertion of the sheathed
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Fig. 34-15 Rabbit urogenital endoscopy. A, View within the anterior vagina demonstrating the urethra (1) and caudal vaginal vault (2). B, Urethral carcinoma (arrow) seen within the anterior vagina. C, Cervices (arrows) seen from within the caudal vagina. D, Urinary sand seen within a saline-infused bladder.
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Fig. 34-16 Ferret gastrointestinal endoscopy. A, Flexible gastroscopy using a 8.6-mm gastroscope. Such endoscopes provide greater flexibility but are limited to larger mammals weighing more than 1 to 2 kg. B, Esophagostomy and gastroscopy being performed with a 2.7-mm telescope and sheath with sterile infusion provided by the attached syringe. Such techniques can be used in mammals weighing less than 1 kg.
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Fig. 34-17 Gastrointestinal endoscopy in ferrets and rodents. Flexible endoscopic views of the normal gastric mucosa (A) and gastric ulceration due to Helicobacter mustelae (B) in ferrets. C, Telescopic view of a carcinoma protruding into the lumen of the descending colon in a mouse. D, Early neoplastic transformation in the colon (arrow) can be detected more easily using methylene blue dye.
telescope, tie a mattress suture to create an airtight seal; however, this is seldom necessary if the incision through the linea alba is small. For larger rabbits and rodents, CO2 insufflation is required; for small rodents, however, it is often possible to simply attach a syringe containing air to one of the sheath ports and manually inject air into the abdomen. Risks associated with air embolism have not been observed in rodents but should be considered. Single-entry techniques are simple and easy to perform, but the variety of instruments is limited; therefore tissue manipulation and endosurgery are rudimentary. Nevertheless, visceral evaluation and biopsy are practical even in small rodents (Fig. 34-18, B-D). For larger animals, multiple-entry techniques using 2- and 3-mm human pediatric instruments are preferred and provide greater opportunities for endosurgery.6 There is a wider selection of equipment for 3-mm instruments; these are used with click-line interchangeable handles and connected to a radiosurgery unit for hemostasis. Access to the abdomen is achieved using lightweight 3.5-mm (for instruments) and 3.9-mm (sheathed telescope) threaded graphite cannulas. Cannula placement is determined by the organ of interest, the preference of the surgeon, and the anatomy of the animal in question (Fig. 34-19). CO2
insufflation using a dedicated endoflator is essential for multiple-entry techniques. Some prefer the use of a Veress needle, while others, concerned over the risk of damage to internal viscera (especially the voluminous gastrointestinal tract of small herbivores), prefer to place the initial cannula or telescope surgically. The ability to rotate the animal from dorsal into lateral can greatly assist with the location of ovaries, kidneys, and other dorsolateral structures. While tilting endoscopy tables are commercially available for domesticated animals, a specifically designed and constructed a small-mammal tilting table has undergone trials at the University of Georgia. For procedures involving dorsolateral structures (e.g., ovariectomy), insert the telescope through the umbilicus with two additional cannulas placed 2- to 3-cm cranial and caudal to the telescope along the linea alba. In this way the instruments can be directed to one side. When required to examine the other side, the surgeon simply moves around the table. It is often helpful to have a second slave monitor located on the other side of the operating table rather than moving the entire endoscopy tower. For access to the liver, gastrointestinal tract, spleen, pancreas, and bladder, again pass the telescope through the linea alba or
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Fig. 34-18 Single-entry rodent laparoscopy. A, Rat intubated, positioned in dorsal recumbency, and prepared for laparoscopy. The linea alba is indicated and the preferred insertion point is the umbilicus (arrow). B, View of the right cranial quadrant illustrating the pancreas (1) and duodenum (2) of a guinea pig. C, Normal rabbit kidney (1) partially obscured by retroperitoneal fat (2). Obesity is commonly encountered, complicating laparoscopy. D, Biopsy forceps (1) harvesting a liver sample (2) from a rat.
umbilicus; but the instruments must be inserted transversely across the abdomen rather than longitudinally. In this manner, the telescope and instruments can be advanced into the cranial abdomen for access to the liver, stomach and intestinal tract, liver, spleen, and pancreas or advanced caudally toward the large intestine and bladder (Fig. 34-20). It is important to aspirate all abdominal gas following surgery, as the presence of residual gas is a source of postoperative discomfort. Close cannula holes using a single suture.
THORACOSCOPY Thoracoscopy remains within its infancy and few clinical applications have been realized in rabbits, rodents, and ferrets. Given the recent advances in human pediatric equipment, the surgeon’s ability is probably the major limiting factor, although the relatively small thoraces of rodents and rabbits will always present greater challenges. To date, a handful of thoracoscopy procedures have been performed in small mammals. All cases involved evaluation and biopsy of an intrathoracic mass where less invasive ultrasound-guided fine-needle aspirate cytology had proved inconclusive. In all cases, a single-entry system was used, with targeted telescope entry based upon preoperative imaging. In general, the paraxiphoid approach is easier in most rabbits and rodents weighing less than 2 kg, while an intercostal approach is preferred for ferrets and larger rabbits/rodents.
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Fig. 34-19 Multiple-entry laparoscopy in a ferret. The telescope (1) has been inserted through the umbilicus, cranial to the Veress needle (2) and CO2 insufflation line. A 3-mm instrument (3) has been inserted through one of two 3.5-mm cannulas (4).
Again, close the ports using single sutures. While the placement of a chest drain is not always necessary if all air has been aspirated from the pleural cavity, it should certainly be considered in cases of poor oxygen status and abnormal blood gas values. When such tubes are used, intermittent negative pressure is
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Fig. 34-20 Multiple-entry ferret laparoscopy. A, Using a palpation probe to examine between the left lateral (1) and left medial (2) lobes of the liver. B, Identification of an abnormally thickened gall bladder (1) between the quadrate and right medial liver lobes (2). C, Endoscopic biopsy of the pancreas (1) using 3-mm biopsy forceps that confirmed the presence of an insulinoma. Note the use of 3-mm atraumatic tissue forceps (arrow) to retract the mesentery. D, View of the spleen (1) demonstrating a nodular abnormality (black arrow) and the previous site of attempted ultrasoundguided fine needle aspiration (white arrow). E, Biopsy of the spleen (1) using 3-mm biopsy forceps (arrow). F, Postbiopsy view of the spleen (1); note the absence of severe hemorrhage.
CHAPTER 34 Exotic Mammal Diagnostic and Surgical Endoscopy applied and the tubes are removed once they are nonproductive for more than 30 minutes.
COMPLICATIONS The major complications encountered are typically associated with anesthesia and related to issues of debilitation, poor ventilation, lack of vascular access, and hypothermia. The importance of a thorough preoperative evaluation, endotracheal intubation, intravenous or intraosseous catheterization, and warm air/water blankets cannot be overemphasized. Hemorrhage following rhinoscopy or tissue biopsy is common but seldom severe, although it is wise to have hemostatic agents available. Most endoscopy issues are related to operator error, which may sometimes occur until experience and ability have been gained. To facilitate the endoscopy caseload without compromising clients or patients, it is recommended that the surgeon retains the option to convert to a traditional surgical approach if required.
POSTOPERATIVE CARE The immediate postoperative period following extubation is usually the most critical, and oxygen and thermal support should be continued until the animal is fully conscious and ambulatory. Opioid analgesics may be continued following major procedures; however, nonsteroidal anti-inflammatory drugs (e.g., meloxicam) are used routinely. Rabbits and rodents are expected to resume feeding within 2 hours; otherwise assisted feeding is initiated and fluid therapy continued. In addition, auscultation of the gastrointestinal tract is routine, with ileus promptly treated using cisapride.
OUTCOME Return to normal behaviors and weight gain are often the most useful indicators of improvement. Clinicopathology can also be useful if abnormalities were detected preoperatively. Use serial endoscopic evaluations of the ears, nose, and mouth to monitor patients with otitis, rhinitis, and dental disease.
ACKNOWLEDGMENTS I am indebted to Drs. Clarence Rawlings, Mary-Ann Radlinsky, Michael Taylor, Michael Murray, Scott Stahl, and Christopher Chamness for their mentorship, support, and encouragement, and to Betsy Kurimo, Ellen Gladden, Carol McElhannon, and
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Rob Miller, whose technical skills have ensured that equipment is always available and well maintained. Thanks also to Mr. Doug Merker of Karl Storz Veterinary Endoscopy for supporting zoological endoscopy research and development at the University of Georgia.
References 1. AVMA. U.S. pet ownership & demographics sourcebook. American Veterinary Medical Association. 2007. 2. Becker C, Fantini MC, Wirtz S, et al. In vivo imaging of colitis and colon cancer development in mice using high resolution chromoendoscopy. Gut. 2005;54:950-954. 3. Carpenter JW. Exotic animal formulary. 3rd ed. St. Louis, MO: WB Saunders; 2005. 4. Heard DJ. Anesthesia, analgesia, and sedation of small mammals. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. Philadelphia, PA: WB Saunders; 2004:356-369. 5. Heard DJ. Lagomorphs (rabbits, hares and pikas). In: West G, Heard D, Caulkett N, eds. Zoo animal & wildlife immobilization and anesthesia. Ames, Iowa: Blackwell Publishing; 2007:647-653. 6. Hernandez-Divers SJ. Minimally-invasive endoscopic surgery of birds. J Avian Med Surg. 2005;19:107-120. 7. Hernandez-Divers SJ. Clinical techniques: dental endoscopy of rabbits and rodents. J Exot Pet Med. 2008;17:87-92. 8. Hernandez-Divers SJ, Murray M. Small mammal endoscopy. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. Philadelphia, PA: WB Saunders; 2004:392-394. 9. Martinez-Jimenez D, Hernandez-Divers SJ, Dietrich U, et al. Endosurgical treatment of a retrobulbar abscess in a rabbit. J Am Vet Med Assoc. 2007;230:869-872. 10. McCarthy TC. Veterinary endoscopy for the small animal practitioner. St. Louis, 2005. 11. Monnet E, Twedt DC. Laparoscopy. Vet Clin North Am Small Anim Pract. 2003;33:1147-1163. 12. Tams TR. Small animal endoscopy. 2nd ed. St. Louis: Mosby; 1999. 13. Taylor M. Endoscopy as an aid to the examination and treatment of the oropharyngeal disease of small herbivorous mammals. Sem Avian Exot Pet Med. 1999;8:139-141.
CHAPTER
35
Diagnostic Imaging
Anthony J. Fischetti, DVM, MS, Diplomate ACVR
General Considerations Which Imaging Examination to Perform Image Resolution The Modalities Advanced Treatments Radiation Therapy Interventional Radiology
Veterinary radiology has undergone exciting changes in the past few years. The incorporation of digital radiography into veterinary practices has improved image dissemination, archiving, and overall quality. Additionally, the greater availability of ultrasound, computed tomography (CT), and magnetic resonance imaging (MRI) gives practitioners an enormous armamentarium of diagnostic options. This chapter touches on advantages and limitations of the different imaging modalities available in small-mammal practice. The chapter ends with a few comments on advanced treatment modalities related to radiation therapy and interventional radiology.
GENERAL CONSIDERATIONS Some radiographic technical principles are ubiquitous regardless of the species involved. These include proper restraint, maintenance of radiation safety, practical considerations for unstable patients, and proper positioning. Chemical restraint is recommended for most small mammal radiographic examinations. Isoflurane or sevoflurane delivered by face mask is convenient and safe. In excitable or fractious animals, induce anesthesia in an induction chamber and use a face mask for maintenance. In other instances, injectable agents can be used for short examinations. Sedative protocols are detailed in Chapter 31. For lengthy procedures, intubating the animal is preferable, but this can be difficult or impossible in small species.26 Depending on local and state laws, manual patient restraining techniques can sometimes be implemented for making 502
radiographs. Whether the patient is manually or chemically restrained, strict adherence to radiation safety principles should always be maintained. The basics of radiation safety can be boiled down to time, distance, and shielding.27 Efforts should be made to reduce the patient and restrainer’s time exposed to ionizing radiation. Lower exposure settings and reduced number of retakes are effective ways to reduce time of exposure (see “Digital Radiography,” below). Increasing the distance from the primary source of radiation is the single most effective method to reduce exposure to the restrainer. If a patient is properly positioned without manual restraint, there is no reason for anyone to be close to the radiology table during exposure, even if a lead apron is worn. One step back from the table exponentially reduces one’s exposure to the radiation.6 Barriers (shielding)—such as lead gloves, aprons, and walls—are effective means of radiation protection. However, it must be stressed that lead gloves will not protect the restrainer from the primary beam radiation (the radiation outlined by the collimator light). Leaded garments will protect only from the scatter radiation emanating from the tube, patient, and table (Fig. 35-1). Proper restraint allows for proper positioning. Improper positioning is a common source of radiographic misinterpretation. In radiography, orthogonal (90-degree) projections allow us to pinpoint the exact location of an abnormality (is it on the patient’s skin or in the body?). The more views we obtain, the more confident we can be of the presence of a radiographic abnormality (Fig. 35-2). Obliquity should be minimized, especially for ventrodorsal (VD) or dorsoventral (DV) projections. Properly positioned VD and DV radiographs are ideal for assessing symmetry from left to right.11,26 Symmetry is also crucial for interpreting cross-sectional imaging modalities like CT and MRI.
WHICH IMAGING EXAMINATION TO PERFORM Uncommon species presented to our facility’s exotic animal department can pose diagnostic challenges. Facing this challenge, the radiologist and primary care veterinarian work as a team to determine which imaging exam will provide the most useful information. A review of the advantages and drawbacks Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
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THE MODALITIES Radiography
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B Fig. 35-1 Lateral projection (A) of a rabbit. A gloved hand is within the primary beam, in violation with radiation safety. Leaded shielding does not protect the restrainer from primary beam radiation within the confines of collimation. B, A lateral projection of a ferret with a field of view that is beyond the confines of collimation (removed border mask). The gloved hands are visible because of off-focus and scatter radiation, not from the primary beam. This demonstrates proper use of a leaded glove to protect from lowerenergy scatter radiation and not the high-energy primary beam.
of various imaging modalities is presented below to illustrate this decision-making process. Practical considerations like anesthetic requirements, cost, and availability are crucial to the decision-making process. Other considerations that may not be readily obvious are the objective criteria used to assess image resolution (Table 35-1).
IMAGE RESOLUTION
Contrast Resolution. Contrast resolution refers to the ability to distinguish between shades of gray in an image. For example, MRI, which has exquisite contrast resolution for soft tissues, is the modality of choice for identifying subtle changes in gray and white matter of the brain (Fig. 35-3). Spatial Resolution. Spatial resolution refers the ability to distinguish two small, closely spaced objects as separate. Small animals can have small changes in anatomy when they are diseased. If the modality can resolve only 5 mm of tissue, disease processes smaller than 5 mm will be averaged as part of the surrounding tissues and appear indistinct or invisible. Spatial resolution is perhaps the greatest challenge posed to radiologists seeking to improve lesion detection and image quality in small patients. Temporal Resolution. An imaging modality’s ability to record anatomy in real time is termed temporal resolution. This is best depicted with ultrasound—for example, in assessing cardiac contraction.
Radiography is the most common imaging modality utilized by veterinarians because of its high availability, low cost, and greatest familiarity regarding technique and interpretation. Additionally, whole-body assessments are easy to obtain in small mammals, allowing for complete imaging of the thorax, abdomen, and musculoskeletal system without excessive geometric distortion. A few points should be made in regard to the radiographic imaging of very small patients. In all veterinary patients but particularly in small mammals with increased respiratory rates, motion during exposure is a problem. Therefore x-ray equipment capable of producing 300 milliamperes (mA) in 0.008 second is a minimum requirement.25,26 Most radiographs can be exposed for 0.008 to 0.016 second with excellent results. Incremental adjustments of kilovoltage are necessary through a range of 45 to 70 kilovoltage peak (kVp). A unit’s focal spot can be thought of as an adjustable narrowing of the beam of radiation emanating from the tube. A small focal spot allows for better spatial resolution.6,25 However, small focal spots limit the total milliamperes and are not recommended in larger patients. A telescoping x-ray stand that can be moved along the x-ray table and adjusted in height is helpful. The ability to change the focal-film distance (distance from the x-ray tube to the film) is useful to produce magnified views of anatomic areas of interest. Effective magnification is possible only with small focal spots. Horizontally directed x-ray beams are helpful in evaluating gravity-dependent (fluid and sediment) and non-gravity-dependent (gas) structures (Fig. 35-4). A horizontally directed beam can also provide safe, effective restraint for fractious patients or patients that can remain standing.12,29 Strategies for optimizing spatial resolution in film-based radiography focus on appropriate film-screen combinations. Highspeed film or cassettes used in most veterinary practices allow for less exposure but do not provide the resolution desired for small patients. Slower-speed film and cassettes provide better detail but require greater exposure. Fine or detail-intensifying screens can be used for radiographs of smaller patients (e.g., Curix Fine, Agfa, Orangeburg, NY; Quanta Detail, E.I. DuPont, Wilmington, DE; Lanex Fine, Eastman Kodak, Rochester, NY). These screens, combined with the proper x-ray film (Curix Detail RPIL, Agfa; Cronex 10, E.I. DuPont; TMG film, Eastman Kodak) maximize anatomic detail. For ultrafine detail in exotic pet radiography, the Min R mammography system (Eastman Kodak) can be used. Mammography may not be as available as general radiography but has superior spatial and contrast resolution (see Table 35-1).25,26
Contrast Radiography An important strategy for improving contrast resolution is the use of a contrast medium (like barium and iodine-based compounds). Barium studies are important for assessing the anatomy and motility of the esophagus and gastrointestinal tract. Barium has excellent coating properties for evaluating mucosal abnormalities. The barium GI series can yield important information regarding the origin of a mass, mass effect, or obstruction.16 A contraindication of barium for GI studies is in cases of suspected viscous rupture. Barium leakage into the peritoneum can have catastrophic consequences.10 Iodine-based contrast medium is less effective orally for evaluating the GI tract but crucial for urogenital imaging. Intravenous urography increases the conspicuity of the kidneys, ureters, and urinary bladder. Direct injection through the urethra is useful for assessing the urethra
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Fig. 35-2 Lateral (A), ventrodorsal (B), and lateral oblique (C,D) projections of the skull of a rabbit. The ventrodorsal view allows for direct comparison of the tympanic bullae on one projection. The marker on the obliqued views denotes the more ventrally directed bulla. Obliqued projections should be positioned similarly (in this case, latero-40 degrees ventral-laterodorsal) to make the comparisons. Despite slight differences in obliquity, the additional views (including the lateral projection) support mild sclerosis and thickening of the left tympanic bulla relative to the right.
Table 35-1 Comparison of the Common Imaging Modalities Available to Small Mammal Practitioners
Radiography Mammography Ultrasound CT MRI Scintigraphy
Cost
Availability
Anesthesia/ Sedation
Radiation Producing
Spatial Resolution
Contrast Resolution
+ + +(+) +++ ++++ +
++++ + +++ ++ + +
± ± Seldom Usually Always ±
++ + None +++(+) None ++
++ +++ +++(+) +++(+) ++(+) +
++ +++ ++ +++ ++++ +
(+) Indicates order of magnitude from least (+) to most (++++).
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Fig. 35-3 Transverse and sagittal reformatted CT images (A,B) compared with T2-weighted transverse and sagittal MRIs (C,D) of a normal ferret’s brain. Notice the excellent contrast resolution in the MR relative to CT image, differentiating gray matter, white matter, and ventricles. MRIs are acquired in different planes, whereas CT images are usually acquired in a single plane (transverse) and then reformatted to produce other planes, like the sagittal view in (B).
(positive contrast urethrogram) and urinary bladder (doublecontrast or positive-contrast cystograms).16 Readers are encouraged to follow proper technique in performing contrast studies.16,24 Common mistakes include (1) forgetting to make survey radiographs (radiographs made immediately before contrast administration), (2) too little contrast medium administered, and (3) too few radiographic views made after the contrast medium is added.
Digital Radiography The advantages of digital radiography (DR) over film-based (analog) radiography are numerous and, in this author’s opinion, outweigh the disadvantages. Dissemination of images to different people in different places can affect patient management. Internet connections and CDs allow emergency clinics, specialty hospitals, and private practices to interact and share imaging information at the same time. Image archiving is also improved with DR. Images can be stored and retrieved from a server without using a dedicated room to house bulky films, which can be misplaced. The space, breaking parts, and toxic chemicals of a darkroom/ processor are things of the past with DR. Digital radiography also provides a wide range of exposures for an acceptable image, thus decreasing the number of retakes for over- and underexposure. Finally, contrast resolution is improved, allowing for exquisite detail to evaluate all anatomic areas, from lungs to bones.1,5
A disadvantage of DR over film is a general reduction in spatial resolution. As stated previously, spatial resolution is a crucial parameter of image quality, particularly for small patients. Slow speed, high-detail radiographs, and mammography units provide optimal spatial resolution that is unmatched by most digital solutions.1,5 However, the differences in spatial resolution may not be clinically relevant.14,22 The technology of DR is improving rapidly, thus further blurring the clinical relevance of the reduced spatial resolution of DR over film.
Ultrasound Ultrasound provides an enormous amount of diagnostic information while at the same time being safe, seldom requiring sedation or anesthesia, and not producing any ionizing radiation. Ultrasound provides real-time information for important physiologic processes such as cardiac motility. The components that influence the image quality of ultrasound for small patients include high frequency, broad- bandwidth transducers, electronic focusing/beam steering, tissue harmonics, and spatial compounding capabilities. Higher- frequency transducers provide better spatial resolution at the cost of diminished depth penetration. Inability to image tissues deep to the skin is generally not a problem with small mammals. A high-frequency transducer can have a large, bulky footprint (surface that touches the patient’s skin), which may be
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impractical on very small patients. A broad-bandwidth transducer allows the operator to switch to lower frequencies when deeper structures are being imaged. Broad-bandwidth technology and electronic focusing/beam steering also provide the capabilities for image optimization with harmonics and spatial compounding, the details of which are described elsewhere.20 An important application of ultrasound is needle guidance for aspirating fluid or masses and for biopsy (Fig. 35-5). Aspirates can be extremely diagnostic for distinguishing inflammatory or infectious conditions from neoplastic processes.3 A caveat to this is the thick fluid within some abscesses, which can be extremely difficult to pull into a thin needle. Neoplastic processes that exfoliate well include round cell tumors and some carcinomas. Peritoneal and pleural fluid aspirates can help to identify a cause of the fluid accumulation.3,17 Definitive histopathologic diagnoses can be made with an ultrasound-guided biopsy sample, but this requires chemical restraint and operator skill. Ultrasound provides the best images of organs that are largely composed of fluid and soft tissue. The abdomen is ideally suited for this. Gas and bone adversely reflect ultrasound waves, decreasing overall image quality. As a result, lung and gas-filled bowel are common barriers to lesion identification. In general, the ferret gastrointestinal ultrasound examination can provide relatively better image quality and potentially more diagnostically useful information than some gas-filled rabbit abdomens.
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B Fig. 35-4 Lateral (A) and ventrodorsal (B) projections, the latter a horizontally directed projection with the patient in left lateral recumbency. The patient is an adult male guinea pig with anorexia and abdominal distention. The horizontally directed beam allows for confirmation of pneumoperitoneum, a surgical emergency for a ruptured viscus. In this case the stomach was ruptured. Notice the gas cap and air-fluid interface in the nondependent portion of the abdomen in (B). This type of radiograph must be made with the patient in left lateral recumbency and not right lateral recumbency, where a gas-cap could be confused with normal fundic gas.
Computed Tomography Computed tomagraphy has become more available in exotic animal veterinary practice; as a result, more literature is available to substantiate its diagnostic utility.7,13 The process of acquiring CT images usually requires general anesthesia. Helical and multidetector CT technology has increased acquisition speed to a point where immobilization devices alone may be used for some acquisitions. These faster CT units are expensive.
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B Fig. 35-5 A rabbit presenting with right-sided exophthalmos, confirmed on contrast-enhanced CT (A). A rim-enhancing retro-orbital mass related to a tooth root abscess was suspected from the CT exam. Color Doppler ultrasound confirmed that the mass was mainly fluid-filled with a peripheral vascular supply. Most importantly, ultrasound aided in directing an 18-gauge needle into the mass for cytologic confirmation of an abscess (B). The arrow denotes the needle in the center of the mass.
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Fig. 35-6 Lateral (A) and ventrodorsal (B) projections of the lumbar vertebral column in a rabbit with progressive lumbar pain and a previous diagnosis of uterine carcinoma. Compare the radiographs with the transverse CT image at the level of the last thoracic vertebral body (C). Notice the extensive bony lysis and associated soft tissue mass of the right pedicle and body. Compare this with the normal L1 vertebra caudally (D). CT removes superimposed structures that would otherwise obscure changes in anatomy on radiographs.
However, slower-speed scanners are economically priced. At our institution, the price of a non-contrast-enhanced skull CT (with our single detector) approximates the price of a four-view radiographic examination of the skull. Both studies require anesthesia or heavy sedation, but the CT scan is technically easier to perform and provides much more information. CT uses a beam of ionizing radiation opposite a detector rotating around a patient. The result is a computer mapping of the different x-ray-attenuating characteristics of tissues. With radiography, the three dimensions of the patient are summated to produce a two-dimensional shadow of x-ray attenuation characteristics. With CT, the third dimension (slice thickness) is thin and minimally contributes to the summed image. A major advantage of CT over standard radiography is removal of superimposed tissues that would obscure anatomy (Fig. 35-6). A CT image includes more shades of gray (greater contrast resolution) than a radiographic one. While radiography provides five shades of gray (gas, fat, fluid/soft tissue, bone, metal), CT can distinguish between thousands of shades of gray, including a distinction between soft tissue and fluid. Contrast resolution
is further improved with the addition of an intravenous contrast medium, increasing the conspicuity of lesions with greater blood flow or capillary permeability (see Fig. 35-5, A).5,6,13 Clinicians can utilize CT for a variety of clinical situations. Subtle bone fractures are better defined with CT than with other modalities. CT also defines extent of disease and the effect of disease on surrounding normal anatomy.5,7,13 A well-documented benefit of CT lies in evaluating dental disease in rabbits (see Chapter 33).2,13 The anatomy of the rabbit’s mouth limits oral examination and radiography lacks sensitivity for ruling out dental disease. CT can help differentiate which specific tooth root is affected in cases of severe tooth root abscessation and osteomyelitis (Fig. 35-7). Images can be reformatted into different scan planes other than the transverse plane. This allows the observer to appreciate disease, as it is oriented relative to surrounding normal tissues. Three-dimensional and multiplanar reformatting help the radiologist to communicate with surgeons, owners, and other veterinarians and help to orientate surgeons for surgical planning, including approach and regional vasculature.2,5,7,23 At our institution, CT is an important tool for surgical planning
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Fig. 35-7 A common indication for CT is for the assessment of dental disease in rabbits. From left to right, transverse, dorsal, and parasagittal images of a rabbit with expansile bony lysis of the right hemimandible secondary to tooth root abscess. (Images courtesy of Dr. Peter V. Scrivani, Cornell University, Ithaca, NY.)
in adrenalectomies in ferrets (to assess tumor thrombus) and any thoracic surgery (to determine surgical approach and vascular environment in mediastinal masses, such as thymomas).
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Magnetic Resonance Imaging Clinical use of MRI in small mammals is limited, despite the enormous library of literature dedicated to MR of rabbits and rodents in research settings. Availability, cost, and long scan times are the major limiting factors. Exquisite spatial resolution of very small patients is possible with high-field-strength magnets (7 Tesla and up).4 However, these magnets are primarily used in research settings. Most veterinary clinics practice with 1.5 T magnets or lower.19,21 With all magnets, image quality tends to be proportional to scan time. Scanning protocols try to strike a balance between image quality and scan time. Notice that the MRI in Figure 35-3 has a stippled appearance, secondary to a low signal-to-noise ratio. Longer scan time can improve on this. MRI takes advantage of the resonance properties of protons within organs, allowing for subtle differentiation in the contrast of soft tissues. Assessment of the brain and spinal cord by MRI has revolutionized the practice of veterinary neurology. MRI also has the advantage of CT in reducing superimposition with thin scan slices. Unlike CT, MRI scans can be acquired in different planes. As a result, an MRI has a similar or even superior capacity to assess disease in relation to surrounding anatomy and to aid surgical planning.
Nuclear Scintigraphy With nuclear scintigraphy, a radioactive substance is injected into the patient; this then localizes to a tissue of interest and emits radiation. Tissue localization of a radionuclide or radiopharmaceutical gives this modality an important advantage over all other modalities: assessment of physiologic processes. For example, technetium pertechnetate, the most common radionuclide used, has similar physiologic properties to negatively charged ions in the body, like iodine. Technetium can localize into tissues like the thyroid lobes, which take up
Fig. 35-8 One-year-old male guinea pig evaluated for a neck mass that was palpated on physical examination. This ventral image of the neck was obtained after technetium pertechnetate was injected intravenously (the patient’s nose is at the top of the image). Pertechnetate localizes to areas of iodine uptake, as in the thyroid lobes. Notice the large amount of uptake in the area of the left thyroid lobe relative to the right. A left thyroid adenoma was diagnosed histologically. (Image courtesy of Drs. Federica Morandi and Silke Hecht, University of Tennessee.)
iodine, to assess ion-trapping as a step in thyroid hormone production (Fig. 35-8). Radionuclides can also be bound to molecules that mimic physiologic processes. Renal scintigraphy with technetium bound to diethylenetriamine pentaacetic acid (DTPA) can quantitatively assess glomerular filtration rate from each kidney. Methylene diphosphonate (MDP)-labeled
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Fig. 35-9 Pre- (A) and post- (B) radiation therapy (RT) in a rabbit with a large thymoma. Radiation therapy reduced the tumor size by approximately 80% and greatly improved the patient’s clinical signs of dyspnea. Notice the severe dorsal and caudal displacement of the trachea before RT and the improvement of this mass effect after treatment. (Images courtesy of Dr. John Farrelly, Animal Medical Center, New York, NY.)
radiopharmaceuticals localize in areas of increased bone metabolism, pinpointing areas of bone metastasis and sources of orthopedic lameness. Nuclear scintigraphy has been used clinically to evaluate tissue ischemia and necrosis of the extremities of a ferret.8 A limitation of nuclear scintigraphy is poor contrast and spatial resolution relative to other imaging modalities.
ADVANCED TREATMENTS Thus far, our discussion has been limited to the use of radiation in imaging for diagnostic and prognostic purposes. Radiation oncology and interventional radiology are two veterinary specialties that work closely with diagnostic imaging to provide novel treatment options.
RADIATION THERAPY Radiation therapy (RT) targets high-energy radiation at diseased tissue to destroy cancerous or other space-occupying masses. RT has been described alone or in conjunction with surgery for the treatment of various tumors in ferrets.9,15,18 At our institution, the most common application of RT in small mammals is in treatment of thymomas in rabbits (Fig. 35-9).
INTERVENTIONAL RADIOLOGY Interventional radiology (IR) uses diagnostic imaging to direct minimally invasive procedures for diagnosis and treatment. Small needles and other tiny instruments are guided to areas of interest primarily by using fluoroscopy (to obtain a real-time radiographic image). Treatments are generally focused on providing an opening (e.g., stenting) or a closing (e.g., embolization) for diseased organs with a lumen. IR has long been used by veterinary cardiologists to treat congenital heart anomalies and
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Fig. 35-10 VD fluoroscopic image of a ferret after a stent was placed in the pyloric sphincter and duodenum. Contrast medium outlines the stomach and intestines with most of the contrast medium settled in the fundus. The stent provides a lumen for a stenosis, caused by lymphoma, at the pyloroduodenal junction. (Image courtesy of Dr. Chick Weisse, Animal Medical Center, New York, NY.)
place of pacemakers. A recent report describes the extraction of heartworms from a ferret with caval syndrome.28 The specialty is gaining momentum for many other applications, including those in small mammals, such as the placement of stents in animals with urethal blockage and pyloric stenosis (Fig. 35-10).
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References 1. Armbrust LJ. Digital images and digital radiographic image capture. In: Thrall DE, ed. Textbook of veterinary diagnostic radiology. 5th ed. Philadelphia: WB Saunders; 2007:22-37. 2. Arzi B, Sinclair KM. Diagnostic imaging in veterinary dental practice. Periapical lesions associated with the left maxillary molar teeth and associated cellulitis. J Am Vet Med Assoc. 2010;236:405-407. 3. Bonfanti U, Bussadori C, Zatelli A, et al. Percutaneous fine-needle biopsy of deep thoracic and abdominal masses in dogs and cats. J Small Anim Pract. 2004;45:191-198. 4. Bushberg JT, Seibert JA, Leidholdt EM, et al. Essential physics of medical imaging. 2nd ed. Philadelphia, Lippincott: Williams & Wilkins; 2002. 5. Capello V, Lennox AM, Widmer WR. The basics of radiology. In: Capello V, Lennox AM, eds. Clinical radiology of exotic companion mammals. Ames: Wiley-Blackwell; 2008:2-40. 6. Curry TS, Dowdey JE, Murray RC. Christensen’s physics of diagnostic radiology. 4th ed. Philadelphia: Lippincott Williams & Wlikins; 1990. 7. De Voe RS, Pack L, Greenacre CB, et al. Radiographic and CT imaging of a skull associated osteoma in a ferret. Vet Radiol Ultrasound. 2002;43:346-348. 8. Goggin JM, Hoskinson JJ, Carpenter JW, et al. Scintigraphic assessment of distal extremity perfusion in 17 patients. Vet Radiol Ultrasound. 1997;38:211-220. 9. Graham J, Fidel J, Mison M. Rostral maxillectomy and radiation therapy to manage squamous cell carcinoma in a ferret. Vet Clin North Am Exot Anim Pract. 2006;3:701-706. 10. Grobmyer AJ, Kerlan RA, Peterson CM, et al. Barium peritonitis. Am Surg. 1984;50:116-120. 11. Hammond G, Sullivan M, Posthumus J, et al. Assessment of three radiographic projections for detection of fluid in the rabbit tympanic bulla. Vet Radiol Ultrasound. 2010;51:48-51. 12. Kottwitz J. Horizontal beam radiography in ferrets. Exot DVM. 2004;6:37-41. 13. Mackey EB, Hernandez-Divers SJ, Holland M, et al. Clinical technique: application of computed tomography in zoological medicine. J Exot Pet Med. 2008;17:198-209. 14. Marolf A, Blaik M, Ackerman N, et al. Comparison of computed radiography and conventional radiography in detection of small volume pneumoperitoneum. Vet Radiol Ultrasound. 2008;49:227-232.
15. McBride M, Mosunic CB, Barron GH, et al. Successful treatment of a retrobulbar adenocarcinoma in a ferret (Mustela putorius furo). Vet Rec. 2009;165:206-208. 16. Morgan JP. Techniques in veterinary radiography. 5th ed. Ames: Iowa State University Press; 1993. 17. Morrison WB, DeNicola. Advantages and disadvantages of cytology and histopathology for the diagnosis of cancer. Sem Vet Med Surg (Small Anim). 1993;8:222-227. 18. Nakata M, Miwa Y, Nakayama H, et al. Localised radiotherapy for a ferret with possible anal sac apocrine adenocarcinoma. J Small Anim Pract. 2008;49:476-478. 19. Neuwirth L, Isaza R, Bellah J, et al. Adrenal neoplasia in seven ferrets. Vet Radiol Ultrasound. 1993;34:340-346. 20. O’Brien RT, Holmes SP. Recent advances in ultrasound technology. Clin Tech Small Anim Pract. 2007;22:93-103. 21. Pye GW, Bennett RA, Roberts GD, et al. Thoracic vertebral chordoma in a domestic ferret (Mustela putorius furo). J Zoo Wildl Med. 2000;31:107-111. 22. Schaefer CM, Greene R, Oestmann JW, et al. Digital storage phosphor imaging versus conventional film radiography in CTdocumented chest disease. Radiology. 1990;174:207-210. 23. Schultz RM, Wisner ER, Johnson EG, et al. Contrast-enhanced computed tomography as a preoperative indicator of vascular invasion from adrenal masses in dogs. Vet Radiol Ultrasound. 2009;50:625-629. 24. Schwarz LA, Solano M, Manning A, et al. The normal upper gastrointestinal examination in a ferret. Vet Radiol Ultrasound. 2002;44:165-172. 25. Silverman S. Diagnostic imaging of exotic pets. Vet Clin North Am Sm Anim Pract. 1993;23:1287-1299. 26. Steffanaci JD, Hoefer HL. Small mammal radiology. In: Hillyer EV, Quesenberry KE, eds. Ferrets, rabbits, and rodents: Clinical medicine and surgery. 1st ed. Philadelphia: WB Saunders; 1997:358-377. 27. Thrall DE, Widmer WR. Radiation physics, radiation protection, and darkroom theory. In: Thrall DE, ed. Textbook of veterinary diagnostic radiology. 5th ed. Philadelphia: WB Saunders; 2007:2-21. 28. Weisse CW, Berent AC, Todd KL, et al. Potential applications of interventional radiology in veterinary medicine. J Am Vet Med Assoc. 2008;10:1564-1574. 29. Williams J. Orthopedic radiography in exotic animal practice. Vet Clin North Am Exot Anim Pract. 2002;5:1-22.
CHAPTER
36
Hematology and Cytology of Small Mammals
Andrea Siegel, DVM, Andrew S. Loar, DVM, Diplomate ACVIM, and James Walberg, DVM, Diplomate ACVP
This chapter provides photomicrographs and other information that may be useful during the examination of blood smears and tissue aspirates from selected small mammals. More comprehensive discussions are available elsewhere regarding the evaluation of blood smears from many of these species; these texts are listed among the references.* While knowledge of the plausible differential diagnoses is often critical in the interpretation of cytologic findings, the same general principles of cytology used in dogs and cats can be applied to small mammals. The reference list below includes several excellent veterinary cytology references.† Some of the conditions described here can be accurately diagnosed in house without the delay and expense involved in using a commercial laboratory. Wright’s-Giemsa (modified Romanowsky), new methylene blue, and acid-fast stains were used on the specimens in these photomicrographs. Diff-Quiktype processing or other forms of three-step quick stains, while acceptable for the evaluation of up to several specimens per day, is considered less favorable for use in a busier in-house laboratory for the following reasons: (1) if a large number of slides are to be processed daily, Diff-Quik-type stains are expensive and labor-intensive; (2) jars containing Diff-Quik stains can easily be contaminated with bacteria and fungal elements, particularly when slides containing material from skin, otic, or fecal samples are processed; (3) in many nonneoplastic tissue cells, Diff-Quik staining may create nuclear textures that inaccurately *References 2-4, 7, 11, 18, 19. †References 1-3, 5, 6, 10, 14-16.
Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
indicate immature changes; (4) polychromasia of red blood cells (RBCs), an important manifestation of erythroid regeneration, is generally not evaluable on Diff-Quik-stained blood smears; and (5) these stains produce variable staining of mast cell granules, occasionally rendering them invisible. Most of the photomicrographs in this chapter were taken under immersion oil by using a 100× objective and 10× eyepiece, equaling 1000× magnification. For some lesions, evaluation of cytologic specimens can readily be performed by using the microscope’s 40× objective (high dry, 400× magnification). In fact, this lens can be more useful than the 100× lens because it provides a larger field of view, allowing the examiner to evaluate a larger portion of the slide in less time while revealing compelling cellular details and other features. Nonetheless, many clinicians and technicians are not satisfied with the quality of the resolution of this lens. In defense of the 40× objective, optically it is designed to be used principally with slides that have been cover-slipped. A good technique for temporary placement of a cover slip is to deposit a very small drop of immersion oil on the stained smear and then place a rectangular glass cover slip on the drop of oil. If resolution is then not improved upon examination of cover-slipped material, suspect that the lens may have been inadvertently soiled with immersion oil during previous use. A thorough cleaning of the external surface of the lens can be successful, using several cotton swabs thickly moistened with lens cleaner, followed by extended buffing with several dry cotton swabs. On microscopes used by numerous operators, and particularly those where the 100× (oil-immersion) lens receives heavy use, the 40× lens may require frequent cleaning.
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Fig. 36-1 Neutrophils. Peripheral blood smear—chinchilla.
Fig. 36-2 Lymphocytes. Peripheral blood smear—chinchilla.
These photographs show two mature (segmented) neutrophils, which contain faint acidophilic cytoplasmic granules. Compared with other mammalian species, neutrophils of chinchillas manifest a variation in segmentation; mature neutrophils frequently appear hyposegmented and can be misinterpreted as immature and erroneously classified as band or metamyelocyte forms. Moreover, automated analyzers may incorrectly identify these cells as being of mononuclear origin. (Modified Wright’s-Giemsa stain, 1000×-100× objective with 10× eyepiece.)
These photographs show a reactive, or stimulated, lymphocyte (principal field) and two smaller mature lymphocytes (inset, lower right). Generally a stimulated lymphoid cell appears slightly enlarged compared with a nonreactive form and shows increased cytoplasm while maintaining a mature, densely compacted/condensed (smudged) nuclear chromatin pattern. (Modified Wright’sGiemsa stain, 1000×.)
Fig. 36-3 Leukocytes. Peripheral blood smear—chinchilla. These
Fig. 36-4 Leukocytes. Peripheral blood smear—ferret. This field
photographs show a neutrophil (left) and an eosinophil (right, principal field) as well as a basophil (inset). Eosinophils generally contain prominent large pink- to orange-staining granules within clear to mildly basophilic cytoplasm. In most mammalian species, basophils tend to be the largest of the granulocytic cell types and reveal a blue-gray cytoplasm with variable numbers of darker-staining intracytoplasmic granules; these granules appear dark orange in this species, while in others they tend to be more basophilic, staining blue-black. (Modified Wright’s-Giemsa stain, 1000×.)
shows two mature segmented neutrophils in the upper right and left corners as well as a band form (lower right corner). Immature (band) neutrophils reveal less condensed nuclear chromatin patterns compared with mature neutrophils, without nuclear lobulation or segmentation; they generally show minimal constriction of the nuclear width. Note: Data derived from automated analyzers, including inclinic instruments or those at large commercial reference laboratories, do not identify band neutrophils, instead relying on manual review by technicians or pathologists. Often these reviews are not performed if other CBC parameters (e.g., WBC/RBC) are within or near normal reference intervals. Bands are misidentified as either monocytes or segmented neutrophils by automated hematology analyzers. (Modified Wright’s-Giemsa stain, 1000×.)
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Fig. 36-5 Leukocytes. Peripheral blood smear—ferret. These
Fig. 36-6 Platelet clumping. Peripheral blood smear—ferret.
photographs show a monocyte (principal field) and a mature segmented neutrophil (inset). In ferrets and most other mammalian species, monocytes are larger than neutrophils and reveal variable nuclear size and shape, ranging from ameboid to multilobular. Monocyte cytoplasm appears blue-gray and grainy and may contain vacuoles. Large mononuclear cells are often distributed toward the feathered edges and sides of blood smears and may be underrepresented within the monolayered body of the smear during manual evaluation. Automated analyzers often incorrectly identify, or interchange, monocytes, neutrophils (particularly immature forms), and lymphocytes. Mature lymphocytes (not present) are generally smaller than neutrophils and reveal densely stained, compacted to smudged nuclear chromatin with scant, basophilic-staining cytoplasm. (Modified Wright’s-Giemsa stain, 1000×.)
This field shows a large platelet clump (aggregation). The presence of large numbers of platelet clumps within a blood specimen may result in artifactually decreased platelet counts generated by automated methods, even if macroscopic clots are not observed. Moreover, clumps on a blood smear will falsely decrease manual platelet estimates. Platelets are oval to round, blue to gray, and may vary in size from one-fifth to greater than three-fourths the diameter of the RBCs. Larger platelets may be present in animals with increased platelet usage, such as that due to hemorrhage or exuberant systemic coagulation. In smears without platelet clumps, an acceptable method to estimate platelet numbers (per μL) using a 100× (oilimmersion) objective requires determining the average number of platelets in 10 microscopic fields (within the monolayer) and multiplying by 15,000. (Modified Wright’s-Giemsa stain, 1000×.)
Fig. 36-7 Leukocytes. Peripheral blood smear—guinea pig.
Fig. 36-8 Leukocytes. Peripheral blood smear—guinea pig. This
These photographs show a basophil (principal field) and an eosinophil (inset). In this species, basophil granules generally stain reddish-purple, while eosinophils typically contain large spherical orange granules that together occupy much of the cytoplasmic space. (Modified Wright’s-Giemsa stain, 1000×.)
field shows a neutrophil/heterophil (right) and a monocyte (left). The neutrophils of guinea pigs, hamsters, and gerbils are often referred as heterophils or pseudoeosinophils because they contain granules that stain eosinophilic in color with Romanowsky stains, as with the Wright’s-Giemsa method. Heterophils in these species perform similar functions as neutrophils; some clinicians interchange their names. In the guinea pig, eosinophils and heterophils are easily differentiated because of the size and number of the granules (see inset, Fig. 36-7); eosinophils have more numerous and larger round to rod-shaped granules. Eosinophils in many small mammal species often reveal a U-shaped nucleus. Automated hematology analyzers generally incorrectly identify heterophils/pseudoeosinophils as eosinophils. (Modified Wright’s-Giemsa stain, 1000×.)
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Fig. 36-9 Leukocytes. Peripheral blood smear—rabbit. These
Fig. 36-10 Lymphocytes with Kurloff’s bodies. Peripheral blood
photographs show, from top to bottom (principal field), a monocyte (note intracytoplasmic vacuoles), a lymphocyte, a neutrophil/ heterophil, and an eosinophil (inset). The neutrophils of rabbits are also referred to as heterophils because their cells do not stain neutral with Romanowsky stains. Note the more similar characteristics of eosinophils and heterophils in this species compared with guinea pigs. (Modified Wright’s-Giemsa stain, 1000×.)
smear—guinea pig. These photographs (principal field and inset) show individual lymphocytes, each containing an intracytoplasmic Kurloff’s body, which is reddish-purple and often larger than the adjacent nuclei. Lymphocytes with Kurloff’s bodies (also known as Foa-Kurloff cells) are unique to this species and may account for 3% to 4% of the circulating leukocytes and occasionally more. These structures are found in higher proportions of circulating lymphocytes in young guinea pigs as well as in adult females. Lymphocytes containing Kurloff’s bodies are believed to be analogous to large granular lymphocytes, or natural killer (NK) cells, in other mammals.2,19 (Modified Wright’s-Giemsa stain, 1000×.)
Fig. 36-11 Erythrocytes with Howell-Jolly bodies. Peripheral
Fig. 36-12 Leukocytes. Peripheral blood smear—mouse. These
blood smear—mouse. This field shows several red cells containing intracytoplasmic Howell-Jolly bodies (arrows). These small darkblue to black structures represent remnants of red cell nuclei and may be observed in low numbers within mature (anucleated) erythrocytes of healthy mammals. This photograph also reveals moderate polychromasia of the red cells. The proportion of red cells containing Howell-Jolly bodies may increase as a result of regenerative anemia or splenic dysfunction. Their presence should not be misinterpreted to suggest red cell parasites or Heinz bodies. (Modified Wright’s-Giemsa stain, 1000×.)
photographs show two neutrophils and an eosinophil (principal field) and a monocyte (inset) as well as scattered polychromatic red cells. Mature neutrophils of rats and mice generally have a clearstaining cytoplasm, but they may contain a few dust-like reddish granules and appear diffusely pink with Romanowsky-type stains. Granulocytes of rodents often have nonlobulated nuclei; these may be horseshoe- or ring-shaped. Rodent eosinophils show considerably fewer and less distinct granules than those of other small mammals. In rats and mice, an age-dependent variation exists in the neutrophil:lymphocyte (N:L) ratio, with the lymphocyte concentration decreasing and the neutrophil concentration increasing as the animal ages.4,19 In rodents, automated analyzers often incorrectly identify a variable proportion of neutrophils and/or lymphocytes as monocytes. (Modified Wright’s-Giemsa stain, 1000×.)
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Fig. 36-13 Erythrocyte polychromasia and reticulocytes. Peripheral blood smear—rabbit. These photographs show several polychromatic red cells (arrows) stained with a modified Wright’sGiemsa stain (principal field) and aggregate reticulocytes stained with new methylene blue stain (inset). Polychromasia, a bluish-gray coloration, is caused by higher quantities of ribosomal RNA present in immature, anucleated red cells. Compared with the red cell life span in other mammals, the relatively shorter half-life of rabbit erythrocytes (57-67 days) results in a higher proportion of polychromatic erythrocytes and reticulocytes. Reticulocytes in adult rodents, rabbits, and hystricomorphs (guinea pigs and chinchillas) range between 2% to 7%. Reticulocyte staining requires a 15-minute incubation of any volume of whole fresh blood with an equal quantity of new methylene blue (0.5% in saline). Smears are then prepared and allowed to air-dry prior to examination. (1000×.)
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Fig. 36-14 Reticulocytes. Peripheral blood smear—ferret. These photographs show two aggregate (A) and three punctate (B) forms of reticulocytes. Ferrets are similar to cats and unique in having two varieties of reticulocytes. With new methylene blue stain (see Fig. 36-13), aggregate reticulocytes reveal clumped or coalescing particulate reticulum, while punctate forms contain smaller and fewer discrete dots without clumping. Increased numbers of aggregate forms represent the initial regenerative response to moderate or severe blood loss. Punctate reticulocytes generally increase later but persist for longer in circulation and may be the primary response to mild anemia. On a Wright’s-stained blood smear, only aggregate reticulocytes will be polychromatophilic. Thus, in most species, polychromatic red cells represent aggregate reticulocytes; however, in ferrets and cats, not all reticulocytes may be polychromatic. (New methylene blue stain, 1000×.)
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Fig. 36-15 Saccharomyces species yeast organisms. Feces— rabbit. These photographs show unstained (principle field) fecal smear preparations using standard zinc solution flotation; the yeast organism Cyniclomyces guttulatus (also referred to as Saccharomyces guttulata) is demonstrated. In addition, the inset shows a Wright’sGiemsa preparation of a fecal smear in the same animal. These are very large structures measuring 10 to 30 μm in length. This yeast is a normal inhabitant of the cecum in rabbits and is occasionally identified from fecal preparations in other species, including dogs.8 While their clinical significance is uncertain, yeast organisms from the Saccharomyces family are unlikely to represent pathogens in the gastrointestinal tract of most mammalian species. (400×.)
Fig. 36-17 Lymphosarcoma. Fine-needle aspirate of a subcutaneous mass at the abdominal wall—hamster. This photograph shows a homogenous population of large, generally round cells each revealing a relatively small amount of basophilic cytoplasm with coarse granular nuclear chromatin patterns and one or more nucleoli. The cells most resemble medium-sized lymphoblasts and are generally as large as or larger than neutrophils/heterophils— in this species, approximately twice the diameter of a RBC. Blastic lymphocytes can be verified based on size, the presence of nucleoli and/or the predominance of an immature nuclear chromatin pattern. Aspirates from lymphoid tissue showing greater than 50% of the intact lymphoid elements as blast forms confirm lymphoma. Lower proportions of blast cells (greater than 35%-40%) are consistent with lymphoma in aspirates from nonlymphoid sites. (Modified Wright’s-Giemsa stain, 1000×.)
Fig. 36-16 Lymphoid hyperplasia. Fine-needle aspirate of peripheral lymph node—ferret. These photographs show a heterogenous population of lymphoid cells predominantly consisting of small mature lymphocytes (smaller in diameter than neutrophils) with low numbers of medium-sized reactive forms, scattered mature plasma cells, and occasional large lymphoblasts. The vacuolated cells are macrophages that contain an admixture of chylous material and phagocytized red cells. Rare extracellular lymphoglandular bodies are seen; these are small spherical gray structures representing variable-sized portions of unused cytoplasmic material. Lymphoglandular bodies are characteristic of reactive lymphoid tissue but do not suggest neoplasia unless they are present in nonlymphoid tissue. Finding that far less than 50% of the intact lymphoid cells are immature precludes a diagnosis of lymphoproliferative disease. Biopsy verified a diagnosis of lymphoid hyperplasia. (Modified Wright’s-Giemsa stain, 200×.)
Fig. 36-18 Lymphosarcoma. Fine-needle aspirate of peripheral lymph node—ferret. The photograph reveals a population of variably sized large round cells with immature, generally homogenous, and finely reticular nuclear chromatin patterns. Few cells contain obvious nucleoli. The cytoplasm of most cells is basophilic and granular. These elements most resemble immature blastic lymphocytes, although, based on a significant variation in cell and nuclear sizes, they clearly do not represent a monomorphic population. Numerous lymphoglandular bodies are also noted. Only a very few small mature lymphocytes are identified (arrows), although these assist in estimating the size of the much larger blast cells. Demonstrating that more than 50% of the intact lymphoid cells present are blast forms is sufficient to confirm lymphoma from this aspirate. (Modified Wright’s-Giemsa stain, 1000×.)
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Fig. 36-20 Well-differentiated malignant mesenchymal neo-
Fig. 36-19 Septic suppurative inflammatory infiltrate. Fine- needle aspirate of a subcutaneous mass on a limb—hamster. This field shows primarily degenerative neutrophils with some activated macrophages against a particulate basophilic background of extracellular protein material. Note the loss of nuclear chromatic texture and lighter staining characteristics of the (neutrophil) nuclei, typical of degenerative changes. Within the relatively clear-staining cytoplasm of several neutrophils are numerous, generally amorphous oval to elongated coccobacilli, verifying a septic process. (Modified Wright’s-Giemsa stain, 1000×.)
plasm (probable nerve sheath tumor). Fine-needle aspirate of subcutaneous mass at facial region—rabbit. These photographs show a low-power (principal field) magnification of numerous loosely cohesive, well-differentiated spindle-shaped fusiform to stellate (tricornered) mesenchymal cells. These cells reveal relatively uniform nuclear sizes and shapes, frequently with elongated tendrils and diaphanous projections of cytoplasm. High-power magnification (inset) shows variably homogenous, often stippled nuclear chromatin patterns, frequently with multiple nucleoli and occasional vacuolated cytoplasm. This view also emphasizes the linear extensions of the cytoplasm as well as the presence of an amorphous eosinophilic matrix. Biopsy confirmed a malignant nerve sheath tumor. (Modified Wright’s-Giemsa stain, 200× and 1000×.)
Fig. 36-22 Benign mammary epithelial hyperplasia. Fine-needle
Fig.
36-21 Malignant epithelial tumor (adenocarcinoma). Fine-needle aspirate of subcutaneous mass—ferret. These photographs show a high-power magnification (principal field) of tightly cohesive, variably sized round to oblong epithelial cells with large, often irregular and occasionally multiple nucleoli. Some cells also reveal molding of adjacent cell nuclei and minimal regulation of neighboring cells’ nuclear borders (crowding)—a strong malignant feature. The lower-power field (inset) reveals dense cell clumping and absence of discernible cytoplasmic borders with a circular cell organization that resembles a rosette or acinar structure and nuclei arranged around a central lumen; this is characteristic of glandular epithelial tissue. Biopsy confirmed an adenocarcinoma, likely of apocrine gland origin. (Modified Wright’s-Giemsa stain, 200× and 1000×.)
aspirate of a subcutaneous mass in the mammary region—guinea pig (female). These photographs show a low-power magnification (principal field) of densely cohesive, generally uniform mammary epithelial elements characteristic, in this species, of mammary hyperplasia. The high-power field (inset) reveals activated macrophages that contain basophilic granules and amorphous intracytoplasmic material suggestive of mammary secretions, including milk proteins. Biopsy confirmed benign mammary epithelial hyperplasia with a cystic component. (Modified Wright’s-Giemsa stain, 1000×.)
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Fig.
Fig. 36-23 Benign sebaceous epithelial structure. Fine-needle aspirate of a subcutaneous mass on the body wall—ferret. The field shows a uniform population of loosely cohesive epithelial cells that are large and round, with small nuclei. These cells contain numerous intracytoplasmic opaque vacuoles or may reveal an amorphous foamy cytoplasmic appearance characteristic of sebaceous-type glandular products. Fine-needle aspirates of benign basal cell lesions also frequently yield these forms of sebaceous glandular elements. Biopsy confirmed a benign sebaceous epithelioma. (Modified Wright’s-Giemsa stain, 200×.)
36-24 Poorly differentiated malignant mesenchymal tumor (sarcoma). Fine-needle aspirate of a subcutaneous mass in the cervical region—hamster. These photographs show a low-power magnification (principal field) containing numerous fusiform and spindle-shaped cells in loosely cohesive clusters with moderate variation in nuclear size and shape. The higher-power magnification (inset) shows several anaplastic round to oval cells revealing significant variation in size and shape, with generally oval nuclei. The nuclei contain multiple irregular and occasionally very large nucleoli. Biopsy confirmed a high-grade, poorly differentiated sarcoma. (Modified Wright’s-Giemsa stain, 200× and 1000×.)
Fig. 36-25 Poorly differentiated malignant tumor (sarcoma).
Fig. 36-26 Malignant melanoma. Fine-needle aspirate of a
Fine-needle aspirate of subcutaneous mass on a limb—rabbit. This field reveals large anaplastic round to angular to fusiform cells that contain irregular nuclei with coarse to finely reticular nuclear chromatin patterns. Some binucleated cells are present. While loosely adherent, the cells do not show the characteristics of cohesive clumping suggestive of epithelial tissue. One bizarre mitotic figure is also noted. Biopsy findings confirmed a high-grade anaplastic sarcoma. (Modified Wright’s-Giemsa stain, 1000×.)
cutaneous mass on the truncal region—hamster. This photograph shows a population of large, discrete, round to oblong cells with several ominous features of malignancy, including finely stippled nuclear chromatic patterns, irregular and often multiple nucleoli, and variable nuclear sizes and shapes. In addition, there are abundant intracellular and extracellular blue-black granules, representing melanin pigment. The volume of pigmented material is sufficiently high to conceal many of the cells; also, the excess pigment absorbs much of the stain, often obscuring the cells’ structural details, including delineation of the cytoplasmic borders. Two cells containing larger granules represent melanophages. Melanomas are the most frequently reported cutaneous tumors in hamsters.17 (Modified Wright’s-Giemsa stain, 1000×.)
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Fig. 36-28 Extramedullary hematopoiesis (EMH). Fine-needle
Fig. 36-27 Malignant neuroendocrine neoplasm, adrenal gland. Fine-needle aspirate of adrenal mass—ferret. This photograph represents a high-power-field of a loose cluster of well-differentiated, generally uniform epithelial cells. The epithelial elements reveal indistinct cell borders, suggesting stripped nuclei within a pool of cytoplasmic material. There is mild to minimal evidence of dysplasia or atypia, with only modest variation in cell size (anisocytosis) or nuclear size (anisokaryosis). These features are characteristic of tumors originating in neuroendocrine organs, including those from adrenal, pancreatic, gastrointestinal, and vasoactive tissues. In ferrets, neuroendocrine tumors of the adrenal cortex and pancreas (insulinoma) are relatively common13 (see Chapter 7). (Modified Wright’s-Giemsa stain, 1000×.)
Fig. 36-29 Cryptosporidium species. Fecal smear. This field shows vast numbers of oocysts of the protozoan Cryptosporidium. The specimen was prepared by using an acid-fast stain method (TB Quick Stain, Becton Dickinson Microbiology Systems, Sparks MD), often referred to as a “cold” acid-fast stain. The cysts are 4 to 7 μm in diameter (slightly smaller than a RBC) and are weakly acid-fastpositive. Their presence in high numbers within a fecal smear is considered diagnostic for the infection. (1000×.)
aspirate from the spleen—ferret. These photographs show a mixed population of nucleated hematopoietic precursor cells predominantly consisting of late-stage erythroid elements. Note several stages of metarubricytes, which show densely clumped/smudged (dark staining) nuclear chromatic patterns, spherical nuclei, and variable amounts of polychromatic cytoplasm. Contrast these cells with the numerous small lymphocytes (lighter staining nuclei) and relatively fewer numbers of mature plasma cells present, the latter showing more abundant basophilic cytoplasm and more eccentric nuclear placement. Also note a large megakaryocyte (inset). In the authors’ experience, EMH is by far the most common cytologic finding from ferret splenic aspirates and is frequently associated with splenic enlargement, splenic nodules, and suspicious ultrasonographic textural changes. (Modified Wright’s-Giemsa stain, 400×.)
Fig. 36-30 Mycobacteriosis. Fine-needle aspirate of a peripheral lymph node—cat. The field shows numerous thin rod-shaped organisms, both extracellularly and within macrophages, compatible with Mycobacterium species. This specimen was initially stained with a Wright’s preparation and subsequently counterstained using a “cold” acid-fast method (see previous Fig. 36-29). The organisms, being acid-fast-positive, reveal a reddish color. Counterstaining previously stained specimens is an option when no unstained slides are available. In the absence of an acid-fast stain, these organisms generally appear as negative (clear) thread-like structures, often seen thickly packed within macrophages. Although definitive culture techniques are necessary for identification, when acid-fast organisms are so numerous they are typically Mycobacterium avium. Infection with this bacterium has been documented in most small mammal species. (Modified Wright’s-Giemsa counterstained via acidfast method, 1000×.)
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Fig. 36-31 Mast cell neoplasm. Fine-needle aspirate of a cutaneous mass on a limb—ferret. This field shows many large, discrete round cells associated with variable numbers of purple or metachromatic granules. As noted for melanin-containing specimens (Fig. 36-26), the abundant granules often absorb stain, rendering other cellular features, particularly cytoplasmic borders and nuclear chromatin textures, less obvious. Conversely, mast cell granules from some of these tumors may not stain. Lastly, specimens stained with many of the Diff-Quik-type techniques do not reveal sufficient distinctive mast cell granules to confirm the diagnosis. Thus, in each of the latter two groups, cytology may simply suggest an undistinguished form of round cell neoplasia. (Modified Wright’s-Giemsa stain, 1000×.)
Fig. 36-33 Lipoma. Fine-needle aspirate of subcutaneous mass at the inguinal region—rabbit. This photograph shows several loosely cohesive lipocytes characterized by large vacuolated cells with thin membranes that frequently appear wrinkled or creased. Lipocyte nuclei are quite small and ovoid and are often displaced to the periphery of the cell by the intracytoplasmic lipid material. Lipomas have been demonstrated in most small mammalian species, including guinea pigs, ferrets, and small rodents. Slides made using material aspirated from a lipoma, or normal fat, consist predominately of oily fatty droplets and debris. While the greasy appearance of these unstained slides is readily visible, routine staining generally dissolves extracellular fat, rendering many specimens virtually acellular. (Modified Wright’s-Giemsa stain, 1000×.)
Fig. 36-32 Helicobacter species. Impression smear from a gastric mucosal biopsy—cat. This field shows numerous coil-shaped and often curved bacterial structures characteristic of all species of Helicobacter. The organism is gram-negative and the spiral-shaped forms are 7 to 10 μm in length. While the coil-spring variety are relatively common in gastric specimens from cats and dogs, it may be less frequently observed in specimens from ferrets or other small mammalian species (authors’ personal observations). The presence of abundant organisms from gastric specimens implies an overgrowth and may be of pathologic significance; however, finding cytologic evidence of Helicobacter species does not necessarily confirm the primary etiology of a gastrointestinal disorder.9 (Modified Wright’s-Giemsa stain, 1000×.)
Fig. 36-34 Chordoma. Fine-needle aspirate of a subcutaneous mass from the dorsal tail base—ferret. This high-power field shows several large, foamy-appearing cells with variably placed nuclei often containing multiple prominent nucleoli. These are characteristic of physaliferous cells, elements distinctive to chordomas. These tumors are more prevalent in ferrets than in other mammals and often contain an amphophilic eosinophilic cartilaginous matrix. Chordomas likely originate from primitive notochord tissue, possibly associated with the intravertebral disc. Although most commonly found adjacent to the tail, they have also been identified near the cervical spine in ferrets.15,16 (Modified Wright’s-Giemsa stain, 1000×.)
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Fig. 36-35 Mammary carcinoma. Fine-needle aspirate of a sub-
Fig. 36-36 Benign basal cell proliferation (basal cell tumor).
cutaneous mass at the mammary region—rabbit (female). This photograph shows a cohesive cluster of anaplastic round to polygonal epithelial cells. The cells exhibit variable amounts of cytoplasm, coarsely granular nuclear chromatin patterns, and moderate anisokaryosis as well as prominent, irregular, and often extremely large nucleoli. Moreover, many of the cells show nuclear crowding and molding. Biopsy confirmed an anaplastic mammary adenocarcinoma. (Modified Wright’s-Giemsa stain, 1000×.)
Fine-needle aspirate of a subcutaneous mass in the head-and-neck region—rabbit. This low-power field contains many densely cohesive round epithelial cells with oval nuclei and scant cytoplasm. While most of these cells are uniform, a few show mild dysplasia to modest atypia. Some cells are found in small papillae and ribbon-like projections, while others are organized in a curled or bent-knuckle arrangement. In other fields (not shown), some cells contain blackish-brown granules characteristic of melanin pigment. These features indicate a benign basal cell proliferation characteristic of a basal cell tumor. Frequently these lesions contain hyperplastic sebaceous cell elements. Interestingly, evaluation of the specimen at higher magnification often suggests a less well differentiated lesion. In other species, this tumor has also been classified histologically as a trichoblastoma.12 (Modified Wright’s-Giemsa stain, 200×.)
Fig. 36-37 Plasma cell tumor. Fine-needle aspirate of cutaneous
Fig. 36-38 Modified transudate. Abdominal effusion—ferret.
mass on limb—ferret. These photographs reveal a population of discrete medium-sized round cells containing eccentrically placed nuclei that show thickly stippled to smudged nuclear chromatin patterns but no obvious nucleoli. These features most resemble neoplastic plasma cells, with other forms of round cell tumor (including lymphoma, melanoma, histiocytoma and mast cell tumor) being considerably less likely. Occasional cells show a perinuclear clearing characteristic of plasma cells and probably representing the presence of the Golgi apparatus. The smaller field (inset) shows a binucleated cell, which is also frequently noted with plasma cell proliferation. Biopsy confirmed a benign cutaneous plasmacytoma. These tumors are generally cured with excision and rarely manifest paraproteinemia (hypergammaglobulinemia). (Modified Wright’sGiemsa stain, 1000×.)
These photographs represent a concentrated (cytospin) preparation of an abdominal fluid specimen. The nucleated cell count was 5,100/μL (centrifuging of the sample suggests a higher cellularity) and the protein level was 3.6 g/dL. The specimen contains an admixture of primarily nondegenerate neutrophils with fewer activated macrophages and rare small lymphocytes. Reactive mesothelial elements with pseudopod-like cell membrane projections and occasional binucleated forms are seen in the inset photo (arrows). No organisms are identified. The cell count and protein concentration classify the effusion as a modified transudate, a relatively nonspecific diagnosis. This form of abdominal fluid may be due to virtually any cause of local lymphatic/venous obstruction, including cardiac failure, certain forms of hepatic disease, and mass lesions (including nonexfoliative tumors). (Modified Wright’s-Giemsa stain, 1000×.)
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References
Fig. 36-39 Artifact: ultrasound gel (lubricant). Fine-needle aspirate of the liver (ultrasound-directed)—rabbit. This photograph reveals moderate numbers of red blood cells as well as a large amount of amorphous particulate magenta-staining material characteristic of virtually any form of water-soluble lubricant, including ultrasound gel. No hepatocytes are identifiable. This is an extremely common artifact derived from ultrasound-directed aspirates and occurs when the operator inserts the needle through a cutaneous site where the gel has been deposited. The material tends to attach avidly to cellular elements within the smears and greatly obscures large portions of this field. In addition, the gel often absorbs relatively more stain than the cellular components do. (Modified Wright’s-Giemsa stain, 1000×.)
Fig. 36-40 Artifact: formalin contamination. Impression smear of an excised cutaneous mass—guinea pig. This photograph shows an artifact typical of smears that have been in close contact with formalin. This often occurs when unstained glass slides are placed in an airtight container with one or more biopsy jars. The formalin fumes from the biopsy specimen coat the slides, which can severely inhibit the subsequent ability to stain the specimen. The cells and extracellular background tend to develop an homogenous blue hue, generally preventing evaluation of cellular features. Clinicians are urged to package unstained glass cytology slides in at least two sealed plastic bags before placing them in the same shipping container with biopsy samples. (Modified Wright’s-Giemsa stain, 100×.)
1. Baker R, Lumsden JH. Color atlas of cytology of the dog and cat. St. Louis: Mosby; 2000. 2. Campbell TW, Ellis CK. Avian and exotic animal hematology and cytology. 3rd ed. Ames, Iowa: Blackwell Publishing; 2007; 119-127. 3. Cowell RL, Tyler RD, Meinkoth JH. Diagnostic cytology and hematology of the dog and cat. 3rd ed. Canada: Mosby; 2008. 4. Everds NE. Hematology of the laboratory mouse. In: Fox JG, Barthold SW, Davisson MT, et al, eds. The mouse in biomedical research. Volume III. 2nd ed. Burlington, MA: Academic Press; 2007:133-170. 5. Garner MM. Cytologic diagnosis of diseases of rabbits, guinea pigs, and rodents. Vet Clin North Am Exot Anim Pract. 2007;10:25-49. 6. Greene CE. Infectious diseases of the dog and cat. 3rd ed. Philadelphia: WB Saunders; 2006. 7. Hadley LT, ed. Hematology and related disorders. Vet Clin North Am Exot Anim Pract. 2008;11:423–610. 8. Hersey-Benner C. Diarrhea in a rabbit. Cyniclomyces guttulatus yeast. Lab Anim (NY). 2008;37:347-349. 9. Johnson-Delaney CA. The ferret gastrointestinal tract and Helicobacter mustelae infection. Vet Clin North Am Exot Animal Pract. 2005;8:197-212. 10. Li X, Fox J, Erdman S, et al. Spontaneous neoplasms in ferrets: a review of 204 cases. (Abstract). Proceedings. 47th Annu Meet Am Coll Vet Pathol. 1996;33:590. 11. Marshall KL. Rabbit hematology. Vet Clin North Am Exot Anim Pract. 2008;11:551-567. 12. Meuten DJ. Tumors in domestic animals. 4th ed. Ames, Iowa: Blackwell; 2002;58-60. 13. Pilny AA, Chen S. Ferret insulinoma: diagnosis and treatment. Comp Cont Ed Pract Vet. 2004;26:722-729. 14. Pinches MD, Liebenberg G, Stidworthy MF. What is your diagnosis? Preputial mass in a ferret. Vet Clin Path. 2008; 37:443-446. 15. Rakich PM, Latimer KS. Cytologic diagnosis of diseases of ferrets. Vet Clin North Am Exot Anim Pract. 2007;10:61-78. 16. Raskin RW, Meyer DJ. Canine and feline cytology: A color atlas and interpretation. 2nd ed. St. Louis: Elsevier Saunders; 2009. 17. Scott DW, Miller WH, Griffin CE. Dermatosis of pet rodents, rabbits and ferrets. In: Scott DW, Miller WH, Griffin CE, eds. Muller and Kirk’s small animal dermatology. 6th ed. Philadelphia: WB Saunders; 2001:1415-1458. 18. Thrall MA. Veterinary hematology and clinical chemistry. Lippincott: Williams & Wilkins; 2004;19:214-221. 19. Zimmerman KL, Moore DM, Smith SA. Species specific hematology. In: Weiss DJ, Wardrop KJ, eds. Schalm’s veterinary hematology. 6th ed. Ames, Iowa: Wiley-Blackwell; 2010:852-917.
CHAPTER
37
Ophthalmologic Diseases in Small Pet Mammals
Alexandra van der Woerdt, DVM, MS, Diplomate ACVO, Diplomate ECVO
Rabbits Conjunctivitis and Epiphora Cornea Uveitis and Diseases of the Lens Glaucoma Orbit Ferrets Guinea Pigs Chinchillas Rats, Mice, and Hamsters Sugar Gliders
RABBITS Ophthalmic examination of rabbits can be performed easily.69,70 The eyes are laterally located and have a round pupil. Evaluation of a menace response is difficult, but most rabbits will react to bright light by squinting. The dorsal rectus muscle can usually be seen as a large striated band of tissue under the conjunctiva. Some rabbits do not respond to topical application of mydriatic agents because of the natural presence of atropinase. In these rabbits, the addition of 10% phenylephrine may help to obtain mydriasis. Rabbits have a merangiotic fundus. The well-myelinated optic nerve is present above the visual axis and has a deep optic cup. Retinal blood vessels are present in a linear streak medial and lateral to the optic nerve. An extensive venous plexus is present in the orbit. Tear production in rabbits can be measured by using Schirmer tear test strips. Average tear production is 5 mm/min (standard deviation, ±2.4 mm/min)1; however, very low values can be measured in some normal rabbits. Normal intraocular pressure measured by applanation tonometry is between 10 and 20 mm Hg. The nasolacrimal system of rabbits has a single nasolacrimal punctum. The punctum is located in the ventral eyelid 3 mm from the eyelid margin, near the medial canthus and ventral to the lacrimal caruncle (Fig. 37-1).9,38 The lacrimal sac is immediately rostral to the punctum and caudal to the nasolacrimal duct aperture. The nasolacrimal duct extends from the orbit to the nasal fossa and runs within the part of the maxilla that forms Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
the lateral wall of the maxillary sinus.38 Approximately 5 to 6 mm within the maxilla, the duct curves sharply and decreases in diameter.9 At the level of the palatine bone, the nasolacrimal duct leaves the bony nasolacrimal canal and makes a sharp turn at the nasolacrimal duct flexure, which is located just caudal to the caudal limit of the incisor tooth roots. The nasolacrimal duct narrows at this flexure in normal rabbits. The duct then follows the ventral margin of the nasoturbinates and exits on the ventromedial aspect of the alar fold just caudal to the mucocutaneous junction of the nares.
CONJUNCTIVITIS AND EPIPHORA Conjunctivitis in rabbits is common. In normal rabbits with no ocular or respiratory disease, the most frequently isolated organisms from the conjunctival cul de sac include Staphylococcus, Micrococcus, and Bacillus species. Less common organisms include Bordetella, Stomatococcus, Neisseria, Pasteurella, Corynebacterium, Streptococcus, and Moraxella species.14 However, Pasteurella multocida is a cause of conjunctivitis, epiphora, nasolacrimal duct obstruction, and dacryocystitis in rabbits.27,38 A wide variety of other infectious agents also have been associated with conjunctivitis in rabbits, including Staphylococcus aureus, Pseudomonas species, Haemophilus species, Treponema paraluiscuniculi, mycoplasmas, chlamydiae, and myxoma virus.27,45,60 In New Zealand white rabbits with conjunctivitis, upper respiratory disease, and pneumonia, bacterial isolates consisted of Bordetella broncheptica, P. multocida, S. aureus, and Pseudomonas alcaligenes.50 Mucopurulent conjunctivitis and blepharitis with corneal ulceration have been associated with S. aureus infection in a rabbit.45 Treatment with topical gentamicin ophthalmic ointment and systemic gentamicin was curative. Methicillin-resistant S. aureus was isolated in a rabbit with severe bilateral conjunctivitis. (K. Quesenberry, personal communication). Other causes of conjunctivitis in rabbits include foreign bodies, entropion, distichia, trichiasis, and high ammonia or dust content in the environment. Dental disease, including root elongation and dental abscesses, is also associated with conjunctivitis. Unilateral or bilateral epiphora can be present in rabbits without conjunctivitis. The discharge often has a white, gritty appearance and may be intermittent and resistant to treatment with topical antibiotics. Root elongation of the maxillary 523
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Fig. 37-1 Diagram of the rabbit nasolacrimal duct. A, Lateral view with inset. The two sharp bends, the proximal maxillary bend (pb), and the bend at the incisor tooth (ib), are indicated. The inset shows the canaliculus (C) and the lacrimal sac (S). B, Dorsoventral view. (1) Proximal portion of the duct extending from the punctum through the proximal maxillary curve; (2) portion of the duct extending from the proximal maxillary curve to the base of the incisor tooth; (3) portion of the duct extending from the base of the incisor tooth to the end of the lacrimal canal; (4) distal portion of the duct extending from the end of the lacrimal canal to the nasal meatus. C, The nasal meatus of the nasolacrimal duct (arrow). The opening is enlarged for diagrammatic purposes. incisors is a common underlying cause.27 The elongated roots can cause an obstruction of the nasolacrimal duct at its flexure just caudal to the roots of the incisors. Radiographs of the skull are needed to assess the incisors; excess curvature of the incisor roots is abnormal. In one report describing two affected rabbits and 13 normal rabbits, radiographs revealed a cystic dilation of the nasolacrimal duct immediately caudal to the duct flexure, and the incisors were more arched than in normal rabbits.38 Irrigation of the nasolacrimal duct in affected rabbits yielded opaque, white, gritty fluid which, on cytologic examination, showed numerous macrophages, lipid-laden mesothelial cells, lipid droplets, and small numbers of bacteria and erythrocytes. Occlusion of the ducts was presumed to be attributed to fat droplets. Bacteriologic culture of fluids used to irrigate the nasolacrimal ducts of both normal and affected rabbits yielded similar bacterial isolates; therefore microorganisms may not be important in the pathogenesis of epiphora in rabbits. The most common bacterial isolates were S. aureus, coagulase-negative
Staphylococcus species, Moraxella and Neisseria species, Oligella urethralis, and Streptococcus viridans. In a study of 28 cases of dacryocystitis in rabbits, 89% of cases were unilateral. The cause was determined to be dental malocclusion in 50%, no apparent cause in 35%, rhinitis in 7%, both rhinitis and dental malocclusion in 4%, and panophthalmitis in 4%. Most animals (98%) were treated with topical antibiotics, with a mean duration of 5.8 weeks.21 In rabbits with epiphora, the diagnostic value of bacterial culture of the irrigation fluid is questionable. However, bacterial culture is recommended if nasal discharge is present in conjunction with epiphora. Skull radiographs are useful to detect underlying dental disease. Dacryocystorhinography using contrast material injected into the nasolacrimal system can help localize the site of obstruction, differentiate between a complete and partial obstruction, and identify any dilation. Treatment of epiphora in rabbits can be frustrating. Irrigation of the nasolacrimal duct is important to restore the patency
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Fig. 37-3 Progressive occlusion of the cornea with conjunctivaFig. 37-2 Irrigation of the nasolacrimal duct in a rabbit with a 24-gauge Teflon intravenous catheter. Rabbits have a single nasolacrimal punctum in the ventral eyelid.
of the nasolacrimal system. After instilling a topical ophthalmic anesthetic, use a 23-gauge lacrimal cannula or a 22- to 24-gauge Teflon intravenous catheter to flush the duct (Fig. 37-2). Recurrence of the obstruction is common, and duct irrigation may need to be repeated every 2 to 3 days or weekly until a few consecutive clear irrigations are obtained. If topical antibiotic therapy is used, a broad-spectrum medication such as triple antibiotic solution is recommended. Topical nonsteroidal, antiinflammatory ophthalmic medications, such as 0.03% flurbiprofen or 0.1% diclofenac, may help minimize irritation caused by the procedure. In rabbits with chronic or severe infections, concurrent topical ophthalmic and systemic antibiotic therapy may be needed. Suggested combinations include systemic enrofloxacin (Baytril, Bayer Corporation, Shawnee Mission, KS) or marbofloxacin (Zeniquin, Pfizer Animal Health, Exton, PA) and topical ciprofloxacin or gentamicin ophthalmic solution. In rabbits with evidence of underlying incisor root elongation, removing the incisors can be considered in severe cases.
CORNEA Corneal dystrophy is the accumulation of cholesterol or lipid crystals in the cornea.4 This may develop spontaneously, as has been reported in American Dutch belted rabbits,47 or be due to high dietary cholesterol. It also occurs in breeds that are predisposed to hypercholesterolemia, such as the Watanabe rabbit with heritable hyperlipidemia.22,35 In Watanabe rabbits, yellowish-white granules can develop along the corneal-scleral junction and in the iris. In rabbits without systemic lipid abnormalities, spontaneous corneal dystrophy is usually bilateral and symmetric and does not progress to visual impairment. In any rabbit with corneal dystrophy, carefully evaluate the fat content of the diet. Progressive occlusion of the cornea with a conjunctiva-like membrane is occasionally seen in rabbits.3,18,57,59 Membranous corneal occlusion, or pseudopterygium, is a pain-free condition that may affect one or both eyes (Fig. 37-3). Ophthalmic examination reveals a circular membrane that originates at the limbus (the junction of the cornea and sclera) and gradually
like tissue. This tissue is not adherent to the cornea. The disease is not painful. (Courtesy David Wilkie, DVM, MS.)
advances over the cornea. In severe cases, only a small central opening is present, allowing visibility of an otherwise normal globe. The membrane does not adhere to the cornea. The cause of this condition is unknown, although trauma has been suggested. Progressive membranous occlusion in rabbits has been compared with pterygium in humans. However, in humans the membrane is triangular and adherent to the cornea, whereas in rabbits it is nonadherent and circumferential from the limbus. Treatment with topical antibiotic or antibiotic-steroid medications has no effect. The membrane can be resected surgically and treated with topical antibiotics postoperatively; this usually results in quick recurrence of the membrane. However, if the membrane is resected a few millimeters beyond the limbus and the eye is then treated with a topical antibiotic-steroid combination, recurrence may be prevented. Good results have also been obtained with surgical resection and the use of topical cyclosporine with or without corticosteroids.57 Another described surgical technique is to incise the membrane into four to six quadrants and suture each quadrant of the membrane to the inside of the eyelid.3,59 With this technique, recurrence may be prevented for at least 1 year. Superficial nonhealing corneal ulcers are occasionally seen in rabbits. Clinical signs are usually mild and include epiphora, conjunctival hyperemia, and blepharospasm. The ulcer is usually located in the paracentral cornea, is very superficial, and has redundant epithelial edges. The clinical appearance is that of an indolent ulcer, as seen in boxer dogs. Carefully examine the eyes of affected rabbits to eliminate potential causes such as abnormal hairs, lagophthalmos (inability to fully close the eyelid), facial nerve paralysis, or a foreign body. Treatment with a topical antibiotic solution or ointment usually fails to resolve the ulcer. Additional therapies such as corneal debridement, grid keratotomy, use of topical serum, application of corneal glue, tarsorrhaphy, or superficial keratectomy are usually necessary for the ulcer to heal.
UVEITIS AND DISEASES OF THE LENS Encephalitozoon cuniculi may cause granulomatous encephalitis and renal lesions in rabbits. Many rabbits infected with E. cuniculi are asymptomatic, but neurologic signs can include
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SECTION VI General Topics unrelated to the virus. Cataract and subsequent glaucoma can develop subsequent to ocular trauma, such as cat-scratch injury. Intraocular sarcomas have been described in two adult rabbits; one had a nonvisual eye associated with chronic inflammation and the second had chronic uveitis, cataract formation, and glaucoma.42 In both rabbits, intraocular spindle cell neoplasms closely associated with lens and lens capsular fragments were described. The histologic features of the tumors closely resembled posttraumatic ocular sarcomas in cats; chronic inflammation and trauma were considered as probable causes.
GLAUCOMA
Fig. 37-4 Rabbit infected with Encephalitozoon cuniculi. A white lesion is present in the iris, protruding into the anterior chamber. Lens involvement with cataract formation is present underneath the iridial lesion. convulsions, tremors, torticollis, paresis, and coma. Encephalitozoon cuniculi infection has also been associated with phacoclastic uveitis.20,23,63 Most affected rabbits are young (less than 2 years of age), and dwarf rabbits appear predisposed to this disease. Clinically, a white mass is often seen protruding into the anterior chamber (Fig. 37-4). Careful examination of the anterior segment of the eye with slit-lamp biomicroscopy may reveal a break in the anterior lens capsule. The break is frequently hidden by inflammatory material and it may appear as if only the iris is involved in the inflammatory process. A focal cataract is often present in the area of the anterior lens capsule break. Signs of a severe pyogranulomatous anterior uveitis are usually present, such as conjunctival hyperemia, a swollen hyperemic iris, miosis, aqueous flare, and low intraocular pressure. The posterior segment of the eye is initially normal; however, if left untreated, severe uveitis and cataract formation can lead to blindness and possible phthisis bulbi or glaucoma. An abscess in the iris caused by P. multocida initially may resemble phacoclastic uveitis. Measurement of serum antibody titers for E. cuniculi and P. multocida may aid in the differential diagnosis. The treatment of choice is surgical removal of the lens by phacofragmentation. Because of the rabbit’s ability to regenerate a lens after this procedure, insertion of an artificial lens after phacofragmentation is not recommended. Systemic treatment of E. cuniculi with albendazole (30 mg/kg PO q24h for 30 days, then 15 mg/kg PO q24h for an additional 30 days) has been reported.63 Fenbendazole (20 mg/kg q24h for 28 days) has proved effective in both preventing experimental E. cuniculi infection in rabbits and treating naturally infected seropositive rabbits.64 If the lens is not removed surgically, control of the uveitis with topical steroidal (such as 1% prednisolone acetate) and nonsteroidal anti-inflammatory medications as well as systemic fenbendazole or albendazole is necessary. Enucleation may be indicated if the uveitis cannot be controlled medically and a chronically painful eye is present.23,74 Spontaneous cataract formation is rare in rabbits. One report describes a low incidence of juvenile cataracts in laboratory New Zealand white rabbits.49 Keratitis caused by Shope fibroma virus has been reported in one rabbit.32 Cataracts developed 3 months after the keratitis, which may have been hereditary in origin and
Congenital glaucoma is inherited as an autosomal recessive trait in rabbits. In those with this condition, the intraocular pressure is high as early as 3 months of age.10 With increasing age, progressive buphthalmos with a markedly enlarged cornea, structural abnormalities of the iridocorneal angle, atrophy of the ciliary processes, and excavation of the optic nerve develop. Topical glaucoma medications used in dogs, such as 0.5% timolol maleate and 2% dorzolamide, may also be used in rabbits. Because response to therapy is unpredictable in rabbits, carefully monitor the intraocular pressure during treatment. Enucleation, insertion of an intrascleral prosthesis, and laser cycloablation with a diode laser have also been used to manage glaucoma in pigmented pet rabbits. However, laser cycloablation cannot be used in albino rabbits. If left untreated in chronic cases, pressure-induced atrophy of the ciliary body may lead the intraocular pressure to return to normal.
ORBIT Retrobulbar disease processes are occasionally seen in rabbits. Clinical signs include progressive exophthalmos, protrusion of the third eyelid, and inability to retropulse the globe. Exposure keratitis may be present if the ability of the eyelids to close properly has been affected (Fig. 37-5). Abscesses are the common cause of retrobulbar disease in rabbits; dental disease with tooth root abscessation is often a predisposing factor (see Chapter 32). Infection is caused by both aerobic and anaerobic bacterial species. A thorough dental examination and skull radiographs are indicated in any rabbit with a suspected retrobulbar mass (see Chapter 35). If available, a computed tomography (CT) scan is especially helpful in diagnosis Taenia serialis coenurus formation caused exophthalmos in a pet rabbit. Surgical removal was curative.51 Retrobulbar neoplasia is uncommon in rabbits. An abscess in the retrobulbar space of a rabbit can be very difficult to treat. Because of the thick nature of the abscessed material and the anatomy of the alveolar bulla, drainage of the abscess through the mouth, as performed in dogs and cats, may or may not be successful. If the abscess is caused by an abscessed tooth root, the tooth or teeth must be extracted to allow drainage and the rabbit treated with long-term systemic antibiotic therapy. In some cases aggressive surgical debridement may be necessary. This may include exenteration of the orbit and sacrifice of a sighted eye. Even with aggressive surgical and medical management, the prognosis for recovery is always guarded. Stomatoscopy-aided dental trimming, tooth removal, and debridement successfully treated a retrobulbar abscess in a rabbit.39 Anecdotal reports suggest that some rabbits with retrobulbar abscesses respond to medical therapy with long-term (3-month) administration of benzathine/procaine penicillin G
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B
A
Fig. 37-5 Rabbit with retrobulbar disease. Note the exophthalmos, hyperemia of the eyelids, discharge, and exposure keratitis.
(for rabbits < 2.5 kg, 75,000 U per rabbit SC q48h; for rabbits > 2.5 kg, 150,000 U per rabbit SC q48h).56 Bilateral exophthalmos in rabbits is commonly associated with compromised vascular drainage of the head. Periodic exophthalmos is common in rabbits with thymomas (see Chapter 20).34,53,68 In one report, a localized myasthenia gravis associated with the thymoma was suggested as a cause.68 More probably, the presence of a large intrathoracic mass causes cranial vena cava syndrome, in which the mass compresses vessels in the cranial thorax.53 This compression impedes blood flow in the right and left cranial vena cava as well as the external jugular veins and causes decreased vascular drainage from the head. Exophthalmos was also reported in a rabbit after the chronic placement of an external jugular catheter,29 and bilateral exophthalmos can develop with thrombosis of both jugular veins. The external jugular veins are the largest vessels draining the head, whereas the internal jugular veins of rabbits are relatively small and primarily drain the brain, throat, and neck regions. Therefore occlusion of the external jugular veins will compromise vascular drainage of the head. In clinical cases, exophthalmos secondary to venous thrombosis often resolves spontaneously, presumably after the jugular veins become patent again. Exophthalmos in obese rabbits may be caused by excessive fat deposition in the orbit. Rabbits have several glands in the orbit. The lacrimal gland is located dorsolaterally, and the accessory lacrimal gland, divided into three lobes, is located along the caudal and ventral orbital margin. The superficial gland of the third eyelid is a small gland located near the third eyelid’s cartilage. The deep gland of the third eyelid, also known as the harderian gland, consists of two parts: dorsal (white) and ventral (pink) lobes. Prolapse of the deep gland of the third eyelid has been described in rabbits. Surgical correction with a pocket technique, as has been described in dogs, was successful in reducing the gland in one rabbit.31
FERRETS Ferrets have prominent globes placed laterally in the skull; they have very limited binocular vision.24 The ferret’s pupil is a horizontal slit that quickly responds to light. Topical 1% tropicamide may have to be applied to evaluate the fundus. Like dogs and cats, ferrets have a holangiotic retinal vascular pattern. The projection of retinal ganglion cells from the temporal area of
the retina in albino ferrets differs from that of pigmented ferrets.48 In pigmented ferrets, 6,000 retinal ganglion cells project ipsilaterally to the brain, whereas in albino ferrets, only 1,500 retinal ganglion cells project ipsilaterally. The significance of this difference has not been established. Intraocular pressure can be measured with a tonopen (Tono-Pen XL, Reichart, Inc., Dewpew, NY). Normal values for intraocular pressure range from 14.5 ± 3.346 to 22.8 ± 5.5 mm Hg.58 Tear production in normal ferret eyes is less than in dogs, with an average of 5.3 ± 1.3 mm/min measured by commercially available strips.46 Conjunctivitis in ferrets can be caused by viral or bacterial infection. Ocular signs of canine distemper virus, a fatal disease in ferrets, are mucopurulent oculonasal discharge, blepharitis, corneal ulcers, and keratoconjunctivitis sicca.33 Conjunctival swelling and a proliferative lesion of the nictitans caused by infection with Mycobacterium genavense have been described in two ferrets. Other clinical signs in these ferrets included peripheral lymph node enlargement.37 Degeneration of corneal endothelial cells leading to progressive corneal edema and cloudiness of the cornea is seen in older mink (8-11 years of age).26 Royal pastel females are predisposed. Unlike the disease in dogs, these mink do not develop corneal ulceration, pigmentation, or vascularization. There is no specific treatment for this condition, but symptomatic treatment with 5% sodium chloride solution or ointment two to four times a day may or may not improve corneal clarity. A lymphoplasmacytic keratitis has been reported in a ferret with lymphoma.55 An infiltrative lesion resembling corneal lesions reported in mink with Aleutian disease was present in the cornea. Cataracts are common in ferrets.67 Progressive cataract formation has been reported in two genetically unrelated populations of ferrets.43 In 1-year-old ferrets, cataracts were observed in 47% of animals examined. Severity ranged from clinically insignificant, small cataracts in the posterior cortex of the lens to blinding, complete cataracts. By 18 months of age, cataracts were detected in virtually every animal, and in animals previously diagnosed, the cataracts had progressed. A genetically separate group had a combination of blinding cataract, microphthalmos, abnormal iris formation, and retinal detachment. In another ferret colony, microphthalmos, cataract, retinal dysplasia, and a persistent hyperplastic primary vitreoustype membrane were shown to be inherited as an autosomal
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dominant defect.17 Dietary factors may play a role in the development of cataracts in ferrets. A diet high in fat or deficient in vitamin E or protein may promote cataract formation.43 In ferrets with cataracts, monitor the eyes regularly for the onset of secondary complications. Lens-induced uveitis can usually be controlled with topical 1% prednisolone acetate applied once or twice daily. Other complications caused by cataracts are lens subluxation or luxation and glaucoma. Ferrets that are blind because of cataracts usually adjust well in a home environment. However, cataract surgery can be performed successfully in ferrets. The lenses can be removed by phacofragmentation or by an extracapsular technique. Artificial lenses are not available in a size suitable for ferrets. Before cataract surgery, make sure the ferret becomes accustomed to frequent application of eye medications to facilitate easy treatment after surgery. Retinal degeneration is seen in ferrets. Clinical signs are progressive loss of vision, which may not be noticed until the disease is advanced. Ophthalmic examination reveals mydriasis with a very poor pupillary light reflex. Cataracts may or may not be present. Retinal vascular attenuation and tapetal hyperreflectivity are seen in the fundus. There is no treatment for retinal degeneration. Exophthalmos can occur in ferrets from several causes. Lymphosarcoma is a common disease in ferrets, and although orbital involvement has been reported in only two ferrets,41 it is occasionally seen clinically. Exophthalmos is often the presenting complaint. Ophthalmic examination reveals unilateral or bilateral exophthalmos, decreased retropulsion of the globe, and protrusion of the third eyelid. Lagophthalmos may result in exposure keratitis with corneal ulceration and vascularization. In clinical cases, lymphosarcoma is usually present in other areas of the body. In the two ferrets described,41 peripheral lymph nodes were affected in one ferret and involvement of the liver, spleen, intestines, kidneys, and adrenals was present in the other. Retrobulbar adenocarcinoma has been reported in one ferret,40 and we have seen this clinically as a rare cause of unilateral exopthalmos (A. van der Woerdt, K. Quesenberry, personal communication, 2006). In ferrets with exophthalmos, a CT scan of the head is the best method of diagnosing a retrobulbar mass and determining the extent of the lesion. Cytologic examination of a sample from the retrobulbar area obtained by ultrasound-guided fine-needle aspiration can confirm the diagnosis of neoplasia. However, this procedure can be difficult because of the limited size of the retrobulbar space. Instead, in cases of suspected orbital lymphosarcoma, diagnosis may be confirmed by obtaining samples for diagnostic tests elsewhere in the body, such as a fine-needle aspirate or wedge biopsy of an enlarged lymph node. In ferrets with lymphosarcoma, therapy is directed at treating the disease systemically with prednisone or chemotherapeutic agents. While treating the ferret for lymphosarcoma, temporary tarsorrhaphy may be necessary to protect the cornea if pronounced exophthalmos is present. If the corneal epithelium is intact, protect the eye of a ferret with exophthalmos with lubricating ophthalmic ointment applied two to four times daily. If an ulcer is present, treat with an antibiotic ophthalmic ointment, such as triple antibiotic or gentamicin ointment, applied three to four times daily. In cases of retrobulbar adenocarcinoma, treatment may include exenteration and radiation therapy.40 Zygomatic salivary gland mucocele is another reported cause of exophthalmos in ferrets.44 Fine-needle aspiration of a soft fluctuant swelling dorsotemporal to the eye yields a tenacious, blood-tinged fluid. Surgical excision is usually curative (see Chapters 3 and 11).
Fig. 37-6 Conjunctivitis and keratitis in a guinea pig. Note the abundant mucopurulent discharge, corneal vascularization, and fibrosis.
GUINEA PIGS Guinea pigs have a paurangiotic retina that appears avascular on examination. Their eyelids are open from birth, and they have a rudimentary third eyelid. Guinea pigs produce a very small amount of tears, and measurement of tear production by using commercially available strips is not possible.65 The phenol red thread tear test is a better option to assess tear production in guinea pigs. Conjunctivitis is common in guinea pigs (Fig. 37-6). One common cause is Chlamydophila caviae (formerly C. psittaci),33 which causes a self-limited disease manifesting as mild chemosis, ocular discharge, and follicle formation. Cytologic examination of a specimen from a conjunctival scraping may reveal intracytoplasmic inclusion bodies in epithelial cells. Treatment is generally considered unnecessary. Vitamin C deficiency in guinea pigs causes conjunctivitis with a flaky discharge. Treatment is directed at correcting the dietary deficiency. A spontaneous outbreak of listerial keratoconjunctivitis has been reported in hairless guinea pigs.13 Clinical signs ranged from serous lacrimation with hyperemic conjunctiva to purulent, ulcerative keratoconjunctivitis with corneal neovascularization. Listeria monocytogenes was cultured from the ocular discharge. Treatment was not attempted. Blepharitis caused by dermatophyte infection may be seen in young guinea pigs.5 Topical antifungal therapy is usually effective. Lipid deposition within the conjunctiva of the eyelids, named “fatty eye” by guinea pig breeders is most commonly seen in overweight animals. There is no specific treatment for this condition. Lymphosarcoma is rare in guinea pigs but has been reported to infiltrate the cornea.62 Lymphosarcoma should also be considered as a differential diagnosis of conjunctival masses in guinea pigs.2 Another differential diagnosis for conjunctival nodules in guinea pigs is a syndrome known as “pea eye.” These nodules are protrusions of portions of the lacrimal or zygomatic glands and appear pale or pink. Treatment is not necessary because animals are usually not bothered by this condition. A dermoid is a congenital lesion in which skin-like tissue is present in an abnormal location. Corneal and conjunctival
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study. Average intraocular pressure measured with rebound tonometry was 2.9 ± 1.8 mm Hg.48a Cataracts and asteroid hyalosis have been reported in older animals.33 The premolar, molar, and incisor teeth in chinchillas, as in guinea pigs, continue to grow and erupt throughout the animal’s life. Insufficient wear can result in elongated roots of the premolar and molar teeth, resulting in progressive orbital disease including epiphora, decreased retropulsion of the globe, and proptosis (see Chapter 32).16 Computed tomography is more sensitive than radiography in detecting early lesions.15 The prognosis for advanced disease is poor. Exophthalmos secondary to an orbital Taenia coenurus has been reported. Surgical excision was curative.28
RATS, MICE, AND HAMSTERS
Fig. 37-7 Osseous metaplasia of the mesectodermal trabecular meshwork in a guinea pig. White opaque material is present in the iridocorneal angle. (From Brown C, Donnelly T: What’s your diagnosis? Heterotopic bone in the eyes of a guinea pig. Lab Anim 2002; 31:23-25.)
dermoids have been reported in guinea pigs.52,71 Irritation from abnormal hairs causes conjunctivitis, irritation, and possibly corneal ulceration. Surgical resection is curative. Cataracts have been reported in guinea pigs. Mature cataract can be associated with diabetes mellitus.73a They can be removed surgically, but the procedure is difficult because of the small size of the globe, the large size of the lens, and the difficulty of intubating a guinea pig for general anesthesia. Osseous metaplasia of the mesectodermal trabecular meshwork occurs in guinea pigs (Fig. 37-7).8,25 Clinically, an arc of white, opaque material is visible in the anterior chamber, covering the iridocorneal angle. Vessels may be present overlying the osseous choristoma. Hematopoietic active bone marrow is present. This is usually an incidental finding and no specific treatment is necessary. As in other rodents and rabbits, exophthalmos in guinea pigs may be related to dental disease. A tooth root abscess of a molar may result in maxillary sinusitis and orbital disease. Careful examination of the teeth is indicated in any guinea pig with exophthalmos.
CHINCHILLAS Chinchillas have fine ciliae on the upper and lower eyelids.35a The third eyelid is rudimentary. The cornea is large and highly curved. The pupil is a slit or vertical oval when constricted. The lens is very large, occupying approximately 50% of the axial diameter of the globe. Chinchillas have prominent choroidal vasculature and an anangiotic retina. The retina has no fovea and no cones and a choroidal tapetum lucidum is absent. The optic nerve can be myelinated or nonmyelinated. Tear production is low, with an average reported value of 1.07 mm/min. The phenol red thread tear test can be used to measure tear production. An average of 14.0 mm/15 seconds has been reported.48a A menace response and dazzle reflex is absent in most chinchillas with normal eyes. Gram positive bacteria are predominant in the conjuntival flora of normal chinchillas. The most frequent bacteria found are Streptococcus species and S. aureus. Average intraocular pressure measured by applanation tonometry was 18.5 ± 5.75 mm Hg in one study, and 17.71 ± 4.17 in another
The retinas of rats, mice, and hamsters are holangiotic, with arteries and venules radiating from the optic nerve like spokes on a wheel.73 Rats have three lacrimal glands: intraorbital, extraorbital, and harderian. Inbred strains of rats and mice are commonly used in commercial laboratories to study naturally occurring ophthalmologic diseases. Diseases involving all parts of the eye have been described. Common abnormalities include retinal degeneration, as in the Royal College of Surgeons (RCS) rat strain66; microphthalmos; and cataracts. In addition to specific genetically determined ocular abnormalities in inbred strains, other spontaneous abnormalities occur. Ophthalmic examination of 6,000 Sprague-Dawley rats revealed a focal linear retinopathy in 3% and a fundic coloboma in 0.5%.30 Spontaneous corneal degeneration has been described in Sprague-Dawley and Wistar rats,7 and corneal dystrophy has been described in Fischer 344 rats.36 Experimental infections also lead to ophthalmic abnormalities. Blepharitis with crust formation in the medial canthus and partial periocular alopecia were observed in mice experimentally infected with Trypanosoma brucei.61 Conjunctivitis in mice can be caused by numerous infectious agents, including Pseudomonas aeruginosa, P. pneumotropica, Salmonella species, Streptobacillus moniliformis, Corynebacterium kutscheri, Lancefield group C streptococci, Mycoplasma pulmonis, mousepox or ectromelia virus, Sendai virus, and lymphocytic choriomeningitis virus.6,33 Bacteriologic culture and sensitivity testing may be indicated in individual rats and mice with persistent conjunctivitis. Epiphora in rats and mice can be caused by dental problems. Nasolacrimal duct obstruction can result from overgrowth or malocclusion of the incisors. Chromodacryorrhea is red staining around the eyes seen in rats and mice. Inflammation of the harderian gland causes secretion of tears pigmented with porphyrin. Sialodacryoadenitis virus is a highly contagious coronavirus that replicates in the respiratory tract epithelium, causing rhinotracheitis, bronchitis, and alveolitis. The virus also causes sialoadenitis of the submandibular and parotid salivary glands and necrotizing dacryoadenitis of orbital and harderian lacrimal glands. Exophthalmos, epiphora, and keratoconjunctivitis may result. The infection usually resolves within 1 week in immunocompetent animals. In a study of athymic rats, infection persisted for more than 3 months, indicating that normal T-cell function is required for host defenses against the virus.72 Infection with sialodacryoadenitis virus may also result in uveitis and multifocal retinal degeneration.33 Complications from infection include corneal opacification, anterior and posterior synechiae, cataract, and glaucoma. Specific therapy is not available, and treatment is
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supportive only. Other causes for red tears include infection with parainfluenza virus type 3 or Sendai virus as well as pain or stress. Ammonia vapor from soiled bedding can act as an ocular irritant, predisposing animals to secondary infection. Keeping the housing areas well ventilated is important in preventing infection with sialodacryoadenitis virus. Of clinical significance is the effect of xylazine on the lens in rats and mice. A reversible cataract has been observed after systemic use of xylazine. Transcorneal water loss and altered aqueous humor composition caused by corneal exposure have been suggested as a pathogenesis of cataract formation.12 In hamsters, keratoconjunctivitis can result from ammonia vapor from soiled bedding. Dental problems including tooth root infection may result in facial or retrobulbar abscesses, with hemifacial swelling, proptosis, and exposure keratitis as common sequelae. Treatment with systemic antibiotics is often unrewarding, and such abscesses frequently lead to the animal’s death.33 Insidious globe enlargement with loss of vision has been reported in four hamsters.19 Ophthalmic examination of these hamsters revealed enlarged globes in both eyes, widely dilated pupils, lack of pupillary light reflex, a small optic nerve, and pale retinas. Histopathologic results suggested chronic openangle glaucoma. Treatment of the suspected glaucoma was not attempted.
SUGAR GLIDERS Sugar gliders have an avascular retina. Only a small residual tuft of fluorescein-impermeable vessels projects from the optic disk into the vitreous.11 Sugar gliders have prominent globes that are susceptible to trauma. Corneal ulcers may result from intraspecies fighting.54 A retrobulbar abscess can result from a bite wound to the face or a molar root abscess. Corneal lipid infiltration may form in juvenile sugar gliders when the mother is fed a diet that is too high in fat. Although not reported, cataract formation is seen clinically.
References 1. Abrams KL, Brooks DE, Funk RS, et al. Evaluation of the Schirmer tear test in clinically normal rabbits. Am J Vet Res. 1990;51:1912-1913. 2. Allgoewer I, Ewringmann A, Pfleghaar S. Lymphosarcoma with conjunctival manifestation in a guinea pig. Vet Ophthalmol. 1999;2:117-119. 3. Allgoewer I, Malho P, Schulze H, et al. Aberrant conjunctival stricture and overgrowth in the rabbit. Vet Ophthalmol. 2008;11:18-22. 4. Andrew SE. Corneal diseases of rabbits. Vet Clin North Am Exot Anim Pract. 2002;5:341-356. 5. Bauck L. Ophthalmic conditions in pet rabbits and rodents. Compend Contin Educ Pract Vet. 1989;11:258-266. 6. Beaumont SL. Ocular disorders of pet mice and rats. Vet Clin North Am Exot Anim Pract. 2002;5:311-324. 7. Bellhorn RW, Korte GE, Abrutyn D. Spontaneous corneal degeneration in the rat. Lab Anim Sci. 1988;38:42-50. 8. Brown C, Donnelly T. What’s your diagnosis? Heterotopic bone in the eyes of a guinea pig. Lab Anim. 2002;31:23-25. 9. Burling K, Murphy CJ, da Silva Curiel J, et al. Anatomy of the rabbit nasolacrimal duct and its clinical implications. Progr Vet Comp Ophthalmol. 1991;1:33-40.
10. Burrows AM, Smith TD, Atkinston CS, et al. Development of ocular hypertension in congenitally buphthalmic rabbits. Lab Anim Sci. 1995;45:443-444. 11. Buttery RG, Haight JR, Bell K. Vascular and avascular retinae in mammals. A funduscopic and fluorescein angiographic study. Brain Behav Evol. 1990;35:156-175. 12. Calderone L, Grimes P, Shalev M. Acute reversible cataract induced by xylazine and by ketamine-xylazine anesthesia in rats and mice. Exp Eye Res. 1986;42:331-337. 13. Colgin LMA, Nielsen RE, Tucker FS, et al. Case report of listerial keratoconjunctivitis in hairless guinea pigs. Lab Anim Sci. 1995;45:435-436. 14. Cooper SC, McLellan GJ, Rycroft AN. Conjunctival flora observed in 70 healthy domestic rabbits (Oryctolagus cuniculus). Vet Record. 2001;149:232-235. 15. Crossley DA, Jackson A, Yates J, et al. Use of computed tomography to investigate cheek tooth abnormalities in chinchillas (Chinchilla laniger). J Sm Anim Pract. 1998;39:385-389. 16. Crossley DA. Dental disease in chinchillas in the UK. J Small Anim Pract. 2001;42:12-19. 17. Dubielzig RR, Miller PE. The morphology of autosomal dominant microphthalmia in ferrets. Proceedings. Am Col Vet Ophthalmol. 1995;vol 2B:62. 18. Dupont C, Carrier M, Gauvin J. Bilateral precorneal membranous occlusion in a dwarf rabbit. J Sm Anim Exot Anim Med. 1995;3:41-44. 19. Ekesten B, Dubielzig RR. Spontaneous buphthalmos in the Djungarian hamster (Phodopus sungorus campbelli). Vet Ophthalmol. 1999;2:251-254. 20. Felche LM, Sigler RL. Phacoemulsification for the management of Encephalitozoon cuniculi-induced phacoclastic uveitis in a rabbit. Vet Ophthalmol. 2002;5:211-215. 21. Florin M, Rusanen E, Haessig M, et al. Clinical presentation, treatment, and outcome of dacryocystitis in rabbits: a retrospective study of 28 cases (2003-2007). Vet Ophthalmol. 2009;12:350-356. 22. Garibaldi BA, Goad ME. Lipid keratopathy in the Watanabe (WHHL) rabbit. Vet Pathol. 1988;25:173-174. 23. Giordano C, Weigt A, Vercelli A, et al. Immunohistochemical identification of Encephalitozoon cuniculi in phacoclastic uveitis in four rabbits. Vet Ophthalmol. 2005;8:271-275. 24. Good KL. Ocular disorders of pet ferrets. Vet Clin North Am Exot Anim Pract. 2002;5:325-339. 25. Griffith JW, Sassani JW, Bowman TA, et al. Osseous choristoma of the ciliary body in guinea pigs. Vet Pathol. 1988;25:100-102. 26. Hadlow WJ. Chronic corneal edema in aged ranch mink. Vet Pathol. 1987;24:323-329. 27. Harcourt-Brown F. Textbook of rabbit medicine. Oxford: Butterworth-Heinemann; 2002. 28. Holmberg BJ, Hollingsworth SR, Osofsky A, et al. Taenia coenurus in the orbit of a chinchilla. Vet Ophthalmol. 2007;10:53-59. 29. Hoyt Jr RF, Powell DA, Feldman SH. Exophthalmia in the rabbit after chronic external jugular catheter placement [Abstract P24]. Proceedings. Annu Meet Am Assoc Lab Anim Sci. 1994;8(4):A-19. 30. Hubert MF, Gillet JP, Durand-Cavagna G. Spontaneous retinal changes in Sprague Dawley rats. Lab Anim Sci. 1994;44:561-567. 31. Janssens G, Simoens P, Muylle S, et al. Bilateral prolapse of the deep gland of the third eyelid in a rabbit: diagnosis and treatment. Lab Anim Sci. 1999;49:105-109. 32. Keller RL, Hendrix DVH, Greenacre C. Shope fibroma virus keratitis and spontaneous cataracts in a domestic rabbit. Vet Ophthalmol. 2007;10:190-195. 33. Kern TJ. Ocular disorders of rabbits, rodents, and ferrets. In: Kirk RW, Bonagura JD, eds. Current veterinary therapy X. Philadelphia: WB Saunders; 1989:681-685.
CHAPTER 37 Ophthalmologic Diseases in Small Pet Mammals 34. Kostolich M, Panciera RJ. Thymoma in a domestic rabbit. Cornell Vet. 1992;82:125-129. 35. Kouchi M, Uead Y, Horie H, et al. Ocular lesions in watanabe heritable hyperlipidemic rabbits. Vet Ophthalmol. 2006;9(3):145-148. 35a. Lima L, Montiani-ferreira F, Tramontin M, et al. The chinchilla eye: morphologic observations, echobiometric findings and reference values for selected ophthalmic diagnostic tests. Vet Ophthalmol. 2010;13:14-25. 36. Losco PE, Troup CM. Corneal dystrophy in Fischer 344 rats. Lab Anim Sci. 1988;38:702-710. 37. Lucas J, Lucas A, Furber H, et al. Mycobacterium genavense infection in two aged ferrets with conjunctival lesions. Aust Vet J. 2000;78:685-689. 38. Marini RP, Foltz CJ, Kersten D, et al. Microbiologic, radiographic, and anatomic study of the nasolacrimal duct apparatus in the rabbit (Oryctolagus cuniculus). Lab Anim Sci. 1996;46:656-662. 39. Martinez-Jiménez D, Hernández-Divers SJ, Dietrich UM, et al. Endosurgical treatment of a retrobulbar abscess in a rabbit. J Am Vet Med Assoc. 2007;230:868-872. 40. McBride M, Mosunic CB, Barron GH, et al. Successful treatment of a retrobulbar adenocarcinoma in a ferret (Mustela putorius furo). Vet Rec. 2009;165:206-208. 41. McCalla TL, Erdman SE, Kawasaki TA, et al. Lymphoma with orbital involvement in two ferrets. Vet Comp Ophthalmol. 1997;7:36-38. 42. McPherson L, Newman SJ, McLean N, et al. Introcular sarcomas in two rabbits. J Vet Diagn Invest. 2009;21:547-551. 43. Miller PE, Marlar AB, Dubielzig RR. Cataracts in a laboratory colony of ferrets. Lab Anim Sci. 1993;43:562-568. 44. Miller PE, Pickett JP. Zygomatic salivary gland mucocele in a ferret. J Am Vet Med Assoc. 1989;194:1437-1438. 45. Millichamp NJ, Collins BR. Blepharoconjunctivitis associated with Staphylococcus aureus in a rabbit. J Am Vet Med Assoc. 1986;189:1153-1154. 46. Montiani-Ferreira F, Mattos BC, Russ HH. Reference values for selected ophthalmic diagnostic tests of the ferret (Mustela putorius furo). Vet Ophthalmol. 2006;9:209-213. 47. Moore CP, Dubielzig R, Glaza SM. Anterior corneal dystrophy of American Dutch belted rabbits: biomicroscopic and histopathologic findings. Vet Pathol. 1987;24:28-33. 48. Morgan JE, Henderson Z, Thompson ID. Retinal decussation patterns in pigmented and albino ferrets. Neuroscience. 1987;20:519-535. 48a. Müller K, Mauler DA, Eule JC. Reference values for selected ophthalmic diagnostic tests and clinical characteristics of chinchilla eyes (Chinchilla lanigera). Vet Ophthalmol. 2010;13:29-34. 49. Munger RJ, Langevin N, Podval J. Spontaneous cataracts in laboratory rabbits. Vet Ophthalmol. 2002;5:177-181. 50. Okewale EA, Olubunmi PA. Antibiograms of pathogenic bacteria isolated from laboratory rabbits in Ibadan, Nigeria. Lab Anim. 2008;42:511-514. 51. O’Reilly A, McCowan C, Hardman C, et al. Taenia serialis causing exophthalmos in a pet rabbit. Vet Ophthalmol. 2002;5:227-230. 52. Otto G, Lipman NS, Murphy JC. Corneal dermoid in a hairless guinea pig. Lab Anim Sci. 1991;41:171-172. 53. Pignon C, Jardel N. Bilateral exophthalmos in a rabbit. Lab Animal. 2010;39:262-265. 54. Pye GW, Carpenter JW. A guide to medicine and surgery in sugar gliders. Vet Med. 1999;94:891-905.
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55. Ringle MJ, Lindley DM, Krohne SG. Lymphoplasmacytic keratitis in a ferret with lymphoma. J Am Vet Med Assoc. 1993;203:670-672. 56. Rosenfield ME. Successful eradication of severe abscesses in rabbits with long-term administration of penicillin G benzathine/penicillin G procaine. Available at http://moorelab. sbs.umass.edu/~mrosenfield/bicillin/. Accessed May 17, 2002. 57. Roze M, Ridings B, Lagadic M. Comparative morphology of epicorneal conjunctival membranes in rabbits and human pterygium. Vet Ophthalmol. 2001;4:171-174. 58. Sapienza JS, Porcher D, Collins BR, et al. Tonometry in clinically normal ferrets (Mustela putorius furo). Progr Vet Comp Ophthalmol. 1991;1:291-294. 59. Schoofs S, Hanssen P. Epicorneal conjunctiva syndroom bij het konijn: een klinisch geval en chirurgische behandeling ervan. Vlaams Dierg Tijdschr. 1998;67:344-346. 60. Srivastava KK, Pick JR, Johnson PT. Characterization of a Haemophilus sp. Isolated from a rabbit with conjunctivitis. Lab An Sci. 1986;36(3):291-293. 61. Shapiro SZ, Thulin JD, Morton DG. Periocular and urogenital lesions in mice (Mus musculus) chronically infected with Trypanosoma brucei. Lab Anim Sci. 1994;44:76-78. 62. Steinberg H. Disseminated T-cell lymphoma in a guinea pig with bilateral ocular involvement. J Vet Diagn Invest. 2000;12:459-462. 63. Stiles J, Didier E, Ritchie B, et al. Encephalitozoon cuniculi in the lens of a rabbit with phacoclastic uveitis: confirmation and treatment. Vet Comp Ophthalmol. 1997;7:233-238. 64. Suter C, Müller-Doblies UU, Hatt J-M, et al. Prevention and treatment of Encephalitozoon cuniculi infection in rabbits with fenbendazole. Vet Rec. 2001;148:478-480. 65. Trost K, Skalicky M, Nell B. Schirmer tear test, phenol red thread tear test, eye blink frequency and corneal sensitivity in the guinea pig. Vet Ophthalmol. 2007;10:141-146. 66. Tso MOM, Zhang C, Abler AS, et al. Apoptosis leads to photoreceptor degeneration in inherited retinal dystrophy of RCS rats. Invest Ophthalmol Vis Sci. 1994;35:2693-2699. 67. Utroska B, Austin WL. Bilateral cataracts in a ferret. Vet Med/ Sm Anim Clin. 1979:1176-1177. 68. Vernau KM, Grahn BH, Clarke-Scott HA, et al. Thymoma in a geriatric rabbit with hypercalcemia and periodic exophthalmos. J Am Vet Med Assoc. 1995;206:820-822. 69. Wagner F, Heider HJ, Gorig C, et al. Augenkrankheiten beim Zwergkaninchen. Teil 1: Anatomie, untersuchungsgang, erkrankungen der Augenlider, der Konjunktiva und des Tranennasengangs. Tierarztl Prax. 1998;26:205-210. 70. Wagner F, Heider HJ, Gorig C, et al. Augenkrankheiten beim Zwergkaninchen. Teil 2: Erkrankungen der kornea, intraokulare und retrobulbare erkrankungen sowie neoplasien. Tierartztl Prax. 1998;26:345-350. 71. Wappler O, Allgoewer I, Schaeffer EH. Conjunctival dermoid in two guinea pigs: a case report. Vet Ophthalmol. 2002;5:245-248. 72. Weir EC, Jacoby RO, Paturzo FX, et al. Persistence of sialodacryoadenitis virus in athymic rats. Lab Anim Sci. 1990;40:138-143. 73. Williams DL. Ocular disease in rats: a review. Vet Ophthalmol. 2002;5:183-191. 73a. Williams D, Sullivan A. Ocular disease in the guinea pig (Cavia porcellus): a survey of 1000 animals. Vet Ophthalmol. 2010;13:54-62. 74. Wolfer J, Grahn B, Wilcock B, et al. Phacoclastic uveitis in the rabbit. Progr Vet Comp Ophthalmol. 1993;3:92-97.
CHAPTER
38
Emergency and Critical Care of Small Mammals
Marla Lichtenberger, DVM, Diplomate ACVECC, and Angela M. Lennox, DVM, Diplomate ABVP (Avian)
Identification and Triage of the Critically Ill Patient Cardiopulmonary-Cerebral Resuscitation Principles CPCR in Small Mammals: Respiratory Arrest CPCR in Small Mammals: Cardiac Arrest Determining the Effectiveness of CPCR Anesthesia-Related Arrest Shock and Fluid Therapy Three Phases of Hypovolemic Shock Types of Fluids Fluid Resuscitation of the Critically Ill Small Mammal Blood Transfusion Use of Glucocorticoids in Shock Routes of Fluid Administration Maintenance of Normothermia Indirect Measurement of Systolic Blood Pressure Critical Care Clinical Pathology Lactate Monitoring Use of Prothrombin and Partial Thromboplastin Times Nutritional Support Sedation and Anesthesia of the Critically Ill Small Mammal Treatment of Selected Common Emergencies Acute Renal Failure Urinary Obstruction Respiratory Distress Anorexia and Gastric Stasis
Treatment of critically ill exotic companion mammals is complicated by their small size, physiologic diversity, and the lack of research and clinical data on response to therapy. Despite these impediments, the same principles and techniques used in domestic animals can be applied to the small mammal. Keep in 532
mind that single-product fluid regimens and fixed administration volumes based on simple milliliter-per-kilogram formulas are in most instances outdated, inappropriate, and often inadequate. Appropriate fluid therapy based on rational product choice, combined with frequent patient evaluation and periodic blood pressure monitoring, can greatly improve patient response to therapy and survival rates.
IDENTIFICATION AND TRIAGE OF THE CRITICALLY ILL PATIENT Most exotic companion mammals do not show symptoms of illness in the early stages of disease. Thus patients with chronic disease often present as emergencies because of an innate ability to mask clinical signs of the disease until the condition is severe. For these reasons owners suspecting illness in their small mammal pets should be advised to present them as soon as possible for examination. Identify critical patients as soon as they come into the veterinary hospital. Admit to the hospital immediately any patient demonstrating extreme respiratory difficulty or open-mouthed breathing, collapse, marked weakness, seizure, or uncontrolled bleeding. In many cases, complete physical examination is delayed in favor of emergency stabilization.
CARDIOPULMONARY-CEREBRAL RESUSCITATION PRINCIPLES The ABCs (airway, breathing, and circulation) of human emergency medicine are universal and apply as well to the exotic companion mammal patient.21 The general goal of cardiopulmonary resuscitation is restoration of spontaneous circulation. In the 2000s, the American Heart Association changed the guidelines to include preservation of neurologic function as a goal of successful resuscitation. Therefore the term cardiopulmonarycerebral resuscitation (CPCR) was adopted.2,33 Guidelines have been reviewed and modified for veterinary patients.8-10 Basic life support consists of the ABC approach, while advanced life Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
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Box 38-1 CPCR Protocol for Use in Small Mammals No respirations (respiratory No respirations (cardiac arrest), but pulse present arrest), and no pulse and and heart rate present no heartbeat
Fig. 38-1 Positive-pressure ventilation in a rabbit using a tightfitting anesthetic mask. This form of ventilation may be effective in cases of respiratory arrest when intubation is not possible.
support consists of electrocardiographic (ECG) identification of the abnormal rhythm, defibrillation, fluid and drug administration, and postresuscitative care.2 To facilitate resuscitation, a crash cart or specific crash area should be established and readily available with necessary supplies to maximize the chances of a successful outcome. The cart should also contain a list that documents the supplies it should contain, thus facilitating restocking.
CPCR IN SMALL MAMMALS: RESPIRATORY ARREST In the event of respiratory arrest, the ideal treatment is immediate endotracheal intubation. Intubation can be challenging in exotic companion mammals, as many are obligate or dependent nasal breathers with limited oral airway access.20 Intubation of the ferret and other exotic carnivores is straightforward, using direct visualization, similar to the technique used in the domestic cat. Rabbits may be intubated using the blind orotracheal or nasotracheal technique. An alternative for rabbits is otoscope- or endoscope-guided orotracheal intubation, using the endotracheal tube over the endoscope (over-the-top method) or side by side.20 Larger guinea pigs can also be intubated using this method.6,17 It must be emphasized that all intubation techniques require practice, and proficiency should be gained prior to presentation of a patient in respiratory arrest. In reality, most smaller mammals and rodents are difficult to intubate without significant practice.5,20,38,40 Therefore the following options should be considered: • Forced high-flow oxygen ventilation using a tight-fitting mask over the nose and mouth. Positive-pressure ventilation should be provided using 100% oxygen at a rate of 20 to 30 breaths per minute (Fig. 38-1). A disadvantage of this technique is accumulation of gastric air and bloat, which can limit movement of the diaphragm. • Tracheostomy. This procedure is similar to that described in dogs and cats.7 Make a 2- to 3-cm skin incision on the ventral midline parallel to the trachea, just proximal to the larynx. Bluntly dissect the subcutaneous fat and fascia, taking care to avoid blood vessels within the fat. Blunt
Turn off anesthesia if applicable. Reverse narcotics or analgesics if applicable.a Establish airway if possible or apply tight fitting mask and ventilate with 100% oxygen at 10-12 breaths per minute and 10 mm Hg airway pressure. Administer doxapram at 1-2 mg/kg IM, IV, IO. If bradycardic, administer atropine at 0.02 mg/kg IV, IO (or glycopyrrolate in rabbits/rats at 0.01 mg/kg). If successful, check blood pressure and correct fluid deficits. Check temperature and correct if necessary.
Begin diagnostic workup. Treat underlying disorders.
Turn off anesthesia if applicable. Reverse narcotics or analgesics if applicable.a Establish airway if possible or use tight-fitting mask and ventilate with 100% oxygen at 10-12 breaths per minute and 10 mm Hg airway pressure. Administer doxapram at 1-2 mg/kg IV, IO, IM. Begin chest compressions 100-120 per minute. Use vasopressin 0.8 U/kg IV, IO; double the dose if used via endotracheal tube. If no response in 1 minute, consider epinephrine 0.01 mg/kg IV, IO, double the dose if used via endotracheal tube. If ventricular fibrillation: defibrillate at 5-10 joules/ kg x3. If unsuccessful, consider open-chest CPR. If successful, check blood pressure and correct fluid deficits. Check temperature and correct if necessary. Begin diagnostic workup. Treat underlying disorders.
aAtipamezole for medetomidine. Butorphanol or buprenorphine for mu-receptor analgesics (i.e., hydromorphone, morphine, fentanyl); these drugs are preferred to naloxone as they reverse mu-receptor respiratory and central depression, leaving kappa receptor analgesia effects intact; naloxone reverses both mu and kappa receptor and is very short acting.
dissection is continued through the sternohyoid and sternothyroid muscles to isolate the trachea. Make a transverse incision between the tracheal rings, which should not exceed 50% of the circumference of the trachea. Place stay sutures in the trachea cranial and caudal to the tracheostomy site. Insert an endotracheal tube or cannula into the trachea and secure it in place.
CPCR IN SMALL MAMMALS: CARDIAC ARREST Cardiac arrest is cessation of effective circulation and is recognized by loss of consciousness and collapse. A palpable pulse is absent, the mucous membranes are pale or cyanotic, and respirations commonly cease (i.e., cardiopulmonary arrest).2 Box 38-1
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Table 38-1 CPCR Drug Dosages Suggested for Use in Exotic Companion Mammals Drug (Concentration)
Dose
mL/50 g
mL/100 g
mL/kg
mL/2 kg
Epinephrine (1:1,000) (1 mg/mL) Atropinea (0.54 mg/mL) Glycopyrrolate (0.2 mg/mL) 50% Dextrose (diluted 50% w/saline) Calcium gluconate (100 mg/mL) Doxapram (20 mg/mL) Vasopressin (20 U/mL) External defibrillation Butorphanolb (10 mg/mL) Buprenorphineb (0.3 mg/mL) Naloxoneb (0.4 mg/mL)
0.01 mg/kg 0.02 mg/kg 0.02 mg/kg 0.25 mL/kg 50 mg/kg 2.0 mg/kg 0.8 U/kg 2-10 J/kg 0.2-0.4 mg/kg 0.04 mg/kg 0.02 mg/kg IV 0.04 mg/kg IM Same volume as medetomidine; IM only
0.0005 0.002 0.005 0.05 0.025 0.005 0.002 N/A 0.001 0.006 0.0025 0.005 —
0.001 0.004 0.01 0.1 0.05 0.01 0.004 1 0.002-0.004 0.013 0.005 0.01 —
0.01 0.037 0.1 1.0 0.5 0.1 0.04 2 0.02-0.04 0.13 0.05 0.1 —
0.02 0.074 0.2 2.0 1.0 0.2 0.08 4 0.04-0.08 0.26 0.1 0.2 —
Atipamezole (reversal of medetomidine)
aAtropine (onset of action 15-30 seconds) is not recommended in rabbits, as many possess serum atropinesterase and the dose is unpredictable. Increasing the dose of atropine increases the risk of severe tachycardia as well as the risk of ventricular arrhythmias. Use glycopyrrolate (onset of action 30-45 seconds) in rabbits. bSee comments on anesthetic/analgesic reversal in Box 38-1.
presents a CPCR flow chart for use in exotic companion mammals. Intubate the patient and ventilate with 100% oxygen or, alternatively, deliver forced high-flow oxygen as directed above. Chest compressions at the rate of 100 to 120 times per minute directly compress the myocardium, resulting in increased cardiac output. It is important that both hands be placed on both sides of the chest with compressions done at the widest portion of the chest. The duration of the compression should be half of the total compression-release cycle.2 The team should continually assess their efforts at CPCR. Check to see if the efforts are generating a palpable pulse. If no pulse is felt, increase the force of chest compressions and assess the ECG. Different cardiac arrhythmias may require specific treatments. Consider intravenous or intraosseous access at this time (see individual species chapters for more information on catheterization techniques). Epinephrine has routinely been the vasopressor of choice for ventricular fibrillation, asystole, and pulseless electrical activity (PEA), the condition of cardiac arrest with no mechanical activity of the heart but normal/slow electrical activity and ECG findings. However, epinephrine and other catecholamines lose much of their effectiveness as vasopressors when the body is in a state of hypoxia and acidosis. Vasopressin may be more effective under these conditions, and may improve rates of restoration of spontaneous circulation and survival.2,24 Vasopressin should be given first, followed by epinephrine. Both drugs may be given intravenously, intraosseously, or via the endotracheal tube. Dosages for CPCR drugs are given in Table 38-1.
gas measurement, can provide a more accurate assessment of organ perfusion.2 End-tidal CO2 should be measured during CPR, as a steady rise is more likely to be associated with a successful outcome. When end-tidal CO2 does not rise above 10 mm Hg after a resuscitation time of 15 to 20 minutes, the resuscitative effort is unlikely to be successful. End-tidal CO2 measurements are practical only in the intubated patient weighing more than 50 g.1,12 The common practice of monitoring arterial blood gases during CPR should be abandoned in favor of monitoring venous blood gases, as the latter represents the oxygenation and acidbase status of peripheral tissues.2 In traditional pet species, failure to regain consciousness in the first few hours after CPR does not necessarily indicate prolonged or permanent neurologic impairment. However, coma longer than 4 hours after CPR carries a poor prognosis for full neurologic recovery.2,24 Brainstem reflexes, most importantly the pupillary light reflex (PLR), can have prognostic value in patients that do not regain consciousness after CPCR. Absence of the pupillary light reflex after one or more days of coma indicates little to no chance of neurologic recovery. This reflex has no prognostic value in the first 6 hours after CPCR because it can be transiently lost and then reappear. It should be noted that atropine and epinephrine can produce pupillary dilation, but these do not interfere with the pupillary response to light.24
DETERMINING THE EFFECTIVENESS OF CPCR
Anesthesia-related arrests represent one of the more treatable causes of arrest in veterinary patients. In the authors’ experience, exotic companion mammals under inhalant anesthesia often become bradycardic just prior to respiratory and cardiac arrest. Therefore use of an ultrasonic Doppler with concurrent measurement of blood pressure with or without ECG monitoring should be used during any anesthetic procedure, with measurements taken every 5 minutes.
The following parameters found useful in humans and traditional pet species may be useful in small mammals as well. The presence of palpable pulses is not an indication of adequate blood flow. Although palpable pulses may evaluate response to resuscitation, they do not indicate adequacy of organ perfusion. Two other measurements, end-tidal CO2 and blood
ANESTHESIA-RELATED ARREST
CHAPTER 38 Emergency and Critical Care of Small Mammals In the event of respiratory arrest, discontinue any inhalant anesthesia, intubate the patient if this has not already been done, and initiate positive-pressure ventilation with 100% oxygen at a rate of 20 to 26 breaths per minute. If intubation is not possible, begin mask ventilation as described above. If other anesthetic/analgesic agents were given prior to anesthesia, administer reversal agents if applicable25 (see Table 38-1, Box 38-1). Doxapram is often useful as a respiratory stimulant.41 For cases of bradycardia, administer atropine intravenously, intraosseously, or via the endotracheal tube. Glycopyrrolate may be indicated in rabbits and rodents because of the presence of serum atropinase.29 Drug dosages are given in Table 38-1.
SHOCK AND FLUID THERAPY Shock is defined as poor tissue perfusion from either low blood flow or unevenly distributed flow, resulting in an inadequate delivery of oxygen to the tissues.3 Hypovolemic shock is caused by absolute or relative inadequate blood volume. Absolute hypovolemia occurs with actual loss of blood—for example, arterial bleeding, gastrointestinal ulcers, or coagulopathies.3 With relative hypovolemia, there is no direct blood loss (hemorrhage) from the intravascular space. Examples include severe dehydration from gastrointestinal tract loss, significant loss of plasma (burns), or extensive loss of intravascular fluids into a body space such as the peritoneal cavity. In any case, there is decreased blood volume and venous return to the right side of the heart. This causes a reduction in return to the left side of the heart and consequently cardiac output.3 Studies in rabbits and rats show baroreceptor response to hypovolemia begins at 30% loss of blood volume. In other pet species, 30% loss also causes a decrease in blood pressure to below 60 mm Hg (diastolic) or less than 90 mm Hg (systolic). Carotid and aortic artery baroreceptors detect a decrease in stretch due to the decrease in cardiac output. This sends a neural signal to the vasomotor center in the medulla oblongata, resulting in inhibition of the vagal parasympathetic center and stimulation of the sympathetic center. The result is vasoconstriction of the veins and arterioles throughout the peripheral circulatory system and increases heart rate and strength of heart contraction. The humoral response to shock results in increased circulating catecholamines, which stimulates rennin release via adrenergic receptors on cells of the juxtaglomerular apparatus (specialized smooth muscle cells in the afferent arterioles). The release of renin stimulates activation of the renin-angiotensin-aldosterone system. These combined effects lead to a restoration of blood pressure, increased cardiac performance, and maximal venous return in the face of blood loss. Continued loss of blood volume results in hypovolemic shock and hypotension. Fluid therapy is required to optimize patient outcome.3
THREE PHASES OF HYPOVOLEMIC SHOCK Early or Compensatory Phase The early or compensatory stage of shock occurs because of the baroreceptor-mediated release of catecholamines.3,34 Blood pressure increases because of the increase in cardiac output and systemic vascular resistance. This is the stage seen commonly in dogs with blood loss less than 20% of total blood volume. In the authors’ experience, exotic companion mammals rarely present in this stage of shock. Clinical signs in dogs include increased heart rate, normal or increased blood pressure, and normal or increased flow (bounding pulses and capillary refill
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in less than 1 second). The increased heart rate and normal or increased blood pressure is the key indicator of compensatory shock. Volume replacement at this stage is usually associated with a good outcome.
Early Decompensatory Phase The second stage of shock occurs when fluid losses continue. There is a reduction in the blood flow to the kidneys, gastrointestinal tract, skin, and muscles. There is an uneven distribution of blood flow.3 This appears to be the most commonly encountered phase of shock in exotic companion mammals. Clinical signs include hypothermia, cool limbs and skin, tachycardia, normal or decreased blood pressure, pale mucous membranes, prolonged capillary refill time (CRT), and mental depression.3 Aggressive fluid therapy using crystalloids and colloids to support blood pressure and heart rate is required at this stage.
Decompensatory Phase When a large volume of blood is lost, the neuroendocrine responses to hypovolemia become ineffective and irreversible organ failure begins. This is the final common pathway of all forms of shock in all species.3 Clinical signs are bradycardia with low cardiac output, severe hypotension, pale or cyanotic mucous membranes, absent capillary refill time, weak or absent pulses, hypothermia, oliguric to anuric renal failure, pulmonary edema, and a stuporous to comatose state. Cardiopulmonary arrest commonly occurs at this stage.3
TYPES OF FLUIDS Fluid choices include crystalloids and colloids. Individual characteristics of fluids influence the type and volume of fluid administered.14 Crystalloids include lactated Ringer’s solution, normal saline, and hypertonic saline (7.2%-7.5%). Hypertonic saline draws fluid into the intravascular space from all body compartments rapidly; therefore this can be extremely useful in selected cases.39 Natural colloids are blood, plasma, or albumin. Synthetic colloids include hetastarch (HES) (Hespan, Jorgensen Labs, Loveland, CO) and Oxyglobin (OPK Biotech, Cambridge, MA). Oxyglobin has the added advantage of carrying oxygen on the hemoglobin molecule to all the small vessels. Isotonic crystalloid solutions are commonly used together with colloids in the resuscitation phase.14 Warm all fluids to the body temperature of the patient regardless of the route of administration (see “Maintenance of Normothermia,” below). Fluids can be warmed to 100°F to 103°F (38°C-39°C) without affecting their composition.14 Dextrose solutions may be added to crystalloid solutions for the treatment of hypoglycemia confirmed via blood glucose measurement. Give an initial bolus of 50% dextrose at 0.25 mL/kg as a 1:1 dilution with isotonic saline intravenously; determine blood glucose 1 hour later. Administer additional dextrose as 1.25% with crystalloids, with frequent rechecks of blood glucose. Administer dextrose with care, as it may induce compartmental shifts in electrolytes and water, resulting in worsening hypovolemia.14
FLUID RESUSCITATION OF THE CRITICALLY ILL SMALL MAMMAL Fluid therapy is used to correct life-threatening abnormalities in volume, electrolyte, and acid-base status. The goals of fluid therapy include resuscitation (correction of perfusion deficits), rehydration (correction of interstitial deficits), and maintenance.14
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It is important to administer the least amount of fluids necessary to reach the desired endpoints of resuscitation. Clinical markers important to determine response to therapy and include restoration of normal mentation, mucous membrane color, CRT, and establishment of normothermia, normovolemia, and normal heart rate and urine output.14 Resuscitation implies an urgent need to restore tissue perfusion and oxygenation. The type, quantity, and rate of fluid administration required to reach the desired resuscitation endpoints are determined based on the phase of shock. These commonly include crystalloids with the addition of colloids.14 As mentioned earlier, exotic companion mammals typically present in the decompensatory phases of shock. Earlier recommendations for shock therapy included crystalloids administered quickly in volumes equivalent to the patient’s blood volume. However, it should be kept in mind that resuscitation with crystalloids alone can result in a significant accumulation of pulmonary and pleural fluid. The resultant hypoxemia contributes to shock pathophysiology.14 The authors have found success with the following procedures for the treatment of hypovolemic shock in various exotic companion mammals (Box 38-2): administer a bolus infusion of 7.2% to 7.5% hypertonic saline (3 mL/kg IV or IO as a slow bolus over 10 minutes). The rapid effects of hypertonic saline are maintained with the addition of the colloid HES, which is given at 3 mL/kg IV or IO over 5 to 10 minutes. Indirect systolic blood pressure should be checked frequently (see “Measurement of Indirect Systolic Blood Pressure,” below). Once it is above 40 mm Hg systolic and/or there is improvement in other clinical markers of shock, administer maintenance isotonic crystalloids while aggressively rewarming the patient (see “Maintenance of Normothermia,” below). In many species, once rectal temperature approaches 98°F (36.7°C), adrenergic receptors begin to respond to catecholamines and fluid therapy.32 Temperatures during this rewarming phase must be checked frequently in all patients to prevent inadvertent hyperthermia. Blood pressure is rechecked when the temperature is greater than 98°F (36.7°C). Isotonic crystalloids (10 mL/kg) with HES in increments of 3 to 5 mL/kg can be repeated over 15 minutes until the systolic blood pressure rises above 90 mm Hg and/or there is again improvement in clinical markers. When the systolic blood pressure is above 90 mm Hg, the rehydration phase of fluid resuscitation begins. If the patient is known to be hypoproteinemic, administer a constant-rate infusion (CRI) of HES at 0.8 mL/kg per hour during rehydration, which will help maintain oncotic pressures in the intravascular space. If improvement is not seen after three to four boluses of HES and crystalloids, as outlined above, the patient is evaluated and treated for causes of nonresponsive shock (i.e., excessive vasodilation or vasoconstriction, hypoglycemia, electrolyte imbalances, acid-base disorder, cardiac dysfunction, hypoxemia). If cardiac function is normal and glucose, acid-base, and electrolyte abnormalities have been corrected, continue treatment for nonresponsive shock. If packed cell volume (PCV) and total protein are low (e.g., PCV < 20%), whole blood may be required (see “Blood Transfusion in Small Mammals,” below). Oxyglobin has not been approved for use in cats or exotic companion mammals but has been used successfully in small-volume boluses.21 Administer 2 mL/kg boluses over 10 to 15 minutes until normal heart rate and blood pressure (systolic blood pressure > 90 mm Hg) are obtained; then follow with a CRI of Oxyglobin at 0.2 to 0.4 mL/kg per hour. Vasopressors
Box 38-2 Correction of Perfusion Deficits Decompensatory phase of shock (bradycardia, hypotension, hypothermia): Slow IV or IO bolus over 10 minutes of hypertonic saline 7.2%-7.5% (3 mL/kg) + hetastarch (3 mL/kg) ↓ Begin external and core body temperature warming over 1-2 hours Begin crystalloids at maintenance rate (3-4 mL/kg per hour) ↓ When patient is warmed to 98°F (36.6°C), begin correction of hypovolemia to indirect systolic blood pressure > 90 mm Hg (recheck pressure after each bolus) Repeat boluses 3-4 times until blood pressure is normal: 1. Crystalloids (LRS, Normasol, Plasmalyte) at 10 mL/kg 2. Hetastarch at 3-5 mL/kg ↓
Positive response: indirect systolic blood pressure > 90 mm Hg: Crystalloids to correct dehydration plus ongoing losses (Box 38-3) ↓ Add maintenance fluids (3-4 mL/kg per hour)
Negative response: indirect systolic blood pressure < 90 mm Hg: Oxyglobin 2 mL/kg slow IV bolus ↓ No response: Check blood glucose, electrolytes, PCV and total protein, ECG ↓ If hypoglycemic: Give 50% dextrose diluted 50:50 with saline at 0.25 mg/kg If PCV < 20% and low total protein: Consider whole blood transfusion If abnormal cardiac function: Correct contractility (nitroglycerin 1⁄8 in./2.5 kg) ↓ No response: Consider vasopressor at small-animal dose
such as dopamine or norepinephrine are used to treat refractory hypotension in traditional pet species and could be considered in exotic companion mammals as well.32 Determine hydration status after successful resuscitation. As in traditional pet species, dehydration is often indicated and losses estimated by decreased skin turgor and dry mucous membranes14 (Box 38-3). Rehydration is best accomplished using an isotonic replacement fluid. The rate of fluid administration depends primarily on the rate of fluid losses and clinical status of the animal, as indicated by the physical examination and laboratory parameters. For animals with evidence of interstitial dehydration on physical examination but stable cardiovascular parameters, replace fluid deficits over 12 to 24 hours. If the interstitial volume was lost rapidly, replace the interstitial fluid deficit rapidly
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Box 38-3 Correction of Dehydration Deficits Estimating percent dehydration: 4%-5% Dry mucous membranes 5%-7% Dry mucous membranes, mild skin tenting 7%-9% Dry mucous membranes, moderate skin tenting ≥10% Dry mucous membranes, sunken eyes, altered mentation, significant skin tenting Calculate deficits: % dehydration x kg x 1,000 mL = fluid requirements in mL Add maintenance requirements: 3-4 mL/kg per hour plus ongoing losses (diarrhea/vomiting) Replace losses over a set period of time. Acute loss <24 hours: replace deficit over 6-8 hours. Chronic loss over 24-72 hours: replace over 24 hours.
Fig. 38-2 Blood collection for transfusion from a donor fer(4-6 hours).14 Fluid requirements for dehydration are calculated as percent dehydration × kg × 1,000 mL = fluid deficit (mL). Maintenance fluids replace ongoing losses (vomiting, diarrhea, polyuria), meet metabolic demands, and restore intracellular water balance until the animal is eating and drinking on its own.14 Maintenance requirements are higher in smaller patients because of their higher metabolic rate. Therefore the authors use a maintenance rate of 3 to 4 mL/kg per hour, which is double that used in cats and dogs.21 An objective way to assess whether the fluid volume is adequate is to evaluate body weight regularly throughout the day. Acute weight loss is commonly associated with fluid loss and can be used to determine if the patient is at risk of becoming dehydrated again.14 While urinary catheterization and measurement of urine can be used to objectively determine urinary output, this is not usually practical in many exotic companion mammals. An alternative approach is to weigh bedding or towels before and after placing them into the cage.
BLOOD TRANSFUSION Administer blood products when albumin, antithrombin, coagulation factors, platelets, or red blood cells are required.14 Most fluid-responsive shock patients will tolerate acute hemodilution to a 20% PCV. Most animals can tolerate an acute blood loss of 10% to 15% blood volume without requiring a blood transfusion. Acute hemorrhage exceeding 20% blood volume often requires transfusion therapy in addition to initial fluid resuscitation. When promoting cardiac output is the first priority in the management of acute hemorrhage, blood is not the ideal resuscitation fluid. Blood products do not promote blood flow as well as acellular fluids (i.e., HES, Oxyglobin, crystalloids). Therefore blood is rarely used for initial resuscitation unless the patient is exsanguinating or there is a coagulopathy. The density of erythrocytes impedes the ability of blood products to promote blood flow (a viscosity effect). HES, Oxyglobin, and crystalloids are less viscous and therefore promote blood flow. As in other species, continued blood loss, nonregenerative anemia with PCV 12% to 15% or below, and clotting disorders (such as rodenticide toxicosis) are indicators used to determine the potential need for a whole-blood transfusion. Whole blood can be administered at 10 to 20 mL/kg intravenously or intraosseously. The availability of blood products in sufficient quantities to meet the needs of exotic patients is often the limiting factor in
ret using a 22-gauge butterfly catheter and syringe with manual restraint only. Isoflurane can cause a drop in PCV in the ferret and should be avoided.
survival. Commercial blood banks do not carry exotic pet blood products with the possible exception of the ferret. The authors maintain a relationship with a local exotic animal rescue and keep a list of other clients willing to bring in patients for blood donation in exchange for clinic credit. Blood collection is performed in the carefully restrained or sedated patient via the cranial vena cava or other large, accessible vein using a needle and syringe or butterfly catheter (Fig. 38-2). It should be kept in mind that isoflurane can cause a reduction in PCV in the ferret and should be avoided.12 Larger needle size prevents red cell hemolysis; therefore the authors prefer to use a 22-gauge needle in rabbits and ferrets. Collect blood into syringes with sodium citrate at product-recommended doses. Ferrets are unique in that they appear to have a single blood type; therefore multiple donor transfusions are possible and commonly performed.29 Blood groups have not been identified in most other exotic companion mammals. A simplified cross match can be performed in these small patients by mixing 2 drops of plasma from the recipient with 1 drop of whole blood from the donor on a slide at room temperature.14 The development of macroscopic agglutination within 1 minute would suggest incompatibility. A successful major cross match does not imply that reactions will not occur but helps to minimize potential complications resulting from agglutination. The blood-administration set must include a filter to remove most of the aggregated debris. Administer donor blood by slow bolus or by infusion with a syringe pump into a catheter placed in the jugular, saphenous, or cephalic vein or into an intraosseous catheter. In cases of massive hemorrhage, administer blood more rapidly, within minutes. Administer blood transfusions within 4 hours of collection, according to standards set by the American Association of Blood Banks, to prevent the growth of bacteria.
USE OF GLUCOCORTICOIDS IN SHOCK The use of glucocorticoids in the treatment of shock is controversial. These drugs have been extensively investigated in the shock syndrome. Although they have repeatedly shown promise
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A Fig. 38-3 A constant-read-out thermometer (Electronic Thermometer, Veterinary Specialty Products, Mission, KS) with a flexible temperature probe used in a rabbit. The probe should be secured with tape to the tail to prevent it from slipping out.
in some experimental studies, they have not shown consistent efficacy in clinical shock syndromes. The side effects of immunosuppression, increased risk of infection, hyperglycemia, and gastric ulceration may outweigh their benefits. Use of corticosteroids in exotic companion mammals for the treatment of shock is not currently recommended.
ROUTES OF FLUID ADMINISTRATION Placement of IV or IO catheters in exotic companion mammals is discussed elsewhere in this text.28 Ideally, oral or subcutaneous fluids should be reserved for stable, standing patients that are suspected to be less than 5% dehydrated. In reality, the risks of catheter placement in some patients may outweigh its benefits. Therefore it may be beneficial to administer subcutaneous fluids while gently rewarming the patient prior to catheterization. Oral fluids should not be administered if there is significant risk of aspiration or evidence of gastrointestinal dysfunction.
MAINTENANCE OF NORMOTHERMIA All small animals experience some degree of hypothermia in the early decompensatory stages of shock. Body temperature is easily recorded in most species using a standard rectal thermometer, a constant-readout temperature probe, or an esophageal temperature probe (Fig. 38-3). New methods using infrared thermometry methods have been developed and tested in human patients and are currently under investigation for small mammals.8,15,19 The normal body temperature of exotic companion mammals varies. Some easily experience heat stress—for example, guinea pigs and chinchillas; therefore rewarming should proceed cautiously in these species.37 Monitor temperature constantly during the rewarming process, as severely depressed animals may not be able to move away from the heat source in case of overheating. External rewarming methods include the use of warming blankets and pads (Hot Dog Warming Unit, Augustine Biomedical and Design, Eden Prairie, MN), circulating-warm-air blowers (Bair Hugger, Arizant Healthcare, Eden Prairie, MN; Thermacare,
B Fig. 38-4 Hospitalized patients. A, Rabbit. B, Ferret. Both animals catheterized with fluids delivered via a pediatric infusion pump, intravenously in the cephalic vein. Note measurement of indirect systolic blood pressure in each. Rewarming with a warm-air circulating device (Thermacare, Gaymar Industries, Orchard Park, NY) (A) and a flexible heating blanket (Hot Dog Warmer, Augustine Biomedical and Design, Eden Prairie, MN) (B).
Gaymar Industries, Orchard Park, NY), or hot-water-filled examination gloves (Fig. 38-4). Warming of core body temperature warming is accomplished through warming of the intravenous or intraosseous fluids using an intravenous warmer (Meditemp III, Gaymar Industries; Hotline Fluid Warmer, Smiths Medical PM, Inc., Waukesha, WI) or by running the IV line under warming devices or through hot water. Inspired oxygen can be warmed by placing warmed fluid bags directly on top of the oxygen line. In severely hypothermic patients, attempts to rewarm via external methods only can result in the shunting of colder blood to the body core, causing rebound hypothermia.32
INDIRECT MEASUREMENT OF SYSTOLIC BLOOD PRESSURE Indirect measurement of systolic blood pressure is commonly used in veterinary medicine. In general, it is thought that, to ensure adequate organ perfusion in both the conscious and anesthetized patient, mammalian diastolic pressure should be above 60 mm Hg and systolic pressure above 90 mm Hg.
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A
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B
C Fig. 38-5 Measurement of indirect blood pressure using an ultrasonic Doppler and pediatric cuff in a ferret and rabbit using the thoracic limb (A, B) and a guinea pig using the pelvic limb (C).
Because of difficulties inherent in the measurement of diastolic (direct) blood pressure, measurement of indirect pressure is most commonly used in clinical practice.23 Methods for detection include use of a sphygmomanometer with an ultrasonic Doppler and newer oscillometric blood pressure measurement techniques.42 Advantages of the Doppler method include relatively low cost and portability. New developments in oscillometric methods may eventually make this a reasonable alternative for use in exotic companion mammals as well. To obtain blood pressure using the ultrasonic Doppler, place the patient in lateral or sternal recumbency. Place a pneumatic cuff above the carpus or tarsus or on the tail (Fig. 38-5). The rear leg can be used for blood pressure recording but appears to be less sensitive than the front leg. The cuff size should ideally be about 40% of the diameter of the limb or tail. Finger cuffs used in human medicine are available (Parks Medical, Aloha, OR). Shave the hair on the ventral carpus, tarsus, or medial midshaft of the radius-ulna or tail. Place the transducer probe crystal on the shaved area in a bed of ultrasonic gel and tape or hold it in place. Inflate the cuff to a suprasystemic pressure until the Doppler signal is extinguished. The first sound heard as the cuff is deflated denotes the systolic pressure. This technique requires practice and becomes more difficult with decreasing patient size. Studies on the use of blood pressure measurement in small mammals are lacking. One study indicated that the fore limb was more reliable than the pelvic limb for indirect measurement
of pressure in the rabbit.42 Another study in ferrets showed that larger-sized cuffs resulted in a decrease of 28 to 30 mm Hg in indirect systolic pressures when compared with direct systolic pressures.26 Keep in mind that studies in many species demonstrate some degree of disagreement between indirect and direct systolic blood pressure measurement; this is attributed to variations in cuff sizes and sensitivities as well as in arterial waveforms between anatomic sites. For this reason, the greatest benefit of blood pressure measurement may be as a trend monitor. The authors routinely use indirect blood pressure measurement along with other parameters to gauge response to therapy in the patient in hypovolemic shock and as part of monitoring during anesthesia. The authors have found indirect systolic blood pressures in normal exotic companion mammals to range between 80 and 120 mm Hg.
CRITICAL CARE CLINICAL PATHOLOGY It is extremely important that clinical pathology data be available as soon as possible to help guide decision making for the critical care patient. For this reason, in-house testing is preferred. A number of manufacturers produce equipment that enables in-house analysis of samples from exotic companion mammals. One biochemical analyzer can produce a panel with 0.13 mL of high-quality whole blood (VetScan, Abaxis, Union City, CA).
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LACTATE MONITORING Blood lactate concentrations are considered by some to be accurate indicators of inadequate tissue perfusion in humans and traditional pet species.4 Lactate concentrations have been shown to be a superior index of hypoxia as compared with oxygen delivery (DO2), oxygen consumption (VO2), oxygen extraction ratio, and cardiac index (cardiac output per minute per square meter of body surface area) in clinical studies of critically ill humans. In studies where no significant differences existed between survivors and nonsurvivors for DO2 and VO2, blood lactate concentrations were closely correlated with survival in humans and are therefore thought to be a more accurate prognostic indicator.4 Lactate monitoring in critically ill patients is especially important in those presenting in shock. Lactate values become elevated (i.e., >2.5 mmol/L) in domestic animals when perfusion parameters are poor; they usually return to normal when fluids are given to correct perfusion parameters to normal (i.e., heart rate, blood pressure, temperature, CRT). Lactate is used as a noninvasive critical care monitoring device—along with heart rate, blood pressure, and temperature—to help the clinician correct perfusion deficits. Normal lactate in a recent study in rabbits is 7.0 (± 2.6) mmol/L for the Nova analyzer (Nova Biomedical, Waltham, MA), 7.3 (± 2.9) mmol/L for the IDEXX analyzer (IDEXX Veterinary Chemistry Analyzer, Westbrook, MA), and 6.6 (± 3.7) mmol/L for the Point of Care Lactate Analyzer (Point of Care Portable Lactate, Arkray, Kyoto, Japan).36
USE OF PROTHROMBIN AND PARTIAL THROMBOPLASTIN TIMES Coagulopathy is not commonly reported in small mammals, although the number of cases of warfarin poisoning in ferrets may be similar to that seen in dogs and cats (ASPCA, personal communication). The use of coagulation parameters in smallanimal critical care medicine is very important; most coagulopathies in such patients occur with liver disease, toxicity, and disseminated intravascular coagulation. Unpublished data (M. Lichtenberger and Z. Schwartz) for normal ferrets comparing reference laboratory values (Antech Diagnostics, Irvine, CA) for prothrombin time (PT) and partial thromboplastin time (PTT) and values derived from the in-house SCA2000 (Synbiotics Corporation, Kansas City, MO) gave results of 12 ± 1.5 seconds from the reference laboratory and 20 ± 1 seconds for the SCA2000 device. PTT from the reference laboratory was 18 ± 2 seconds and for the SCA2000 was 52 ± 19 seconds. There was a good correlation between the PT results from the two different testing methods but not for the PTT. To the authors’ knowledge, no studies have been done in ferrets to determine normal activated clotting times (ACT). The ACT of ferrets may be similar to the PTT measurements in this study.
NUTRITIONAL SUPPORT Enteral nutrition should be delivered as rapidly as possible to prevent negative consequences of prolonged anorexia. Anorexia causes breakdown of the tight junctions between gastric and intestinal cells, leading to bacterial overgrowth and possible sepsis. Nutritional support is a vital component of treatment and is crucial for resolution and prevention of gastric stasis, especially in herbivores. Fluid replacement therapy must also play a
role in nutritional support, as the gastrointestinal tract must be hydrated to facilitate motility and function. Many small mammals tolerate syringe feeding well in general, but when syringe feeding is refused, placement of a nasogastric or esophagostomy tube should be considered (see “Treatment of Selected Common Emergencies,” and “Anorexia and Gastric Stasis,” below). The most suitable enteral diets for syringe feeding contain a high percentage of indigestible fiber as well as little fat and relatively low carbohydrates. Herbivore enteral diets are commercially available from Oxbow Pet Products (Critical Care, Oxbow Animal Health, Murdock, NE). The Oxbow products have been specifically formulated for nutritional support of herbivorous small mammals and rodents. The kilocalories are given on a dry-matter basis with 2.69 kcal/g of dry weight of the powder. Critical Care powder when mixed 1:1 with water provides approximately 1.9 kcal/mL, which can be used for syringe feeding of stable patients in hospital or for home feeding by the owner. Place the syringe into the diastema, or the large space between the incisors and cheek teeth. Perform syringe feeding slowly and carefully in order to avoid aspiration, especially in animals in respiratory distress. Unfortunately this formulation is not suitable for nasogastric tube feeding. Two diets designed for nasogastric tube feeding have been evaluated: Oxbow Critical Care Fine Grind and Lafeber Emeraid Herbivore (Lafeber, Cornell, IL). These diets should be fed as recommended by the manufacturers.
SEDATION AND ANESTHESIA OF THE CRITICALLY ILL SMALL MAMMAL Anesthesia should be avoided in the critically ill patient; however, some diagnostic testing and procedures can produce discomfort and stress, negatively affecting patient survival. The risk of sedation and/or anesthesia should be weighed against the risk of foregoing diagnostics and treatments or attempting procedures in the conscious animal. Hemodynamic abnormalities (shock, hypovolemia) and hypothermia should be corrected prior to administration of sedatives/anesthetic agents. It should be noted that many minor procedures (catheterization, sample collection, diagnostic imaging) can be accomplished in ill exotic companion mammals with the use of sedation alone or with the addition of local analgesia if indicated. For example, for intravenous or intraosseous catheterization, the authors prefer a combination of low-dose midazolam (0.25 mg/kg) plus an opioid, with topical application or infiltration of lidocaine over the catheterization site. Isoflurane induction via face mask has historically been favored by exotic mammal practitioners. While isoflurane and sevoflurane have a wider margin of safety than many older anesthetic agents, they can produce apnea and hypotension, especially at the higher concentrations required when they are used as sole agents. Multimodal analgesic/anesthetic protocols greatly reduce the needed amount of any single agent, thus enhancing patient safety and survival.22 Avoid injectable induction agents in critically ill patients. An exception may be etomidate (Amidate, BenVenue Laboratories, Bedford, OH), an imidazole derivative that undergoes rapid redistribution and hepatic metabolism, resulting in rapid recovery following a single bolus.16 Etomidate induces minimal cardiovascular depression and has a wide margin of safety. It has a respiratory depressant effect, but this is dose-dependent; therefore patients should be intubated if possible or supplied
CHAPTER 38 Emergency and Critical Care of Small Mammals with oxygen via face mask. Give the drug to effect, using the lower end of the recommended dose (1-2 mg/kg IV). This drug is used in combination with diazepam or midazolam to prevent myoclonic twitching, rigidity, and seizure16 For longer surgical procedures expected to produce discomfort, deliver analgesia via CRI. Delivery of analgesics via CRI allows reduction of the amount of anesthetic agent required, which is of extreme benefit in critically ill patients.22
TREATMENT OF SELECTED COMMON EMERGENCIES ACUTE RENAL FAILURE
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Box 38-4 Diuresis for Acute Renal Failure in Small Mammals Correct perfusion deficits (shock) as indicated in Box 38-2. Correct dehydration deficits as indicated in Box 38-3. Implement diuresis until azotemia and electrolyte abnormalities have resolved: Maintain at 3-4 mL/kg per hour. Add measured urinary output (or estimate at 2-4 mL/kg per hour). Add additional losses (vomiting/diarrhea). Add additional 3%-5%. Continue maintenance fluids until the patient is eating and drinking well.
Acute renal failure (ARF) is characterized by a sudden onset of filtration failure, accumulation of uremic toxins, and dysregulation of fluid, electrolytes, and acid-base balance. It is potentially reversible if diagnosed quickly and treated aggressively. Azotemia is defined as an elevation in blood urea nitrogen (BUN) and creatinine. The serum urea nitrogen is usually severely elevated in an azotemic exotic companion mammal, whereas creatinine may be severely elevated, mildly elevated, or normal. Acute renal failure is categorized as prerenal (shock, hypovolemia), renal (intrinsic renal disease), or postrenal (obstruction). Prerenal azotemia is characterized by mild to moderate azotemia, clinical dehydration, hypovolemia or hypotension in conjunction with a concentrated urine specific gravity (>1.030).29 Differentiation between ARF and chronic renal failure is important in terms of prognosis, since chronic renal failure may not be completely reversible. Both acute and chronic renal failure will often show laboratory data with elevations in BUN and creatinine. However, with chronic renal failure, there is often loss of body condition, polydipsia and polyuria, anemia, and isosthenuria (1.010 to 1.015).29 Treatment of ARF involves correction of perfusion deficits and dehydration (see “Shock and Fluid Therapy,” above) and diuresis to correct azotemia, electrolyte, and acid-base status (Box 38-4). Treatment also includes correction of the underlying cause. Once the patient is normotensive and rehydrated, record hourly urine production via urinary catheterization or pre/post weighing of towels or bedding. The volume of fluid to be administered in each 1-hour period is therefore the sum of the calculated maintenance requirements (3-4 mL/kg per hour) plus estimated losses via vomiting/diarrhea plus urine volume produced from the previous hour. In general, assume that most patients with ARF become 3% to 5% dehydrated each day because of ongoing losses. During the polyuric phase of ARF, the patient may produce very large volumes of urine, necessitating aggressive fluid administration. In some cases, urine loss may be as high as 5 to 10 mL/kg per hour. Gradually discontinue fluids when hydration is maintained, urine production is normalized, BUN and creatinine are normal, and the patient is eating and drinking. Taper the fluids by 50%/day.
Hyperkalemia is a common consequence of prolonged urinary tract obstruction. If hyperkalemia is present without cardiac or ECG abnormalities and perfusion is good, forced diuresis and relief of the obstruction is generally effective in correcting the potassium excess. Hyperkalemia in the presence of ECG abnormalities (loss of the P wave, widening of the QRS complex, peaked T wave, and a short QT interval; note as the QRS and T waves merge, a sine wave is recognized) requires treatment. Administer calcium gluconate at 50 to 100 mg/kg IV slowly while monitoring the ECG. For prolonged ECG abnormalities, administer regular insulin at 0.2 U/kg IV. Follow insulin therapy with a glucose bolus of 1 to 2 g for each unit of insulin administered. This treatment should begin to lower potassium concentration within 10 minutes to 1 hour. If ECG abnormalities persist beyond 1 hour, administer a second dose of insulin. Additional fluids should contain 1.25% dextrose solution for at least 24 hours after insulin therapy. Check blood glucose regularly.30 Relief of obstruction is imperative and must be accomplished rapidly.13 Urinary catheterization is extremely difficult in male exotic companion mammals with the exception of the ferret, which is covered in detail in Chapters 4 and 7. Cystocentesis to relieve bladder overextension temporarily can be performed once but should not be repeated. The authors have not encountered urinary obstruction in the female ferret. Urethroliths in female rabbits tend to produce partial obstruction. In most cases, these can be removed manually if within reach or gently pushed into the bladder for later retrieval. Manipulation of uroliths in female rabbits and guinea pigs is greatly facilitated with rigid 2.7- to 1.9-mm endoscopy with insufflation of sterile fluid. When catheterization is impossible and surgical resolution must be delayed, an alternative is placement of a temporary percutaneous catheter (Fig. 38-6). These specialized catheters are available in sizes from 5 to 8 Fr and can be placed using minimally invasive techniques. Complete instructions are available from the manufacturer (Infiniti Medical, Orlando, FL).
URINARY OBSTRUCTION
RESPIRATORY DISTRESS
Urinary obstruction can be seen in any small mammal but is most commonly reported in male ferrets, rabbits, and guinea pigs. In male ferrets, urethral obstruction is linked to poor-quality diets or is caused by prostatomegaly and urinary obstruction due to adrenal disease.30 Treatment of ARF is described above.
Exotic companion mammals in distress but not arrest should immediately be placed into a quiet, oxygen-enriched environment (oxygen flow rate at 5 L/min for the first 12-24 hours). Commercial oxygen cages are ideal, but if such a cage is not available, one can be made from a pet carrier in a plastic bag
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ANOREXIA AND GASTRIC STASIS
Fig. 38-6 Temporary percutaneous catheter for management of urethral obstruction (Infiniti Medical, Orlando, FL).
or, in the case of a small patient, a large anesthetic face mask. Oxygen can also be delivered via an appropriately sized anesthetic mask or nasal cannula, but this is often stressful. Small mammals with respiratory disease frequently benefit from sedation to reduce anxiety. The authors prefer midazolam at 0.25 to 0.5 mg/kg combined with butorphanol at 0.4 to 0.5 mg/kg or buprenorphine at 0.04 to 0.06 mg/kg IM. Humidification of oxygenated air helps clear respiratory secretions and foreign material within the trachea and bronchi. Commercially manufactured intensive care units can provide oxygen, heat, and humidity. Alternatively, humidification can be obtained by bubbling the oxygen line through an isotonic fluid solution. It is important to distinguish between upper and lower airway disease. Upper airway disease involves the nares and nasal cavities, sinuses, larynx, and trachea, while lower airway disease involves the small airways, lung parenchyma, or pleural space. The following generalizations are useful in traditional pet species and in many cases can be applied to exotic companion mammals as well. In carnivores and many rodents, upper airway lesions often create a normal breathing pattern with a loud inspiratory wheeze.30 However, in obligate nasalbreathing species (rabbits, chinchillas, guinea pigs), diseases of the nares and nasal cavities can produce severe respiratory distress.11 Referred upper airway sounds are heard on auscultation of the chest but are loudest over the affected area. Tracheal obstruction can also present with pronounced gagging and swallowing. Small-airway disease is often associated with a soft expiratory wheeze and synchronous movement of the chest and abdomen.30 There is a wheezing sound on auscultation of the chest. Exotic companion mammals with these findings may benefit from bronchodilators—for example, terbutaline at 0.01 mg/kg IM or SC and oxygen. With parenchymal disease, there are increased lung sounds with respiratory wheeze and synchronous movement of the chest and abdomen. Differentials include pulmonary edema secondary to heart disease and pneumonia. If cardiac disease is suspected, administer furosemide at 3 to 4 mg/kg SC or IM and a vasodilator (nitroglycerin ointment 1⁄8 in. per 2.5 kg body weight on the tongue or gums). Cardiac disease should
Anorexia is a common presenting sign in all small mammals but is particularly common in rabbits and other herbivores. Anorexia in these species may be due to pain, systemic disease, gastrointestinal stasis, or even psychological stress. Any period of anorexia lasting more than 1 or 2 days is a potential emergency. Anorexia produces dehydration, which slows gastrointestinal motility and eventually leads to hypovolemia and hepatic lipidosis. Anorexia is particularly dangerous in rabbits and other herbivores; it is discussed in more detail in those respective chapters. Many anorectic rabbits are dehydrated and hypovolemic. Motility drugs are helpful; the authors have found cisapride and cisapride-like drugs to be extremely beneficial. Cisapride is currently available in the United States through compounding pharmacies. Oral cisapride in rabbits is absorbed rapidly from the gastrointestinal tract, with a plasma half-life similar to that in dogs.27 Other data shows that cisapride may modify the contractile responses of the rabbit intestine to ranitidine, resulting in enhanced effects when these agents are administered together.18 Trimebutine is a cisapride-like drug available in Canada and parts of Europe. Anecdotal reports of the use of oral and injectable forms of the drug are extremely promising. The use of metoclopramide as a motility agent, either subcutaneously or as a CRI, has been reported with variable results. Early enteral feeding decreases pain, helps with motility of the gastrointestinal tract, and decreases bacterial translocation. Hepatic lipidosis develops quickly, often within 36 to 48 hours. Start nutritional support as soon as possible. Hand-feeding in small mammals is well described. When hand-feeding is difficult, painful, or stressful, nutritional support is facilitated by placement of a feeding tube. In carnivores, esophagostomy tube placement is straightforward and accomplished under general anesthesia, as in traditional pet species. Use nasogastric tubes in rabbits and larger guinea pigs. The authors recommend a 3.5- to 8-Fr Argyle tube (Surgivet, Waukesha, WI), which is softer than a red rubber tube and less susceptible to stomach acid degradation. Length to reach the stomach is determined by measuring from the tip of the nose to the last rib. Sedation is extremely useful for placement of a nasogastric tube, along with 2% lidocaine gel placed into the patient’s nostril. Restrain the patient carefully, with the head ventrally flexed and the neck straight to avoid compression of the trachea. Pass the tube ventrally and medially into the ventral nasal meatus. Advance the end of the tube until it reaches the stomach. Suture the tube to the margin of the nares and to skin of the forehead, between the eyes
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Box 38-5 Treatment of Gastric Stasis in the Critically Ill Herbivorous Small Mammal Correct perfusion deficits (shock) as indicated in Box 38-2. Correct dehydration deficits if present as indicated in Box 38-3. Place nasogastric tube. Sedate with midazolam plus an opioid. Begin feeding (Oxbow Critical Care Fine Grind or Lafeber’s Herbivore Hand Feeding formula) at manufacturers’ instructions. • Decrease the amount of maintenance fluids by amount of fluid fed by nasogastric tube. • Begin offering regular diet, remove the tube when the animal is eating on its own, and supplement syringe feeding as needed.
Fig. 38-7 Nasogastric tube in place in an anorexic rabbit (5 Fr). The tube is secured by suturing to the skin at the nare and between the ears.
Fig. 38-8 Nasogastric tube (3.5 Fr) in an anorexic guinea pig secured as in the rabbit in Figure 38-7. The ideal tube-feeding product (i.e., Critical Care Fine Grind, Oxbow Animal Health, Murdock, NE) must be appropriate for the species in question and capable of passing through small-diameter tubes without risk of occlusion. Delivery is enhanced with smaller-size syringes (1 mL for 3.5 Fr and 3 mL for 5 Fr).
(Fig. 38-7). Tape the remainder of the tube end around the neck. Verification of tube placement can be determined radiographically and/or by aspiration of gastric contents. Smaller nasogastric tubes (3.5-5 Fr) can be placed in very small dwarf rabbits or guinea pigs (Fig. 38-8). Products for use with nasogastric tubes must be chosen carefully. Oxbow Critical Care is useful for syringe feeding. Critical Care Fine Grind is specially manufactured to pass through the nasogastric tube. Regardless of the level of enteral support selected, food should be available to the animal at all times for voluntary consumption. Box 38-5 summarizes fluid and nutritional support for patients with gastric stasis.
References 1. Aldrich J, Haskins S. Monitoring the critically ill patient. In: Bonagura JD, Kirk RW, eds. Current veterinary therapy. Vol X11. Philadelphia: Harcourt Publishers; 2004:465-469. 2. American Heart Association guidelines for cardiopulmonary resuscitation and emergency cardiovascular care. Circulation. 2005;112(suppl 24):IVI-203. 3. Astiz ME. Pathophysiology and classification of shock states. In: Fink MP, Abraham EA, Vincent J-L, Kochanek PM, eds. Textbook of critical care. Philadelphia: Elsevier Saunders; 2005:897-997. 4. Bernardin G, Pradier C, Tiger F, et al. Blood pressure and arterial lactate level are early indicators of short-term survival in human septic shock. Intensive Care Med. 1996;22:17-25. 5. Bihun C, Bauck L. Small rodents: basic anatomy, physiology, husbandry, and clinical techniques. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2004:286-298. 6. Blouin A, Cormier Y. Endotracheal intubation in guinea pigs by direct laryngoscopy. Lab Anim Sci. 1987;37:244-245. 7. Briscoe J, Sring R. Techniques for emergency airway and vascular access in special species. Sem Avian Exot Pet Med. 2004;13:118-131. 8. Carroll D, Finn C, Judge B, et al. A comparison of measurements from a temporal artery thermometer and a pulmonary artery catheter thermistor [abstract]. Am J Crit Care. 2004;13:258. 9. Costello MF. Principles of cardiopulmonary cerebral resuscitation in special species. Sem Avian Exot Pet Med. 2004; 13:132-141. 10. Crowe DT. Cardiopulmonary resuscitation in the dog: a review and proposed new guidelines (part II). Sem Vet Med Surg (Small Anim). 1988;3:328-348. 11. Donnelly TM. Disease problems of small rodents. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. St. Louis: Elsevier Saunders; 2004:299-315. 12. Edling TM. Advances in anesthesia monitoring in birds, reptiles and small mammals. Exot DVM. 2003;5:15-20. 13. Fossum TW, Hedlund CS, Hulse DA, Johnson AL, et al. Surgery of the bladder and urethra. In: Fossum TW, Hedlund CS, Hulse DA, Johnson AL, et al. eds. Small animal surgery. 2nd ed. St. Louis: Mosby; 2002:572-609. 14. Haskins S. Fluid therapy. In: Kirk R, ed. Handbook of veterinary procedures and emergency treatment. 5th ed. Philadelphia: WB Saunders; 1990:574-600.
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15. Hebbar K, Fortenberry JD, Rogers K, et al. Comparison of temporal artery thermometer to standard temperature measurements in pediatric intensive care unit patients. Pediat Crit Care Med. 2005;6:557-561. 16. Janssen PA, Niemegeers CJ, Marsboom RP. Etomidate, a potent non-barbituate hypnotic. Intravenous etomidate in mice, rats, guinea pigs, rabbits and dogs. Arch Int Pharmacodyn Ther. 1975;214:92-132. 17. Kujime K, Natelson B. A method for endotracheal intubation of guinea pigs (Cavia porcellus). Lab Anim Sci. 1981;31:715-716. 18. Langer JC, Bramlett G. Effect of prokinetic agents on ileal contractility in a rabbit model of gastroschisis. J Pediatr Surg. 1997;32:605-608. 19. Lawson L, Bridges EJ, Ballou I, et al. Accuracy and precision of non-invasive temperature measurement in adult intensive care patients. Am J Crit Care. 2007;16:485-496. 20. Lennox AM, Capello V. Trachea intubation in exotic companion mammals. J Exot Pet Med. 2008;17:221-227. 21. Lichtenberger M. Principles of shock and fluid therapy in special species. Semin Avian Exot Pet Med. 2004;13:142-153. 22. Lichtenberger M, Ko J. Anesthesia and analgesia for small mammals and birds. Vet Clin North Am Exot Anim Pract. 2007;10:293-315. 23. Marino PL. Arterial blood pressure. In: Marino PL, ed. The ICU book. 2nd ed. Philadelphia: Lippincott Williams & Wilkins; 1997:143-153. 24. Marino PL. Cardiac arrest. In: Marino PL, ed. The ICU book. 2nd ed. Philadelphia: Lippincott Williams & Wilkins; 1997:260-298. 25. Muir WW. Drug antagonism and antagonists. In: Gaynor JS, Muir WW, eds. Handbook of veterinary pain management. St. Louis: Mosby; 2002:393-404. 26. Olin JM, Smith TJ, Talcott MR. Evaluation of noninvasive monitoring techniques in domestic ferrets (Mustela putorius furo). Am J Vet Res. 1997;58:1065-1069. 27. O’Rourke D, Micheals M, Monbalie J, et al. Pharmacokinetics and tissue distribution of the new gastrokinetic agent cisapride in rat, rabbit, and dog [abstract]. Arzneimittelforschung. 1987;37:59-67. 28. Otto C, Crowe D. Intraosseous resuscitation techniques and applications. In: Kirk R, Bonagura J, eds. Current veterinary therapy XI: small animal practice. Philadelphia: WB Saunders; 1992:107-112.
29. Paul-Murphy J. Critical care of the rabbit. Vet Clin North Am Exot Anim Pract. 2007;10:437-461. 30. Pollock C. Emergency medicine of the ferret. Vet Clin North Am Exot Anim Pract. 2007;10:463-500. 31. Richtarik A, Woolsey TA, Valdivia E. Method for recording ECGs in unanesthetized guinea pigs. J Appl Physiol. 1965;20:1091-1093. 32. Rudloff E, Kirby R. Colloid and crystalloid resuscitation. Vet Clin North Am Small Anim Pract. 2001;31:1207-1229. 33. Safar P. On history of modern resuscitation. Crit Care Med. 1996;24(suppl 2):S3-S11. 34. Schadt JC, Ludbrook J. Hemodynamic and neurohumoral responses to acute hypovolemia in conscious mammals. Am J Physiol. 1991;260:H305-H318. 35. Schoemaker NJ, Zandvliet MM. Electrocardiograms in selected species. Sem Avian Exot Pet Med. 2005;14:26-33. 36. Schwartz Z, Lichtenberger MK, Thamm DH, Kirby R. Lactate normals in the healthy rabbits comparing three different analyzers. Proc Int Vet Crit Care (abstract). 2006;1038. 37. Strake J, Davis L, LaRegina M, et al. Chinchillas. In: Laber-Laird K, Swindle M, Flecknell PA, eds. Handbook of rodent and rabbit medicine. Exeter: BPC Wheatons; 1996:151-181. 38. Tran DQ, Lawson D. Endotracheal intubation and manual ventilation of the rat. Lab Anim Sci. 1986;36:540-541. 39. Velasco IT, Rocha e Silva M, Oliveira MA, Silva RIN. Hypertonic and hyperoncotic resuscitation from severe hemorrhagic shock in dogs: a comparative study. Crit Care Med. 1989;17:261-264. 40. Yasaki S, Dyck PJ. A simple method for rat endotracheal intubation. Lab Anim Sci. 1991;41:620-622. 41. Yost CS. A new look at the respiratory stimulant doxapram. CNS Drug Rev. 2006;12:236-249. 42. Ypsilantis P, Didilis VN, Politou M, et al. A comparative study of invasive and oscillometric methods of arterial blood pressure measurement in the anesthetized rabbit. Res Vet Sci. 2005;78:269-275.
CHAPTER
39
Behavior of Small Mammals
Teresa Bradley Bays, DVM, CVA
Rabbits Social Behaviors Play Behaviors Behavior Problems Litter-Box Training Introduction of New Conspecifics Mourning the Death of a Bonded Mate Ferrets Social Behaviors Play and Sleeping Behaviors Behavior Problems Introduction of New Conspecifics Litter-Box Training Guinea Pigs Social Behaviors Play Behaviors Communication Behaviors Behavior Problems Introduction of New Conspecifics Chinchillas Social Behaviors Play Behaviors Communication Behaviors Behavior Problems Introduction of New Conspecifics Sugar Gliders Social Behaviors Communication Behaviors Play Behaviors Behavior Problems Introduction of New Conspecifics Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
Hedgehogs Social Behaviors Communication Behaviors Play Behaviors Behavioral Training Techniques for Small Mammals Conclusions
In many cases, the behavior of small mammals in captivity is very similar to that seen in the wild. Therefore, because of the constraints of confinement and unrealistic expectations of owners, behavioral problems often arise from the owners’ inability to provide for normal behaviors. Most small mammals that are kept domestically are also prey species, which causes them to be more susceptible to stressors associated with actual or perceived predators. Overall, if their psychosocial needs are not addressed and their instinctual personalities cannot be expressed, the result may be additional behavior problems and even failure to thrive. It is imperative that veterinary practitioners understand what is normal for each species, so that behavioral cues associated with fear or stress as well as abnormal behavior are noted and addressed as soon as possible. Veterinary staff must be educated on what signs to look for in the daily care of these pets and what questions to ask during anamnesis. Equally important is to educate clients about these subtleties so that anxiety, discomfort, pain, and disease are detected and alleviated more promptly. The behavior of small mammals is a very large topic covering many species. For this chapter, therefore, only those behaviors that are not associated with the medical care of rabbits, ferrets, and some other more common species are covered. Many of the behaviors associated with the medical care of these animals are 545
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covered in other chapters. For a more complete reference on the behavior of small mammals, the reader may refer to Exotic Pet Behavior—Birds, Reptiles, and Small Mammals.3,4,11,21-23
RABBITS SOCIAL BEHAVIORS Rabbits are a very social species and, in the wild, live in large stable groups or warrens of up to several hundred animals.29 Because of their highly social nature, rabbits do best if they are kept in groups of two or three or more and given the right combination of companions and the proper introductory time (see “Introduction of New Conspecifics,” below). By ensuring these conditions, inseparable bonds can be created. In the laboratory setting, domesticated rabbits grouped in small social groups were found to have increased social contact and exercise34 and less stereotypic behaviors, including pawing at the corners of the cage, overgrooming, wire biting, overeating, and playing with the water supply.4 In the hospital setting, allowing bonded mates to stay together may help to decrease stress and increase the success of treatment.5,20 Rabbits can also form strong social bonds with human companions and with other domesticated pets. When not socialized with different animals and people and handled extensively at an early age, rabbits tend to be more shy and fearful and have more trouble adapting to new situations for the rest of their lives. As with other pets, it is important that introductions be performed in a careful and safe manner. Additionally, all interactions with other pets should be supervised by an adult, because domestication does not abolish the natural instincts and behaviors of predator species such as dogs and cats. Rabbits communicate with each other and with human companions with marking behaviors (see “Territorial and Destructive Behaviors,” below) and with verbal and postural forms of communication. Verbal and postural communication behaviors with which the rabbit caregiver should be familiar are listed in Table 39-1.
PLAY BEHAVIORS Normally, rabbits are intelligent, inquisitive, and inventive; research has shown that rabbits kept continuously caged exhibit more nervous behaviors than rabbits kept in an open area.12 These attributes are often difficult to appreciate when a rabbit is caged most of the time; therefore it is important for rabbits to be given time out of the cage in a safe area for at least several hours a day. Rabbits will nudge with their noses or beg on hind legs for attention from human companions as well as from bonded rabbit mates and other pets they have learned to trust. Both male and female rabbits will also weave in and out of their owner’s feet to gain attention and if “courting” them. They will push and toss objects around5 and jump on and off the couch. If a paper is wadded and thrown, a rabbit may chase after it and sometimes will retrieve it. Rabbits like to instigate chase games with other rabbits and with humans, but a human companion who chases a rabbit may be more likely to be perceived as a predator than a playmate. Juvenile rabbits are particularly playful and need lots of space to run, jump, and twist. An extremely happy or excited rabbit may perform what has been fondly called a “binkie”—a jump high in the air with a twist similar to what an excited lamb will do.
When rabbits are through with play or being handled, they may nudge with their noses to say “enough attention.” An irritated shake of the head or vigorous ear shaking may also signal displeasure at unwanted handling or the need for time to settle down. A rabbit will also shake its body (much as a dog might shake off water) as it settles down, and as if to shake off unwanted attention. When not playing or eating, rabbits spend a lot of time resting sternally or sleeping on their sides.
BEHAVIOR PROBLEMS Undesirable behavior in rabbits includes fecal and urine marking, chewing, digging, and aggression—either rabbit to rabbit or rabbit to human aggression. All of these negative behaviors are most obvious at about 3½ to 6 months of age, when rabbits become sexually mature. Boundaries will be tested as rabbits exhibit more instinctual behaviors, become more assertive and mischievous, and develop a stronger drive to exhibit more territorial behaviors as well as to perform other behaviors that humans may find undesirable. It is also at this time that they are establishing a social order within their group of humans, other rabbits, and in some cases other pets. This strong will for autonomy and independence may be viewed as a nuisance by unprepared owners as the rabbits become less cute and cuddly almost overnight. Clients must understand that these are normal instinctive behaviors and not meant to be “spiteful.” These negative behaviors are a temporary function of adolescence. With careful handling, time, and having the rabbit surgically altered, these behaviors will usually lessen. New owners often have unrealistic expectations for their rabbit pets and must realize how important it is to understand that, as individuals, they may not all react in the same way to similar situations. It is important to understand that each rabbit has a unique personality. It is also important to realize that how the caregiver responds will determine his or her future relationship with the pet rabbit. The rabbit caregiver must reward desirable behaviors instead of accidentally rewarding undesirable ones. Alternatively, he or she can control the environment so that the undesirable behaviors do not occur or occur less.
Aggressive Behaviors Aggression that rabbits exhibit towards humans is primarily motivated by fear.9 As with other domestic pets, negative behaviors can be inadvertently reinforced as caregivers pull away from the pet and interact with it less in response to an adverse interaction. For instance, if a rabbit bites a human and that human stops handling the animal, it may learn that biting will keep humans away. Also, any rough handling in response to mischievous behavior can cause damage to the trust relationship that a rabbit has with its caregiver, thereby creating and/or reinforcing an adversarial relationship. Behavior modification (see “Behavioral Training Techniques for Small Mammals,” below) is recommended for rabbits that have developed fear aggression.9 Aggression between rabbits is usually due to defense of territory, fear, and/or the desire to establish dominance (fighting, for instance, to establish rank or priority access to resources such as food or potential mates), all of which can become worse as these animals reach sexual maturity. Aggression is particularly prevalent between males at puberty, and serious injury may follow if such rabbits are not separated. Bucks will spray urine and begin mounting objects, people, and other pets. It is best
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Table 39-1 Verbal and Postural Communication Behaviors in Rabbits4,7,19 Name of Behavior
Description of Behavior
Function of Behavior
Fear posture
Make the rabbit appear smaller in order to be less likely to be noticed by a predator
Alert posture
Will lie motionless in a crouched position with feet beneath body, head extended, ears flattened against head and eyes bulging Ears are held forward or laterally
Erect tail
Tail is held in an upright position away from the body
Licking a companion
Lick a bonded mate, a human companion, or other trusted pet Biting gently Biting more assertively Weaving in between feet or circling the feet of the human companion Moving the head horizontally from side to side especially when being carried The tail is twitched rapidly back and forth
Nipping Weaving/circling Scanning Tail twitching Presenting Begging Nudging gently Aggressive nudging/digging Tooth purring Teeth grinding Chinning Urine spraying Head shaking, ear shaking, and/or body shudder Aggressive posture
Lying flat on floor with feet tucked, head extended, and chin on the floor Sitting vertically with front legs elevated off the ground A push using the nose on a human or another rabbit Using the nose to push and front feet to dig at objects A low-pitched hum with teeth lightly vibrating and whiskers quivering A slower, louder tooth crunching with eyes bulging Rubbing underside of chin on objects or bonded mates or humans
Allow the rabbit to hear better and to be able to bolt if needed To exhibit excitement or anticipation of a happy event or if threatened A sign of affection To solicit attention Signals anger To solicit attention or a courting behavior Evidence of impaired vision or difficulty focusing When urine spraying or to exhibit sexual interest To solicit petting by human companions or for grooming by another trusted rabbit To solicit food, treats, or attention To gain attention or to signal “enough attention” Signals anger or irritation A sign of contentment A sign of pain To mark territory or possession of objects or companions with secretions from scent glands on chin To mark territory, also a sexual behavior Signals unwanted handling or when settling down To signal anger
Thumping or foot stomping
To spray urine on objects, people, or other pets To vigorously shake the head, the ears, or the whole body An upright stance with ears flat and tail stretched out, also may kick high and backward A single or repetitive stomp with the hind foot or feet
Vocalization
Description of Sound
Function of Vocalization
Grunt, growl
A growling or snorting sound sometimes like a bark
Honking/oinking Scream
A honking sound A high-pitched repetitive scream
Wheezing/sniffing
An intermittent nasal sound that is often mistaken for a respiratory infection
Signals anger, annoyance, or territorial protection To solicit food, attention, or courtship Signals fear, terror, or pain; may make this sound when seizuring or when dying More vocal rabbits will make this sound to show irritation
to neuter them early, before this aggression begins, to quell this behavior. Does will often have intense mood swings, and they will also mount companions, spray urine, and begin digging and displaying nesting behaviors. They may become more aggressive toward people, other rabbits, and other pets. Groups of female rabbits that are bonded and have been grouped together from a young age tend to continue to get along despite hormonal changes associated with puberty. However, they should still be spayed to prevent medical problems often seen in intact females (see Chapter 17).
Signals anger or an alarm or warning of danger to other rabbits
Rabbits will exhibit anger and annoyance with a grunt, growl, snort, or short barking sound. An upright stance will be assumed, with ears flat and tail stretched out. An angry, aggressive rabbit may also kick high and backwards and will often lunge and bite. If a rabbit nips or bites, the caregiver should respond with a short, high-pitched yip to indicate that the behavior is not acceptable. This mimics what a rabbit might do if bitten too hard by a conspecific and is therefore a response that the rabbit will likely understand. Improper socialization and boredom can lead to aggression in rabbits. Providing for mental stimulation and exercise
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Box 39-1 Behavioral Enrichment for Rabbits4,7 These recommendations are made to simulate the natural environment and behavior as much as possible in order to provide for the emotional and psychosocial needs of pet rabbits. • Allow rabbit safe space for play and exercise • Keep in bonded pairs or trios whenever possible • Provide UVB lighting • Provide foraging opportunities • Dangle treats from a high place • Hide food treats in paper cup/cardboard box • Compressed alfalfa cubes and free choice of grass hays • Cardboard boxes with holes cut in them for tunnels • Large PVC tubes and/or dryer hose to use as a tunnel • Straw mats and baskets to chew on, dig at and hide in • Low carpeted ramps to get to higher places • Telephone books for them to chew on and dig at • Rabbit safe toys (see www.rabbit.org /links/mail-orderresources.html for links to retailers that sell toys for rabbits) • Paper bags to hide in or eat hay in • Toilet paper/paper towel tubes
can help to decrease boredom (Box 39-1). Previous traumatic events may also create aggressive behaviors, and it takes time and patience for some rabbits to regain confidence and trust that they will not get hurt again. In general, animals that are well socialized and handled gently often recover from traumatic events more quickly, whereas rabbits that are poorly socialized are more likely to perceive everything as a traumatic event. Aggression or other behavior changes that may be related to a medical problem should be assessed by a veterinarian skilled in rabbit medicine. These behaviors might include pain-related problems such as soft tissue trauma or osteoarthritis, urinary tract infections, and gastrointestinal stasis. Medical implications of abnormal behaviors and clinical signs associated with pain in rabbits have been previously reported.4,6 Failure to recognize behavior changes associated with medical problems from those that occur in response to environmental stimuli may lead to misdiagnosis and continued suffering or discomfort of the rabbit.
Territorial and Destructive Behaviors Territorial behaviors include aggression, as previously addressed, as well as chewing, digging, and marking behaviors. Marking is accomplished by scattering fecal pellets at perceived territorial boundaries and urinating on caregivers or their personal items or, again, at perceived boundaries. Chinning, another marking behavior, is done on objects, other pets, and people in order to mark them with secretions from scent glands, which have been found to contain different volatile components, depending on geographic location,16 and in the wild as a means of maintaining dominance hierarchies in a warren.16,17 Territorial and destructive behaviors can be quelled with surgical altering, which helps to eliminate the need to be territorial and redirecting the behaviors (see “Behavioral Training Techniques for Small Mammals,” below). Increasing the number of litter boxes and moving them to the areas where they will be most used can also be helpful (see “Litter-Box Training,”
below). Remember that the rabbit may be marking his favored “person” by urinating in the clothes basket or on the bed, so this is actually a “compliment.” The only way to control it may be to keep the rabbit away from these personal items. Chewing and digging are natural rabbit behaviors; since they cannot be controlled, the caregiver should try to divert the rabbit’s attention and provide areas that are rabbit-safe as well as offering appropriate items to dig at or chew on.9,10 Digging boxes can be created by providing a covered box with a hole in the side and filling it with shredded paper, a basket filled with hay or straw, or a cardboard box with towels in it.9 Female rabbits tend to be more prone to chewing even if they are spayed.19 Safe choices for chewing include apple, willow, and aspen branches, untreated pine lumber (molding, for example), and untreated straw baskets. These behaviors will also be seen when rabbits are bored and/or seeking attention.
Managing Behavior Problems Much of the behavior exhibited by rabbits is instinctive, but it may also be learned. Behavior problems are usually a temporary function of puberty and will often diminish over time unless they are reinforced by mishandling or by not being addressed at all. It is highly recommended that rabbits be neutered and spayed early in order to help to quell the natural hormonally directed behaviors previously discussed. Decreased hormonal influence on behavior after neutering will often not be appreciated for 30 days or more after altering, and it may take up to 6 months in larger breeds for negative sexual behaviors to decrease.19 It is also important to decrease confinement and increase exercise, which will help to decrease stress and anxiety15,19—often exhibited in rabbits as polydipsia, mutual and self-barbering, and carpet digging. The caregiver should provide interactive items that stimulate instinctive behaviors and decrease boredom. This can be as simple as offering a free choice grass hays, a high-fiber diet, and allowing for foraging behaviors by hiding food and treats for the rabbits to find; it can also be as elaborate as suggestions listed in Box 39-1, “Behavioral Enrichment,” which will help to stimulate the animals mentally. Providing a consistent schedule—including feeding, exercise, and a day/night cycle—will also decrease stress-related behavior problems. By providing pet rabbits with twice yearly veterinary exams and exams whenever subtle or overt behavior changes are noted, medical issues can also be ruled out as a cause of behavioral problems. As with children and all other species, it is important to divert their attention to acceptable behaviors9 (see “Behavioral Training Techniques for Small Mammals,” below). Rabbits are intelligent and can be very sensitive emotionally; they should be treated and respected as the individuals that they are.
LITTER-BOX TRAINING In puberty both males and females that were previously litterbox trained may begin to urinate and defecate randomly to mark their territory and by marking the caregivers’ personal items. To help to minimize this problem, redirect the behaviors (keep rabbit away from personal items, provide distractions and enrichment) and eliminate the need of the rabbit to be territorial. The most obvious choice would be to have the rabbit surgically altered to minimize hormonal influence. Make litterboxes more appealing with easier access.
CHAPTER 39 Behavior of Small Mammals In the wild, rabbits eliminate in specific areas, referred to as latrines or scrapes, which signal the territory of their warren.4,9,10 It is therefore best to place multiple litter boxes in the areas where the most frequent soiling occurs, as these are the sites that the rabbit will likely continue to use. Provide organic litters such as alfalfa, oat, citrus, or recycled paper products. It is often recommended to place a handful of hay in each box to encourage its use, as rabbits will often defecate as they sit and eat. Use white vinegar to clean the boxes and to remove any urine from inappropriate places. Additional information on the litter-box training of rabbits is available.4,9,10,14,19
INTRODUCTION OF NEW CONSPECIFICS Because rabbits are a highly intelligent and social species, it is very important to provide bonded mates, especially if the rabbit has limited social contact with human companions or other pets. Introducing a new rabbit should always be done slowly and carefully and with direct adult supervision. This is best accomplished by diverting the established rabbit’s attention to the owner, so that it does not become overly interested in the new rabbit, which is likely to be fearful. The supervisors should be prepared to separate the rabbits before they begin to fight, as rabbits can inflict serious injuries on each other. Because of their instinctual social behavior, a social hierarchy or pecking order must be established when rabbits that are new to each other are brought together; on such an occasion, some fighting and aggressive displays should be expected. Not all individuals, however, will get along well, and the caregiver should be prepared to house them separately for an extended period of time or possibly even permanently. For social, behavioral, and medical reasons, adult rabbits should be neutered or spayed before they are introduced, as mentioned previously. The House Rabbit Society recommends that at least 20 minutes per day should be dedicated to the bonding process.19 Introductions should be made in neutral territory; “coerced closeness”14,15,19 may help them to bond if they are placed together in situations that are new and slightly stressful to them (such as a car ride or being placed in a bathtub10) in order to create artificial situations where they are distracted from fighting. In such circumstances, as a prey species, they are more likely to be comforting or greeting each other in a more neutral fashion.10 Equal time should be allotted for less stressful situations, such as placing them in a large room where they can get away from each other and find places to hide15,16,20 (like a run-through cardboard tunnel or hide boxes in their own open cages) if necessary. Additional information on bonding rabbits can be obtained in other references.*
MOURNING THE DEATH OF A BONDED MATE Rabbits form very strong pair bonds, and the loss of cage mate or house mate can lead to what is perceived as grieving in the surviving mate. They will often eat less and become lethargic and polydipsic, which can be signs of stress. Some rabbits may isolate themselves, others demand more attention,
*References 4, 9, 10, 14, 15, 19.
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and some may engage in misbehavior, including digging and chewing.19 The caregiver should provide more support and pay extra attention (e.g., extra time, treats, and patience) to a rabbit that has lost its mate. Monitor the surviving rabbit closely to ensure that it is eating and drinking and producing an adequate amount of feces. The rabbit may need to have more physical contact with the owner and to be tempted to eat with favored food items. Although subjective, it is the author’s experience as well as that of many rabbit caregivers that it can be helpful to allow the surviving rabbit to “view” the dead mate’s body after it has passed or been euthanized, thus helping to shorten the bereavement period. If this is not done, the survivor will often continue to look for or anticipate the return of the lost mate. As a result, lethargy and depression may be extended. Because it is important for rabbits to have bonded mates, the owner, if possible, should try to provide the surviving rabbit with a new mate as soon as possible. This process takes time and patience, however, as not all rabbits will get along (see “Introduction of New Conspecifics,” above). Veterinary care should be sought if signs of grief are extensive or if the rabbit is not eating, not defecating, or losing weight. Other sources of information on grieving in rabbits are available.14,19 The foregoing suggestions are similar for other small mammal species (including guinea pigs, ferrets, sugar gliders, and chinchillas) that are social in nature and form strong pair bonds.
FERRETS SOCIAL BEHAVIORS In the wild, ferrets tend to be solitary. With domestication, however, they have become very social and do best when they are living with other ferrets. This is thought to be due to a juvenilization of existing natural behavior (the retention of juvenile behaviors or neotenization) that can occur with domestication in species that have been studied.9,30 Ferrets can live well alone, however, if they are given a lot of playtime and human interaction. Regular gentle handling is necessary for the socialization of young ferrets2; the sensitive period of socialization is from the time their eyes open at 4 weeks to about 10 weeks of age.11 Ferrets will also bond with other companion pets, but as with other species, introductions should be made slowly and with supervision (Fig. 39-1). Ferrets communicate through various vocalizations, postural changes, and scent marking.4,11 Common ways in which ferrets communicate are listed in Box 39-2.
PLAY AND SLEEPING BEHAVIORS In general ferrets are either in constant motion or sleeping. Intense play sessions will be intermittently interrupted by “slumping,” whereby the ferret will suddenly slump down flattened to the ground with eyes open and back legs splayed for several minutes and then as suddenly begin playing again. Ferrets spend about 12 to 16 hours a day sleeping4 and like to do so in warm, dark areas similar to their underground nests in the wild. A sleeping area can be as simple as soft clean cloths such as towels or old shirts to burrow under or elaborate hammocks, tents, hide boxes, and tubes. Excessive sleeping or difficulty in waking them may indicate a medical problem, such
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SECTION VI General Topics Box 39-3 Understanding Aggressive Play and Biting Behavior Motivations in Ferrets11
Fig. 39-1 Ferrets will readily bond with other companion pets, but their interactions should be supervised. (Courtesy of Amanda McGuire.)
• Play aggression—mostly seen in young ferrets that are overly aroused and have not learned that play immediately stops when they get too aggressive • Possessive aggression—to keep another ferret away from an object, food, or favored toy • Fear-related aggression—can be seen as a result of poor socialization or trauma • Redirected aggression—if ferrets are overly aroused and redirect their arousal or aggression to an object or person near them (e.g., when separating ferrets that are playing) • Maternal aggression—jill can be aggressive to protect kits • Pain-induced aggression or disease-induced aggression—can be seen when trauma or disease create pain • Predatory aggression—can be seen as a stalking, chasing, grasping, and biting sequence and reproduced in play • Sexual aggression—intense neck biting of intact ferrets and reproduced in play with intact or altered ferrets
Box 39-2 Examples of Play Behaviors of Ferrets11 • Exaggerated approach or ambush • Chase, veering off, and reciprocal chase • Mounting, rolling, wrestling • Inhibited neck biting accompanied by dooking (excitement) and hissing (anger) • Digging—which is derived from burrowing behavior • “Ferreting away”—objects of interest put away in small dark places
as hypoglycemia as seen in ferrets with insulinoma; in such instances they should be evaluated by a veterinarian. Ferrets love to play, and it is best to provide them with items that are safe if chewed on and that simulate the natural behaviors associated with hunting and burrowing. Avoid items that are made of rubber or foam rubber as well as any toys that contain fillers such as Styrofoam pellets, because ferrets love to chew on and ingest them, which can lead to gastrointestinal obstruction. Adolescent ferrets tend to be intensely playful and exuberant and will interact with each other by biting, mouthing, and even dragging each other by the scruff of the neck (Box 39-3). This behavior tends to be more intense in young males than in females.1 Ferrets often hop or gallop when playing. The “dance of joy” or “weasel war dance” consists of rough play whereby ferrets (especially young ferrets) will hop and spin wildly with backs arched, swaying their heads from side to side with mouths open and teeth bared, making a hissing noise. Although this can be frightening and misinterpreted as aggressive by new owners, it is an expression of joy and excitement. Like their wild counterparts, ferrets love to dig and “tunnel” and will burrow under clothing or towels and hide in holes in the furniture or in drawers (Fig. 39-2) and cabinets if left unsupervised. They also love to steal and stash objects that they find, similar to the stashing of food and prey items in their burrows in the wild.
Fig. 39-2 Ferrets love to get into small, dark places similar to their burrows in the wild. (Courtesy of Amanda McGuire.)
BEHAVIOR PROBLEMS Nipping and biting are the most common behavior problems noted in ferrets and are especially common in young animals. Ferrets tend to bite down, hold on, and shake their heads as they might in the wild with a prey item. Biting behaviors usually decrease at about 8 to 9 months of age if ferrets are handled gently. New owners, however, may inadvertently encourage or exacerbate the problem by handling them less or by overexuberant reprimands and mishandling when bitten. Often a ferret will lick its owner and then nip at him or her, which is a natural grooming behavior. It is therefore wise, with a new or young ferret, to proactively consider the licking behavior as a precursor to nipping and divert the animal’s attention with a distraction such as a toy or food treat. Interestingly, in a survey of ferrets that were gonadectomized after adulthood and then adopted out, behavioral issues were noted in 36% of the
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Box 39-4 Communication Behaviors of Ferretsa • Anal drag, wiping, body rubbing (provides information on gender and hormonal status) • Bark—loud chirp when fearful or excited • Hiss—fear, anger, frustration, warning • Sniffing—ferrets greet new ferrets by sniffing anal, neck, and shoulder area • Scream—high-pitched screech depicting fear or pain, also seen during seizures • Chuckling, buck—series of chortles depicting excitement, happiness (often referred to as “dook” by ferret owners and fanciers) • Piloerection of tail—indication of arousal which can be due to anger, fear, or excitement, also known as “brush tail”; often with arched back and a hiss or screech Fig. 39-3 Young ferrets are active chewers and biters; patience is needed to get them through adolescence. (Courtesy of Amanda McGuire.)
aFurther detail on ferret communication is available in references 2 and 11.
43 ferrets, including problems with nipping and failure to litter train, indicating that young ferrets are not the only ones with behavioral problems.13 Chewing and digging are also very prevalent among ferrets less than a year of age; ferrets should therefore be allowed access only to areas that have been ferret-proofed, and they should be supervised by an adult when they are outside of their enclosures (Fig. 39-3).
Managing Behavior Problems Behavior modification in the form of frequent gentle handling and passive correction such as distracting the ferret with a ferretsafe toy will help to redirect overly aggressive play behavior (see “Behavioral Training Techniques for Small Mammals,” below). Gentle handling may then be gradually introduced while the ferret is otherwise distracted. Behavior can also be modified by using a repeated firm warning, such as “Stop that!” or another sharp noise to distract the animal whenever biting behavior occurs. If the ferret does not heed the warning, then it is put in “time out” for a few minutes in a pet carrier devoid of towels or toys; at minimum, the social contact is discontinued to decrease the animal’s level of arousal. When this is done consistently, the ferret may begin to realize that biting decreases social interaction. Avoid provoking aggression or fear (Box 39-4). This includes not hitting ferrets, not provoking aggressive play with one’s hands, such as playing tug-of-war games, and not tapping them on the nose if they bite. It may be necessary to keep one’s fingers curled when handling a ferret. If bitten, some handlers will give a high-pitched yip that mimics what a litter mate might do during overly aggressive play, which causes play to stop. Another suggestion to try if a ferret bites is to make a low hissing sound, mimicking the behavior of a jill scolding a kit that is too aggressive, and then putting the ferret down. The ferret can also be placed in a towel for gentle cuddling and then given a food treat when it has calmed down. As with behavior modification with any species, it is important to be patient, consistent, and to end a play or training session on a positive note. It is imperative, therefore, that sessions should be relatively short (approximately 5-10 minutes). If a
Fig. 39-4 New places are exciting to ferrets; a tub is a great place to explore and good for mental stimulation and enrichment. (Courtesy of Amanda McGuire.)
negative behavior is exhibited, the owner should try to redirect the ferret to a more positive behavior and reward that behavior with a small food treat. Ending the session on a positive note such as this is helpful in reinforcing the desired behaviors. In managing behavioral problems in any species, it is important to provide mental stimulation and exercise and free-play time outside of the enclosure (Fig. 39-4). An extensive list of environmental enrichment ideas for ferrets can be found in Exotic Pet Behavior—Birds, Reptiles, and Small Mammals.11 Keep in mind that whenever a major change in behavior is noted (such as if a normally gentle ferret starts to bite) or a previously established behavior suddenly worsens, that it is wise to have a full medical workup performed. For instance, adult, sexually altered ferrets that become aggressive should be checked by a veterinarian for adrenal disease, because nipping, mounting, and even dragging other ferrets by the scruff of the neck may occur with the masculinization that can be seen with this disease process.18 If behavioral changes occur, however subtle, it is imperative that the ferret be examined to rule out any illness or disease process or a problem that might be causing pain.
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INTRODUCTION OF NEW CONSPECIFICS Although ferrets tend not to be overtly aggressive to each other in situations other than when at play, initial introduction of a new ferret can be challenging. Fighting is seen more often when new ferrets are introduced to older ferrets that have been housed alone for a period of time, a group of ferrets for which the social structure has been established for a while, and in same-sex ferrets.8 After initial sniffing of the anal area and then the neck, where scent glands are located, the ferrets will usually begin neck biting and fighting to establish dominance.11 A prospective trial using 56 ferrets showed that factors that help to determine levels of aggression between ferrets include gender, neutering status, and the level of familiarity of the ferrets.32 In the case of neutered ferrets that were not familiar with each other, it was found that the introduction of two males or a male and a female would likely result in the lowest levels of aggression.32 In this same study it was noted that caging ferrets next to each other for 2 weeks prior to introduction did not significantly alter the aggression exhibited when the ferrets were introduced,32 which is contrary to advice that is often given. In another study, however, where intact juvenile ferrets were used, it was determined that new male-to-male interactions were generally more aggressive than new female-to-female interactions, and that animals raised together exhibited less negative interactions overall.25 As seen with other species, it is recommended to introduce ferrets under direct adult supervision in a neutral space that is large enough to allow for escape if needed. Some handlers recommend putting Ferretone (Ferretone Skin and Coat Liquid, 8 in 1, Islandia, NY) on the backs of the necks of ferrets that are being introduced, as they often become so interested in the Ferretone that it distracts them from fighting while they investigate each other.11 Ferret shelter managers have found that introducing a new ferret to the most congenial ferret (usually an older laid-back male) in a group first and then slowly introducing the other established ferrets in the group facilitates introductions.11
LITTER-BOX TRAINING Ferrets prefer to urinate and defecate in corners and on vertical surfaces. They do not cover their urine or feces and will often perform an anal drag after defecating.11 Multiple boxes should be provided in preferred elimination areas. Boxes should be lowsided, with a small amount of litter, which should be of the pelleted variety or consist of shredded recycled paper. A litter box is best placed in the corner of a cage and wired into place to keep it from being overturned. Young ferrets are likely to dig and play in the litter, so paper strips may be used at first and litter added slowly to them in order to introduce the animals to the litter.2 Behavioral changes associated with litter training can be very significant and may signal medical problems such as enteritis, urinary tract infection, cystic calculi, adrenal disease, or prostatic disease. For a more extensive list of litter-box training tips the reader is referred to Exotic Pet Behavior—Birds, Reptiles, and Small Mammals.11
GUINEA PIGS SOCIAL BEHAVIORS In the wild, guinea pigs live in small groups of five to ten27 with a dominant male and female. They are docile and mostly affectionate pets. Since they are extremely social in nature, they are
Fig. 39-5 Guinea pigs are less stressed in the hospital exam room if they are handled quietly and gently and given a towel for traction, fresh hay and greens, and a place to hide.
Fig. 39-6 Guinea pigs can be trained to live in certain areas of the house without caging as long as they are provided with adequate food and water as well as hiding places. (Courtesy of Paul and Debbie Ladas.)
easily stressed when separated from bonded mates.3 It is important that guinea pigs be socialized to handling when they are young, otherwise they can be exceedingly shy and may stress so much as to injure themselves in attempts to escape handling (often referred to as stampeding3). They can also be stressed by loud noises in the home or hospital environment (Fig. 39-5) and tend to do better in the hospital if they are kept with bonded mates whenever possible.
PLAY BEHAVIORS Guinea pigs are very intelligent and playful and will initiate play with humans and other companion pets. Like rabbits, they will initiate chase games with each other and with humans. A guinea pig-safe area should be provided for at least several hours of exercise per day (Fig. 39-6), preferably in the morning or evening (when they are naturally most active) or both. Young guinea pigs will jump up or “popcorn” when excited.
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Fig. 39-8 Because chinchillas will lose their fur if grabbed by a Fig. 39-7 Grass can be organically grown for guinea pigs, rabbits, and chinchillas to provide for behavioral enrichment. (Courtesy of Rebecca Howell.)
COMMUNICATION BEHAVIORS Guinea pigs communicate using short impulse sounds including the chutter, chip, or purr and the slow long call or whistle.33 A more complete list of vocalizations and postural forms of communication in guinea pigs is available.3
BEHAVIOR PROBLEMS Although guinea pigs have few real behavior problems, occasionally you may be presented with a biter or one with barbering issues. As a species they are generally gentle and not usually prone to biting or scratching unless provoked or mishandled. Some individuals, however, will bite with unwanted handling, as for toenail trims. Others may nip gently (or not so gently) to encourage or discourage interaction with humans and other companion pets. As with other species, early socialization and frequent gentle handling help to decrease biting behaviors. Mutual or self-barbering can be quelled by decreasing stress, including that associated with overcrowding and boredom and by increasing environmental enrichment.3 Providing free choice grass hays and allowing for foraging (Fig. 39-7) and other natural behaviors will help to increase mental stimulation, and other forms of behavioral enrichment will increase exercise. Self-barbering, as in other species, can also be related to medical problems, which should be ruled out.
predator (known as “fur slip”), it is best to hold them at the base of the tail with one hand and support the chest with the other when they are being carried or examined in the clinic.
CHINCHILLAS SOCIAL BEHAVIORS Chinchillas live in large groups in the wild, with several hundred living in burrows and rock crevices. They are nocturnal in the wild24 but can acclimate to daytime activity in captivity; however, they will eat and defecate mainly at night. They tend to be quiet and can be shy if not well socialized to humans (Fig. 39-8); they are easier to socialize when they are young. Because of their social nature, it is best to house them in pairs or colonies, although they can do well as single pets if given a lot of human attention.
PLAY BEHAVIORS Chinchillas like to jump and climb and are very agile. They should be provided with multilevel cages or a chinchilla-safe room in which to play. They will often use a solid running wheel, which may need to be removed periodically to keep them from using it too much.21 They are mostly nocturnal and are normally less active during the day, so they should be allowed a quiet time during the day with no handling. Polyvinylchloride (PVC) pipes and cardboard boxes can be provided as well as other places to hide, and bird-safe toys (toys not easily chewed and ingested) and balls can be provided.
INTRODUCTION OF NEW CONSPECIFICS
COMMUNICATION BEHAVIORS
Intraspecies aggression can be seen when introducing new guinea pigs together and also when two intact males are in the presence of females.7 Many guinea pig owners have mistakenly put a newly acquired cavy in the enclosure of an already established guinea pig with dire consequences. Without a gradual, adult-supervised introductory period, they can inflict serious bite wounds on each other. Placing guinea pigs in a neutral area with ample distractions (hay, greens, carrots), room to escape, and hiding spots (boxes, tunnels, hide boxes) for gradually increasing lengths of time can be helpful. Expect a lot of rumbling and chortling noises and “rumba” type movements, and be prepared to separate them quickly or distract them before they begin to fight.
Chinchillas communicate with each other by soft, highpitched grunting noises.27 If they are startled by loud noises, they will often stand on their hind limbs and elicit a warning cry, which is a whistling sound. They will honk for a treat or to elicit attention. Hissing will usually be a precursor to fighting.
BEHAVIOR PROBLEMS Chinchillas are usually not aggressive, rarely bite, tend to use flight as their defense mechanism, and normally do not like to be held for extended periods.21 When they bite, however, the
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bite is very sharp and strong. Intraspecies aggression is most likely seen during the reproductive season and during introductions, where barking, chasing, and biting are common and serious injuries can occur.21 Females tend to be the more aggressive sex and have been known to stand up and spray urine at a potential or perceived threat.21 They will make a barking noise if they become agitated or angry, may growl, and chatter, and scent glands near the anus may be expressed when they are frightened. Fear-related and abnormal behaviors often arise because of stress and lack of social interaction. Perceived or actual predators and lack of places to hide can also lead to stress. Chinchillas are also easily stressed by loud noises and external commotion; they should be kept in a room apart from where family activity occurs. Excessive stress can cause anorexia, gastrointestinal stasis, barbering, and failure to thrive.
INTRODUCTION OF NEW CONSPECIFICS Severe and even fatal injuries may occur if introductions are not made slowly and with adult supervision. Caging in close proximity is recommended for a week or more before introductions are made. Camphor or other odiferous substances can be applied to each chinchilla prior to introductions; many handlers will create a form of “coerced closeness”19 by placing them together for about 15 minutes in a cage small enough that they can’t fight and then allowing them together in a larger, neutral space with two dust baths and two hide boxes.21
Fig. 39-9 Sugar gliders should be provided with solid exercise wheels; when given the ability to play and explore, they are active at night. (Courtesy of Tracy Brandow.)
SUGAR GLIDERS SOCIAL BEHAVIORS Wild groups of sugars gliders are arboreal and nocturnal and nest in colonies of up to seven adult males, females, and their young; such groups are exclusive and territorial.23 Because of their highly social nature, it is best to keep them in groups of two or more and allow for a socialization period with people of at least 2 hours or more per day, preferably at night when they are most active.26
Fig. 39-10 Sugar gliders can jump long distances with the aid of their gliding membranes. (Courtesy of Tracy Brandow.)
COMMUNICATION BEHAVIORS Sugar gliders have a complex system of chemical communication based on scents produced in the glands of the skin and, in the female, in the pouch.23 The scents identify the group and its territory. The dominant male actively marks the others in the group, and urine is also used to mark territory.23 Sugar gliders are very vocal and make a variety of yapping, barking, buzzing, droning, hissing, and screaming sounds.23 They may make a crying sound when they are separated from cage mates. They will make a purring sound when they feel content, and females with young will make a singing sound. They will chatter quietly for attention and make a “crabbing” sound when they are irritated or angry.23 If frightened, they will produce a white, oily secretion from their paracloacal glands, which has the odor of soured fruit.23 Sugar gliders will assume a defensive posture by standing on their hind legs with head extended, often with the mouth open. If really frightened, they may lie on their backs with their feet up in the air while vocalizing.
PLAY BEHAVIORS As a nocturnal species, sugar gliders are most active in the early evening and throughout the night (Fig. 39-9). They like to glide and jump and, as an arboreal species, to be in high places (Fig. 39-10). During the day they will spend time in their nesting sites or in pouches often carried by their owners around their necks.
BEHAVIOR PROBLEMS As in the case of other small mammals, it is important to provide mental stimulation, behavioral enrichment in the form of foraging for food, bird-safe toys, nontoxic tree branches, perches, slings, and solid exercise wheels. Sugar gliders are the most likely species to self-mutilate in times of stress, anxiety, and pain (including pain from trauma and surgery); these issues must be addressed proactively and aggressively.
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INTRODUCTION OF NEW CONSPECIFICS Sugar gliders are very territorial and will be aggressive to new cage mates unless gradual supervised introductions are made during the day, when they are less active. It can be helpful to place cages near each other and trade nest boxes at night, thus allowing the animals to become familiar with each others’ smells prior to introduction.23
HEDGEHOGS SOCIAL BEHAVIORS African pygmy hedgehogs (Fig. 39-11) are mostly solitary and territorial and usually kept as single pets or maintained in separate enclosures.22 They tend to be shy and reclusive, especially if not well socialized, and should be provided with hide boxes.22
COMMUNICATION BEHAVIORS A mostly quiet species, hedgehogs make squeaking and hissing sounds during courting and will communicate with chattering, snorting, and snuffling noises.24 They will nudge each other and human companions to gain attention. When frightened, they will sniff and cough and even scream.22 Hedgehogs erect the spines on their head or curl up when they are feeling defensive.
PLAY BEHAVIORS Hedgehogs tend to eat and explore more at night, but they can easily be trained to interact more during the day. They like to swim and climb and will spend hours on a solid exercise wheel.22 Given plenty of environmental enrichment items, including cardboard toilet paper rolls and large bird-safe toys, they will spend a lot of time playing.
BEHAVIORAL TRAINING TECHNIQUES FOR SMALL MAMMALS The following methods can be used with small mammals, especially with rabbits and ferrets, in order to help to train them using positive reinforcement. These methods also help to stimulate them mentally, provide avenues of exercise, reduce stress, and increase quality of life. Clicker Training.10,28,31 This technique is used to make the connection between doing something, a click, and a reward (e.g., a food treat). This method has been used to help tame feral rabbits, help rabbits in shelters become more adoptable, and help pet rabbits and ferrets learn tricks. Targeting.31 Like other species, many small mammals can be taught to touch a target with their noses as a method of behavior modification to distract or to redirect behavior. While targeting usually involves training a pet to touch a place or object with its nose, a caregiver can also train a rabbit or ferret to touch or follow a stick or spoon. Examples of how this can be used include having them touch a spot repeatedly on command to focus the animal’s attention away from a source of aggression, as when cleaning a cage or replacing food bowls. Rabbits can also be taught to do some agility-type skills by following a target. As with the training of other species, the pet is rewarded with a food treat when it touches or follows the desired target.
Fig. 39-11 To examine the underside of a hedgehog (to check an incision, for example), the hedgehog can be placed in a seethrough container and held above the examiner’s head. (Courtesy Parrish Antoine, RVT.)
Desensitization and Counterconditioning.35 Desensitization allows the pet to slowly become accustomed to low level stimuli that previously produced a negative response. This is implemented by gradually increasing the stimulus, or moving it closer, over time so that the pet does not react fearfully or barely notices the stimulus. For the pet to get used to it, the stimulus is always presented at a level below the animal’s fear threshold. This is a process that requires a lot of patience, whereby a ferret or rabbit is rewarded for allowing the caretaker to get closer and closer to the object that stimulates aggression or fear, such as a food bowl or toy, or to allow petting or picking the animal up. Desensitization is most effective when used with counterconditioning. For example, with classic counterconditioning, food treats are used so that the pet associates the stimulus with something positive, so that the stimulus becomes less fearprovoking and eventually elicits a positive emotional response. With operant counterconditioning, the pet is trained to perform an alternative behavior when the stimulus is present and then is rewarded for that behavior. Because the pet is rewarded, it is likely to develop a positive emotional response to the stimulus.
CONCLUSIONS The most important thing that clients can do for their small mammal pets is to allow for natural behaviors by providing suitable behavioral enrichment. Practitioners can decrease stress and anxiety in small mammal patients by catering to their natural behaviors whenever they are hospitalized and by providing multimodal and preemptive pain management. In any species, whenever a new behavior is noted or a previously established behavior suddenly worsens, it is wise to have a full medical workup performed. Even subtle behavior changes can indicate medical problems. This is especially true in small mammals that are prey species. This chapter is not comprehensive for behavior in general or the behavior of all small mammal species. For more detailed information on behavior-related topics in small mammals, the reader may refer to Exotic Pet Behavior—Birds, Reptiles and Small Mammals.3,4,11,21-23 When managing behavioral problems in any
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species, the author recommends using only positive reinforcement techniques and following guidelines recommended by the American Veterinary Society of Animal Behavior (www.avsabon line.org).
ACKNOWLEDGMENTS The author wishes to express gratitude to Sophia Yin, MS, DVM, for her review of and suggestions for this chapter.
References 1. Baum MJ. Use of the ferret in behavioral research. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Philadelphia: Lipp incott, Williams & Wilkins; 1998:511-520. 2. Boyce SW, Zingg BM, Lightfoot TL. Behavior of Mustela putorius furo. Vet Clin North Am Exot Anim Pract. 2001;4:697-712. 3. Bradley Bays T. Guinea pig behavior. In: Bradley Bays T, Lightfoot T, Mayer J, eds. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: Elsevier; 2006:207-238. 4. Bradley Bays T. Rabbit behavior. In: Bradley Bays T, Lightfoot T, Mayer J, eds. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: Elsevier; 2006:1-49. 5. Bradley T. Rabbits: understanding normal behavior. Exot DVM. 2000;2:19-24. 6. Bradley TA. Rabbits: medical implications of selected abnormal behaviors. Exot DVM. 2000;2:27-31. 7. Bradley TA. Normal behavior and clinical implications of abnormal behavior in guinea pigs. Vet Clin North Am Exot Anim Pract. 2001;4:681-696. 8. Church B. Ferret-polecat domestication: genetic, taxonomic and pylogenetic relationships. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. 2nd ed. New York: WB Saunders; 2007:122-150. 9. Crowell-Davis SL. Rabbit behavior. Compend Vet Med. 2006;28:715-718. 10. Crowell-Davis SL. Behavior problems in pet rabbits. J Exot Pet Med. 2007;16:38-44. 11. Fisher P. Ferret behavior. In: Bradley Bays T, Lightfoot T, Mayer J, eds. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: Elsevier; 2006:163-206. 12. Hansen LT, Berthelsen H. The effect of environmental enrichment on the behavior of caged rabbits (Oryctolagus cuniculus). Appl Anim Behav Sci. 2000;68:163-178. 13. Harms CA, Stoskopf MK. Outcomes of adoption of adult laboratory ferrets after gonadectomy during a veterinary student teaching exercise. J Am Assoc Lab Anim Sci. 2007;46:50-54. 14. Harriman M. House rabbit handbook: how to live with an urban rabbit. 4th ed. Alameda, CA: Drollery Press; 2005. 15. Harriman M. Introducing rabbits—bonding techniques for matchmakers. 2nd ed. Alameda, CA: Drollery DVD; 2008. 16. Hayes RA, Richardson BJ, Claus SC, et al. Semiochemicals and social signaling in the wild European rabbit in Australia: II. Variations in chemical composition of chin gland secretion across sampling sites. J Chem Ecol. 2002;28:2613-2625.
17. Hayes RA, Richardson BJ, Wyllie SG. To fix or not to fix: the role of 2-phenoxyethanol in rabbit, Oryctolagus cuniculus, chin gland secretion. J Chem Ecol. 2003;29:1051-1064. 18. Hoefer HL. Small mammal behavior. Proceedings. Atlantic Coast Veterinary Conference. 2001. 19. House Rabbit Society Web site. Rabbit behavior. Available at http://www.rabbit.org. Accessed July 15, 2008. 20. Jenkins JR. Rabbit behavior. Vet Clin North Am Exot Anim Pract. 2001;4:669-679. 21. Johnson DH. Miscellaneous small mammal behavior—chinchillas. In: Bradley Bays T, Lightfoot T, Mayer J, eds. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: Elsevier; 2006:263-280. 22. Johnson DH. Miscellaneous small mammal behavior—hedgehogs. In: Bradley Bays T, Lightfoot T, Mayer J, eds. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: Elsevier; 2006:293-304. 23. Johnson DH. Miscellaneous small mammal behavior—sugar gliders. In: Bradley Bays T, Lightfoot T, Mayer J, eds. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: Elsevier; 2006:328-344. 24. Lightfoot TL. Clinical examination of chinchillas, hedgehogs, prairie dogs and sugar gliders. Vet Clin North Am Exot Anim Pract. 1999;2:447-469. 25. Lode T. Kin recognition versus familiarity in a solitary mustelid, the European polecat Mustela putorius. C R Biol. 2008;331:248-254. 26. Ness R, Booth R. Sugar gliders. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2004:330-338. 27. Nowak RM, ed. Walker’s mammals of the world. 6th ed. Vol I. Baltimore: John Hopkins Press; 1999. 28. Orr J. What? Train a rabbit? Available at www.clickertraining. org. Accessed Feb 24, 2009. 29. Parer I. The population ecology of the wild rabbit (Oryctolagus cuniculus) in a Mediterranean-type climate in New South Wales. Aust Wildl Res. 1977;4:171-205. 30. Price EO. Animal domestication and behavior. New York: CABI Publishing; 2003. 31. Pryor K. Clicker training bunnies gain confidence. Available at www.clickertraining.org. Accessed Feb 24, 2009. 32. Staton VW, Crowell-Davis SL. Factors associated with aggression between pairs of domestic ferrets. J Am Vet Med Assoc. 2003;222:1709-1712. 33. Suta D, Kvasnak E, Popelar J, et al. Representation of speciesspecific vocalizations in the inferior colliculus of the guinea pig. J Neurophysiol. 2003;90:3794-3808. 34. Whary M, Peper R, Borkowski G, et al. The effects of group housing on the research use of the laboratory rabbit. Lab Anim. 1993;27:330-341. 35. Yin S. Low stress handling, restraint and behavior modification of dogs & cats. Davis, CA: Cattledog Publishing; 2009.
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40
Zoonotic Diseases
Mark A. Mitchell, DVM, MS, PhD, Diplomate ECZM (Herpetology), and Thomas N. Tully, Jr., DVM, MS, Diplomate ABVP (Avian), Diplomate ECZM (Avian)
Bacterial Diseases Viral Diseases Parasitic Diseases Mycotic Diseases Allergic Reactions
Nontraditional small mammals are popular as companion animals. The most common species treated by veterinarians include rabbits (Oryctolagus cuniculus), ferrets (Mustela putorius), mice (Mus musculus), rats (Rattus norvegicus), chinchillas (Chinchilla lanigera), golden hamsters (Mesocricetus auratus), Mongolian gerbils (Meriones unguiculatus), guinea pigs (Cavia porcellus), African pygmy hedgehogs (Atelerix albiventris), and sugar gliders (Petaurus breviceps). Small mammals are relatively inexpensive pets that often do not require the intensive care or attention that is required of dogs and cats. Because of these perceived advantages, ferrets, rabbits, and rodents are frequently purchased as first pets for children. Unfortunately, most pet owners are unaware of the zoonotic risks associated with owning nontraditional pets. However, this also extends to medical and veterinary professionals, who are also often unfamiliar with the common zoonotic agents associated with these animals. Specific reports of zoonotic diseases attributed to nontraditional species are rare. However, because of their increasing popularity and the amplified exposure to children and individuals with compromised immune systems (e.g., those who have undergone organ transplantation or immunosuppressive drug therapy or who have immunosuppressive diseases), the incidence of zoonotic diseases correlated with small mammal ownership may increase. Most disease conditions associated with nontraditional mammalian pets are related to allergies, bites, and scratches. Dander from these nontraditional species can cause cutaneous and respiratory allergies in susceptible individuals. Many bite and scratch wounds inflicted by these pets can develop into bacterial or viral infections. Pet owners can avoid bite injuries if they understand the behavior of their pets. These animals Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
should be primarily handled during peak activity hours. Certain rodents, such as hamsters, will assume a defense posture by rolling on their backs and will vocalize to ward off any perceived dangerous advances from humans. Most bite injuries are related to nocturnal/crepuscular species, such as hamsters, and can be avoided if owners are educated as to the most appropriate times to handle their pets. This chapter focuses on the zoonotic potential of disease transmission to the typical companion animal owner. Individuals who work with large collections of ferrets, rabbits, and rodents in laboratory animal settings or breeding facilities have a higher potential for exposure.
BACTERIAL DISEASES Bordetella bronchiseptica is a gram-negative rod to which high morbidity and mortality rates in guinea pigs have been attributed (Fig. 40-1).3 Hamsters and mice do not appear to be as sensitive to this pathogen as other rodents.52 Bordetella bronchiseptica can be isolated from clinically normal rodents and lagomorphs. Clinical signs in affected rodents and lagomorphs are generally associated with the respiratory system and are characterized by nasal and ocular discharge, coughing, sneezing, and pneumonia. This pathogen has been associated with respiratory tract infections in humans. Although rodents and lagomorphs pose minimal risk in the dissemination of this bacterium to humans, at-risk individuals should take specific precautions to reduce exposure. Pet dogs are a greater risk to humans than nontraditional mammals for B. bronchispetica through increased mouth-to-face contact. Campylobacteriosis is a serious zoonotic disease responsible for the annual expenditure of millions of dollars in medical expenses. Although contaminated food (e.g., poultry) and water are the primary methods of exposure to Campylobacter species in the United States, infected pets (e.g., dogs, cats, horses, hamsters) may also be a source of this pathogenic bacterium.39 A very small percentage of the nearly 2 million confirmed cases a year of campylobacteriosis in the United States are attributed to pet exposure.39 While nontraditional small mammals may harbor Campylobacter jejuni or C. pylori, of the 557
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Fig. 40-1 Congested lungs removed from a guinea pig that died from infection with Bordetella bronchiseptica. (Courtesy of Bruce Williams, DVM.)
risk that they will transmit this organism to their owners is low. A pet animal with diarrhea is considered a risk factor for shedding of Campylobacter species, which resulted in 6.3% of the cases of campylobacteriosis reported in a 1993 study.44 Incubation of Campylobacter species is approximately 2 to 5 days, and the disease is generally self-limiting (10-14 days) in the immunocompetent host.19 Ferrets that have recovered from campylobacteriosis may shed the bacteria for more than 140 days.19 A 9-month bacteriologic survey of biomedical research ferrets suggests that ferrets may pose a special threat to individuals working with ferret colonies. In this study, C. jejuni and C. coli were isolated from 18% of the ferrets being tested.19 Although no cases of ferret-associated campylobacteriosis have been reported, the prevalence of Campylobacter species in these ferrets suggests that they may serve as a source of infection. When infected, humans usually suffer from a mild, self-limiting gastroenteritis, but the disease may develop into a generalized septicemia that can result in death. Corynebacterium kutscheri is a gram-positive bacillus that is routinely isolated from the nasopharynx of rats. This microbe has also been isolated from clinically normal mice and hamsters. Most rodents that harbor this microbe are asymptomatic; however, acute clinical disease has been reported in susceptible animals. Affected rodents are generally in poor condition and may develop cutaneous abscesses, nasal and ocular discharge, and arthritis. Because this pathogen has been isolated from a human case of chorioamnionitis, rodents should be considered a potential reservoir for the disease. Francisella tularensis is the causative agent of rabbit fever, or tularemia. This gram-negative coccobacillus is primarily a pathogen of wild rodents and lagomorphs. Captive rodents and lagomorphs generally are not considered important reservoirs for this microbe. The domestic cat is actually more likely to serve as a reservoir than other domestic species because of possible contact with infected wildlife (Fig. 40-2). Francisella tularensis can be transmitted by aerosolization, direct contact, or ectoparasites (fleas and ticks). Affected individuals may develop cutaneous lesions, respiratory disease, or meningitis. Leptospira interrogans is frequently isolated from wild rodents. Individuals working with wild rodents or in areas where high densities of wild rodents are concentrated may be at risk of contracting leptospirosis. Leptospira interrogans, L. grippotyphosa, and L. icterohaemorrhagiae have been isolated from ferrets. Individuals who have contact with ferrets used for hunting may also
Fig. 40-2 Spleen and internal organs of beaver that died from infection with Franciella tularensis. (Courtesy of Bruce Williams, DVM.)
be predisposed to leptospirosis. The bacteria are shed in the urine, and transmission may occur from direct contact or aerosolization of urine-contaminated food, water, bedding, or soil. Affected individuals may have chills, fever, and malaise or septicemia and generalized organ dysfunction. Although nontraditional small mammals kept as pets are not generally considered to be a source of L. interrogans for humans, these animals can serve as reservoirs for the spirochete, and appropriate precautions should be taken to prevent exposure. The incubation period for a human exposed to Leptospira organisms until infection occurs is 2 days to 4 weeks.6 The first signs associated with a human infection are a sudden fever, chills, headaches, vomiting, myalgia, and/or diarrhea.6 The human patient may recover from the initial disease condition and have no other problems from the infection. If more severe disease problems develop after the initial recovery, this is also called Weil’s disease (i.e., the development of kidney or liver failure or meningitis). Listeria monocytogenes is a gram-positive, non-spore-forming rod. Ferrets may serve as a source of the Listeria organism because it has been isolated from the liver and lung tissue of infected animals. Listeriosis was also identified in ferrets experimentally infected with canine distemper virus. The common finding in many cases is that the infected ferrets were immunocompromised animals. Because adrenal gland disease is a common finding in geriatric ferrets and L. monocytogenes is ubiquitous in the environment, caretakers should take appropriate precautions to minimize their animals’ exposure to this pathogen. Ferrets are extremely sensitive to mycobacteriosis. Mycobacterium avium, M. bovis, and M. tuberculosis have all been isolated from ferrets. Most mycobacteriosis cases reported in ferrets are from Europe and New Zealand, where ferrets are used for hunting and are often fed uncooked meat and meat by-products. Mycobacterium species can be transmitted to humans from ferrets through aerosolization of contaminated respiratory secretions or direct skin contact. Ferret owners should be instructed to offer only cooked meat to their animals, thereby reducing the likelihood of introducing pathogens to their colony. Pasteurella multocida, a gram-negative rod, is an opportunistic pathogen of lagomorphs and rodents. In rabbits, P. multocida can cause multisystemic disease and is difficult to eradicate. Because this organism is ubiquitous in the environment, inoculation into bite injuries or scratches received from lagomorphs or rodents can occur. In humans, P. multocida has also been
CHAPTER 40 Zoonotic Diseases
Lymphadenopathy
2
Rash
3
1
Fig. 40-3 Rat-bite fever in humans is initiated with a bite that compromises the protective epithelial barrier. 1, Entry wound (rat bite); 2, spread of infection; 3, disease ulceration at entry site.
associated with multisystemic disease. Susceptible individuals, including young children and immunocompromised adults, should practice strict hygiene after handling these animals. Bite or scratch injuries should be immediately disinfected and a physician contacted for additional information for wound management. Pseudomonas aeruginosa, a gram-negative bacterium, is ubiquitous in the environment. While many exotic small animals are considered to be commensal hosts, few studies have characterized the prevalence of this opportunistic pathogen. A 2010 study in chinchillas reported the prevalence of P. aeruginosa to be nearly 42% in animals maintained as pets or in the laboratory setting.27 The P. aeruginosa isolated from chinchillas showed patterns of resistance to antibiotics commonly used to treat these animals in captivity. Because this organism is a common cause of nosocomial infections in humans, routine disinfection methods should be practiced after handling or cleaning small mammals and their cages to minimize the potential of exposure to this organism. Rat-bite fever (RBF) is a rare but serious zoonosis caused by Streptobacillus moniliformis and Spirillum minus (Fig. 40-3). These gram-positive microbes are considered indigenous flora in rats and may be isolated from the nasal and oropharyngeal areas of these animals.40 Laboratory and pet rats are considered the primary reservoir for RBF, although wild rats and mice may also serve as competent reservoirs. These bacteria are primarily transmitted from a bite. The incubation for streptobacillary RBF in humans is approximately 3 to 10 days, whereas it is 1 to 6 days for spirillary RBF. The clinical signs attributed to RBF are generally observed at approximately the same time the bite wound is healing; they include relapsing fever, chills, vomiting, myalgia, and regional lymphadenopathy. Individuals infected with S. moniliformis frequently have a maculopapular rash on their extremities. This rash does not occur with spirillary RBF. In 2003, two cases were reported of previously healthy adult women dying from S. moniliformis septicemia 2 to 3 days
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after the initial onset of disease symptoms (e.g., headache, diarrhea, lethargy, myalgia, lymphadenopathy).23 In another case reported in 2004, a 24-year-old male pet shop employee died after becoming infected through a superficial skin wound that occurred from contact with a contaminated rat cage.47 Clinical signs were similar to those described for the women in the earlier report, although endocarditis and multiorgan failure ultimately led to the demise of the male patient. Rat-bite wounds should be immediately disinfected to reduce the likelihood of transmitting these pathogens, and care should be taken in working around enclosures with positive animals. Affected individuals should seek medical attention and report their exposure history.40 Physicians should be informed of a patient’s exposure to a rat if an owner is being examined for an unexplained febrile illness or sepsis.23,47 Although the disease is called rat-bite fever, approximately 30% of the patients confirmed with RBF have not reported being bitten or scratched by a rat.23,47 Wearing gloves, handwashing after handling a rat, and avoiding hand-to-mouth contact with the animal or when cleaning its cage will help reduce the risk of being infected with RBF or other potential rodent-transmitted zoonotic infections.40 Salmonella species, which are gram-negative facultative anaerobes, are ubiquitous in the environment. This microbe has been associated with both enzootic and epizootic outbreaks in captive rodent and lagomorph colonies.14,24,38,48 Salmonella have also been isolated from other nontraditional small mammals, including sugar gliders, African hedgehogs, and ferrets.52 All vertebrate species should be considered susceptible to salmonella infection. There are more than 2,400 different Salmonella serotypes and all are potentially pathogenic, although most infections in humans and animals are caused by less than 12 serotypes. Salmonella enterica ser. Enteritidis (S. enteritidis) and S. enterica ser. Typhimurium are the two most common serotypes isolated from rodents and lagomorphs, whereas S. enterica ser. Tilene appears to be the most common serotype isolated from sugar gliders and hedgehogs.26,53 S. enterica serovars Typhimurium, Hadar, Kentucky, and Enteritidis have been isolated from ferrets.26,53 Most rodents and ferrets appear to be asymptomatic reservoirs for this microbe. One exception is the guinea pig, which can develop life-threatening septicemia. Salmonella infections are primarily transmitted by the fecal-oral route or from contaminated fomites, but they can also can be transmitted by aerosol through the conjunctiva in some species.35 Affected vertebrates may have anorexia, weight loss, enteritis, lymphadenopathy, or septicemia. Reproductive disorders, including abortions and premature birth, have also been associated with Salmonella infections in rodents and lagomorphs. Humans who contract salmonellosis from these animals may have headaches, nausea, vomiting, abdominal pain, enteritis, or septicemia.50 The incubation period in humans is approximately 6 to 48 hours. Humans can acquire Salmonella from either direct or indirect exposure to mammal reservoirs. In 1994, a case of hedgehog-associated salmonellosis in a 10-month-old girl from Washington was attributed to indirect contact with the family’s breeding colony of hedgehogs.30 From December 2003 through October 2004, there were 28 reported cases in the United States of multi-drug-resistant S. enterica ser. Typhimurium matched by pulsed-field gel electrophoresis to isolates from rodents (e.g., hamsters, mice) purchased at retail pet stores.49 The median age of those affected was 16, and a large number of affected individuals (40%) were hospitalized. In addition to having been isolated from humans and rodents, the organism was also
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found in rodent cages and transport boxes. Veterinarians should remind clients to be aware of these risks when they are cleaning their pet rodents and cages. Frozen rodents intended as food for reptiles have also been found to serve as a potential zoonotic source of S. enterica ser. Typhimurium.21 Because of the inherent zoonotic risks associated with the ownership of these and other pets, pet ownership should always involve adult supervision. Strict hygiene, including handwashing with soap, should be practiced to reduce the risk of exposure. The recommendations for reducing the risk of reptile-associated salmonellosis formulated by the Association of Reptile and Amphibian Veterinarians and the Centers for Disease Control and Prevention would also be appropriate to distribute to owners of nontraditional small mammals.4 Streptococcus pneumoniae, a gram-positive coccus, has been isolated from several different mammalian species. However, the rat and guinea pig appear to be the most susceptible to this pathogen. Several different serotypes affect rats (II, III, VIII, XVI, and XIX) and guinea pigs (III, IV, and XIX); some of these serotypes can also infect humans. Transmission of this pathogen is generally through aerosolization of contaminated respiratory secretions or by direct contact. Affected rodents may have pneumonia and torticollis. Humans infected with S. pneumoniae may have respiratory and meningeal disease. There have been three known major historic pandemics of plague. Yersinia pestis, the causative agent of plague, is generally transmitted by flea-borne vectors, although it can also be spread by contaminated respiratory secretions with the pneumonic form and direct contact with infected tissues or fluids from handling sick or dead animals. The three forms of disease associated with Y. pestis infection in humans are (1) bubonic plague (enlarged sensitive lymph nodes, fever, chills), (2) septicemic plague (fever, chills, prostration, shock, and bleeding into the skin and other organs), (3) pneumonic plague (fever, chills, cough and difficulty breathing, rapid shock and death if not treated early).7 The primary vector for Y. pestis is the rodent flea (Xenopsylla cheopis), although Thrasis, Diamanus, Hoplopsyllus, and Odropsylla species and Orchopeas sexentatus are also competent vectors. In the United States, Y. pestis is primarily a problem in southwestern states (e.g., New Mexico, Arizona, Colorado, California) and may be spread by domestic species. Because captive nontraditional pet mammals (e.g., chipmunks, prairie dogs) can also serve as competent hosts for many of the vectors reported for this pathogen, precautions should be taken to prevent the introduction of flea vectors into the home. Although there are no reported cases of plague associated with nontraditional pets in the United States, guinea pigs have been associated with plague in Andean countries, where they are raised for food.42 Yersinia pseudotuberculosis and Y. enterocolitica, opportunistic gram-negative coccobacilli, are routinely isolated from clinically normal rodents and lagomorphs. Y. pseudotuberculosis has been associated with epizootics in guinea pig colonies worldwide, in rats from Japan, and in hares from Europe.2 Affected animals generally suffer from weight loss and enteritis. Mortality rates can approach 75%. Wild rodents are considered to play an important role in the transmission of this pathogen. Although captive rodents have not been identified as an important source of infection in humans, they certainly could serve as competent reservoirs of the bacteria in the human environment. Yersinia enterocolitica has been associated with epizootics in chinchillas from Europe, Mexico, and the United States.2
Septicemia was a common finding, and affected animals had enteritis and weight loss. Studies evaluating the prevalence of enterocolitic yersiniosis in wild rodent populations from the Czech Republic, Slovakia, Scandinavia, and Chile suggest that the prevalence of this pathogen in wild rodents is low (4%26%). Although pathologic lesions, including abscesses of the liver and intestinal lesions, have been reported in wild rodents, most such animals screened for this pathogen are negative. The biotypes characterized in rodents are not the same as those that infect humans. Although rodents are not considered an important source of enterocolitic yersiniosis in humans, veterinarians and animal caretakers should practice strict hygiene and take appropriate precautions in working with infected rodents and lagomorphs.
VIRAL DISEASES Type A and B influenza viruses from the family Orthomyxoviridae are common pathogens of both ferrets and humans (see Chapter 6). These viruses can be transmitted between humans and ferrets by aerosolization. The disease is characterized in both humans and ferrets by a mucopurulent nasal and ocular discharge, sneezing, lethargy, fever, and inappetence. Pneumonia can develop in immunocompromised individuals. The influenza virus is apparently widespread in some ferret colonies. To reduce the likelihood of transmission of the virus between the two hosts, infected ferrets should be isolated from humans and vice versa. Ferrets are so susceptible to the same influenza viruses that infect humans that they are commonly used as laboratory animal models in developing influenza vaccines.51 In October 2009, a ferret tested positive for the H1N1 virus after exhibiting flu-like clinical signs.17 The owner of the ferret had suffered from flu symptoms before the ferret became ill. The ferret’s nasal secretions were tested at an Oregon State University diagnostic laboratory, where the genetic markers for the strain of H1N1 influenza virus were identified.17 Although uncommon, other small exotic mammal species may be infected with influenza viruses. In 2009, H5N1, a highly pathogenic avian influenza virus, was isolated from wild pikas in China.54 Hantavirus infections have been associated with epidemics in the United States, Japan, and Eurasia. There are at least 30 hantaviruses worldwide (genus Bunyavirus), many of which can cause a severe, often fatal illness in humans. This virus is transmitted through direct contact and the aerosolization of infected rodent urine, droppings, and saliva. Infected rodents are generally asymptomatic. Although this virus is generally associated with wild rodents, outbreaks have also been attributed to laboratory rodents in Europe and Japan. Infected humans reported flu-like symptoms, fever, myalgia, and oliguria. The Sin Nombre hantavirus was isolated from the Four Corners region of the southwestern United States in 1993.46 This virus caused an acute respiratory disease, subsequently named hantavirus pulmonary syndrome, in humans.46 From 1993 to 1997, infection of otherwise healthy individuals with the Sin Nombre hantavirus had a mortality rate of 45%.34 Rodents should be screened for the virus before they are incorporated into a colony. Lymphocytic choriomeningitis (LCM) is caused by an RNA arenavirus. The mouse is the primary reservoir for it, although the hamster and guinea pig are also competent reservoirs. This virus is transmitted by horizontal and vertical routes in rodents.2 Affected rodents are generally asymptomatic, although clinical disease—including weight loss, photophobia, tremors, and
CHAPTER 40 Zoonotic Diseases convulsions—may occur.26 Humans may become infected with the LCM virus from exposure to contaminated feces, urine, aerosolized dust or droplets, or nesting material or from a bite. People with normal immune systems may have no symptoms with LCM viral infection.31 Affected humans frequently report clinical symptoms consistent with a flu, including malaise, headaches, fever, myalgia, and arthritis. Pregnant women who become infected with LCM virus can pass the infection to their fetuses.31 Fatal aseptic meningitis or meningoencephalitis is rare. In a serologic study, about 5% of adults had a positive blood test showing that they were infected with LCM virus at some time in their lives.8 In 1973-1974 epidemic of LCM with more than 181 human cases reported from 12 states, morbidity was associated with hamsters being produced by a single facility in Birmingham, Alabama.9 Humans working closely with large populations of domestic rodents or wild rodents should practice strict hygiene and wear protective clothing to reduce the likelihood of exposure. Although the risk of infection from LCM virus is low, any woman who is pregnant or planning to become pregnant should avoid contact with rodents, including pet rodents (e.g., hamsters, guinea pigs) and rodent droppings.31 Rabies is caused by a negative-strand RNA lyssavirus. All mammals should be considered competent reservoirs for this virus; however, most (more than 90%) reported cases are associated with wildlife.36 Rabies has been isolated from ferrets, lagomorphs, and various rodent species, including rats, squirrels, and woodchucks. Although lagomorphs and pet rodents are not considered an important reservoirs for rabies, a recent report of rabies in seven domestic rabbits and a guinea pig suggests that pet owners and veterinarians should take appropriate precautions when animals show clinical signs associated with rabies infection.15 The pet rabbits and guinea pig were exposed and infected with the raccoon variant of rabies virus in the northeastern United States. A rabies vaccination (Imrab 3, Merial, Athens, GA) has been approved for use in ferrets but not rabbits or rodents. Veterinarians should strongly recommend vaccinating ferrets against rabies. Although the risk of contracting rabies is extremely low, an unvaccinated ferret that bites a human must be confined for a 10-day period to be evaluated for signs of disease. If no disease signs are noted after the 10-day confinement period, the animal is released back to the owner.15 At this time, there is no known shedding period or clinical signs of rabies infection in pet rabbits and rodents.15 If an owner is bitten and the owner is unsure of potential rabies exposure by that animal, a strong recommendation for definitively determining the animal’s rabies status through euthanasia should be given.15 Pet owners should take the appropriate precautions to prevent contact between pets housed outdoors and wildlife, especially in the northeastern United States. Reoviruses are found in several different vertebrate hosts. Mouse reovirus type 3 has been associated with morbidity and mortality in suckling mice. Affected animals may be jaundiced, stunted, develop oily hair, and have diarrhea. This virus is spread through both horizontal and vertical routes. Reovirus type 3 has been associated with disease in humans, causing enteritis in susceptible and particularly immunocompromised people who come into contact with infected mice. Immunocompromised individuals should therefore avoid contact with reovirus-infected mice. In 2003, a rare zoonotic viral disease outbreak occurred in the United States. Monkeypox virus (MPV), a member of the orthopoxvirus group, was responsible for the outbreak. MPV
561
Fig. 40-4 Lesions on the face of primate infected with monkeypox virus. (Courtesy of Bruce Williams, DVM.)
infection has similar clinical manifestations as smallpox and primarily occurs in the rainforest countries of central and west Africa.20 This was the first report of the disease in the United States, and community-acquired MPV infection had never been reported outside of Africa. Until the latest outbreak, most of the concerns for zoonotic infection have been in primate research and laboratory animal facilities, especially those housing Asian macaques (Fig. 40-4).16 Human-to-human contact has been reported but rarely occurs.16 In the 2003 U.S. outbreak, most people who contracted clinical illness were exposed through close contact with sick captive prairie dogs (Cynomys species). The prairie dogs were thought to have became infected after they were exposed to MPV-positive animals from a group of about 800 animals imported from Ghana to Texas.12 The animals were transported to an Iowa animal vendor and subsequently to a Chicago area pet distributor; they included at least one MPV-positive Gambian giant pouched rat (Aterurus species), two rope squirrels (Funisciurus species), and three dormice (Graphiurus species).12 MPV was transmitted from a diseased prairie dog to a rabbit at a veterinary clinic in the 2003 U.S. outbreak.12 The rabbit was considered the source of MPV infection in a human and confirmed the transmissibility of the virus from a common pet mammal to humans.12 Animal species that are potential carriers of MPV and/or showing disease signs of a nonspecific nature should be quarantined, stabilized, and reported to state public health officials. The clinical signs of MPV infection reported in animals during the 2003 U.S. outbreak included fever, cough, sneezing, blepharoconjunctivitis, lymphadenopathy, and nodular rash. Traditionally, MPV has been associated with a high morbidity rate and a low to variable mortality rate in humans (1%-10% case fatality rate in African outbreaks). There were 81 reported cases of MPV infection in the 2003 U.S. outbreak with no fatalities.12 Humans can be exposed from direct contact with an infected animal’s body fluids or bite wounds.20 There are currently no proven treatments for human MPV infection.12 Morbidity and mortality rates for animals are variable.
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If MPV infection is suspected in an animal, the clinic should be contacted before its arrival. The owners should protect themselves against potential exposure by quarantining the animal and wearing masks and gloves when the animal needs to be handled. Currently, all animals suspected of being infected with MPV or exhibiting clinical illness should be euthanatized and undergo necropsy. If the animal is not exhibiting clinical signs, there is no recommendation by the Centers for Disease Control and Prevention for euthanasia, even with the knowledge of an animal’s possible exposure to a confirmed case of MPV infection. The 2003 MPV outbreak is an example of a rare emerging viral disease in the United States. Veterinarians should be aware that they are often the first line of defense when an unusual form of morbidity and mortality in animals is encountered. Communication with local, regional, state, and national public health officers is important in trying to identify a series of unusual animal disease presentations and conditions. In 2009, four human cases of cowpox virus (CPV) infection were reported in France. The source of infection was exposure to infected pet rats (Rattus norvegicus) purchased at the same pet store.37 The rats had been purchased from a rat breeder in the Czech Republic.37 All four patients reported being scratched by their rats but not bitten.37 Clinical signs included cutaneous pustular lesions and lymphangitis. All of the patients recovered from the CPV infection with no apparent complications. As with MPV, the CPV infections reemphasize the possibility of unusual infectious diseases being transmitted from animals to humans. Purchasing small mammals commonly maintained as pets from reputable sources will help reduce the potential of exposure to a novel infectious disease.
PARASITIC DISEASES Cryptosporidiosis is a common zoonotic disease. Cryptosporidium parvum does not appear to be host-specific and may infect a wide range of mammals.5 The infective dose is low (1-10 oocysts) and they are immediately infective when excreted in the feces. The oocyts are very stable within the environment, which can lead to contamination of food and water supplies.5 A 6-month study evaluating three serial fecal samples from a group of ferrets being received at a biological research facility found that 66% of the animals had cryptosporidiosis.41 Ferrets with cryptosporidiosis are generally asymptomatic, although postmortem examination has revealed mild pathologic changes in some clinically normal animals. Individuals with compromised immune systems, such as children and those with acquired immunodeficiency virus (AIDS), should limit their contact with Cryptosporidium-positive ferrets. Fatal intestinal cryptosporidiosis has also been reported in African hedgehogs, suggesting that these animals may also pose a zoonotic threat to human caretakers.22 Recent reports appear to indicate that since household pets have rarely been implicated as a source of human infection, they may not serve as an important reservoir for Cryptosporidium species infection.5 Although there are no serious zoonotic diseases that can be spread from rabbits to healthy humans, immunocompromised (e.g., AIDS) patients may contract Encephalitozoon cuniculi.10 In one report, three patients with human immunodeficiency virus (HIV) had E. cuniculi, with one of these having clinical manifestations of disease with the parasite (severe interstitial
Fig. 40-5 Photomicrograph of eggs of Hymenolepis species. These tapeworms infect pet rats, mice, and hamsters. (Courtesy of Bruce Williams, DVM.)
pneumonitis).5 The symptoms abated and excretion of the parasite ceased when these patients were treated with albendazole. The two main Giardia species that infect rodents are Giardia duodenalis and Giardia muris.29 G. duodenalis has also been isolated from ferrets.1 While both of these parasites have been isolated from pet animals, limited epidemiologic investigations have been done to determine the prevalence of this parasite in this group of animals. It is important to collect this type of information in determining the potential risk for exotic pet mammal-associated giardiasis in humans. A 2010 study in chinchillas from Belgium found that 66.3% of animals tested shed G. duodenalis cysts. Shedding was more common in young animals and animals participating in shows. These results suggest that the risk to humans may be greatest in those animals experiencing a form of physiologic stress. Clinical signs vary depending on the species of Giardia and the species of animal infected but may include either no outward signs of clinical disease or severe dehydration and diarrhea. The clinical disease in humans primarily affects the gastrointestinal tract, but other body systems may be involved, including the joints, specifically the synovial surfaces; there may also be hypersensitivity reactions.33 Transmission is direct through fecal-oral contamination of the trophozoite and cyst form of the organism, primarily in water. Diagnosis and treatment of the affected animals to prevent further shedding and proper hygiene after handling animals will prevent the spread and transmission of this protozoal parasite to humans. Hymenolepsis is a cestode that infects rodents. Hymenolepsis diminuta has been identified in rats, hamsters, and mice, whereas Hymenolepsis nana is found mainly in mice and rats. Humans can become infected with this parasite by ingesting contaminated feces. Indirect infection can occur when insects ingest the tapeworm eggs, which then develop into cysticercoid larvae in the indirect host’s body (Fig. 40-5). In a survey of pet stores in southern Connecticut, a fecal flotation analysis revealed a 9.1% overall prevalence of H. nana in 110 samples collected weekly from cages holding group-housed small exotic mammals from three stores for 4 weeks.13 Only cages holding rats, mice, and prairie dogs were positive.13 Necropsies were performed on 38 rats (31.6% prevalence, mean worm load 66), 72 domestic mice (22.2% prevalence, mean worm load 14), 39 golden
CHAPTER 40 Zoonotic Diseases hamsters (10.3% prevalence, mean worm load 15) purchased from nine different pet stores.13 The findings of the pet store survey indicates the need for pet owners of small exotic mammals to practice good hygiene after handling their pets and the potential of public health significance of H. nana. Although most infected humans are asymptomatic, particularly with H. diminuta, clinical disease can occur with heavy infections because of the absorption of metabolic waste produced by a heavy worm burden and gastrointestinal irritation.28 Children are more likely to be clinically affected than adults. Affected individuals may have abdominal cramping, nausea, vomiting, and diarrhea. The cestodes Taenia taeniaeformis of rats and mice and Taenia serialis in rabbits can cause clinical illness in people when infected animals are consumed. Sarcoptic mange, caused by Sarcoptes scabiei, affects many mammalian species. Affected animals often have severe, intense pruritus, scales, crusts, erythema, and generalized alopecia. Pododermatitis and self-mutilation are common findings in ferrets. Humans who become infected with sarcoptic mange may develop wheals, vesicles, papules, and intense pruritus. Infected animals should be quarantined from other animals and handled only with gloves to reduce exposure. Lime sulfur shampoos, ivermectin, and selamectin may be used to treat affected animals. Trixacarus caviae is an ectoparasite of guinea pigs. In general, guinea pigs infested with T. caviae are asymptomatic. However, animals that are maintained in poor conditions or are stressed may have generalized clinical signs, including alopecia, intense pruritus, scales, crusts, and hyperkeratosis. Pet owners, especially children, may become infected with T. caviae from direct contact with infected guinea pigs. Affected individuals generally have mild dermatitis. T. caviae is very difficult to eliminate in affected guinea pigs. Intensive environmental cleaning and management along with therapeutic treatment with ivermectin or selamectin have been effective. Liponyssus bacoti, the tropical rat mite, will bite humans. Although this parasite mainly feeds on the rat host and lives within the animal’s bedding, it is the transfer of other infectious diseases through the bite of the mite that is of medical concern. Liponyssus bacoti can transmit murine typhus, Q fever, and Y. pestis.26 Treatment and prevention follow the same environmental and therapeutic protocols as described for other skin mite species. Cheyletiella parasitovorax is an ectoparasite that infests domestic rabbits (Fig. 40-6). This mite is generally found on the hair shaft and in the keratin layer of the skin. Infested rabbits may be asymptomatic or have alopecia and scaling. In severe cases, rabbits with cheyletiellosis appear to have “walking dandruff.” The life cycle of this mite is 35 days, and the parasite generally does not survive for more than 10 days off of the rabbit host. Humans who become infested with this mite generally develop focal to multifocal dermatitis. Papular and pruritic eruptions on the arms, thorax, waist, and legs have been described. The mite does not reproduce on the human host; therefore the infestation is generally self-limiting. Pet owners should be instructed to limit contact with infected rabbits during the treatment period, and gloves should be worn when contact is necessary.
MYCOTIC DISEASES Trichophyton species (usually T. mentagrophytes) is an opportunistic fungal disease of mammals. “Ringworm” has been isolated from rodents, lagomorphs, sugar gliders, ferrets, hedgehogs, and marsupials. Affected animals may be asymptomatic or have
563
Fig. 40-6 Photomicrograph of the mite Cheyletiella parasitovorax, which commonly infests rabbits and can transiently infest humans. (Courtesy of Bruce Williams, DVM.) focal to multifocal alopecia. Depending on the degree of pruritus associated with the infection, crusts, scabs, and excoriations may also be present on all of the animals or groups of animals mentioned above. Approximately 25% of the human ringworm cases reported annually are attributed to animal contact.43 Affected humans may be asymptomatic or have alopecia and crusts, similar to the lesions identified in pet animals. To a lesser extent, Microsporum canis may affect rodents, with active lesions in humans most often found on the scalp. A Wood’s lamp may be used to diagnose Microsporum canis through fluorescence of the fungal organism, while M. gypseum rarely fluoresce and T. mentagrophytes do not fluoresce.32 Trichophyton rubrum and T. mentagrophytes produce mannans and other compounds that reduce cell-mediated immunity, thereby predisposing animals to recurrent infections.32 Topical application of antimycotics is the recommended treatment in infected humans. Griseofulvin is commonly used as a systemic treatment for all small exotic mammals diagnosed with fungal dermatopathies.32 Most of the topical antifungal shampoos and treatments approved for cats are safe for small exotic mammals.32 Always check an exotic animal formulary for drug dosages and comments relating to treatment of a specific animal species with any therapeutic agent. Paecilomyces species, saprophytic fungi, are ubiquitous in the environment.45 Although these fungal organisms rarely cause clinical disease in mammals, infections have been reported in horses, dogs, cats, and reptiles.11,18 A case of paecilomycosis was recently described in an African pygmy hedgehog.25 The animal presented for marked hyperkeratosis on the right side of the face. Treatment with itraconazole was curative. It is generally assumed that these infections are secondary to some immunecompromising event; however, this could not be determined in the hedgehog in this case. Human caretakers for exotic pet mammals should be reminded of the potential for exposure to fungal agents, especially when animals are being treated with immunosuppressant drugs or have an immune-compromising disease. The practice of strict hygiene should minimize the potential for exposure to this organism.
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SECTION VI General Topics
ALLERGIC REACTIONS It has been estimated that between 11% and 15% of people exposed to rodent and lagomorph species have or develop allergic reactions to antigens produced by these animals.26 Human allergic reactions to small-mammal skin and fur antigens can present as respiratory (e.g., dyspnea, congestion), ocular (e.g., itchy tearing eyes), skin (e.g., dermal rash, pruritus), or anaphylactic clinical signs. Many people are allergic to one animal’s antigen, although multiple allergies involving small mammals may occur. The most common small-animal antigens are, in descending order, those of rats, guinea pigs, rabbits, mice, hamsters, and gerbils. Respiratory reactions to aerosolized rat urine are particularly severe because of the life-threatening pulmonary effects of the tissue response to the antigenic components of the urinary compounds.26 Antihistamine treatment is recommended for people with allergic reactions; reducing exposure to the antigenic compounds by washing hands and/or wearing gloves and a face mask will help decrease hypersensitivity reactions. Although zoonotic diseases rarely result from our interactions with nontraditional small mammals maintained as companion animals, it is an area of importance to veterinarians. Understanding the zoonotic potential of small-animal diseases is essential not only to protect the health of the veterinarians and staff but also to educate and inform their pet-owning clients and to serve as a source of information for the human medical community.
References 1. Abe N, Tanoue T, Noguchi E, et al. Molecular characterization of Giardia duodenalis isolates from domestic ferrets. Parasitol Rev. 2010;106(3):733-736. 2. Acha F, Szyfres B. Zoonoses and communicable diseases common to man and animals. 2nd ed. Washington, DC: Pan American Health Organization; 1987. 3. Anderson LC. Guinea pig medicine and husbandry. Vet Clin North Am Small Anim Pract. 1987;17:1045-1060. 4. Bradley T, Angulo F, Mitchell MA. Public health education on Salmonella spp and reptiles. J Am Vet Med Assoc. 2001;219:754-755. 5. Cacciò SM, Thompson RC, McLauchlin J, et al. Unravelling Cryptosporidum and Giardia epidemiology. Trends Parasit. 2005;21:430-437. 6. CDC on leptospirosis. http//www.cdc.gov/ncidod/dbmd/disea seinfo/leptospirosis_g.htm. Accessed October 19, 2009. 7. CDC plague fact sheet. http://www.cdc.gov/ncidod/dvbid/plag ue/facts.htm. Accessed October 19, 2009. 8. Chiles JE, Glass GE, Ksiazek TG, et al. Human rodent contact and infection with lymphocytic choriomeningitis and Seoul viruses in an inner-city population. Am J Trop Med Hyg. 1991;44:117-121. 9. Deibel R, Woodall JP, Decher WJ, et al. Lymphocytic choriomeningitis virus in man. Serologic evidence of association with pet hamsters. J Am Med Assoc. 1975;232:501-504. 10. Deplazes P, Mathis A, Baumgartner R, et al. Immunologic and molecular characteristics of Encephalitozoon-like microsporidia isolated from humans and rabbits indicate that Encephalitozoon cuniculi is a zoonotic parasite. Clin Infect Dis. 1996;22:557-559. 11. Diaz-Figueroa O, Mitchell MA, Ramirez S, et al. Paecilmomyces lilacinus pneumonia in a free-ranging Louisiana gopher tortoise. J Herp Med Surg. 2008;18(2):52-60.
12. DiGiulio DB, Eckburg PB. Human monkeypox: an emerging zoonosis. Lancet Infect Dis. 2004;4:15-25. 13. Duclos LM, Richardson DJ. Hymenolepis nana in pet store rodents. Compar Parasit. 2000;67:197-201. 14. Duthie RC, Mitchell CA. Salmonella enteritidis infections in guinea pigs and rabbits. J Am Vet Med Assoc. 1931;78:27-41. 15. Eidson M, Matthews SD, Willsey AI, et al. Rabies virus infection in a pet guinea pig and seven pet rabbits. J Am Vet Med Assoc. 2005;227:932-935. 16. Fenner F, Wallace P. Rowe lecture. Poxviruses of laboratory animals. Lab Anim Sci. 1990;40:469-480. 17. Ferret gets swine flu from its owner. http://www.oregonlive. com/pets/index.ssf/2009/10/ferret_gets_swine_flu_from_its. html. Accessed October 22, 2009. 18. Foley JE, Norris CR, Jang SS. Paecilomycosis in dogs and horses and a review of the literature. J Vet Intern Med. 2002;16:238-243. 19. Fox JG, Ackerman JL, Taylor N, et al. Campylobacter jejuni in the ferret: a model of human campylobacteriosis. Am J Vet Res. 1987;48:85-90. 20. Fox JG, Newcomer CE, Rozmiarek H. Selected zoonoses. In: Fox JG, Anderson LC, Loew FM, et al. eds, Laboratory Animal Medicine. 2nd ed. San Diego: Academic Press; 2002:1059-1105. 21. Fuller CC, Jawahir SL, Leano FT, et al. A multi-state Salmonella Typhimurium outbreak associated with frozen vacuum-packed rodents used to feed snakes. Zoonoses Public Health. 55(8-10): 481-487. 22. Graczyk TK, Cranfield MR, Dunning C, et al. Fatal cryptosporidiosis in a juvenile captive African hedgehog (Atelerix albiventris). J Parasitol. 1998;84:178-180. 23. Graves MH, Janda MJ. Rat-bite fever (Streptobacillus moniliformis): a potential emerging disease. Int J Infect Dis. 2001; 5:151-154. 24. Habermann RT, Williams Jr FP. Salmonellosis in laboratory animals. J Natl Cancer Inst. 1958;20:933-947. 25. Han J, Na KJ. Cutaneous paecilomycosis caused by Paecilomyces variottii in an African pygmy hedgehog (Atelerix albiventris). J Exot Pet Med. 2010;19(4):309-312. 26. Harkness JE, Wagner JE. The biology and medicine of rabbits and rodents. 4th ed. Baltimore: Lea & Febiger; 1995;103-284. 27. Hirakawa Y, Sasaki H, Kawamoto E, et al. Prevalence and analysis of Pseudomonas aeruginosa in chinchillas. BMC Vet Res. 6:52. 28. Jueco NL. Hymenolepis nana infection. In: Steele JH, Jacobs L, Arambulo P, eds. Handbook series in zoonosis. Section C: parasitic zoonosis. Vol 1. Boca Raton: CRC Press; 1982:283-287. 29. Levecke B, Meulemans L, Dalemans T, et al. Mixed Giardia duodenalis assemblage A, B, C, and E infections in pet chinchillas (Chinchilla laniger) in Flanders (Belgium). Vet Parasitol. 2010. 30. Lipsky S, Tanino T. African pygmy hedgehog-associated salmonellosis—Washington. MMWR Morb Mortal Wkly Rep. 1995;44:462-463. 31. Lymphocytic choriomeningitis virus (LCMV) and pregnancy: facts and prevention. www.cdc.gov/ncbddd. Accessed October 19, 2009. 32. Marshall KL. Fungal disease in small mammals: therapeutic trends and zoonotic considerations. Vet Clin N Am Exot Anim Pract. 2003;6:415-427. 33. Meyer EA, Jarroll EL. Giardiasis. In: Steele JH, Jacobs L, Arambulo P, eds. Handbook series in zoonosis. Section C: parasitic zoonosis. Vol 1. Boca Raton: CRC Press; 1982:25-40. 34. Mills JN, Yates TL, Ksiazek TG, et al. Long-term studies of hantavirus reservoir populations in the southwestern United States: rationale, potential, and methods. Emerg Infect Dis. 1999;5:95-101. 35. Moore B. Observations pointing to the conjunctiva as the portal of entry in salmonella infection of guinea pigs. J Hyg (Lond). 1957;55:414-433.
CHAPTER 40 Zoonotic Diseases 36. Morrison G. Zoonotic infections from pets. Understanding the risks and treatment. Postgrad Med. 2001;110:24-26, 29-30, 35-36. 37. Ninove L, Domart Y, Vervel C, et al. Cowpox virus transmission from pet rats to humans, France. Emerging Infect Dis. 2009;15:781-784. 38. Olson GA, Shields RP, Gaskin JM. Salmonellosis in a gerbil colony. J Am Vet Med Assoc. 1977;171:970-972. 39. Plant M, Zimmerman EM, Goldstein RA. Health hazards to humans associated with domestic pets. Annu Rev Public Health. 1996;17:221-245. 40. Pollock WJ, Lanza J, Williams PA, et al. Fatal rat-bite fever– Florida and Washington 2003. MMWR. 2005;53(51&52): 1198-1202. 41. Rehg JE, Gigliotti F, Stokes DC. Cryptosporidiosis in ferrets. Lab Anim Sci. 1988;38:155-158. 42. Ruiz A. Plague in the Americas. Emerg Infect Dis. 2001;7:539-540. 43. Ryan CP. Animals in schools and rehabilitation facilities. So Calif Vet Med Assoc. 1998;9(6):16. 44. Saeed AM, Harris NV, DiGiamcomo RF. The role of exposure to animals in the etiology of Campylobacter jejuni/coli enteritis. Am J Epidemiol. 1993;137:108-114. 45. Salle V, Lecuyer E, Chouaki T, et al. Paecilomyces variottii fungemia in a patient with multiple myeloma: case report and literature review. J Infect. 2005;51:93-95.
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46. Schmaljohn C, Hjelle B. Hantaviruses: a global disease problem. Emerg Infect Dis. 1997;3:95-104. 47. Shvartsblat S, Kochie M, Harber P, et al. Fatal rat bite fever in a pet shop employee. Am J Ind Med. 45(4):357-360. 48. Steffen EK, Wagner JE. Salmonella enteritidis serotype Amsterdam in a commercial rat colony. Lab Anim Sci. 1983;33:454-456. 49. Swanson SJ, Snider C, Braden CR, et al. Multidrug-resistant Salmonella enterica serotype Typhimurium associated with pet rodents. N Engl J Med. 2007;356(91):21-28. 50. Turnbull PCB. Food poisoning with special reference to Salmonella—its epidemiology, pathogenesis, and control. Clin Gastroenterol. 1979;8:663-713. 51. Webby RJ, Perez DR, Coleman JS. Responsiveness to a pandemic alert: use of reverse genetics for rapid development of influenza vaccines. The Lancet. 2004;363:1099-1103. 52. Winsser J. A study of Bordetella bronchiseptica. Proc Anim Care Panel. 1960;10:87-104. 53. Woodward DL, Khakhria R, Johnson WM. Human salmonellosis associated with exotic pets. J Clin Micro. 1997;35:2786-2790. 54. Zhou J, Sun W, Wang J, Guo J, et al. Characterization of the H5N1 highly pathogenic avian influenza virus derived from wild pikas in China. J Virol. 2009;83:8957-8964.
APPENDIX
Formulary
James K. Morrisey, DVM, Diplomate ABVP (Avian), and James W. Carpenter, MS, DVM, Diplomate ACZM
As in the previous editions, this formulary presents the antimicrobial agents (Table A-1); antifungal agents (Table A-2); antiparasitic agents (Table A-3); chemical restraint, anesthetic, and analgesic agents (Table A-4); and miscellaneous agents (Table A-5) most commonly used in ferrets, rabbits, guinea pigs, chinchillas, hamsters, rats, mice, hedgehogs, and sugar gliders. There are many specific circumstances and drug types that cannot be included in this chapter. The reader is referred to another, more comprehensive reference1 and the literature for this information. Additionally, specific chapters in this book may suggest dosages that are different or not mentioned in this chapter. Most of the dosages presented in this chapter are adapted from the Exotic Animal Formulary,1 and the reader should refer to this reference for more details of the drugs, dosages, indications, and precautions used in exotic animals. It should be noted that most of the dosages listed in this formulary are off-label and based on empiric data, observation, and experience. To date, very few drugs are approved by the Food and Drug Administration (FDA) for use in small mammals kept as nontraditional pets, such as the animals discussed in this book. The Animal Medicinal Drug Use Clarification Act enables veterinarians to use drugs approved for human and other animal use on animals other than those for which they are approved if the reasoning is sound. This act requires that owners sign an informed consent form for off-label drug use. Because most drugs are not approved by the FDA for use in these animals, it may be warranted for the owners to sign a consent form in registering their pet. Because pharmacokinetic studies are lacking in these pet species, it is important to know some of the pharmacobiologic, physiologic, and anatomic characteristics of these animals. Drug uptake and use depend on many factors, including age, body fat, sex, physiology (gastrointestinal, hepatic, renal, etc.), illness, diet, fasting state, and others. For example, it is well known that hindgut fermenters, such as rabbits and guinea pigs, do not tolerate some oral antibiotics, such as penicillins, because they can cause a fatal enterotoxemia. The complete list of this
566
information is beyond the scope of this chapter; the reader is referred to the chapters on anatomy and physiology of each species elsewhere in this book. Because these dosages are often empiric, it is helpful to know, for maximum safety and efficacy, the basic pharmacologic features and side effects of the drugs being used. It is important to inform the owner or technical staff of clinical signs that may indicate toxicity. This information is available in the drug inserts and with other references and formularies. Allometric scaling may also be helpful in prescribing drugs in this formulary to other species. Most of the drugs given to these nontraditional pet species are given parenterally or orally in the form of suspensions. There has been little research on the efficacy of various drug suspensions in these patients. It is essential for practitioners to have a good working relationship with a licensed compounding pharmacist. These pharmacists will ensure that the drug is placed in the appropriate medium to remain viable in suspension and can warn of potential risks associated with this medium. It is important to obtain a definitive diagnosis whenever possible to avoid the problems associated with the empiric dosages and improve efficacy in these species. Antibiotics should be selected based on culture and sensitivity results whenever possible. The use of more pathogen-specific antimicrobials will decrease the potential for resistant bacteria and often narrow the potential side effects. We have attempted to verify and double-check all dosages presented in this formulary. However, despite these efforts, errors in the original sources or in the preparation of this chapter may have occurred. All users of this reference should, therefore, empirically evaluate all dosages to determine that they are reasonable before use. We assume no responsibility for and make no warranty regarding results obtained from the dosages listed.
Reference 1. Carpenter JW, ed. Exotic animal formulary. 4th ed. St. Louis: Saunders/Elsevier; 2012.
Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
Table A-1 Antimicrobial Agents Commonly/Occasionally Used in Small Mammals Agent
Ferrets
Rabbits
Guinea Pigs (G)/ Chinchillas (C)
Gerbils (G)/ Hamsters (H)
Rats (R)/Mice (M)
Hedgehogs
Sugar Gliders
Amikacina
10-15 mg/kg SC, IM q12h 20-30 mg/kg PO q8-12h 12.5-25 mg/kg PO q8-12h 5-30 mg/kg SC, IM, IV q8-12h
5-10 mg/kg SC, IM, IV divided q8-24h Do not use
16 mg/kg SC, IM, IV divided q8-24h Do not use
16 mg/kg SC, IM divided q8-24h Do not use
16 mg/kg SC, IM q8-12h 20 mg/kg PO q24h
3 mg/kg IM q12h
Do not use
Do not use
Do not use
20 mg/kg PO q12h
Do not use
Do not use
20-100 mg/kg PO, SC, IM q8h
Azithromycin
5 mg/kg PO q24h
15-30 mg/kg PO q24h
Cephalexin
15-30 mg/kg PO q8-12h
5 mg/kg IM q48h; 15-30 mg/kg PO q24h 15 mg/kg SC q12h
6-30 mg/kg PO q8h (G); Do not use (H) 15-30 mg/kg PO q24h
2.5-5.0 mg/kg IM q8-12h 15 mg/kg PO, SC, IM q12h 12.5 mg/kg PO q12h 10 mg/kg IM q12h (use with caution) —
Best to avoid
25 mg/kg SC q24h
25 mg/kg PO q8h
30 mg/kg PO, SC divided q12-24h
Chloramphenicol
25-50 mg/kg PO, SC, IM q12h —
25-50 mg/kg PO, SC, IM q12h
30-50 mg/kg PO, SC, IM q8-12h
30-50 mg/kg PO, SC, IM q8-12h
60 mg/kg PO q12h (M); 15 mg/kg SC q12-24h (R) 30-50 mg/kg PO, SC, IM q8-12h
30-50 mg/kg PO, SC, IM, IV q12h
—
50 mg/kg PO q12-24h
50 mg/kg PO q12h (C)
20 mg/kg PO, SC, IM q12h
5-20 mg/kg PO q12h
—
5-15 mg/kg PO q12h 12.5-25 mg/kg PO q12h 5-10 mg/kg PO q12h —
5-20 mg/kg PO q12-24h —
5-20 mg/kg PO q12-24h —
5-20 mg/kg PO q12-24h —
10 mg/kg SC, IM q12h (R): 25 mg/kg PO, SC, IM q12h (M) 10 mg/kg PO q12h
10 mg/kg PO q12h —
Do not use
7.5 mg/kg SC q12h Do not use PO 2.5-5.0 mg/kg PO q12h
7.5 mg/kg SC q12h Do not use PO 2.5-5.0 mg/kg PO q12h
5-20 mg/kg PO q12h 5.5 mg/kg PO q12h 5.5-10 mg/kg PO q12h 2.5-10 mg/kg PO, SC, IM q12h
Enrofloxacinb
10-20 mg/kg PO, SC, IM q12-24h
5-10 mg/kg PO, SC, IM, IV q12h
5-20 mg/kg PO, SC, IM q12h
5 mg/kg PO, SC, IM q12h
5 mg/kg PO, SC, IM q12h
Erythromycin
10 mg/kg PO q6h —
Do not use
Do not use 4 mg/kg PO, SC q24h
10 mg/kg PO, IM q12h —
—
2 mg/kg IM, IV q24h; 5 mg/kg PO q24h
5-20 mg/kg PO, SC, IM q12h; 0.05-0.2 mg/mL drinking water Do not use or use with caution 4 mg/kg PO, SC q24h
Amoxicillin Amoxcillin/ clavulanic acid Ampicillin
Chlortetracycline
Ciprofloxacin Clarithromycin Clindamycin Doxycycline
— 7.5 mg/kg SC q12h Do not use PO 2.5-5.0 mg/kg PO q12h; 70-100 mg/kg SC, IM q7d (long-acting form) 5-20 mg/kg PO, SC, IM q12h; 0.05-0.2 mg/mL drinking water 20 mg/kg PO q12h 4 mg/kg PO, SC q24h
—
— —
Formulary
Marbofloxacin
2.5 mg/kg PO q12h
15-30 mg/kg PO q24h
30 mg/kg PO, IM divided q12-24h 12.5 mg/kg PO, SC divided q12-24h —
— 567
Continued
568
Table A-1 Antimicrobial Agents Commonly/Occasionally Used in Small Mammals—cont’d Ferrets
Rabbits
Metronidazole
20 mg/kg PO q12h
20 mg/kg PO q12h
Neomycin
10-20 mg/kg PO q6h 40,000 IU/kg SC q24h
Best to avoid
25 mg/kg PO q12h 15-30 mg/kg PO, SC q12h 10 mg/kg PO, SC q8-12h
50 mg/kg PO q8-12h 30 mg/kg PO, SC q12h 10 mg/kg PO, SC q12h
Penicillin G procaine Tetracycline Trimethoprim/ sulfa Tylosin
60,000 IU/kg SC, IM q8h (use with caution)
Guinea Pigs (G)/ Chinchillas (C)
Gerbils (G)/ Hamsters (H)
20 mg/kg PO q12h (G); 10-20 mg/kg PO q12h (use with caution) (C) Best to avoid Do not use (G); 22,000 IU/kg SC, IM q24h (C) (use with caution) 10 mg/kg PO q8-12h (use with caution) 15-30 mg/kg PO, SC q12h 10 mg/kg PO, SC q12h (use with caution in guinea pigs)
Rats (R)/Mice (M)
Hedgehogs
Sugar Gliders
20 mg/kg PO q12h
10-40 mg/kg PO q24h
20 mg/kg PO q12h
25 mg/kg PO q12h
30 (H) -100 (G) mg/ kg PO q24h 22,000 IU/kg SC, IM q24h
25 mg/kg PO q12h
—
—
22,000 IU/kg SC, IM q24h
40,000 IU/kg SC, IM q24h
22,000-25,000 IU/ kg q12-24h
10-20 mg/kg PO q8-12h 15-30 mg/kg PO, SC q12h 2-10 mg/kg PO, SC q12h (use with caution)
10-20 mg/kg PO q8-12h 15-30 mg/kg PO, SC q12h 10 mg/kg PO, SC q12h
—
—
30 mg/kg PO q12h
15 mg/kg PO q12h
10 mg/kg PO, SC q12h
—
aMost
infectious disease clinicians now recommend that aminoglycosides should be dosed q24h in most mammals. This dosing regimen yields higher peak levels with a resultant greater bacterial kill. However, incorporating this recommendation with the published dosages is up to the individual clinician. Aminoglycosides should be used with fluid support. bThe oral route of administration is preferable; avoid subcutaneous (unless antibiotic is diluted) and intramuscular routes of administration because of potential tissue necrosis at injection site.
Table A-2 Antifungal Agents Commonly/Occasionally Used in Small Mammals Agent
Ferrets
Rabbits
Guinea Pigs (G)/ Chinchillas (C)
Gerbils (G)/ Hamsters (H)
Amphotericin B
0.4-0.8 mg/kg IV q7d (total dose, 7-25 mg) 50 mg/kg PO q12h
—
—
38 mg/kg PO q12h 12.5-25 mg/kg PO q12-24h 20-40 mg/kg PO q24h 10-40 mg/kg PO q24h Dip q7d x 4 treatments 100 mg/kg PO q12-24h
16 mg/kg PO q24h x 14 days 15-50 mg/kg PO q24h (G); 25 mg/kg PO q24h (C) 2.5-10 mg/kg PO q24h (G)
Fluconazole Griseofulvin Itraconazole Ketoconazole Lime sulfur dip (2.5%) Terbinafine
25 mg/kg PO q12-24h 15 mg/kg PO q24h 10-30 mg/kg PO q12-24h Dip q7d —
10-40 mg/kg PO q24h Dip q7d (G); Dip q7d x 4 treatments (C) 10-30 mg/kg PO q24h x 4-6 wk
Rats (R)/Mice (M)
Hedgehogs
Sugar Gliders
1 mg/kg SC q24h 5 days/wk x 3 wk (H) —
0.11 mg/kg SC; 0.43 mg/kg PO (M)
—
—
—
—
—
25 mg/kg PO q24h
25 mg/kg PO q24h
—
—
2.5-10 mg/kg PO q24h (R); 50 mg/kg PO q24h (M) 10-40 mg/kg PO q24h
50 mg/kg PO q24h 5-10 mg/kg q12-24h 10 mg/kg PO q24h Apply topically —
—
10-40 mg/kg PO q24h Apply topically q7d 10-30 mg/kg PO q24h x 4-6 wk
Apply topically q7d 10-30 mg/kg PO q24h x 4-6 wk
5-10 mg/kg PO q12h — —
Formulary
Agent
Table A-3 Antiparasitic Agents Commonly/Occasionally Used in Small Mammals Agent
Ferrets
Rabbits
Albendazole
—
7.5-20 mg/kg PO q24h (use with caution; deaths reported)
Amitraz
0.03% solution topically q7d Apply topically q7d
—
20 mg/kg PO q24h x 5 days 0.2-0.4 mL q30d; 1 pump of spray or 1⁄5-1⁄2 of cat tube q60d 0.1-0.4 mL or 1 cat dose topically q30d 0.2-0.5 mg/kg PO, SC, repeat in 14 days; 0.05 mg/kg PO, SC q30d (heartworm prevention or microfilaricide) Dip 1:40 dilution q7d
Carbaryl powder (5%) Fenbendazole
Fipronil
Imidacloprid Ivermectin
30-45 mg/kg PO q30d
Metronidazole
15-20 mg/kg PO q12h
Milbemycin oxime Piperazine adipate
1.15-2.33 mg/kg PO q30d —
Gerbils/Hamsters
Rats (R)/Mice (M)
Hedgehogs
Sugar Gliders
5 mg/kg PO q12h (G); 25 mg/kg PO q12h x 2 days (giardiasis) (C) 0.3% solution topically q7d (G) Apply topically q7d x 3 treatments
—
—
—
—
1.4 mL/L topically q7-14d Apply topically q7d x 3 treatments
1.4 mL/L topically q7d Apply topically q7d
0.3% solution topically q7d —
—
10-20 mg/kg PO, repeat in 14 days
20-50 mg/kg PO q24h x 5 days
20-50 mg/kg PO q24h x 5 days
20-50 mg/kg PO q24h x 5 days
10-15 mg/kg PO q14d x 2-3 treatments
Toxic; do not use
3 mL /kg topically (use with caution)
7.5 mg/kg topically q30-60d
7.5 mg/kg topically q30-60d
Apply 1 spray over dorsum, repeat in 10 days
20-50 mg/kg PO q24h x 3 days, repeat in 14 days —
10-16 mg/kg or 1 cat dose topically q30d 0.2-0.4 mg/kg SC q10-14d
20 mg/kg topically q30d
20 mg/kg topically q30d
20 mg/kg topically q30d
½ kitten dose topically q30d
—
0.2-0.4 mg/kg SC q7-14d
0.2-0.4 mg/kg SC q7-14d
0.2-0.4 mg/kg SC q7-14d
0.2-0.4 mg/kg PO, SC q10-14d x 3-5 treatments
0.2 mg/kg SC q10-14d
Dip q7d x 4-6 wk
Dip q7d x 6 wk
Dip q7d x 6 wk
Dip q7d x 6 wk
—
—
30 mg/kg PO q30d 20 mg/kg PO q12h —
—
—
—
—
25 mg/kg PO q12h
20-50 mg/kg PO q8h —
10-40 mg/rat PO q24h —
½ kitten dose PO q30d 25 mg/kg PO q12h
3-5 mg/mL drinking water x 7 days, off 7 days, repeat
4-7 mg/mL drinking water x 3-10 days
Apply topically q7d
500 mg/kg PO q24h x 2d
— 4-7 mg/mL drinking water x 3-10 days (G); 500 mg/kg PO q24h (C)
Apply topically sparingly
—
25 mg/kg PO q12h —
—
—
569
Continued
Formulary
Lime sulfur dip (2.5%) Lufenuron
Guinea Pigs (G)/ Chinchillas (C)
570
Table A-3 Antiparasitic Agents Commonly/Occasionally Used in Small Mammals—cont’d Guinea Pigs (G)/ Chinchillas (C)
Ferrets
Rabbits
Piperazine citrate
50-100 mg/kg PO, repeat in 14 days
100-200 mg/kg PO, repeat in 14 days
10 mg/mL drinking water x 7 days, off 7 days, repeat (G); 100 mg/kg PO q24h x 2 days (C)
Praziquantel Pyrantel pamoate
5-10 mg/kg PO, SC, repeat in 10-14 days 4.4 mg/kg PO, repeat in 14 days
Pyrethrin products Selamectin
Topically as directed q7d 10 mg/kg topically
5-10 mg/kg PO, SC, IM, repeat in 10 days 5-10 mg/kg PO, SC repeat in 14 days Topically as directed q7d 20 mg/kg topically q7da
Sulfadi methoxine
50 mg/kg PO once, then 25 mg/kg q24h x 9 days
Sulfamerazine
—
Sulfaqui noxaline Thiabendazole
—
aAlthough
—
50 mg/kg PO once, then 25 mg/kg q24h x 10-20 days 100 mg/kg PO 1 mg/mL in drinking water 50-100 mg/kg PO q24h x 5 days
Gerbils/Hamsters
Rats (R)/Mice (M)
Hedgehogs
Sugar Gliders
200 mg/kg PO q24h x 7 days, off 7 days, on 7 days (R); 4-5 mg/mL drinking water x 7 days, off 7 days, repeat 6-10 mg/kg PO, SC, repeat in 10 days
—
—
6-10 mg/kg PO, SC, repeat in 10 days
200 mg/kg PO q24h x 7 days, off 7 days, on 7 days (G); 10 mg/mL drinking water x 7 days, off 7 days, repeat 6-10 mg/kg PO, SC, repeat in 10 days
7 mg/kg PO, SC, repeat in 14 days
—
50 mg/kg PO
50 mg/kg PO
50 mg/kg PO
—
—
Topical powder q7d x 3 treatments 20 mg/kg topically (G)/6 mg/kg topically (C) 25-50 mg/kg PO q24h x 10 days
Topical powder 3x/ Topical powder 3x/ wk; shampoo q7d wk; shampoo q7d 15-30 mg/kg 15-30 mg/kg topically q21-28d topically q21-28d x 2 treatments x 2 treatments 25-50 mg/kg PO 10-15 mg/kg PO q24h x 10-14 days q12h
—
Topically as directed q7d 6-18 mg/kg topically
1 mg/mL drinking water 1 mg/mL drinking water 50-100 mg/kg PO q24 x 5 days
1 mg/mL drinking water 1 mg/mL drinking water 50-100 mg/kg PO q24h x 5 days
1 mg/mL drinking water 1 mg/mL drinking water 50-100 mg/kg PO q24h x 5 days
6 mg/kg topically
2-20 mg/kg q24h PO, — SC, IM x 2-5 days, off 5 days, on 2-5 days — — —
—
—
—
dosage is based on pharmacokinetic findings, further studies are needed to assess long-term safety in rabbits at this dose following repeated application.
Formulary
Agent
Table A-4 Chemical Restraint, Anesthetic, and Analgesic Agents Commonly/Occasionally Used in Small Mammals Agent
Ferrets
Rabbits
Acepromazine
0.1-0.5 mg/kg SC, IM 10-20 mg/kg PO q24h 1 mg/kg SC, IM, IV
0.25-1.0 mg/kg IM
Acetylsalicylic acid Atipamezolea Atropine
10-100 mg/kg PO q8-24h 1 mg/kg SC, IM, IV 0.1-0.5 mg/kg SC, IMb 0.01-0.05 mg/kg SC, IM, IV q6-12h
Butorphanol
0.05-0.4 mg/kg SC, IM q4-6h
Carprofen
Enflurane
2-5 mg/kg PO, SC q12-24h — 0.5-2.0 mg/kg IM, IV 2% maintenance
0.1-1.0 mg/kg SC, IM, IV q2-6h (doses up to 2 mg/kg may be given) 1-5 mg/kg PO q12-24h — 0.5-2.0 mg/kg IM, IV To effect; MACd = 2.9%
Fentanyl/fluanisone
0.3 mg/kg IM
—
Flunixin meglumine
0.3-2.0 mg/kg SC q12-24h
1.1 mg/kg SC, IM q12h
Gabapentin Glycopyrrolate
3-5 mg/kg PO q8-24h 0.01 mg/kg IM
Ibuprofen
1 mg/kg PO q4-8h
3-5 mg/kg PO q12-24h 0.01-0.02 mg/kg SC 2-7.5 mg/kg PO q12-24h
Buprenorphine
Dexmedetomidinec Diazepam
Gerbils (G)/ Hamsters (H)
0.5-1.0 mg/kg IM 50-100 mg/kg PO q4h (G) 1 mg/kg SC, IM, IV 0.05-0.2 mg/kg SC, IM, IV 0.05-0.1 mg/kg SC q6-12h
0.5-1.0 mg/kg IM 100-150 mg/kg PO q4h — 0.04-0.4 mg/kg SC, IM 0.1-0.2 mg/kg SC q8h (G); 0.5 mg/ kg SC q8h (H)
0.2-2.0 mg/kg SC, IM q2-4h
1-5 mg/kg SC q4h
2-5 mg/kg PO, SC q12-24h — 0.5-3.0 mg/kg IM, IV To effect; MAC = 2.17% (G); to effect (C) 0.5-1.0 mL /kg IM (G) 2.5-5.0 mg/kg SC q12-24h (G); 1-3 mg/kg SC q12h (C) 3-5 mg/kg PO q12-24h 0.01-0.02 mg/ kg SC 10 mg/kg PO q4h (G)
5 mg/kg SC q24h — 1-5 mg/kg IM, IP To effect 0.2-0.6 mL /kg IM 2.5 mg/kg SC q12-24h
50 mg/kg PO q24h (H) 0.01-0.02 mg/kg SC —
Rats (R)/Mice (M)
Hedgehogs
Sugar Gliders
0.5-1.0 mg/kg IM 100-150 mg/kg PO q4h 1 mg/kg SC, IM, IV, IP 0.05-0.4 mg/kg SC, IM 0.02-0.1 mg/kg SC, IM, IP q6-12h (R); 0.05-2.5 mg/kg SC, IM, IP q6-12h (M) 0.2-2 mg/kg SC, IM q2-4h (R); 1-5 mg/kg SC q4h (M) 2-5 mg/kg PO, SC, IM q12-24h — 1-5 mg/kg IM, IV
0.1-1.0 mg/kg PO, SC, IM —
—
0.3-0.5 mg/kg IM 0.01-0.04 mg/kg SC, IM 0.01 mg/kg SC, IM q6-8h
— 0.01-0.02 mg/kg SC, IM 0.01-0.03 mg/kg PO, SC q12h
0.05-0.4 mg/kg SC q6-8h
0.1-0.5 mg/kg SC, IM q6-8h —
To effect
1 mg/kg PO, SC q12-24h — 0.5-2.0 mg/kg IM To effect
0.2-0.6 mL /kg IM
—
2.5 mg/kg SC q12-24h
0.3 mg/kg SC q24h
—
—
0.01-0.02 mg/kg — SC 10-30 mg/kg PO — q4h (R); 7-15 mg/kg PO q4h (M)
—
— 0.5-2.0 mg/kg PO, IM, IV To effect 0.1-1.0 mL /kg IM q12-24h 0.1 mg/kg IM
— 0.01-0.02 mg/kg SC, IM, IV —
Formulary
0.04-0.05 mg/kg SC, IM, IV 0.01-0.05 mg/kg SC, IM, IV q8-12h
Guinea Pigs (G)/ Chinchillas (C)
Continued
571
572
Table A-4 Chemical Restraint, Anesthetic, and Analgesic Agents Commonly/Occasionally Used in Small Mammals—cont’d Gerbils (G)/ Hamsters (H)
Rats (R)/Mice (M)
Hedgehogs
Sugar Gliders
1.50%-1.75% maintenance; MAC = 2.05%
0.25%-4.0% maintenance
0.25-4.0% maintenance
0.25%-4.0% maintenance
0.5%-3.0% maintenance
5-50 mg/kg SC, IM, IV
5-50 mg/kg SC, IM; 15 mg/kg IV
5-40 mg/kg SC, IM, IV
22-44 mg/kg IM
5-20 mg/kg IM
10-35 mg/kg (K)/0.20-0.35 mg/ kg (A) SC, IM 10-35 mg/kg (K)/1-2 mg/kg (D) IM
40 mg/kg (K)/0.5-1.0 mg/kg (A) IM
—
—
50-150 mg/kg (K)/2.5-5.0 mg/ kg IM —
Ketamine/ medetomidine
5 mg/kg (K)/0.08 mg/kg (Me) IM
0.35 mg/kg (Me)/5-20 mg/kg (K) IV 15 min later
75-90 mg/kg (K)/0.5 mg/kg (Me) IM, IP
75-90 mg/kg (K)/ 1 mg/kg (Me) IP (mice)
5 mg/kg (K)/0.1 mg/kg (Me) IM
2-3 mg/kg (K)/0.05-0.1 mg/kg (Me) SC, IM
Ketamine/midazolam
5-10 mg/kg (K)/0.250.5 mg/kg (Mi) IM, IV
25 mg/kg (K)/≤2 mg/kg (Mi) IM, IV
40 mg/kg (K)/0.5 mg/kg (A) IM (C) 20-30 mg/kg (K)/1-2 mg/kg (D) IM (G); 20-40 mg/kg (K)/1-2 mg/kg (D) IM (C) 40 mg/kg (K)/0.5 mg/kg (Me) IM, IP (G); 5 mg/kg (K)/0.06 mg/kg (Me) IM (C) 5-10 mg/kg (K)/0.5-1.0 mg/ kg (Mi) IM
40-100 mg/kg IM (G); 50-100 mg/kg IM (H) —
0.25%-4.0% (generally 1%-3%) maintenance 20 mg/kg IM
—
40-150 mg/kg (K)/3-5 mg/kg (Mi) IV, IP
—
Ketoprofen
1-3 mg/kg PO, SC, IM q12-24h 0.1-0.2 mg/kg SC, IM
1-3 mg/kg IM q12-24h 0.1-0.25 mg/kg IM
1-2 mg/kg SC, IM q12-24h 0.3 mg/kg SC, IM (G)
5 mg/kg SC q24h
2-5 mg/kg PO, SC, IM q12-24h 0.03-0.1 mg/kg SC
—
10-20 mg/kg (K)/0.35-0.5 mg/kg (Mi) SC, IM —
0.1 mg/kg IM
—
Meloxicam
0.2-0.3 mg/kg PO, SC, IM q24h
≥0.3-0.5 mg/kg PO, SC q24h
≥0.5 mg/kg PO, SC q24h
0.2 mg/kg PO, SC q24h
0.2 mg/kg PO, SC q24h
Meperidine
5-10 mg/kg SC, IM, IV q2-4h 0.25-0.5 mg/kg SC, IM, IV
5-10 mg/kg SC, IM q2-3h 0.25-2 mg/kg SC, IM, IV
20 mg/kg SC, IM q2-3h (G) 1-2 mg/kg SC, IM, IV
—
—
0.25-0.5 mg/ kg IM
0.25-0.5 mg/ kg IM
Ferrets
Rabbits
Isoflurane
2%-3% maintenance
Ketamine
Ketamine/ acepromazine Ketamine/diazepam
Medetomidinec
Midazolam
10-15 mg/kg (K)/0.3-0.5 mg/kg (D) IM, IV
0.1-0.2 mg/kg SC, IM (G)/0.1 mg/kg SC (H) ≥0.5 mg/kg PO, SC 1-2 mg/kg PO, SC q24h q24h (R); 1-5 mg/kg PO, SC q24h (M) 20 mg/kg SC, IM 20 mg/kg SC, IM q2-3h q2-3h 1-2 mg/kg IM 1.0-2.5 mg/kg SC, IM (R); 1-5 mg/kg SC, IM (M)
5-20 mg/kg (K)/0.5-2.0 mg/kg (D) IM
10-30 mg/kg (K)/1-2 mg/kg (A) SC —
Formulary
Guinea Pigs (G)/ Chinchillas (C)
Agent
Morphine
0.2-4.0 mg/kg SC, IM q2-4h
0.5-2.0 mg/kg SC, IM q2-4h
Oxymorphone
0.05-0.2 mg/kg SC, IM, IV q6-12h
0.05-0.2 mg/kg SC, IM q6-12h
Propofol
3-6 mg/kg IV
3-6 mg/kg IV
Sevoflurane
To effect
Tiletamine/zolazepam
12-22 mg/kg IM (rarely indicated)
To effect MAC d = 3.7% 3 mg/kg IM (not recommended)
Tramadol
5 mg/kg PO q12-24h
Xylazine Yohimbine
2-5 mg/kg SC, IM q4h
2-5 mg/kg SC q2-4h 2-5 mg/kg SC q4h (R); 2-10 mg/kg SC, IM q4h (M) 0.2-0.5 mg/kg SC, 0.2-0.5 mg/kg SC, 1.2-1.5 mg/kg SC IM q6-12h IM q6-12h (R); 0.2-4.0 mg/kg SC (M) 3-5 mg/kg IV — 7.5-10 mg/kg IV (R); 12-25 mg/kg IV (M) To effect To effect To effect
—
—
—
—
—
—
To effect
To effect
20-40 mg/kg IM (C)
—
50-80 mg/kg IM
Do not use
5 mg/kg PO q4-8h
5-10 mg/kg PO q12-24h
5-10 mg/kg PO q12-24h
1-5 mg/kg IM (seldom indicated; gas anesthesia preferred) —
0.1-0.5 mg/kg SC, IM
1-5 mg/kg SC, IM
2-10 mg/kg SC, IM, IP
0.2-1.0 mg/kg IM
0.2-1.0 mg/kg IM, IV
0.5-1.0 mg/kg IV
2 mg/kg IM (G); 1-5 mg/kg IM, IP (H) 0.5-1.0 mg/kg IV
5-20 mg/kg PO, SC, — IP q12-24h (R); 5-40 mg/kg SC, IP q12-24h (M) 1-5 mg/kg SC, IM, 0.5-1.0 mg/ 5 mg/kg SC, IP (R); 5-10 mg/kg kg IM (rarely IM SC, IP (M) indicated) 0.5-1.0 mg/kg IV 0.5-1.0 mg/kg IM 0.2 mg/kg IV
aGenerally
administer at 1:1 volume reversal of medetomidine (1 mg/mL) or dexmedetomidine (0.5 mg/mL). doses have been as high as 1 to 3 mg/kg SC because many rabbits possess serum atropinase. cAlthough medetomidine is no longer commercially available, it can be compounded at various concentrations through select compounding pharmacies (i.e., Diamondback Drugs, www.dia mondbackdrugs.com; Wildlife Pharmaceuticals, www.wildpharm.com). Dexmedetomidine (0.5 mg/mL) was developed to be used at the same volume as medetomidine (1 mg/mL); however, use and clinical observations in exotic pets and clinical observations have been limited. dMAC, minimum alveolar concentration. bSome
Formulary 573
574
Table A-5 Miscellaneous Agents Commonly/Occasionally Used in Small Mammals Ferrets
Rabbits
Guinea Pigs (G)/ Chinchillas (C)
Gerbils (G)/ Hamsters (H)
Rats (R)/Mice (M)
Hedgehogs
Sugar Gliders
Aminophylline
4 mg/kg PO, IM, IV q12h 0.25-0.5 mg/kg PO q24h 20-30 mg/kg SC q12h 5-10 mg/kg PO, SC, IM, IV q6-8h 0.5 mg/kg PO q8-12h
—
50 mg/kg PO, SC (G)
—
—
—
—
0.25-0.5 mg/kg PO q24h 27 mg/kg SC q6-12h
≤0.1 mg/kg PO q24h
≤0.1 mg/kg PO q24h 30 mg/kg SC q12h (G) 5-10 mg/kg PO, SC, IM, IV q6-12h 0.1-0.5 mg/kg PO q12h
≤0.1 mg/kg PO q24h —
—
—
—
—
Dexamethasone
0.5-1.0 mg/kg SC, IM, IV
0.5-2.0 mg/kg SC, IM, IV
0.5-2.0 mg/kg SC, IM, IV
0.1-1.5 mg/kg IM
Digoxin
0.005-0.01 mg/kg PO q12-24h 1.5-7.5 mg/kg PO q12h 0.5-2.0 mg/kg PO, SC, IM q8-12h
0.5-1.0 mg/kg PO (use with caution or avoid) 0.005-0.01 mg/kg PO q12-24h 0.5-1.0 mg/kg PO q8-24h 2 mg/kg PO, SC q8-12h
0.25 mg/kg q8-24h PO, IM 0.1-0.6 mg/kg SC, IM, IV
0.05-0.1 mg/kg PO q12-24h (H) 0.5-1.0 mg/kg PO q12-24h 1-2 mg/kg PO, SC q12h
—
—
—
0.5-1.0 mg/kg PO q12-24h 1-2 mg/kg PO, SC q12h
—
—
—
—
5-10 mg/kg IV, IP 0.5-1.0 mg/kg PO q24h 0.1 mg/kg IV
5-10 mg/kg IV, IP 0.5-1.0 mg/kg PO q24h 0.1 mg/kg IV
— 0.5 mg/kg PO q24h 0.003 mg/kg IV
2 mg/kg IV 0.22-0.44 mg/kg PO q24h 0.003 mg/kg IV
Benazepril Calcium EDTA Cimetidine Cisapride
Diltiazem Diphenhydramine
Doxapram Enalapril Epinephrine Furosemide
Human chorionic gonadotropin Hydroxyzine Insulin Iron dextran
30 mg/kg SC q12h
5-10 mg/kg PO, SC, 5-10 mg/kg PO, SC, IM, IV q8-12h IM, IV q6-12h 0.5 mg/kg PO q8-12h 0.5 mg/kg PO q8-12h 0.5-2.0 mg/kg SC, IM, IV (use with caution) —
0.5-1.0 mg/kg PO q12-24h 1-5 mg/kg SC, PRN; 2.0-7.5 mg/kg PO (G); 1-2 mg/kg PO, SC q12h (C) 2-5 mg/kg IV — 2-5 mg/kg IV, IP 0.25-0.5 mg/kg PO 0.1-0.5 mg/kg PO 0.5-1.0 mg/kg PO q24-48h q24-48h q24h 0.02 mg/kg SC, IM, 0.2 mg/kg IV (cardiac 0.003 mg/kg IV prn IV, IT arrest) (G); 0.1 mg/kg IV (C) 1-4 mg/kg PO, SC, 1-3 mg/kg PO 2-5 mg/kg PO, SC, IM IM, IV q8-12h q8-24h; q12h 1-4 mg/kg SC, IM, IV PRN 100 IU/animal IM, 20-25 IU/animal IV 1000 IU/animal IM, repeat in 14 days repeat in 7-10 days (G) 2 mg/kg PO q8h 2 mg/kg PO q8-12h 0.5-6 IU/kg SC (or — 1-2 IU/animal SC q12h to effect) (NPH) (G); 1 kg IU SC q12h (C) 10 mg/kg IM once 4-6 mg/kg IM once —
5-10 mg/kg PO, SC, 10 mg/kg PO IM, IV q6-12h q8h 0.1-0.5 mg/kg PO — q12h
—
2-10 mg/kg PO, SC, 2-10 mg/kg PO, SC, 2.5-5.0 mg/kg IM q12h IM q12h PO, SC, IM q8h
1-5 mg/kg PO, SC q12h
—
—
—
—
— 2 IU/animal SC
— 1-3 IU/animal SC (R)
— —
— —
—
—
25 mg/kg IM
—
Formulary
Agent
Lactulose syrup
0.15-0.75 mL /kg PO q12h
—
0.5 mL/kg PO q12h
0.5 mL/kg PO q12h 0.5 mL/kg PO q12h —
—
Lidocaine
0.5-1.0 mg/kg IV q12h 0.2-1.0 mg/kg PO, SC, IM q6-8h
1-2 mg/kg IV PRN
1-2 mg/kg IV
1-2 mg/kg IV
1-2 mg/kg IV
—
—
0.5 mg/kg PO, SC q6-8h
0.2-1.0 mg/kg PO, SC, IM q12h
0.2-1.0 mg/kg PO, SC, IM q12h
0.2-1.0 mg/kg PO, SC, IM q12h
0.2-0.5 mg/kg PO, SC
0.1-3.0 IU/kg SC, IM 1-2 mg/kg PO q12h
0.2-3.0 IU/kg SC, IM, IV 5-20 mg/kg PO, IP (G) 0.2-0.4 mg/kg PO q12h —
— —
—
0.2-0.4 mg/kg PO q12h —
—
—
—
—
Prednisolone
22 mg/kg IV
—
0.2-3.0 IU/kg SC, IM, IV 5-20 mg/kg PO, IV, IP (G) 0.2-0.4 mg/kg PO q12h 10-30 mg/kg PO q12h (G) —
0.2-3.0 IU/kg SC, IM, IV —
Potassium citrate
0.2-3.0 IU/kg SC, IM 1-2 mg/kg PO q8-12h 0.5 mg/kg PO q12h —
0.05-0.1 mg/ kg PO, SC, IM q6-12h prn —
—
—
2.5 mg/kg PO, SC, IM q12h
Prednisone
Variable; refer to reference 1 25-125 mg/kg PO q8-12h 4.25 mg/kg PO q8-12h —
0.5-2.0 mg/kg PO q12h 25 mg/kg PO q8-12h —
0.5-2.2 mg/kg PO, SC, IM 25-100 mg/kg PO q8-12h —
0.5-2.2 mg/kg PO, SC, IM 25-100 mg/kg PO q8-12h —
0.5-2.2 mg/kg PO, SC, IM 25-100 mg/kg PO q8-12h —
0.1-0.2 mg/kg PO, SC, IM q24h — —
500-1000 IU/kg IM once
500 IU/kg IM (G)
500 IU/kg IM
—
1-2 mg (thiamine)/ kg SC, IM 50-100 mg/kg PO q24h
0.02-0.4 mL/kg IM
0.02-0.2 mL/kg SC, IM —
0.02-0.2 mL/kg SC, IM —
—
17 IU vit E /rabbit
Use feline dose
1-10 mg/kg IM prn
0.02-0.2 mL/kg SC, IM 50-100 mg/animal PO, SC q24h (for deficiency) (G); 10-30 mg/kg PO, SC, IM (maintenance) (G) 0.1 mL /100-250 g SC 1-10 mg/kg IM q24h prn
10 mg/kg PO q8-12h 10 mg/kg PO, IM q12h 400 IU/kg IM q24h x 10 days 1 mL/kg SC, IM
0.1 mL /100-250 g SC 1-10 mg/kg IM q24h prn
0.1 mL /100-250 g SC 1-10 mg/kg IM q24h prn
Metoclopramide Oxytocin Phenobarbital Pimobendan
Sucralfate Theophylline Vitamin A Vitamin B complexa Vitamin C
aUse
concentration formulated for small animals.
50-100 mg/kg PO q12h
50-200 mg/kg PO, SC q24h
— 500-5000 IU/ kg IM 0.02 mL/kg SC, IM —
—
10 IU vit E /kg SC
—
2 mg/kg SC q24-72h
Formulary
Vitamin E/selenium SoSe, Schering Vitamin K
0.1-0.3 mg/kg PO q12-24h 33 mg/kg PO q8h
575
Index A
Abdominal masses in small rodents, 385 Abdominal pregnancy in rabbits, 220 Abdominal procedures in small rodents, 384–385 Abortion and resorption in rabbits, 220 Abscesses, subcutaneous in small rodents, 386 Abscesses in small rodents, 386 Acute renal failure (ARF) in ferrets, 47 Acute renal failure (ARF) in small mammals, emergency treatment of, 541, 541b Adenocarcinoma in ferrets, 129, 129f Adrenal gland surgery in ferrets, 145–150, 147f, 149f Adrenocortical neoplasms in ferrets, 105–106, 105f African hedgehogs, 411 anatomy and physiology, 411–414, 412f–413f alimentary, 412 integument, 412–413, 414f musculoskeletal, 412 neurology and behavior, 413–414, 414f reproductive, 413 anesthetic and surgical considerations, 424–425 basic procedures and preventive medicine, 415–417 clinical techniques, 416–417, 417f–418f, 417t preventive medicine, 417 restraint and examination, 415–416, 416f biology and anatomy, 411–414 anatomy and physiology, 411–414 taxonomy and natural history, 411, 412f common diseases, 418–424 cardiovascular and hematologic, 419 gastrointestinal and hepatic, 419–420 integumentary, 421–422, 421f–423f lethargic, weak, and anorectic, 424 musculoskeletal, 420 neoplastic, 422–423, 424f neurologic, 420–421, 421f nutritional, 423–424, 424f ocular injuries, 418 oral and dental, 418–419, 418f otic, 418 reproductive, 420 respiratory, 419 urinary, 420 husbandry, 414–415 breeding and neonatal care, 415 diet, 415 housing, 414–415 zoonoses and suitability as pets, 425 Agalactia in ferrets, 58 Aleutian disease in ferrets, 48, 71–73, 72f, 135–136 Allergic reactions, 564 Alopecia in guinea pigs, 305 Anal sacculectomy in ferrets, 154–155 Analgesia in small mammals, 429 in small rodents, 390
Analgesic agents used in small mammals, 571t–573t Androgen receptor blockers in ferrets, 90 Anemia in ferrets, 73–74 Anesthesia in sugar gliders, surgery and, 408–409 anesthesia, 408–409 castration and scrotal ablation, 409, 409f ovariohysterectomy, 409 removal of paracloacal glands, 409, 410f repair of patagium, 409 soft tissue surgery, 409 Anesthesia of small mammals, 429 Anesthetic agents used in small mammals, 571t–573t Anorectic diseases in African hedgehogs, 424 Anorexia in small mammals, emergency treatment of, 542–543, 543f Antifungal agents used in small mammals, 568t Antimicrobial agents used in small mammals, 567t–568t Antiparasitic agents used in small mammals, 569t–570t ARF. See Acute renal failure (ARF) Aromatase inhibitors in ferrets, 90 Arrhythmia in rabbits, 261 Auditory bulla surgery in rabbits, 276–278 lateral ear canal ablation, 276–277 osteotomy of ventral bulla, 277–278
B
Bacterial and viral pneumonia in guinea pigs, 298–299, 299f Bacterial diseases, 557–560, 558f–559f in sugar gliders, 408 Bacterial enteritis in rabbits, 199–200 enteropathogenic E. coli (EPEC), 199 miscellaneous bacterial enteritides, 200 proliferative enteritis, 199 proliferative enterocolitis, 199 proliferative enteropathy, 199 Tyzzer’s disease, 199–200 Bacterial infections in chinchillas, 320–321 Bacterial infections in rabbits, 232–236, 250–251 bacterial infections of CNS, 251 cellulitis, 234 mastitis, 233–234 methicillin-resistant staphylococcal infection, 234 moist dermatitis, 234–235 necrobacillosis, 236 otitis interna, 250–251, 251f rabbit syphilis, 235–236, 235f subcutaneous abscesses, 232–233, 233f ulcerative pododermatitis, 235 Bacterial infections of CNS in rabbits, 251 Barbering in rabbits, 241 Basal cell tumors in ferrets, 128 Behavior of small mammals, 545 behavioral training techniques for small mammals, 555 chinchillas, 553–554 behavior problems, 553–554 communication behaviors, 553
Behavior of small mammals (Continued) introduction of new conspecifics, 554 play behaviors, 553 social behavior, 553, 553f ferrets, 549–552 behavior problems, 550–551, 551b, 551f introduction of new conspecifics, 552 litter-box training, 552 play and sleeping behaviors, 549–550, 550b, 550f social behaviors, 549, 550b, 550f guinea pigs, 552–553 behavior problems, 553, 553f communication behaviors, 553 introduction of new conspecifics, 553 play behaviors, 552, 552f social behavior, 552, 552f hedgehogs, 555 communication behaviors, 555, 555f play behaviors, 555 social behaviors, 555 rabbits, 546–549 behavior problems, 546–548, 548b introduction of new conspecifics, 549 litter-box training, 548–549 mourning death of bonded mate, 549 play behaviors, 546 social behaviors, 546, 547t sugar gliders, 554–555 behavior problems, 554 communication behaviors, 554 introduction of new conspecifics, 555 play behaviors, 554, 554f social behaviors, 554 Bladder neoplasia in ferrets, 50 Blastomycosis in ferrets, 83 Bowel, prolapsed in hamsters, 387–389
C
Campylobacteriosis in ferrets, 36–37 Canine distemper in ferrets, 138 Canine distemper vaccinations in ferrets, 15 Canine distemper virus (CDV) in ferrets, 37, 78–80, 79f Carcinoma, thymic, in rabbits, 263–265 Cardiac disease in chinchillas, 318 Cardiac disease in rabbits, 257–258 arrhythmia, 261 congenital heart disease, 261 congestive heart failure (CHF), 260 diagnostic methods, 258–260, 258f–259f, 259t–260t diseases and management, 260–262 examination of rabbit, 257–258 myocardial disease, 261, 261f normal cardiovascular structure, 257 valvular disease, 261–262 vascular disease, 262 Cardiovascular and other diseases in ferrets, 62 Aleutian disease, 71–73, 72f anemia, 73–74 cardiac disease, 62 dilated cardiomyopathy, 67–68, 68f general principles, 62–67
Page numbers followed by b, indicate box; f, figure, t, table.
577
578
Index
Cardiovascular and other diseases in ferrets (Continued) diagnosis, 63 echocardiography, 64–66, 66t electrocardiography, 64, 65f, 65t history and clinical signs, 62 physical examination, 62–63 radiography, 63–64, 63f–64f treatment, 66–67, 66t–67t heartworm disease, 70–71, 70f hypertrophic cardiomyopathy (HCM), 68–69 ibuprofen toxicosis, 75 myocarditis, 69–70 neoplasia, 70 splenomegaly, 73 valvular heart disease, 69, 69f Cardiovascular in African hedgehogs, 419 Cardiovascular structure, normal in rabbits, 257 Castration in ferrets, 154 in rabbits, 271–273, 272f–273f Cataracts in sugar gliders, 406–407 Cats Helicobacter species in, 520f mycobacteriosis in, 519f Cavian leukemia in guinea pigs, 309 Cellulitis in rabbits, 234 Cerebral larva migrans in rabbits, 250 Cervical lymphadenitis in chinchillas, 336–337 in degus, 336–337 in guinea pigs, 304, 336–337 Cesarean section in small rodents, 382 Cestodes in rabbits, 202 Cheek pouch eversion in hamster, 387, 388f–389f Chemical restraints used in small mammals, 571t–573t Chinchillas, 279, 280f behavior of, 553–554 behavior problems, 553–554 communication behaviors, 553 introduction of new conspecifics, 554 play behaviors, 553 social behavior, 553, 553f biology and husbandry of, 284–289 anatomy, physiology, and behavior, 284–288, 285f, 285t dust baths, 288 gastrointestinal (gi) system, 285–286 housing, 288 husbandry, 288–289 nutrition and feeding, 288–289 reproductive behavior and breeding, 287–288 sexing, 286–287, 286f–287f urogenital system, 286 clinical techniques for, 289–292 administration of medications, 292 antibiotic therapy, 292 blood collection, 290 clinical laboratory findings, 290–291, 291t fluid therapy, 291–292 handling and restraint, 289, 289f intravenous and intraosseous catheters, 291 physical examination, 289–290 treatment techniques, 291–292 urethral catheterization and cystocentesis, 290 leukocytes in, 512f lymphocytes in, 512f neutrophils in, 512f ophthalmologic diseases in, 529
Chinchillas, disease problems in, 311 dental disorders, 313–314 dermatologic disorders, 317–318 dermatophytosis, 317 fur chewing, 313f, 317 fur slip, 318 matted fur, 318 digestive system disorders, 311–313 constipation, 312 diarrhea and soft feces, 312 dysbacteriosis, 311–312 esophageal disorders, 312–313, 313f gastroenteritis, 311–312 rectal tissue prolapse and intussusception, 312, 313f tympany, 312 disease problems, miscellaneous, 318–319 cardiac disease, 318 diabetes mellitus, 318 fractures, 318–319 hepatic lipidosis and ketosis, 318 neoplasia, 319 ear disorders, 315, 315f eye disorders, 314–315 conjunctivitis, 314 corneal disorders, 314 epiphora, 314 miscellaneous, 314–315 foot disorders, 318 infectious diseases, 319–321 bacterial infections, 320–321 fungal infections, 319 parasitic infections, 319–320 viral infections, 319 neurologic disorders, 317 heat stroke, 317 lead poisoning, 317 seizures, 317 parasitic infections, 319–320 helminthic infections, 320 protozoal infections, 319–320 reproductive system disorders, 315–316 dystocia, 316 endometriosis, 315–316 penile disorders, 316, 316f pyometra, 315–316 respiratory system disorders, 315 urinary system disorders, 316–317 Chinchillas, soft tissue surgery in, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 331f–333f ovariectomy, 327–328 ovariohysterectomy, 328–329 penile prolapse, 333 pyometra, 329 thoracotomy, 337 urolithiasis, 334–336 uterine prolapse, 330 uterine torsion, 329 uterine tumors, 330 Chondromas and chondrosarcomas in ferrets, 136 Chronic intestinal nephritis in guinea pigs, 301 Chronic renal failure in ferrets, 47 in guinea pigs, 301 Coccidia in rabbits, 201–202 Coccidioidomycosis in ferrets, 83
Computed tomography (CT), 506–508, 507f–508f Congenital heart disease in rabbits, 261 Congestive heart failure (CHF) in rabbits, 260 Conjunctivitis in chinchillas, 314 in guinea pigs, 307–308 neonatal in ferrets, 59 in rabbits, 523–525, 525f Constipation in chinchillas, 312 Contrast radiography, 503–505 Corneal disorders in chinchillas, 314 Corneal ulceration in guinea pigs, 307 Corneas in rabbits, 525, 525f Coronavirus in ferrets, 37, 37f in rabbits, 200 Critical care of small mammals, emergency and, 532 cardiopulmonary-cerebral resuscitation (CPCR), 532–535 anesthesia-related arrest, 534–535, 534t determining effectiveness of CPCR, 534 in small mammals: cardiac arrest, 533–534, 533b in small mammals: respiratory arrest, 533, 533f critical care clinical pathology, 539–540 lactate monitoring, 540 use of prothrombin and partial thromboplastin times, 540 identification and triage of critically ill patient, 532 indirect measurement of systolic blood pressure, 538–539, 539f maintenance of normothermia, 538, 538f nutritional support, 540 sedation and anesthesia of critically ill small mammal, 540–541, 542f shock and fluid therapy, 535–538 blood transfusion, 537, 537f fluid resuscitation of critically ill small mammal, 535–537, 536b–537b routes of fluid administration, 538 three phases of hypovolemic shock, 535 types of fluids, 535 use of glucocorticoids in shock, 537–538 treatment of selected common emergencies, 541–543 acute renal failure (ARF), 541, 541b anorexia, 542–543, 543f gastric stasis, 542–543, 543b, 543f respiratory distress, 541–542 urinary obstruction, 541, 543f Cryptococcosis in ferrets, 83 Cryptorchidism in ferrets, 54 in rabbits, 221, 222f Cryptosporidia in rabbits, 202 Crystalluria in sugar gliders, 405 CT. See Computed tomography (CT) Cutaneous dermal masses in chinchillas, 336 in degus, 336 in guinea pigs, 336 Cutaneous lymphoma in ferrets, 129, 129f in rabbits, 263 Cutaneous masses, miscellaneous, in small rodents, 384, 384f Cutaneous myiasis in ferrets, 125–126 Cutaneous neoplasia, in rabbits, 241, 241f
Index Cutaneous neoplasia, surgery of, in ferrets, 141–142 Cuterebra species in rabbits, 250 Cystadenomas in ferrets, 128 Cystic mastitis in rabbits, 223 Cystitis crystalluria, and urolithiasis in sugar gliders, 405 in ferrets, 49–50 in sugar gliders, 405 and urinary tract infection in guinea pigs, 301 Cystotomy in ferrets, 152 in small rodents, 384–385 Cytology of small mammals. See Hematology and cytology of small mammals
D
Degus. See also Small rodents Degus, clinical signs and treatment by species, 368–369 Degus, soft tissue surgery in, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 332f ovariectomy, 327–328 ovariohysterectomy, 328–329 penile prolapse, 333 pyometra, 329 thoracotomy, 337 urolithiasis, 334–336 uterine prolapse, 330 uterine torsion, 329 uterine tumors, 330 Dental abscesses in hamsters, 386 Dental diseases in African hedgehogs, 418–419, 418f in ferrets, 27, 28f, 468 in guinea pigs, 295–296 in rabbits, 206–207, 207f, 457–458, 457f in rodents, 465–467, 466f–467f in sugar gliders, 402, 404f Dental disorders in chinchillas, 313–314 Dentistry, small mammal, 452 diagnostic testing, 453–455 clinical examination, 453 computed tomography (ct), 454–455 imaging, 453–455 oral endoscopy, 454 other diagnostic testing, 455 radiography, 453–454, 454f equipment, 452–453 ferrets, 468–469 anatomy and physiology of skull and teeth, 468, 468f dental disease, 468 treatment and prevention, 468–469 hedgehogs, 469 rabbits, 455–462 anatomy and physiology of skull and teeth, 455 dental disease, 457–458, 457f dental procedures, 458–460, 458f, 459b facial surgery and surgical treatment, 460–462, 462f medical treatment, 458 pathophysiology of dental disease, 455–456, 456f
Dentistry, small mammal (Continued) treatment of dental disease, 458 treatment of periapical infections and abscesses, 460, 461b, 461f rodents, 462–468 anatomy and physiology of skull and teeth, 462–464, 463t, 464f clinical presentation, 465 dental disease, 465–467, 466f–467f pathophysiology of dental disease, 464–467, 464f treatment of dental disease, 467 treatment of periapical infections and abscesses, 467–468 sugar gliders, 469 Dermal fibrosis in rabbits, 242, 242f Dermatologic diseases in ferrets, 122 anatomy, physiology, and husbandry, 122–123 bacterial disease, 127, 127f diseases, 123–130 ectoparasites, 123–126 cutaneous myiasis, 125–126 fleas, 123–124 mites, 124–125 ticks, 125 endocrine disease, 130 fungal disease, 126–127 dermatophytosis, 126–127 miscellaneous fungal infections, 127 miscellaneous, 130, 130f mites, 124–125 demodectic mange, 125, 125f ear, 124 miscellaneous infections, 125 sarcoptic mange, 124–125 neoplasia, 127–130 adenocarcinoma, 129, 129f basal cell tumors, 128 cutaneous lymphoma, 129, 129f cystadenomas, 128 epitheliomas, 128, 128f mast cell tumors, 127–128, 128f miscellaneous neoplasms involving skin or subcutis, 129–130 sebaceous adenomas, 128, 128f squamous cell carcinoma (SCC), 128–129 viral disease, 126 Dermatologic disorders in sugar gliders, 405–406 Dermatological diseases in rabbits, 232 bacterial infections, 232–236 cellulitis, 234 mastitis, 233–234 methicillin-resistant staphylococcal infection, 234 moist dermatitis, 234–235 necrobacillosis, 236 rabbit syphilis, 235–236, 235f subcutaneous abscesses, 232–233, 233f ulcerative pododermatitis, 235 cutaneous neoplasia, 241, 241f fungal infections, 236–237 dermatophytosis, 236–237 parasitic infections, 237–239 ear mites, 237, 237f fleas, 238 fur mites, 237, 237f lice, 239 myiasis, 238–239 pinworms, 239 tapeworm cysts, 239 ticks, 239 skin disease, behavioral causes of, 241
579
Dermatological diseases in rabbits (Continued) barbering, 241 self-mutilation after intramuscular injection, 241 skin disease of unknown cause, 241–242 dermal fibrosis, 242, 242f Ehlers-Danlos-like syndrome, 242 eosinophilic granuloma, 242 sebaceous adenitis, 241–242, 242f viral infections, 239–241 myxomatosis, 239–240 oral papillomavirus, 240 rabbit (Shope) fibroma virus, 240 rabbit (Shope) papillomavirus, 240 rabbitpox, 240–241 Dermatophytosis in chinchillas, 317 in ferrets, 126–127 in guinea pigs, 304 in rabbits, 236–237 Diabetes mellitus in chinchillas, 318 in ferrets, 91–92 in guinea pigs, 308 Diagnostic imaging, 502 advanced treatments, 509 interventional radiology (IR), 509, 509f radiation therapy (RT), 509, 509f general considerations, 502, 503f–504f which imaging examination to perform, 502–509, 504t computed tomography (CT), 506–508, 507f–508f contrast radiography, 503–505 digital radiography, 505 image resolution, 503, 505f magnetic resonance imaging (MRI), 508 modalities, 503–509 nuclear scintigraphy, 508–509, 508f radiography, 503, 506f ultrasound, 505–506, 506f Diarrhea in chinchillas, 312 in ferrets, 36–38, 40–42, 59 in guinea pigs, 297–298 in rabbits, 197–198 Digital radiography, 505 Dilated cardiomyopathy in ferrets, 67–68, 68f DIM. See Disseminated idiopathic myofasciitis (DIM) Disease in rabbits, cardiac, 257–258 arrhythmia, 261 congenital heart disease, 261 congestive heart failure (CHF), 260 diagnostic methods, 258–260, 258f–259f, 259t–260t diseases and management, 260–262 examination of rabbit, 257–258 myocardial disease, 261, 261f normal cardiovascular structure, 257 valvular disease, 261–262 vascular disease, 262 Disease problems in chinchillas, 311 dental disorders, 313–314 dermatologic disorders, 317–318 dermatophytosis, 317 fur chewing, 313f, 317 fur slip, 318 matted fur, 318 digestive system disorders, 311–313 constipation, 312 diarrhea and soft feces, 312 dysbacteriosis, 311–312 esophageal disorders, 312–313, 313f gastroenteritis, 311–312
580
Index
Disease problems in chinchillas (Continued) rectal tissue prolapse and intussusception, 312, 313f tympany, 312 disease problems, miscellaneous, 318–319 cardiac disease, 318 diabetes mellitus, 318 fractures, 318–319 hepatic lipidosis and ketosis, 318 neoplasia, 319 ear disorders, 315, 315f eye disorders, 314–315 conjunctivitis, 314 corneal disorders, 314 epiphora, 314 miscellaneous, 314–315 foot disorders, 318 infectious diseases, 319–321 bacterial infections, 320–321 fungal infections, 319 parasitic infections, 319–320 viral infections, 319 neurologic disorders, 317 heat stroke, 317 lead poisoning, 317 seizures, 317 parasitic infections, 319–320 helminthic infections, 320 protozoal infections, 319–320 reproductive system disorders, 315–316 dystocia, 316 endometriosis, 315–316 penile disorders, 316, 316f pyometra, 315–316 respiratory system disorders, 315 urinary system disorders, 316–317 Disease problems of small rodents, 354 client education, 369 clinical signs and treatment by species, 358–369 degus, 368–369 gerbils, 367–368, 367f–368f hamsters, 364–367, 365f–366f mice, 358–361, 360f rats, 361–364, 361f–364f diagnostic challenge, 354–356 clinical examination, 355–356 medical history, 355 pet rodent etiquette, 354 reception area, 355 scheduling appointments, 354–355 diseases, 357–369 clinical signs and treatment by species, 358–369 prophylaxis for small rodents, 358, 359t seen in practice, 357 significant diseases and life spans, 357–358, 357t medications and antibiotic therapy, 369 Diseases. See also Disorders; Specific animal bacterial, 557–560, 558f–559f congenital heart in rabbits, 261 mycotic, 563 myocardial in rabbits, 261, 261f parasitic, 562–563, 562f–563f valvular in rabbits, 261–262 vascular in rabbits, 262 viral, 560–562, 561f Diseases and syndromes in sugar gliders, 402–408 bacterial diseases, 408 cataracts, 406–407 cystitis, crystalluria, and urolithiasis, 405 dental disease, 402, 404f dermatologic disorders, 405–406
Diseases and syndromes in sugar gliders (Continued) ear margin canker, 406 encephalomalacia and encephalitis, 407 endocrine alopecia, 406, 406f enteritis and enteropathy, 402 failure to thrive in a joey, 405 fractures, 407 gastrointestinal disease, 402–405 impaction of paracloacal gland, 405 infectious disease, 408 infertility, 405 malnutrition, 402 musculoskeletal disease, 407 neoplasia, 408 nephritis and renal failure, 405 neurologic disease, 407–408 nutritional osteodystrophy, 407 obesity, 407 ocular injury, 406 opthalmic disorders, 406–407 parasitic diseases, 408 pouch infection and mastitis, 405 rectal prolapse, 402–404 reproductive disorders, 405 reproductive tract infection, 405 respiratory disease, 405 retrobulbar abscess, 407 self-mutilation, 406, 406f self-mutilation of penis and scrotum, 405 stress-related disorders, 405–406 traumatic injuries, 406 tremors and seizures, 407–408 urinary tract obstruction, 405 urogenital disease, 405 Diseases in African hedgehogs, common, 418–424 cardiovascular and hematologic, 419 gastrointestinal and hepatic, 419–420 integumentary, 421–422, 421f–423f lethargic, weak, and anorectic, 424 musculoskeletal, 420 neoplastic, 422–423, 424f neurologic, 420–421, 421f nutritional, 423–424, 424f ocular injuries, 418 oral and dental, 418–419, 418f otic, 418 reproductive, 420 respiratory, 419 urinary, 420 Diseases in ferrets, dermatologic, 122 anatomy, physiology, and husbandry, 122–123 bacterial disease, 127, 127f diseases, 123–130 ectoparasites, 123–126 cutaneous myiasis, 125–126 fleas, 123–124 mites, 124–125 ticks, 125 endocrine disease, 130 fungal disease, 126–127 dermatophytosis, 126–127 miscellaneous fungal infections, 127 miscellaneous, 130, 130f mites, 124–125 demodectic mange, 125, 125f ear, 124 miscellaneous infections, 125 sarcoptic mange, 124–125 neoplasia, 127–130 adenocarcinoma, 129, 129f basal cell tumors, 128 cutaneous lymphoma, 129, 129f
Diseases in ferrets, dermatologic (Continued) cystadenomas, 128 epitheliomas, 128, 128f mast cell tumors, 127–128, 128f miscellaneous neoplasms involving skin or subcutis, 129–130 sebaceous adenomas, 128, 128f squamous cell carcinoma (SCC), 128–129 viral disease, 126 Diseases in ferrets, musculoskeletal and neurologic, 132 ataxia, 132–133 intracranial disorders, 138 canine distemper, 138 osteoma, 138 rabies, 138 miscellaneous diseases, 139 miscellaneous metabolic diseases, 139 musculoskeletal disorders, 138–139 disseminated idiopathic myofasciitis (DIM), 138–139 myasthenia gravis, 139 neoplasia of central nervous system, 138 neuronal vacuolation, 138 posterior paresis, 132 seizures, 133–135, 134f–137f spinal disorders, 135–138 Aleutian disease, 135–136 intervertebral disk disease, 138 spinal tumors, 136–138 spinal tumors, 136–138 chondromas and chondrosarcomas, 136 fibrosarcoma, 137–138 lymphoma, 137 Diseases in ferrets, respiratory, 78 canine distemper virus (CDV), 78–80, 79f influenza, 80–81, 81t pneumonia, 81–83, 82f pulmonary mycoses, 83 blastomycosis, 83 coccidioidomycosis, 83 cryptococcosis, 83 history and physical examination, 83 respiratory diseases, other causes of respiratory signs, 83, 83f Diseases in guinea pigs, 295 dermatologic diseases, 303–305 alopecia, 305 cervical lymphadenitis, 304 dermatophytosis, 304 ectoparasites, 304, 304f neoplasia, 305, 305f pododermatitis, 305, 305f gastrointestinal (GI) and hepatic diseases, 295–299 bacterial and viral pneumonia, 298–299, 299f dental disease, 295–296 diarrhea, 297–298 enteritis, 297–298 fecal impaction, 298, 298f gastrointestinal (GI) hypomotility or stasis, 296–297, 296f hepatic lipidosis, 298 neoplasia, 298 noninfectious respiratory diseases, 299 respiratory diseases, 298 miscellaneous diseases, 308–309 diabetes mellitus, 308 heat stress, 308–309 lymphosarcoma/cavian leukemia, 309 ototoxicity, 308 musculoskeletal diseases, 305–307 fibrosis osteodystrophy, 306–307 iatrogenic muscle necrosis, 307
Index Diseases in guinea pigs (Continued) metastatic mineralization, 307 nutritional muscular dystrophy, 307 osteoarthritis and osteoarthrosis, 306 scurvy, 305–306, 306f vitamin C deficiency, 305–306, 306f neurologic diseases, 307 lymphocytic choriomeningitis virus (LCMV), 307 mites, 307 otitis media/otitis interna, 307, 308f rabies, 307 ophthalmologic diseases, 307–308 conjunctivitis, 307–308 corneal ulceration, 307 pea eye, 307, 309f reproductive diseases, 301–303 dystocia, 302 epididymitis, 303 mastitis, 303 metritis, 303 neoplasia, 303 orchitis, 303 ovarian cysts, 301–302, 301f–302f pyometra, 303 scrotal plugs, 303 toxemia of pregnancy, 302–303 uterine prolapse, 302 vaginitis, 303 urinary diseases, 299–301 chronic intestinal nephritis, 301 chronic renal failure, 301 cystitis and urinary tract infection, 301 miscellaneous uropathies, 301 urolithiasis, 299–301, 300f Diseases in rabbits, dermatological, 232 bacterial infections, 232–236 cellulitis, 234 mastitis, 233–234 methicillin-resistant staphylococcal infection, 234 moist dermatitis, 234–235 necrobacillosis, 236 rabbit syphilis, 235–236, 235f subcutaneous abscesses, 232–233, 233f ulcerative pododermatitis, 235 cutaneous neoplasia, 241, 241f fungal infections, 236–237 dermatophytosis, 236–237 parasitic infections, 237–239 ear mites, 237, 237f fleas, 238 fur mites, 237, 237f lice, 239 myiasis, 238–239 pinworms, 239 tapeworm cysts, 239 ticks, 239 skin disease, behavioral causes of, 241 barbering, 241 self-mutilation after intramuscular injection, 241 skin disease of unknown cause, 241–242 dermal fibrosis, 242, 242f Ehlers-Danlos-like syndrome, 242 eosinophilic granuloma, 242 sebaceous adenitis, 241–242, 242f viral infections, 239–241 myxomatosis, 239–240 oral papillomavirus, 240 rabbit (Shope) fibroma virus, 240 rabbit (Shope) papillomavirus, 240 rabbitpox, 240–241
Diseases in rabbits, gastrointestinal (GI), 193 acute gastrointestinal (GI) dilation or obstruction, 196–197, 196f aflatoxicosis, 203 cecoliths, 198 cecotrophy and intermittent diarrhea, 197–198 enteritis complex and enterotoxemia, 198–200 bacterial enteritis, 199–200 dysbiosis caused by treatment with antibiotics, 198–199 enteropathogenic E. coli (EPEC), 199 miscellaneous bacterial enteritides, 200 mucoid enteritis, 198 prevention of enterotoxemia, 199 proliferative enteritis, 199 proliferative enterocolitis, 199 proliferative enteropathy, 199 treatment of enteritis, 199 Tyzzer’s disease, 199–200 gastrointestinal (GI) stasis syndrome, 193–196 gastrointestinal (GI) stress stasis syndrome diagnostic testing, 195, 195f effect of diet and cecocolic motility, 194 history and clinical signs, 194–195 physical examination findings, 195 role of fiber, 193–194 treatment, 196 liver lobe torsion, 202–203 neoplasia, 202 parasitic disorders of gastrointestinal (GI) tract, 201–202 cestodes, 202 coccidia, 201–202 cryptosporidia, 202 helminths, 202 hepatic coccidia, 201 intestinal coccidia, 201–202 miscellaneous protozoa, 202 nematodes, 202 trematodes, 202 viral diseases of digestive tract, 200–201 hemorrhagic disease virus (RHDV), 200–201 papillomatosis, 200 rabbit enteric coronavirus, 200 rotavirus, 200 Diseases in rabbits, neurologic and musculoskeletal, 245, 246f, 246t bacterial infection, 250–251 bacterial infections of CNS, 251 otitis interna, 250–251, 251f degenerative/developmental, 252 osteoarthritis, 252 splay leg, 252 spondylosis, 252, 254f metabolic, 254 heat stroke, 254 toxemia of pregnancy, 254 miscellaneous causes, 255 idiopathic, 255 miscellaneous, 255 neoplastic, 255 vascular, 255 nutritional issues, 254–255 parasitic infection, 246–250 cerebral larva migrans, 250 cuterebra species, 250 encephalitozoonosis, 246–250, 247f, 248t toxoplasmosis, 250
581
Diseases in rabbits, neurologic and musculoskeletal (Continued) toxic, 253–254 fipronil toxicosis, 253–254 lead toxicosis, 253 pyrethrin/permethrin toxicosis, 254 traumatic, 251–252 vertebral fracture or luxation, 251–252, 252f–253f viral infection, 251 herpes simplex virus (HSV), 251 rabies, 251 Diseases in rabbits, respiratory, 205 anatomy of respiratory tract, 205 diagnosis and differentiation, 208–209 diagnostic imaging, 209, 209f–211f diseases producing secondary respiratory systems, 208 laboratory analysis, 208–209, 208f lower respiratory tract diseases, 207–208 infectious, 207 neoplastic, 207–208 physical examination, 208 prevention and control of respiratory disease, 214 serology and molecular diagnostic testing, 209 treatment of respiratory disease, 209–214, 212f–215f upper respiratory tract diseases, 205–207 bacterial pathogens, 205–206 dental disease, 206–207, 207f fungal pathogens, 206 infectious, 205–206 miscellaneous conditions, 207 neoplastic, 207 noninfectious, 206–207 trauma, 206, 207f viral pathogens, 206 Diseases in small pet mammals, ophthalmologic, 523 chinchillas, 529 ferrets, 527–528 guinea pigs, 528–529, 528f–529f hamsters, 529–530 mice, 529–530 rabbits, 523–527, 524f conjunctivitis, 523–525, 525f cornea, 525, 525f diseases of, 525–526, 526f epiphora, 523–525, 525f glaucoma, 526 orbit, 526–527, 527f uveitis and diseases of lens, 525–526, 526f rats, 529–530 sugar gliders, 530 Disorders in rabbits, lymphoproliferative, 257, 262–267 chemotherapy, 265–266 cutaneous lymphoma, 263 diagnosis, 265 etiology, 262 leukemia, 263 multicentric lymphoma, 262–263 thymic masses, 263–265 thymoma/thymic lymphoma, thymic carcinoma, 263–265 treatment, 265–267 Disseminated idiopathic myofasciitis (DIM) in ferrets, 138–139 Dysbacteriosis in chinchillas, 311–312
582
Index
Dystocia in chinchillas, 316, 329–330 in degus, 329–330 in ferrets, 57 in guinea pigs, 302, 329–330 in rabbits, 220
E
Ear canal surgery in rabbits, 276–278 lateral ear canal ablation, 276–277 osteotomy of ventral bulla, 277–278 Ear margin canker in sugar gliders, 406 Ear mites in rabbits, 237, 237f Ectoparasites in ferrets, 17, 123–126 cutaneous myiasis, 125–126 fleas, 123–124 mites, 124–125 ticks, 125 in guinea pigs, 304, 304f Ehlers-Danlos-like syndrome in rabbits, 242 Emergency and critical care of small mammals, 532 cardiopulmonary-cerebral resuscitation (CPCR), 532–535 anesthesia-related arrest, 534–535, 534t determining effectiveness of cpcr, 534 in small mammals: cardiac arrest, 533–534, 533b in small mammals: respiratory arrest, 533, 533f critical care clinical pathology, 539–540 lactate monitoring, 540 use of prothrombin and partial thromboplastin times, 540 identification and triage of critically ill patient, 532 indirect measurement of systolic blood pressure, 538–539, 539f maintenance of normothermia, 538, 538f nutritional support, 540 sedation and anesthesia of critically ill small mammal, 540–541, 542f shock and fluid therapy, 535–538 blood transfusion, 537, 537f fluid resuscitation of critically ill small mammal, 535–537, 536b–537b routes of fluid administration, 538 three phases of hypovolemic shock, 535 types of fluids, 535 use of glucocorticoids in shock, 537–538 treatment of selected common emergencies, 541–543 acute renal failure (ARF), 541, 541b anorexia, 542–543, 543f gastric stasis, 542–543, 543b, 543f respiratory distress, 541–542 urinary obstruction, 541, 543f Encephalitozoonosis in rabbits, 228, 246–250, 247f, 248t Encephalomalacia and encephalitis in sugar gliders, 407 Endocrine alopecia in sugar gliders, 406, 406f Endocrine diseases in ferrets, 86 adrenal gland disease, 86–91 adrenal histopathology, 91 androgen receptor blockers, 90 aromatase inhibitors, 90 clinical pathology and diagnostic testing, 88–89, 89t gonadotropin-releasing hormone (GnRH) analogs, 90 history and physical examination, 87, 87f–88f
Endocrine diseases in ferrets (Continued) ketoconanazole, 90 management, 89–90 medical management, 90 melatonin, 90 mitotane, 90 possible concurrent abnormalities, 89 prognosis, 91 surgical therapy, 89–90 clinical signs, 93t diabetes mellitus, 91–92 pancreatic islet cell tumors, 92–99 advanced imaging, 96 baseline insulin and glucose concentrations, 95, 95t beyond Whipple’s triad, 95 clinical features, 93–94 clinical pathologic abnormalities, 94 clinical signs, 93–94 diagnostic approach, 95–96 diagnostic imaging, 96 differential diagnoses for fasting hypoglycemia, 94–95 etiology, 92–93 histopathology, 99 insulin: glucose ratios, 95 management of islet cell tumors, 96–99 medical management of chronic hypoglycemia, 98–99 medical therapies for active hypoglycemic crisis, 97–98 miscellaneous testing, 96 pathophysiology, 93 physical examination, 94, 94f prognosis, 99 provocative testing, 95–96 radiography, 96 signalment, 93 surgical therapy, 96–97, 96t–97t ultrasonography, 96, 96f pheochromocytomas, 91, 91f thyroid disease, 91, 92t Endometrial hyperplasia or uterine polyps in rabbits, 218–219 Endometrial venous aneurysms in rabbits, 221 Endometriosis in chinchillas, 315–316 Endometritis in rabbits, 219 Endoparasites in ferrets, 16–17 Enteritis and enteropathy in sugar gliders, 402 in guinea pigs, 297–298 Enterotomy in rabbits, 273–274, 274f Enucleation and exenteration in small rodents, 386–387 Eosinophilic gastroenteritis in ferrets, 37–38 Eosinophilic granuloma in rabbits, 242 Epididymitis in guinea pigs, 303 in rabbits, 222 Epiphora in chinchillas, 314 in rabbits, 523–525, 525f Epitheliomas in ferrets, 128, 128f Esophageal disease in ferrets, 28–29, 29f Exotic mammal diagnostic and surgical endoscopy, 485 anesthesia, 485–486 complications, 501 instrumentation, 486, 487t, 488f–489f outcome, 501 patient evaluation, 485 patient selection, 485 postoperative care, 501 procedures, 486–501
Exotic mammal diagnostic and surgical endoscopy (Continued) endotracheal intubation, 488, 493f gastroscopy/colonoscopy, 493–496, 497f–498f laparoscopy, 496–499, 499f–500f otoscopy, 486, 489f rhinoscopy, 488–493, 494f–496f stomatoscopy, 486–488, 490f–492f thoracoscopy, 499–501 tracheobronchoscopy, 488, 494f vaginoscopy/cystoscopy, 493, 497f
F
Failure to thrive in joey in sugar gliders, 405 Fecal impaction in guinea pigs, 298, 298f Ferrets behavior of, 549–552 behavior problems, 550–551, 551b, 551f introduction of new conspecifics, 552 litter-box training, 552 play and sleeping behaviors, 549–550, 550b, 550f social behaviors, 549, 550b, 550f benign sebaceous epithelial structure in, 518f chordoma in, 520f diarrhea, emaciation with, 41–42 diarrhea in, 40–41 enteritis and diarrhea, 36–38 campylobacteriosis, 36–37 eosinophilic gastroenteritis, 37–38 inflammatory bowel disease (IBD), 37–38 mycobacteriosis, 36 salmonellosis, 36 viral diarrhea, 37 extramedullary hematopoiesis (EMH) in, 519f leukocytes in, 512f–513f lymphoid hyperplasia in, 516f lymphosarcoma in, 516f malignant epithelial tumor in, 517f malignant neuroendocrine neoplasm in, 519f mast cell neoplasm in, 520f modified transudate in, 521f ophthalmologic diseases in, 527–528 plasma cell tumor in, 521f platelet clumping in, 513f reticulocytes in, 515f viral diarrhea, 37 canine distemper virus, 37 coronavirus, 37, 37f influenza virus, 37 rotavirus, 37 Ferrets, anatomy, physiology, and husbandry of, 1 anatomy and physiology, 3–9, 3f–6f, 7t cardiovascular and lymphatic systems, 8 endocrine system, 8 gastrointestinal system, 6–7 integument, 3 musculoskeletal system, 8–9 neurologic system and special senses, 9 respiratory system, 8 urogenital system, 8 cardiovascular and lymphatic systems, 8 heart and blood vessels, 8 lymphatic structures, 8 domestication history, 1–2 endocrine system, 8 adrenal glands, 8 parathyroid glands, 8 thyroid glands, 8
Index Ferrets, anatomy, physiology, and husbandry of (Continued) gastrointestinal system, 6–7 esophagus, 6–7 gallbladder, 7 intestines, 6–7 liver, 7 pancreas, 7 salivary glands, 6 stomach, 6–7 teeth, 6 husbandry, 9–11 environmental enrichment, 10 housing, 9–10 nutrition, 10–11 integument, 3 anal glands, 3 coat, 3, 7f skin and associated glands, 3 musculoskeletal system, 8–9 neurologic system and special senses, 9 brain, 9 special senses, 9 spinal cord, 9 physiology and reproduction, 9 body size and seasonal weight variation, 9 physiology, 7t, 9 reproduction, 9 special senses, 9 hearing, 9 sight, 9 taste and olfaction, 9 urogenital system, 8 bladder, 8 female reproductive tract, 8 kidneys, 8 male reproductive tract, 8 ureters, 8 uses, 2 Ferrets, cardiovascular and other diseases in, 62 Aleutian disease, 71–73, 72f anemia, 73–74 cardiac disease, 62 dilated cardiomyopathy, 67–68, 68f general principles, 62–67 diagnosis, 63 echocardiography, 64–66, 66t electrocardiography, 64, 65f, 65t history and clinical signs, 62 physical examination, 62–63 radiography, 63–64, 63f–64f treatment, 66–67, 66t–67t heartworm disease, 70–71, 70f hypertrophic cardiomyopathy (HCM), 68–69 ibuprofen toxicosis, 75 myocarditis, 69–70 neoplasia, 70 splenomegaly, 73 valvular heart disease, 69, 69f Ferrets, dentistry of, 468–469 anatomy and physiology of skull and teeth, 468, 468f dental disease, 468 treatment and prevention, 468–469 Ferrets, dermatologic diseases in, 122 anatomy, physiology, and husbandry, 122–123 bacterial disease, 127, 127f diseases, 123–130 ectoparasites, 123–126 cutaneous myiasis, 125–126 fleas, 123–124
Ferrets, dermatologic diseases in (Continued) mites, 124–125 ticks, 125 endocrine disease, 130 fungal disease, 126–127 dermatophytosis, 126–127 miscellaneous fungal infections, 127 miscellaneous, 130, 130f mites, 124–125 demodectic mange, 125, 125f ear, 124 miscellaneous infections, 125 sarcoptic mange, 124–125 neoplasia, 127–130 adenocarcinoma, 129, 129f basal cell tumors, 128 cutaneous lymphoma, 129, 129f cystadenomas, 128 epitheliomas, 128, 128f mast cell tumors, 127–128, 128f miscellaneous neoplasms involving skin or subcutis, 129–130 sebaceous adenomas, 128, 128f squamous cell carcinoma (SCC), 128–129 viral disease, 126 Ferrets, endocrine diseases in, 86 adrenal gland disease, 86–91 adrenal histopathology, 91 androgen receptor blockers, 90 aromatase inhibitors, 90 clinical pathology and diagnostic testing, 88–89, 89t gonadotropin-releasing hormone (GnRH) analogs, 90 history and physical examination, 87, 87f–88f ketoconanazole, 90 management, 89–90 medical management, 90 melatonin, 90 mitotane, 90 possible concurrent abnormalities, 89 prognosis, 91 surgical therapy, 89–90 clinical signs, 93t diabetes mellitus, 91–92 pancreatic islet cell tumors, 92–99 advanced imaging, 96 baseline insulin and glucose concentrations, 95, 95t beyond Whipple’s triad, 95 clinical features, 93–94 clinical pathologic abnormalities, 94 clinical signs, 93–94 diagnostic approach, 95–96 diagnostic imaging, 96 differential diagnoses for fasting hypoglycemia, 94–95 etiology, 92–93 histopathology, 99 insulin: glucose ratios, 95 management of islet cell tumors, 96–99 medical management of chronic hypoglycemia, 98–99 medical therapies for active hypoglycemic crisis, 97–98 miscellaneous testing, 96 pathophysiology, 93 physical examination, 94, 94f prognosis, 99 provocative testing, 95–96 radiography, 96 signalment, 93 surgical therapy, 96–97, 96t–97t ultrasonography, 96, 96f
583
Ferrets, endocrine diseases in (Continued) pheochromocytomas, 91, 91f thyroid disease, 91, 92t Ferrets, gastrointestinal (GI) diseases in, 27 dental disease, 27, 28f eosinophilic gastroenteritis, 37–38 esophageal disease, 28–29, 29f gastritis and ulceration, 29–33, 30f helicobacter mustelae gastritis, 30–33, 31f, 32t gastrointestinal (GI) disorders, 27 gastrointestinal (GI) foreign bodies, 33–35, 34f–35f, 34t gastrointestinal (GI) parasitism, 36 gastrointestinal (GI) polyps, 33 gastric distention (bloat), 33, 33f inflammatory bowel disease (IBD), 37–38 liver disease, 35–36 neoplasia, 40 oral neoplasia, 28, 28f proliferative bowel disease, 38–39, 39f rectal disease, 39–40 salivary mucocele, 28, 28f vomiting, 40 Ferrets, musculoskeletal and neurologic diseases in, 132 ataxia, 132–133 intracranial disorders, 138 canine distemper, 138 osteoma, 138 rabies, 138 miscellaneous diseases, 139 miscellaneous metabolic diseases, 139 musculoskeletal disorders, 138–139 disseminated idiopathic myofasciitis (DIM), 138–139 myasthenia gravis, 139 neoplasia of central nervous system, 138 neuronal vacuolation, 138 posterior paresis, 132 seizures, 133–135, 134f–137f spinal disorders, 135–138 Aleutian disease, 135–136 intervertebral disk disease, 138 spinal tumors, 136–138 spinal tumors, 136–138 chondromas and chondrosarcomas, 136 fibrosarcoma, 137–138 lymphoma, 137 Ferrets, neoplasia in, 103 endocrine system tumors, 104–106 adrenocortical neoplasms, 105–106, 105f insulinoma, 104–105 thyroid neoplasms, 106 etiology, 103–104 gastrointestinal (GI) tract tumors, 116–117, 117f hemolymphatic system tumors, 106–115 ancillary treatments, 114–115 chemotherapy, 110–112, 112t–115t classification of lymphoma, 106–107, 106t–107t, 107f cytologic/histologic description, 108–110, 110f–111f diagnostic imaging, 108, 109f laboratory evaluation, 108 palliative therapy, 112–113 radiation treatment, 113–114 signalment and clinical signs, 107–108, 108f treatment, 110–115 incidence and behavior, 104 malignant peripheral nerve sheath tumor, 119
584
Index
Ferrets, neoplasia in (Continued) miscellaneous neoplasms, 120 musculoskeletal system tumors, 118 nervous system tumors, 118–119, 119f reproductive tract tumors, 118 respiratory system tumors, 119 skin tumors, 115–116, 115f urinary system tumors, 119 vascular neoplasms, 119–120 Ferrets, respiratory diseases in, 78 canine distemper virus (CDV), 78–80, 79f influenza, 80–81, 81t other causes of respiratory signs, 83, 83f pneumonia, 81–83, 82f pulmonary mycoses, 83 blastomycosis, 83 coccidioidomycosis, 83 cryptococcosis, 83 history and physical examination, 83 Ferrets, soft tissue surgery in, 141 caval syndrome, 155 endocrine system, 145–152 gallbladder surgery, 147f pancreatic surgery, 150–151, 151f splenectomy, 151–152, 152f surgery of adrenal gland, 145–150, 147f, 149f exploratory laparotomy, 142–143, 143f gastrointestinal (GI) system, 143–145 gallbladder surgery, 145 intestinal surgery, 143–144, 144f liver biopsy, 144–145, 145f–146f salivary mucocele resection, 143 general surgical principles, 141, 142f heartworm disease, 155 miscellaneous surgical procedures, 154–155 anal sacculectomy, 154–155 surgery of cutaneous neoplasia, 141–142 urogenital system, 152–154 castration, 154 cystotomy, 152 nephrectomy, 152 ovarian remnant, 154 ovariohysterectomy, 154 paraurethral/prostatic cysts, 153–154, 153f perineal urethrostomy, 153, 153f preputial masses, 154, 155f pyometra, 154 Ferrets, urinary and reproductive systems disorders in, 46 female ferret, 54–56 female reproductive tract tumors, 55–56 hydrometra, 56, 56f hyperestrogenism, 55 mucometra, 55–56 pyometra, 55–56 jill diseases, 57–58 agalactia, 58 dystocia, 57 mammary gland neoplasia, 58 mastitis, 58 metritis, 58 postparturient hypocalcemia, 58 pregnancy toxemia, 57 pseudopregnancy, 57 kit diseases, 58–59 caring for ill kits, 58–59, 59t diarrhea, 59 enlarged umbilical cords, 59 neonatal conjunctivitis, 59 neonatal mortality and deformities, 59 normal kit, 58 splay-legged kits, 59
Ferrets, urinary and reproductive systems disorders in (Continued) male ferret, 54 cryptorchidism, 54 male reproductive tract tumors, 54, 54f penile lesions, 54 prostatic lesions, 54 periparturient disease, 56–59 breeding ferrets, management of, 57 jill diseases, 57–58 kit diseases, 58–59 normal breeding, 56–57 polycystic kidney disease, polycystic kidney in ferret with acute renal failure, 47f renal disease and renal failure, 47–48 acute renal failure, 47 chronic renal failure, 47 diagnosis and treatment, 47–48 reproductive system disorders, 54–56 female ferret, 54–56 male ferret, 54 urethral obstruction urethral catheterization of male ferret, 51f urethral obstruction, 50f urinary and reproductive systems disorders periparturient disease, 56–59 reproductive system disorders, 54–56 urinary system disorders, 46–54 urinary system disorders, 46–54 Aleutian disease, 48 bladder neoplasia, 50 cystitis, 49–50 hydronephrosis, 47 nephrocalcinosis, 48 paraurethral cysts or paraurethral disease, 53–54 polycystic kidney disease, 46–47 prostatic cysts, 51–52, 51f–52f prostatic tumors, 53 prostatitis and prostatic abscess, 53, 53f pyelonephritis, 48 renal cysts, 46 renal disease and renal failure, 47–48 renal neoplasia, 48 ureteral rupture, 48 urethral obstruction, 50–51 urinary incontinence, 50 urolithiasis, 49 Ferrets, veterinary care, approaches to, 13 basic approach to veterinary care clinical and treatment techniques, 17–25 hospitalization, 17 preventive medicine, 15–17 clinical and treatment techniques, 17–25 antibiotic and drug therapy, 21 blood pressure monitoring, 24–25 blood transfusion, 24 bone marrow collection, 24, 24f diagnostic peritoneal lavage, 25 fluid therapy, 21 intravenous catheters, 20–21, 21f nutritional support, 22 pain management, 21–22 splenic aspiration, 23 tracheal wash, 24 urinary catheterization, 23 urine collection and urinalysis, 22–23, 23t venipuncture, 17–20, 18f–19f parasites, 16–17 ectoparasites, 17 endoparasites, 16–17
Ferrets, veterinary care, approaches to (Continued) preventive medicine, 15–17 parasites, 16–17 vaccinations, 15–16 restraint and physical examination, 13–15 physical examination, 14–15 restraint, 13–14, 14f vaccinations, 15–16 canine distemper, 15 rabies, 15–16 vaccine-associated adverse events, 16 venipuncture, 17–20, 18f–19f reference ranges, 18–20, 19t–20t Fetuses, retained, in rabbits, 220 Fibrosarcoma, of ferrets, 137–138 Fibrosis osteodystrophy in guinea pigs, 306–307 Fipronil toxicosis in rabbits, 253–254 Fleas in ferrets, 123–124 in rabbits, 238 Formulary, 566 analgesic agents used in small mammals, 571t–573t anesthetic agents used in small mammals, 571t–573t antifungal agents used in small mammals, 568t antimicrobial agents used in small mammals, 567t–568t antiparasitic agents used in small mammals, 569t–570t chemical restraints used in small mammals, 571t–573t miscellaneous agents used in small mammals, 574t–575t Fractures in chinchillas, 318–319 in sugar gliders, 407 Fungal disease in ferrets, 126–127 dermatophytosis, 126–127 miscellaneous fungal infections, 127 Fungal infections in chinchillas, 319 in rabbits, 236–237 Fur chewing in chinchillas, 313f, 317 Fur mites in rabbits, 237, 237f Fur slip in chinchillas, 318
G
Gallbladder surgery in ferrets, 145, 147f Gastric stasis in small mammals, emergency treatment of, 542–543, 543f Gastric trichobezoars in chinchillas, 333–334 in degus, 333–334 in guinea pigs, 333–334 Gastroenteritis in chinchillas, 311–312 Gastrointestinal (GI) diseases in African hedgehogs, 419–420 Gastrointestinal (GI) diseases in rabbits, 193 acute gastrointestinal (GI) dilation or obstruction, 196–197, 196f aflatoxicosis, 203 cecoliths, 198 cecotrophy and intermittent diarrhea, 197–198 enteritis complex and enterotoxemia, 198–200 bacterial enteritis, 199–200 dysbiosis caused by treatment with antibiotics, 198–199 enteropathogenic E. coli (EPEC), 199
Index Gastrointestinal (GI) diseases in rabbits (Continued) miscellaneous bacterial enteritides, 200 mucoid enteritis, 198 prevention of enterotoxemia, 199 proliferative enteritis, 199 proliferative enterocolitis, 199 proliferative enteropathy, 199 treatment of enteritis, 199 Tyzzer’s disease, 199–200 gastrointestinal (GI) stasis syndrome, 193–196 gastrointestinal (GI) stress stasis syndrome diagnostic testing, 195, 195f effect of diet and cecocolic motility, 194 history and clinical signs, 194–195 physical examination findings, 195 role of fiber, 193–194 treatment, 196 liver lobe torsion, 202–203 neoplasia, 202 parasitic disorders of gastrointestinal (GI) tract, 201–202 cestodes, 202 coccidia, 201–202 cryptosporidia, 202 helminths, 202 hepatic coccidia, 201 intestinal coccidia, 201–202 miscellaneous protozoa, 202 nematodes, 202 trematodes, 202 viral diseases of digestive tract, 200–201 hemorrhagic disease virus (RHDV), 200–201 papillomatosis, 200 rabbit enteric coronavirus, 200 rotavirus, 200 Gastrointestinal (GI) diseases in sugar gliders, 402–405 Gastrointestinal (GI) parasitism in ferrets, 36 Gastrointestinal (GI) physiology and nutrition of rabbits, 183 dietary components, 188–191 commercial mixes and pellets, 190–191 edible plants, 190 fresh vegetables, 190 grass, 188 greens, 190 hay, 188–190 miscellaneous feed items, 191 water, 191 dietary recommendations summary, 191 gastrointestinal physiology, 183–188 cecotrophy, 186–187, 186f–187f digestion and absorption, 187 energy requirements, 187–188 hindgut flora and fermentation, 186 ingestion of food, 183–184 large intestine, 184–185, 185f motility, 187 small intestine, 184, 184f stomach, 184, 184f nutrition, 188 carbohydrate, 188 fat, 188 fiber, 188 minerals, 188, 189t protein, 188 vitamins, 188, 189t Gastrointestinal (GI) polyps in ferrets, 33 gastric distention (bloat), 33, 33f
Gastrointestinal (GI) surgery in rabbits, 273–274 anastomosis, 273–274, 274f enterotomy, 273–274, 274f gastrotomy, 273 intestinal resection, 273–274, 274f surgery of large bowel, 274 Gastrointestinal (GI) tract tumors in ferrets, 116–117, 117f Gastrotomy in rabbits, 273 Gerbils. See also Small rodents anatomic and physiologic characteristics, 342 clinical signs and treatment by species, 367–368, 367f–368f general characteristics, 340 Glaucoma in rabbits, 526 Gonadotropin-releasing hormone (GnRH) analogs in ferrets, 90 Guinea pigs, 279, 280f behavior of, 552–553 behavior problems, 553, 553f communication behaviors, 553 introduction of new conspecifics, 553 play behaviors, 552, 552f social behavior, 552, 552f benign mammary eipthelial hyperplasia in, 517f biology and husbandry of, 279–284, 280f anatomy and physiology, 280–283, 281f behavior, 283 gastrointestinal system, 281–282 housing, 283–284 husbandry, 283–284 nutrition and feeding, 284 reproductive behavior and breeding, 282–283, 282t sexing, 281f, 282 urogenital system, 282 clinical techniques for, 289–292 administration of medications, 292 antibiotic therapy, 292 blood collection, 290, 290f clinical laboratory findings, 290–291, 291t fluid therapy, 291–292 handling and restraint, 289 intravenous and intraosseous catheters, 291, 292f physical examination, 289–290 treatment techniques, 291–292 urethral catheterization and cystocentesis, 290 formalin contamination in, 522f leukocytes in, 513f lymphocytes with Kurloff’s bodies in, 514f ophthalmologic diseases in, 528–529, 528f–529f Guinea pigs, diseases of, 295 dermatologic diseases, 303–305 alopecia, 305 cervical lymphadenitis, 304 dermatophytosis, 304 ectoparasites, 304, 304f neoplasia, 305, 305f pododermatitis, 305, 305f gastrointestinal (GI) and hepatic diseases, 295–299 bacterial and viral pneumonia, 298–299, 299f dental disease, 295–296 diarrhea, 297–298 enteritis, 297–298 fecal impaction, 298, 298f gastrointestinal (GI) hypomotility or stasis, 296–297, 296f
585
Guinea pigs, diseases of (Continued) hepatic lipidosis, 298 neoplasia, 298 noninfectious respiratory diseases, 299 respiratory diseases, 298 miscellaneous diseases, 308–309 diabetes mellitus, 308 heat stress, 308–309 lymphosarcoma/cavian leukemia, 309 ototoxicity, 308 musculoskeletal diseases, 305–307 fibrosis osteodystrophy, 306–307 iatrogenic muscle necrosis, 307 metastatic mineralization, 307 nutritional muscular dystrophy, 307 osteoarthritis and osteoarthrosis, 306 scurvy, 305–306, 306f vitamin C deficiency, 305–306, 306f neurologic diseases, 307 lymphocytic choriomeningitis virus (LCMV), 307 mites, 307 otitis media/otitis interna, 307, 308f rabies, 307 ophthalmologic diseases, 307–308 conjunctivitis, 307–308 corneal ulceration, 307 pea eye, 307, 309f reproductive diseases, 301–303 dystocia, 302 epididymitis, 303 mastitis, 303 metritis, 303 neoplasia, 303 orchitis, 303 ovarian cysts, 301–302, 301f–302f pyometra, 303 scrotal plugs, 303 toxemia of pregnancy, 302–303 uterine prolapse, 302 vaginitis, 303 urinary diseases, 299–301 chronic intestinal nephritis, 301 chronic renal failure, 301 cystitis and urinary tract infection, 301 miscellaneous uropathies, 301 urolithiasis, 299–301, 300f Guinea pigs, soft tissue surgery in, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 332f–333f ovariectomy, 327–328 ovariohysterectomy, 328–329, 328f penile prolapse, 333 pyometra, 329 thoracotomy, 337 urolithiasis, 334–336, 335f uterine prolapse, 330, 330f uterine torsion, 329 uterine tumors, 330
H
Hamsters. See also Small rodents anatomic and physiologic characteristics, 342 cheek pouch eversion in, 387, 388f–389f clinical signs and treatment by species, 364–367, 365f–366f dental abscesses in, 386 general characteristics, 340 lymphosarcoma in, 516f
586
Index
Hamsters (Continued) malignant melanoma in, 518f malignant mesenchymal tumor in, 518f ophthalmologic diseases in, 529–530 prolapsed bowel in, 387–389 septic suppurative inflammatory infiltrate in, 517f tumors in, 518f Heartworm disease in ferrets, 70–71, 70f Heat stress in guinea pigs, 308–309 Heat stroke in chinchillas, 317 in rabbits, 254 Hedgehogs. See also African Hedgehogs behavior of, 555 communication behaviors, 555, 555f play behaviors, 555 social behaviors, 555 dentistry of, 469 Helicobacter mustelae gastritis in ferrets, 30–33, 31f, 32t Helminthic infections in chinchillas, 320 Helminths in rabbits, 202 Hematologic diseases in African hedgehogs, 419–420 Hematology and cytology of small mammals, 511 cats Helicobacter species in, 520f mycobacteriosis in, 519f chinchillas leukocytes in, 512f lymphocytes in, 512f neutrophils in, 512f Cryptosporidium species, 519f ferrets benign sebaceous epithelial structure in, 518f chordoma in, 520f extramedullary hematopoiesis (EMH) in, 519f leukocytes in, 512f–513f lymphoid hyperplasia in, 516f lymphosarcoma in, 516f malignant epithelial tumor in, 517f malignant neuroendocrine neoplasm in, 519f mast cell neoplasm in, 520f modified transudate in, 521f plasma cell tumor in, 521f platelet clumping in, 513f reticulocytes in, 515f guinea pigs benign mammary eipthelial hyperplasia in, 517f formalin contamination in, 522f leukocytes in, 513f lymphocytes with Kurloff’s bodies in, 514f hamsters lymphosarcoma in, 516f malignant melanoma in, 518f malignant mesenchymal tumor in, 518f septic suppurative inflammatory infiltrate in, 517f mice erythrocytes with Howell-Jolly bodies in, 514f leukocytes in, 514f rabbits benign basal cell proliferation in, 521f erythrocyte polychromasia and reticulocytes in, 515f fine-needle aspiration of liver in, 522f leukocytes in, 514f
Hematology and cytology of small mammals (Continued) lipoma in, 520f malignant mesenchymal neoplasm in, 517f malignant tumor in, 518f mammary carcinoma in, 521f Saccharomyces species yeast organisms in, 516f Hemolymphatic system tumors in ferrets, 106–115 ancillary treatments, 114–115 chemotherapy, 110–112, 112t–115t classification of lymphoma, 106–107, 106t–107t, 107f cytologic/histologic description, 108–110, 110f–111f diagnostic imaging, 108, 109f laboratory evaluation, 108 palliative therapy, 112–113 radiation treatment, 113–114 signalment and clinical signs, 107–108, 108f treatment, 110–115 Hemorrhagic disease virus (RHDV) in rabbits, 200–201 Hepatic coccidia in rabbits, 201 Hepatic lipidosis in guinea pigs, 298 and ketosis in chinchillas, 318 Herpes simplex virus (HSV) in rabbits, 251 Husbandry of African hedgehogs, 414–415 breeding and neonatal care, 415 diet, 415 housing, 414–415 of chinchillas, 288–289 of ferrets, 9–11 environmental enrichment, 10 housing, 9–10 nutrition, 10–11 of guinea pig, 283–284 of rabbits, 170–171, 171f of small rodents, 342–344 diet and feeding, 343 housing and equipment, 342–343, 343f zoonosis, 344 of sugar gliders, 397–398 Hydrometra in ferrets, 56, 56f in rabbits, 221 Hydronephrosis in ferrets, 47 Hypercalciuria in rabbits, 225–227, 225f–227f Hyperestrogenism in ferrets, 55 Hypertrophic cardiomyopathy (HCM) in ferrets, 68–69 Hypervitaminosis D in rabbits, 228
I
Iatrogenic muscle necrosis in guinea pigs, 307 Ibuprofen toxicosis in ferrets, 75 Imaging, diagnostic, 502 advanced treatments, 509 interventional radiology (IR), 509, 509f radiation therapy (RT), 509, 509f general considerations, 502, 503f–504f which imaging examination to perform, 502–509, 504t computed tomography (CT), 506–508, 507f–508f contrast radiography, 503–505 digital radiography, 505 image resolution, 503, 505f magnetic resonance imaging (MRI), 508
Imaging, diagnostic (Continued) modalities, 503–509 nuclear scintigraphy, 508–509, 508f radiography, 503, 506f ultrasound, 505–506, 506f Impaction of paracloacal gland in sugar gliders, 405 Infectious diseases in sugar gliders, 408 Infertility in sugar gliders, 405 Inflammatory bowel disease (IBD) in ferrets, 37–38 Influenza in ferrets, 80–81, 81t virus in ferrets, 37 Insulinoma in ferrets, 104–105 Integumentary diseases in African hedgehogs, 418f, 421–422, 421f–423f Interventional radiology (IR), 509, 509f Intervertebral disk disease of ferrets, 138 Intestinal coccidia in rabbits, 201–202 Intestinal nephritis, chronic in guinea pigs, 301 Intestinal resection in rabbits, 273–274, 274f Intestinal surgery in ferrets, 143–144, 144f IR. See Interventional radiology (IR)
J
Jill diseases in ferrets, 57–58
K
Ketoconanazole in ferrets, 90 Kit diseases in ferrets, 58–59
L
Lead poisoning in chinchillas, 317 toxicosis in rabbits, 253 Lens diseases in rabbits, 525–526, 526f Leukemias in rabbits, 263 Lice in rabbits, 239 Liver biopsy in ferrets, 144–145, 145f–146f disease in ferrets, 35–36 Lymphocytic choriomeningitis virus (LCMV) in guinea pigs, 307 Lymphoma, of ferrets, 137 Lymphomas in rabbits cutaneous, 263 multicentric, 262–263 thymic, 263–265 Lymphoproliferative disorders in rabbits, 257, 262–267 chemotherapy, 265–266 cutaneous lymphoma, 263 diagnosis, 265 etiology, 262 leukemia, 263 multicentric lymphoma, 262–263 thymic masses, 263–265 thymoma/thymic lymphoma, thymic carcinoma, 263–265 treatment, 265–267 Lymphosarcoma/cavian leukemia in guinea pigs, 309
M
Magnetic resonance imaging (MRI), 508 Male reproductive tract tumors in ferrets, 54, 54f Malignant peripheral nerve sheath tumor in ferrets, 119 Malnutrition in sugar gliders, 402
Index Mammal diagnostic and surgical endoscopy, exotic, 485 anesthesia, 485–486 complications, 501 instrumentation, 486, 487t, 488f–489f outcome, 501 patient evaluation, 485 patient selection, 485 postoperative care, 501 procedures, 486–501 endotracheal intubation, 488, 493f gastroscopy/colonoscopy, 493–496, 497f–498f laparoscopy, 496–499, 499f–500f otoscopy, 486, 489f rhinoscopy, 488–493, 494f–496f stomatoscopy, 486–488, 490f–492f thoracoscopy, 499–501 tracheobronchoscopy, 488, 494f vaginoscopy/cystoscopy, 493, 497f Mammals. See Small mammals Mammary dysplasia in rabbits, 223 Mammary gland neoplasia in chinchillas, 330–331 in degus, 330–331 in ferrets, 58 in guinea pigs, 330–331 in small rodents, 382–384, 383f Mammary glands disorders in rabbits, 223 Mammary tumors in rabbits, 223 Mast cell tumors in ferrets, 127–128, 128f Mastitis in ferrets, 58 in guinea pigs, 303 in rabbits, 233–234 Matted fur in chinchillas, 318 Melatonin in ferrets, 90 Metastatic mineralization in guinea pigs, 307 Methicillin-resistant staphylococcal infection in rabbits, 234 Metritis in ferrets, 58 in guinea pigs, 303 Mice. See also Small rodents anatomic and physiologic characteristics, 342 clinical signs and treatment by species, 358–361, 360f erythrocytes with Howell-Jolly bodies in, 514f general characteristics, 340 leukocytes in, 514f ophthalmologic diseases in, 529–530 Mites in ferrets, 124–125 in guinea pigs, 307 Mitotane in ferrets, 90 Moist dermatitis in rabbits, 234–235 MRI. See Magnetic resonance imaging (MRI) Mucometra in ferrets, 55–56 Multicentric lymphoma in rabbits, 262–263 Musculoskeletal and neurologic diseases in ferrets, 132 ataxia, 132–133 intracranial disorders, 138 canine distemper, 138 osteoma, 138 rabies, 138 miscellaneous diseases, 139 miscellaneous metabolic diseases, 139 musculoskeletal disorders, 138–139 disseminated idiopathic myofasciitis (DIM), 138–139 myasthenia gravis, 139 neoplasia of central nervous system, 138
Musculoskeletal and neurologic diseases in ferrets (Continued) neuronal vacuolation, 138 posterior paresis, 132 seizures, 133–135, 134f–137f spinal disorders, 135–138 Aleutian disease, 135–136 intervertebral disk disease, 138 spinal tumors, 136–138 spinal tumors, 136–138 chondromas and chondrosarcomas, 136 fibrosarcoma, 137–138 lymphoma, 137 Musculoskeletal diseases in African hedgehogs, 420 in sugar gliders, 407 Musculoskeletal diseases in rabbits, neurologic and, 245, 246f, 246t bacterial infection, 250–251 bacterial infections of CNS, 251 otitis interna, 250–251, 251f degenerative/developmental, 252 osteoarthritis, 252 splay leg, 252 spondylosis, 252, 254f metabolic, 254 heat stroke, 254 toxemia of pregnancy, 254 miscellaneous causes, 255 idiopathic, 255 miscellaneous, 255 neoplastic, 255 vascular, 255 nutritional issues, 254–255 parasitic infection, 246–250 cerebral larva migrans, 250 cuterebra species, 250 encephalitozoonosis, 246–250, 247f, 248t toxoplasmosis, 250 toxic, 253–254 fipronil toxicosis, 253–254 lead toxicosis, 253 pyrethrin/permethrin toxicosis, 254 traumatic, 251–252 vertebral fracture or luxation, 251–252, 252f–253f viral infection, 251 herpes simplex virus (HSV), 251 rabies, 251 Musculoskeletal system tumors in ferrets, 118 Myasthenia gravis in ferrets, 139 Mycobacteriosis in ferrets, 36 Mycotic diseases, 563 Myiasis in rabbits, 238–239 Myocardial disease in rabbits, 261, 261f Myocarditis in ferrets, 69–70 Myxomatosis in rabbits, 239–240
N
Necrobacillosis in rabbits, 236 Nematodes in rabbits, 202 Neonatal conjunctivitis in ferrets, 59 Neonatal mortality and deformities in ferrets, 59 Neoplasia in chinchillas, 319 in guinea pigs, 298 in sugar gliders, 408 Neoplasia in ferrets, 40, 70, 103 endocrine system tumors, 104–106 adrenocortical neoplasms, 105–106, 105f insulinoma, 104–105 thyroid neoplasms, 106 etiology, 103–104
587
Neoplasia in ferrets (Continued) gastrointestinal (GI) tract tumors, 116–117, 117f hemolymphatic system tumors, 106–115 ancillary treatments, 114–115 chemotherapy, 110–112, 112t–115t classification of lymphoma, 106–107, 106t–107t, 107f cytologic/histologic description, 108–110, 110f–111f diagnostic imaging, 108, 109f laboratory evaluation, 108 palliative therapy, 112–113 radiation treatment, 113–114 signalment and clinical signs, 107–108, 108f treatment, 110–115 incidence and behavior, 104 malignant peripheral nerve sheath tumor, 119 miscellaneous neoplasms, 120 musculoskeletal system tumors, 118 nervous system tumors, 118–119, 119f reproductive tract tumors, 118 respiratory system tumors, 119 skin tumors, 115–116, 115f urinary system tumors, 119 vascular neoplasms, 119–120 Neoplasia in guinea pigs dermatology, 305, 305f reproductive, 303 Neoplasms involving skin or subcutis in ferrets, 129–130 Neoplastic diseases in African hedgehogs, 422–423, 424f Nephrectomy in ferrets, 152 in rabbits, 275 Nephritis chronic intestinal in guinea pigs, 301 and renal failure in sugar gliders, 405 Nephrocalcinosis in ferrets, 48 Nephrotomy in rabbits, 275 Nephrotoxicity in rabbits, 228 Nerve sheath tumors in ferrets, 119 Nervous system tumors in ferrets, 118–119, 119f Neurologic and musculoskeletal diseases in rabbits, 245, 246f, 246t bacterial infection, 250–251 bacterial infections of CNS, 251 otitis interna, 250–251, 251f degenerative/developmental, 252 osteoarthritis, 252 splay leg, 252 spondylosis, 252, 254f metabolic, 254 heat stroke, 254 toxemia of pregnancy, 254 miscellaneous causes, 255 idiopathic, 255 miscellaneous, 255 neoplastic, 255 vascular, 255 nutritional issues, 254–255 parasitic infection, 246–250 cerebral larva migrans, 250 cuterebra species, 250 encephalitozoonosis, 246–250, 247f, 248t toxoplasmosis, 250 toxic, 253–254 fipronil toxicosis, 253–254 lead toxicosis, 253 pyrethrin/permethrin toxicosis, 254
588
Index
Neurologic and musculoskeletal diseases in rabbits (Continued) traumatic, 251–252 vertebral fracture or luxation, 251–252, 252f–253f viral infection, 251 herpes simplex virus (HSV), 251 rabies, 251 Neurologic diseases in African hedgehogs, 420–421, 421f in sugar gliders, 407–408 Neurologic diseases in ferrets, musculoskeletal and, 132 ataxia, 132–133 intracranial disorders, 138 canine distemper, 138 osteoma, 138 rabies, 138 miscellaneous diseases, 139 miscellaneous metabolic diseases, 139 musculoskeletal disorders, 138–139 disseminated idiopathic myofasciitis (DIM), 138–139 myasthenia gravis, 139 neoplasia of central nervous system, 138 neuronal vacuolation, 138 posterior paresis, 132 seizures, 133–135, 134f–137f spinal disorders, 135–138 Aleutian disease, 135–136 intervertebral disk disease, 138 spinal tumors, 136–138 spinal tumors, 136–138 chondromas and chondrosarcomas, 136 fibrosarcoma, 137–138 lymphoma, 137 Noninfectious respiratory diseases in guinea pigs, 299 Nuclear scintigraphy, 508–509, 508f Nutrition of rabbits, gastrointestinal (GI) physiology and, 183 dietary components, 188–191 commercial mixes and pellets, 190–191 edible plants, 190 fresh vegetables, 190 grass, 188 greens, 190 hay, 188–190 miscellaneous feed items, 191 water, 191 dietary recommendations summary, 191 gastrointestinal physiology, 183–188 cecotrophy, 186–187, 186f–187f digestion and absorption, 187 energy requirements, 187–188 hindgut flora and fermentation, 186 ingestion of food, 183–184 large intestine, 184–185, 185f motility, 187 small intestine, 184, 184f stomach, 184, 184f nutrition, 188 carbohydrate, 188 fat, 188 fiber, 188 minerals, 188, 189t protein, 188 vitamins, 188, 189t Nutritional diseases in African hedgehogs, 423–424, 424f Nutritional muscular dystrophy in guinea pigs, 307 Nutritional osteodystrophy in sugar gliders, 407
O
Obesity in sugar gliders, 407 Ocular injuries in African hedgehogs, 418 in sugar gliders, 406 Ophthalmologic diseases in small pet mammals, 523 chinchillas, 529 ferrets, 527–528 guinea pigs, 528–529, 528f–529f hamsters, 529–530 mice, 529–530 rabbits, 523–527, 524f conjunctivitis, 523–525, 525f cornea, 525, 525f diseases of, 525–526, 526f epiphora, 523–525, 525f glaucoma, 526 orbit, 526–527, 527f uveitis and diseases of lens, 525–526, 526f rats, 529–530 sugar gliders, 530 Opthalmic disorders in sugar gliders, 406–407 Oral and dental diseases in African hedgehogs, 418–419, 418f Oral neoplasia in ferrets, 28, 28f Oral papillomavirus in rabbits, 240 Orbit in rabbits, 526–527, 527f Orchidectomy in chinchillas, 331–333, 331f–333f in degus, 331–333, 332f in guinea pigs, 331–333, 332f–333f in rabbits, 271–273, 272f–273f in small rodents, 378–380, 378f–380f Orchitis in guinea pigs, 303 in rabbits, 222 Orthopedics in small mammals, 472 amputations, 473 complications, 481–482 posttraumatic osteomyelitis, 482 external coaptation, 473–476, 474f intramedullary (IM) pinning, 474–475, 475f fracture fixation methods, 473 fracture healing and postoperative management, 481 fracture incidence, 482 initial fracture management/first aid, 472–473 intramedullary (IM) pinning bone plating, 475–476 external skeletal fixation, 476 joint luxations, 480, 481f pectoral limb, 476–478 humerus, 477 metacarpals, 478 radius/ulna, 477–478, 477t, 478f scapula, 476–477 pelvic limb, 478–479 femur, 478–479, 479f metatarsals, 479 pelvis, 478 tibia/fibula, 479, 480f repair of specific fracture types, 476 skull fractures, 480 spinal fractures and luxations, 479 Osteoarthritis and osteoarthrosis in guinea pigs, 306 in rabbits, 252 Osteoma in ferrets, 138 Otic diseases in African hedgehogs, 418 Otitis interna in rabbits, 250–251, 251f
Otitis media/otitis interna in guinea pigs, 307, 308f Ototoxicity in guinea pigs, 308 Ovarian cysts in guinea pigs, 301–302, 301f–302f Ovarian disease in small rodents, 381 Ovarian remnant in ferrets, 154 Ovariectomy in chinchillas, 327–328 in degus, 327–328 in guinea pigs, 327–328 and ovariohysterectomy in small rodents, 381–382 in small rodents, 381–382 Ovariohysterectomy in chinchillas, 328–329 in degus, 328–329 in ferrets, 154 in guinea pigs, 328–329, 328f in rabbits, 271, 271f in small rodents, 382
P
Pancreatic islet cell tumors in ferrets, 92–99 advanced imaging, 96 baseline insulin and glucose concentrations, 95, 95t beyond Whipple’s triad, 95 clinical features, 93–94 clinical pathologic abnormalities, 94 clinical signs, 93–94 diagnostic approach, 95–96 diagnostic imaging, 96 differential diagnoses for fasting hypoglycemia, 94–95 etiology, 92–93 histopathology, 99 insulin: glucose ratios, 95 management of islet cell tumors, 96–99 medical management of chronic hypoglycemia, 98–99 medical therapies for active hypoglycemic crisis, 97–98 miscellaneous testing, 96 pathophysiology, 93 physical examination, 94, 94f prognosis, 99 provocative testing, 95–96 radiography, 96 signalment, 93 surgical therapy, 96–97, 96t–97t ultrasonography, 96, 96f Pancreatic surgery in ferrets, 150–151, 151f Papillomatosis in rabbits, 200 Paracloacal gland, impaction, in sugar gliders, 405 Parasites. See also Ectoparasites; Endoparasites Parasites in ferrets, 16–17 ectoparasites, 17 endoparasites, 16–17 Parasitic diseases, 562–563, 562f–563f Parasitic diseases in sugar gliders, 408 Parasitic disorders of gastrointestinal (GI) tract in rabbits, 201–202 cestodes, 202 coccidia, 201–202 cryptosporidia, 202 helminths, 202 hepatic coccidia, 201 intestinal coccidia, 201–202 miscellaneous protozoa, 202 nematodes, 202 trematodes, 202
Index Parasitic infections in rabbits, 237–239, 246–250 cerebral larva migrans, 250 cuterebra species, 250 ear mites, 237, 237f encephalitozoonosis, 246–250, 247f, 248t fleas, 238 fur mites, 237, 237f lice, 239 myiasis, 238–239 pinworms, 239 tapeworm cysts, 239 ticks, 239 toxoplasmosis, 250 Paraurethral cysts or paraurethral disease in ferrets, 53–54 Paraurethral/prostatic cysts in ferrets, 153–154, 153f Pasteurellosis in rabbits, 205 Pea eye in guinea pigs, 307, 309f Penile lesions in ferrets, 54 Penile prolapse in chinchillas, 333 in degus, 333 in guinea pigs, 333 Perineal urethrostomy in ferrets, 153, 153f Periparturient disease in ferrets, 56–59 Pheochromocytomas in ferrets, 91, 91f Pinworms in rabbits, 239 Pneumonia in ferrets, 81–83, 82f Pododermatitis in guinea pigs, 305, 305f Polycystic kidney disease in ferrets, 46–47 in ferret with acute renal failure in ferrets, 47f Postparturient hypocalcemia in ferrets, 58 Pouch infection and mastitis in sugar gliders, 405 Pregnancy abdominal in rabbits, 220 toxemia in ferrets, 57 toxemia in rabbits, 219 Preputial masses in ferrets, 154, 155f Prolapsed bowel in hamsters, 387–389 Prolapsed vagina in rabbits, 220–221 Proliferative bowel disease in ferrets, 38–39, 39f Prostatic cysts in ferrets, 51–52, 51f–52f Prostatic lesions in ferrets, 54 Prostatic tumors in ferrets, 53 Prostatitis and prostatic abscess in ferrets, 53, 53f Protozoa in rabbits, miscellaneous, 202 Protozoal infections in chinchillas, 319–320 Pseudopregnancy in ferrets, 57 in rabbits, 219–220 Psychogenic polyuria and polydipsia in rabbits, 229 Pulmonary lobectomy in small rodents, 385–386 Pulmonary mycoses in ferrets, 83 blastomycosis, 83 coccidioidomycosis, 83 cryptococcosis, 83 history and physical examination, 83 Pyelolithotomy in rabbits, 275 Pyelonephritis in ferrets, 48 Pyometra in chinchillas, 315–316, 329 in degus, 329 in ferrets, 55–56, 154 in guinea pigs, 303, 329 in rabbits, 219 Pyrethrin/permethrin toxicosis in rabbits, 254
R
Rabbit (Shope) fibroma virus, 240 papillomavirus, 240 Rabbitpox, 240–241 Rabbits behavior of, 546–549 behavior problems, 546–548, 548b introduction of new conspecifics, 549 litter-box training, 548–549 mourning death of bonded mate, 549 play behaviors, 546 social behaviors, 546, 547t benign basal cell proliferation in, 521f erythrocyte polychromasia and reticulocytes in, 515f fine-needle aspiration of liver in, 522f leukocytes in, 514f lipoma in, 520f malignant mesenchymal neoplasm in, 517f malignant tumor in, 518f mammary carcinoma in, 521f ophthalmologic diseases in, 523–527, 524f conjunctivitis, 523–525, 525f cornea, 525, 525f epiphora, 523–525, 525f glaucoma, 526 lens, diseases of, 525–526, 526f orbit, 526–527, 527f uveitis, 525–526, 526f Saccharomyces species yeast organisms in, 516f Rabbits, anatomy, physiology, and husbandry of, 157 anatomy, 159 behavior, 169–170 auditory signals, 170 drinking, 169–170 eating, 169–170 elimination behavior, 169–170 group behavior, 170 visual signals, 170 vocalization, 170 breeds and varieties, 159 cardiovascular system, 164–165 digestive system, 161–163 abdominal cavity, 162 gallbladder, 163 large intestine, 162–163, 163f liver, 163 oral cavity, 162 pancreas, 163 small intestine, 162 spleen, 163 stomach, 162 teeth, 161–162, 161f ears, 160–161 etymology, 157–158, 158f eyes, 160 female reproductive system, 166–168 anatomy and physiology, 166–167, 166f female sexual behavior, 167 hand-rearing of baby rabbits, 168, 168b pregnancy and nursing behavior, 167–168 hematology, 164 husbandry, 170–171, 171f lifespan, 159 male reproductive system, 168–169 anatomy and physiology, 168–169 male sexual behavior and reproduction, 169 muscles and skeleton, 161 puberty and breeding life, 166, 166f respiratory system and thymus, 163–164
589
Rabbits, anatomy, physiology, and husbandry of (Continued) scent marking glands, 159–160 sense organs and nervous system, 160–161 skin and hair, 159–165 taxonomy and similarities to rodents, 158–159 urinary system, 164–165, 165f Rabbits, cardiac disease in, 257–258 arrhythmia, 261 congenital heart disease, 261 congestive heart failure (CHF), 260 diagnostic methods, 258–260, 258f–259f, 259t–260t diseases and management, 260–262 examination of rabbit, 257–258 myocardial disease, 261, 261f normal cardiovascular structure, 257 valvular disease, 261–262 vascular disease, 262 Rabbits, cardiovascular disease in, 257 Rabbits, dentistry of, 455–462 anatomy and physiology of skull and teeth, 455 dental disease, 457–458, 457f dental procedures, 458–460, 458f, 459b facial surgery and surgical treatment, 460–462, 462f medical treatment, 458 pathophysiology of dental disease, 455–456, 456f treatment of dental disease, 458 treatment of periapical infections and abscesses, 460, 461b, 461f Rabbits, dermatological diseases in, 232 bacterial infections, 232–236 cellulitis, 234 mastitis, 233–234 methicillin-resistant staphylococcal infection, 234 moist dermatitis, 234–235 necrobacillosis, 236 rabbit syphilis, 235–236, 235f subcutaneous abscesses, 232–233, 233f ulcerative pododermatitis, 235 cutaneous neoplasia, 241, 241f fungal infections, 236–237 dermatophytosis, 236–237 parasitic infections, 237–239 ear mites, 237, 237f fleas, 238 fur mites, 237, 237f lice, 239 myiasis, 238–239 pinworms, 239 tapeworm cysts, 239 ticks, 239 skin disease, behavioral causes of, 241 barbering, 241 self-mutilation after intramuscular injection, 241 skin disease of unknown cause, 241–242 dermal fibrosis, 242, 242f Ehlers-Danlos-like syndrome, 242 eosinophilic granuloma, 242 sebaceous adenitis, 241–242, 242f viral infections, 239–241 myxomatosis, 239–240 oral papillomavirus, 240 rabbit (Shope) fibroma virus, 240 rabbit (Shope) papillomavirus, 240 rabbitpox, 240–241
590
Index
Rabbits, gastrointestinal (GI) diseases in, 193 acute gastrointestinal (GI) dilation or obstruction, 196–197, 196f aflatoxicosis, 203 cecoliths, 198 cecotrophy and intermittent diarrhea, 197–198 enteritis complex and enterotoxemia, 198–200 bacterial enteritis, 199–200 dysbiosis caused by treatment with antibiotics, 198–199 enteropathogenic E. coli (EPEC), 199 miscellaneous bacterial enteritides, 200 mucoid enteritis, 198 prevention of enterotoxemia, 199 proliferative enteritis, 199 proliferative enterocolitis, 199 proliferative enteropathy, 199 treatment of enteritis, 199 Tyzzer’s disease, 199–200 gastrointestinal (GI) stasis syndrome, 193–196 gastrointestinal (GI) stress stasis syndrome diagnostic testing, 195, 195f effect of diet and cecocolic motility, 194 history and clinical signs, 194–195 physical examination findings, 195 role of fiber, 193–194 treatment, 196 liver lobe torsion, 202–203 neoplasia, 202 viral diseases of digestive tract, 200–201 hemorrhagic disease virus (RHDV), 200–201 papillomatosis, 200 rabbit enteric coronavirus, 200 rotavirus, 200 Rabbits, gastrointestinal (GI) physiology and nutrition of, 183 dietary components, 188–191 commercial mixes and pellets, 190–191 edible plants, 190 fresh vegetables, 190 grass, 188 greens, 190 hay, 188–190 miscellaneous feed items, 191 water, 191 dietary recommendations summary, 191 gastrointestinal physiology, 183–188 cecotrophy, 186–187, 186f–187f digestion and absorption, 187 energy requirements, 187–188 hindgut flora and fermentation, 186 ingestion of food, 183–184 large intestine, 184–185, 185f motility, 187 small intestine, 184, 184f stomach, 184, 184f nutrition, 188 carbohydrate, 188 fat, 188 fiber, 188 minerals, 188, 189t protein, 188 vitamins, 188, 189t Rabbits, lymphoproliferative disorders in, 257, 262–267 chemotherapy, 265–266 cutaneous lymphoma, 263 diagnosis, 265 etiology, 262 leukemia, 263 multicentric lymphoma, 262–263
Rabbits, lymphoproliferative disorders in (Continued) thymic masses, 263–265 thymoma/thymic lymphoma, thymic carcinoma, 263–265 treatment, 265–267 Rabbits, neurologic and musculoskeletal diseases in, 245, 246f, 246t bacterial infection, 250–251 bacterial infections of CNS, 251 otitis interna, 250–251, 251f causes, miscellaneous, 255 idiopathic, 255 miscellaneous, 255 neoplastic, 255 vascular, 255 degenerative/developmental, 252 osteoarthritis, 252 splay leg, 252 spondylosis, 252, 254f metabolic, 254 heat stroke, 254 toxemia of pregnancy, 254 nutritional issues, 254–255 parasitic infection, 246–250 cerebral larva migrans, 250 cuterebra species, 250 encephalitozoonosis, 246–250, 247f, 248t toxoplasmosis, 250 toxic, 253–254 fipronil toxicosis, 253–254 lead toxicosis, 253 pyrethrin/permethrin toxicosis, 254 traumatic, 251–252 vertebral fracture or luxation, 251–252, 252f–253f viral infection, 251 herpes simplex virus (HSV), 251 rabies, 251 Rabbits, reproductive and urinary systems disorders in, 217 cystic mastitis, 223 mammary dysplasia, 223 mammary glands disorders, 223 septic mastitis, 223 mammary tumors, 223 urinary system disorders, 223–229, 224f encephalitozoonosis, 228 hypercalciuria, 225–227, 225f–227f hypervitaminosis D, 228 nephrotoxicity, 228 psychogenic polyuria and polydipsia, 229 red urine, 229 renal adipose deposition, 228 renal agenesis, 228 renal cysts, 228 renal failure, 227–228 urinary bladder inversion, 229 urinary incontinence, 229 urinary tract tumors, 229 urolithiasis, 225–227, 225f–227f Rabbits, respiratory diseases and pasteurellosis in, 205 anatomy of respiratory tract, 205 diagnosis and differentiation, 208–209 diagnostic imaging, 209, 209f–211f diseases producing secondary respiratory systems, 208 laboratory analysis, 208–209, 208f lower respiratory tract diseases, 207–208 infectious, 207 neoplastic, 207–208 physical examination, 208
Rabbits, respiratory diseases and pasteurellosis in (Continued) prevention and control of respiratory disease, 214 serology and molecular diagnostic testing, 209 treatment of respiratory disease, 209–214, 212f–215f upper respiratory tract diseases, 205–207 bacterial pathogens, 205–206 dental disease, 206–207, 207f fungal pathogens, 206 infectious, 205–206 miscellaneous conditions, 207 neoplastic, 207 noninfectious, 206–207 trauma, 206, 207f viral pathogens, 206 Rabbits, soft tissue surgery in, 269 abscesses, 275 anorectal papilloma removal, 274 common procedures, 270–271 castration, 271–273, 272f–273f orchidectomy, 271–273, 272f–273f ovariohysterectomy, 271, 271f correcting problems of urine scald, 275–276, 276f–277f dermoplasty and tail amputation, 275–276, 276f–277f equipment, 269 evaluating rabbit as surgical patient, 269 gastrointestinal (GI) surgery, 273–274 anastomosis, 273–274, 274f enterotomy, 273–274, 274f gastrotomy, 273 intestinal resection, 273–274, 274f surgery of large bowel, 274 postsurgical monitoring, 270 blood loss, 270 pain and analgesics, 270 presurgical treatment, 270 surgery of auditory bulla, 276–278 lateral ear canal ablation, 276–277 osteotomy of ventral bulla, 277–278 surgery of ear canal, 276–278 lateral ear canal ablation, 276–277 osteotomy of ventral bulla, 277–278 surgical techniques, 270–271 adhesion formation, 270 choice of suture material, 270 skin closures, 270–271 Rabbits, thymomas in, 257, 262–267 chemotherapy, 265–266 diagnosis, 265 etiology, 262 thymic carcinoma, 263–265 thymic lymphoma, 263–265 thymic masses, 263–265, 264f–265f treatment, 265–267 treatment options, 266–267 Rabbits, veterinary care, basic approach to, 174 handling and restraint, 175, 175f housing, 174–175 miscellaneous procedures, 181–182 anesthetic delivery, 181 ear cleaning, 182 nasolacrimal cannulation, 181, 181f–182f physical examination, 175–176, 176f sample collection, 176–178 blood collection, 176–177, 177f, 177t– 178t cerebrospinal fluid (CSF), 178, 179t collection of urine and feces, 177–178, 178t dermatologic samples, 178
Index Rabbits, veterinary care, basic approach to (Continued) treatment techniques, 178–181 catheterization and fluid therapy, 178–179, 179f external support, 180, 180f injection techniques, 179–180, 179f oral medications, 180, 180f pain control, 181 vaccinations, 181 Rabies in ferrets, 138 in guinea pigs, 307 in rabbits, 251 vaccinations in ferrets, 15–16 Radiation therapy (RT), 509, 509f Radiography, 503, 506f Rats. See also Small rodents anatomic and physiologic characteristics, 342 clinical signs and treatment by species, 361–364, 361f–364f general characteristics, 339–340 ophthalmologic diseases in, 529–530 Rectal disease in ferrets, 39–40 Rectal prolapse in sugar gliders, 402–404 Red urine in rabbits, 229 Reduced fertility in rabbits, 220 Renal adipose deposition in rabbits, 228 Renal agenesis in rabbits, 228 Renal cysts in ferrets, 46 Renal cysts in rabbits, 228 Renal disease and renal failure in ferrets, 47–48 Renal failure chronic in guinea pigs, 301 in rabbits, 227–228 in sugar gliders, 405 Renal neoplasia in ferrets, 48 Reproductive and urinary systems disorders in rabbits, 217 cystic mastitis, 223 mammary dysplasia, 223 mammary glands disorders, 223 septic mastitis, 223 mammary tumors, 223 urinary system disorders, 223–229, 224f encephalitozoonosis, 228 hypercalciuria, 225–227, 225f–227f hypervitaminosis D, 228 nephrotoxicity, 228 psychogenic polyuria and polydipsia, 229 red urine, 229 renal adipose deposition, 228 renal agenesis, 228 renal cysts, 228 renal failure, 227–228 urinary bladder inversion, 229 urinary incontinence, 229 urinary tract tumors, 229 urolithiasis, 225–227, 225f–227f Reproductive diseases in African hedgehogs, 420 Reproductive disorders in sugar gliders, 405 Reproductive system disorders in ferrets, 54–56 Reproductive systems disorders in ferrets, urinary and, 46 female ferret, 54–56 female reproductive tract tumors, 55–56 hydrometra, 56, 56f hyperestrogenism, 55 mucometra, 55–56 pyometra, 55–56
Reproductive systems disorders in ferrets, urinary and (Continued) jill diseases, 57–58 agalactia, 58 dystocia, 57 mammary gland neoplasia, 58 mastitis, 58 metritis, 58 postparturient hypocalcemia, 58 pregnancy toxemia, 57 pseudopregnancy, 57 kit diseases, 58–59 caring for ill kits, 58–59, 59t diarrhea, 59 enlarged umbilical cords, 59 neonatal conjunctivitis, 59 neonatal mortality and deformities, 59 normal kit, 58 splay-legged kits, 59 male ferret, 54 cryptorchidism, 54 male reproductive tract tumors, 54, 54f penile lesions, 54 prostatic lesions, 54 periparturient disease, 56–59 breeding ferrets, management of, 57 jill diseases, 57–58 kit diseases, 58–59 normal breeding, 56–57 polycystic kidney disease, polycystic kidney in ferret with acute renal failure, 47f renal disease and renal failure, 47–48 acute renal failure, 47 chronic renal failure, 47 diagnosis and treatment, 47–48 reproductive system disorders, 54–56 female ferret, 54–56 male ferret, 54 urethral obstruction urethral catheterization of male ferret, 51f urethral obstruction, 50f urinary and reproductive systems disorders periparturient disease, 56–59 reproductive system disorders, 54–56 urinary system disorders, 46–54 urinary system disorders, 46–54 Aleutian disease, 48 bladder neoplasia, 50 cystitis, 49–50 hydronephrosis, 47 nephrocalcinosis, 48 paraurethral cysts or paraurethral disease, 53–54 polycystic kidney disease, 46–47 prostatic cysts, 51–52, 51f–52f prostatic tumors, 53 prostatitis and prostatic abscess, 53, 53f pyelonephritis, 48 renal cysts, 46 renal disease and renal failure, 47–48 renal neoplasia, 48 ureteral rupture, 48 urethral obstruction, 50–51 urinary incontinence, 50 urolithiasis, 49 Reproductive tract infection in sugar gliders, 405 Reproductive tract tumors in ferrets, 118 Respiratory diseases in African hedgehogs, 419 in guinea pigs, 298 in sugar gliders, 405
591
Respiratory diseases and pasteurellosis in rabbits, 205 anatomy of respiratory tract, 205 diagnosis and differentiation, 208–209 diagnostic imaging, 209, 209f–211f diseases producing secondary respiratory systems, 208 laboratory analysis, 208–209, 208f lower respiratory tract diseases, 207–208 infectious, 207 neoplastic, 207–208 physical examination, 208 prevention and control of respiratory disease, 214 serology and molecular diagnostic testing, 209 treatment of respiratory disease, 209–214, 212f–215f upper respiratory tract diseases, 205–207 bacterial pathogens, 205–206 dental disease, 206–207, 207f fungal pathogens, 206 infectious, 205–206 miscellaneous conditions, 207 neoplastic, 207 noninfectious, 206–207 trauma, 206, 207f viral pathogens, 206 Respiratory diseases in ferrets, 78 canine distemper virus (CDV), 78–80, 79f influenza, 80–81, 81t pneumonia, 81–83, 82f pulmonary mycoses, 83 blastomycosis, 83 coccidioidomycosis, 83 cryptococcosis, 83 history and physical examination, 83 respiratory diseases, other causes of respiratory signs, 83, 83f Respiratory diseases, noninfectious in guinea pigs, 299 Respiratory distress in small mammals, emergency treatment of, 541–542 Respiratory system disorders in chinchillas, 315 tumors in ferrets, 119 Retrobulbar abscess in sugar gliders, 407 Rodents. See also Small rodents Rodents, dentistry of, 462–468 anatomy and physiology of skull and teeth, 462–464, 463t, 464f clinical presentation, 465 dental disease, 465–467, 466f–467f pathophysiology of dental disease, 464–467, 464f treatment of dental disease, 467 treatment of periapical infections and abscesses, 467–468 Rotavirus in ferrets, 37 in rabbits, 200 RT. See Radiation therapy (RT)
S
Saccharomyces species yeast organisms of rabbits, 516f Salivary mucocele in ferrets, 28, 28f Salivary mucocele resection in ferrets, 143 Salmonellosis in ferrets, 36 SCC. See Squamous cell carcinoma (SCC) Scrotal plugs in guinea pigs, 303 Scurvy in guinea pigs, 305–306, 306f Sebaceous adenitis in rabbits, 241–242, 242f Sebaceous adenomas in ferrets, 128, 128f
592
Index
Sedation in small mammals, 429 Seizures in chinchillas, 317 Self-mutilation after intramuscular injection in rabbits, 241 of penis and scrotum in sugar gliders, 405 in sugar gliders, 406, 406f Septic mastitis in rabbits, 223 Skin disease in rabbits, behavioral causes of, 241 barbering, 241 self-mutilation after intramuscular injection, 241 Skin tumors in ferrets, 115–116, 115f Small mammal dentistry, 452 diagnostic testing, 453–455 clinical examination, 453 computed tomography (ct), 454–455 imaging, 453–455 oral endoscopy, 454 other diagnostic testing, 455 radiography, 453–454, 454f equipment, 452–453 ferrets, 468–469 anatomy and physiology of skull and teeth, 468, 468f dental disease, 468 treatment and prevention, 468–469 hedgehogs, 469 rabbits, 455–462 anatomy and physiology of skull and teeth, 455 dental disease, 457–458, 457f dental procedures, 458–460, 458f, 459b facial surgery and surgical treatment, 460–462, 462f medical treatment, 458 pathophysiology of dental disease, 455–456, 456f treatment of dental disease, 458 treatment of periapical infections and abscesses, 460, 461b, 461f rodents, 462–468 anatomy and physiology of skull and teeth, 462–464, 463t, 464f clinical presentation, 465 dental disease, 465–467, 466f–467f pathophysiology of dental disease, 464–467, 464f treatment of dental disease, 467 treatment of periapical infections and abscesses, 467–468 sugar gliders, 469 Small mammals African hedgehogs, 411 antifungal agents used in, 568t antimicrobial agents used in, 567t–568t antiparasitic agents used in, 569t–570t behavior of, 545 chemical restraints used in, 571t–573t miscellaneous agents used in, 574t–575t sugar gliders, 393 Small mammals, analgesic agents used in, 571t–573t Small mammals, anesthesia, analgesia, and sedation of, 429 anesthesia, analgesia, 429–430 anesthetizing small exotic mammals, 429 clinical techniques, 430–432 incubation, 431–432, 432f–433f vascular access, 430–431, 431f equipment, 430 breathing circuits, 430 ventilators, 430
Small mammals, anesthesia, analgesia, and sedation of (Continued) injectable medications, 435 preanesthetic considerations, 430 nutritional status and fasting, 430 patient evaluation, 430 preanesthetic medications, 432, 434t Small mammals, anesthetic agents used in, 571t–573t Small mammals, behavior of behavioral training techniques for small mammals, 555 chinchillas, 553–554 behavior problems, 553–554 communication behaviors, 553 introduction of new conspecifics, 554 play behaviors, 553 social behavior, 553, 553f ferrets, 549–552 behavior problems, 550–551, 551b, 551f introduction of new conspecifics, 552 litter-box training, 552 play and sleeping behaviors, 549–550, 550b, 550f social behaviors, 549, 550b, 550f guinea pigs, 552–553 behavior problems, 553, 553f communication behaviors, 553 introduction of new conspecifics, 553 play behaviors, 552, 552f social behavior, 552, 552f hedgehogs, 555 communication behaviors, 555, 555f play behaviors, 555 social behaviors, 555 rabbits, 546–549 behavior problems, 546–548, 548b introduction of new conspecifics, 549 litter-box training, 548–549 mourning death of bonded mate, 549 play behaviors, 546 social behaviors, 546, 547t sugar gliders, 554–555 behavior problems, 554 communication behaviors, 554 introduction of new conspecifics, 555 play behaviors, 554, 554f social behaviors, 554 Small mammals, emergency and critical care of, 532 cardiopulmonary-cerebral resuscitation (CPCR), 532–535 anesthesia-related arrest, 534–535, 534t determining effectiveness of cpcr, 534 in small mammals: cardiac arrest, 533–534, 533b in small mammals: respiratory arrest, 533, 533f critical care clinical pathology, 539–540 lactate monitoring, 540 use of prothrombin and partial thromboplastin times, 540 identification and triage of critically ill patient, 532 indirect measurement of systolic blood pressure, 538–539, 539f maintenance of normothermia, 538, 538f nutritional support, 540 sedation and anesthesia of critically ill small mammal, 540–541, 542f shock and fluid therapy, 535–538 blood transfusion, 537, 537f fluid resuscitation of critically ill small mammal, 535–537, 536b–537b routes of fluid administration, 538
Small mammals, emergency and critical care of (Continued) three phases of hypovolemic shock, 535 types of fluids, 535 use of glucocorticoids in shock, 537–538 treatment of selected common emergencies, 541–543 Small mammals, hematology and cytology of, 511 cats Helicobacter species in, 520f mycobacteriosis in, 519f chinchillas leukocytes in, 512f lymphocytes in, 512f neutrophils in, 512f Cryptosporidium species, 519f ferrets benign sebaceous epithelial structure in, 518f chordoma in, 520f extramedullary hematopoiesis (EMH) in, 519f leukocytes in, 512f–513f lymphoid hyperplasia in, 516f lymphosarcoma in, 516f malignant epithelial tumor in, 517f malignant neuroendocrine neoplasm in, 519f mast cell neoplasm in, 520f modified transudate in, 521f plasma cell tumor in, 521f platelet clumping in, 513f reticulocytes in, 515f guinea pigs benign mammary eipthelial hyperplasia in, 517f formalin contamination in, 522f leukocytes in, 513f lymphocytes with Kurloff’s bodies in, 514f hamsters lymphosarcoma in, 516f malignant melanoma in, 518f malignant mesenchymal tumor in, 518f septic suppurative inflammatory infiltrate in, 517f mice erythrocytes with Howell-Jolly bodies in, 514f leukocytes in, 514f rabbits benign basal cell proliferation in, 521f erythrocyte polychromasia and reticulocytes in, 515f fine-needle aspiration of liver in, 522f leukocytes in, 514f lipoma in, 520f malignant mesenchymal neoplasm in, 517f malignant tumor in, 518f mammary carcinoma in, 521f Saccharomyces species yeast organisms in, 516f Small mammals, orthopedics in, 472 amputations, 473 complications, 481–482 posttraumatic osteomyelitis, 482 external coaptation, 473–476, 474f intramedullary (im) pinning, 474–475, 475f fracture fixation methods, 473 fracture healing and postoperative management, 481 fracture incidence, 482
Index Small mammals, orthopedics in (Continued) initial fracture management/first aid, 472–473 intramedullary pinning (IM) bone plating, 475–476 external skeletal fixation, 476 joint luxations, 480, 481f pectoral limb, 476–478 humerus, 477 metacarpals, 478 radius/ulna, 477–478, 477t, 478f scapula, 476–477 pelvic limb, 478–479 femur, 478–479, 479f metatarsals, 479 pelvis, 478 tibia/fibula, 479, 480f repair of specific fracture types, 476 skull fractures, 480 spinal fractures and luxations, 479 Small pet mammals, ophthalmologic diseases in, 523 chinchillas, 529 ferrets, 527–528 guinea pigs, 528–529, 528f–529f hamsters, 529–530 mice, 529–530 rabbits, 523–527, 524f conjunctivitis, 523–525, 525f cornea, 525, 525f diseases of, 525–526, 526f epiphora, 523–525, 525f glaucoma, 526 orbit, 526–527, 527f uveitis and diseases of lens, 525–526, 526f rats, 529–530 sugar gliders, 530 Small rodents anatomic and physiologic characteristics, 340–342 general, 340–341, 341t gerbils, 342 hamsters, 342 mice, 342 rats, 342 sexing, 341–342, 342f clinical techniques, 344–352 blood sample collection, 345–350, 350f bone marrow collection, 350 diagnostic testing procedures, miscellaneous, 351 fecal sample collection, 350 handling and restraint, 344–345, 344f–346f hospitalization, 351, 351f sample collection, 345–351, 346t–350t sample collection, miscellaneous, 350–351 therapeutics, 351–352, 351f–352f urine collection, 350 general characteristics, 339–340 gerbils, 340 hamsters, 340 mice, 340 rats, 339–340 husbandry of, 342–344 diet and feeding, 343 housing and equipment, 342–343, 343f Small rodents, disease problems of, 354 clinical signs and treatment by species, 358–369 hamsters, 364–367 mice, 358–361, 360f rats, 361–364, 361f–364f
Small rodents, disease problems of (Continued) diagnostic challenge, 354–356 clinical examination, 355–356 medical history, 355 pet rodent etiquette, 354 reception area, 355 scheduling appointments, 354–355 diseases, 357–369 clinical signs and treatment by species, 358–369 prophylaxis for small rodents, 358, 359t seen in practice, 357 significant diseases and life spans, 357–358, 357t Small rodents, soft tissue surgery in, 373 abdominal masses, 385 abdominal procedures, 384–385 abscesses, 386 analgesia, 390 cesarean section, 382 cheek pouch eversion in hamsters, 387, 388f–389f cutaneous masses, 384, 384f cystotomy, 384–385 dental abscesses in hamsters, 386 enucleation and exenteration, 386–387 focal light, 376 hemostatic aids, 377, 377f instrumentation, 374–376, 375f magnification, 376, 376f mammary gland neoplasia, 382–384, 383f orchidectomy, 378–380, 378f–380f ovarian disease, 381 ovariectomy, 381–382 ovariectomy and ovariohysterectomy, 381–382 ovariohysterectomy, 381–382 patient preparation, 377 patient support, 373–374, 374f presurgical considerations, 373–378 prolapsed bowel in hamsters, 387–389 pulmonary lobectomy, 385–386 recovery, 389–390 subcutaneous abscesses, 386 sutures, needles, and closure, 378 tail amputation, 389 uterine disease, 381 Soft tissue surgery in chinchillas, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 331f–333f ovariectomy, 327–328 ovariohysterectomy, 328–329 penile prolapse, 333 pyometra, 329 thoracotomy, 337 urolithiasis, 334–336 uterine prolapse, 330 uterine torsion, 329 uterine tumors, 330 Soft tissue surgery in degus, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 332f ovariectomy, 327–328 ovariohysterectomy, 328–329
593
Soft tissue surgery in degus (Continued) penile prolapse, 333 pyometra, 329 thoracotomy, 337 urolithiasis, 334–336 uterine prolapse, 330 uterine torsion, 329 uterine tumors, 330 Soft tissue surgery in ferrets, 141 caval syndrome, 155 endocrine system, 145–152 gallbladder surgery, 147f pancreatic surgery, 150–151, 151f splenectomy, 151–152, 152f surgery of adrenal gland, 145–150, 147f, 149f exploratory laparotomy, 142–143, 143f gastrointestinal (GI) system, 143–145 gallbladder surgery, 145 intestinal surgery, 143–144, 144f liver biopsy, 144–145, 145f–146f salivary mucocele resection, 143 general surgical principles, 141, 142f heartworm disease, 155 miscellaneous surgical procedures, 154–155 anal sacculectomy, 154–155 surgery of cutaneous neoplasia, 141–142 urogenital system, 152–154 castration, 154 cystotomy, 152 nephrectomy, 152 ovarian remnant, 154 ovariohysterectomy, 154 paraurethral/prostatic cysts, 153–154, 153f perineal urethrostomy, 153, 153f preputial masses, 154, 155f pyometra, 154 Soft tissue surgery in guinea pigs, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 332f–333f ovariectomy, 327–328 ovariohysterectomy, 328–329, 328f penile prolapse, 333 pyometra, 329 thoracotomy, 337 urolithiasis, 334–336, 335f uterine prolapse, 330, 330f uterine torsion, 329 uterine tumors, 330 Soft tissue surgery in rabbits, 269 abscesses, 275 anorectal papilloma removal, 274 common procedures, 270–271 castration, 271–273, 272f–273f orchidectomy, 271–273, 272f–273f ovariohysterectomy, 271, 271f correcting problems of urine scald, 275–276, 276f–277f dermoplasty and tail amputation, 275–276, 276f–277f equipment, 269 evaluating rabbit as surgical patient, 269 gastrointestinal (GI) surgery, 273–274 anastomosis, 273–274, 274f enterotomy, 273–274, 274f gastrotomy, 273 intestinal resection, 273–274, 274f surgery of large bowel, 274
594
Index
Soft tissue surgery in rabbits (Continued) postsurgical monitoring, 270 blood loss, 270 pain and analgesics, 270 presurgical treatment, 270 surgery of auditory bulla, 276–278 lateral ear canal ablation, 276–277 osteotomy of ventral bulla, 277–278 surgery of ear canal, 276–278 lateral ear canal ablation, 276–277 osteotomy of ventral bulla, 277–278 surgical techniques, 270–271 adhesion formation, 270 choice of suture material, 270 skin closures, 270–271 Soft tissue surgery in small rodents, 373 abdominal masses, 385 abdominal procedures, 384–385 abscesses, 386 analgesia, 390 cesarean section, 382 cheek pouch eversion in hamsters, 387, 388f–389f cutaneous masses, 384, 384f cystotomy, 384–385 dental abscesses in hamsters, 386 enucleation and exenteration, 386–387 focal light, 376 hemostatic aids, 377, 377f instrumentation, 374–376, 375f magnification, 376, 376f mammary gland neoplasia, 382–384, 383f orchidectomy, 378–380, 378f–380f ovarian disease, 381 ovariectomy, 381–382 ovariectomy and ovariohysterectomy, 381–382 ovariohysterectomy, 381–382 patient preparation, 377 patient support, 373–374, 374f presurgical considerations, 373–378 prolapsed bowel in hamsters, 387–389 pulmonary lobectomy, 385–386 recovery, 389–390 subcutaneous abscesses, 386 sutures, needles, and closure, 378 tail amputation, 389 uterine disease, 381 Spinal tumors in ferrets, 136–138 Splay leg in rabbits, 252 Splenectomy in ferrets, 151–152, 152f Splenomegaly in ferrets, 73 Spondylosis in rabbits, 252, 254f Squamous cell carcinoma (SCC) in ferrets, 128–129 Stress-related disorders in sugar gliders, 405–406 Subcutaneous abscesses in rabbits, 232–233, 233f Subcutaneous abscesses in small rodents, 386 Sugar gliders, 393 behavior of, 554–555 behavior problems, 554 communication behaviors, 554 introduction of new conspecifics, 555 play behaviors, 554, 554f social behaviors, 554 biology, 393–398 anatomic characteristics, 394, 394f–395f anatomy and physiology, 393–396 behavior, 396–397 caging, 397 color variations and genetics, 394–395, 395f
Sugar gliders (Continued) hand-rearing, 398 husbandry, 397–398 marsupials, 393–394 natural history, 393 nutrition and feeding, 397–398, 399t–400t reproduction, 395–396, 396f, 396t clinical techniques, 398–402 blood collection, 398–400, 399t–401t drug dosages, 401–402, 403t–404t handling and restraint, 398 radiography, 400–401, 401f–402f treatment techniques, 400 dentistry of, 469 diseases and syndromes, 402–408 bacterial diseases, 408 cataracts, 406–407 cystitis, crystalluria, and urolithiasis, 405 dental disease, 402, 404f dermatologic disorders, 405–406 ear margin canker, 406 encephalomalacia and encephalitis, 407 endocrine alopecia, 406, 406f enteritis and enteropathy, 402 failure to thrive in a joey, 405 fractures, 407 gastrointestinal disease, 402–405 impaction of paracloacal gland, 405 infectious disease, 408 infertility, 405 malnutrition, 402 musculoskeletal disease, 407 neoplasia, 408 nephritis and renal failure, 405 neurologic disease, 407–408 nutritional osteodystrophy, 407 obesity, 407 ocular injury, 406 opthalmic disorders, 406–407 parasitic diseases, 408 pouch infection and mastitis, 405 rectal prolapse, 402–404 reproductive disorders, 405 reproductive tract infection, 405 respiratory disease, 405 retrobulbar abscess, 407 self-mutilation, 406, 406f self-mutilation of penis and scrotum, 405 stress-related disorders, 405–406 traumatic injuries, 406 tremors and seizures, 407–408 urinary tract obstruction, 405 urogenital disease, 405 ophthalmologic diseases in, 530 surgery and anesthesia, 408–409 anesthesia, 408–409 castration and scrotal ablation, 409, 409f ovariohysterectomy, 409 removal of paracloacal glands, 409, 410f repair of patagium, 409 soft tissue surgery, 409 Surgery in chinchillas, soft tissue, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 331f–333f ovariectomy, 327–328 ovariohysterectomy, 328–329 penile prolapse, 333 pyometra, 329
Surgery in chinchillas, soft tissue (Continued) thoracotomy, 337 urolithiasis, 334–336 uterine prolapse, 330 uterine torsion, 329 uterine tumors, 330 Surgery in degus, soft tissue, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 332f ovariectomy, 327–328 ovariohysterectomy, 328–329 penile prolapse, 333 pyometra, 329 thoracotomy, 337 urolithiasis, 334–336 uterine prolapse, 330 uterine torsion, 329 uterine tumors, 330 Surgery in ferrets, soft tissue, 141 caval syndrome, 155 endocrine system, 145–152 adrenal gland surgery of, 145–150, 147f, 149f gallbladder surgery, 147f pancreatic surgery, 150–151, 151f splenectomy, 151–152, 152f exploratory laparotomy, 142–143, 143f gastrointestinal (GI) system, 143–145 gallbladder surgery, 145 intestinal surgery, 143–144, 144f liver biopsy, 144–145, 145f–146f salivary mucocele resection, 143 general surgical principles, 141, 142f heartworm disease, 155 miscellaneous surgical procedures, 154–155 anal sacculectomy, 154–155 surgery of cutaneous neoplasia, 141–142 urogenital system, 152–154 castration, 154 cystotomy, 152 nephrectomy, 152 ovarian remnant, 154 ovariohysterectomy, 154 paraurethral/prostatic cysts, 153–154, 153f perineal urethrostomy, 153, 153f preputial masses, 154, 155f pyometra, 154 Surgery in guinea pigs, soft tissue, 326 cervical lymphadenitis, 336–337 cutaneous dermal masses, 336 dystocia, 329–330 gastric trichobezoars, 333–334 mammary gland neoplasia, 330–331 miscellaneous procedures, 337 orchidectomy, 331–333, 332f–333f ovariectomy, 327–328 ovariohysterectomy, 328–329, 328f penile prolapse, 333 pyometra, 329 thoracotomy, 337 urolithiasis, 334–336, 335f uterine prolapse, 330, 330f uterine torsion, 329 uterine tumors, 330 Surgery in rabbits, soft tissue, 269 abscesses, 275 anorectal papilloma removal, 274 common procedures, 270–271
Index Surgery in rabbits, soft tissue (Continued) castration, 271–273, 272f–273f orchidectomy, 271–273, 272f–273f ovariohysterectomy, 271, 271f correcting problems of urine scald, 275–276, 276f–277f dermoplasty and tail amputation, 275–276, 276f–277f equipment, 269 evaluating rabbit as surgical patient, 269 gastrointestinal (GI) surgery, 273–274 anastomosis, 273–274, 274f enterotomy, 273–274, 274f gastrotomy, 273 intestinal resection, 273–274, 274f surgery of large bowel, 274 postsurgical monitoring, 270 blood loss, 270 pain and analgesics, 270 presurgical treatment, 270 surgery of auditory bulla, 276–278 lateral ear canal ablation, 276–277 osteotomy of ventral bulla, 277–278 surgery of ear canal, 276–278 lateral ear canal ablation, 276–277 osteotomy of ventral bulla, 277–278 surgical techniques, 270–271 adhesion formation, 270 choice of suture material, 270 skin closures, 270–271 thoracotomy, 278 urinary tract surgery, 274–275 nephrectomy, 275 nephrotomy, 275 pyelolithotomy, 275 Surgery in small rodents, soft tissue, 373 abdominal masses, 385 abdominal procedures, 384–385 abscesses, 386 analgesia, 390 cesarean section, 382 cheek pouch eversion in hamsters, 387, 388f–389f cutaneous masses, 384, 384f cystotomy, 384–385 dental abscesses in hamsters, 386 enucleation and exenteration, 386–387 focal light, 376 hemostatic aids, 377, 377f instrumentation, 374–376, 375f magnification, 376, 376f mammary gland neoplasia, 382–384, 383f orchidectomy, 378–380, 378f–380f ovarian disease, 381 ovariectomy, 381–382 ovariectomy and ovariohysterectomy, 381–382 ovariohysterectomy, 381–382 patient preparation, 377 patient support, 373–374, 374f presurgical considerations, 373–378 prolapsed bowel in hamsters, 387–389 pulmonary lobectomy, 385–386 recovery, 389–390 subcutaneous abscesses, 386 sutures, needles, and closure, 378 tail amputation, 389 uterine disease, 381 Surgical endoscopy, exotic mammal diagnostic and, 485 anesthesia, 485–486 complications, 501 instrumentation, 486, 487t, 488f–489f outcome, 501
Surgical endoscopy, exotic mammal diagnostic and (Continued) patient evaluation, 485 patient selection, 485 postoperative care, 501 procedures, 486–501 endotracheal intubation, 488, 493f gastroscopy/colonoscopy, 493–496, 497f–498f laparoscopy, 496–499, 499f–500f otoscopy, 486, 489f rhinoscopy, 488–493, 494f–496f stomatoscopy, 486–488, 490f–492f thoracoscopy, 499–501 tracheobronchoscopy, 488, 494f vaginoscopy/cystoscopy, 493, 497f Syphilis in rabbits, 235–236, 235f
T
Tail amputation in small rodents, 389 Tapeworm cysts in rabbits, 239 Testicular neoplasms in rabbits, 222 Thoracotomy in chinchillas, 337 Thoracotomy in degus, 337 Thoracotomy in guinea pigs, 337 Thoracotomy in rabbits, 278 Thymic carcinoma in rabbits, 263–265 Thymic lymphoma in rabbits, 263–265 Thymic masses in rabbits, 263–265, 264f–265f Thymoma/thymic lymphoma, thymic carcinoma in rabbits, 263–265 Thyroid disease in ferrets, 91, 92t Thyroid neoplasms in ferrets, 106 Ticks in ferrets, 125 Ticks in rabbits, 239 Toxemia of pregnancy in guinea pigs, 302–303 in rabbits, 254 Toxoplasmosis in rabbits, 250 Traumatic injuries in sugar gliders, 406 Trematodes in rabbits, 202 Tremors and seizures in sugar gliders, 407–408 Tumors, basal cell, in ferrets, 128 Tumors, malignant, in rabbits, 518f Tumors, mast cell, in ferrets, 127–128, 128f Tumors, spinal, in ferrets, 136–138 Tumors, uterine in chinchillas, 330 in degus, 330 in guinea pigs, 330 Tumors in ferrets basal cell, 128 gastrointestinal (GI) tract tumors, 116–117, 117f hemolymphatic system, 106–115 ancillary treatments, 114–115 chemotherapy, 110–112, 112t–115t classification of lymphoma, 106–107, 106t–107t, 107f cytologic/histologic description, 108–110, 110f–111f diagnostic imaging, 108, 109f laboratory evaluation, 108 palliative therapy, 112–113 radiation treatment, 113–114 signalment and clinical signs, 107–108, 108f treatment, 110–115 malignant peripheral nerve sheath, 119 mast cell, 127–128, 128f musculoskeletal system, 118 nervous system, 118–119, 119f reproductive tract, 118
595
Tumors in ferrets (Continued) respiratory system, 119 skin, 115–116, 115f spinal, 136–138 urinary system, 119 Tumors in ferrets, pancreatic islet cell, 92–99 advanced imaging, 96 baseline insulin and glucose concentrations, 95, 95t beyond Whipple’s triad, 95 clinical features, 93–94 clinical pathologic abnormalities, 94 clinical signs, 93–94 diagnostic approach, 95–96 diagnostic imaging, 96 differential diagnoses for fasting hypoglycemia, 94–95 etiology, 92–93 histopathology, 99 insulin: glucose ratios, 95 management of islet cell tumors, 96–99 medical management of chronic hypoglycemia, 98–99 medical therapies for active hypoglycemic crisis, 97–98 miscellaneous testing, 96 pathophysiology, 93 physical examination, 94, 94f prognosis, 99 provocative testing, 95–96 radiography, 96 signalment, 93 surgical therapy, 96–97, 96t–97t ultrasonography, 96, 96f Tumors in hamsters, malignant mesenchymal, 518f Tumors of endocrine system in ferrets adrenocortical neoplasms, 105–106, 105f insulinoma, 104–105 thyroid neoplasms, 106 Tympany in chinchillas, 312 Tyzzer’s disease in rabbits, 199–200
U
Ulcerative pododermatitis in rabbits, 235 Ultrasound, 505–506, 506f Umbilical cords, enlarged in ferrets, 59 Ureteral rupture in ferrets, 48 Urethral catheterization of male ferret in ferrets, 51f Urethral obstruction in ferrets, 50–51, 50f Urinary and reproductive systems disorders in ferrets, 46 female ferret, 54–56 female reproductive tract tumors, 55–56 hydrometra, 56, 56f hyperestrogenism, 55 mucometra, 55–56 pyometra, 55–56 jill diseases, 57–58 agalactia, 58 dystocia, 57 mammary gland neoplasia, 58 mastitis, 58 metritis, 58 postparturient hypocalcemia, 58 pregnancy toxemia, 57 pseudopregnancy, 57 kit diseases, 58–59 caring for ill kits, 58–59, 59t diarrhea, 59 enlarged umbilical cords, 59 neonatal conjunctivitis, 59 neonatal mortality and deformities, 59
596
Index
Urinary and reproductive systems disorders in ferrets (Continued) normal kit, 58 splay-legged kits, 59 male ferret, 54 cryptorchidism, 54 male reproductive tract tumors, 54, 54f penile lesions, 54 prostatic lesions, 54 periparturient disease, 56–59 breeding ferrets, management of, 57 jill diseases, 57–58 kit diseases, 58–59 normal breeding, 56–57 polycystic kidney disease, polycystic kidney in ferret with acute renal failure, 47f renal disease and renal failure, 47–48 acute renal failure, 47 chronic renal failure, 47 diagnosis and treatment, 47–48 reproductive system disorders, 54–56 female ferret, 54–56 male ferret, 54 urethral obstruction urethral catheterization of male ferret, 51f urethral obstruction, 50f urinary and reproductive systems disorders periparturient disease, 56–59 reproductive system disorders, 54–56 urinary system disorders, 46–54 urinary system disorders, 46–54 Aleutian disease, 48 bladder neoplasia, 50 cystitis, 49–50 hydronephrosis, 47 nephrocalcinosis, 48 paraurethral cysts or paraurethral disease, 53–54 polycystic kidney disease, 46–47 prostatic cysts, 51–52, 51f–52f prostatic tumors, 53 prostatitis and prostatic abscess, 53, 53f pyelonephritis, 48 renal cysts, 46 renal disease and renal failure, 47–48 renal neoplasia, 48 ureteral rupture, 48 urethral obstruction, 50–51 urinary incontinence, 50 urolithiasis, 49 Urinary bladder inversion in rabbits, 229 Urinary diseases in African hedgehogs, 420 Urinary incontinence in ferrets, 50 in rabbits, 229 Urinary obstruction in small mammals, emergency treatment of, 541, 543f Urinary system disorders in ferrets, 46–54 tumors in ferrets, 119
Urinary systems disorders in rabbits, reproductive and reproductive system disorders, 217–223 abdominal pregnancy, 220 abortion and resorption, 220 cryptorchidism, 221, 222f dystocia, 220 endometrial hyperplasia or uterine polyps, 218–219 endometrial venous aneurysms, 221 endometritis, 219 epididymitis, 222 hydrometra, 221 orchitis, 222 pregnancy toxemia, 219 prolapsed vagina, 220–221 pseudopregnancy, 219–220 pyometra, 219 reduced fertility, 220 retained fetuses, 220 testicular neoplasms, 222 uterine adenocarcinoma, 218, 218f uterine atresia, 221, 221f uterine torsion, 221 uterus unicornis, 221 venereal spirochetosis, 222–223 urinary system disorders, 223–229, 224f encephalitozoonosis, 228 hypercalciuria, 225–227, 225f–227f hypervitaminosis D, 228 nephrotoxicity, 228 psychogenic polyuria and polydipsia, 229 red urine, 229 renal adipose deposition, 228 renal agenesis, 228 renal cysts, 228 renal failure, 227–228 urinary bladder inversion, 229 urinary incontinence, 229 urinary tract tumors, 229 urolithiasis, 225–227, 225f–227f Urinary tract infection in guinea pigs, 301 obstruction in sugar gliders, 405 tumors in rabbits, 229 Urinary tract surgery in rabbits, 274–275 nephrectomy, 275 nephrotomy, 275 pyelolithotomy, 275 Urogenital disease in sugar gliders, 405 Urolithiasis in chinchillas, 334–336 in degus, 334–336 in ferrets, 49 in guinea pigs, 299–301, 300f, 334–336, 335f in rabbits, 225–227, 225f–227f in sugar gliders, 405 Uropathies in guinea pigs, 301 Uterine adenocarcinoma in rabbits, 218, 218f Uterine atresia in rabbits, 221, 221f
Uterine disease in small rodents, 381 Uterine prolapse in chinchillas, 330 in degus, 330 in guinea pigs, 302, 330, 330f Uterine torsion in chinchillas, 329 in degus, 329 in guinea pigs, 329 in rabbits, 221 Uterus unicornis in rabbits, 221 Uveitis in rabbits, 525–526, 526f
V
Vaccinations in ferrets, 15–16 canine distemper, 15 rabies, 15–16 vaccine-associated adverse events, 16 Vaccine-associated adverse events in ferrets, 16 Vaginitis in guinea pigs, 303 Valvular disease in rabbits, 261–262 Valvular heart disease in ferrets, 69, 69f Vascular disease in rabbits, 262 Venereal spirochetosis in rabbits, 222–223 Vertebral fracture or luxation in rabbits, 251–252, 252f–253f Viral diarrhea in ferrets, 37 canine distemper virus, 37 coronavirus, 37, 37f influenza virus, 37 rotavirus, 37 Viral diseases, 560–562, 561f Viral diseases of digestive tract in rabbits, 200–201 hemorrhagic disease virus (RHDV), 200–201 papillomatosis, 200 rabbit enteric coronavirus, 200 rotavirus, 200 Viral infections in chinchillas, 319 Viral infections in rabbits, 239–241, 251 herpes simplex virus (HSV), 251 myxomatosis, 239–240 oral papillomavirus, 240 rabbit (Shope) fibroma virus, 240 rabbit (Shope) papillomavirus, 240 rabbitpox, 240–241 rabies, 251 Viral pneumonia in guinea pigs, 298–299, 299f Vitamin C deficiency in guinea pigs, 305–306, 306f Vomiting in ferrets, 40
Z
Zoonotic diseases, 557 allergic reactions, 564 bacterial diseases, 557–560, 558f–559f mycotic diseases, 563 parasitic diseases, 562–563, 562f–563f viral diseases, 560–562, 561f