Advances in
PARASITOLOGY VOLUME
66
EDITORIAL BOARD M. COLUZZI Department of Public Health Sciences, Section of Parasitology ‘Ettore Biocca’ ‘Sapienza – Universita` di Roma’ P. le Aldo Moro, 5, 00185 Roma, Italia
C. COMBES Laboratoire de Biologie Animale, Universite´ de Perpignan, Centre de Biologie et d’Ecologie Tropicale et Me´diterrane´enne, Avenue de Villeneuve, 66860 Perpignan Cedex, France
D. D. DESPOMMIER Division of Tropical Medicine and Environmental Sciences, Department of Microbiology, Columbia University, 630 West 168th Street, New York, NY 10032, USA
J. J. SHAW Instituto de Cieˆncias Biome´dicas, Universidade de Sa˜o Paulo, av. Prof Lineu Prestes 1374, 05508-990, Cidade Universita´ria, Sa˜o Paulo, SP, Brazil
K. TANABE Laboratory of Malariology, International Research Center of Infectious Diseases. Research Institute for Microbial Diseases, Osaka University, 3-1 Yamada-Oka, Suita, 565-0871. Japan
Advances in
PARASITOLOGY VOLUME
66 Edited by
D. ROLLINSON Department of Zoology The Natural History Museum Cromwell Road, London, UK
S. I. HAY Senior Research Fellow Malaria Public Health & Epidemiology Group Centre for Geographic Medicine KEMRI/University of Oxford/Wellcome Trust Collaborative Programme, Nairobi, Kenya
AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier
Academic Press is an imprint of Elsevier 84 Theobald’s Road, London WC1X 8RR, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, UK 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA First edition 2008 Copyright # 2008 Elsevier Ltd. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher. Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865853333; email:
[email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material. Notice No responsibility is assumed by the publisher 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. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made. ISBN: 978-0-12-374229-2 ISSN: 0065-308X For information on all Academic Press publications visit our website at books.elsevier.com Printed and bound in The Netherlands 07 08 09 10 11 10 9 8 7 6 5 4 3 2 1
CONTENTS
Contributors Preface Obituary
1. Strain Theory of Malaria: The First 50 Years
vii ix xi
1
F. Ellis McKenzie, David L. Smith, Wendy P. O’Meara, and Eleanor M. Riley Introduction Background Clinical Virulence Reaction to Anti-malarial Remedies Infectivity Antigenic Properties Latency and Relapse Summary and Discussion Acknowledgements References 1. 2. 3. 4. 5. 6. 7. 8.
2. Advances and Trends in the Molecular Systematics of Anisakid Nematodes, with Implications for their Evolutionary Ecology and Host–Parasite Co-evolutionary Processes
2 3 6 9 12 16 28 33 38 38
47
Simonetta Mattiucci and Giuseppe Nascetti Introduction Molecular Systematics of Anisakid Nematodes The Current Taxonomy Phylogenetic Systematics of Anisakid Nematodes Genetic Differentiation in Anisakids Host–Parasite Cophylogeny Host Preference, Ecological Niche and Competition Anisakids as Biological Indicators Conclusions and Identification of Gaps in Our Knowledge of Anisakids to be Filled by Future Research Acknowledgements References 1. 2. 3. 4. 5. 6. 7. 8. 9.
49 51 55 91 100 108 114 119 130 137 137
v
vi
Contents
3. Atopic Disorders and Parasitic Infections
149
Aditya Reddy and Bernard Fried Introduction Atopic Disorders Relationship of Parasites to Atopic Disorders Laboratory Studies on Atopy Using Selected Parasites and Rodent Models 5. Concluding Remarks References 1. 2. 3. 4.
4. Heartworm Disease in Animals and Humans
150 151 154 175 182 184
193
John W. McCall, Claudio Genchi, Laura H. Kramer, Jorge Guerrero, and Luigi Venco Introduction (Biology and Life Cycle) Epidemiology in Domestic and Wild Hosts Pathogenesis, Immunology and Wolbachia Endosymbiosis Canine Heartworm Disease Feline Heartworm Disease Heartworm Disease in Ferrets Human Dirofilariosis Emerging Strategies in Heartworm Treatment and Control Acknowledgments References 1. 2. 3. 4. 5. 6. 7. 8.
195 199 209 222 233 253 258 263 265 265
Index
287
Contents of Volumes in This Series
295
See Colour Plate Section in the back of this book
CONTRIBUTORS
Bernard Fried Department of Biology, Lafayette College, Easton, Pennsylvania 18042, USA. Claudio Genchi DIPAV, Sezione di patologia Generale e Parassitologia, Universita` degli Studi di Milano, Via Celoria 10, 20133 Milano, Italy. Jorge Guerrero Department of Pathobiology, School of Veterinary Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104, USA. Laura H. Kramer Dipartimento di Produzione Animale, Universita` di Parma, via del Taglio 8, 43100 Parma, Italy. Simonetta Mattiucci Department of Public Health Sciences, Section of Parasitology, ‘‘Sapienza’’—University of Rome, P.le Aldo Moro, 5, 00185 Rome, Italy. John W. McCall Department of Infectious Diseases, College of Veterinary Medicine, University of Georgia, Athens, Georgia, 30602, USA. F. Ellis McKenzie Fogarty International Center, Building 16, National Institutes of Health, Bethesda, Maryland 20892, USA. Giuseppe Nascetti Department of Ecology and Sustainable Economic Development—Tuscia University—Via S. Giovanni Decollato, 1, 01100 Viterbo, Italy. Wendy P. O’Meara Fogarty International Center, Building 16, National Institutes of Health, Bethesda, Maryland 20892, USA. Aditya Reddy Department of Biology, Lafayette College, Easton, Pennsylvania 18042, USA.
vii
viii
Contributors
Eleanor M. Riley Department of Infectious and Tropical Diseases, London School of Hygiene and Tropical Medicine, London WC1 E 7HT, United Kingdom. David L. Smith Zoology Department and Emerging Pathogens Institute, 223 Bartram Hall, University of Florida, Gainesville, Florida 32611, USA. Luigi Venco Clinica Veterinaria Citta` di Pavia, Viale Cremona 179, 27100 Pavia, Italy.
PREFACE
Ellis McKenzie, of the Fogarty International Center at the National Institutes of Health, and colleagues open this volume with a review of the history of strain theory in malaria research. This comprehensive review looks back to the origins of the concept of ‘varieties, strains or races’ of the Plasmodium species that cause human disease. The historical perspective allows insight not only to how these ideas developed but also how they affected clinical practice and epidemiological study. The development of these theories is then explored in relation to several themes: parasite phenotypes related to clinical virulence, reactions to anti-malarial drugs, infectivity to mosquitoes, antigenic properties and host immunity, latency and relapse. The authors conclude by discussing how these definitions of strain have evolved in relation to discoveries around each of these themes and by commenting on where ambiguity in working definition of a malaria strain remains. The next paper concerns the molecular systematics of the anisakid nematodes of the genera Anisakis, Pseudoterranova and Contracaecum. Simonetta Mattiucci from the University of Rome and Giuseppe Nascetti from Tuscia University, Italy, draw on their wealth of experience on working with this group and present a detailed account of the current understanding relating to the relationships of the different species involved and the implications for evolutionary ecology. It has long been known that morphological characters alone are insufficient to differentiate the species and each genus includes a number of sibling species. Once good molecular markers are available, it is possible to examine questions relating to host specificity and distribution. Comparing host and parasite phylogenies can be a powerful way of determining co-divergence and host-switching events, and the authors provide interesting insights by comparing Anisakis and the cetaceans and Contracaecum and the pinnipeds. Aditya Reddy and Bernard Fried of the Department of Biology, Lafayette, Easton, USA provided an interesting overview of the relationship between atopic disorders and parasitic infections. This is a topic that attracts much interest, especially in relation to helminth infections, but the associated literature tends to be scattered in diverse journals. Both helminth infections and allergy are common diseases, and the general observation is that helminth infections tend to be negatively associated with atopy, prevalence of allergic diseases and the severity of asthma.
ix
x
Preface
This review explores the protective mechanisms against atopic disorders which may be associated with parasites. This volume concludes with a chapter reviewing the biology and life cycle of the parasite Dirofilaria immitis, the causative agent of dog and cat heartworm. The chapter is contributed by John W. McCall, of the College of Veterinary Medicine of the University of Georgia, and an international assembly of co-authors. Advances in the understanding of its global distribution and prevalence of heartworm are outlined and the pathogenesis and immunology of infection discussed. The current understanding of the potential role of the Wolbachia endosymbiont in inflammatory and immune responses are discussed, along with the antibiotic treatment of infected animals. A large part of the chapter reviews the clinical presentation, diagnosis, prevention, therapy and management of the disease in dog, cats and ferrets. There is also a discussion of heartworm infection in humans, with notes on other epizootic filarial infections, particularly D. repens in Europe. The chapter concludes by examining novel treatments and highlighting the potential role of tetracycline antibiotics in adulticidal therapy. D. ROLLINSON S. I. HAY
OBITUARY Ralph Muller (1933–2007)
Ralph Muller, who died on 11 October 2007 as a result of prostate cancer, was a co-editor of Advances in Parasitology for 29 years. In 1978, Ralph (and I) joined the late Professor Russell Lumsden, then at the London School of Hygiene and Tropical Medicine, as assistant editors when Professor Lumsden took over as senior editor following the death of the founder, Professor Ben Dawes. When Professor Lumsden himself retired in 1981, Ralph and I continued to edit the publication; in 1994, we were joined by Dr. David Rollinson, of the British Museum (Natural History) in London. xi
xii
Obituary
I retired from this role at the end of 2006, to be replaced by Dr. Simon Hay (University of Oxford, UK), but Ralph continued in harness until shortly before his death. Some years before becoming an editor, in 1971, Ralph had begun his association with Advances by contributing a lengthy article on dracunculiasis to volume 9 (pp. 73–151). Ralph was one of the foremost parasitologists of the twentieth century in the UK. Beginning his life’s work as a parasitologist in 1955, when he graduated in zoology from Queen Mary College (University of London), Ralph moved to King’s College (London) where he worked on the maintenance in vitro of Haplometra cylindracea, a trematode living in the lungs of frogs, thus obtaining his Doctor of Philosophy (PhD) degree in 1958. Ralph was awarded Fellowship of the Institute of Biology in 1972 and the degree of Doctor of Science by London University in 1989. Ralph continued to work in King’s College as a research fellow until 1960. He first obtained ‘hands on’ experience of parasitology in the tropics during the two years, from 1960 to 1962, as a scientific officer working for the UK Overseas Development Administration on the control of schistosomiasis. Although based in London, this work involved visits to Kenya, Tanzania and Uganda, all then part of British East Africa. Following this, Ralph crossed the continent to become lecturer and chief of the subdepartment of parasitology in the University of Ibadan in Nigeria from 1962 to 1966. Here he became acquainted with the guinea worm Dracunculus medinensis, in which he retained a life-long interest. In 1966, he joined the Department of Parasitology of the London School of Hygiene and Tropical Medicine, where his continued study of guinea worm infection helped to draw attention to the importance of this infection which, though not of itself fatal, was a cause of considerable morbidity in affected populations and could often be the portal of entry of various pathogenic bacteria, including Clostridium tetani. Ralph later put his experience of dracunculiasis to good use through his association with the World Health Organization’s global programme to eradicate guinea worm infection—a target achieved in Asia and many, but not yet all, African countries; the current target date (not the first!) for global eradication is 2009. After 15 years at the London School, Ralph left in 1981 to become director of the International Institute of Parasitology (IIP) in St Albans, Hertfordshire, in the UK (where Ralph also lived). The Institute was part of an inter-governmental organization founded as the Commonwealth Agricultural Bureaux (CAB), and later was renamed as CAB International. The institute’s major function was to disseminate information concerning helminth parasites, primarily through the abstracting journal Helminthological Abstracts. This journal was later joined by the complementary Protozoological Abstracts. IIP was also instrumental in setting up relevant research projects in tropical countries. Ralph remained in this
Obituary
xiii
post until 1993; to the regret of many parasitologists, the Institute was closed in 1998, though publication of the abstracting journals continues. Ralph served as a Council member of the Royal Society of Tropical Medicine and Hygiene from 1982 to 1985. From 1993, after his retirement from the IIP, until the end of his life, Ralph continued to put his wide knowledge of helminthology at the disposal of students as a Visiting Lecturer at Imperial College in London and an Honorary Senior Lecturer at the London School of Hygiene and Tropical Medicine. In addition to editing Advances in Parasitology for many years, Ralph edited the primary research publication Journal of Helminthology (originally published by the London School of Hygiene and Tropical Medicine, now by Cambridge University Press), a post which he filled from 1972 to 1980 and again from 1987 to 1995. Ralph was the author or co-author of well over 100 scientific papers and also of two books: Worms and Disease: A Manual of Medical Helminthology (William Heinemann, 1975) and Medical Parasitology, with myself as coauthor (J. P. Lippincott Company/Gower Medical Publishing, 1990). The former book, re-titled Worms and Human Disease and with additional material by Derek Wakelin, achieved a second edition (CABI Publishing, 2002). The later version regrettably was devoid of the first edition’s delightful frontispiece, a reproduction of a detailed drawing of a sorrowful case of multiple guinea worm infection, reprinted from a publication of 1674, Exercitationes de Vena Medinensis et de Vermiculis capillaribus infantium by G. H. Velschius (Augsburg). The inclusion of this frontispiece seems to me to embody two of Ralph’s many good qualities—his dry humour and also his wide-ranging interest in and beyond the basics of helminthology. While being the director of the IIP, Ralph also edited a computerized Bibliography of Onchocerciasis (1841–1985), published by CAB International in 1987. Having known Ralph, mainly as an editorial colleague on Advances in Parasitology, for some 30 years, like many others I shall greatly miss, as well as happily remember, him as a distinguished helminthologist and a true friend. Apart from his knowledge of helminthology, which was both broad and deep, I remember in particular his calmness, his good humour and his generally ‘laid back’ attitude, all of which were of great value in the occasional moments of hustle and crisis. These qualities he courageously continued to manifest throughout his final illness. In the 1970s and 1980s, Ralph and I made several visits to Libya as external examiners in the Medical School at Garyounis University in Benghazi and the Medical Faculty of Al Fatah University in Tripoli. Ralph’s sense of humour contributed greatly to my enjoyment of these visits. I remember his delight on one such visit when he found that, by bounding up the stairs to our shared hotel room, he would be just in time to watch the televised broadcast of a World Cup football international game in which England was participating; regrettably, being perhaps rather less
xiv
Obituary
FIGURE 1
Ralph Muller PhD, DSc, CBiol, FIBiol.
FIGURE 2 Ralph Muller in the field, somewhere in West Africa (photo copyright # CAB International; reproduced by permission).
Obituary
xv
interested in sporting events than Ralph (who was himself a keen soccer player), I cannot remember whether they won. He was somewhat less delighted to find, on another occasion, that we were expected to share not only a room but also a double bed. I should add that a second bed was hastily procured. I am very grateful to Barnaby and Harriet Muller for their helpfulness in providing me with some of the above information. JOHN BAKER
This page intentionally left blank
CHAPTER
1 Strain Theory of Malaria: The First 50 Years F. Ellis McKenzie,* David L. Smith,† Wendy P. O’Meara,* and Eleanor M. Riley‡
Contents
Introduction Background Clinical Virulence Reaction to Anti-malarial Remedies Infectivity Antigenic Properties 6.1. Homologous and heterologous response 6.2. Clinical and parasitological response 6.3. Superinfection 7. Latency and Relapse 8. Summary and Discussion Acknowledgements References
Abstract
From the 1920s to the 1970s, a large body of principles and evidence accumulated about the existence and character of ‘strains’ among the Plasmodium species responsible for human malaria. An extensive research literature examined the degree to which strains were autonomous, stable biological entities, distinguishable by clinical, epidemiological or other features, and how this knowledge could be used to benefit medical and public health practice. Strain theory in this era was based largely on parasite phenotypes related to
2 3 6 9 12 16 16 19 23 28 33 38 38
1. 2. 3. 4. 5. 6.
* Fogarty International Center, Building 16, National Institutes of Health, Bethesda, Maryland 20892, USA {
{
Zoology Department and Emerging Pathogens Institute, 223 Bartram Hall, University of Florida, Gainesville, Florida 32611, USA Department of Infectious and Tropical Diseases, London School of Hygiene and Tropical Medicine, London WC1 E 7HT, United Kingdom
Advances in Parasitology, Volume 66 ISSN 0065-308X, DOI: 10.1016/S0065-308X(08)00201-7
#
2008 Elsevier Ltd. All rights reserved.
1
2
F. Ellis McKenzie et al.
clinical virulence, reactions to anti-malarial drugs, infectivity to mosquitoes, antigenic properties and host immunity, latency and relapse. Here we review the search for a definition of ‘strain’, suggest how the data and discussion shaped current understandings of many aspects of malaria and sketch a number of specific connections with perspectives from the past 30 years.
1. INTRODUCTION In the early 1920s, as debates about the number and nature of species that cause human malaria were receding, the idea emerged that each of these species consists of ‘varieties, strains or races’. Over the next 50 years, a complex set of ideas developed about such entities—whether they were discrete, independent, mutable; what traits distinguished them; how they affected clinical and epidemiological observations and interventions. By the late 1970s, cloning and cultivation of P. falciparum were possible, serologic and molecular techniques were developing rapidly, and concepts emerging from the rise of laboratory-based studies were accompanied by a shift in language: ‘strain’ and ‘race’ were largely displaced by the new terms ‘clone’ and ‘isolate’. In the mid-1990s, the concept of a ‘strain’ reemerged in anticipation of a vaccine protective against a subset of circulating parasites. This ‘strain theory’ assumed that malaria comprised discrete, independently transmitted, immutable entities, and concluded that ‘control of malaria through vaccination may be far easier than previously assumed’ (Gupta and Day, 1994a). Related theory was invoked to explain immunity to clinical malaria (Gupta and Day, 1994b), and, more recently, to describe immunity to var gene products (Gatton and Cheng, 2004). Early investigations of ‘strains’ in malaria were devoted to understanding parasite phenotypes, but, as rapid and reliable molecular techniques for determining parasite genotypes were developed, it seemed clear that ‘strains’ could be distinguished with unprecedented precision, either directly or by mapping genotypes to some smaller set of phenotypes. To do so, however, would require at least a provisional definition of ‘strain’. Here we review the initial search for a definition of ‘strain’, as the theory developed in interplay with practice in the 1920s–1970s. That search is not only of historical interest, but is also relevant as contemporary studies begin to revisit and rediscover important aspects of parasite phenotypes. This historical review points to important themes for contemporary malariology, and reminds us that sorting and classifying the objects of study critically determines how research is framed and pursued. Theories do not simply catalogue observations: they formulate general principles explaining many specific observations in terms of relatively few underlying entities, forces and relationships, so the
Strain Theory of Malaria: The First 50 Years
3
evidence accumulating for or against a theory often informs a wider field. Thus the pursuit of ‘strains’ was integral to the understanding developed in the 1920s–1970s of many aspects of malaria, notably its immunology— the now-familiar distinction between clinical and parasitological aspects of response, for instance. Here, we quote from the original literature as much as possible, to emphasize the richness of the resource and the degree to which some key points have been echoed and others neglected in recent literature. In a box at the end of each main section, we sketch several such connections between historical perspectives and current knowledge.
2. BACKGROUND ‘Shall we not be obliged to say that tertian and quartan agues are divine too, for nothing can be more regular than the process of their recurrence? But all such phenomena call for rational explanation’ (Cicero, 45BC). For millennia, the ‘intermittent’ fevers associated with chills (‘agues’) were distinguished and classified on the basis of periodicity—quotidian (daily), tertian (every other day) or quartan (every third day). In 1880, Laveran described crescent-shaped parasites in the blood of a soldier suffering the fevers of ‘paludism’ (malaria). He later distinguished an additional 46 parasite forms, and over the next 40 years maintained that ‘the different forms in which the haematozoa of paludism present themselves belong to one and the same polymorphic parasite’ (Laveran, 1893). In the late 1880s, Golgi linked periodicity in malaria fevers to parasite replication cycles, showed that morphologically distinct parasites were responsible for tertian and quartan fevers, attributed quotidian fevers to the presence of ‘different parasite generations reaching complete development at a day’s interval’ and, while noting that ‘the numerous varieties of intermittent malarial fevers reported are, in the very great majority simple varieties or combinations of the two fundamental types (tertian and quartan fevers)’, posited a third distinct type, a tertian parasite characterized by crescent forms, quinine resistance and ‘irregular’ fevers (Golgi, 1889). In the 1890s, Marchiafava, Bignami, Celli and their colleagues argued that ‘there is no group of fevers which are naturally and per se irregular, but fevers of every class may become irregular, and in different ways’ (Marchiafava and Bignami, 1894). Like Golgi, and Laveran (with respect to ‘parasitic elements’), they recognized that many problems in interpretation arose from concurrent infections by different parasite species and ‘colonies’ (Golgi’s ‘generations’) within a species, given that fevers in some way followed the relative densities, transitions or transformations of parasites: ‘difficulty is met with in the study of the mixed malarial
4
F. Ellis McKenzie et al.
infections . . . it is easy to predict the possibility of a very large series of combinations, caused by the number of the parasitic colonies, by the way in which their life cycles are, so to say, interwoven, &c.; as a resultant therefrom, the fever takes the most different courses’ (Marchiafava and Bignami, 1894). But they insisted on the existence of four distinct entities, including a ‘true’ quotidian parasite. They differentiated springtime tertians and quartans from ‘malignant’ aestivo-autumnal (summer-fall) crescent producers, and in the latter distinguished tertians and quotidians: ‘The spring tertian and quartan parasites give rise to the mild forms; the aestivo-autumnal tertian and, more rarely, aestivo-autumnal quotidian . . . give rise to the severe forms’ (Celli, 1901). In 1910, Ross who had demonstrated that malaria parasites are transmitted by mosquitoes, wrote of ‘three different types of fever, the quartan, the tertian, and the irregular or malignant type . . . Since the time of Golgi, all observers admit . . . that these three types are different species . . . Some authors consider that there are two if not three varieties (or ? species) of malignant parasites. I am inclined to agree with them, but have not yet satisfied myself sufficiently on the point to admit it in my classification’ (Ross, 1910). He too noted that some patients had concurrent infections with different parasite species, each of which might consist of ‘two or three sets sporulating [i.e. completing schizogony] on different days . . . The rule generally accepted is that each set of parasites continues it own evolution independently of other sets which may be present. But much more precise work requires to be done on this point’. The ‘mild’ tertian and quartan parasites are now known as Plasmodium vivax and Plasmodium malariae, respectively, the ‘malignant’ tertians as Plasmodium falciparum (‘falciform’ means ‘crescent-shaped’, a reference to the distinctive gametocytes—the forms transmissible to mosquitoes—of P. falciparum). Quotidian and irregular fevers are rarely mentioned in recent literature, but would be attributed to double tertian, triple quartan or other combinations of infections. The last of the four species known to cause malaria infections in humans in nature, the ‘tertian’ Plasmodium ovale, was fully described in 1922 (Stephens, 1922). In that same year, citing several observations in the literature—notably on the low gametocyte production in P. falciparum infections on the coast of West Africa, compared to those in Italy and Macedonia, and on the development of clinical malaria in ‘resistant’ West Africans, who moved from the coast inland, or between countries— Marchoux made a speculative leap: ‘There are not only 3 species or varieties of Plasmodium, but 3 groups within each of which there exist a considerable number of varieties’ (Marchoux, 1922). Laveran, Golgi, Marchiafava, Bignami, Celli, Ross and their contemporaries were familiar with experimental transfers of malaria parasites by blood inoculation, and had noted, for instance, that ‘one species of
Strain Theory of Malaria: The First 50 Years
5
parasite often disappears from the blood upon inoculation with a different species’ (Deaderick, 1909). Wagner-Jauregg observed that patients with severe syphilis were sometimes ‘healed through intercurrent infectious diseases’, which led him to ‘intentionally imitate this experiment of nature’ through blood inoculations of malaria (WagnerJauregg, 1922), which in turn led to his 1927 Nobel Prize. Before the advent of effective antibiotics, tens of thousands of neurosyphilis patients were treated in malariatherapy clinics in the United Kingdom, the United States, continental Europe and elsewhere. A 1984 review concluded that ‘malariatherapy was less expensive and produced clinical improvements more frequently and more rapidly than did the best drug treatments’ (Chernin, 1984). Contemporaneous descriptions of procedures are given in Badenski (1936), de Rudolf (1927), Kupper (1939) and Lomholt (1944). A recent, independent, explicit analysis of relevant ethical issues accompanies extensive information about patient participation and treatment, and descriptions of procedures used in two of the major US facilities, in Collins and Jeffery (1999). Parasites were transferred to patients by mosquito bite or by blood inoculation. Because most experts thought that the probability of the neurosyphilis cure was related in some way to the number and intensity of fevers, clinic staff often gave subcurative doses of anti-malarial drugs to manage infections accordingly, and then full therapeutic doses at the end of the treatment regimen. Patients not cured of syphilis by initial infections were sometimes re-infected, typically with some variation in material or procedure. As clinics tried new methods and combinations, their results appeared in research papers and at conferences; much of our current knowledge of malaria is founded on these observations. With respect to Marchoux’s ‘varieties’ and their classification, the practice of malariatherapy meant that ‘now strains of widely diverse origins can be brought together and inoculated into patients lying side by side in the same hospital’ (Hackett, 1937). In the 1920s and 1930s, the results reported from malariatherapy, in conjunction with several influential malaria experiments in birds and non-human primates, and field observations in malaria-endemic countries, produced ‘general agreement that within each species of malaria parasite there are races or strains which can be recognised as being distinct by their clinical virulence, their infectivity, their reaction to antimalarial remedies and their antigenic properties’ (James and Ciuca, 1938). This characterization of ‘races’ or ‘strains’ had a strong practical bent: ‘clinical virulence’ and ‘reaction to antimalarial remedies’ were important because therapeutic malaria infections should produce fevers and other symptoms at sufficient but not excessive levels, and should be manageable; ‘infectivity’ and ‘antigenic properties’ were important because parasite transfer and patient re-infection should be reliable and
6
F. Ellis McKenzie et al.
predictable. It soon became clear that ‘the latency and relapse patterns of malarial infections vary with the type and geographic origin of the specific parasites’ (Schwartz et al., 1950), and that this fifth distinguishing characteristic had a similar practical importance. In this paper, we use these five characteristics—clinical virulence, reaction to anti-malarial remedies, infectivity, antigenic properties, and latency and relapse—as a framework for reviewing the development of strain theory from the 1920s to 1970s.
3. CLINICAL VIRULENCE By the 1920s, it was ‘well known that infections vary greatly in severity, and that in some localities nearly every infection is pernicious, while in others pernicious symptoms do not develop, although the same plasmodium is presumably the cause of both’ (Craig, 1926). In India, for instance, ‘malarial fevers are usually most severe and persistent in low-lying coast districts’ (Hehir, 1927). If differences between parasites were responsible for ‘the common observation that in some parts of the world malignant malaria is more malignant than in others’ (James et al., 1932), then, for malariatherapy, ‘pure tested strains, if available for inoculation, are far and away preferable, since not all strains are equally suitable. Some cause only quite mild reactions, whilst others produce severe reactions with more pronounced clinical symptoms’ (Nocht and Mayer, 1937). It was also thought ‘possible that different strains of Plasmodium vivax give different clinical types of malaria’ (Grant, 1923), and that ‘the tendency also of P. falciparum to select certain organs—the intestinal wall, the omentum, the brain—for localization has been explained by tropisms developed by certain strains’ (Hackett, 1937). Though differences in virulence were often said to be dramatic, the differences were seldom specified in terms of symptoms, and, when specified—for example, ‘the strains under review differed in regard to manner of onset, character of the fever and highest pulse-rates’ (de Rudolf, 1924)—were seldom quantified. The clinical differences most often specified and quantified between strains related to fevers—‘a very marked difference between the two with respect to the height of temperature produced’, for example (Bunker and Kirby, 1925). Later, in the 1940s, the periodicity of fevers with the Baltimore, St. Elizabeth and New Hebrides strains of P. vivax was reported as 41.5, 43.4 and 45.8 h, respectively, which ‘suggests that each strain might have a characteristic periodicity . . . [which] will be a valuable point in distinguishing strains’ (Young, 1944). Strains were typically identified by their geographic site or clinic of origin, the former sometimes assumed rather than known. For instance, Horton Hospital, in England, most often used the ‘Madagascar’ strain of P. vivax, obtained in 1925 from an Indian seaman at an English port
Strain Theory of Malaria: The First 50 Years
7
and named for the putative source of his infection. Compared to a ‘Holland’ strain of P. vivax judged inappropriate for malariatherapy, the Madagascar strain had a shorter incubation period (interval between inoculation and first fever), a lower frequency of spontaneous recovery and higher frequencies of quotidian fever and fevers above 106.4oF (James and Ciuca, 1938). With P. falciparum, Horton Hospital staff found that ‘the Italian and Sardinian strains are more severe than any we have obtained from India and Africa’, based largely on their observation that ‘severe clinical symptoms’ developed much more rapidly after inoculation. Roumanian strains of P. falciparum were found to be less severe than the Italian-Sardinian, but more severe than the Indian–African (Shute and Maryon, 1954). Another ‘means of comparing the severity of the cases’ at Horton Hospital was the ‘longest period of fever without a fall to the normal temperature’, which averaged 74 h with the P. falciparum Rome I and Sardinia strains, 37 h with the Indian I and Indian II strains. Further evidence of these ‘striking clinical differences’ was that the Italian strains required eight times as much quinine for treatment as did the Indian strains, on average, and ‘continued to relapse for a much longer time’ (James et al., 1932). The application of these studies was seen as urgent and direct: ‘physicians in all malarious parts of the world should endeavour as soon as possible to add to existing information on the clinical virulence of the particular strains of P. falciparum prevalent in the countries where they work. Are all the strains in India as mild as those reported in the paper and are all the Italian strains as virulent? If the patient is able to say in what place he contracted the infection, shall we have at hand information indicating what will be the probable course of his illness?’ (James et al., 1932). As the comparisons based on quinine and ‘relapses’ suggest, however, studies of differing virulence were entangled with questions about the nature of parasite drug response and latency, the role of the host and many other factors under investigation, including ‘whether the terms ‘‘immunity’’, ‘‘virulence’’, ‘‘infectivity’’, ‘‘epidemic strains’’, etc. current in work on immunology as it relates to bacterial infections should be used in work on the immunology of malaria’ (James and Ciuca, 1938). This succession of quotes from the Horton Hospital points to the interesting context of their use of the term ‘virulence’ during the 1930s. They maintained that ‘within the species there are various geographical races which . . . can be recognized as being distinct by their clinical virulence’ (James et al., 1932), and that ‘we have no evidence that the inherent virulence of a particular strain can be increased or diminished’ (James et al., 1936). However, they also revealed that ‘at Horton between 1925 and 1930 we succeeded in increasing the physical vigour and activity of an endemic strain of P. vivax from Madagascar to the degree in which it
8
F. Ellis McKenzie et al.
caused this severe ‘‘epidemic type’’ of the disease in 80% of our cases’ (James et al., 1936). The procedure was not described (see Shute, 1951 for a schematic), but ‘careful selection of donors and recipients’ (James and Ciuca, 1938) had produced ‘a strain which multiplies freely and vigorously’ and thereby increased severity because, all else equal, ‘the number of parasites govern the severity of the case’ (James et al., 1936). This increase in severity was ‘not an increase in virulence’, however, given ‘that ‘‘virulence’’ implied the possession or elaboration of a poisonous or venomous active principle (‘‘toxin’’). If so, it was not a good word to apply to the malaria parasite in which . . . no ‘‘toxin’’ had been found’ (James et al., 1936). Furthermore, it had become ‘evident that ‘‘infectivity’’, ‘‘virulence’’, and so on are relative terms depending as much on the receptivity of the host as on the biological properties of the parasite’ (James and Ciuca, 1938). Thus, at Horton Hospital, it appeared that ‘virulence’ might refer to an interaction between a particular host and a fixed property of a strain, in the form of a toxin, but that the clinical severity of a malaria infection might be determined by malleable as well as ‘inherent’ properties of the infecting strain. The contention that clinical features of a strain could change was not unique to Horton Hospital; however, the claim that ‘virulence tends to be increased with each successive transmission’ (Macbride, 1924) had been supported (Bunker and Kirby, 1925; Grant, 1923), contradicted (Fiertz, 1926; Yorke and Macfie, 1924) and confounded by mixed results (Lilly, 1925) in the mid-1920s. Furthermore, Wagner-Jauregg not only held the unnatural transmission [via blood inoculation] in malariatherapy responsible for ‘mildness’ and drug sensitivity, but seemed to associate the two traits: ‘inoculated malaria showed itself much more sensitive toward quinine than the natural [mosquito-transmitted] malaria . . . The mildness of this inoculation malaria may be explained thus, that the plasmodia which always reproduce themselves only in the asexual way are less capable of resistance’ (Wagner-Jauregg, 1922). Thus—despite early speculation ‘that a race of malarial parasites that is immune to quinine may be developed [as] fresh water amoebae may be gradually habituated to salt water’. (Leslie, 1910)—in the 1930s parasite response to a therapeutic drug was often taken as a token or reflection of ‘virulence’ as well as a marker of strain identity: ‘The virulence of any strain of parasite may be manifested by the toxic symptoms it produces in the host, and this influences the degree of ease or difficulty with which a clinical cure can be produced . . . but it may also be manifest by the power of the parasite to resist such means and to survive in the host’ (Sinton, 1931). At Horton Hospital, and elsewhere, the importance of distinguishing between clinical-severity and drug-response traits emerged gradually through the 1940s: ‘Failure of a strain of parasite to respond to a given drug is not, however, in itself evidence of virulence; it may be that the
Strain Theory of Malaria: The First 50 Years
9
strain, though resistant to this particular drug, is highly susceptible to others. The virulence or toxicity of a strain can best be seen by its effect on the host in the absence of drug treatment’ (Shute and Maryon, 1954). The search for a definition of virulence and the underlying causes continues, but ‘virulence’ has been used to describe so many different aspects of malaria that a specific operational definition remains elusive. The same historical difficulties and ambiguities dominate the current search, including the mutability of ‘virulence’ and the role of host factors in determining disease presentation. Because it is now considered unethical to allow experimental infections to progress to severe clinical symptoms, and because natural infections are difficult to observe, definitions of virulence have moved away from detailed clinical observations to focus on severe manifestations such as cerebral malaria and life-threatening anaemia. Although, selection studies in a laboratory rodent-malaria model imply that virulence (as indicated by host weight loss) is a heritable trait and may involve parasite replication rate (Mackinnon and Read, 1999), there are still no robust molecular markers of parasite ‘virulence’. Genetic association studies have failed to find any reproducible markers of parasites causing cerebral malaria or severe anaemia (A-Elbasit et al., 2007; Dobano et al., 2007), although some differences in the distribution of var gene alleles have been reported between severe malaria cases and controls (Bull et al., 2005; Kyriacou et al., 2006). Parasites sequestering in placenta and giving rise to pregnancy-associated malaria do share a common phenotype (adherence of infected red blood cells to chondroitin sulphate A via PfEMP-1) and some candidate var genes responsible for this phenotype have been identified (Francis et al., 2007; Salanti et al., 2004; Viebig et al., 2005).
4. REACTION TO ANTI-MALARIAL REMEDIES Uncertainty about drug resistance as a stable strain characteristic was evident in the extensive studies of anti-malarial drugs conducted during World War II: ‘there are two main types of strain—one relatively insusceptible to atebrin [mepacrine] and the other normally susceptible . . . [but] variation in the degree of susceptibility to atebrin amongst the relatively insusceptible strains . . . suggests that there may have been several strains of P. falciparum occurring naturally in the Aitaipe-Wewak area or that the phenomenon of atebrin insusceptibility was not so much an inherent characteristic of the strains as one which had been, or was, in the process of being acquired’ (Fairley, 1946). The initial description of the
10
F. Ellis McKenzie et al.
Chesson strain of P. vivax, also from New Guinea, also implies doubt about relying on drug response alone for strain definition: it ‘reacted differently to certain drugs than did the St. Elizabeth strain . . . This and other characteristics suggest that it might be a strain distinct from some of the American malarias’ (Ehrman et al., 1945). In 1949, using a strain of P. vivax from Hong Kong and a strain of P. falciparum from West Africa, ‘resistance to proguanil was produced . . . by giving small doses of the drug to consecutive patients in a series in whom the strain was being maintained by blood-inoculation’ (Seaton and Adams, 1949; Seaton and Lourie, 1949). Nonetheless, ‘the accidental production in the field of proguanil-resistant strains . . . is thought unlikely’, because high-level resistance had taken 20 months to arise in P. vivax and proguanil had a ‘sterilizing effect’ on P. falciparum gametocytes. At exactly the same time, proguanil resistance was detected in the field, in Malaya: ‘the widespread use of proguanil in the Tampin district for the past two years . . . we believe, is related to the emergence of resistant strains of P. falciparum’ (Edeson and Field, 1950). The authors noted that Rollo et al. (1948), based on avian malaria experiments, had offered the ‘likely explanation . . . that resistant parasites occur spontaneously as rare mutants in a normally sensitive strain; and, as a result of selective survival of these resistant mutants when the parasites are exposed to the drug, a stable resistant strain emerges’. One of the proguanil-resistant strains of P. falciparum from Malaya was later reported to be cross-resistant to pyrimethamine (Robertson et al., 1952), and another not (Davey and Robertson, 1957). The appearance and spread of pyrimethamine resistance in P. falciparum was demonstrated in field experiments in the mid-1950s in East Africa—‘during the course of the treatments the resistant varieties spread, replacing sensitive strains; upon cessation of the treatments resistant varieties regressed and became submerged’ (Clyde and Shute, 1954)—but whether their rise and fall occurred through competitive interactions or through ‘conjugation with other strains . . . could not readily be determined’. Macdonald’s summary of the situation, in 1957, was that ‘the reactions of strains of parasite are unpredictable and in consequence drugs enjoy very different reputations in various parts of the world’ (Macdonald, 1957). Soon after chloroquine resistance was detected in the early 1960s, in Colombia (Moore and Lanier, 1961) and Thailand (Young et al., 1963), a series of studies began to compare and differentiate ‘strains of Plasmodium falciparum resistant not only to chloroquine but to other groups of synthetic anti-malarial drugs’ on the basis of cross-resistance patterns, determining, for instance, that ‘the Malayan I strain differs from the Thailand strain in its susceptibility to pyrimethamine; the Malayan III strain differs in its susceptibility to mepacrine’ (Contacos et al., 1963).
Strain Theory of Malaria: The First 50 Years
11
Though all strains appeared susceptible to quinine, an experimental transfer of a Malayan chloroquine-resistant strain between quininetreated prison volunteers gave ‘strong evidence indicating the emergence of a variant strain characterized by lessened sensitivity to quinine compared to that of the parent strain’ (McNamara et al., 1967), an ‘exceptional’ result cited in support of the hypothesis that ‘new strains may continually be emerging in the endemic area and adapting to the intermittent and irregular drug pressure encountered there’ (Clyde et al., 1969). The World Health Organization’s conclusion, in 1969, was that ‘most of the drugs commonly used to treat malaria have shown variations in effect that are related to the strain of parasite . . . [but] changes in sensitivity may be transient or permanent. . . . The problem is particularly complex if one attempts to distinguish between the inherent differences in the response to drugs of various strains of the same species and the changing pattern of response induced by previous contact with drugs’ (WHO, 1969). In the 1930s, differences in drug response had already suggested that ‘each strain of tertian or of aestivo-autumnal malaria is a problem in therapeutics by itself’ (Hackett, 1937), and that ‘isolation, selection, and other factors known to bring about change in other organisms would act upon malarial parasites . . . [though] malariologists have been slow in acknowledging this possibility’ (Huff, 1938). By the 1980s, the challenge had become that ‘isolates that exhibit multiple drug resistance may, in fact, be mixed infections of parasites exhibiting resistance to each drug separately’ (Rosario, 1981), and the role of drug response in characterizing ‘strains’ had become less clear. The determination of the molecular/genetic basis of resistance to several anti-malarial drugs (Gregson and Plowe, 2005; Wellems and Plowe, 2001) has allowed many of these hypotheses to be tested. For example, a single point mutation (pfcrt K76T) provides high-level chloroquine resistance in P. falciparum infections, but the mutation is found with nine other mutations. Discrete point mutations in a single bi-functional molecule confer resistance to pyrimethamine and to sulphadoxine. These mutations occur with relatively high frequency but impose real fitness costs on the parasite that are observed when selection pressure is relaxed. Following the switch away from sulphadoxine–pyrimethamine as a first-line anti-malarial in East Africa, the regression of resistant varieties first described by Clyde and Shute (1954) and the re-emergence of sensitive strains has been documented (Hastings and Donnelly, 2005), confirming the partial loss of fitness associated with drug-resistance mutations. In Malawi, the loss of chloroquine resistance has been dramatic (Kublin et al., 2003). As postulated by Rollo et al. (1948), mutant parasites are selected and increase in frequency by spreading among
12
F. Ellis McKenzie et al.
hosts in populations exposed to drugs (Anderson and Roper, 2005; Pearce et al., 2005; Mita et al., 2006), but ‘conjugation with other strains’ (Clyde and Shute, 1954), that is, recombination, allows highly- or multiply-drug-resistant strains (carrying multiple point mutations) to emerge and allows spread into many different parasite genetic backgrounds. Thus, in the case of resistance to anti-malarial drug resistance, there appears to be a simple map between genotype and phenotype, and there is a clear basis for defining resistant strains.
5. INFECTIVITY Whatever their other characteristics, parasite strains in the field and in most malariatherapy clinics had to be transmissible from humans to mosquitoes (and back to humans) to persist. Gametocytemia was taken as a marker of transmissibility to mosquitoes, and gametocyte production seemed to vary between strains. At the Horton Hospital, for instance, the Roumanian P. vivax Apostol ‘strain was unsatisfactory because it did not produce a sufficient number of gametocytes’, in contrast to the ‘large number of gametocytes which are usually produced by this [Madagascar] strain’ (Shute, 1937). Dramatic changes in gametocyte production were associated with transfer solely by blood inoculation: ‘malaria strains when inoculated from patient to patient more or less rapidly lose the capacity of producing gametes, and thus of infecting mosquitoes’ (Pijper and Russell, 1926), and, though such ‘gametocyteless’ strains were sometimes propagated (Jeffery, 1951), clinics were cautioned to ‘keep patients undergoing malariatherapy in screened rooms’ because ‘one can never be sure that such loss of power to produce sexual forms is permanent, and strains of malaria are known which retain the ability to infect mosquitoes more or less indefinitely’ (Russell et al., 1946). While studies increasingly indicated that ‘gametocyte density is not a reliable guide to the probably resulting qualitative infection of mosquitoes’ (Boyd, 1942a), it was the most reliable gametocyte producers that became the strains commonly used in most clinics. As for the actual infection of mosquitoes, Anopheles species often showed differing susceptibilities not only to different Plasmodium species (Boyd and Stratman-Thomas, 1934; Boyd et al., 1935), but also to different strains within each species: ‘we failed entirely to infect our English maculopennis with the Indian strains of P. falciparum [but] with the European strains from Rome and Sardinia we succeeded . . . the capacity of each strain to infect a named species of anopheles must be separately worked out’ (James et al., 1932). Further studies with ‘English maculopennis’ at
Strain Theory of Malaria: The First 50 Years
13
Horton Hospital reported successes with P. falciparum strains from Roumania, and failures with strains from East and West Africa (Shute, 1940). European vectors transported to Africa gave a similar result: ‘experiments in Garki, Nigeria, with A. atroparvus from Italy have shown a refractoriness to infection with the local strain of P. falciparum’ (Ramsdale and Coluzzi, 1975). However, ‘A. gambiae, originating from the Nyanza province of Kenya, was able to transmit the Malayan strains of P. falciparum with no more difficulty than African strains’ (Davey and Robertson, 1957). Accordingly, even the broadest of summaries mentioned exceptions: ‘European strains of P. falciparum require a European mosquito to transmit them, and tropical strains tropical mosquitoes, but in the latter case it may be noted that the Indian A. stephensi is a good vector of the far distant West African P. falciparum’ (Garnham, 1966). In the United States, it appeared that A. quadrimaculatus and A. punctipennis ‘vary widely in their susceptibility to different strains of P. falciparum [but] are approximately equally susceptible to both strains of P. vivax’, and, as one result of such findings, the Rockefeller Foundation clinic ‘abandoned the propagation of the Coker strain’ but continued the Holzendorf, Long and Manuel strains of P. falciparum (Boyd and Kitchen, 1936a). As results of vector–parasite transmission tests accumulated, it appeared that ‘local strains of parasites may or may not show a high degree of adaptation to anophelines which are coindigenous to their own faunal regions, and . . . anophelines may or may not show a high degree of susceptibility to exotic strains of parasite’ (Boyd, 1940a), and the practical implications of strain infectivity were seen to extend well beyond malariatherapy clinics. At Horton Hospital, ‘results indicate that persons carrying gametocytes of P. falciparum of tropical origin would be unlikely to cause any outbreak of fresh cases of malaria in this country through the agency of our English maculopennis. On the other hand, our four English anophelines become infected when fed on tropical strains of P. vivax and our A. maculopennis is a very efficient carrier of P. ovale’ (Shute, 1940). In the United States, extensive transmission experiments with indigenous species of Anopheles and ‘exotic’ strains of P. vivax and P. falciparum were undertaken during World War II (e.g. Boyd, 1949) in part because ‘studying foreign malaria imported by returning servicemen, it has been shown that these strains are infective to and can be transmitted by our native malaria vectors’ (Young et al., 1947). A particular concern was that ‘returning soldiers with such infections may be responsible for the establishment in this country of epidemic or endemic foci for imported vivax strains’ (Watson, 1945). Extensive studies of P. vivax infections from the ‘Solomon Islands, New Hebrides islands, New Guinea, Tunisia, Liberia, Trinidad, and the China-Burma-India theater’ with the major malaria vectors of the Western
14
F. Ellis McKenzie et al.
and Eastern United States found that, based on the fraction of fed mosquitoes infected, ‘A.m. freeborni is more susceptible to these foreign malarias than A. quadrimaculatus’ (Young and Burgess, 1948), the same result as with ‘a domestic strain of P. vivax’, St. Elizabeth (Burgess and Young, 1950). P. vivax strains from the Mediterranean and India, however, infected a lower percentage of A. freeborni than A. quadrimaculatus (Young et al., 1949). Similar studies with P. falciparum showed that the Malayan IV strain was much more infective, and the domestic McLendon strain slightly more infective to A. freeborni than to A. quadrimaculatus (Coatney et al., 1971), that A. albimanus was highly susceptible to the Panama strain, but almost refractory to the Thailand strain (Collins et al., 1963), and so forth, thus that ‘the infectivity of isolates of P. falciparum to different anophelines is dependent to some extent on the geographical origin of either the parasite or the mosquito’ (Coatney et al., 1971). The usual inference was that, while ‘there may exist, between particular strains of parasites and their definitive hosts, a very high degree of local adaptation, which under certain conditions may conceivably be a natural barrier to the extension of the range of a given strain of the parasite . . . the human intermediate host is not likely to be a factor in limiting the extension of the range of strains of these parasites’ (Boyd et al., 1938a). In what was apparently the only published study of its kind, the results suggested that ‘anophelines infected on a person concurrently infected with two strains of a single species (P. vivax) may either: (a) become infected with but one of the strains, or (b) may possibly become infected with both of the strains and simultaneously propagate both’ (Boyd et al., 1941). Thus, it was plausible that ‘the happy adjustment of parasite to vector in any area has come about through a weeding-out process, in which ill-adapted strains (and species) have failed to be transferred at proper intervals and have consequently become extinct’ (Hackett, 1937). If there was any correlation between infectivity and gametocytemia, however, it seemed certain that ‘the accidental introduction of a strain of parasite producing large numbers of gametocytes would lead to an increase in incidence’ (Bishop, 1955), and, since ‘gametocyte output is considerably greater in European strains than in Indian or African strains’ of P. falciparum (Shute and Maryon, 1954), for instance, it was not clear what, other than strict local matching of vectors and parasites, might constrain such introductions, or selection for increased gametocyte production. This question led directly to another: ‘are the male gametocytes of one strain of a species of parasite able to fertilize the females of a different strain? . . . inability to do so would be essential if strains are to retain their identity where more than one occurs in a given locality’ (Shute and Maryon, 1954). If cross-fertilization were possible, ‘‘‘How long would an imported strain
Strain Theory of Malaria: The First 50 Years
15
retain its identity when it was introduced into an endemic area?’’ And the answer, we think, would be: ‘‘not very long providing the indigenous anopheles were susceptible to infection’’. . . . [This] helps to explain why it is fairly obvious that there are separate geographical strains, but so very much less obvious that there are separate ‘locality strains’ (Shute and Maryon, 1954). By the 1980s, in vitro cultivation of P. falciparum made it possible to distinguish between a ‘clone’ (‘genetically identical organisms derived from a single cell by asexual division’; Walliker, 1983) and an ‘isolate’ (‘a sample of parasites, not necessarily genetically homogeneous, collected from a naturally infected host on a single occasion’; Walliker, 1983). Studies soon demonstrated that gametocyte production and infectivity might differ between clones grown from a single isolate (Burkot et al., 1984; Graves et al., 1984a), and that ‘for those strains that appear to lose gametocyte formation in culture this is a result of selection operating on a mixed population’ (Bhasin and Trager, 1984). Indeed, inability of cultured asexual forms to produce gametocytes was found to be due to complete and irreversible loss of large segments of genetic material (Day et al., 1993). Host immunity that reduces gametocyte densities and that blocks infection in mosquitoes suggests a role for the host in determining infectivity. High gametocyte production in response to sulphadoxine–pyrimethamine suggests that gametocyte production can respond to some cues within the host, and is not a fixed trait (Barnes and White, 2005). Incompatibility between parasite strains and various species and strains of Anopheles is now believed to reflect an interplay between polymorphic innate immune response genes of the mosquito that promote or retard parasite killing (melanization) and unknown melanization resistance genes of the parasite (Vlachou and Kafatos, 2005), hence current discussions often invoke evolutionary costs (Lambrechts et al., 2005; Sinden et al., 2004). Simultaneous propagation of multiple strains of malaria by a single mosquito has been confirmed by molecular genetic studies (Babiker et al., 1994; Huber et al., 1998; Ranford-Cartwright et al., 1993), but the high frequency of recombination during the sexual phase of the parasite life-cycle (confirming that ‘male gametocytes of one strain’ can indeed ‘fertilize the females of a different strain’) may mean that outcrossing, between strains, is as common as ‘selfing’ and thus, as predicted by Shute and Maryon (1954), that strains rapidly lose their identity in endemic areas. Hence, as with drug response, the role of infectivity in maintaining and characterizing strains began to appear highly complex.
16
F. Ellis McKenzie et al.
6. ANTIGENIC PROPERTIES 6.1. Homologous and heterologous response In 1898, an army doctor observed that ‘the contrast between West Indians and the natives composing the Sierra Leone Frontier Police when serving in the field in the same column was most marked: the former much troubled by fever, the latter having none’, and inferred ‘that the malarial organism of the West Coast is of a different variety to the plasmodium of the West Indies, and that immunity acquired to the first mentioned confers no protection from the second’ (Smith, 1898). In India, ‘there is always a certain amount of malaria of local origin in most towns, but this affects mainly children and new-comers; it is probable that adults sometimes acquire a certain measure of immunity to local infection, but are not immune to the more virulent type found at certain places in the interior of the districts’ (Kenrick, 1910). By the 1920s, it was recognized that ‘immunity developed against one species of Plasmodium does not confer a similar protection against the other species’ (Yorke and Macfie, 1924) and, retrospectively, with respect to strains, that in the ‘intermingling of men coming from malarial regions’ during and soon after World War I, ‘inhabitants more or less resistant to local strains showed little or no immunity to foreign strains and the immigrants had no resistance against the local virus [parasite]’ (League of Nations Health Organisation Malaria Commission, 1934). With such intermingling, and ‘numerous bites from innumerable mosquitoes . . . the chance of a person receiving an infection with one or more strains of parasite of varying degrees of virulence was considerable . . .. [and] with infection by each new strain the sufferer would be liable to a recurrence of clinical manifestations of greater or lesser intensity’ (Sinton, 1931). Thus, it began to appear that ‘persons can easily be immunized against a particular strain . . . but that the resistance breaks down if they are inoculated . . . even with a different strain of the same species’ (Thomson, 1931). Deciphering the nature of this ‘immunity’ or ‘resistance’ required more precise information about how it developed, and how it might affect ‘virulence’, drug response and transmission. Such studies might explain pronounced differences in the initial responses of malariatherapy patients: ‘the patients of our series were not of a homogeneous susceptibility to the strains . . . some clearly possessed a pristine susceptibility, others exhibited evidence of previous autochthonous infection, varying from partial immunity to complete refractoriness, the latter interpreted as an immunity homologous to the strain we employed’ (Boyd, 1942b). Similarly, the results might help to explain differences in responses to re-infections, in that ‘the usual criterion of adequate malariatherapy is the actual number of paroxysms reaching a certain febrile height . . . [so]
Strain Theory of Malaria: The First 50 Years
17
heterologous immunity exists when reinfection of a patient with a strain of P. vivax differing in geographic origin from the original strain results in definite clinical activity sufficient for the completion of a course of malaria therapy’ (Kaplan et al., 1946). Many of the early re-infection studies led to straightforward conclusions: ‘re-inoculation of a patient who has recovered from a P. falciparum attack with the same strain of the parasite will not result in a second clinical attack . . . Re-inoculation of a patient who has recovered from a P. falciparum attack with a strain of the parasite different from that which caused the primary attack, will result in a second clinical attack which may be as severe as the first’ (Boyd et al., 1936). Thus, homologous and heterologous responses were seen to reveal underlying similarities and dissimilarities between strains: a patient ‘exhibits a resistance when reinoculated homologously with the same line of parasites, but if reinoculated with parasites of the same species from a different source and presumably of a different (heterologous) line, he acquires an infection. From this it is inferred that the parasites of the second inoculation represent a different race or strain’ (Brumpt, 1949). Accordingly, crossinoculation became a common means of strain differentiation: ‘the derivation of two lines of parasites from presumably unrelated patients cannot be taken as a basis for their characterization as different strains . . . [especially] when applied to lines of parasites of obviously local indigenous origin. Their antigenic dissimilarity must be proven by the cross inoculation of convalescents’ (Boyd, 1940a). Cross-inoculation studies determined not only that ‘the White, Wilson Dam, and Mayo strains are immunologically distinct from the McCoy strain . . . [and] the Cuban, Mexican, and Panamanian strains are different from the Long strain’ (Boyd, 1940a), for instance, but, by comparing responses of patients from different regions of the United States, that ‘West Florida and contiguous Alabama are indicated as the indigenous habitat of the McCoy strain’ (Boyd and Kitchen, 1936b). Further afield, ‘little heterologous immunity was shown between vivax infections from the South Pacific, China-Burma-India theatres and from the United States. In fact, one infection originating from New Guinea exhibited little immunity to another infection from the same area, suggesting multiplicity of strains in small areas’ (Young et al., 1949). Other studies introduced complications, however. Some patients still developed fever at a fourth and parasitemia at a tenth infection with a homologous strain of P. falciparum (Ciuca et al., 1934). One possible explanation was that ‘immunity to a malarial infection exists when the subject is cured and then challenged to a reinfection with the homologous organism . . . this residual humoral immunity, however, is rapidly lost . . . [with] no evidence of cross immunity with a heterologous strain . . . [so] it is more than apparent that there is little reason to hope for an effective
18
F. Ellis McKenzie et al.
vaccine for malaria’ (Yount and Coggeshall, 1949). However, if this ‘rapid loss’ existed, the rate of loss was uncertain—for example, ‘homologous immunity to superinfection persists for as long as a year and may last seven years’ (Boyd, 1949)—and might differ with different methods of parasite transfer, for example, as ‘a temporary immunity to reinfection by the homologous organism, lasting up to two months with trophozoite inoculation and up to 13½ months with sporozoite inoculation’ (Schwartz et al., 1950). Furthermore, it began to appear that ‘convalescents from artificially induced falciparum infections usually exhibit a distinct clinical tolerance when artificially reinoculated with a heterologous strain of this parasite, manifested by a shortened period of clinical activity . . . [thus] the immunity developed during convalescence from a falciparum infection has an appreciable heterologous value’ (Boyd and Kitchen, 1945). Thus, residents of malaria-endemic countries might experience within a ‘period of time a series of episodes successively involving different strains. With a progressive series of inoculations, while the original clinical reaction might be characteristic of complete susceptibility, later episodes would exhibit characteristics of heterologous-strain immunity until the individual had acquired experience with all of the locally prevalent strains. Any inoculation subsequent to this period would result in a homologous-strain reaction’ (Kitchen, 1949). Another set of difficulties arose with ‘premunition’ (Sergent et al., 1924, 1925), the doctrine that ‘resistance to a malaria infection is dependent upon an existing infection, either active or subclinical’ (Yount and Coggeshall, 1949): that is, some responses to seeming re-infection might actually be responses to superinfection. One implication of latency or sub-detectable parasitemia (see below) was that, at least with respect to homologous immunity, ‘we never know when any one is cured. Our only proof is a renewed susceptibility to reinfection by the same strain of parasite’ (Hackett, 1937). Moreover, conclusions about homologous and heterologous responses with one species of Plasmodium might not be directly applicable to other species: ‘Superinfection with the same strain of P. vivax rarely, if ever, takes place [whereas] with P. falciparum, superinfection is apparently possible, but the course of the second infection is very greatly modified’ (Earle et al., 1939) and ‘there is less indication that acquired immunity to P. falciparum has heterologous value than in the case of acquired immunity to P. vivax’ (Boyd et al., 1936). Perhaps ‘the immunity acquired to the three species is of a different order in each case . . . and immunity to P. falciparum is the least complete. The alternative explanation is that there exists a greater variety of strains of P. falciparum, and that, while immunity to one strain is reaching a high titre, infection with a new strain, or with an older strain to which immunity has already largely disappeared, intervenes . . . [though] such a multiplicity of strains in one area seems improbable’ (Wilson, 1936).
Strain Theory of Malaria: The First 50 Years
19
Wilson (1936) would likely be astounded by the level of genetic diversity that is now known to occur within P. falciparum. Diversity is linked to levels of malaria endemicity, such that parasites from unrelated patients in Africa or New Guinea are likely to represent different strains whereas parasites from unrelated patients in South America have a higher chance of belonging to the same strain. Species-specific and strain-specific immunity is believed (although far from being formally demonstrated) to be due to (allelic) polymorphism of genes encoding the major surface proteins of each stage of the parasite, such that antibodies raised to one form of the protein bind less efficiently to heterologous forms present in other parasite species or strains. That allelic diversity is greater among P. falciparum compared to other human malarias, predicted by Wilson (1936), is being confirmed by molecular genotyping but only partially explains the lack of homologous immunity. The ability of a single strain of P. falciparum to cause numerous bouts of fever or parasitemia in a single patient (Ciuca et al., 1934) and the shorter duration of immunity following trophozoite inoculation compared with sporozoite inoculation (Schwartz et al., 1950), are likely explained by clonal antigenic variation (Dzikowski et al., 2006; Kraemer and Smith, 2006). Sequential expression among genetically identical blood stage parasites of heterologous var genes that encode for PfEMP1 expressed on the surface of infected red blood cells may enable a single parasite strain to escape effects of the developing immune response; strains with inherently high switching rates (allowing rapid sequential expression of novel antigens) would be expected to be able to cause repeated infections in the same patient. The gradual reduction in severity of disease with successive reinfections by the same strain might be explained by increasing immunity to non-variant antigens or gradual exhaustion of the repertoire of clonally variant antigens.
6.2. Clinical and parasitological response Given this accumulation of studies suggesting that ‘homologous’ protection was not necessarily immediate, absolute or permanent, that ‘heterologous’ responses might include partial protection, and so forth, more nuanced interpretations began to emerge, for example, that ‘a considerable degree of resistance to reinfection and superinfection with a homologous strain of malaria parasite, may be acquired after a single infection with such a plasmodium, and that, in some instances at least, this resistance may be increased by successive inoculations with the same strain of parasite’ (Sinton, 1940). Homologous resistance was increasingly
20
F. Ellis McKenzie et al.
described in terms of clinical severity and duration: ‘If a patient has an acute attack with a certain strain prevalent in one area and recovers from it, it is found that inoculation again with that strain shows an immunity. The attack is not so acute and he recovers more rapidly. Now if he is inoculated with a different strain brought from an area several hundred miles away, he has another acute attack similar to the first one. In other words he has some homologous immunity but no heterologous immunity’ (Bispham, 1944). By the late 1940s, it appeared that cross-inoculation might not always distinguish between heterologous strains, at least not unambiguously, without careful elaboration of the terms of reference: ‘reinoculation . . . after the chronic infection is no longer microscopically patent, with the same species and strain involved in the previous infection, is not likely to result in more than a transitory subclinical parasitemia. . . . (Boyd, 1949) Successive reinfections . . . with the same strain . . . result in progressively milder infections than the initial infection, i.e. they progressively enhance the homologous immunity . . . [and] a number of races or strains . . . may be immunologically similar. Others may not only vary in virulence and respond differently to treatment, but also fail to protect against another strain . . . An immunity may exist between strains of the same species although to a less extent than against the homologous strain . . . partial heterologous immunity between some strains and not between other strains in man has been substantiated’. One critical refinement in the terms of reference built on the observation that there were two major kinds of effects of this ‘immunity’, providing ‘some protective mechanism against both the multiplication of the parasites and their pathogenic effects. The rate of development, the efficacy, and the duration of this ‘‘immunity’’ appear to vary with (a) the species or strain of Plasmodium responsible for the infection and (b) the degree of resistance or susceptibility possessed at the time by the host’ (Sinton, 1939). There seemed to be an incomplete correspondence between these effects on symptoms and parasitemia in re-infection: ‘upon recovery from an attack of malaria . . . the patient possesses a very potent homologous immunity to that strain . . . [which] manifests itself by two characteristics: (a) the acquirement of a tolerance to densities of the parasite that in a susceptible person would produce a clinical reaction, and (b) the acquirement by the body of an ability to destroy and remove the parasites. As immunity becomes established the former characteristic is first acquired; the latter develops more slowly’ (Boyd et al., 1938b). Thus, responses to homologous and heterologous re-infection were seen as more sharply distinct in clinical than in parasitological terms, in both magnitude and duration: with ‘the reaction of ‘‘heterologous (strain) immunity’’ . . . the latent state . . . is attained as a rule in 3 to 7 days’ after the peak parasitemia, and the attacks ‘are relatively brief, usually
Strain Theory of Malaria: The First 50 Years
21
not exceeding 14–16 days in duration of clinical activity’; with ‘the ‘‘homologous (strain) immunity’’ reaction . . . the infection provokes no febrile reaction . . . [and] parasitemia is usually brief’ (Kitchen 1949). If patients could acquire ‘an immunity to fever but not to parasites’ (James and Ciuca, 1938), there might be ‘two independent kinds of immunity, one to the parasite itself, leading to its corporal destruction, and one neutralizing the pathological effects of its growth and activity upon the host’ (Hackett, 1937). In the 1930s, as the more complicated results from re-infection studies were beginning to suggest new interpretations, some authors began to use the term ‘tolerance’ specifically to refer to reduced or absent clinical response: ‘inhabitants of an area may become tolerant to the local strain of parasite, yet at the same time be susceptible to the pathogenic effects of strains present in other areas’ (Sinton, 1931). And, while the details would require investigation, ‘that the time taken to produce this tolerance is prolonged may possibly be explained if it might be assumed that there are numerous strains of parasites of malaria specifically differing in their antigenic properties’ (Thomson, 1933). Not surprisingly, premunition and other emerging issues often obscured the distinction: ‘during an attack of malaria a person acquires a tolerance to the parasite of the strain harbored, which renders him refractory to reinoculation with the same strain’ (Boyd and Stratman-Thomas, 1933a), but, while ‘a patient with a latent benign tertian infection [P. vivax] does not possess a heterologous tolerance to other strains’ (Boyd and Stratman-Thomas, 1933b), ‘following superinfection by a heterologous strain . . . we have found the attack caused by the superinfection to be less severe, indicating the actual existence of some degree of heterologous tolerance’ (Boyd and Stratman-Thomas, 1933c). However, as distinctions between clinical and parasitological aspects of response were further elaborated—for example, ‘heterologous immunity to malaria is rarely strong enough to prevent infection but it may so modify a second attack that it closely resembles a relapse’ (Russell et al., 1946)—some authors began to use the term ‘immunity’ strictly to refer to anti-parasite response, and, by the late 1940s, studies of homologous and heterologous re-infections were investigating the two aspects of response in parallel, in corresponding terms: given ‘less reaction by the host to a given bulk of infection—tolerance . . . [and] an increased ability of the host to limit the bulk of infection developed— immunity . . . [we found] that the development of tolerance preceded the development of immunity and that immunity was strain specific . . . [while] tolerance was not strictly strain specific’ (Blackburn, 1948). With respect to the ‘antigenic properties’ of parasite strains reflected in these responses, ‘the only explanation would seem to be that each of these strains contains immunological elements in common with each of the
22
F. Ellis McKenzie et al.
others’ (Hackett, 1937) and ‘that the antigenic constitution of the malaria parasites is analogous to that of some pathogenic bacteria . . . [i.e.] in all specimens of a particular strain of a malarial parasite there are parasites of two types, one containing specific antigen, the other group antigens . . . [and] that the antigens may be subdivided into 1o group antigens which are common to all members of the group, 2o group antigens which are common only to certain members of the group, 3o specific antigens peculiar to each type’ (James and Ciuca, 1938). Thus, in an attempt to integrate the evidence, the observation that ‘differences between the results of homologous and heterologous reinoculations lie mainly in the number of febrile infections produced . . . suggests that possibly the common antigenic factor in these strains may be related more to the antiparasitic element than to the anti-toxic one . . . [while the] more rapid decline of the percentage of cases showing febrile reactions after successive reinoculations than of the recorded parasitic infections . . . might suggest that the antitoxic immunity factor was developed more rapidly than the anti-parasitic one’ (Sinton, 1940). Then, ‘in the condition of premunition, while both defensive factors are in operation, the antitoxic element is probably the more efficient . . . [but] the duration of the efficacy of the anti-parasitic element persists for a longer time than does the anti-toxic one’ (Sinton, 1940). As with ‘virulence’, and drug response, several authors asked whether antigenic features of a strain might change with time. Most concluded that, since observations ‘do not suggest that extensive sexual reproduction has altered the antigenic composition of one strain . . . the antigenic characteristics of the parasites upon which immunological differentiation of strains is based, are evidently firmly fixed and retained through an indefinite number of passages through the definitive and intermediate hosts’ (Boyd, 1940a). Huff, however, argued that with ‘parasites which are distinguishable only on immunological grounds . . . whether they constitute separate races, possibly incapable of cross-breeding or whether they are simply manifestations of variations within a variety or species due to sexual reproduction can only be guessed at the present time . . . Individuals in endemic areas probably build up an immunity to one strain of malaria only to suffer from another immunologically different strain. And since there is the possibility that strains of malaria may change genetically in immunological characteristics, man in these areas is being subjected to reinfection by a multiplicity of genetic stocks of parasites. If, on the other hand, these strains of malaria have evolved far enough that they no longer cross breed it ought to be possible for individuals living in a given region to develop eventually an immunity to superinfection to all of the strains, providing the number of these strains is not unreasonably large’ (Huff, 1938).
Strain Theory of Malaria: The First 50 Years
23
The molecular basis for the two types of anti-malarial immunity— immunity against the parasite itself and tolerance to the pathology it causes—is still somewhat unclear, although the distinction is still widely appreciated (Schofield and Grau, 2005) and the former is still regarded to be somewhat more strain-specific but longer-lasting than the latter. One theory, that tolerance is provided by short-lived, T-cellindependent antibodies to non-polymorphic glycophospholipid ‘toxins’, is consistent with these observations as is the notion that immunity to parasites is provided by longer-lived and boostable antibodies to polymorphic protein antigens. Of these, conserved or relatively conserved proteins such as the circumsporozoite protein would fit the James and Ciuca (1938) definition of 1o group antigens, proteins such as merozoite surface protein 1 (MSP-1) and MSP-2 which exist as a small number of allelic families defined by conserved family-specific sequences would meet the definition of 2o group antigens, whilst the individual variants within the MSP-1 and MSP-2 families—or indeed the clonally variant PfEMP-1 proteins—could be classified as 3o group antigens. Whilst diversity within the MSP-1 and MSP-2 families seems to have arisen by a combination of both point mutation and intragenic recombination (Ferreira and Hartl, 2007; Ferreira et al., 2003), thus refuting the notion of Boyd (1940a) that antigenic characteristics are firmly fixed, it does still seem to be the case that the antigenic properties of a strain do appear to be rather more durable than—for example—drug sensitivity, suggesting perhaps that the selective forces imposed by the immune system are rather less than those imposed by chemotherapy. Regardless of the mechanism, mathematical models of epidemiological data support the view that immunity to clinical disease [tolerance] develops earlier in life than does anti-parasite immunity (Filipe et al., 2007) and that immunity to severe malaria is acquired quite rapidly (Gupta et al., 1999).
6.3. Superinfection Thus, again, there arose the question of distinguishing between re-infection and superinfection, now complicated by questions about the stability of antigens that might give rise to immunity. Using the available clinical, parasitological and immunological methods, it would be ‘necessary to know whether the presence of one parasite influences the course of infection with the others. Further, we must know if superinfection can take place and if so, at what stage of the previous infection this is possible. Finally we will need to know how many strains of each parasite there are . . .
24
F. Ellis McKenzie et al.
In nature either the number of strains or the times that they will infect in the presence of other parasites must be limited. Otherwise in endemic areas adults would have difficulty in developing immunity to all strains that existed’ (Earle et al., 1939). Sequential, single-strain infections might be just one among many sets of possibilities to be considered in interpreting responses, whatever the number of strains present. Re-infections following concurrent infections with multiple strains suggested that ‘the homologous immunity to either of two strains of P. vivax which follows simultaneous inoculation with the two strains is not so effective as that acquired after inoculation by a single strain. The immunity is characterized by the heterologous rather than the homologous properties . . . [i.e.] the simultaneous presence of two strains delays the development of an adequate homologous immunity to either’. (Boyd et al., 1938b) In World War II, it was ‘recognized that Pacific vivax malaria represents a complex situation in that each patient frequently harbors multiple strains which are not synchronized . . . the clinical features of this group of cases should be interpreted as the characteristics of multiple strain infections and possibly superposed infections of homologous strains . . . the complexity of the presence of multiple strains is such that the features of this study are only significant in a clinical light, and cannot be applied to the duration or study of immunity of a single strain infection’ (Hill and Amatuzio, 1949). The concept of multiple concurrent, superinfecting, interacting strains had implications for epidemiology and control as well as for clinical and immunological understanding. For an individual, the presence of ‘an unknown number of strains without any cross-immunity to speak of among them [implied that] only by chance is he infected twice in succession with the same kind of parasite, and his individual malaria season draws to a close only when he has solidified anew his resistance to the principal strains which are in the air around him every night. We have no idea how many strains of each parasite there are in any one locality . . . [but] any reduction in anopheline density begins by removing layer after layer of these superimposed infections before it cuts down the amount of malaria, or number of infected persons’ (Hackett, 1937). Because ‘each new strain, like a new species, finds the host defenceless and initiates a train of events culminating in an acute attack, and a period of gametocyte production . . . chronic malaria, then, is due to overlapping infections of different species and heterologous strains of plasmodia. Mixed infections must be the rule and not the exception in localities with even a moderate transmission rate . . . [and thus] children can not grow up in a malarious locality of even moderate endemicity without acquiring a representative assortment of all the species and many of the strains of plasmodia with which the local anopheles are infected . . . The transmission rate, thus determined, creates a corresponding tolerance which is made up of highly
Strain Theory of Malaria: The First 50 Years
25
specific reactions to the numerous and immunologically independent strains and species of the parasite. The picture is one of growing multiplicity of mixed infections, resulting in chronic malaria and the attendant phenomenon of mutual parasite antagonisms. The mass effect, however, of group immunity at high levels of intensity is one of powerful protection of the older age groups and the shifting of the struggle to childhood or even infancy’ (Hackett, 1941). Rates at which protective responses were acquired with age were uncertain, however, even in hyperendemic regions, as were the rates at which those responses were lost (if they were): ‘attacks are less and less frequent until the age of twelve, when, if the child survives, an immunity is produced which lasts the entire life of the native if he does not leave the locality in which he was reared. Leaving the area per se does not lessen the child’s immunity, but he would be subject to attacks of other strains of the parasite’ (Bispham, 1944) or, ‘because of the variety of strains and species, a more or less stable immunity may develop somewhere between 20 and 30 years’ (Boyd, 1949). However, given that ‘the greater the number of bites the greater the chance of the introduction of multiple strains of parasite, [which] might account for the presence of both the more virulent and the more cureresistant features recorded’ (Sinton, 1931), and that many protective responses appeared to be strain-specific, strain introductions were invoked to ‘explain the numerous observations of sudden, severe, and acute outbreaks of malaria, which can be called ‘‘malaria epidemics,’’ in endemic districts’ (Nocht and Mayer, 1937). In more general terms, the diversity of strains, clinical and parasitological effects, and frequencies of superinfection and reinfection were all seen to vary with the prevalence of infection and intensity of transmission: ‘a single or at most very few strains of parasites would be prevalent where endemicity is at a low level. The acquired immunity will render further clinical activity improbable in the event of homologous re-inoculation . . . At higher levels of endemicity the number of strains prevalent may be expected to be greater [and] through repeated reinoculations in the course of time with other strains, the individual’s immunity will become polyvalent to all of the species and strains of parasites which are locally prevalent . . . A person may be reinoculated one or more times with the same or a different species or strain of parasite, after an interval which should be expected to vary with the endemic or epidemic level prevailing . . . [if] with the same species and strain of parasite which caused a previous infection, it may result in a superinfection if homologous immunity is as yet incomplete. If effected with a different species and strain, it is distinguished as a reinfection . . . Where endemicity is at low or moderate levels, it is likely that reinoculations would be infrequent within a short interval, so that the first infection would have become latent, or even have been
26
F. Ellis McKenzie et al.
eradicated, before reinoculation. . . . Under conditions of high or hyperendemicity, or during epidemics, reinoculation may be experienced at shorter intervals . . . [which] produces a protracted or continuous infection and clinical activity’ (Boyd, 1949). While the number of strains at a given site would be expected to correlate with basic entomologic and epidemiologic variables, the same strain might be present at different sites: ‘The immunity of persons living in regions where malaria is widespread is due to the fact that ever since birth they have been inoculated by an unpredictable number of strains of the different plasmodia, some of these strains having the same antigenic properties as those which they might contract in other endemic regions, even far removed from their original surroundings’ (Brumpt, 1949). Cross-inoculation experiments in Liberia produced equivocal results: ‘we are unable therefore, to support the hypothesis that a number of immunologically differing strains exist in relatively small areas and its commonly held practical consequence that a semi-immune who travels relatively short distances in Africa is particularly liable to a ‘‘foreign’’ malaria infection with symptoms’ (Bray et al., 1962). Resistance to a seemingly novel strain might reflect some common antigenic property, or, perhaps, some innate or acquired host factors not yet identified: ‘whether the African volunteers were infected with an African strain or a Malayan strain there was no obvious difference in the symptoms or course of the malaria . . . [our results] all suggest that if there has been intensive infection throughout childhood a very definite immunity is built up which extends to a strain of P. falciparum from several hundred miles away or even from another continent . . .. the different response to infection appeared to be much more dependent on the origin, and on the quantitative degree of immunity of the patient, than on the origin of the strains’ (Davey and Robertson, 1957). Summarizing the evidence in the mid-1950s, Macdonald agreed that ‘in nature there are probably a number of strains and species of human parasite transmitted at any one moment’, but, at least in children in hyperendemic regions, ‘the occurrence of one infection with falciparum makes no material alteration to the probability of another during its course, and that fresh infections during this time are marked by a fresh onset of parasitemia materially unaffected by the previous one’ (Macdonald, 1957). He noted that responses differed between Plasmodium species, and that responses to different strains were difficult to distinguish in nature: ‘infection with P. vivax confers a homologous immunity preventing superinfection or subsequent reinfection with parasites of the same strain though not necessarily with other strains of the same species. A small degree of heterologous immunity to other strains is produced and repeated infection with several strains may ultimately produce a firm heterologous immunity to all. In the case of P. falciparum the general
Strain Theory of Malaria: The First 50 Years
27
picture is somewhat similar, but the degree of immunity conferred is considerably less. The clear-cut effects of infection with particular strains cannot be observed in the field, but only the general effect of inoculation with parasites of two or more species and of an unknown number of strains of each. In such circumstances there can be little doubt that superinfection, that is the imposition of a second infection on a first before it has died out, commonly occurs’ (Macdonald, 1957). Soon after, Bruce-Chwatt wrote that ‘it is surprising, however, that apparent superinfection should be so easy in African adults who, having been exposed to malaria since childhood could presumably have a considerable ‘‘multi-strain’’ resistance to infection . . . [so] until we have the immunological means of distinguishing between two strains of the same species of Plasmodium the explanation of such happenings and their follow-up will be difficult’ (Bruce-Chwatt, 1963). Hence, even amidst the technical advances of the 1960s, ‘it is unwise to say more than that residual immunity and not premonition [premunition] follows recovery from some infections, that the immunity is strain specific, and lasts for at least 3 years’ (Garnham, 1966). The challenge remained that ‘immunological differences exist . . . among geographically isolated strains of the same plasmodial species; but the extensive available evidence for this is based on the degree of cross-immunity among species and strains rather than on clear-cut antigenic definition’ (Zuckerman, 1964), and thus it was ‘further hoped that by serological means it will be possible to say which malaria parasites an individual has been infected with in the past, which ones he still carries, and to which strains or species he is immune’ (Voller, 1964). Though experimental infections with rodent malarias indicate withinhost competition between parasite clones (Mackinnon et al., 2002), such that one drops to sub-detectable levels when the other is introduced, no data are available on the quality of the immune response induced in such situations. Longitudinal studies in humans suggest that competition and immunity mainly affect parasite density, not the time to clearance (Sama et al., 2006); apparent loss of particular genotypes is common, as densities drop below detection limits. Molecular genotyping has confirmed the expectations that ‘mixed infections must be the rule . . . in localities with even moderate transmission rate’ (Babiker et al., 2000; Hackett, 1941; Peyerl-Hoffmann et al., 2001; Sallenave-Sales et al., 2000), that superinfection occurs more commonly in areas of higher transmission (Arnot, 1998), that novel genotypes infecting otherwise immune children give rise to symptomatic infections (Contamin et al., 1996), and that introductions of new phenotypes can cause epidemics of clinical malaria (Arez et al., 1999; Laserson et al., 1999). There is less evidence from these studies of small-scale variations in genotype
28
F. Ellis McKenzie et al.
frequencies that might lead to exposure to new strains after travelling ‘relatively short distances in Africa’ (Bray et al., 1962), but such variations have been observed in remote South American villages (Machado et al., 2004), suggesting that human population movements can create mosaics of local parasite diversity at various spatial scales. Surprisingly, despite the characterization of vast numbers of antigenic variants, the use of serological methods to determine patterns of prior exposure to different parasite strains is still in its infancy (Gray et al., 2007).
7. LATENCY AND RELAPSE With the development of malariatherapy, blood samples were sometimes taken before the onset of fever as well as after, and from these it appeared that parasites could usually be detected before the first fever occurred. Clinicians and researchers remained more focused on the incubation period (the interval between parasite inoculation and first fever in the host) than the pre-patent period (the interval between parasite inoculation and first detected parasitemia in the host blood), but noted considerable variation in the duration of each, even more with P. vivax than P. falciparum. Because the time of inoculation was known, it seemed likely that during pre-patency the parasites were not only present but multiplying at sub-detectable levels, and that the lag to the first fever involved a ‘pyrogenic threshold’. Furthermore, these intervals and levels might reflect properties of strains, arising, for instance, because ‘the number of merozoites formed at schizogony varies with these different strains of P. vivax’—referring to an average 17–18 observed with Madagascar, 16 with McCoy, 12–13 with Dutch (Boyd, 1941). Instances in which the initial latency was protracted for months were particularly striking: thus, based on ‘those very common benign tertian [P. vivax] infections of northern Europe in which even the primary attack is suppressed . . . this primary latency or prolonged incubation seems a character which belongs particularly to certain strains’ (Hackett, 1937). The pronounced differences between P. vivax strains in average incubation period—for example, 13.5 days with Madagascar and McCoy, 16.5 days with St. Elizabeth, 21 days with Dutch, 282 days with Roumanian—typically reflected differing proportions of patients with this protracted initial latency (Boyd, 1941, 1949; Kitchen, 1949). In contrast, there seemed to be relatively little variation between P. falciparum strains in average incubation period—for example, 12 days with Roman, Sardinian and West African strains, 13 days with Coker, Costa and Long strains—despite sometimes wide ranges within strains, for example, 6–25 days with Coker (Boyd, 1941; Coatney and Young, 1941; Kitchen, 1949).
Strain Theory of Malaria: The First 50 Years
29
There were complications in interpreting strain differences in incubation period, however, for instance that ‘many more sporozoites of P. vivax than of P. falciparum are required to ensure the onset of the malarial attack within the usual incubation period’ (James and Ciuca, 1938), and that ‘the duration of the incubation period tends to vary inversely with the dose of sporozoites received [and] the duration of the clinical attack appears to vary directly with the dose of sporozoites’ (Boyd, 1940b). At Horton Hospital, it appeared that though ‘latency in BT [P. vivax] malaria . . . occurs more frequently with temperate region strains than it does with tropical strains . . . true latency occurs only when the sporozoites injected are too few to set up an immediate attack’; however, with patients seemingly recovered from Madagascar-strain infections, ‘if they were infected with a different strain of the same species, some developed fever and parasites but with a protracted incubation period, usually of several months duration’ (Shute, 1946). Initial latency might be influenced not only by dose, and parasite strain, but individual host response: ‘the time which elapses between the date of being bitten by an infected mosquito and the date when the earliest clinical symptoms are felt by the patient varies with the dose of sporozoites injected, the virulence of the particular strain of parasite, and the different factors which tend to lessen or to increase the patient’s resisting power’ (James, 1920). Thus it might be, in contrast to those who ‘suggest that there is not a constant pyrogenic threshold for all strains . . . that varying susceptibility of patients rather than varying virulence of different strains of parasites is chiefly responsible for the variations in density noted at the onset’ (Boyd, 1941), and thereby the variations in incubation and pre-patent periods. When re-infection or superinfection could be excluded as possibilities, it became difficult to interpret the renewed fevers and parasitemia that sometimes followed an initial attack after latent periods of varying lengths. Several types of renewed activity could be differentiated in accord with their timing: ‘for our own purposes, and quite arbitrarily, we distinguish between the returns of fever and parasites which may follow recovery from a primary attack, thus: Recrudescence . . . Relapse . . . Recurrence’ (James, 1931), here with the first category applied to renewed activity within 8 weeks after recovery, the second 8–24 weeks after, the third more than 24 weeks after. It gradually emerged that the last of these categories—now considered ‘relapse’—might occur with P. vivax, but not P. falciparum: ‘neither latency nor long-term relapses occur in malignant tertian malaria . . . based on a study of several geographical strains of both tropical and sub-tropical origin’ (Shute, 1946). Frequencies of relapse appeared to vary dramatically between P. vivax strains, for example, ‘we have observed renewed clinical activity after cessation of the primary attack approximately ten times more often in patients inoculated with the McCoy strain than in those inoculated with
30
F. Ellis McKenzie et al.
other strains’ (Boyd, 1940a). The frequency of protracted initial latency with the Dutch strain was five-fold higher than with the Madagascar strain, and the frequency of relapses eight-fold lower (James and Ciuca, 1938). A geographical pattern emerged: ‘strains of P. vivax that originate in tropical areas characteristically produce clinical attacks of malaria at frequent intervals throughout the year . . . [but] Korean vivax malaria exhibits a bimodal pattern of clinical activity and a period of long-term latency similar to other strains of P. vivax originating in temperate climates . . . similar in all major respects to the pattern of the St. Elizabeth strain’ (Hankey et al., 1953). Thus ‘the likelihood of secondary attacks may vary with the strain . . . [and] New World strains have exhibited a decidedly less frequent tendency to become reactivated after long intervals of quiescence than have the Old World strains’ (Boyd, 1941), a difference that may have arisen because ‘in areas where transmission by mosquitoes can occur during only a short period of each year and where the infective inoculum per patient is often small, strains of P. vivax which can hibernate for many months within the human host would be much more likely to survive until the next transmission season than would strains which relapse promptly’ (Coatney et al., 1950a). Frequencies of relapse were important in evaluating potential introductions of ‘exotic’ strains of P. vivax, so studies during World War II compared relapse rates and infectivity ‘by origin of strains’—sometimes by whether the infections had most likely been acquired in islands ‘A, B or C’—reporting, for instance, that patients were most infectious during their 6th–15th relapse, and when asymptomatic (Watson, 1945). One large US clinic reported relapses in 80% of P. vivax cases returning from the Pacific, with an average latent period between recurrences of 4.2 months, compared with relapses in 30% of cases from the Mediterranean and 2% with the US St. Elizabeth strain (Schwartz et al., 1950). An extensive series of studies conducted on P. vivax from returning soldiers reported that the average pre-patent and incubation periods were shorter, and parasitemia at first fever was higher, in Mediterranean than Pacific strains, and that ‘Mediterranean malarias had a higher gametocyte density and a higher parasite level at clinical relapse than the Pacific malarias. However, the Pacific malarias showed a higher proportion of patients relapsing after treatment and a greater relative prevalence of parasitemia . . . [and therefore] the Pacific malarias might be considered as being more virulent in man than the Mediterranean malarias’ (Young et al., 1949). Thus, as with initial latency, it had become ‘apparent that the pattern of relapse in P. vivax infections is determined by the strain of parasite, as well as by immunity, chemotherapy, and the size of the infective inoculum’ (Coatney et al., 1950a), and, again, that several of these effects might be intertwined, for instance through ‘a stage of the parasite living in fixed tissue cells which intervenes between the sporozoite and the trophozoite,
Strain Theory of Malaria: The First 50 Years
31
the development of which may either be retarded, or inhibited or narcotized by a drug’ (Boyd, 1941). It appeared that ‘the frequency, duration and severity of ‘‘relapses’’ (including recrudescences) depend on the amount of ‘‘tolerance’’ or ‘‘immunity’’ which the patient may have acquired during the primary attack. Patients who are treated with quinine very early in the primary attack acquire little or no tolerance’ (James, 1931). One set of studies showed that ‘after a single attack of Chesson strain vivax malaria, cured with pentaquine–quinine, there was no appreciable homologous strain immunity. After four to seven attacks, followed by chemotherapeutic cure, homologous strain immunity was present, but was inadequate to prevent an abortive attack following sporozoite inoculation . . . [the data] strongly suggest that a large proportion of vivax infections resulting from small numbers of sporozoites will display short courses and few relapses and that they may subside under non-curative therapy without the development of homologous strain immunity’ (Coatney et al., 1950b). Another study with the Chesson strain confirmed that ‘immunological phenomena have a profound effect on the intervals between attacks and on the frequency of relapses . . . the immune cases reinoculated with the homologous parasite . . . almost invariably experienced some symptoms early . . . and also almost invariably experienced one parasite recurrence, always asymptomatic . . . relapses occurred later and less frequently after treatment of extended clinical attacks than was the case in those attacks terminated prior to extensive clinical malaria’ (Jeffery, 1956). Furthermore, ‘an important factor in relapsing malaria may be multiple mosquito bites involving perhaps a diversity of strains of the parasite . . . [and] the greater the infection dosage the greater the liability to relapse and to do so for a longer period’ (Russell et al., 1946). If so, perhaps, ‘every second, third or fourth infected Anopheles mosquito bite which is prevented means the avoidance of one, two or three relapses later on’ (Horing, 1947), with different strains. An attempt to integrate the evidence hypothesized that ‘when pre-erythrocytic parasites are discharged from an exo-erythrocytic depot, there may be several strains amongst them but one strain predominates . . . [and] immunity develops against the predominating strain, but, since there will be some overlapping of the antigenic patterns, there may also be some, possibly transient, cross-immunity against other strains. When later there is another discharge of mixed strains from the depot, the immunity that has developed against the first strain will prevent the development of the erythrocytic schizogony cycle by that strain, and another strain will predominate; this goes on until immunity has developed against all the strains present . . . The concept of multiple strains being in some way responsible for relapses is gaining ground [though] the therapeutic strains hitherto in use were probably mixed (multiple) strains; that it would be possible to initiate an infection from a single sporozoite seems improbable, but it should be possible by a
32
F. Ellis McKenzie et al.
process of dilution to induce a single-strain infection which would, if the above hypothesis is correct, be a nonrelapsing one’ (Napier, 1947). During and just after World War II, studies began using the clinical, parasitological and immunological methods available to investigate these hypotheses about connections between relapses and concurrent multiple strain infections, initially by comparing cross-inoculation responses to parasites (A, B, C, D) taken from different relapses in the same soldier. Given that ‘different strains vary characteristically in the frequency with which secondary episodes occur’ and that the effects of ‘cross inoculations with a heterologous strain’ were recognizable, ‘the results suggest that ‘‘Strains’’ C and D are closely similar antigenically, if not identical, but that ‘‘Strain’’ A substantially differs from ‘‘Strains’’ C and D’ (Boyd and Kitchen, 1948). Another study, having noted that ‘successive attacks of vivax malaria in an individual exposed in a malarious area need not necessarily be relapses caused by the same strain of parasite [and that] it is theoretically possible for the fixed-tissue parasites of two or more strains of Plasmodium vivax to coexist in a host and produce malarial attacks independently’, took advantage of ‘the characteristically different relapse patterns of the Chesson and St. Elizabeth strains of P. vivax . . . [i.e.] the Chesson strain usually produces an infection with several closely-spaced attacks, whereas the St. Elizabeth strain infection exhibits an early primary attack, several months of latency and a series of late attacks in close succession . . . Volunteers infected with the Chesson and St. Elizabeth strains of P. vivax displayed a relapse pattern consistent with a combination of the relapse patterns which were exhibited by the two strains when present separately’ (Cooper et al., 1950). Thus, with relapse as with re-infection and superinfection, it seemed reasonable that ‘if several strains were present more attacks of malaria would be required before tolerance and immunity to all strains would develop’ (Cooper et al., 1950), and, accordingly, ‘when many strains are superimposed, in individuals of differing ages and states of nutrition, who are subject to repeated reinfections as occurs in malaria in its natural habitat, there is little wonder that the pattern of relapsing vivax malaria can appear to be hopelessly complex’ (Coatney et al., 1950b). It is disconcerting to realize how recently the hypnozoite stage of P. vivax was discovered (see below) and how little progress has been made since then in understanding how hypnozoite formation and reactivation is controlled; the inability to culture P. vivax in vitro has severely limited opportunities for molecular research on this parasite. Nevertheless, the ability to genotype relapsing infections has led to confirmation that relapses show a strong tendency to be clonal and that multiple relapses in a single patient reflect reactivation of different
Strain Theory of Malaria: The First 50 Years
33
parasite genotypes (Chen et al., 2007; Imwong et al., 2007), in line with the predictions noted above. The notion that the time between being bitten by an infectious mosquito and the onset of either parasitemia or clinical symptoms is dependent, amongst others, on the number of sporozoites inoculated (James, 1920) has been supported by molecular techniques allowing accurate quantification of subpatent parasitemia (Bejon et al., 2005).
8. SUMMARY AND DISCUSSION ‘‘‘The question is,’’ said Alice, ‘‘whether you can make words mean so many different things.’’ ‘‘The question is,’’ said Humpty Dumpty, ‘‘which is to be master—that’s all’’’ (Dodgson, 1872). The historical study of strain theory provides a basis for framing contemporary debate, but the context of the debate has changed. The genetic bases for resistance to chloroquine and sulphadoxine–pyramethamine are now reasonably well-understood. The questions of infectivity and relapse remain underinvestigated. The goal of distinguishing strains in terms of clinical virulence and immunity remains as elusive as ever, in part because it is now possible to see more genetic variability than is expressed in the phenotype. The genes for merozoite stage proteins are known to be highly polymorphic, for example, but much of the observed genetic variation is neutral, or nearly so, such that multiple genotypes have essentially the same phenotype. The clinical, immunological and epidemiological relevance of genetic variability remains poorly understood. It is now evident that each P. falciparum genotype can express 50–60 different phenotypes of the PfEMP1 protein through var gene switching, and that the genes are distributed across all of the P. falciparum genome. It is plausible that heterologous/homologous immunity to P. falciparum is explained by different msp or var gene families and their patterns of crossimmunity. Human immune response may also be heterogeneous, in that the state of clinical immunity in different humans could be conferred by immunity to different sets of immunogens. Continual sexual reassortment of the genes for merozoite-stage proteins and the var genes during meiosis in the mosquito, and strong disruptive selection provide new variation. What, if anything, is a strain? If a strain could be clearly defined in one parasite generation, it might not exist in the next, and an operational definition of a strain with respect to one trait (e.g. drug resistance) might not be coherent with respect to other traits (e.g. infectivity). Aside from the limited experience of malariatherapy and laboratory experiments, the existence of a strain may be too transient for any definition to be useful. Therefore, it is unclear whether increasingly detailed genetic identification
34
F. Ellis McKenzie et al.
of strains will ever converge with the clinical definition of strains pioneered during the 1920s–1970s. The idea that each of the species that cause human malaria consists of ‘varieties, strains or races’ emerged from the observation that malaria infections seemed to differ in severity from place to place and the inference that these differences might arise from biological differences between morphologically identical parasites. When efforts to control and improve responses to malariatherapy made it necessary to distinguish and compare the parasites in clinical use, most such ‘strains’ were named on a geographic basis. The common assumption was that, in nature, ‘in each region, malaria has a character of its own conferred upon it by the peculiarities of the local parasites. No doubt there are a multitude of strains of each species of plasmodium, differing as widely in virulence, response to treatment and tendency to relapse as though they were separate species’ (Hackett, 1937). If, worldwide, ‘each parasite has many strains . . . [but] a strain prevalent in one area is frequently not found in another several hundred miles away’ (Bispham, 1944), then ‘some strains may have a localized habitat or geographical distribution’ (Boyd, 1940a). More drastically, researchers at the Horton Hospital concluded that their three decades of studies had given evidence for several strains each of P. falciparum and P. vivax, ‘from widely separated geographical areas’, but only one each of P. malariae and P. ovale, and that ‘while it is fairly certain that there are different geographical strains of malaria parasites it is extremely difficult to detect variations sufficiently well defined to justify the conclusion that different strains can exist within a single circumscribed locality’ (Shute and Maryon, 1954). The number and geographical distribution of strains—along with their infectivity and relapse characteristics—became topics of wider practical concern with the rising potential for introductions and reintroductions of malaria during World War II. Though the introduction of a particular ‘new’ strain might be probabilistic, its persistence and spread might be constrained by the relative susceptibilities of local mosquitoes or local humans. In humans, presumably, protective responses to endemic strains developed with continued exposure. Thus, ‘among the foreign infections brought into this country, there must be many different and distinct strains. As these are propagated in nature they are added to the various strains already indigenous . . . where there is little malaria now, outbreaks due to importation of foreign strains would be fairly obvious . . . [but elsewhere] the spread of foreign malaria, except under unusual and rare circumstances, could not be detected . . . As these foreign strains are immunologically distinct, it means that no protection is gained by previous infections with native malarias . . . [so] we can expect additional strains to be added to those already present in this country’ (Young et al., 1949).
Strain Theory of Malaria: The First 50 Years
35
During this same period, concurrent multiple strain infections became accepted as common and significant. Though it was well known that multiple Plasmodium species co-existed at almost all malaria-endemic sites, and that co-infecting Plasmodium species appeared to interact in suppressing each other’s parasitemia (Boyd and Kitchen, 1937, 1938), the idea that strains might similarly co-exist, co-infect and interact took hold slowly and unevenly. Understanding strain co-infections posed complex challenges: sequential infection or superinfection might elicit homologous or heterologous responses, in clinical or parasitological terms, and might influence infectivity, relapse or other features. Thus, ‘considering the opportunities for mixed infections of an unknown number of strains, it would be remarkable if any case of malaria resembled another . . . and each of these strains in turn may not be pure but may comprise a different assortment of immunological and other elements, part of which it owns in common with other strains of the same region . . . We are only at the beginning of these studies . . . [and] it is only natural that the imagination of workers in this field has seized upon the existence of such strains to explain all kinds of obscure phenomena in malaria’ (Hackett, 1937). If ‘strains’ could not be considered independent or immutable, but collections of changeable, exchangeable elements, expressed as antigenic and other properties, how could they be defined? Thus, ‘the problem of strains within species is both interesting and important, but it is by no means easy to define what is meant by this term; indeed, we have been compelled to ask ourselves, ‘‘What is a strain?’’ . . . if there are no insurmountable barriers which would prevent the spread of the parasite either by man or by the mosquito, it is the persistence of separate strains within a locality which are so difficult to comprehend, especially in hyperendemic areas . . . [since] if each strain is to retain its individuality, it must be immune to cross breeding with other strains. If several strains of a species were present in a given locality, presumably it would not be long before a large proportion of the population would be infected with two or more strains . . . [so if] a strain retains its identity in a locality where other strains occur . . . the gametocytes of one strain must be resistant to fertilization by another strain’ (Shute and Maryon, 1954). In the late 1960s, a WHO expert committee noted that ‘a parasite strain has been defined as ‘‘a population of common stock descending from a single ancestor or derived from a single source and maintained without intermixture from other sources through a number of generations’’ [WHO, 1963]. This definition may be appropriate for experimentally selected and maintained laboratory strains, but it is too precise for the present purpose’ (WHO, 1969). With ‘parasites recovered from natural infections in the field . . . the term ‘‘strain’’ is used for a population of parasites, recovered from a source in a given geographical area, that possesses confirmed or suspected
36
F. Ellis McKenzie et al.
distinctive characteristics that may be the results of the pressure of natural selection . . . [thus] biological differences in the behaviour of P. falciparum may be apparent in (a) the susceptibility to drugs, (b) the pattern of human infection, (c) the immunological response of the host and (d) the ability to infect mosquitos’ (WHO, 1969). In addition, it was thought that three types of P. vivax strains could be distinguished on the basis of their relapse patterns (WHO, 1969). That is, while a new concern about precision was noted, and discounted, the listing of strain characteristics remained identical to that given in the early 1930s. In the 1970s and early 1980s, when it became possible to clone and cultivate P. falciparum (Rosario, 1981; Trager and Jensen, 1976; Trager et al., 1981), it became clear that virtually every natural ‘isolate’ contained a mixture of parasite entities, each of which when cloned and cultivated might demonstrate strikingly different phenotypic properties with respect to growth rate, drug susceptibility, gametocyte production, antigen and enzyme variants (Burkot et al., 1984; Graves et al., 1984b). This wide range of ‘clones’ was seen to reflect ‘the extent of the genetic diversity which can exist within a single isolate, or ‘‘strain,’’ of these parasites’; thus, were an isolate cultured in conditions which favored the growth of some clones over others, ‘some ‘‘strains’’ of P. falciparum might well undergo changes in such characters as drug resistance or antigenic phenotype’ (Thaithong et al., 1984). Similarly, the report of a 1981 WHO expert committee on ‘malaria parasite strain characterization’ considered ‘isolate’ and ‘strain’ synonymous, noting that ‘the advent of cloning of asexual blood forms of P. falciparum is expected to provide a number of well-characterized uniform strains’ with respect to ‘the available biological markers, including drug sensitivity, isoenzymes, antigenic determinants, plasmodial infectivity to insect vectors, and DNA and other biochemical characteristics’ (WHO, 1981). If an ‘isolate’ or ‘strain’ was a community of parasite entities, however, it was not at all clear what constraints preserved its character from one host to the next. If ‘phenotype’ was a matter of proportions within a mixture, then it might be the uniformity or diversity of hosts and transmission between them that determined phenotypic stability by shaping those proportions: hosts were mixing vessels, sampled by mosquitoes. But, for parasites with an obligate sexual phase in the mosquito, it was not clear how ‘clones’ could be the constituent entities in nature. The questions have since shifted to (multi-locus) genotypes, but they still echo aspects of strain theory from long ago, for example, that ‘partial immunities, partial tolerances might be explained by the loss of certain elements during the passages of composite strains’ (Hackett, 1937). Much of our current knowledge of malaria derives from investigations into the ‘obscure phenomena’ of strains. This is most obviously so in
Strain Theory of Malaria: The First 50 Years
37
malaria immunology—for example, in distinguishing between homologous and heterologous, clinical and parasitological aspects of response, or in proposing that responses might differ due to premunition or inherent and acquired differences between hosts. Not until the mid-1970s, for instance, could the insusceptibility of black patients to P. vivax malariatherapy, and the rarity of P. vivax infections in Sub-Saharan Africa, be attributed to an absence of Duffy receptor (Miller et al., 1976)—a finding counter to the suggestion, noted above, that ‘the human intermediate host is not likely to be a factor in limiting the extension of the range of strains of these parasites’ (Boyd et al., 1938a). Similarly, over time, the observation that while incubation and prepatent periods might vary between strains, they were generally shorter with blood than with sporozoite inoculation—that ‘the method of inoculation (natural or artificial) appears to have some influence on subsequent events’ (Stratman-Thomas, 1941)—led to the hypothesis that the natural parasite life-cycle included an intermediate stage in fixed tissue: it ‘may be that by blood inoculation only the forms of the parasite which live in red blood corpuscles are introduced, whilst by the natural method of infection a form of the parasite is introduced which has always lived, not in red blood corpuscles, but in tissue cells . . . true (long interval) relapses, and recurrences are not observed (so far as we can ascertain) in inoculated cases, while they occur in 50 per cent of mosquito-infected cases. This difference has led to various suggestions about what happens to sporozoites when they are injected by the mosquito’ (James, 1931). Naturally-induced infections often appeared less responsive to drugs, less infectious to mosquitoes and less resistant to re-infection during the initial latent period than later (James and Ciuca, 1938), but hepatic forms of P. vivax and P. falciparum were not detected until the late 1940s (Shortt and Garnham, 1948; Shortt et al., 1951), and P. vivax hypnozoites not until the 1980s (Krotoski et al., 1982). Yet it can still be asked, as it was 50 years ago, ‘Is not the word ‘‘strain’’ in regard to malaria used much too loosely?’ (Shute, 1958). Beginning in the 1920s, the theory was that ‘strains’ exist, defined by distinct, observable characteristics. Over the next 50 years, attempts to tighten ‘loose’ definitions produced important insights into clinical virulence, infectivity, reaction to anti-malarial remedies, antigenic properties, latency and relapse—among them the insights that, upon closer examination, some characteristics that had seemed evident might prove too elusive to be useful in definition. Strain theory had seemed to make testable predictions, but the insights it produced arose from its ramifying ambiguities, and from the recognition of ever more complex confounding variables. What, if anything, will be said of the second 50 years of strain theory? The first dictionary of the English language defined ‘theory’as ‘speculation; not practice; scheme; plan or system yet subsisting only in the mind’
38
F. Ellis McKenzie et al.
(Johnson, 1755). Atoms, genes and germs were once ‘only in the mind’, with definitions that were sometimes ‘too loose’ or ‘too precise’, but those terms have persisted as the concepts were refined and explanations tested, with profound practical consequences. Other terms—for example, ‘phlogiston’ and ‘humours’—have been abandoned. If the four species that cause human malaria contain coherent units that behave as ‘strains’, interventions would likely be improved by understanding them. Progress towards that understanding requires a testable, falsifiable theory of Plasmodium ‘strains’, no less now, than in the 1920s.
ACKNOWLEDGEMENTS We gratefully acknowledge the contributions of J. Makulowich, B. C. Sorkin and M. Taylor.
REFERENCES A-Elbasit, I. E., Elghazali, G., A-Elgadir, T. M., Hamad, A. A., Babiker, H. A., Elbashir, M. I., and Giha, H. A. (2007). Allelic polymorphism of MSP2 gene in severe P. falciparum malaria in an area of low and seasonal transmission. Parasitol. Res. 102, 29–34. Anderson, T. J., and Roper, C. (2005). The origins and spread of antimalarial drug resistance: Lessons for policy makers. Acta Trop. 94, 269–280. Arez, A. P., Snounou, G., Pinto, J., Sousa, C. A., Modiano, D., Ribeiro, H., Franco, A. S., Alves, J., and do Rosario, V. E. (1999). A clonal Plasmodium falciparum population in an isolated outbreak of malaria in the Republic of Cabo Verde. Parasitology 118, 347–355. Arnot, D. (1998). Unstable malaria in Sudan: The influence of the dry season. Clone multiplicity of Plasmodium falciparum infections in individuals exposed to variable levels of disease transmission. Trans. R. Soc. Trop. Med. Hyg. 92, 580–585. Babiker, H. A., Ranford-Cartwright, L. C., Currie, D., Charlwood, J. D., Billingsley, P., Teuscher, T., and Walliker, D. (1994). Random mating in a natural population of the malaria parasite Plasmodium falciparum. Parasitology 109, 413–421. Babiker, H. A., Abdel-Muhsin, A. A., Hamad, A., Mackinnon, M. J., Hill, W. G., and Walliker, D. (2000). Population dynamics of Plasmodium falciparum in an unstable malaria area of eastern Sudan. Parasitology 120, 105–111. Badenski, G. T. (1936). ‘‘A Note Describing the Technique Employed in Malariatherapy in the Centres at Rome and Horton (England).’’ League of Nations Health Organisation Malaria Commission, Geneva(document 237). Barnes, K. I., and White, N. J. (2005). Population biology and antimalarial resistance: The transmission of antimalarial drug resistance in Plasmodium falciparum. Acta Trop. 94, 230–240. Bejon, P., Andrews, L., Andersen, R. F., Dunachie, S., Webster, D., Walther, M., Gilbert, S. C., Peto, T., and Hill, A. V. (2005). Calculation of liver-to-blood inocula, parasite growth rates, and preerythrocytic vaccine efficacy, from serial quantitative polymerase chain reaction studies of volunteers challenged with malaria sporozoites. J. Infect. Dis. 191, 619–626. Bhasin, V. K., and Trager, W. (1984). Gametocyte-forming and non-gametocyte-forming clones of Plasmodium falciparum. Am. J. Trop. Med. Hyg. 33, 534–537. Bishop, A. (1955). Problems concerned with gametogenesis in Haemosporidiidea, with particular reference to the genus Plasmodium. Parasitology 45, 163–185. Bispham, W. N. (1944). ‘‘Malaria.’’ Williams and Wilkins Company, Baltimore.
Strain Theory of Malaria: The First 50 Years
39
Blackburn, C. R. B. (1948). Observations on the development of resistance to vivax malaria. Trans. R. Soc. Trop. Med. Hyg. 42, 117–162. Boyd, M. F. (1940a). On strains or races of the malaria parasites. Am. J. Trop. Med. 20, 69–80. Boyd, M. F. (1940b). The influence of sporozoite dosage in vivax malaria. Am. J. Trop. Med. 20, 279–286. Boyd, M. F. (1941). The infection in the intermediate host: Symptomatology, general considerations. In ‘‘A Symposium on Human Malaria’’ (F. R. Moulton, ed.), pp. 163–182. American Association for the Advancement of Science, Washington DC. Boyd, M. F. (1942a). On the varying infectiousness of different patients infected with vivax malaria. Am. J. Trop. Med. 22, 73–81. Boyd, M. F. (1942b). Criteria of immunity and susceptibility in naturally induced vivax malaria infections. Am. J. Trop. Med. 22, 217–226. Boyd, M. F. (1949). Epidemiology of malaria: Factors related to the intermediate host. In ‘‘Malariology’’ (M. F. Boyd, ed.), pp. 551–607. W. B. Saunders Company, Philadelphia. Boyd, M. F., and Kitchen, S. F. (1936a). The comparative susceptibility of Anopheles quadrimaculatus, Say, and Anopheles punctipennis, Say, to Plasmodium vivax, Grassi, and Plasmodium falciparum, Welch. Am. J. Trop. Med. 16, 67–71. Boyd, M. F., and Kitchen, S. F. (1936b). On the localization of the geographical distribution of the McCoy strain of Plasmodium vivax. Am. J. Trop. Med. 16, 583–587. Boyd, M. F., and Kitchen, S. F. (1937). Simultaneous inoculation with Plasmodium vivax and Plasmodium falciparum. Am. J. Trop. Med. 17, 855–861. Boyd, M. F., and Kitchen, S. F. (1938). Vernal vivax activity in persons simultaneously inoculated with Plasmodium vivax and Plasmodium falciparum. Am. J. Trop. Med. 18, 505–514. Boyd, M. F., and Kitchen, S. F. (1945). On the heterologous value of acquired immunity to Plasmodium falciparum. J. Natl. Malar. Soc. 4, 301–306. Boyd, M. F., and Kitchen, S. F. (1948). On the homogeneity or heterogeneity of Plasmodium vivax infections acquired in highly endemic regions. Am. J. Trop. Med. 28, 29–34. Boyd, M. F., and Stratman-Thomas, W. K. (1933a). Studies on benign tertian malaria 1. On the occurrence of acquired tolerance to Plasmodium vivax. Am. J. Hyg. 17, 55–59. Boyd, M. F., and Stratman-Thomas, W. K. (1933b). Studies on benign tertian malaria 3. On the absence of a heterologous tolerance to Plasmodium vivax. Am. J. Hyg. 18, 482–484. Boyd, M. F., and Stratman-Thomas, W. K. (1933c). Studies on benign tertian malaria 6. On heterologous tolerance. Am. J. Hyg. 20, 482–487. Boyd, M. F., and Stratman-Thomas, W. K. (1934). The comparative susceptibility of A. quadrimaculatus, Say, and A. crucians, Wied (inland variety) to the parasites of human malaria. Am. J. Hyg. 20, 247–257. Boyd, M. F., Stratman-Thomas, W. K., and Kitchen, S. F. (1935). On the relative susceptibility of Anopheles quadrimaculatus to Plasmodium vivax and Plasmodium falciparum. Am. J. Trop. Med. 15, 485–493. Boyd, M. F., Stratman-Thomas, W. K., and Kitchen, S. F. (1936). On acquired immunity to Plasmodium falciparum. Am. J. Trop. Med. 16, 139–145. Boyd, M. F., Carr, H. P., and Rozeboom, L. E. (1938a). On the comparative susceptibility of certain species of Nearctic and Neotropical anophelines to certain strains of P. vivax and P. falciparum from the same regions. Am. J. Trop. Med. 18, 157–168. Boyd, M. F., Kupper, W. H., and Matthews, C. B. (1938b). A deficient homologous immunity following simultaneous inoculation with two strains of Plasmodium vivax. Am. J. Trop. Med. 18, 521–524. Boyd, M. F., Kitchen, S. F., and Matthews, C. B. (1941). On the natural transmission of infection from patients concurrently infected with two strains of Plasmodium vivax. Am. J. Trop. Med. 21, 645–652.
40
F. Ellis McKenzie et al.
Bray, R. S., Gunders, A. E., Burgess, R. W., Freeman, J. B., Etzel, E., Guttuso, C., and Colussa, B. (1962). The inoculation of semi-immune African with sporozoites of Laverania falcipara (Plasmodium falciparum) in Liberia. Riv. Malariol. 41, 199–210. Bruce-Chwatt, L. J. (1963). A longitudinal survey of natural malaria infection in a group of West African adults, II. West Afr. Med. J. 12, 199–217. Brumpt, E. (1949). The human parasites of the genus Plasmodium. In ‘‘Malariology’’ (M. F. Boyd, ed.), pp. 65–121. W. B. Saunders Company, Philadelphia. Bull, P. C., Pain, A., Ndungu, F. M., Kinyanjui, S. M., Roberts, D. J., Newbold, C. I., and Marsh, K. (2005). Plasmodium falciparum antigenic variation: Relationships between in vivo selection, acquired antibody response, and disease severity. J. Infect. Dis. 192, 1119–1126. Bunker, H. A., Jr., and Kirby, G. H. (1925). Treatment of general paralysis by inoculation with malaria. J. Am. Med. Assoc. 84, 563–568. Burgess, R. W., and Young, M. D. (1950). The comparative susceptibility of Anopheles quadrimaculatus and Anopheles freeborni to infection by Plasmodium vivax (St. Elizabeth strain). J. Natl. Malar. Soc. 9, 218–221. Burkot, T. R., Williams, J. L., and Schneider, I. (1984). Infectivity to mosquitoes of Plasmodium falciparum clones grown in vitro from the same isolate. Trans. R. Soc. Trop. Med. Hyg. 78, 339–341. Celli, A. (1901). ‘‘Malaria.’’ Longmans, Green, and Co., London. Chen, N., Auliff, A., Rieckmann, K., Gatton, M., and Cheng, Q. (2007). Relapses of Plasmodium vivax infection result from clonal hypnozoites activated at predetermined intervals. J. Infect. Dis. 195, 934–941. Chernin, E. (1984). The malariatherapy of neurosyphilis. J. Parasitol. 70, 611–617. Cicero, M. T. (45BC) De Natura Deorum. In ‘‘Cicero XIX’’ (H. Rackham, transl.), part 3. Harvard University Press, Cambridge, MA, 1933. Ciuca, M., Ballif, L., and Chelarescu-Vieru, M. (1934). Immunity in malaria. Trans. R. Soc. Trop. Med. Hyg. 27, 619–622. Clyde, D. F., and Shute, G. T. (1954). Resistance of East African varieties of Plasmodium falciparum to pyrimethamine. Trans. R. Soc. Trop. Med. Hyg. 48, 495–500. Clyde, D. F., Dawkins, A. T., Jr., Heiner, G. G., McCarthy, V. C., and Hornick, R. B. (1969). Characteristics of four new drug-resistant strains of Plasmodium falciparum from South-east Asia. Mil. Med. 134, 787–794. Coatney, G. R., and Young, M. D. (1941). The taxonomy of the human malaria parasites with notes on the principal American strains. In ‘‘A Symposium on Human Malaria’’ (F. R. Moulton, ed.) pp. 19–24. American Association for the Advancement of Science, Washington DC. Coatney, G. R., Cooper, W. C., Ruhe, D. S., Young, M. D., and Burgess, R. W. (1950a). Studies in human malaria. XVIII. The life pattern of sporozoite-induced St. Elizabeth strain vivax malaria. Am. J. Hyg. 51, 200–215. Coatney, G. R., Cooper, W. C., and Young, M. D. (1950b). Studies in human malaria. XXX. A summary of 204 sporozoite-induced infections with the Chesson strain of Plasmodium vivax. J. Natl. Malar. Soc. 9, 381–396. Coatney, G. R., Collins, W. E., Warren, Mc W., and Contacos, P. G. (1971). ‘‘The Primate Malarias.’’ U. S. Department of Health Education and Welfare, Washington DC. Collins, W. E., and Jeffery, G. M. (1999). A retrospective examination of sporozoite-and trophozoite-induced infections with Plasmodium falciparum. Am. J. Trop. Med. Hyg. 61, 4–48. Collins, W. E., Jeffery, G. M., and Burgess, R. W. (1963). Comparative infectivity of strains of P. falciparum to Anopheles quadrimaculatus. Mosq. News 23, 102–104. Contacos, P. G., Lunn, J. S., and Coatney, G. R. (1963). Drug-resistant falciparum malaria from Cambodia and Malaya. Trans. R. Soc. Trop. Med. Hyg. 57, 417–424.
Strain Theory of Malaria: The First 50 Years
41
Contamin, H., Fandeur, T., Rogier, C., Bonnefoy, S., Konate, L., Trape, J. F., and MercereauPuijalon, O. (1996). Different genetic characteristics of Plasmodium falciparum isolates collected during successive clinical malaria episodes in Senegalese children. Am. J. Trop. Med. Hyg. 54, 632–643. Cooper, W. C., Coatney, G. R., Culwell, W. B., Eyles, D. E., and Young, M. D. (1950). Studies in human malaria. XXVI. J. Natl. Malar. Soc. 9, 187–194. Craig, C. F. (1926). ‘‘Parasitic Protozoa of Man.’’ J. P. Lippincott Company, Philadelphia. Davey, D. G., and Robertson, G. I. (1957). Experiments with antimalarial drugs in man II. A description of methods. Trans. R. Soc. Trop. Med. Hyg. 51, 450–456. Day, K. P., Karamalis, F., Thompson, J., Barnes, D. A., Peterson, C., Brown, H., Brown, G. V., and Kemp, D. J. (1993). Genes necessary for expression of a virulence determinant and for transmission of Plasmodium falciparum are located on a 0.3-magabase region of chromosome 9. Proc. Natl. Acad. Sci. USA 90, 8292–8296. de Rudolf, G. M. (1924). The temperature-chart in artificially-inoculated malaria. J. Trop. Med. Hyg. 27, 259–263. de Rudolf, G. M. (1927). ‘‘Therapeutic Malaria.’’ Oxford University Press, London. Deaderick, W. H. (1909). ‘‘A Practical Study of Malaria.’’ W. B. Saunders Company, Philadelphia. Dobano, C., Rogerson, S. J., Taylor, T. E., McBride, J. S., and Molyneux, M. E. (2007). Expression of merozoite surface protein markers by Plasmodium falciparum-infected erythrocytes in peripheral blood and tissues of children with fatal malaria. Infect. Immun. 75, 643–652. Dodgson, C. L. (1872). ‘‘Through the Looking-Glass.’’ Macmillan and Co, London. Dzikowski, R., Templeton, T. J., and Deitsch, K. (2006). Variant antigen gene expression in malaria. Cell. Microbiol. 8, 1371–1381. Earle, W. C., Perez, M., del Rio, J., and Arzola, C. (1939). Observations on the course of naturally acquired malaria in Puerto Rico. Puerto Rican J. Public Health Trop. Med. 14, 391–406. Edeson, J. F. B., and Field, J. W. (1950). Proguanil-resistant falciparum malaria in Malaya. Br. Med. J. ii, 147–151. Ehrman, F. C., Ellis, J. M., and Young, M. D. (1945). Plasmodium vivax Chesson strain. Science 101, 377. Fairley, N. H. (1946). Atebrin susceptibility of the Aitaipe-Wewak strains of P. falciparum and P. vivax. Trans. R. Soc. Trop. Med. Hyg. 40, 229–273. Ferreira, M. U., and Hartl, D. L. (2007). Plasmodium falciparum: Worldwide sequence diversity and evolution of the malaria vaccine candidate merozoite surface protein-2 (MSP-2). Exp. Parasitol. 115, 32–40. Ferreira, M. U., Ribeiro, W. L., Tonon, A. P., Kawamoto, F., and Rich, S. M. (2003). Sequence diversity and evolution of the malaria vaccine candidate merozoite surface protein-1 (MSP-1) of Plasmodium falciparum. Gene 304, 65–75. Fiertz, C. O. (1926). The malarial treatment of general paralysis. State Hosp. Q. xi, 626–643. Filipe, J. A. N., Riley, E. M., Drakeley, C. J., Sutherland, C. J., and Ghani1, A. C. (2007). Determination of the processes driving the acquisition of immunity to malaria using a mathematical transmission model. PLoS Comput. Biol. 3, e255. Francis, S. E., Malkov, V. A., Oleinikov, A. V., Rossnagle, E., Wendler, J. P., Mutabingwa, T. K., Fried, M., and Duffy, P. E. (2007). Six genes are preferentially transcribed by the circulating and sequestered forms of Plasmodium falciparum that infect pregnant women. Infect. Immun. 75, 4838–4850. Garnham, P. C. C. (1966). ‘‘Malaria Parasites and other Haemosporidia.’’ Blackwell Scientific Publications, Oxford. Gatton, M. L., and Cheng, Q. (2004). Modeling the development of acquired clinical immunity to Plasmodium falciparum malaria. Infect. Immun. 72, 6538–6545.
42
F. Ellis McKenzie et al.
Golgi, C. (1889). On the cycle of development of malarial parasites in tertian fever: Differential diagnosis between the intracellular malarial parasites of tertian and quartan fever. In ‘‘Tropical Medicine and Parasitology: Classic Investigations’’ (B. H. Kean, K. E. Mott, and A. J. Russell, eds.), Volume I, pp. 26–35. Cornell University Press, Ithaca, NY, 1978. Grant, A. R. (1923). The treatment of general paralysis by malaria. Br. Med. J. ii, 698–700. Graves, P. M., Carter, R., and McNeill, K. M. (1984a). Gametocyte production in cloned lines of Plasmodium falciparum. Am. J. Trop. Med. Hyg. 33, 1045–1050. Graves, P. M., Carter, R., Keystone, J. S., and Seeley, D. C., Jr. (1984b). Drug sensitivity and isoenzyme type in cloned lines of Plasmodium falciparum. Am. J. Trop. Med. Hyg. 33, 212–219. Gray, J. C., Corran, P. H., Mangia, E., Gaunt, M. W., Li, Q., Tetteh, K. K., Polley, S. D., Conway, D. J., Holder, A. A., Bacarese-Hamilton, T., Riley, E. M., and Crisanti, A. (2007). Profiling the antibody immune response against blood stage malaria vaccine candidates. Clin. Chem. 53, 1244–1253. Gregson, A., and Plowe, C. V. (2005). Mechanisms of resistance of malaria parasites to antifolates. Pharmacol. Rev. 57, 117–145. Gupta, S., and Day, K. P. (1994a). A strain theory of malaria transmission. Parasitol. Today 10, 476–481. Gupta, S., and Day, K. P. (1994b). A theoretical framework for the immunoepidemiology of Plasmodium falciparum malaria. Parasite Immunol. 16, 361–370. Gupta, S., Snow, R. W., Donnelly, C. A., Marsh, K., and Newbold, C. (1999). Immunity to non-cerebral severe malaria is acquired after one or two infections. Nat. Med. 5, 340–343. Hackett, L. W. (1937). ‘‘Malaria in Europe.’’ Oxford University Press, London. Hackett, L. W. (1941). Malaria and the community. In ‘‘A Symposium on Human Malaria’’ (F. R. Moulton, ed.), pp. 148–156. American Association for the Advancement of Science, Washington DC. Hankey, D. D., Jones, R., Jr., Coatney, G. R., Alviing, A. S., Coker, W. G., Garrison, P. L., and Donovan, W. N. (1953). Korean vivax malaria. I. Am. J. Trop. Med. Hyg. 2, 958–969. Hastings, I. M., and Donnelly, M. J. (2005). The impact of antimalarial drug resistance mutations on parasite fitness, and its implications for the evolution of resistance. Drug Resist. Updat. 8, 43–50. Hehir, P. (1927). ‘‘Malaria in India.’’ Oxford University Press, London. Hill, E., and Amatuzio, D. S. (1949). Southwest Pacific vivax malaria. Am. J. Trop. Med. 29, 203–214. Horing, R. O. (1947). Induced and war malaria. J. Trop. Med. Hyg. 50, 150–160. Huber, W., Haji, H., Charlwood, J. D., Certa, U., Walliker, D., and Tanner, M. (1998). Genetic characterization of the malaria parasite Plasmodium falciparum in the transmission from the host to the vector. Parasitology 116, 95–101. Huff, C. G. (1938). The significance of different strains of malaria and mosquitoes in the epidemiology of the disease. Am. J. Med. Technol. 4, 41–46. Imwong, M., Snounou, G., Pukrittayakamee, S., Tanomsing, N., Kim, J. R., Nandy, A., Guthmann, J. P., Nosten, F., Carlton, J., Looareesuwan, S., Nair, S., Sudimack, D., et al. (2007). Relapses of Plasmodium vivax infection usually result from activation of heterologous hypnozoites. J. Infect. Dis. 195, 927–933. James, S. P. (1920). ‘‘Malaria at Home and Abroad.’’ John Bale, Sons & Danielsson, Ltd., London. James, S. P. (1931). Some general results of a study of induced malaria in England. Trans. R. Soc. Trop. Med. Hyg. 27, 477–538. James, S. P., and Ciuca, M. (1938). Species and races of human malaria parasites and a note on immunity. In ‘‘Acta Conventus Tertii de Tropicis Atque Malariae Morbis.’’ Volume 2, pp. 269–281. C. A. Spin & Zoon N. V., Amsterdam.
Strain Theory of Malaria: The First 50 Years
43
James, S. P., Nicol, W. B., and Shute, P. G. (1932). A study of induced malignant tertian malaria. Proc. R. Soc. Med. 25, 1153–1186. James, S. P., Nicol, W. B., and Shute, P. G. (1936). Clinical and parasitological observations on induced malaria. Proc. R. Soc. Med. 29, 879–894. Jeffery, G. M. (1951). Observations on a gametocyteless strain of Plasmodium falciparum. J. Natl. Malar. Soc. 10, 337–344. Jeffery, G. M. (1956). Relapses with Chesson strain Plasmodium vivax following treatment with chloroquine. Am. J. Trop. Med. Hyg. 5, 1–13. Johnson, S. (1755). ‘‘A Dictionary of the English Language.’’ James Maxwell, London. Kaplan, L. I., Read, H. S., and Becker, F. T. (1946). Homologous and heterologous strains of Plasmodium vivax: A cross-inoculation study of malaria strain immunity. J. Lab. Clin. Med. 31, 400–408. Kenrick, W. H. (1910). Malaria in the Central Provinces. Proceedings of the Imperial Malaria Conference, pp. 22–24. Government Central Branch Press, Simla. Kitchen, S. F. (1949). Vivax malaria. In ‘‘Malariology’’ (M. F. Boyd, ed.), pp. 1027–1045. W. B. Saunders Company, Philadelphia. Kraemer, S. M., and Smith, J. D. (2006). A family affair: var genes, PfEMP1 binding, and malaria disease. Curr. Opin. Microbiol. 9, 374–380. Krotoski, W. A., Collins, W. E., Bray, R. S., Garnham, P. C., Cogswell, F. B., Gwadz, R. W., Killick-Kendrick, R., Wolf, R., Sinden, R., Koontz, L. C., and Stanfill, P. S. (1982). Demonstration of hypnozoites in sporozoite-transmitted Plasmodium vivax infection. Am. J. Trop. Med. Hyg. 31, 1291–1293. Kublin, J. G., Cortese, J. F., Njunju, E. M., Mukadam, R. A., Wirama, J. J., Kazembe, P. N., Djimde, A. A., Kouriba, B., Taylor, T. E., and Plowe, C. V. (2003). Reemergence of chloroquine-sensitive Plasmodium falciparum malaria after cessation of chloroquine use in Malawi. J. Infect. Dis. 187, 1870–1875. Kupper, W. H. (1939). ‘‘The Malarial Therapy of General Paralysis and Other Conditions.’’ Edwards Brothers, Inc., Ann Arbor MI. Kyriacou, H. M., Stone, G. N., Challis, R. J., Raza, A., Lyke, K. E., Thera, M. A., Kone, A. K., Doumbo, O. K., Plowe, C. V., and Rowe, J. A. (2006). Differential var gene transcription in Plasmodium falciparum isolates from patients with cerebral malaria compared to hyperparasitaemia. Mol. Biochem. Parasitol. 150, 211–218. Lambrechts, L., Halbert, J., Durand, P., Gouagna, L. C., and Koella, J. C. (2005). Host genotype by parasite genotype interactions underlying the resistance of anopheline mosquitoes to Plasmodium falciparum. Malar. J. 11, 3. Laserson, K. F., Petralanda, I., Almera, R., Barker, R. H., Jr., Spielman, A., Maguire, J. H., and Wirth, D. F. (1999). Genetic characterization of an epidemic of Plasmodium falciparum malaria among Yanomami Amerindians. J. Infect. Dis. 180, 2081–2085. Laveran, A. (1893). ‘‘Paludism.’’ The New Syndenham Society, London. League of Nations Health Organisation Malaria Commission (1934). Ten years of activity of the Commission Geneva (document 212). Leslie, J. W. T. (1910). Malaria in India. In ‘‘Proceedings of the Imperial Malaria Conference’’, pp. 3–11. Government Central Branch Press, Simla. Lilly, G. A. (1925). The treatment of general paralysis at Hanwell Mental Hospital. J. Ment. Sci. 71, 267–278. Lomholt, M. (1944). ‘‘Clinic and Prognosis of Malaria-Treated Paralysis.’’ Ejnar Munksgaard, Copenhagen. Macbride, H. J. (1924). The treatment of general paralysis of the insane by malaria. J. Neurol. Psychopathol. 5, 13–27. Macdonald, G. (1957). ‘‘The Epidemiology and Control of Malaria.’’ Oxford University Press, London.
44
F. Ellis McKenzie et al.
Machado, R. L., Povoa, M. M., Calvosa, V. S., Ferreira, M. U., Rossit, A. R., dos Santos, E. J., and Conway, D. J. (2004). Genetic structure of Plasmodium falciparum populations in the Brazilian Amazon region. J. Infect. Dis. 190, 1547–1555. Mackinnon, M. J., and Read, A. F. (1999). Selection for high and low virulence in the malaria parasite Plasmodium chabaudi. Proc. R. Soc. London Ser. B 266, 741–748. Mackinnon, M. J., Gaffney, D. J., and Read, A. F. (2002). Virulence in rodent malaria: Host genotype by parasite genotype interactions. Infect. Genet. Evol. 1, 287–296. Marchiafava, E., and Bignami, A. (1894). ‘‘On Summer-Autumn Malarial Fevers.’’ The New Syndenham Society, London. Marchoux, E. (1922). Multiplicite des races dans les trois formes de parasites du paludisme. Bulletin de la Socie´te´ de Pathologie Exotique 15, 108–109. McNamara, J. V., Rieckmann, K. H., Frischer, H., Stockert, T. A., Carson, P. E., and Powell, R. D. (1967). Acquired decrease in sensitivity to quinine observed during studies with a strain of chloroquine-resistant Plasmodium falciparum. Ann. Trop. Med. Parasitol. 61, 386–395. Miller, L. H., Mason, S. J., Clyde, D. F., and McGinniss, M. H. (1976). The resistance factor to Plasmodium vivax in blacks. The Duffy-blood-group genotype, FyFy. N. Engl. J. Med. 295, 302–304. Mita, T., Kaneko, A., Hwaihwanje, I., Taukahara, T., Takahashi, N., Osawa, H., Tanabe, K., Kobayakawa, T., and Bjorkman, A. (2006). Rapid selection of dhfr mutant allele in Plasmodium falciparum isolates after the introduction of sulfadoxine/pyrimethamine in combination with 4-aminoquinolones in Papua New Guinea. Infect. Genet. Evol. 6, 447–452. Moore, D. V., and Lanier, J. E. (1961). Observations on two Plasmodium falciparum infections with an abnormal response to chloroquine. Am. J. Trop. Med. Hyg. 10, 5–9. Napier, L. E. (1947). The malaria relapse problem. J. Trop. Med. Hyg. 50, 147–150. Nocht, B., and Mayer, M. (1937). ‘‘Malaria.’’ John Bale Medical Publications, London. Pearce, R., Malisa, A., Kachur, S. P., Barnes, K., Sharp, B., and Roper, C. (2005). Reduced variation around drug-resistant dhfr alleles in African Plasmodium falciparum. Mol. Biol. Evol. 22, 1834–1844. Peyerl-Hoffmann, G., Jelinek, T., Kilian, A., Kabagambe, G., Metzger, W. G., and von Sonnenburg, F. (2001). Genetic diversity of Plasmodium falciparum and its relationship to parasite density in an area with different malaria endemicities in West Uganda. Trop. Med. Int. Health 6, 607–613. Pijper, A., and Russell, E. D. (1926). Malaria treatment of general paralysis: A report on 44 cases. S. Afr. Med.l Rec. 35, 292–305. Ramsdale, C. D., and Coluzzi, M. (1975). Studies on the infectivity of tropical African strains of Plasmodium falciparum to some southern European vectors of malaria. Parassitologia 17, 39–48. Ranford-Cartwright, L. C., Balfe, P., Carter, R., and Walliker, D. (1993). Frequency of crossfertilization in the human malaria parasite Plasmodium falciparum. Parasitology 107, 11–18. Robertson, G. I., Davey, D. G., and Fairley, N. H. (1952). Cross-resistance between ‘‘Daraprim’’ and Proguanil. Br. Med. J. ii, 1255–1256. Rollo, I. M., Williamson, J., and Lourie, E. M. (1948). Acquired paludrine-resistance in Plasmodium gallinaceum. II. Failure to produce such resistance by prolonged treatment of latent infections. Ann. Trop. Med. Parasitol. 42, 241–248. Rosario, V. (1981). Cloning of naturally occurring mixed infections of malaria parasites. Science 212, 1037–1038. Ross, R. (1910). ‘‘The Prevention of Malaria.’’ E. P. Dutton & Company, New York. Russell, P. F., West, L. S., and Manwell, R. D. (1946). ‘‘Practical Malariology.’’ W. B. Saunders Company, Philadelphia. Salanti, A., Dahlback, M., Turner, L., Nielsen, M. A., Barfod, L., Magistrado, P., Jensen, A. T., Lavstsen, T., Ofori, M. F., Marsh, K., Hviid, L., and Theander, T. G. (2004). Evidence for the involvement of VAR2CSA in pregnancy-associated malaria. J. Exp. Med. 200, 1197–1203.
Strain Theory of Malaria: The First 50 Years
45
Sallenave-Sales, S., Daubersies, P., Mercereau-Puijalon, O., Rahimalala, L., Contamin, H., Druilhe, P., Daniel-Ribeiro, C. T., and Ferreira-da-Cruz, M. F. (2000). Plasmodium falciparum: A comparative analysis of the genetic diversity in malaria-mesoendemic areas of Brazil and Madagascar. Parasitol. Res. 86, 692–698. Sama, W., Owusu-Agyei, S., Felger, I., Dietz, K., and Smith, T. (2006). Age and seasonal variation in the transition rates and detectability of Plasmodium falciparum malaria. Parasitology 132, 13–21. Schofield, L., and Grau, G. E. (2005). Immunological processes in malaria pathogenesis. Nat. Rev. Immunol. 5, 722–735. Schwartz, R., Mansuy, M. M., and John, W. C. (1950). Current concepts on malaria. Arch. Intern. Med. 86, 837–856. Seaton, D. R., and Adams, A. R. D. (1949). Acquired resistance to proguanil in Plasmodium falciparum. Lancet ii, 323–324. Seaton, D. R., and Lourie, E. M. (1949). Acquired resistance to proguanil (paludrine) in Plasmodium vivax. Lancet i, 394–395. Sergent, E., Parrot, L., and Donatien, A. (1924). Une question de terminologie, immuniser et premunir. Bulletin de la Socie´te´ de Pathologie Exotique 17, 37–38. Sergent, E., Parrot, L., and Donatien, A. (1925). On the necessity of having a term to express the resistance of carriers of germs to superimposed infections. Trans. R. Soc. Trop. Med. Hyg. 18, 383–385. Shortt, H. E., and Garnham, P. C. C. (1948). The pre-erythrocytic development of Plasmodium cynomolgi and Plasmodium vivax. Trans. R. Soc. Trop. Med. Hyg. 41, 785–795. Shortt, H. E., Fairley, N. H., Covell, G., Shute, P. G., and Garnham, P. C. C. (1951). The pre-erythrocytic stage of Plasmodium falciparum. Trans. R. Soc. Trop. Med. Hyg. 44, 405–419. Shute, P. G. (1937). ‘‘Report on a third visit to Roumania for the Study of Malaria.’’ League of Nations Health Organisation Malaria Commission, Geneva (document 250). Shute, P. G. (1940). Failure to infect English specimens of Anopheles maculopennis var. atroparvus with certain strains of Plasmodium falciparum of tropical origin. J. Trop. Med. Hyg. 43, 175–178. Shute, P. G. (1946). Latency and long-term relapses in benign tertian malaria. Trans. R. Soc. Trop. Med. Hyg. 40, 189–200. Shute, P. G. (1951). Mosquito infection in artificially induced malaria. Br. Med. Bull. 8, 56–63. Shute, P. G. (1958). Thirty years of malaria-therapy. J. Trop. Med. Hyg. 61, 557–561. Shute, P. G., and Maryon, M. (1954). A contribution to the problem of strains of human plasmodium. Riv. Malariol. 33, 1–21. Sinden, R. E., Alavi, Y., and Raine, J. D. (2004). Mosquito-malaria interactions: A reappraisal of the concepts of susceptibility and refractoriness. Insect Biochem. Mol. Biol. 34, 625–629. Sinton, J. A. (1931). Studies in malaria, with special reference to treatment. Part XV. Does the strain of parasite influence cure? Indian J. Med. Res. 18, 845–853. Sinton, J. A. (1939). A summary of our present knowledge of the mechanism of immunity in malaria. J. Malaria Inst. India 2, 71–83. Sinton, J. A. (1940). Studies of infections with Plasmodium ovale. V. Trans. R. Soc. Trop. Med. Hyg. 33, 585–595. Smith, F. (1898). Malaria: Immunity: Absence of negro immunity: Variety. Br. Med. J. ii, 1807. Stephens, J. W. W. (1922). A new malaria parasite of man. Ann. Trop. Med. Parasitol. 16, 383–388. Stratman-Thomas, W. K. (1941). The infection in the intermediate host. Symptomatology: Vivax malaria. In ‘‘A Symposium on Human Malaria’’ (F. R. Moulton, ed.), pp. 183–189. American Association for the Advancement of Science, Washington DC. Taliaferro, W. H. (1949). Immunity to the malaria infections. In ‘‘Malariology’’ (M. F. Boyd, ed.), pp. 935–965. W. B. Saunders Company, Philadelphia.
46
F. Ellis McKenzie et al.
Thaithong, S., Beale, G. H., Fenton, B., McBride, J., Rosario, V., Walker, A., and Walliker, D. (1984). Clonal diversity in a single isolate of the malaria parasite Plasmodium falciparum. Trans. R. Soc. Trop. Med. Hyg. 78, 242–245. Thomson, J. G. (1931). The question of immunity in man to protozoal diseases. Proc. R. Soc. Med. 24, 499–513. Thomson, J. G. (1933). Immunity in malaria. Trans. R. Soc. Trop. Med. Hyg. 26, 483–514. Trager, W., and Jensen, J. B. (1976). Human malaria parasites in continuous culture. Science 193, 673–675. Trager, W., Tershakovec, M., Lyandvert, L., Stanley, H., Lanners, N., and Gubert, E. (1981). Clones of the malaria parasite Plasmodium falciparum obtained by microscopic selection: Their characterization with regard to knobs, chloroquine sensitivity, and formation of gametocytes. Proc. Natl. Acad. Sci. USA 78, 6527–6530. Viebig, N. K., Gamain, B., Scheidig, C., Lepolard, C., Przyborski, J., Lanzer, M., Gysin, J., and Scherf, A. (2005). A single member of the Plasmodium falciparum var multigene family determines cytoadhesion to the placental receptor chondroitin sulphate A. EMBO Rep. 6, 775–778. Vlachou, D., and Kafatos, F. C. (2005). The complex interplay between mosquito positive and negative regulators of Plasmodium development. Curr. Opin. Microbiol. 8, 415–421. Voller, A. (1964). Comments on the detection of malaria antibodies. Am. J. Trop. Med. Hyg. 13, 204–208. Wagner-Jauregg, J. (1922). The treatment of general paresis by inoculation of malaria. J. Nerv. Ment. Dis. 55, 369–375. Walliker, D. (1983). The genetic basis of diversity in malaria parasites. Adv. Parasitol. 22, 217–259. Watson, R. B. (1945). On the probability of soldiers with Pacific Plasmodium vivax malaria infecting Anopheles quadrimaculatus. J. Natl. Malar. Soc. 4, 183–188. Wellems, T. E., and Plowe, C. V. (2001). Chloroquine-resistant malaria. J. Infect. Dis. 184, 770–776. Wilson, D. B. (1936). Rural hyper-endemic malaria in Tanganyika Territory. Trans. R. Soc. Trop. Med. Hyg. 29, 583–618. World Health Organization (1963). ‘‘Terminology of Malaria and of Malaria Eradication.’’ Geneva. World Health Organization (1969). ‘‘Parasitology of Malaria.’’ Technical Report 433. Geneva. World Health Organization (1981). Malaria parasite strain characterization, cryopreservation, and banking of isolates: A WHO memorandum. Bull. World Health Organ. 59, 537–548. Yorke, W., and Macfie, J. W. S. (1924). Observations on malaria made during treatment of general paralysis. Trans. R. Soc. Trop. Med. Hyg. 18, 13–33. Young, M. D. (1944). Studies on the periodicity of induced Plasmodium vivax. J. Natl. Malar. Soc. 3, 237–240. Young, M. D., and Burgess, R. W. (1948). Studies on imported malarias 9. The comparative susceptibility of Anopheles quadrimaculatus and Anopheles maculopennis freeborni to foreign vivax malarias. J. Natl. Malar. Soc. 7, 134–137. Young, M. D., Ellis, J. M., and Stubbs, T. H. (1947). Some characteristics of foreign vivax malaria induced in neurosyphilis patients. Am. J. Trop. Med. 27, 585–596. Young, M. D., Eyles, D. E., and Burgess, R. W. (1949). Studies on imported malarias. 10. An evaluation of the foreign malarias introduced into the United States by returning troops. J. Natl. Malar. Soc. 7, 171–185. Young, M. D., Contacos, P. G., Stitcher, J. E., and Millar, J. W. (1963). Drug resistance in Plasmodium falciparum from Thailand. Am. J. Trop. Med. Hyg. 12, 305–314. Yount, E. H., Jr., and Coggeshall, L. T. (1949). Status of immunity following cure of recurrent vivax malaria. Am. J. Trop. Med. Hyg. 29, 701–705. Zuckerman, A. (1964). The antigenic analysis of plasmodia. Am. J. Trop. Med. Hyg. 13, 209–213.
CHAPTER
2 Advances and Trends in the Molecular Systematics of Anisakid Nematodes, with Implications for their Evolutionary Ecology and Host–Parasite Co-evolutionary Processes Simonetta Mattiucci* and Giuseppe Nascetti†
Contents
1. Introduction 2. Molecular Systematics of Anisakid Nematodes 2.1. Current methods for anisakid identification 3. The Current Taxonomy 3.1. The current taxonomy of Anisakis 3.2. The current taxonomy of Pseudoterranova decipiens (sensu lato) 3.3. The current taxonomy of Contracaecum species from pinnipeds 4. Phylogenetic Systematics of Anisakid Nematodes 4.1. Genetic relationships between Anisakis spp. 4.2. Genetic relationships between Pseudoterranova spp. 4.3. Genetic relationships between Contracaecum spp.
49 51 54 55 55 76 83 91 92 95 96
* Department of Public Health Sciences, Section of Parasitology, ‘‘Sapienza’’—University of Rome, {
P.le Aldo Moro, 5, 00185 Rome, Italy Department of Ecology and Sustainable Economic Development—Tuscia University—Via S. Giovanni Decollato, 1, 01100 Viterbo, Italy
Advances in Parasitology, Volume 66 ISSN 0065-308X, DOI: 10.1016/S0065-308X(08)00202-9
#
2008 Elsevier Ltd. All rights reserved.
47
48
Simonetta Mattiucci and Giuseppe Nascetti
5. Genetic Differentiation in Anisakids 5.1. Genetic differentiation at interspecific level 5.2. Genetic differentiation at the intraspecific level and gene flow 6. Host–Parasite Cophylogeny 7. Host Preference, Ecological Niche and Competition 8. Anisakids as Biological Indicators 8.1. Anisakis spp. larvae as biological tags of fish stocks 8.2. Anisakids as indicators of trophic web stability and habitat disturbance of marine ecosystems 9. Conclusions and Identification of Gaps in Our Knowledge of Anisakids to be Filled by Future Research 9.1. Molecular systematics 9.2. Identification of human infections 9.3. Molecular ecology and life cycle 9.4. Host–parasite co-evolutionary aspects 9.5. Genetic variability of anisakids as an indicator of habitat disturbance Acknowledgements References
Abstract
100 100 106 108 114 119 120 123
130 131 132 132 133 134 137 137
The application of molecular systematics to the anisakid nematodes of the genera Anisakis, Pseudoterranova and Contracaecum, parasites of aquatic organisms, over the last two decades, has advanced the understanding of their systematics, taxonomy, ecology and phylogeny substantially. Here the results of this effort on this group of species from the early genetic works to the current status of their revised taxonomy, ecology and evolutionary aspects are reviewed for each of three parasitic groups. It has been shown that many anisakid morphospecies of Anisakis, Contracaecum and Pseudoterranova include a certain number of sibling species. Molecular genetic markers provided a rapid, precise means to screen and identify several species that serve as definitive and intermediate and or/paratenic hosts of the so far genetically characterized species. Patterns of differential distribution of anisakid nematodes in various definitive and intermediate hosts are presented. Differences in the life history of related species can be due both to differential host–parasite co-adaptation and co-evolution, and/or to interspecific competition, that can reduce the range of potential hosts in sympatric conditions. Phylogenetic hypotheses attempted for anisakid nematodes and the possible evolutionary scenarios that have been proposed inferred from molecular data, also with respect to the phylogeny of their hosts are presented for the parasite–host associations Anisakis-cetaceans and
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
49
Contracaecum-pinnipeds, showing that codivergence and hostswitching events could have accompanied the evolution of these groups of parasites. Finally, examples in which anisakid nematodes recognized genetically at the species level in definitive and intermediate/paratenic hosts from various geographical areas of the Boreal and Austral regions and their infection levels have been used as biological indicators of fish stocks and food-web integrity in areas at high versus low levels of habitat disturbance (pollution, overfishing, by-catch) are presented.
1. INTRODUCTION Adult nematodes of the genera of most species of the anisakid nematodes Anisakis, Dujardin, 1845, Pseudoterranova Krabbe, 1878 and Contracaecum Railliet and Henry, 1913 are parasites of the alimentary tract of aquatic vertebrates. They display indirect life cycles in aquatic ecosystems and involve various hosts at different levels in food webs. Marine mammals (cetaceans and pinnipeds) and fish-eating birds serve as definitive hosts; fish, squids and other invertebrates serve as intermediate or paratenic hosts; and crustaceans serve as first intermediate hosts. In humans, several larval anisakid nematodes cause the zoonotic disease, presently known as ‘anisakidosis’ or ‘anisakiosis’, when consumed in raw or undercooked fish. Anisakis is considered as the most important anisakid genus with regard to human infection (Audicana et al., 2002, and references therein), but some species of Pseudoterranova also have been implicated in human infections (Adams et al., 1997; Oshima, 1987), and a few species of Contracaecum are potentially infective (Vidal-Martinez et al., 1994). This zoonosis has a history starting from the first report (Van Thiel, 1960) of a larval nematode from herring in the gastro-intestinal tract of humans in the Netherlands. Some aspects of human anisakidosis have recently been reviewed by Audicana et al. (2002). Because of the existence of several excellent reviews by others authors on the history and clinical aspect of the anisakidosis (Audicana et al., 2002; Chai et al., 2005; Umehara et al., 2007), this introduction will not summarize the major events that mark the history of anisakid nematodes as causative agents of anisakidosis, but instead will highlight major landmarks involved in our understanding of their taxonomic status and ecology based on genetic markers. This field has had a major impact during the last 20 years on our knowledge of these parasites, including their host-specificity, geographical range and the possible identification of human cases of anisakidosis. Starting from the knowledge summarized in the exhaustive revision by Smith and Wootten (1978), this review treats the explosion in the
50
Simonetta Mattiucci and Giuseppe Nascetti
literature that has accompanied research activities on these nematodes and resulted in a series of discoveries over the last 25 years on the systematics and ecology of anisakid nematodes using molecular genetic tools applied to their taxonomy. Indeed, based on the advances in our understanding of the basic biology of Anisakis, reviewed by Smith and Wootten (1978), this introduction starts from their concluding remarks highlighting knowledge gaps and perspectives for investigations over the next 30 years. One major gap concerned the identification of species. Indeed, in the case of the species belonging to the genus Anisakis, they wrote: ‘. . . despite Davey’s revision (1971), there are still taxonomic problems, including the distinction between A. simplex and A. typica . . .. . . The generic diagnosis of Anisakis should be re-examined . . .. Anisakis larva (I) from North Atlantic waters has been cultured in vitro and shown to develop into A. simplex: There is need to culture the other ‘‘larval types’’ in order to confirm that they do, in fact, represent Anisakis and, if so, to determine which species they represent . . ..’. In 1978, Smith and Wootten underlined the major gap in our knowledge of nematode biology at that date, as represented by the genus Anisakis but also relevant to other anisakid species, that is, the identification of biological species. Indeed, species identification, based on morphological characters only, is difficult for adults, but even more difficult for larval stages. The need for species identification was especially important for larval stages of Anisakis because they have been implicated as causative agents of human anisakidosis. Indeed, Smith and Wootten (1978) concluded their review as follows: ‘. . . in these circumstances there is need for reliable methods of differential diagnosis. There is much scope for future investigation’. Thus, this ‘future investigation’ started with the application of molecular genetic methodologies in an attempt to address the systematics of these anisakids. The prospect of assessing anisakid nematode biodiversity based on molecular genetic markers as the preferred diagnostic tools seemed promising because the unambiguous identification of specimens causing human disease (anisakidosis) is essential for a proper epidemiological survey. The power of resolution of molecular genetic methodologies in detecting anisakid species revolutionized the taxonomy of these species during the following 25 years. Most descriptions of parasite species conformed with what can be regarded as the ‘morphological or typological, species concept’. Because genetic speciation is not always accompanied by corresponding morphological change, the actual number of biological species is likely to be greater than the current tally of nominal species, most of which are delineated based on morphological grounds. Detecting biological species of anisakid nematodes challenged parasitologists via an expected genetic variation and heterogeneity within the nominal species and has led to the definition of anisakid species according to the Mayr’s (Mayr, 1963) ‘biological species concept’ (BSC).
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
51
The aim of this review is to gather together the many and varied aspects concerning the biology and evolutionary ecology of those anisakid species detected genetically and recognized as members of Anisakis, Pseudoterranova and Contracaecum, including (a) species presently accepted as senior synonyms based on the application of different molecular genetic markers; (b) current molecular genetic approaches to identify anisakid species; (c) ecological evidence relating to the geographical distribution of detected species, their host preferences and life cycles; (d) the use of anisakid nematodes as biological tags for fish stock identification in a multidisciplinary approach; (e) correlation between the values of genetic variability and the levels of parasitic infection in definitive and intermediate/paratenic hosts as indicators of the integrity of marine food webs; (f) estimates of genetic divergence at intraspecific and interspecific levels; and (g) estimates of their genetic relationships based on different clustering approaches inferred from different molecular genetic data sets and their use to infer phylogenetic hypothesis and possible co-evolutionary events with their definitive hosts.
2. MOLECULAR SYSTEMATICS OF ANISAKID NEMATODES The nematode superfamily Ascaridoidea contains 52 genera; species are mainly parasites of the alimentary tract of vertebrates. Several are of medical and veterinary concern, and some are of economic significance. The classification scheme for members of the Anisakidae proposed by Hartwich (1974) is based largely on features of the ‘excretory system’ and alimentary tract. According to the ‘systematic keys’ of Hartwich (1974), the family Anisakidae contains the subfamilies Anisakinae and Contracaecinae. The Anisakinae includes (cf. Hartwich, 1974) the genera Anisakis and Pseudoterranova plus several others. The Contracaecinae includes three genera, Contracaecum, Phocascaris and Galeiceps. Species of Anisakis, Pseudoterranova, Contracaecum and Phocascaris have aquatic life cycles and homeothermic final hosts. Anisakid nematodes have been extensively studied with respect to their alpha-taxonomy (Berland, 1961, 1964; Bruce and Cannon, 1990; Davey, 1971; Deardoff and Overstreet, 1979; Fagerholm, 1989, 1991; Fagerholm and Gibson, 1987; Gibson, 1983; Osche, 1963; Sprent, 1977, 1978, 1979, 1983) and life cycles (Huizinga, 1967; Klo¨ser et al., 1992; Kie and Fagerholm, 1995). Much of the controversy in the systematics of anisakid species, when dealing with identifications based solely on morphological differences, revolved around their confused taxonomic position. Indeed, morphological characters of taxonomic significance in this group are very few (i.e. features of the excretory system, alimentary canal, number and
52
Simonetta Mattiucci and Giuseppe Nascetti
distribution of male caudal papillae, position of the vulva and length of the spicules) and are applicable to adult specimens only. Furthermore, these are often only relevant to male individuals, making identification difficult for many worms at the species level. In recent decades, molecular genetic techniques have been used to demonstrate that structural features are not always adequate for recognizing ‘true’ species within Anisakis, Pseudoterranova and Contracaecum. In the last two decades, the diversity of anisakid species has increased due to the detection by genetic markers of several sibling species with reproductively isolated gene pools that are morphological very similar, and thus correspond to the ‘biological species’. There are now morphospecies, or species complexes, based on previously recognized cosmopolitan species (sensu lato), that may comprise several recognized species. This solved one of the major problems in the systematics of anisakid nematodes: the occurrence of the parallelism and convergence of morphological features, which confound the systematic value of some morphological criteria and often accompany a high genetic and ecological divergence between the species. The lack of morphological differences in these parasites may be due to factors such as similar selection pressures causing the conservation of morphology; consequently, some morphological characters have little or no taxonomic value because of the evolutionary co-adaptation of these endoparasites to the stable habitat represented by their definitive hosts. Indeed morphospecies may appear to have multiple host species, that is, parasite populations isolated in their hosts have diverged genetically but have conserved morphological features. Moreover, species identification based on morphological characters only makes identification very difficult, especially for larval stages that lack reliable diagnostic features at the species level. Thus, the inconsistency in morphological characteristics of anisakid nematodes impeded the development of a credible scheme for their taxonomy. This prompted the need to classify these nematodes using genetic and/or biochemical methods. The assessment of anisakid nematodes biodiversity based on molecular genetic markers as preferable tools for specific diagnosis was an important prospect because the unambiguous identification of those anisakids with a zoonotic potential is an essential requirement for a proper epidemiological survey. Initial attempts to apply population genetics to the study of genetic variation among large samples of anisakids collected from different intermediate/paratenic and definitive hosts in nature employed the use of multilocus allozyme electrophoresis (MAE) (19–24 enzyme loci). These tools revealed the existence of high genetic heterogeneity within certain anisakid morphospecies, such as those of Anisakis, Pseudoterranova and Contracaecum (Bullini et al., 1986; Mattiucci et al., 1986, Nascetti et al., 1986;
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
53
Paggi et al., 1991). The species concept (BSC) (Mayr, 1963) was well supported by the application of allozyme markers for several anisakid species. Indeed, after the first such application, the diversity of species belonging to Anisakis, Pseudoterranova and Contracaecum quickly increased after the detection of several sibling species and leads to the discovery and description of several new species. Reproductive isolation and absence of gene flow were demonstrated by these allozymes between sympatric and allopatric sibling species, establishing their specific status (Mattiucci et al., 1997, 2001, 2003, 2005; Nascetti et al., 1993; Paggi et al., 1991). Allozyme markers have allowed us to (a) genetically characterize different species of anisakid nematodes, (b) estimate their genetic differentiation, (c) establish their genetic relationships, and (d) identify their larval stages which lack morphological characters (Arduino et al., 1995; Bullini et al., 1986, 1994, 1997; Mattiucci and Nascetti, 2006; Mattiucci et al., 1986, 1997, 1998, 2001, 2002a, 2003, 2004, 2005, 2006, 2007a, 2008a,b,c; Nascetti et al., 1986, 1993; Orecchia et al., 1986a, b, 1994; Paggi and Bullini, 1994; Paggi et al., 1991, 1998b, 2000, 2001). The introduction of the polymerase chain reaction (PCR)-derived molecular methodologies later confirmed taxonomic decisions involving anisakid species previously based on allozyme markers. Reference individuals initially characterized by allozymes have been used to develop DNA-based approaches for species identification, such as PCR-RFLP and direct sequencing of ITS rDNA or mitochondrial DNA. Ascaridoid classifications and inferred patterns of character evolution have been investigated previously, and some evolutionary hypotheses for representative ascaridoids, some of which are anisakids, have been proposed based on phylogenetic methods, including ribosomal DNA sequence data and comparative analysis of morphological and life-history characters (Nadler, 1992, 1995, 2005; Nadler and Hudspeth, 1998; Zhu et al., 2000a). Results from these studies identified some consistent, putative, shared-derived morphological features, strongly suggesting that some morphological features represent ancestral states or highly homoplastic characteristics (Nadler and Hudspeth, 1998). Phylogenetic analysis indeed provided a new perspective for the delimitation of anisakid sibling species, including hierarchical relatedness and relative rates of evolution. An evolutionary perspective provides a conceptual approach to view species as independent evolutionary lineages and provides another approach for delimitating species (Adams, 1998; Nadler, 2002, 2005). Indeed, based on phylogenetic DNA analysis, sibling anisakid species have been confirmed by methods that can test the hypothesis of lineage independence analysing many individual specimens and sometimes detecting new genotypes and species (D’Amelio et al., 2000, 2007; Mattiucci et al., 2008b, c; Nadler et al., 2000; 2005; Valentini et al., 2006).
54
Simonetta Mattiucci and Giuseppe Nascetti
2.1. Current methods for anisakid identification Except for some morphological differences between anisakid species (Mattiucci et al., 1998, 2005, 2008c; Paggi et al., 1998b, 2000), all the species genetically characterized are to date indistinguishable at all developmental stages (larvae and adults); consequently, only molecular genetic markers can be used reliably to identify these species. Of all the recently developed tools, and despite the useful PCR-derived molecular markers, allozymes continue to remain the best methodological approach for demonstrating reproductive isolation between anisakids. Starting from the study of genetic variation at several enzyme loci, the main argument in favour of using allozymes is the ability to test (using the Hardy– Weinberg equilibrium) for reproductive isolation of anisakid populations among large numbers of individuals. Accordingly, allozymes provide the best choice for determining molecular genetic markers applied to the systematics of anisakids and for confirming the BSC. Based on allozymes diagnostic for different anisakid taxa, easy and rapid identification of large numbers of individuals can be performed; this method is particularly valuable for identifying larval individuals collected from several intermediate/paratenic hosts and often in mixed infections. Accordingly, such identifications have been demonstrated to be very informative tools for answering epidemiological questions involving geographical range, host preference and life cycles. Moreover, because numerous allozymes (20–24 enzyme loci) have been applied to thousands of anisakid individuals, they have contributed greatly to our knowledge of the genetic diversity of anisakid populations collected from various ecosystems in the Boreal and Austral hemispheres (see also Section 8.2). However, the tool is limited to frozen-preserved or fresh individuals. This constraint has been resolved by DNA-based diagnostic techniques, which have the advantage of also being able to use alcohol or formalin-preserved specimens. In contrast with allozymes, the DNAbased techniques have increased our ability to study phylogenetic relationships between related anisakids based on the evolutionary lineage concept (Adams, 1998). The PCR-DNA molecular derived methodologies so far applied to the systematics of anisakid nematodes include PCRrestriction fragment length polymorphism (PCR-RFLPs of ITS-DNA) (D’Amelio et al., 2000; Kijewska et al., 2002; Pontes et al., 2005), singlestrand conformational polymorphism SSCP-DNA (ITS) of PCR products (Zhu et al., 1998, 2000b, 2007), direct sequencing of PCR-amplified DNA of 28S (LSU) and complete internal transcribed spacer (ITS-1, 5.8S, ITS-2) ribosomal DNA (Hu et al., 2001; Li et al., 2005; Nadler et al., 2000, 2005; Zhu et al., 1998, 2000b, 2001, 2002), mitochondrial cytochromoxidase b (mtDNA cytb) (Mattiucci et al., 2003) and mitochondrial cytochromoxidase 2 (mtDNA cox2) sequence analyses (Mattiucci and Nascetti, 2006;
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
55
Mattiucci et al., 2008b, c; Valentini et al., 2006). PCR-DNA markers have so far confirmed the existence of the sibling species previously detected by allozymes, establishing their taxonomic status. The only exception is represented by the two Antarctic sibling species of the Contracaecum osculatum complex (i.e. C. osculatum D and C. osculatum E), where PCR-DNA markers based on ITS-rDNA (Zhu et al., 2000b) and three mitochondrial DNA genes (i.e. mtDNA cox1, ssrRNA and lsrDNA) (Hu et al., 2001) were unable to identify species previously detected by allozyme markers. In contrast, the existence of the two sibling species, C. osculatum D and C. osculatum E, first detected by allozymes, was confirmed by sequencing the mtDNA cox2 gene (Mattiucci et al., 2008b). A scheme of allozymes and PCR-derived methods which can be used routinely to identify a single individual belonging to genetically characterized species and gene pools of anisakids as belonging to the genera Anisakis, Pseudoterranova and Contracaecum is provided in Tables 2.1–2.3, respectively.
3. THE CURRENT TAXONOMY This section summarizes the current taxonomy of anisakid species of the genera Anisakis, Pseudoterranova and Contracaecum (here considering only those species maturing in pinnipeds) which have been genetically characterized to date. They can be recognized, at any life-history stage, by different molecular genetic approaches, as reported in Tables 2.1–2.3. A synopsis of each recognized anisakid species, including data on both the definitive and intermediate hosts and the geographical range, is also presented.
3.1. The current taxonomy of Anisakis The taxonomy of Anisakis species has traditionally relied on adult morphology. According to Davey (1971), the primary characters are the length and shape of the ventriculus, length and shape of the male spicules, and arrangement of the male caudal papillae. According to Berland (1961), larval morphological features (i.e. length of ventriculus and presence/ absence of caudal spine) could distinguish Anisakis type I and Anisakis type II larvae. In the revision by Davey (1971), the generic diagnosis for Anisakis states: ‘three lips each bearing a bilobed anterior projection which carries the single dentigerous ridge; interlabia absent; excretory gland with duct opening between ventrolateral lips; oesophagus with anterior muscular portion (proventriculus) and posterior ventriculus, the latter being oblong and sometimes sigmoid or else as broad as long; no oesophageal appendix or intestinal caecum; vulva in middle or first third of the body, spicules of male unequal; preanal papillae numerous; postanal papillae including a group of three or four
56 TABLE 2.1 Molecular genetic markers for the identification of the species of Anisakis Molecular genetic marker
Method
Identified speciesa
References
22 allozyme loci 20 allozyme loci 25 allozyme loci 20 allozyme loci 5.8S rDNA, ITS rDNA 18S and 5.8S rDNA, ITS rDNA 22 allozyme loci
MAE MAE MAE MAE PCR-RFLP; SSCP PCR-RFLP
Nascetti et al., 1986 Mattiucci et al., 1997 Paggi et al., 1998b Mattiucci et al., 1998 Zhu et al., 1998 D’Amelio et al., 2000
21 allozyme loci
MAE
20 allozyme loci
MAE
18S, 28S and 5.8S rDNA, ITS rDNA ITS rDNA
PCR and sequencing
Asstr, Ape Asstr, AsC, Ape Azi, Ape, AsC, Aph Asstr Asstr Ape, AsC, Aty, Azi, Aph, Abr, Asc Abr, Aph, Azi, Ape, AsC, Aph Aty, Asstr, Ape, Azi, Ape, AsC, Aph Apa, Asstr, Ape, Aty, Az, Aph, Abr AsC, Ape, Asp Asstr, Ape, Aty, Azi, Aph, AspA
Pontes et al., 2005
MAE
PCR-RFLP
Mattiucci et al., 2001 Mattiucci et al., 2002a Mattiucci et al., 2005 Nadler et al., 2005
a
20 allozyme loci
MAE
Mattiucci and Nascetti, 2006 Valentini et al., 2006
PCR-RFLP
Asp, Asstr, Ape, AsC, Aty, Azi, Aph, Ab, Apa Asstr, Ape, AsC, Aty, Azi, Aph, Abr, Apa, Asp Asstr, Ape
Mitochondrial cytochrome c-oxidase subunit 2 (mtDNA cox2) ITS rDNA, 5.8S rDNA, mitochondrial cytochrome c-oxidase subunit 1 (mtDNA cox1) ITS rDNA, 5.8S rDNA Mitochondrial cytochrome c-oxidase subunit 1 (mtDNA cox1), NADH dehydrogenase subunit 1 ITS-2 rDNA ITS rDNA
PCR and sequencing
PCR-RFLP PCR and sequencing
Asstr, Ape Asstr
Abe et al., 2006 Cross et al., 2007
PCR-SSCP Multiplex-PCR
Ape, Aty Asstr, Ape, Aph
Zhu et al., 2007 Umehara et al., 2008
Umehara et al., 2006
Codes for Anisakis spp.: Asstr, A. simplex (s.s.); Ape, A. pegreffii; AsC, A. simplex C; Aty, A. typica; Azi, A. ziphidarum; Aph, A. physeteris; Abr, A. brevispiculata; Apa, A. paggiae; Asp, Anisakis sp.; Asc, A. schupakovi; AspA, Anisakis sp. A.
57
58
Simonetta Mattiucci and Giuseppe Nascetti
TABLE 2.2 Molecular genetic markers for the identification of the Pseudoterranova decipiens species complex Molecular genetic marker
Identified speciesa
References
MAE
Pdss, Pkr, Pbu
Paggi et al., 1991
MAE
Pca
16 allozyme loci
MAE
20 allozyme loci ITS rDNA
MAE
Pdss, Pkr, Paz, Pbu, PdE Pbu, Paz, Pdss, Pkr Pdss, Pkr, Paz, Pbu, Pca, PdCa1 Pdss, Pkr, Paz, Pbu, Pca, PdCa1
George-Nascimento and Llanos, 1995; George-Nascimento and Urrutia, 2000 Bullini et al., 1997
16 allozyme loci 9 allozyme loci
18S, 28S and 5.8S rDNA, ITS rDNA a
Method
PCR-SSCP
PCR and sequencing
Mattiucci et al., 1998 Zhu et al., 2002
Nadler et al., 2005
Codes for Pseudoterranova spp.: Pdss, P. decipiens (s.s.) (=P. decipiens B); Pkr, P. krabbei (=P. decipiens A); Pbu, P. bulbosa (=P. decipiens C); Paz, P. azarasi (=P. decipiens D); PdE, P. decipiens E; Pca, P. cattani; PdCa1, P. decipiens from Chaenocephalus aceratus.
pairs set close to the tip of the tail on ventral side’. After a critical revision of the 21 included species, Davey (1971) concluded that many of the species had to be considered as junior synonyms of three main species, the only ones accepted in his revision, that is, A. simplex (Rudolphi, 1809, det. Krabbe, 1878) with 10 synonyms, A typica (Diesing, 1860) with one synonym and A. physeteris (Baylis, 1923) with three synonyms. He also retained four others as species inquirendae because of the lack of sufficient data: A. dussurmierii (van Beneden, 1870) reported from a dolphin in the Indian Ocean, A. schupakovi Mozgovoi, 1951 from the Caspian seal, Pusa caspica (later redescribed and accepted by Delyamure et al., 1964), A. alexandri Hsu¨ and Hoeppli, 1933 from Sotalia sinensis and A. insignis (Diesing, 1851) from Inia geoffrensis in South America rivers (redescribed and accepted by Petter, 1972). According to Davey’s revision, the only morphological characters of systematic value in recognizing the taxonomic status of the species of Anisakis were the length and shape of ventriculus, length and shape of the
TABLE 2.3
Molecular genetic markers for the identification of the species of Contracaecum and Phocascaris from pinnipeds
Molecular genetic marker
Method
Identified speciesa
References
17 allozyme loci 24 allozyme loci
MAE MAE
Nascetti et al., 1993 Orecchia et al., 1994
25 allozyme loci
MAE
20 allozyme loci 28S rDNA
MAE PCR and sequencing
ITS rDNA ITS rDNA 18S and 28S rRNA, mitochondrial cytochrome c-oxidase subunit 1 (mtDNA cox1) 18 allozyme loci
PCR-SSCP PCR-RFLP; PCR-SSCP PCR-SSCP
CoA, CoB, Coss CoD, CoE, CoA, CoB, Coss Cra, CoA, CoB, Coss, CoD, CoE CoA, CoB CoA, CoB, Coss, Cba, Crad, Cmir, Pcy, Pph CoA, CoB, Coss, Cbai Cogm CoA, CoB, Coss, Cbai
MAE
Cogm, Cmar
Arduino et al., 1995 Mattiucci et al., 1998 Nadler et al., 2000 Zhu et al., 2000 Zhu et al., 2001 Hu et al., 2001
Mattiucci et al., 2003 (continued)
TABLE 2.3 (continued)
a
Molecular genetic marker
Method
Identified speciesa
References
18S, 28S and 5.8S rDNA, ITS rDNA
PCR and sequencing
Nadler et al., 2005
20 allozyme loci
MAE
Mitochondrial cytochrome c-oxidase subunit 2 (mtDNA cox2)
PCR and sequencing
CoA, CoB, Coss, Crad, Cmir, Cbai, Cmar, Cogm, Pcys, Pph CoA, CoB, Coss, CoD, CoE, Cbai, Crad, Cmir, Cmar, Cogm, Pcys CoA, CoB, Coss, CoD, CoE, Cbai, Crad, Cmir, Cmar, Cogm, Pcys
Mattiucci et al., 2008b
Mattiucci et al., 2008b
Codes for Contracaecum spp.: CoA, C. osculatum A; CoB, C. osculatum B; Coss, C. osculatum (s.s.) (=C. osculatum C); CoD, C. osculatum D; CoE, C. osculatum E; Cbai, C. baicalensis; Cmar, C. margolisi; Crad, C. radiatum; Cogm, C. ogmorhini (s.s.); Cmir, C. mirounga; Pph, Phocascaris phocae; Pcys: Ph. Cystophorae.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
61
male spicules, arrangement of the male caudal papillae and position of the vulva in females. Morphological features such as the length of the ventriculus and presence/absence of caudal spine were proposed for the recognition of larval stages of Anisakis, but these are too inconsistent and inaccurate for use at the specific level (Berland, 1961; Shiraki, 1974). Despite Davey’s valuable discussion, when considering the main morphological characters of the species Anisakis, instability of their systematics, including their identification at the species level (especially in the case of females and larvae), remained. The resolution of some of the taxonomic issues, highlighted in this critical revision, has been achieved during the last 20 years facilitated by the adoption of new methodologies, allozymes first and DNA-based molecular methodologies later. Starting from that critical revision, and based on our growing understanding on the systematic of species Anisakis, we can state that Davey’s revision failed in considering only three species as valid and several species as synonyms, as some of the latter have been subsequently validated by genetic methodology (Mattiucci et al., 2001). However, this revision anticipated, in some way, a future cladistic analysis by indicating morphological characters likely to be useful for the systematics of the species belonging to Anisakis following their genetic identification. In addition, in his revision, Davey did not consider Skryabinisakis Mosgovoi, 1951, proposed for the species A. physeteris, as an accepted genus. However, the high level of genetic differentiation of both A. physeteris, A. brevispiculata and A. paggiae (see Mattiucci et al., 2005) from those species of Anisakis studied genetically using different nuclear and mitochondrial markers (Mattiucci and Nascetti, 2006; Mattiucci et al., 2001; Valentini et al., 2006) indicates that Anisakis is indeed polyphyletic and highly heterogeneous. Today, the existence of two main clades within Anisakis is clearly inferred from the phylogenetic analysis based on nuclear data sets from allozymes and mtDNA cox2 sequence analysis of all the genetically characterized species (Mattiucci and Nascetti, 2006; Mattiucci et al., 2005; Valentini et al., 2006). One clade encompasses species with the larval stage known as Anisakis type I (sensu Berland, 1961) and the second sharing the larval morphology of Anisakis type II (sensu Berland, 1961) (see Mattiucci et al., 2005, 2007a; Orecchia et al., 1986a). The first clade includes the species of the A. simplex complex [i.e. A. simplex (sensu stricto), A. pegreffii and A. simplex C], A. typica, A. ziphidarum and Anisakis sp. The second includes the species A. physeteris, A. brevispiculata and A. paggiae (see Mattiucci et al., 2005; Valentini et al., 2006). The clade including A. simplex (s.s.) and A. simplex C and A. pegreffii is also well supported by a phylogenetic analysis inferred from ITS rDNA sequence data sets (Nadler et al., 2005). This analysis is congruent with both allozymes and mtDNA cox2 analysis in depicting A. physeteris + A. brevispiculata as the sister group to the remaining Anisakis spp.
62
Simonetta Mattiucci and Giuseppe Nascetti
3.1.1. Anisakis spp. included in clade I According to the genetic data, six species can currently be included in this clade: three species of the A. simplex complex [i.e. A. simplex (s.s.), A. pegreffii and A. simplex C], A. typica, A. ziphidarum and a new gene pool referred to as Anisakis sp. (see Valentini et al., 2006, unpublished data). As far as is known, all of these species have the larval form known as type I (sensu Berland, 1961). A synopsis of the ecological aspects of each species, including their known geographical distribution and both definitive and intermediate/paratenic hosts, is presented below. A. simplex (Rudolphi, 1809) (sensu stricto) (see Nascetti et al., 1986): A. simplex (s.s.) is widespread between 35 N and the Arctic Circle; it is present in both the western and eastern Atlantic and Pacific Oceans (Abe et al., 2005, 2006; Abollo et al., 2001; Mattiucci et al., 1997, 1998; Nadler et al., 2005; Paggi et al., 1998a; Umehara et al., 2006, 2008) (Fig. 2.1). The southern limit of this species in the north east Atlantic Ocean is the waters around the Gibraltar area. A. simplex (s.s.) is also occasionally present in western Mediterranean waters due to the migration of pelagic fish species into the Alboran Sea from the Atlantic (Mattiucci and Nascetti, 2006; Mattiucci et al., 2004, 2007a). A. simplex (s.s.) has been so far genetically recognized from nine species of cetacean hosts. Several squid and fish species have been found harbouring larvae of this species throughout its geographical range (Table 2.5). A sympatric area between A. simplex (s.s.) and A. pegreffii has been identified along the Spanish and Portuguese Atlantic coasts (Abollo et al., 2001; Marques et al., 2006; Mattiucci et al., 1997, 2004, 2007a; Pontes et al., 2005), in the Alboran Sea (Mattiucci et al., 2004, 2007a) and recently also in Japanese Sea waters (Umehara et al., 2006). A. simplex (s.s.) also occurs with A. simplex C in the eastern Pacific Ocean, where it has been identified in definitive and intermediate/paratenic hosts (Mattiucci et al., 1997, 1998, unpublished data; Paggi et al., 1998c) (Tables 2.4 and 2.5; Fig. 2.1). Although it has sympatric and syntopic occurrences in mixed infections at both larval and adult stage with other Anisakis species (Mattiucci et al., 2004, 2005, 2007a), reproductive isolation between A. simplex (s.s.) and both A. pegreffii and A. simplex C was proved by the lack of adult F1 hybrids and/or backcross genotypes clearly demonstrated at the nuclear level by allozyme markers (Mattiucci et al., 1997, 2005). Only a few F1 hybrid larval individuals A. pegreffii–A. simplex (s.s.), of the thousands identified, were detected by allozymes in some fish host from the sympatric area off the Atlantic Iberian coast (Mattiucci et al., 2004). However, back-crossing of F1 hybrids with parental species has not been detected. Based on PCR-RFLP of the ITS, Umehara et al. (2007) recognized A. simplex (s.s.) as the main source of infections in humans in Japan.
60⬚
i olar C Ar tic P
rcle
30⬚
0⬚
30⬚
Antarc
60⬚
tic Po
60⬚
120⬚ A. pegreffii
A. simplex (s.s. )
C. osculatum (s.s.) P. decipiens (s.s.) Ph. phocae
A. simplex C
C. osculatum A P. krabbei
A. typica
C. osculatum B
P. bulbosa
P. azarasi
A. physeteris
C. o. baicalensis P. decipiens E
0⬚ A. brevispiculata C. osculatum D
lar Cir
cle
60⬚ A. ziphidarum C. osculatum E
120⬚ A. paggiae C. radiatum
180⬚
Anisakis sp. C. mirounga
C. ogmorhini (s.s)
C. margolisi
P. cattani
Ph. cystophorae
FIGURE 2.1 World map showing the so far known distribution areas of anisakid species of Anisakis (□), Pseudoterranova (△), Contracaecum (○) and Phocascaris (?). The geographical areas indicated are related to the sampling localities for their definitive and intermediate hosts.
64
TABLE 2.4
Definitive hosts so far detected, by molecular genetic markers, for the Anisakis spp.
Cetaceans Balaenopteridae Balaenoptera acutorostrata Delphinidae Delphinus delphis Globicephala melaena Globicephala macrorhynchus Lagenorhynchus albirostris Lissodelphis borealis Orcinus orca Pseudorca crassidens Stenella coeruleoalba Tursiops truncatus Sotalia fluviatilis Stenella attenuata Stenella longirostris Steno bredanensis
A. simplex (s.s.)
A. pegreffii A. simplex C A. typica
A. ziphidarum
Anisakis sp. A. physeteris
A. brevispiculata A. paggiae
NEA
–
–
–
–
–
–
–
–
IC IC, SA
IC, WM –
– SA
– –
– –
– –
– –
– –
– –
–
–
–
FL
–
–
–
–
–
NEA
–
–
–
–
–
–
–
–
–
–
NEP
–
–
–
–
–
NEP NEP
–
– NEP
–
– –
– –
– –
– –
– –
IC
WM
–
EM
–
–
–
–
–
–
CM, SA
–
FL, CS
–
–
–
–
–
– – –
– – –
– – –
BR FL, CS BR
– – –
– – –
– – –
– – –
– – –
–
–
–
CS
–
–
–
–
–
Kogiidae Kogia breviceps Kogia sima Monodontidae Delphinapterus leucas Neobalaenidae Caperea marginata Phocoenidae Phocoena phocoena Physeteridae Physeter catodon Ziphiidae Mesoplodon densirostris Mesoplodon europaeus Mesoplodon grayi Mesoplodon mirus Ziphius cavirostris
– –
– –
– –
– –
– –
– –
– –
SA, IC, FL –
SA, FL FL
NWA
–
–
–
–
–
–
–
–
–
SA
–
–
–
–
–
–
–
NEP
–
–
–
–
–
–
–
–
–
–
–
–
–
–
CM
–
–
–
–
–
–
SA
–
–
–
–
–
–
–
–
CS
–
–
–
–
– –
– –
– –
– –
– –
SA SA, NZ
– –
– –
– –
–
–
–
–
CM, SA
–
–
–
–
65
Sampling locality codes: AZ: Azores Islands; BE: Bering Sea; BR: Brazil Atlantic coast; BS: Barents Sea; CM: Central Mediterranean Sea; CS: Caribbean Sea; EM: East Mediterranean Sea; FA: Falkland Islands; FL: Florida coast; IC: Iberian Atlantic Coast; JA: Japan Sea; MA: Mauritanian coast; MD: Madeira Island; NAM: North African Mediterranean coast; NEA: NorthEast Atlantic; NEP: North-East Pacific; NWA: North-West Atlantic; NZ: New Zealand; PC: Portuguese coast; SA: South Africa coast; SC: Somali coast; SI: off Sakhalin Islands; TA: Tasman Sea; WM: West Mediterranean Sea (data from Mattiucci and Nascetti, 2006, 2007; Mattiucci et al., 1986, 1997, 2001, 2002, 2004, 2005; Nadler et al., 2005; Nascetti et al., 1986; Paggi et al., 1998a,b,c). Hosts listed by alphabetical order of the family.
66
TABLE 2.5
Intermediate/paratenic hosts so far detected, by molecular genetic markers, for the Anisakis spp.
Cephalopods Sepiidae Sepia officinalis Ommastrephidae Todaropsis eblanae Todarodes sagittatus Todarodes angolensis Illex coindettii Fishes Belonidae Belone belone Bothidae Arnoglossus laterna Arnoglossus imperialis Bramidae Brama brama Carangidae Trachurus capensis Trachurus mediterraneus Trachurus picturatus Trachurus trachurus
Selar crumenophthalmus
A. simplex (s.s.) A. pegreffii A. simplex C A. typica
Anisakis A. ziphidarum sp.
A. physeteris
A. brevispiculata A. paggiae
IC
–
–
–
–
–
–
–
–
IC, SA IC – IC
IC, SA NAM, CM SA –
– – – –
– – – –
– – – –
– – – –
– CM – –
– – – –
– – – –
IC
IC
–
–
–
–
–
–
–
PC –
– PC
– –
– –
– –
– –
– –
– –
– –
–
SA
–
–
–
–
–
–
–
– –
SA CM
– –
– –
– –
– –
– –
– –
– –
AZ, MD
AZ, MD
AZ, MD
–
NEA, IC, MA, WM –
CM, EM, WM, IC, MA –
–
EM
–
AZ
CM
–
–
–
CHS
–
–
–
–
–
Citharidae Citharus linguatula Clupeidae Clupea harengus
PC
NEA, BS, NEP Etrumeus whiteheadi – Congridae Conger conger IC Astroconger – myriaster Coryphaenidae Coryphaena – hippurus Emmelicththydae Emmelicththys – nitidus nitidus Engraulidae Engraulis – encrasicolus Gadidae Boreogadus saida IC Micromesistius IC poutassou Gadus morhua BS, IC Theragra NEP, JA, BE chalcogramma Trisopterus luscus IC Gempylidae Thyrsites atun –
PC
–
–
–
–
–
–
–
–
–
–
–
–
–
–
–
SA
–
–
–
–
–
–
–
CM CHS
– –
– –
– –
– –
– –
– –
– –
–
–
SC
–
–
–
–
–
SA
–
–
–
–
–
–
–
CM
–
–
–
–
–
–
–
– CM, IC, NAM – –
– –
– –
– –
– –
– –
– –
– –
– –
– –
– –
– –
– –
– –
– –
–
–
–
–
–
–
–
–
–
SA
–
–
–
–
–
–
67
(continued)
68
TABLE 2.5
(continued)
Hexagrammidae Pleurogrammus azonus Lophiidae Lophius piscatorius Lophius vomerinus Lotidae Molva dypterygia Brosme brosme Merluccidae Merluccius capensis Merluccius hubbsi Merluccius merluccius
Muraenidae Muraena helena Moridae Pseudophycis bachus Nemipteridae Nemipterus virgatus Nemipterus bathybius Ophidiidae Genypterus capensis
A. simplex (s.s.) A. pegreffii A. simplex C A. typica
Anisakis A. ziphidarum sp.
A. physeteris
A. brevispiculata A. paggiae
NEA
–
–
–
–
–
–
–
–
IC –
NAM SA
– –
– –
– –
– –
– –
– –
– –
IC NEA
– –
– –
– –
– –
– –
– –
– –
– –
– – NEA, IC, MA
SA FA CM, EM, WM, IC, NEA, MA
– – –
– – MA, EM, NAM
– – MA
– – IC
– – CM, MA, WM, IC, EM, NAM
– – MA
– – IC
–
NAM
–
–
–
–
–
–
–
–
NZ
NZ
–
–
–
–
–
–
– –
– –
– –
CHS CHS
– –
– –
– –
– –
– –
–
SA
–
–
–
–
–
–
–
Osmeridae Hypomesus pretiosus japonicus Phycidae Phycis phycis Phycis blennoides Pinguipedidae Parapercis colias Pleuronectidae Hippoglossus hippoglossus Platichthys flesus Salmonidae Oncorhynchus gorbuscha Oncorhynchus keta Salmo salar Scophtalmidae Scomberesocidae Scomberesox saurus Scombridae Lepidorhombus boscii Scomber japonicus Scomber scombrus
JA
–
–
–
–
–
–
–
–
– –
NAM –
– –
NAM –
– –
– –
NAM NAM
– –
– –
–
NZ
NZ
–
–
–
–
–
–
BE
–
–
–
–
–
–
–
–
–
–
–
PC
–
–
–
–
–
SI
–
–
–
–
–
–
–
–
SI NWA
– –
– –
– –
– –
– –
– –
– –
– –
NWA
–
–
–
–
–
–
–
–
IC
IC, NAM
–
–
–
–
–
–
–
AZ, MD NAM
AZ, MD –
– –
MD NAM
– –
– –
AZ, MD, JA AZ, MD, JA – NEA, IC, CM, IC – NAM
(continued)
69
70
TABLE 2.5
(continued)
Thunnus thynnus Auxis thazard Euthynnus affinis Sarda orientalis Scomberomorus commerson Scorpaenidae Scorpaena scrofa Sebastidae Helicolenus dactylopterus Soleidae Dicologlossa cuneata Solea senegalensis Sparidae Spondyliosoma cantharus Sternoptychidae Maurolicus muelleri Trachichthyidae Hoplostethus atlanticus Hoplostethus mediterraneus Trachinidae Echiichthys vipera Trichiuridae Lepidopus caudatus
A. simplex (s.s.) A. pegreffii A. simplex C A. typica
Anisakis A. ziphidarum sp.
A. physeteris
A. brevispiculata A. paggiae
JA – – – –
CM, BR – – – –
– – – – –
BR BR SC SC SC
– – – – –
– – – – –
– – – – –
– – – – –
– – – – –
IC
IC
–
–
–
–
–
–
–
–
CM, SA
–
–
–
–
–
–
–
– PC
PC –
– –
– –
– –
– –
– –
– –
– –
IC
–
–
–
–
–
–
–
–
NEA
–
–
–
–
–
–
–
–
–
–
TA
–
–
–
–
–
–
–
CM
–
–
–
–
–
–
–
–
NAM
–
–
–
–
–
–
–
–
CM, SA
–
–
–
–
–
–
–
Aphanopus carbo Trichiurus lepturus Triglidae Eutrigla gurnardus Xiphiidae Xiphias gladius
MD –
MD NAM
–
–
MD
AZ
AZ
AZ
AZ
IC
–
–
–
–
–
–
–
–
NEA
CM
–
CA
CA
–
CM, IC, NEA, CA, EM
AZ, CA
AZ
Sampling locality codes: AZ: Azores Islands; BE: Bering Sea; BR: Brazil Atlantic coast; BS: Barents Sea; CHS: China Sea; CA: Central Atlantic Ocean; CM: Central Mediterranean Sea; CS: Caribbean Sea; EM: East Mediterranean Sea; FA: Falkland Islands; FL: Florida coast; IC: Iberian Atlantic Coast; JA: Japan Sea; MA: Mauritanian coast; MD: Madeira Island; NAM: North African Mediterranean coast; NEA: North-East Atlantic; NEP: North-East Pacific; NWA: North-West Atlantic; NZ: New Zealand; PC: Portuguese coast; SA: South Africa coast; SC: Somali coast; SI: off Sakhalin Islands; TA: Tasman Sea; WM: West Mediterranean Sea (data from: Abollo et al., 2001; Farjallah et al., 2008; Klimpel et al., 2007; Marques et al., 2006; Mattiucci and Nascetti, 2006, 2007; Mattiucci et al., 1986, 1997, 2001, 2002a, 2004, 2005, 2007a; Nascetti et al., 1986; Orecchia et al., 1986a; Paggi et al., 1998a, b, c; Pontes et al., 2005; Umehara et al., 2006, 2008; Zhu et al., 2007). Hosts listed by alphabetical order of the family.
71
72
Simonetta Mattiucci and Giuseppe Nascetti
A. pegreffii Campana-Rouget and Biocca, 1955: Previously indicated as A. simplex A (see Nascetti et al., 1986), A. pegreffii is the dominant species of Anisakis in the Mediterranean Sea, being widespread in all the fish species. Indeed, it is presently the most important anisakid nematode in several pelagic and demersal fish from Mediterranean waters (Farjallah et al., 2008; Mattiucci et al., 1997, 2007a; Paggi et al., 1998a). It is also widely distributed at both adult and larval stage in the Austral Region between 30 N and 55 S (Mattiucci et al., 1997). In Atlantic waters, the northerly limit of its geographical range is represented by the Iberian coast (Abollo et al., 2001; Marquez et al., 2006; Mattiucci et al., 1997, 2004, 2007a; Pontes et al., 2005), and has not so far been reported from the western Atlantic (our unpublished data). It has been detected, using the PCR-RFLP of the ITS region, at the larval stage in some fish hosts from Japanese waters (Abe et al., 2006; Umehara et al., 2006, 2008) (Table 2.5; Fig. 2.1). SSCPbased identification of A. pegreffii larvae in fish from China waters, using genetic markers in the ITS-2 rDNA was also reported (Zhu et al., 2007) (Table 2.5; Fig. 2.1). To date, it has been recorded as an adult in three species of dolphins, belonging to the family Delphinidae, and in several species of fish and three squids as a larva (Tables 2.4 and 2.5). Among these, two definitive and sixteen intermediate/paratenic hosts were found to be shared with A. simplex (s.s.) in the contact area between the two species [Iberian Atlantic coast, western Mediterranean Sea (Alboran Sea) and Japan Sea waters] (Fig. 2.1). Whereas, two definitive and few intermediate/paratenic hosts are shared by A. pegreffii and A. simplex C in the Austral waters off New Zealand, the South African coast and the southern Chilean coast (Table 2.5; Fig. 2.1). Based on PCR-RFLP of the ITS, D’Amelio et al. (1999) and direct sequencing of the mtDNA cox2 (Mattiucci et al., 2007b), Anisakis larvae removed by endoscopy from humans in Italy were recognized as belonging to A. pegreffii. A. simplex C of Mattiucci et al. (1997): A. simplex C currently exhibits a discontinuous range, including the Canadian and Chilean Pacific coasts, New Zealand waters and the South African Atlantic coast (Mattiucci et al., 1997, unpublished data; Nadler et al., 2005). This species has been identified at the adult stage in cetaceans and as a larva it occurs syntopically with A. pegreffii in some fish species (Tables 2.4 and 2.5; Fig. 2.1). It has been occasionally identified also in Mirounga leonina from sub-Antarctic area (our unpublished data) and in M. angustirostris from North-East Pacific Ocean (Nadler et al., 2005). A. typica (Diesing, 1860): According to the data from the A. typica populations so far detected genetically, its range extends from 30 S to 35 N in warmer temperate and tropical waters (Tables 2.4 and 2.5; Fig. 2.1) (Mattiucci et al., 2002a). In these areas it was found by genetic markers as an adult in dolphin species and as a larva in several fish
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
73
species (Tables 2.4 and 2.5). A. typica has also been identified in the striped dolphin, Stenella coeruleoalba, and in the European hake, Merluccius merluccius, from the eastern Mediterranean Sea (off Cyprus). Its presence in these waters could be the result of the ‘Lesseptian migration’ (through the Suez Canal) (Mattiucci et al., 2004) of its intermediate/paratenic hosts from the Indian Ocean. It was recently recognized using RFLPPCR of the ITS rDNA in the flatfish Platichthys flesus captured in central Portuguese waters of the NE Atlantic Ocean (Marques et al., 2006), and only rarely in some fish caught along the North African coast of the Mediterranean Sea (Farjallah et al., 2008). SSCP-based identification of Anisakis spp. larvae, using genetic markers in the ITS-2 rDNA, detected A. typica larvae in fish from China waters (Zhu et al., 2007) (Table 2.5; Fig. 2.1). Anisakis sp. 1: Recently, a new gene pool, referred to as Anisakis sp. 1 was genetically detected by both allozyme and mtDNA analysis, at the larval stage, as a parasite of the fish Nemipterus japonicus caught off the Malaysian coast (Berland and Mattiucci, unpublished data). This new taxon is genetically distinct from all the known species of Anisakis but most closely related to A. typica from central Atlantic waters. Although known only at the larval stage, the third-stage larva of this undescribed taxon is a type I larva (sensu Berland, 1961). The preliminary results appear to indicate that this taxon may be a sibling species of A. typica occurring in central Pacific waters (Berland and Mattiucci, unpublished data). A. ziphidarum Paggi, Nascetti, Webb, Mattiucci, Cianchi and Bullini, 1998: A. ziphidarum was first described, both genetically and morphologically, as an adult in the beaked whales Mesoplodon layardii and Ziphius cavirostris from the South Atlantic Ocean (off the South African coast). Subsequently, it has also been recorded in the Mediterranean Sea, also in Z. cavirostris. Since its first morphological description and genetic characterization (Paggi et al., 1998b), it has recently been identified genetically as an adult in other species of beaked whale, such as M. mirus and M. grayi, in South Atlantic waters and in Mesoplodon sp. and Z. cavirostris in Caribbean waters (Mattiucci and Nascetti, 2006). Thus, its geographical range appears to be wide (Fig. 2.1) and related to that of its definitive hosts. Only scanty data are available concerning its infection in fish and/or squid, but it is responsible for a low prevalence of infection in some fish species, such as in Merluccius merluccius (see Mattiucci et al., 2004) and Aphanopus carbo (see Pontes et al., 2005; Saraiva et al., 2007) in central Atlantic waters. However, it seems that this species may involve other intermediate hosts, such as squid, rather than fish in its life cycle, as these represent the main food source of beaked whales. Anisakis sp. of Valentini et al. (2006): This species has been detected only at a larval stage (L4) in the beaked whales Mesoplodon mirus and M. grayi from South African and New Zealand waters (Mattiucci and
74
Simonetta Mattiucci and Giuseppe Nascetti
Nascetti, 2006) (Fig. 2.1; Table 2.4). The gene pool was found to be reproductively isolated from the sympatric species A. ziphidarum occurring in the same hosts and geographical region. It is genetically very distinct from the other species of Anisakis, but is most closely related to A. ziphidarum (see Section 5.1; Table 2.8). The third-stage larva of this undescribed taxon is apparently of type I and has on rare occasions been identified in the fishes (see Mattiucci et al., 2007a; Saraiva et al., 2007) caught in North East Atlantic waters. Conversely, this species has been genetically identified, at the larval stage, heavily infecting the squid Moroteuthis ingens in Tasman Sea waters (our unpublished data). This appears to support the hypothesis that this species involves squid rather than fish in its life cycle. Recently, Pontes et al. (2005) detected the occurrence of a new taxon at larval stage in Aphanopus carbo and Scomber japonicus from Madeira waters (Atlantic Ocean) indicated as Anisakis sp. A, genetically closely related to A. ziphidarum. However, the exact correspondence between this taxon and Anisakis sp. of Valentini et al. (2006) has not yet been assessed.
3.1.2. Anisakis spp. included in clade II
Three species and one new gene pool of Anisakis so far comprises this clade, as clearly demonstrated by allozymes (Mattiucci et al., 2005) and mtDNA cox2 sequence analysis (Valentini et al., 2006). They are A. physeteris, A. brevispiculata and A. paggiae, which represent a complex of sibling species readily recognized genetically at both nuclear and mitochondrial levels. The existence of A. physeteris clustering with A. brevispiculata in the same clade, as a sister group to the other species included in the clade I, was also supported by rDNA ITS sequence phylogenetic analysis (Nadler et al., 2005) (see also Section 4.1). As far as is known, all these species share at the third larval stage the morphology known as type II (sensu Berland, 1961). A summary of each follows, including a synopsis of some ecological aspects of each species in relation to their known geographical distribution and both definitive and intermediate/paratenic hosts (Tables 2.4 and 2.5). A. physeteris (Baylis, 1920): A population of this species was first genetically characterized in its main definitive host, the sperm whale, Physeter macrocephalus, from Mediterranean waters (Mattiucci et al., 1986); no genetically identified adults have been recorded in other cetacean hosts. Type II larvae of A. physeteris have been genetically identified, occurring rarely in only a very few fish host species (Mattiucci et al., 1986, 2001, 2004), except for the swordfish, Xiphias gladius, from Mediterranean and Atlantic waters in which it represents the main Anisakis species (Mattiucci et al., 2007a) (Tables 2.4 and 2.5; Fig. 2.1). Despite the fact that the swordfish might only represent an accidental host in the life cycle of
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
75
Anisakis spp., acquiring the infection by preying on infected invertebrates or squid (main prey of the swordfish), suggests that other intermediate hosts, mainly squids rather than fish are involved in the life cycle of this parasite. A. brevispiculata Dollfus, 1966: A population of A. brevispiculata was initially characterized genetically using allozymes based on material from a pygmy sperm whale, Kogia breviceps, in South African and North East Atlantic waters (Iberian coast) (Mattiucci et al., 2001) (Table 2.4; Fig. 2.1). Reproductive isolation from the morphologically closely related A. physeteris was demonstrated, establishing the validity of A. brevispiculata (see Mattiucci et al., 2001), which has been synonymized with A. physeteris by Davey (1971). Later, the same species was sequenced at the mtDNA cox2 gene and its genetic relationships with respect to the other Anisakis spp. was established, confirming that A. brevispiculata clusters well with those Anisakis species forming the second clade, as also indicated by nuclear markers (allozymes). A congruent result was inferred from the ITS rDNA sequence analysis (Nadler et al., 2005) (see Section 4.1). Anisakis larvae of type II corresponding to A. brevispiculata were recognized by allozyme markers as a rare parasites of the fish Merluccius merluccius (see Mattiucci et al., 2004, 2007a) and the swordfish Xiphias gladius in Atlantic waters (Mattiucci et al., 2007a). A. paggiae Mattiucci et al. (2005): In the cluster formed by A. physeteris and A. brevispiculata, this third species has been recently demonstrated by both allozymes (Mattiucci et al., 2005) and mtDNA cox2 sequence analysis (Valentini et al., 2006) (Fig. 2.1). A. paggiae was first genetically characterized and described morphologically as an adult parasite of the pygmy sperm whale, Kogia breviceps, and the dwarf sperm whale, K. sima, from off both Florida and the South African Atlantic coast (Mattiucci et al., 2005). Scanty data are so far available regarding the identification of the intermediate hosts in the life cycle of A. paggiae. A very few larvae of type II have been identified as belonging to these species in fish from Atlantic waters (i.e. M. merluccius and X. gladius) (see Mattiucci et al., 2007a, unpublished data), thus suggesting that other hosts, not yet detected, are involved in the life cycle of this Anisakis species. Anisakis sp. 2: A further gene pool, referred to as Anisakis sp. 2, has been detected genetically by means of allozyme markers and mtDNA cox2 sequence analysis based on larvae of type II from the swordfish X. gladius in the equatorial area (Mattiucci et al., 2007a, unpublished data). The new taxon was shown to be genetically distinct from all the other Anisakis species, and that it is mostly closely related to A. physeteris. Preliminary phylogenetic analysis showed that it clusters with the clade formed by A. physeteris, A. brevispiculata and A. paggiae, thus suggesting that this new gene pool might represent an undescribed sibling species belonging to this same complex.
76
Simonetta Mattiucci and Giuseppe Nascetti
The high genetic heterogeneity of the Anisakis spp. is now also supported by some differential morphological features detected in the species belonging to this genus, where the two major clades can be delineated as follows: (i) the ventriculus, in the adult stage, is short, never sigmoid and broader than long in A. physeteris, A. brevispiculata and A. paggiae (see Mattiucci et al., 2005), and longer than broad and often sigmoid in shape in those species included in clade I (the species of the A. simplex complex, A. typica and A. ziphidarum) (see Mattiucci et al., 2005; Paggi et al., 1998b); (ii) male spicules that are short, stout and of similar length can be observed in A. physeteris, A. brevispiculata and A. paggiae (Mattiucci et al., 2005) but are long and often unequal (equal in A. ziphidarum; see Paggi et al., 1998b) in species of clade I; and (iii) type II larval morphology (sensu Berland, 1961) is characteristic of A. physeteris, A. brevispiculata, A. paggiae and Anisakis sp. 2 (Mattiucci et al., 2001, 2004, 2005, 2007a) (clade II), whereas a type I morphology (sensu Berland, 1961), can be found in all the species of the A. simplex complex, A. typica, A. ziphidarum and Anisakis sp. and Anisakis sp. 1 (clade I). While no morphological characters are so far known which help in distinguishing the sibling species of the A. simplex complex, some morphological features, of diagnostic value, available in male and female adult specimens, were used to help in distinguishing A. paggiae from A. physeteris and A. brevispiculata (see Mattiucci et al., 2005). Indeed a morphological key to the recognized adults of Anisakis spp. so far included in clade II was provided by Mattiucci et al. (2005).
3.2. The current taxonomy of Pseudoterranova decipiens (sensu lato) The species decipiens was first described as Ascaris decipiens by Krabbe (1878). A. decipiens was later linked with Terranova Leiper and Atkinson, 1914 (a genus erected for parasites of elasmobranchs) by Baylis (1916), because of the presence of an intestinal caecum. Subsequently, Baylis linked this species within Porrocaecum Railliet and Henry, 1912 (a group of ascarids now mainly restricted to terrestrial birds), because he considered it as a senior synonym of Terranova. Terranova was resurrected at full generic level by Johnston and Mawson (1945) and as a subgenus by Karokhin (1946), the latter being the first to use the combination T. decipiens. T. decipiens was accepted by Mozgovoi (1951, 1953), Hartwich (1957) and Yamaguti (1961). However, because Terranova included species maturing in different definitive hosts (elasmobranchs, reptiles, marine mammals), Myers (1959) erected Phocanema (with decipiens as the type and only species) for those species included in Terranova which were parasites of marine mammals on the base of some morphological features at the cephalic end (structure of the labia) and the male caudal end.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
77
Accordingly, the species decipiens and Terranova kogiae Johnston and Mawson, 1945 (described from the pygmy sperm whale, Kogia breviceps) were included in Phocanema. Later, Gibson (1983) considered Phocanema as a synonym of Pseudoterranova, previously erected by Mozgovoi (1951; listed in Mozgovoi, without indication) for parasites from sperm whales, and decipiens was transferred to this genus (combination first used in Gibson and Colin, 1982). According to the same authors, the group of species of Pseudoterranova from marine mammals included P. kogiae (Johnston and Mawson, 1945), P. ceticola (Deardoff and Overstreet, 1981) and P. decipiens. This is the status quo which has been widely accepted. Although several nominal species related to Pseudoterranova decipiens (i.e. occurring in seals and possessing only an intestinal caecum) having been described, these species have tended to be considered as synonyms of P. decipiens. This species appeared to have a low host specificity, with up to 18 seal species been recorded as definitive hosts (e.g. Hartwich, 1975; King, 1983; Myers, 1959), and exhibited some morphological variation between specimens collected from the bearded seal, Erignathus barbatus, in Canadian waters (Baylis, 1916). Population genetic analysis, first performed by allozyme markers, on specimens of P. decipiens (s.l.) recovered from 10 fish species, and from four seal species, collected at several locations of the North Atlantic Ocean from the Canadian Atlantic eastwards to the Barents Sea, demonstrated the existence of a remarkable genetic heterogeneity with striking variation in allele frequencies among the samples (Paggi et al., 1991). Indeed, the observed genotype frequencies at various loci showed significantly deviations from those expected using the Hardy–Weinberg equilibrium, with a complete absence or marked deficiency of various heterozygous classes in material recovered from several sites and also between worms recovered from one host individual. The only possible explanation was that three distinct biological species occurred sympatrically in the samples of P. decipiens (s.l.) collected in seals hosts from those geographical areas, with no gene flow between them: the three taxa genetically recognized were thus provisionally designated as P. decipiens A, P. decipiens B and P. decipiens C (see Paggi et al., 1991). Morphological analysis carried out on male specimens identified by allozyme markers as P. decipiens A and B allowed the detection of significant differences in a number of characters between these two members; on the basis of such differences the nomenclature designation for P. decipiens A and P. decipiens B was proposed (see Paggi et al., 2000). The names Pseudoterranova krabbei Paggi et al., 2000 and P. decipiens (s.s.) were proposed, respectively, for species A and B, and a formal description of the two taxa was provided (see Paggi et al., 2000). The name of P. bulbosa (Cobb, 1888) was proposed for the taxon P. decipiens C (see Mattiucci et al., 1998), as the latter taxon was demonstrated
78
Simonetta Mattiucci and Giuseppe Nascetti
to correspond morphologically with Ascaris bulbosa described by Cobb (1888) from the collection of Dr. Kukenthal collected from the bearded seal, Erignathus barbatus, at Spitzbergen (NE Atlantic Ocean). Furthermore, a sample of P. decipiens C from that same locality was studied both genetically and morphologically by Paggi et al. (1991, 2000). Previously, Ascaris bulbosa had been considered a synonym of Phocanema decipiens [later included in Pseudoterranova by Gibson and Colin (1982)] by Myers (1959). Therefore, Mattiucci et al. (1998) proposed the name P. bulbosa (Cobb, 1888) n. comb. for P. decipiens C. A further taxon, provisionally designated as P. decipiens D (see Mattiucci et al., 1998), was later included in the P. decipiens complex; this was detected by exhibiting several fixed differences at some enzyme loci with respect to P. decipiens A, P. decipiens B and P. decipiens C. It was found to occur sympatrically with P. bulbosa in the same geographical areas (Japanese waters) and occasionally in the same definitive host, the bearded seal Erignathus barbatus, from which it was demonstrated to be reproductively isolated (Mattiucci et al., 1998). P. decipiens D was found to correspond to the measurements and tail drawing of Porrocaecum azarasi Yamaguti and Arima, 1942, a species described by Yamaguti and Arima (1942) based on specimens recovered in the ribbon seal Phoca (=Histriophoca) fasciata on the islands of Sakhalin and Hokkaido. This taxon was synonymized by Margolis (1956) with ‘Phocanema decipiens’. Therefore, Mattiucci et al. (1998) proposed the name Pseudoterranova azarasi (Yamaguti and Arima, 1942) n. comb. for species P. decipiens D. Using allozyme markers on larval and adult population of P. decipiens (s.l.) collected from four fish species and the Austral fur seal, Otaria byronia, in the SE Pacific Ocean, a further member of the P. decipiens complex has been shown to exist (George-Nascimento and Llanos, 1995). In the formal description, this taxon was named P. cattani GeorgeNascimento and Urrutia, 2000. A summary of the sibling species P. decipiens (s.l.) recognized by using several molecular genetic methodologies are reported below with ecological data on their host preference (Table 2.5) and geographical range (Fig. 2.1). P. decipiens (Krabbe, 1868) (sensu stricto) (=P. decipiens B): This species was initially recognized as P. decipiens B by Paggi et al. (1991); its formal description was later given based on morphological features considered to be of diagnostic value with respect to the other members included in the P. decipiens complex (see Paggi et al., 2000). Its geographical range appears to be wide and mainly in the Arctic and sub-Arctic regions, including the North East Atlantic (comprising Scottish, Faroes, southern Icelandic and Norwegian waters), the Canadian Atlantic (including Newfoundland waters and the Gulf of S. Lawrence) (Brattey and Stenson, 1993; Paggi et al., 1991) and the Canadian Pacific waters (Mattiucci et al.,
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
79
1998; Nadler et al., 2005; Paggi et al., 1998c). Areas of sympatry between P. decipiens (s.s.) and other species of the P. decipiens complex have been detected; in the North Eastern Atlantic waters in mixed infections with both P. krabbei and P. bulbosa and in Canadian Atlantic waters with P. bulbosa (see Paggi et al., 1991) (Fig. 2.1). P. decipiens (s.s.) was recognized, using genetic markers, as a parasites at the adult stage in the common seal, Phoca vitulina, grey seal, Halichoerus grypus, in both North and West Atlantic waters, and in Phoca vitulina richardsii and Zalophus californianus in northern Pacific waters (Paggi et al., 1998c). Its larval forms have been identified in some gadoid fish species of the North Atlantic Ocean (Mattiucci et al., 1998; Paggi et al., 1991) (Table 2.6). P. krabbei Paggi, Mattiucci et al., 2000: This species was previously referred to as P. decipiens A (Paggi et al., 1991). To date it has been found only in the North-East Atlantic (including off Scotland, the Faroes, southern Iceland and Norway) (Fig. 2.1). It is mainly an adult parasite of the grey seal, Halichoerus grypus, and the common seal, Phoca vitulina, often in mixed infections in the same individual host with P. decipiens (s.s.) (Table 2.6). However, in the eastern Atlantic waters, when detected in sympatry with P. decipiens (s.s.), P. krabbei prevails in the grey seal rather than in the common seal. The ecological significance of this finding is discussed in Section 7. P. krabbei has been recognized genetically at the larval stage as a parasite of Gadus morhua, Melanogrammus aeglefinus and Pollachius virens (see Paggi et al., 1991) (Table 2.6; Fig. 2.1). P. krabbei exhibits, in male specimens with respect to P. decipiens (s.s.), the following diagnostic morphological features: shorter spicules; a proximal papilla (nomenclature according to Fagerhom, 1989) smaller than d1 versus the same papilla size in P. decipiens (s.s.); distal papillae 1, 2 and 4 closer to each other; and caudal plates of a similar width and narrower than in P. decipiens (s.s.) where plates (wp) 1 (wp1) and 2 (wp2) are of a similar width but plate 3 (wp3) is narrower (Paggi et al., 2000). P. bulbosa (Cobb, 1888): Previously referred to as P. decipiens C (Paggi et al., 1991) (see Mattiucci et al., 1998), this species has been recorded from the Barents and Norwegian Seas, the Canadian Atlantic and the Sea of Japan, between 40 N and 80 N (Brattey and Stenson, 1993; Mattiucci et al., 1998; Paggi et al., 1991). Its main definitive host so far detected is the bearded seal, Erignathus barbatus. In this phocid host from the Otaru Sea (Sea of Japan), P. bulbosa has been identified as occurring in mixed infections with P. azarasi (see Mattiucci et al., 1998). The benthic flatfishes Hippoglossoides platessoides and Reinhardtius hippoglossoides are reported as its intermediate hosts (Paggi et al., 1991) (Table 2.6; Fig. 2.1). P. bulbosa differs from the other members of the P. decipiens complex in having the following morphological features in adult male specimens: longer spicules; caudal plates (wp) of unequal width, with plate 2 (wp2)
80 TABLE 2.6 Definitive and intermediate/paratenic hosts so far detected, by molecular genetic markers, for the Pseudoterranova decipiens species complex
Pinnipeds Otariidae Eumetopias jubatus Otaria byronia Zalophus californianus Phocidae Phoca vitulina richardsii Phoca vitulina Erignathus barbatus Halichoerus grypus Cystophora cristata Mirounga angustirostris Leptonychotes weddellii Fishes Channichthydae Chaenocephalus aceratus
P. krabbei
P. decipiens (s.s.)
P. bulbosa
P. azarasi
P. decipiens E
P. cattani
–
–
–
–
–
– –
– NEP
– –
NWP, JA – NWA
– –
SEP –
– NEA
– –
NWA –
– –
– –
–
NEP NEA, NWA, NEP –
–
NEA, NWA NWA, NEA NEP –
NWP, JA – – – –
–
NEA – – –
NEP, NEA, LS – – – –
– – – AN
– – – –
–
–
–
–
AN
– (continued)
TABLE 2.6 (continued)
Cottidae Myxocephalus scorpius Myxocephalus quadricornis Gadidae Gadus morhua macrocephalus Gadus morhua Gadus ogac Pollachius virens Melanogrammus aeglefinus Boreogadus saida Lotidae Brosme brosme Merluccidae Merluccius gayi Ophidiidae Genypterus maculatus Osmeridae Osmerus eperlanus
P. krabbei
P. decipiens (s.s.)
P. bulbosa
P. azarasi
P. decipiens E
P. cattani
– –
NWA –
– NWP
– BE, NWP
– –
– –
–
NWA
NWP
–
–
NEA – FI FI
NEA NWA NWA –
– – – –
JA, NWP – – – –
– – – –
– – – –
–
NWA
–
–
–
–
–
NEA
–
–
–
–
–
–
–
–
–
SEP
–
–
–
–
–
SEP
–
NEA
–
–
–
–
81
(continued)
TABLE 2.6 (continued)
Notothenidae Notothenia coriiceps Notothenia neglecta Trematomus newnesi Paralichthydae Paralichthys microps Pleuronectidae Hippoglossus hippoglossus Hippoglossoides platessoides Reinhardtius hippoglossoides Scophthalmidae Psetta maxima
P. krabbei
P. decipiens (s.s.)
P. bulbosa
P. azarasi
P. decipiens E
P. cattani
– – –
– – –
– – –
– – –
AN AN AN
– – –
–
–
–
–
–
SEP
–
–
NWP
–
–
NEA
NEA
BS
BE, NWP –
–
–
–
–
NWA
–
–
–
NEA
–
–
–
–
–
Codes: AN: Antarctica; BE: Bering Sea; BS: Barents Sea; FI: Faeroe Islands; JA: Japan Sea; LS: Labrador Sea; NEA: North-East Atlantic; NEP: North-East Pacific; NWA: NorthWest Atlantic; NWP: North-West Pacific; SEP: South-East Pacific (Chilean coast) (data from George-Nascimento and Llanos, 1995; George-Nascimento and Urrutia, 2000; Mattiucci et al., 1998; Nadler et al., 2005; Paggi et al., 1991, 1998c; Zhu et al., 2002). Hosts listed by alphabetical order of the family.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
83
narrower than 1(wp1) and 3 (wp3); and a different pattern of distal caudal papillae, with d2 far apart from d4 (Mattiucci et al., 1998). P. azarasi (Yamaguti and Arima, 1942): This species was previously referred to as P. decipiens D (see Mattiucci et al., 1998). Its main definitive host has been identified as the Steller’s sea lion, Eumetopias jubatus. Its geographical distribution appears limited to Japanese and Sakhalinese waters of the North Pacific Ocean (Mattiucci et al., 1998) (Table 2.6; Fig. 2.1). Both P. bulbosa and P. azarasi were resurrected, on both allozyme and morphological bases (see Mattiucci et al., 1998), from synonymy with P. decipiens (see Margolis, 1956; Myers, 1959). P. azarasi differs from the other members of the P. decipiens complex in having shorter spicules; caudal plates (wp) of increasing width from 1 to 3; and a different pattern of distal papillae (sensu Fagerholm, 1989), with d2 closer to d4 than in P. bulbosa but more distant than in P. krabbei and P. decipiens (s.s.) (see Mattiucci et al., 1998). P. decipiens E of Bullini et al., 1997. This taxon was genetically detected in the Antarctic Weddell seal, Leptonychotes weddellii (see Bullini et al., 1997). Scanty data are so far available in relation to the nature of its intermediate host, although larvae corresponding to this sibling species were found as parasites of benthic fish hosts in subantartic waters (see Section 7.2) (Table 2.6). P. cattani George-Nascimento and Urrutia, 2000: As stated above, this species was found as an adult in Otaria byronia on the Chilean coast (Table 2.5; Fig. 2.1). Using molecular markers in the internal transcribed spacers of ribosomal DNA (ITS-rDNA), this species was recently shown to cluster with the P. decipiens complex (Zhu et al., 2002) (Section 4.2).
3.3. The current taxonomy of Contracaecum species from pinnipeds Contracaecum was originally defined by Railliet and Henry (1912) based on the morphology of the oesophago-intestinal region. C. spiculigerum (Rudolphi, 1809) (=Ascaris spiculigerum, Rudolphi, 1809) was designated as the type-species. The original type-material was later identified as C. microcephalum (Rudolphi, 1809) by Hartwich (1964), and this species was accordingly defined as the new type-species of the genus. Presently, the genus comprises some 50 nominal species, most of which are adults in pinnipeds and fish-eating birds. Hartwich (1975) defined the genus based on morphological details; important features are the presence of interlabia and the position of the excretory pore at the base of the ventral interlabium. However, several later authors suggested that the genus should be divided or amended. Mozgovoi (1951) had previously divided the genus
84
Simonetta Mattiucci and Giuseppe Nascetti
into three subgenera: Contracaecum (Erschovicaecum) [clearly representing Hysterothylacium (see Deardoff and Overstreet, 1980) and including species maturing in fish], Contracaecum (Contracecum) and Contracaecum (Ornitocaecum). These two new subgenera were considered as synonyms of Contracaecum by Hartwich (1975). Phocascaris Ho¨st, 1932 included the following definition ‘interlabia present, reduced or absent . . . . parasites of the digestive tract of seals’, P. phocae being the type-species of the genus. According to that definition, Berland (1964), when describing P. cystophorae from the seal Cystophora cristata, suggested that the species of Contracaecum ‘with opposite caeca from seals’ should be transferred to the genus Phocascaris. His proposal was to retain within Contracaecum species from fish-eating birds. Berland’s (1964) hypothesis was mainly based on the life-cycle patterns of these anisakid nematodes and their definitive hosts, irrespective of their morphological phenotype. However, a taxonomic proposal based on those biological features was not generally accepted by taxonomists, and Hartwich (1975) did not take into account that observation when he produced the ‘Key of Ascaridoidea’, still considering a morphological character (relating to the presence/absence of interlabia) as a valid criterion for distinguishing the species belonging to Contracaecum and Phocascaris. Several years later, the same morphological character was shown to be inconsistent and not of useful systematic value for differentiating the two groups of species included in Contracaecum. Indeed, based on allozymes (22 enzyme loci), Orecchia et al. (1986b) and Nascetti et al. (1990) used species Contracaecum from seals and fish-eating birds to show that of all the species of the Contracaecum osculatum complex (see Section 4.3) are genetically much more closely related to species of Phocascaris (i.e. Phocascaris phocae and P. cystophorae) from seals than to the congeneric species from fish-eating birds, demonstrating high levels of differentiation (Contracaecum species from pinnipeds share no alleles in common with those from fish-eating birds). Clustering methods based on allozyme markers showed that P. phocae and P. cystophorae form a clade with the species of Contracaecum from seals (Nascetti et al., 1990) (Section 4.3), suggesting an evolutionary hypothesis for the systematic status of these species. Nadler et al. (2000), based on nuclear rDNA sequence data of several taxa of the genera Contracaecum and Phocascaris, demonstrated the validity of the evolutionary hypothesis previously suggested by allozyme markers. A phylogenetic hypotheses based on different clustering analyses of the ITS-rDNA (Nadler et al., 2000) and the mtDNA cytochromoxidase-2 (mtDNA cox2) (Mattiucci et al., 2008b) sequences data strongly supported the hypothesis based on allozymes, according to which species of Phocascaris are nested within the clade formed by the Contracaecum species hosted by phocid seals and are thus closely related to species of the Contracaecum osculatum complex. All of the phylogenetic analyses also support the hypothesis
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
85
advanced by Berland (1964), based on the biology of these anisakid nematodes, in terms of the monophyly of the species of Contracaecum and Phocascaris from phocids (see also Section 4.3). On the other hand, the molecular genetic approach has been demonstrated to be very fruitful in the detection of sibling species and disclosing new taxa of Contracaecum parasites of fish-eating birds (Bullini et al., 1986; D’Amelio et al., 1990, 2007; Li et al., 2005; Mattiucci et al., 2002b, 2008c), which are not however treated in details in this review.
3.3.1. The Contracaecum osculatum species complex
To date, five members of the C. osculatum complex have been recognized genetically by the use of allozyme markers. They are the three Arctic sibling species referred to as C. osculatum A, C. osculatum B and C. osculatum (s.s.) (see Nascetti et al., 1993), and the two Antarctic members, C. osculatum D and C. osculatum E (see Orecchia et al., 1994). C. osculatum A of Nascetti et al. (1993): This species occurs in the Norwegian and Barents Seas, Canadian Atlantic, Icelandic, Canadian Pacific waters and the Sea of Japan, between 40 N and 80 N. Its main definitive host is the bearded seal, Erignathus barbatus, in both eastern and western part of the North Atlantic Ocean, but the grey seal, Halichoerus grypus, has also been recorded as a host (Brattey and Stenson, 1993; Nascetti et al., 1993). Subsequently, it has been detected genetically in the Steller’sea lion, Eumetopias jubatus, in the Western Pacific Ocean (Sea of Japan) (Mattiucci et al., 1998; Paggi et al., 1998c). Its larvae have been identified in the gadoid fish Theragra chalcogramma from the Sea of Japan (Mattiucci et al., 1998) (Table 2.7; Fig. 2.1). C. osculatum B of Nascetti et al. (1993): This species was first detected genetically as an adult in the phocid seals Pagophilus groenlandicus, Phoca vitulina and Halichoerus grypus from the North-eastern and north-western Atlantic Ocean (Brattey and Stenson, 1993; Nascetti et al., 1993); reproductive isolation was demonstrated by allozyme markers between this species and the other two Arctic members of the C. osculatum complex (Nascetti et al., 1993). Later, it was identified based on diagnostic allozyme markers as a parasite of the Phoca vitulina and Zalophus californianus in northern Pacific waters (Mattiucci et al., 1998) (Table 2.7; Fig. 2.1). C. osculatum (Rudolphi, 1802) (sensu stricto): This species, named C. osculatum C by Nascetti et al. (1993), was genetically characterized from the grey seal, Halichoerus grypus; its reproductive isolation from C. osculatum B was shown in the sympatric situation of individual seal hosts (i.e. the grey seal) from Iceland waters (Nascetti et al., 1993). It is the only species of the C. osculatum complex present in the Baltic Sea. So far, this species has not been reported from the western part of the Atlantic or from Pacific waters (Table 2.7; Fig. 2.1).
86
TABLE 2.7
Definitive and intermediate/paratenic hosts so far detected, by molecular genetic markers, for the Contracaecum spp. from pinnipeds
Pinnipeds Phocidae Phoca vitulina
C. osculatum A
C. osculatum B
C. osculatum (s.s.)
C. osculatum D
C. osculatum E
C. o. baicalensis
C. ogmorC. radiatum C. mirounga hini (s.s.)
C. margolisi
–
NWA, NEP, JA, BE NEA, NWA, JA, BS – BS
–
–
–
–
–
–
–
–
–
–
–
–
–
–
–
–
– –
– –
– –
BL –
– –
– –
– –
– –
Pagophilus groenlandicus
–
Phoca sibirica Erignathus barbatus
– NEP, NWA, NEA, JA, BS –
–
–
–
–
–
–
AN
–
–
–
NEP
–
–
–
–
–
–
–
–
NWA, NEA
NWA, NEA
–
–
–
–
–
–
–
–
–
BS, NEA, NWA, BA –
RS, WS
RS, WS
–
RS, WS
–
–
–
–
NWA
–
–
–
–
–
–
–
–
–
–
–
–
–
–
–
–
Mirounga leonina Eumetopias jubatus Halichoerus grypus Leptonychotes weddellii Cystophora cristata Otariidae Zalophus californianus
NEP
Arctocephalus australis Arctocephalus pusillus Fishes Bathydraconidae Gymnodraco acuticeps Cygnodraco mawsonii Channichthydae Cryodraco antarcticus Chionodraco hamatus Pagetopsis macropterus Chaenodraco wilsoni Cottidae
–
–
–
–
–
–
–
AR
AR
–
–
–
–
–
–
–
–
–
SA, AU
–
–
–
–
RS
RS
–
–
–
–
–
–
–
–
RS
RS
–
–
–
–
–
–
–
–
RS
RS
–
RS
–
–
–
–
–
–
RS
RS
–
RS
–
–
–
–
–
–
RS
RS
–
–
–
–
–
–
–
–
RS
RS
–
–
–
–
–
–
–
BS
–
Myoxocephalus quadricornis – Gadidae Gadus morhua macrocephalus Theragra chalcogramma Notothenidae
–
–
–
–
–
BE
–
–
–
–
–
–
–
–
–
BE
–
–
–
–
–
–
–
–
–
–
–
–
RS
RS
–
–
–
–
–
87 (continued)
TABLE 2.7
(continued) C. osculatum A
C. osculatum B
C. osculatum (s.s.)
C. osculatum D
C. osculatum E
C. o. baicalensis
C. ogmorC. radiatum C. mirounga hini (s.s.)
C. margolisi
–
–
–
RS
RS
–
–
–
–
–
–
–
–
RS
RS
–
–
–
–
–
BE
–
–
–
–
–
–
–
–
–
Notothenia neglecta Trematomus bernacchii Trematomus pennelli Pleuronectidae Hippoglossus hippoglossus
Sampling locality codes: AN: Antarctica; AR: Argentine coast; AU: Australian coast; BA: Baltic Sea; BE: Bering Sea; BL: Baikal Lake; BS: Barents Sea; JA: Japan Sea; NEA: North-East Atlantic; NEP: North-East Pacific; NWA: North-West Atlantic; NWP: North-West Pacific; RS: Ross Sea; SA: South Atlantic Ocean (off South Africa coast); SEA: South-East Atlantic; SWA: South-West Atlantic; WS: Weddell Sea (data from Mattiucci and Nascetti, 2007; Mattiucci et al., 1998, 2003, 2008b; Nadler et al., 2000, 2005; Nascetti et al., 1993; Orecchia et al., 1994; Paggi et al., 1998c). Hosts listed by alphabetical order of the family.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
89
C. osculatum D and C. osculatum E of Orecchia et al. (1994): The Antarctic members of the C. osculatum complex, C. osculatum D and C. osculatum E were demonstrated to be reproductively isolated by allozyme markers. They occur sympatrically in the same definitive host, the Weddell seal, Leptonychotes weddellii, and have so far been reported from both the Weddell and the Ross Seas (Antarctica) (Orecchia et al., 1994). The larval stages of the two sibling species have been identified by allozyme markers from several fish species belonging to the families Channicthydae, Bathydraconidae and Nototheniidae, in which a differential distribution of the two sibling species is reported (Mattiucci and Nascetti, 2007; see also Section 7.2) (Table 2.7; Fig. 2.1).
3.3.2. The Contracaecum ogmorhini species complex
The pinniped parasite Contracaecum ogmorhini Johnston and Mawson, 1941, first described from the leopard seal, Hydrurga leptonyx, in South Australian waters, was later synonymized with C. osculatum (see Johnston and Mawson, 1945). However, it was considered valid by Fagerholm and Gibson (1987). The species was found to be genetically heterogenous using allozyme markers (18 enzyme loci), indicating the existence of two reproductively isolated taxa (sibling species) included within the morphospecies. A formal description of the two taxa was given by Mattiucci et al. (2003), and they were named C. ogmorhini Johnston and Mawson (1941) (sensu stricto) and C. margolisi Mattiucci et al. (2003). A morphological description of C. ogmorhini (s.s.) from Arctocephalus australis was given by Timi et al. (2003). C. ogmorhini (s.s.) has been detected as an adult in the otariid seals Arctocephalus pusillus pusillus, A. pusillus doriferus and A. australis in the Austral region, while C. margolisi was detected in the otariid Zalophus californianus in the Boreal region (Table 2.6; Fig. 2.1) (see also Section 4.3). To date, the allopatric distribution of these two sibling species appears to be related to that of their definitive hosts, ranging from 20 to 55 N in Boreal Pacific waters, but from 20 to 50 S in South Atlantic and South Pacific waters.
3.3.3. Validity of species of Contracaecum, parasitic in seals, using molecular markers
C. osculatum baicalensis Moszgovoi and Ryzhykov, 1950: The specific status of C. osculatum baicalensis was established genetically using allozyme markers (22 enzyme loci) with respect to the species of the C. osculatum complex listed above (D’Amelio et al., 1995). It was found to be genetically well distinct from the other members of this complex, as well as from the other congeneric taxa (Mattiucci et al., 2008b). C. osculatum baicalensis is an adult parasite of the freshwater Baikal seal, Phoca sibirica, endemic to Lake Baikal. The genetic relationships between C. osculatum baicalensis and other congeneric taxa were inferred from
90
Simonetta Mattiucci and Giuseppe Nascetti
LSU rDNA sequences (Nadler et al., 2000) and from the mtDNA cox2 sequence (Mattiucci et al., 2008b) analyses. Both phylogenetic analyses were congruent with the allozyme clustering in showing C. osculatum baicalensis nested within the clade formed by the species of the C. osculatum complex (see Section 4.3). The genetic relationships found between C. osculatum baicalensis and the members of the C. osculatum complex suggest that the evolutionary divergence of this taxon likely took place following its host’s isolation during a Pliocene-Pleistocene refuge (Deme´re´ et al., 2003). At a larval stage, C. osculatum baicalensis was recognized by allozymes in the endemic fish species Cottomephorus grewingki, C. inermis, Comephorus baicalensis, Coregonus lavaretus, C. autumnalis migratorius and Thymallus arcticus, which represent prey items of Phoca sibirica in Lake Baikal (our unpublished data). C. radiatum (v. Linstow, 1907) Baylis, 1920: The taxonomic status of this species was confirmed genetically by Arduino et al. (1995) on the basis of 24 enzyme loci. Several allozymes were found to be diagnostic between C. radiatum and the other taxa so far characterized as belonging to Contracaecum species from seals (Arduino et al., 1995; Mattiucci et al., 2008b, unpublished data). Reproductive isolation from the two Antarctic members of the C. osculatum complex (i.e. C. osculatum D and C. osculatum E) occurring sympatrically in the same definitive hosts (the Weddell seal) was proved by the lack of F1 hybrids and recombinant or introgressed individuals between the Antarctic taxa in the sympatric areas of the Weddell and Ross Seas (Arduino et al., 1995). The genetic relationships between C. radiatum and other congeneric taxa were later inferred from LSU rDNA sequences (Nadler et al., 2000) and mtDNA cox2 sequence analyses (Mattiucci et al., 2008b). Morphological distinction between C. radiatum and C. osculatum (s.l.) was given by Klo¨ser and Plo¨tz (1992). C. radiatum has been genetically identified as an adult in Leptonychotes weddellii and as a larva in the pelagic channichthyd fishes Chionodraco hamatus and Criodraco antarcticus (see Arduino et al., 1995). This finding supports a previous report by Klo¨ser et al. (1992), according to which C. radiatum has become adapted to a pelagic food web. Other definitive hosts recorded for this species in Antarctic waters are the leopard seal, Hydrurga leptonyx, and the Ross seal, Ommatophoca rossi (see Baylis, 1937; Dailey, 1975). Genetic investigations on this parasite of Antarctic seals are needed in order to determine any host preference of this species. C. mirounga Nikolskii, 1974: The taxonomic status of the species was confirmed genetically by allozyme markers (20 enzyme loci) (Mattiucci et al., 2008b, unpublished data). It was detected genetically as an adult in Mirounga leonina from the Antartic and sub-Antarctic area (Mattiucci et al., 2008b) and also in the otariid Arctocephalus australis (see Mattiucci et al., 2003). Several allozymes were found to be diagnostic between C. mirounga and other Contracaecum species from seals (Mattiucci et al., 2008b,
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
91
unpublished data). Reproductive isolation from the two Antarctic members of the C. osculatum complex (i.e. C. osculatum D and C. osculatum E) occurring sympatrically in the same definitive host (the Weddell seal), and from C. ogmorhini (s.s.) which occurs syntopically in the same host (the fur seal, Arctocephalus australis), was demonstrated by the lack of F1 hybrids and recombinant or introgressed individuals. The genetic relationships between C. mirounga and other congeneric taxa were later inferred from LSU rDNA sequences (Nadler et al., 2000) and from the mtDNA cox2 sequences analyses (Mattiucci et al., 2008b). No data on the larvae of this species which have been identified genetically are available.
4. PHYLOGENETIC SYSTEMATICS OF ANISAKID NEMATODES Allozyme data, typically consisting of allele frequencies obtained from starch gel electrophoresis of proteins, are an important source of characters for understanding the genetic structure of a population of anisakid nematodes and, consequently, for reconstructing phylogenies among conspecific populations and closely related species. Despite the increasing use of DNA sequence data in phylogenetics, allozyme data remain widely used in systematic and evolutionary studies of anisakid nematodes and have some advantages. For example, allozyme data consist of multiple unlinked nuclear loci, with each locus providing independent estimate of the species differentiation and phylogeny. Therefore, in contrast to results of nuclear and mitochondrial DNA data sets, allozyme data are less likely to be systematically missed by mismatches between gene trees and species trees. Furthermore, it is relatively cheap and easy to survey allozyme variation for a large number of individual anisakid nematodes. A problematic aspect using allozyme data, however, is that there is a longstanding and continuing controversy as to the preferred method for their phylogenetic analysis. Doubt remains as to the validity of allozyme data as a basis for inference of genetic relationship among related taxa. Although this issue is still debated in the literature, aspects could be addressed with empirical data recently obtained for anisakid nematodes. Genetic relationships inferred from allozyme markers have been largely obtained, in this group of parasites, by different methods such as phenograms from genetic distance methods, Neighbour Joining (NJ) from genetic matrix distances, spatial representation [Multidimensional Scale Ordination (MDS), Principal Component Analysis (PCA)] based on distance values or allele frequencies data, between several conspecific anisakid nematodes. From a phylogenetic point of view, anisakid species have been recognized based upon evidence of independent evolutionary lineages
92
Simonetta Mattiucci and Giuseppe Nascetti
reflected in the form of character states unique to the same individuals of a species. Indeed, genetically defined clades, corresponding to distinct evolutionary lineages, are consistent with their recognition as separate species. Congruence studies so far carried out on anisakid nematodes provided, in general, that clades supported by many different lines of evidence (allozymes, DNA sequences) can effectively be considered to be ‘known’ clades (as determined and supported by Felsenstein, 1985). As it has been presented in previous paragraphs, several molecular systematic studies of anisakid nematodes have employed more than one gene to define the taxonomic status of the species so far known. Indeed, generally, concordant evidence from independent gene analysis inspires much more confidence than the patterns that reflect separate lineage (and, thus, species existence) inferred from data obtained from a single gene. Congruence and incongruence evidences between allozyme data sets and not-allozyme data (i.e. DNA sequence analysis at both nuclear and mitochondrial genes) in anisakid nematode phylogenies are here reviewed. The opportunity to compare independent data sets supporting a given clade could affirm the existence of the clade with greater authority. In other words, shared hypothetical phylogenetic history seems to be the most likely explanation when congruence between diverse genetic data sets is observed. In this sense we are revising the phylogenetic relationships so far attempted for each group of anisakid nematodes belonging to the genera Anisakis, Pseudoterranova and Contracaecum. Indeed, phylogenetic analysis of partial 28S (LSU) sequences carried out by Nadler et al. (2005) of Anisakis, Pseudoterranova and Contracaecum taxa revealed strong support for the monophyly of the Anisakinae, Contracaecum plus Phocascaris, Pseudoterranova and Anisakis. Parsimony and ML analyses indicated that the Raphidascarididae, Contracaecum plus Phocascaris, and the Anisakinae (here considering only Pseudoterranova and Anisakis) are each monophyletic, the latter two groups with consistently strong bootstrap support at MP and ML analyses (Nadler et al., 2005).
4.1. Genetic relationships between Anisakis spp. Work performed on the phylogenetic studies within this genus was done initially using the unweighted pair group method using arithmetic averages (UPGMA phenograms) on the studied populations and species of Anisakis based on distance values inferred from DNei (Nei, 1972) and Dc (chord distance by Cavalli-Sforza and Edwards, 1967). Among the first extensive UPGMA phenograms were those generated by Mattiucci et al. (1997, 2001) and Paggi et al. (1998b) showing the genetic relationships between the species of Anisakis. This suggested that Anisakis is polyphyletic and highly heterogeneous, with a high genetic differentiation of A. physeteris and A. brevispiculata from the rest of the Anisakis taxa
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
93
recognized genetically. Other elaborations to infer genetic relationships between seven taxa of Anisakis (i.e. the A. simplex complex, A. typica, A. ziphidarum, A. brevispiculata and A. physeteris) included the MDS (Guttman, 1968) from chord-distance values (Dc, Cavalli-Sforza and Edwards, 1967), demonstrating that A. physeteris and A. brevispiculata are forming a well-distinct clade with respect to that formed by A. simplex complex, as well as from both A. typica and A. ziphidarum (see Mattiucci et al., 2002a). A consensus NJ tree was later inferred from Cavalli-Sforza and Edwards chord-distance values estimated at 19 enzyme loci between eight Anisakis taxa, including the new species A. paggiae (see Mattiucci et al., 2005). This showed consistently the existence of two distinct clades supported by high bootstrap values, including, respectively: (1) the three members of A. simplex complex, A. typica and A. ziphidarum within clade I and (2) A. physeteris, A. brevispiculata and A. paggiae within clade II (Fig. 2.2). Both A. ziphidarum and A. typica form two distinct subclades, closely related to the A. simplex species complex (Mattiucci et al., 2005). Maximum parsimony (MP) analysis of the ITS data set of 724 characters in seven taxa out of those genetically recognized in Anisakis, was performed by Nadler et al. (2005). This produced a strict consensus of the MP trees which also depicted A. physeteris plus A. brevispiculata as the sister group to the remaining Anisakis taxa. This clade was well supported in the MP bootstrap tree. According to this analysis, the remaining ingroup Anisakis were monophyletic in maximum likelihood (ML) and MP consensus trees, with 100% bootstrap support (Nadler et al., 2005). Later, Valentini et al. (2006) provided a phylogenetic hypothesis for all the nine taxa currently recognized species in Anisakis, using MP, NJ and Bayesian analysis (BA) inferred from mtDNA cox2 sequences data (629 bp). Phylogenetic trees generated show high congruence with the UPGMA and NJ trees obtained from allozyme data sets performed on the same species of Anisakis. An overall high congruence was indeed found between the tree topologies obtained from consensus NJ inferred from mtDNA cox2 sequence data and consensus NJ tree generated from allozyme data sets of the same taxa (Fig. 2.2). Both depicted the species A. physeteris, A. brevispiculata and A. paggiae as a sister group, highly supported, with respect to the other Anisakis taxa. In addition, A. paggiae appears to share a common ancestor with A. brevispiculata, and this is well supported by the NJ allozyme data. In addition, allozyme tree topology clearly demonstrated that Anisakis sp. formed a monophyletic group with A. ziphidarum; this was supported by the MP, BI and NJ of mtDNA cox2 sequences (Valentini et al., 2006) (Fig. 2.2). Finally, consistent tree topologies were observed between nuclear gene products and mitochondrial genes for the position occupied by A. typica as forming a separate clade—as depicted by mtDNA cox2-derived NJ (Valentini et al., 2006),
A (allozyme data)
B (mtDNA cox2) A. simplex (s.s.)
A. simplex (s.s.) 71
A. pegreffii
100 96
A. pegreffii
A. simplex C
99
100
A. simplex C
A. typica
A. typica
A. ziphidarum
73
A. ziphidarum
71
76 Anisakis sp.
Anisakis sp.
A. physeteris
A. paggiae 50
A. brevispiculata
A. brevispiculata A. physeteris
A. paggiae P. decipiens
70
(outgroup) P. decipiens Clade I Clade II
FIGURE 2.2 Genetic relationships among Anisakis spp. depicted by (A) neighbour-joining (NJ) tree inferred from chord-distance values (Dc, Cavalli-Sforza and Edwards, 1967) from allozyme data; (B) NJ inferred from K2P distance values obtained by mtDNA cox2 sequences analysis (data from Valentini et al., 2006). Bootstrap values 60 are shown at the internal nodes. Pseudoterranova ceticola as outgroup. The two data sets are congruent in depicting the existence of two main clades: (I) includes the A. simplex species complex [A. pegreffii, A. simplex (s.s.), A. simplex C], A. typica, A. ziphidarum and Anisakis sp.; (II) comprises the species A. physeteris, A. brevispiculata and A. paggiae. The species included in the two main clusters show a different larval morphology: type I (sensu Berland, 1961) in the first group and type II in the second one.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
95
allozyme data, and inferred from ITS rDNA sequence analysis (Nadler et al., 2005) (Fig. 2.2). Therefore, phylogenies inferred from different sets of nuclear and mitochondrial data (i.e. allozymes, ITS rDNA sequences, mtDNA cox2) (Mattiucci et al., 2005; Nadler et al., 2005; Valentini et al., 2006) supported the group of species formed by A. physeteris, A. brevispiculata and A. paggiae as sister group to the remaining Anisakis taxa. Finally, the clade formed by the sibling species of the A. simplex complex received strong support in all the phylogenetic elaborations from allozymes (NJ), ITS-rDNA (MP) and mtDNA cox2 sequences (MP, NJ, BI). They were concordant to show a sister-group relationship for A. simplex (s.s.) and A. pegreffii, with A. simplex C as a sister to this clade (Mattiucci and Nascetti, 2006; Nadler et al., 2005; Valentini et al., 2006) (Fig. 2.2).
4.2. Genetic relationships between Pseudoterranova spp. Genetic relationships between Pseudoterranova species have included analyses based on allozyme genetic distance methods (Bullini et al., 1997, Paggi et al., 1991, 1998c). The allozyme analysis inferred from Dc chorddistance values between the five sibling species of the P. decipiens complex indicated the following topology: [P.decipiens (s.s.), P. azarasi, P. krabbei, P. bulbosa, P. decipiens E], with a close genetic relationship between P. decipiens (s.s.) and P. azarasi (Fig. 2.3). An UPGMA phenogram of uncorrected ITS rDNA distances (Zhu et al., 2002) was also produced for individual specimens, previously characterized by allozymes, belonging to the taxa: P. decipiens (s.s.), P. krabbei
P. decipiens (s.s.)
P. azarasi
P. bulbosa P. decipiens E 1.00
0.80
0.60
0.40
0.20
Nei’s D 0.00
FIGURE 2.3 Genetic relationships among Pseudoterranova spp. depicted by neighbourjoining (NJ) tree inferred from chord-distance values (Dc, Cavalli-Sforza and Edwards, 1967) from allozyme data (redrawn from Bullini et al., 1997).
96
Simonetta Mattiucci and Giuseppe Nascetti
P. krabbei, P. bulbosa, P. azarasi, P. cattani, and a taxon indicated as P. decipiens Ca1. This analysis showed the following relationship: [P. decipiens (s.s.), P. azarasi, P. bulbosa, P. cattani, P. krabbei, P. decipiens Ca1]. Zhu et al. (2002) also suggested that, because of the geographical origin, the P. decipiens Ca1 from Chaenocephalus aceratus, it is likely to correspond to P. decipiens E, but this correspondence has not been verified. An overall congruence was found between the topology produced by the allozyme phenogram (except for the absence of P. cattani from the allozyme analysis) produced by Bullini et al. (1997) and that obtained by both MP and ML analyses of ITS rDNA sequences by Nadler et al. (2005) performed on all the P. decipiens species complex. These topologies were congruent in depicting the close relationship between P. decipiens (s.s.) and P. azarasi. Nadler et al. (2005) showed that P. cattani and P. decipiens Ca1 are not closely related and they do not represent sister taxa with respect to the other taxa. According to those authors, this finding suggests a more complex evolutionary scenario than might be explained by a simple biogeographical scenario of their definitive hosts.
4.3. Genetic relationships between Contracaecum spp. A first phylogenetic hypothesis of the species included in the genus Contracaecum was that attempted using allozyme data on several species of Contracaecum from pinnipeds and fish-eating birds as definitive hosts, plus Phocascaris phocae and P. cystophorae (see Nascetti et al., 1990; Orecchia et al., 1986b). It was shown that Contracaecum species from seals were genetically most similar to each other rather than to the avian Contracaecum, with no alleles found in common between the two groups of species. In addition, it was clearly demonstrated that species of Phocascaris (i.e. P. phocae and P. cystophorae), despite their morphological characters (interlabia absent and/or reduced), were clustering within the clade formed by the species of the C. osculatum complex (interlabia present), which now includes five sibling species [C. osculatum A, C. osculatum B, C. osculatum (s.s.), C. osculatum D and C. osculatum E] (Nascetti et al., 1993; Orecchia et al., 1994) (see also Section 3.3.) Nadler et al. (2000) supported this first phylogenetic hypothesis using the nuclear-encoded large subunit ribosomal DNA sequences (LSUrDNA) for several taxa of Contracaecum (including seven species from pinnipeds and seven from fish-eating birds) and Phocascaris (two taxa), previously identified by allozyme markers. These data provided high support for the monophyly of all Contracaecum and Phocascaris of phocid seals. This finding is consistent with Berland’s hypothesis (1964) that such species form a group sharing the same life-cycle pathway. However, Berland’s proposal to recognize all species in phocid seals as Phocascaris, with all the species from birds as Contracaecum, was not congruent, according to
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
97
Nadler et al. (2000) with the inferred 28S rDNA trees produced, because it would result in a paraphyletic Contracaecum. Moreover, that phylogenetic elaboration, with respect to Berland’s hypothesis, would not depict avian parasites as monophyletic; some of them, for instance C. rudolphii species A and B, cluster in the same clade with the otariid parasite C. ogmorhini (see Nadler et al., 2000, 2005). A phenetic clustering (UPGMA), inferred from the allozyme data set (22 enzyme loci) only in Contracaecum spp. using pinnipeds as definitive hosts, showing their genetic relationship was produced by Nascetti et al. (1993). Later Arduino et al. (1995) and Bullini et al. (1997) produced a phenetic tree (UPGMA) showing the genetic relationships among six taxa, including the species of the C. osculatum complex [C. osculatum A, C. osculatum B, C. osculatum (s.s.), C. osculatum D, C. osculatum E] and C. radiatum. This depicted C. radiatum as the basal species of the first group of Contracaecum plus Phocascaris. Other graphical representations of the genetic relationships among the same group of Contracaecum taxa were those produced by MDS ordination obtained from Dc values in Orecchia et al. (1994) and the PCA analysis in Arduino et al. (1995), also including C. radiatum. A phenogram depicting the genetic differences observed in the entire ITS rDNA among specimens genetically identified previously by allozyme markers as belonging to the five taxa of the C. osculatum complex [i.e. C. osculatum A, C. osculatum B, C. osculatum baicalensis, C. osculatum (s.s.), C. osculatum D and C. osculatum E] was produced by Zhu et al. (2000b). This cluster analysis showed that C. osculatum A, C. osculatum D, C. osculatum E and C. osculatum baicalensis were genetically more similar to C. osculatum B than each was to C. osculatum (s.s.). Moreover, Zhu et al. (2000b) found C. osculatum (s.s.) the most genetically distinct taxon from the other members of the complex. This latter finding was concordant with that previously presented in Nascetti et al. (1993) and Orecchia et al. (1994). However, in the same genetic studies and phylogenetic representation, Zhu et al. (2000b) did not find any genetic differentiation in the ITS rDNA sequence analysis between C. osculatum D and C. osculatum E, previously detected by allozymes (Orecchia et al., 1994). Similarly, the lack of unequivocal nucleotide differences in any of three mtDNA sequences analysed (mtDNa cox1, ssrRNA and lsrrRNA) between C. osculatum D and C. osculatum E was also shown by Hu et al. (2001). These findings are inconsistent with allozyme data analysis (MAE) which, on the contrary, clearly demonstrated the reproductive isolation between C. osculatum D and C. osculatum in sympatry, with several allozyme loci fixed for alternative alleles in the two species. Moreover, the specific status of C. osculatum D and C. osculatum E was confirmed by the phylogenetic analysis, inferred from mtDNA cox2 sequences, which depicted the two taxa as two separate lineages (phylogenetic units) (Mattiucci et al., 2008b) (Fig. 2.4).
A (allozyme data)
B (mtDNA cox2)
98
C. osculatum A
C. osculatum B
C. osculatum E
C. osculatum (s.s.)
C. osculatum D
P. cystophorae C. osculatum A
C. o. baicalensis
C. osculatum D C. osculatum (s.s.)
90
96
62 94
C. o. baicalensis
C. osculatum B
90
95
96 99
C. osculatum E
85
P. cystophorae C. radiatum
88
C. radiatum
95 C. mirounga
C. mirounga
C. ogmorhini (s.s. )
C. ogmorhini (s.s. )
100 C. margolisi
C. margolisi P. ceticola
100
(outgroup)
P. ceticola
Clade I Clade II
FIGURE 2.4 Genetic relationships between species of Contracaecum and Phocascaris from pinnipeds depicted by (A) NJ tree inferred from chord-distance values (Dc, Cavalli-Sforza and Edwards, 1967) from allozyme data and (B) NJ inferred from mtDNA cox2 sequences analysis. Bootstrap values 60 are shown at the internal nodes. Pseudoterranova ceticola as outgroup. The two data sets are congruent in depicting the existence of two main clades: (I) includes the species of C. osculatum complex, Phocascaris cystophorae, C. radiatum, C. mirounga; (II) comprises C. ogmorhini (s.s.) and C. margolisi. The two phylogenetic trees show P. cystophorae nested in the subclade formed by the species of C. osculatum complex.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
99
The overall concordant results with the allozyme data sets have been found in the tree topology inferred from the mtDNA cox2 sequences data (519 bp) ascertained for all the Contracaecum plus Phocascaris taxa so far recognized by allozyme markers [i.e. C. osculatum A, C. osculatum B, C. osculatum (s.s.), C. osculatum D, C. osculatum E, C. osculatum baicalensis, C. radiatum, C. mirounga, C. ogmorhini (s.s.), C. margolisi, Phocascaris cystophorae] (see Mattiucci et al., 2008b) (Fig. 2.4). A high congruence in the topology of the trees generated by a consensus NJ tree inferred from mtDNA cox2 sequences and that produced by a NJ based on Dc chord distance from allozyme markers was observed (Fig. 2.4). They are highly consistent in showing the existence of two well-supported clades: (1) one including the species of the C. osculatum complex, Phocascaris cystophorae, plus C. radiatum and C. mirounga (clade I), with the last two taxa being genetically closely related and forming a supported subclade; (2) the second clustering of the species C. ogmorhini (s.s.) and C. margolisi (clade II). Moreover, both elaborations from different data sets showed that the species P. cystophorae is nested in the subclade of the species of the C. osculatum complex. This is despite its morphological differences from the species of the C. osculatum complex. Allozyme NJ elaboration clearly depicted the species C. ogmorhini (s.s.) and C. margolisi as sister taxa with respect to the other Contracaecum taxa, and this was also shown by the mtDNA cox2 data. In addition, the Antarctic taxon C. osculatum D forms a monophyletic group with the other Antarctic member C. osculatum E, from which it was also found to be clearly genetically distinct (Table 2.10 and Fig. 2.4; see also Section 6). An overall congruence of this phylogenetic analysis was with that produced by Nadler et al. (2000) based on the LSU rDNA sequences analysis of seven Contracaecum species from pinnipeds [i.e. C. osculatum A, C. osculatum B, C. osculatum (s.s.), C. osculatum baicalensis, C. radiatum and C. mirounga]. Indeed, both NJ tree inferred from the full alignment data set using log-determinant distances and MP produced by Nadler et al. (2000) demonstrated the monophyly of Contracaecum plus Phocascaris from phocids. A strong support in all bootstrap trees also indicated for the subclades formed by C. radiatum and C. mirounga and that formed by C. osculatum A and C. osculatum baicalensis (see Nadler et al., 2005). These subclades were shown to be highly supported at MP and NJ analyses inferred from mtDNA cox2 sequences analysis (Mattiucci et al., 2006, 2008b) (Fig. 2.4). Furthermore, the trees inferred by all the methods, were congruent in demonstrating that Phocascaris spp. are clustering with the C. osculatum complex group of species and are well nested in the clade including the Contracaecum from phocids (see Mattiucci et al., 2008b; Nadler et al., 2000, 2005) (see also Section 6). Meanwhile, C. ogmorhini (s.s.) and C. margolisi cluster in a separate clade (Mattiucci et al., 2008b).
100
Simonetta Mattiucci and Giuseppe Nascetti
5. GENETIC DIFFERENTIATION IN ANISAKIDS Although the level of genetic differentiation is not a suitable parameter or a measurement for establishing species delimitation, values of genetic differentiation between taxa of anisakid nematodes inferred from nuclear loci analysis (19–22 enzyme loci) were found to be of a similar order. In sympatric or partially sympatric sibling species of anisakid nematodes, a significant deficiency or complete lack of some heterozygote classes at polymorphic loci strongly suggests that the sample being dealt with comprises distinct gene pools. This was widely demonstrated in samples of A. simplex (s.l.), P. decipiens (s.l.), and C. osculatum (s.l.) (see Mattiucci et al., 1997; Nascetti et al., 1986, 1993; Paggi et al., 1991). In allopatric anisakid nematodes, allozymes have provided reliable information on their specific status, with clear evidence of alternative and unique allozymes existing in allopatric populations. As the most common mode of speciation of endoparasites is the peripatric model (sensu Mayr, 1963, 1976), the build up of genetic divergence is generally a better predictor of whether two allopatric populations will interbreed upon recontact, than is conventional morphology. Indeed, according to the peripatric model of speciation, this involves geographical isolation of small populations whose genetic structure begins to differ from that of the parental population since the beginning of the process by genetic drift phenomena. In the Anisakis simplex complex and Contracaecum osculatum complex of species, genetic data strongly support the notion that adaptation to different hosts and speciation is strictly related to the geographical isolation of the hosts (see also Section 6.1). At least, in the case of anisakid nematodes, when Nei’s (1972) genetic distance between populations reaches values of 0.2–0.3, gene exchange is interrupted by intrinsic reproductive isolating mechanisms (RIMs). On the other hand, in anisakid nematodes, conspecific populations generally show similar allele frequencies, even when located thousands of kilometrers apart. Accordingly, their DNei values are quite low (0.0001– 0.002). Average genetic distance values inferred from allozymes within and among species of the genera are given in Tables 2.8–2.10.
5.1. Genetic differentiation at interspecific level Between sibling species of anisakid nematodes (sympatric and allopatric), the genetic differentiation, estimated at the nuclear level (allozymes), ranges on average, from values of Nei’s (1972) standard genetic distance (DNei) DNei 0.20 (as observed between the Arctic taxon C. osculatum A and the Antarctic species C. osculatum E) (Table 2.10) to DNei 0.90 (as found between the Antarctic taxon P. decipiens E vs the other species of
TABLE 2.8 Average of standard genetic distance by Nei (1972, DNei, below the diagonal) inferred from 20 enzyme-loci, and by Kimura2-parameter (Kimura, 1980, K2P, above the diagonal) inferred from 629bp of mtDNA cox2 between the species of Anisakis so far genetically detected
Species
A. simplex (s.s.)
A. simplex (s.s.) A. pegreffii A. simplex C A. typica A. ziphidarum Anisakis sp. A. physeteris A. brevispiculata A. paggiae
0.40 0.36 1.16 1.64 2.04 6.90 4.10 1
A. pegreffii
A. simplex C
A. typica
A. ziphidarum
Anisakis sp.
A. physeteris
A. brevispiculata
A. paggiae
0.05
0.06 0.06
0.14 0.14 0.14
0.12 0.12 0.13 0.13
0.13 0.13 0.14 0.12 0.09
0.14 0.14 0.14 0.15 0.13 0.15
0.17 0.17 0.17 0.18 0.14 0.16 0.11
0.14 0.14 0.15 0.15 0.12 0.13 0.12 0.13
0.37 1.45 1.99 2.63 8.30 6.11 1
1.14 1.62 1.92 7.40 5.54 1
1.67 1.62 4.77 3.49 1
0.68 1 1 1
1 1 1
0.95 1.06
(Data from: Mattiucci et al., (2001, 2002a, 2005), Mattiucci and Nascetti, 2006; Paggi et al., 1998b, and Valentini et al., 2006).
0.79
102
Simonetta Mattiucci and Giuseppe Nascetti
TABLE 2.9 Average values of genetic distance, DNei (1972), between species of the Pseudoterranova decipiens complex Species
P. krabbei
P. decipiens (s.s.) P. bulbosa P. azarasi P. decipiens E
0.51 0.93 0.56 0.91
P. decipiens (s.s.)
P. bulbosa
P. azarasi
0.60 0.38 0.96
0.64 0.97
0.93
Data from Bullini et al. (1997), Mattiucci et al. (1998, unpublished data), Paggi et al. (1991, 1998c).
P. decipiens complex), or DNei = 0.93 (as between P. krabbei and P. bulbosa) (Table 2.9). Similar high values were those observed in the comparison of Arctic members of the C. osculatum complex [i.e. C. osculatum D vs C. osculatum (s.s.), DNei = 0.70] and between members of the A. physeteris complex (i.e. A. physeteris vs A. paggiae and A. brevispiculata) (DNei 0.90) (see Mattiucci et al., 2005) (Table 2.8). The highest interspecific differentiation is that found at the allozyme level in the comparison of anisakid species that are morphologically distinct, such as the A. simplex complex and A. ziphidarum (see Paggi et al., 1998b) or the A. physeteris complex (Mattiucci et al., 2005). In the last group of species, spicule length of male specimens and the ventriculus are significantly shorter than those of the members of A. simplex complex. Moreover, in A. paggiae, the spicule length and ventriculus shape were demonstrated to be of diagnostic level not only from the members of A. simplex complex, but also with respect to genetically closely related species, that is, A. physeteris and A. brevispiculata (see Mattiucci et al., 2005). Analogously, a high genetic divergence at the allozyme level was also found between members of the C. osculatum complex with respect to C. mirounga (on average, DNei 1.05) (Mattiucci et al., 2008b) and to the C. ogmorhini species complex (on average, DNei 1.30) (Mattiucci et al., 2003, 2008b), which are morphologically distinct. The highest level of differentiation at allozyme level was found between the A. simplex species complex with respect to the A. physeteris species complex (Mattiucci et al., 2005) (Table 2.8). Nuclear and mitochondrial DNA polymorphisms among species of anisakid nematodes demonstrate significantly high genetic variation among sibling species and morphospecies of those anisakid nematodes previously characterized genetically by allozyme markers. Indeed, it has recently been shown that most of the genetic diversity is strongly structured between species rather than within species. Pairwise comparisons were estimated of the level of sequence differences at the ITS rDNA between 14 Anisakis samples by Nadler et al. (2005). Sequence differences
TABLE 2.10 Average of standard genetic distance by Nei (1972, DNei, below the diagonal) inferred from 20 enzyme-loci, and by Kimura2-parameter (Kimura, 1980, K2P, above the diagonal) inferred from 519 bp of mtDNA cox-2 between Contracaecum spp. from pinnipeds. Species
COSA
C. osculatum A (COSA) C. osculatum B (COSB) C. osculatum (s.s.) (COSS) C. osculatum D (COSD) C. osculatum E (COSE) C. baicalensis (CBAI) P. cystophorae (PCYS) C. radiatum (CRAD) C. mirounga (CMIR) C. ogmorhini (s.s.) (COGM) C. margolisi (CMAR)
0.41 0.57 0.23 0.20 0.30 0.45 0.72 1.00 1.30 1.35
COSB
COSS
COSD
COSE
CBAI
PCYS
CRAD
CMIR
COGM
CMAR
0.09
0.10 0.09
0.09 0.09 0.11
0.09 0.10 0.10 0.05
0.06 0.08 0.10 0.09 0.09
0.09 0.10 0.07 0.11 0.11 0.09
0.14 0.14 0.16 0.13 0.15 0.16 0.15
0.13 0.13 0.15 0.13 0.13 0.12 0.15 0.09
0.16 0.15 0.14 0.16 0.14 0.15 0.16 0.14 0.14
0.15 0.15 0.15 0.16 0.15 0.16 0.16 0.15 0.15 0.05
0.80 0.38 0.36 0.27 0.83 0.76 1.06 1.35 1.40
0.64 0.53 0.41 0.46 1.30 1.13 1.37 1.42
0.25 0.60 0.68 0.78 1.15 1.40 1.39
0.50 0.51 0.67 0.86 1.48 1.45
(Data from: Nascetti et al., 1993; Orecchia et al., 1994; Mattiucci et al., 2008b and unpublished).
0.64 0.79 1.20 1.70 1.70
0.96 1.07 1.90 1.89
0.95 1.60 1.75
1.81 1.80
0.30
104
Simonetta Mattiucci and Giuseppe Nascetti
(absolute differences) of 0–7 existed among individuals belonging to the same species, whereas it ranged from 37 to 149 between individuals belonging to distinct species (Nadler et al., 2005). At the mtDNA level, the genetic divergence estimation among the nine Anisakis taxa, previously characterized on the basis of allozyme data, was calculated at Kimura-2-parameters distance values (K2P) inferred from sequence analysis (629 bp) of the mtDNA cox2 gene by Valentini et al. (2006) (Table 2.8). The mtDNA cox2 fragment in all the Anisakis was found to be A+T rich (60.7%, 64.9% and 74.7% at the first, second and third positions, respectively). The lowest level of interspecific genetic distance was found between sibling species of the Anisakis simplex complex (on average, K2P 0.05). Whereas, values of K2P = 0.14 and 0.13 were observed when comparing the A. simplex complex, on average, with A. typica and A. ziphidarum. Similar values (on average K2P 0.13) were obtained when A. physeteris, A. brevispiculata and A. paggiae are compared to A. ziphidarum (see Valentini et al., 2006) (Table 2.8). At the amino acid level, a total of 31 variable positions were identified in the mtDNA cox2 gene for Anisakis spp. The average variation ranged from 2.4% between the sibling species of A. simplex complex to as high as 7.7% between morphologically distinct species (i.e. A. physeteris, A. brevispiculata and A. paggiae vs the A. simplex complex (Valentini et al., 2006)). Single strand conformation polymorphism (SSCP) analysis of the ITS rDNA, performed by Zhu et al. (2002), has revealed that, while no variation in single stranded ITS profiles existed within each of the five sibling species of P. decipiens complex (with the exception of a slight microheterogeneity evidenced in P. bulbosa and P. cattani), SSCP analysis of the ITS-2 amplicons allowed significant differentiation between them (Zhu et al., 2002). Pairwise comparison of the ITS sequences revealed nucleotide differences ranging from 0 to 6.8%, which were within the range observed in other complex members (0–2.3% in the C. osculatum complex) (see Zhu et al., 2000b). Sequence variation at the same internal transcribed spacers of ribosomal DNA (ITS-rDNA) within and among members of the Contracaecum osculatum complex and SSCP was used to screen the ITS-1 and ITS-2 amplicons separately for sequence variation within and among individuals (Zhu et al., 2000b). The G+C contents of the sequences obtained were 45.7% (ITS-1) and 42.0–43.5% (ITS-2). However, no variation in single-strand profiles was detected between/among samples representing a taxon by Zhu et al. (2000b), except for slight nucleotide polymorphism detected in the ITS for C. osculatum A and C. osculatum B. Indeed, there was no variation in the length of the ITS-1 (499 bp) or ITS-2 (262 bp) sequence for any of the individuals examined, and no-intraspecific variation in the sequence was observed. Whereas, significant inter-taxon
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
105
differences in SSCP profiles were detected between C. osculatum A, C. osculatum B, C. osculatum C and C. osculatum baicalensis. This revealed a reliable genetic differentiation of these taxa from one another, except in the case of C. osculatum D versus C. osculatum E, which exhibited identical sequences (Zhu et al., 2000b). Each of the other four taxa had distinct sequences with interspecific differences ranging from 0.3 to 2.3% (Zhu et al., 2000b). C. osculatum (s.s.) was the most distinct taxon with respect to all the other members. Comparison among the taxa revealed 22 variable nucleotide positions in the ITS sequence. Each of the taxa could be identified by the presence of 1–10 fixed nucleotide differences, except in the case of C. osculatum D and C. osculatum E, which were found at the ITS rDNA gene to show the same nucleotide sequence. At the mitochondrial level, the genetic divergence estimated from sequence analysis (519 bp) of the cox2 gene was given for the five members of the C. osculatum complex [i.e. C. osculatum A, C. osculatum B, C. osculatum (s.s.), C. osculatum D and C. osculatum E] with respect to C. osculatum baicalensis, C. radiatum, C. mirounga, Phocascaris cystophorae and to the otariid parasites, C. ogmorhini (s.s.) and C. margolisi (see Mattiucci et al., 2008b). Genetic divergence has been shown to range from K2P = 0.05 (C. osculatum D vs C. osculatum E) to K2P = 0.11 [C. osculatum (s.s.) vs C. osculatum D] between sibling species of the C. osculatum complex. Higher values of K2P were those estimated between morphologically differentiated species such as in the comparison of C. radiatum and the species of C. osculatum complex (on average K2P 0.14), or C. osculatum complex versus C. ogmorhini complex species (on average K2P 0.15) (Mattiucci et al., 2008b) (Table 2.10). However, despite morphologically distinct characters between Phocascaris spp. and the species so far included in the C. osculatum complex, P. cystophorae is genetically very closely related to this group of species (on average K2P 0.10) (Mattiucci et al., 2008b) (Table 2.10) (see also Section 4.3). The Contracaecum mtDNA cox2 gene was found to be A+T rich. This, however, is consistent with that found in other anisakid nematodes, such as Contracaecum parasites of fish-eating birds (Mattiucci et al., 2008c). Similar distance values were those estimated at the same mitochondrial gene (mtDNA cox2) among species of Contracaecum parasitic in fish-eating birds. Indeed K2P ranged from 0.08 between the two sibling species, C. rudolphii A versus C. rudolphii B, to 0.13 between the two sibling species and species morphologically well differentiated (i.e. the C. multipapillatum complex) (Mattiucci et al., 2008c, unpublished data). The genetic divergence among the anisakid taxa so far evaluated at the mtDNA cox2 locus is of the same order as found between other nematode species (Anderson et al., 1998; Blouin et al., 1998; La Rosa et al., 2001; Pozio and Murrel, 2006; Zarlenga et al., 1998).
106
Simonetta Mattiucci and Giuseppe Nascetti
5.2. Genetic differentiation at the intraspecific level and gene flow In Anisakis spp., most of the genetic diversity has been found within populations rather than among populations (see also Section 8.2). Indeed, for instance, at the intraspecific level, the values of DNei found between populations of A. simplex (s.s.) ranged from 0.003 to 0.0038. Such a pattern of genetic distance is related to that of geographical distance (isolation by distance). Average DNei values is 0.007 among Atlantic populations, DNei 0.015 among Pacific ones, and DNei 0.025 between the two groups. The highest value (DNei 0.038) was found between the most distant populations, from the Baltic Sea and off Hokkaido Island, Japan. This finding is mainly a consequence of the differences in allele frequencies at the loci Sod-2, fEst-2, Lap-2 and Pep B in populations of A. simplex (s.s.), compared with those from the other localities (Mattiucci et al., 1997, 1998). At the interpopulation level, A. pegreffii showed the lowest values of Gst (average 0.02, Mattiucci et al., 1997) and DNei is, on average 0.005. This finding indicates remarkable genetic homogeneity of A. pegreffii throughout its wide range, from the Austral region to the Mediterranean Sea. A more consistent intraspecific distance (DNei, on average, 0.045; Gst = 0.10) was found in A. simplex C between the population from the Canadian Pacific and those from Austral region (South Africa, New Zealand, Tasman Sea and sub-Antarctic area) (Table 2.8). This is mainly revealed by the locus Sod-1, which markedly differentiated the sample from the Canadian Pacific and those from the Austral region (Mattiucci et al., 1997). Also in the case of A. typica (s.l.), remarkable genetic homogeneity was observed in larval and adult samples despite being geographically quite distant. Interpopulational genetic distances values detected among populations from Somali, Mediterranean and Brazilian waters were low, ranging from DNei 0004 (eastern Mediterranean vs Somali waters) to DNei 0.010 (Brazilian vs Somali). The average value of Fst among all the populations was 0.04 (Mattiucci et al., 2002a). Similar low levels of population structuring were those found among Boreal and Austral populations of other taxa, such as DNei 0.002 within A. ziphidarum (see Paggi et al., 1998b), DNei 0.008 and DNei 0.009, respectively, in A. brevispiculata and A. physeteris (Mattiucci et al., 2001) and, finally DNei 0.005 in A. paggiae, despite the geographical distance between the samples, indicating high levels of gene flow in all of these Anisakis taxa (Table 2.8). Similar values, at the intraspecific level, were found in the comparison of populations belonging to the sibling species of the P. decipiens complex. The four members of this complex from the Arctic Boreal region showed DNei among populations ranging from, on average 0.001 (in P. krabbei) to DNei 0.005 [in P. decipiens (s.s.)]. Indeed, the lowest genetic heterogeneity was found within P. bulbosa, despite the geographical isolation of the
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
107
populations (NE and NW Atlantic, Pacific) with an average of 0.004. The highest intraspecific differentiation was detected within P. azarasi, for example, the sample from Anadyr (Bering Sea) showed a value of DNei = 0.026 with respect to conspecific Japanese samples (Mattiucci et al., 1998; Paggi et al., 1998c). Both P. decipiens (s.s.) and P. azarasi showed a strict correlation between geographical and genetic distance, whereas higher levels of gene exchange were detected within P. bulbosa. Analogous average values and ranges of genetic diversity between conspecific populations were reported for the members of the C. osculatum complex, from both Arctic and Antartic regions. At the intraspecific level, the values of DNei within C. osculatum A ranged from 0.002 to 0.017, with an average of DNei 0.009. Average DNei values were 0.004 within Northern Atlantic populations, while they were 0.008 among the conspecific populations and 0.018 between the two groups (Mattiucci et al., 1998; Paggi et al., 1998c). This demonstrates a geographical pattern in the genetic variation, although less marked than that observed in A. simplex (s.s.). Among conspecific populations of C. osculatum B, a lower level of differentiation was found DNei 0.005 on average, with a range of values from 0.001 to 0.014, the last value being reported among Pacific and Atlantic populations (Paggi et al., 1998c). A lower degree of interpopulational genetic differentiation was found between populations of the two Antarctic taxa, C. osculatum D and C. osculatum E, detected in definitive and fish hosts from the Weddell and Ross Seas, on average, DNei 0.003 and DNei 0.002, respectively. The highest levels of interpopulational distance between anisakid populations were those found within the otariid parasite C. ogmorhini (s.s.), ranging from DNei = 0.021 to DNei = 0.029 between samples from Argentine, South African and New Zealand waters (on average, DNei = 0.024) (Mattiucci et al., 2003). Similar values are generally found among races in different organisms (Ayala, 1975; Bullini and Sbordoni, 1980). Allozyme data sets have indicated that genetic variation of anisakids is generally not structured in geographical races or subspecies. Such low values of Nei’s D between populations located thousands of kilometres apart are likely to be caused by the homogenizing effects of gene flow, enhanced by the high dispersal capacity of intermediate/paratenic and definitive hosts. Indeed, levels of interpopulational gene flow, indirectly estimated from allele frequencies and Fst values, have been reported in these marine ascaridoid nematodes to be at high levels (Table 2.11). Despite the geographical distance between sampling locations, high rates of gene flow were detected within species belonging to three genera. The highest values were those observed in the species included in the Anisakis simplex complex. As we have illustrated above (Section 3.1), these worms mature in several species of ‘oceanic dolphins’ whose large geographical distribution could maintain the high level of gene flow observed in these anisakid nematodes (Table 2.11). Similarly, the high
108
Simonetta Mattiucci and Giuseppe Nascetti
TABLE 2.11 Values of intraspecific gene flow, estimated in anisakids from the standardized variance of allele frequencies, Fst (Crow and Aoki, 1984; Slatkin and Barton, 1989); m is the fraction of immigrant individuals in a population of effective size N
Species
A. simplex (s.s.) A. pegreffii A. simplex C A. typica A. physeteris A. brevispiculata P. krabbei P. decipiens (s.s.) P. bulbosa C. osculatum A C. osculatum B C. osculatum (s.s.) C. osculatum D C. osculatum E C. ogmorhini (s.s.)
Gene flow (Nm)
8.00 15.00 9.00 6.00 5.10 6.20 4.38 3.66 9.36 5.20 4.90 3.90 4.60 6.10 4.10
References
Mattiucci et al., 1997 Mattiucci et al., 1997 Mattiucci et al., 1997 Mattiucci et al., 2002 Unpublished data Unpublished data Mattiucci et al., 1998; Paggi et al., 1991 Mattiucci et al., 1998; Paggi et al., 1991 Mattiucci et al., 1998; Paggi et al., 1991 Mattiucci et al., 1998; Nascetti et al., 1993 Nascetti et al., 1993 Nascetti et al., 1993 Orecchia et al., 1994, unpublished data Orecchia et al., 1994, unpublished data Mattiucci et al., 2003
mobility and dispersal capacity of their intermediate/paratenic hosts (fish) could enhance the high degree of gene exchange. Such figures are similar to those found for the Arctic-Boreal members belonging to the C. osculatum and P. decipiens complexes maturing in pinnipeds (Table 2.11). These findings confirm that the complex life cycle of these nematodes does not limit gene exchange, but enhances it through the high mobility of the different hosts, such as fish and marine mammals (see also Section 7). On the other hand, the possible correlation between gene exchange and host mobility has also been suggested for other nematodes with different life cycles, as inferred by other genetic markers (Anderson et al., 1998; Blouin et al., 1995, 1998; Criscione et al., 2005).
6. HOST–PARASITE COPHYLOGENY Uncovering the phylogenetic history of anisakid parasites and their hosts represents a steadily advancing part of their systematics and ecology. Thus, cophylogeny mapping has become a recent development in the study of the relationships among and between ecologically related anisakid nematodes with respect to their hosts. Generally, in cophylogeny studies, a dependent phylogeny, such as that of a taxonomic group of
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
109
parasites, is mapped onto an independent one, such as that of their hosts. Co-evolutionary studies mostly rely on morphological or molecular data which are now becoming increasingly available in both parasites and their hosts, allowing detailed comparison of parasite phylogenies with those proposed for their hosts. Results of such studies have demonstrated how the phylogeny of the parasites comes to ‘mirror’ that of their hosts (sensu Eichler, 1948; Fahrenholz, 1913). In cophylogeny mapping, the map is constructed to provide the best possible explanation for the phylogenies and thus determine whether the parasites have co-diverged with their hosts, or whether they have undergone host switching, duplication by independent speciation or even whether parasites had extinction events (Charleston, 2003; Desdevises, 2007). Another method of uncovering ancient associations between dependent and independent taxonomic groups is the ‘biogeographical method’ also known as Brooks’ Parsimony Analysis (Brooks, 1990; Brooks and Hoberg, 1981). In this model, hosts are considered as ‘geographical regions’ and their parasites are their ‘endemic species’, undergoing codivergence in vicariant speciation, loss by extinction, ‘missing the boat’ (sensu Paterson et al., 1993), sympatric speciation and horizontal transfer by migration. Similar investigations have been made into host–parasite associations between anisakid nematodes and their definitive hosts. Phylogenetic trees, mostly inferred from molecular genetic data sets, are now available for these parasites and for some taxonomic groups of their definitive hosts. Comparison of these trees has allowed identification of cophylogenetic events. Mattiucci and Nascetti (2006) have recently provided evidence for such cophylogenetic events in the host–parasite systems of Anisakis spp. and cetaceans (their main definitive hosts). The presence of the two main clades, as presented in the phylogenetic relationships between Anisakis spp. (Section 4.1), is supported also by ecological data and specific host– parasite relationships. Phylogenetic relationships proposed elsewhere and here revised for species of Anisakis ‘mirror’, in several host–parasite associations, that of their definitive hosts (Arnason et al., 2004; Cassens et al., 2000; Milinkovitch, 1995; Nikaido et al., 2001). The sperm whales, Physeter catodon, Kogia breviceps and K. sima are the main definitive hosts for A. physeteris, A. brevispiculata and A. paggiae, respectively (Mattiucci and Nascetti, 2006; Mattiucci et al., 2001, 2005) (Figs. 2.2 and 2.5). Several oceanic dolphins in the Delphinidae, Arctic dolphins in the Monodontidae and porpoises in the Phocoenidae (Table 2.4) are hosts of the species of the A. simplex complex and of A. typica (see Mattiucci et al., 1997, 2002a, 2005). The beaked whales Ziphius cavirostris, Mesoplodon layardii, M. mirus and M. grayi are hosts of A. ziphidarum (see Paggi et al., 1998b) and Anisakis sp. (see Valentini et al., 2006) that are partitioned in clade I in the parasite phylogenetic tree (Fig. 2.3). The phylogeny of cetaceans
Host Bottlenose dolphin
Parasite A. pegreffii 71
F
Short-finned pilot whale
Mage19
E
A. simplex (s.s.)
Isi14 Isi36 Isi36 Mago24 Mago26 Mago32
G
Narwhal
Amz13 Bando1
D
Amazon river dolphin H
60
A. simplex C
Amz11
La plata dolphin
Mago8 Mago13
Yangtze river dolphin
I
C
Tuti24 Tuti35
Mago21 Mago22
A. typica 73
Baird’s beaked whale Mesoplodon sp.
B
A. ziphidarum
76
Anisakis sp.
Sperm8 Sperm28 Sperm47
Ganges river dolphin J
A Bando1 SP316
100
Dall’s porpoise
K Sp9
Sp2
Pygmy sperm whale
Pm72 Pm52 M11
A. paggiae A. brevispiculata
Sperm whale L Hump20 Hump203 aaa792
70
A. physeteris Humpback whale Fin whale
(outgroup) P. ceticola Dolphins
Minke whale
Hosts of Anisakis spp.
Ziphiids Physeterids
FIGURE 2.5 Tanglegram of phylogenies of Anisakis spp. and their cetacean hosts. It shows the phylogeny of a group of extant cetaceans (adapted from Nikaido et al., 2001) mapped into the phylogeny of Anisakis spp. inferred from mtDNA cox2 sequence analysis. Lines depict the observed host–parasite co-speciation events; the dotted line indicates possible host-switching events (redrawn from Mattiucci and Nascetti, 2006).
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
111
representing all extant cetacean families has recently been proposed by Milinkovitch (1995) based on mtDNA genes (12S, 16S and cytb partial sequences) and on myoglobin sequences; by Cassens et al. (2000) inferred from sequence analyses of 12S, 16S and cytb; by Nikaido et al. (2001) based on retroposon analysis and by Arnason et al. (2004) based on analyses of complete mitochondrial genomes. According to the phylogenetic hypothesis proposed by Arnason et al. (2004), the Cetacea group splits into monophyletic Mysticeti (baleen whales) and monophyletic Odontoceti (toothed whales). The Odontoceti diverged into the four extant lineages, Physeteridae (sperm whales: represented by the sperm and pygmy sperm whales), Ziphiidae (beaked whales), Platanistidae (Indian river dolphins) and Delphinoidea (encompassing the families Iniidae, Monodontidae, Phocoenidae and Delphinidae). Phylogenetic trees provided by Nikaido et al. (2001) and Arnason et al. (2004) were congruent in depicting the branching order of the extant cetacean lineages, where the sperm whale and the pygmy sperm whales (belonging to the families Physeteridae and Kogiidae) represent basal taxa, followed by the beaked whales (belonging to the family Ziphidae) and the freshwater and marine dolphins as the most derived (Fig. 2.5). In accordance with those analyses, the branching order so far proposed for the Anisakis taxa showed that nematodes from the sperm whale and pygmy sperm whales (i.e. A. physeteris, A. brevispiculata and A. paggiae) always occupy a basal lineage, always well supported, followed by those parasitizing the beaked whales (A. ziphidarum and Anisakis sp.). The species of the A. simplex complex and A. typica, parasites of delphinoids, are the most derived (Mattiucci and Nascetti, 2006) (Fig. 2.5). Elaboration of these empirical results to assess the global congruence between host and parasite trees gained by ParaFit (Legendre et al., 2002) was statistically significant (P < 0.05). Individual host–parasite associations which contributed more to the cophylogenetic cetacean—Anisakis spp. mapping were represented by those between A. physeteris–Physeter catodon, A. brevispiculata–Kogia breviceps and A. ziphidarum–Mesoplodon spp., suggesting host–parasite co-speciation events. Whereas, a lower significant contribute to the total test was that formed by the host–parasite association A. simplex (s.s.)–Balenoptera acutorostrata, suggesting a possible host-switching event (our unpublished data). Such parallelism between host and parasite phylogenies is also attempted for the group of Contracaecum taxa and their definitive hosts, the pinnipeds of the Families Phocidae and Otariidae. The presence of the two main clades, as presented in the phylogenetic relationships among Contracaecum spp. (Fig. 2.4), is also supported by the ecological data concerning host preference (see also Section 7.1) and specific host–parasite relationships. Phylogenetic relationships so far proposed and here reviewed for species of Contracaecum parallel that recently reported for their definitive hosts based on molecular data (Arnason et al., 1995;
112
Simonetta Mattiucci and Giuseppe Nascetti
Deme´re´ et al., 2003). Several phocid seals (true seals) in the Phocinae are hosts of the species of the C. osculatum complex and Phocascaris spp. in the Arctic Boreal region. Moreover, seals in the Monachinae are hosts for C. mirounga and C. radiatum in the Antarctic and sub-Antarctic region. These anisakids are included in clade I in the parasite phylogenetic tree (Mattiucci et al., 2008b) (Figs. 2.4 and 2.6). Whereas, the otariid species in the Otariinae (sea lions) Zalophus californianus, and in the Arctocephalinae (fur seals) Arctocephalus spp. are hosts of the C. ogmorhini species complex [C. margolisi and C. ogmorhini (s.s.)]. These anisakids are included in clade II in the parasite phylogenetic tree (Mattiucci et al., 2008b) (Figs. 2.4 and 2.6). Although a complete species-level phylogeny for pinnipeds including fossil and extant taxa is as yet, unavailable, a molecular assessment of pinniped relationships was performed by Arnason et al. (1995) using the complete sequences of the mitochondrial cytochrome b gene (mtDNA cytb) of the Phocidae, Odobenidae and Otariidae. Later, Deme´re´ et al. (2003) used a composite tree inferred from the basic topology of generic level, morphological and molecular data, fossil taxa and consensus phylogeny of the phocid subfamilies to propose an integrated hypothesis for pinniped evolutionary biogeography. According to that data elaboration, the Pinnipedia includes three major monophyletic clades: (1) the Otariidae (fur seals and sea lions), (2) the Odobenidae, and (3) the Phocidae (true seals), plus the extinct desmatophocids. In this combined tree, the fur seals and sea lions comprising the Otariinae (Zalophus californianus) and the Arctocephalinae (Arctocephalus spp. from the southern hemisphere), are represented as well-supported basal groups (Arnason et al., 1995; Deme´re´ et al, 2003). In accordance with that analysis, the branching order so far proposed for the Contracaecum taxa showed that nematodes from the Otariidae [i.e. C. ogmorhini (s.s.) from Arctocephalus spp. and C. margolisi from Zalophus californianus] always occupy a basal lineage of the parasite phylogenetic tree, with the species of the C. osculatum complex from the Phocinae (true seals) as the most derived (Fig. 2.6). Speciation of the members of C. osculatum complex is apparently related to their geographical isolation, through that of their hosts, as well as to a rapid host–parasite adaptation and co-evolution. Such processes apparently occurred in different times during the Plio-Pleistocene, when extreme climatic variation took place. The genetic relationships found between the members of the C. osculatum complex suggest that the evolutionary divergence of the most differentiated species [C. osculatum (s.s.)] started more than 3 million years ago, in a Pleistocene refuge (the Baltic Sea). As to the other C. osculatum species, their evolutionary divergence probably took place during the Pleistocene, when the complex achieved a distribution over both polar regions. This process involved two distinct colonizations of the marine Antarctic region by ancestors of the northern hemisphere, giving rise to C. osculatum D and C. osculatum E,
Host P. vitulina vitulina P. vitulina richardsi
97 65
P. largha H. grypus
Parasite C. osculatum B C. osculatum (s.s.)
95
Ph. cystophorae C. osculatum A
P. hispida C. o. baicalensis
90 87
62 94
89
P. fasciata
C. osculatum D
96 99
P. groenlandicus E. barbatus
74
80
L. weddellii
85
C. osculatum E C. radiatum
95
M. schauinslandi H. leptonyx
97
M. leonina
C. mirounga
C. ursinus Z. californianus
100 100
88
91
A. gazella
C. margolisi
100
C. ogmorhini (s.s.)
A. forsteri (outgroup) P. ceticola
97
93 100
O. rosmarus Brown bear Polar bear
American black bear Domestic cat Lion
Hosts of Contracaecum spp.
Phocinae Monachinae Otariidae
Tiger
FIGURE 2.6 Tanglegram of phylogenies of Contracaecum spp. and their pinnipeds hosts. It shows the phylogeny of a group of extant pinnipeds (adapted from Arnason et al., 1995) mapped into the phylogeny of Contracaecum spp. inferred from mtDNA cox2 sequence analysis (adapted from Mattiucci et al., 2008b). Lines depict possible host–parasite co-speciation events; the dotted line indicates possible hostswitching events.
114
Simonetta Mattiucci and Giuseppe Nascetti
both parasites of Leptonychotes weddellii (see Bullini et al., 1997). This hypothesis seems to fit with the evolutionary biogeography of a pinnipedimorph hypothesis based on both dispersal and vicariant events in the context of a species-level phylogenetic framework proposed by Deme´re´ et al. (2003). This hypothesis supports an eastern North Pacific origin during the late Oligocene coincident with start of glaciation in Antarctica. During the late Miocene, pinnipedimorphs remained restricted to the eastern North Pacific, where they began to diversify. Fur seals remained restricted to the North Pacific until the late Pliocene, with a dispersal and rapid speciation in the Southern Ocean during the Pleistocene. The phocine seal diversification took place in the Arctic and North Atlantic during the late Miocene with a subsequent dispersal into the Paratethys and Pacific during the Pleistocene. Finally, the monachine seals, including Mirounga leonina and Leptonychotes weddellii, seem to have the southern hemisphere as the centre of diversification (Deme´re´ et al., 2003). The mode of speciation which apparently fits well with the anisakid nematodes is the peripatric model proposed by Mayr (1963, 1976) (see also Section 5). This involves the geographical isolation of small populations whose genetic structure begins to differ from the parental one by different genetic mechanisms. In the case of the Anisakis spp., and the C. osculatum and P. decipiens species complexes, molecular genetic data strongly suggest that adaptation to different hosts and speciation is related to the geographical isolation of the hosts. Such processes apparently occurred in different times from the lower Miocene to Pliocene, and Pleistocene, when extreme climatic variation took place. During glacial maxima (a period also of lowest sea level), smaller populations of hosts and their endoparasites would have remained isolated in marine refuges, promoting genetic divergence and co-adaptation. Then during interglacial periods, geographical ranges might have expanded, favouring host range expansion (Bullini et al., 1997). Similar co-evolutionary processes have been proposed by Hoberg (1992, 2005) for other host–parasite interactions, involving Holarctic cestodes and their definitive hosts (fish-eating birds and marine mammals).
7. HOST PREFERENCE, ECOLOGICAL NICHE AND COMPETITION Molecular genetic markers of the so far genetically characterized species of anisakid nematodes provide a rapid, precise means to screen and identify several marine mammals, fish and squids that serve as their definitive and intermediate and or/paratenic hosts. These data give important information regarding the life-history traits and epizootiological aspects of these parasites.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
115
Marked differences in definitive and intermediate host preferences were detected among the species belonging to Anisakis, members of Pseudoterranova decipiens complex and species of Contracaecum from pinnipeds. They have shown distinct host preferences, with respect to their definitive and intermediate hosts, often exhibiting differential distribution in various host species and/or with different ecology. Patterns of differential distribution are the product of ecological processes and therefore interactions between parasite populations could promote and even enhance the complexity of multi-species assemblages observed in a given host (definitive and intermediate). Such differences in the life history and host preference between related species of anisakid nematodes appear to be due to differential host–parasite co-adaptation and co-evolution as well as to interspecific competition. This acts either in reducing the range of potential hosts or promoting a differential use of resources from a single individual host in sympatric conditions. In other words, the realized niche in some anisakid species could be more restricted than the fundamental niche. Some examples of such ecological processes have been found in anisakids of the P. decipiens complex and C. osculatum complex and are described below. In the North Atlantic and Pacific Oceans (Fig. 2.7), P. bulbosa is the only species present in the bearded seal, Erignathus barbatus, both in the eastern and western parts of the Atlantic Ocean, and also in the Pacific (Table 2.5), whereas in the North Eastern Atlantic waters P. decipiens (s.s.) and P. krabbei occur sympatrically as adults and often also syntopically in the same individual host in the common seal, Phoca vitulina, and in the grey seal, Halichoerus grypus (see Mattiucci et al., 1998; Paggi et al., 1991). However, statistically significant differences in relative numbers have been observed between P. decipiens (s.s.) and P. krabbei identified by genetic markers (allozymes) in those hosts, where P. decipiens (s.s.) predominates about tenfold in the common seal, while P. krabbei is the main prevalent species in the grey seal. On the other hand, in the western Atlantic, where P. krabbei is absent, P. decipiens (s.s.) occurs in equal proportions in both grey and common seals (Nascetti, 1992; Paggi and Bullini, 1994) (Fig. 2.7). This finding was also observed by Brattey and Stenson (1993), using the same genetic markers (diagnostic allozymes), to identify to species level anisakid nematodes in seal hosts from Newfoundland and Labrador. The bearded seal was found to be parasitized by P. bulbosa (under its former name, P. decipiens C, by Brattey and Stenson, 1993), while both the grey and common seal were found heavily infected by P. decipiens (s.s.) (under its former name, P. decipiens B, by Brattey and Stenson, 1993). This finding could reflect differential adaptation of the two parasites, which have evolved allopatrically. After their secondary contact and, consequently, their sympatric occurrence in the North East Atlantic seal hosts, interspecific competition between the two
North Atlantic Ocean
Halichoerus grypus
Erignathus barbatus
Pagophilus groenlandicus
Pseudoterranova bulbosa
Contracaecum osculatum A
Pseudoterranova decipiens (s.s.)
Contracaecum osculatum B
Pseudoterranova krabbei
Contracaecum osculatum (s.s.)
Phoca vitulina
FIGURE 2.7 Relative proportions of Pseudoterranova decipiens and Contracaecum osculatum species complexes in definitive hosts (seals) from the North Atlantic Ocean. It shows differences in their definitive host preferences.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
117
Pseudoterranova species could have led to a niche subdivision, evolving by character displacement (Bullini et al., 1997; Nascetti, 1992). Another example of niche subdivision is represented by the two Antarctic members of the C. osculatum complex, that is, C. osculatum D and C. osculatum E. The two sibling species share the same definitive host, the Weddell seal, Leptonychotes weddellii, in both the Weddell and Ross Seas, of Antarctica. However, the two anisakid species appear to be adapted in this sympatric area to pelagic and benthic food webs, respectively. Larval individuals (n = 1306) of C. osculatum D and C. osculatum E, genetically identified to the species level by allozymes, were found to be heavily infecting fish species in the Ross Sea (Mattiucci and Nascetti, 2007, unpublished data). Statistically significant differences were detected in the relative proportions of the two Antarctic species of the C. osculatum complex in benthic and mesopelagic fish intermediate host species. Indeed, C. osculatum D outnumbers C. osculatum E in channichthyid fish (i.e. Chionodraco hamatus, Pagetopsis macropterus) and in bathydraconids (i.e. Gymnodraco acuticeps, Cygnodraco mawsoni); while, higher relative proportions of the infection by C. osculatum E were observed in nototheniid fish (Trematomus bernacchii, T. pennellii, T. newnesi and Notothenia neglecta) (Fig. 2.8). The last group of nototheniid fish are bottom living and feed on benthic invertebrates, such as benthic amphipods (Daniels, 1982; Foster and Montgomery, 1993; Vacchi et al., 1994), whereas channichthyid and bathydraconid fish have a pelagic/mesopelagic mode of life (Daniels, 1982). In the sibling species C. osculatum D and E, competition in the intermediate hosts could be avoided by spatial separation in the pelagic and benthic components of food webs in the Antarctic marine ecosystem. On the other hand, Klo¨ser et al. (1992) suggested that C. radiatum in Antarctic waters uses a pelagic food web. This was also confirmed by the genetic identification of larval stages of C. radiatum, at a high prevalence, in pelagic channichthyid fish (i.e. Chionodraco hamatus) (Arduino et al., 1995). Life-history differences in anisakid nematodes genetically recognized at the species level were also reported in the case of the P. decipiens species complex. In the northern hemisphere, P. bulbosa is a parasite at adult stage of the bearded seal both in the North Atlantic and Pacific Oceans, and at larval stage it occurs in the flatfishes Hippoglossoides platessoides, Reinhardtius hippoglossoides and Hippoglossus hippoglossus, in Myoxocephalus quadricornis, and, rarely, in Gadus morhua macrocephalus (see Brattey and Davidson, 1996; Bristow and Berland, 1992; Mattiucci et al., 1998). The hosts so far detected for P. bulbosa are specialized benthic species, with the exception of G. morhua which is benthopelagic (Paggi et al., 1991, 1998c). This supports the hypothesis that P. decipiens (s.l.) in the northern hemisphere has a benthic life cycle (Bristow and Berland, 1992). A similar benthic life-history pathway can be suggested for P. azarasi in Pacific
118
Simonetta Mattiucci and Giuseppe Nascetti
C. osculatum E C. osculatum D Notothenia neglecta
Bentihic
Trematomus bernacchii Trematomus pennelli
Gymnodraco acuticeps Cygnodraco mawsoni Cryodraco antarcticus Chionodraco hamatus Chaenodraco wilsoni Pagetopsis macropterus 0
10
20
30
40
50
60
70
80
90
100
Pelagic and mesopelagic
Trematomus newnesi
FIGURE 2.8 Relative proportions of larval specimens of the Antarctic anisakids Contracaecum osculatum D and C. osculatum E, genetically identified from fish hosts. It depicts differential distribution of the two species in pelagic, mesopelagic and benthic fish hosts respectively, in Antarctica.
waters. It was detected genetically at the adult stage in the benthic feeding Steller’s sea lion, Eumetopias jubatus, and also in mixed infections with P. bulbosa in the bearded seal; and at larval stage in the flatfish Hippoglossus hippoglossus (see Mattiucci et al., 1998; Paggi et al., 1998c). Benthic and benthopelagic fish species (i.e. Chaenocephalus aceratus, Notothenia coriiceps, N. neglecta, Trematomus newnesi) are found infected by the Antarctic taxon P. decipiens E (our unpublished data), confirming the suggestion of Palm et al. (1994), according to which P. decipiens (s.l.) completes its life cycles in the Weddell Sea environment in organisms of the benthic food web. Interactions in seals between anisakid species of different genera are likely to exist, since they often co-occur in the same host individual. As anisakine nematodes become numerous in seal stomachs, they may be increasingly prone to the consequences of intra- and inter-specific competition. These effects could provoke high mortality rates in one species, retard its maturity or result in lower uterine egg counts in female worms. McClelland et al. (1985) and Burt (1994) speculated that P. decipiens (s.l.) abundance in seal stomachs may be limited as a consequence of ‘competitive exclusion’ by the related nematode C. osculatum (s.l.). McClelland et al. (1985) suggested that C. osculatum (s.l.) interfered with the proliferation of P. decipiens (s.l.), since seasonal and geographical reductions in the numbers of P. decipiens (s.l.) in grey seals often coincided with increases in the abundance of C. osculatum (s.l.). This negative correlation between the abundance of the two anisakid nematodes was not supported by the data
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
119
of Brattey and Stenson (1993) according to which, these changes could be equally well due to differences in diet among grey seals, as these tend to acquire infection with larval P. decipiens (s.l.) by preying more on benthic fish (McClelland et al., 1990); whereas others feed more on pelagic fish species, such as capelin, Mallotus villosus, or cod, Gadus morhua, that harbour larvae of C. osculatum (s.l.) (see Pa`lsson and Beverly-Burton, 1984). Marcogliese (1997), remarking on the relatively small size and low uterine egg counts of female P. decipiens (s.l.) in grey seals from Anticosti Island, suggested that P. decipiens may be suppressed as a consequence of competitive pressure from C. osculatum. McClelland (2002) suggested, however, that significant negative correlation between the abundance of the two nematode species, P. decipiens and C. osculatum, could result when samples taken in a given area also include migrant seals acquiring infections in other areas. However, when members of the P. decipiens and C. osculatum complexes infest the same seal host, a differential use of resources could take place, as the former occupy the seal intestine when the stomach (which is probably the suitable microhabitat for both anisakids) is highly parasitized by C. osculatum (s.l.) (see Berland, 1964; Klo¨ser et al., 1992; Paggi and Bullini, 1994; Mattiucci and Nascetti, unpublished data). Berland (1964) speculated that C. osculatum competes with and displaces Phocascaris spp. from the stomach of harp Pagophilus groenlandicus, and hooded seal Cystophora cristata. This result was later confirmed by genetic identification (allozymes) of several individuals of C. osculatum B and Phocascaris spp. It is likely that C. osculatum B is a better competitor than Phocascaris phocae or P. cystophorae during syntopic occurrence, at the adult stage, in the same individual seal host. Marked differences were indeed observed in their relative proportions when they occur syntopically in the harp and common seals. In these cases, the stomach is mostly occupied by C. osculatum B, while P. phocae and P. cystophorae are displaced to the upper intestine. In contrast, the hooded seal has always been found parasitized solely by P. cystophorae/P. phocae in both sites (stomach and intestine). In this seal, Phocascaris spp., in the absence of a better competitor, usually occupies the stomach. These phenomena of niche subdivision, promoting character displacement, could reduce the inter-specific competition between two closely related anisakid species (i.e. C. osculatum B and P. phocae/P. cystophorae) for a trophic resource (Nascetti, 1992; Paggi and Bullini, 1994).
8. ANISAKIDS AS BIOLOGICAL INDICATORS In recent years, marine parasites have been widely used in biological and ecological surveys of marine ecosystems as biological indicators of food chain structure (Thompson et al., 2005), pollution (Sures, 2004), global
120
Simonetta Mattiucci and Giuseppe Nascetti
climate changes (Brooks and Hoberg, 2007; Poulin, 2006), anthropogenic impacts and environmental stresses (Marcogliese, 2005), fish stock assessment (MacKenzie, 2002) and of general ‘ecosystem health’ (Marcogliese, 2005). Anisakid parasites are an integral part of aquatic ecosystems. They are indeed important components of any ecosystem. Not only do they play key roles in population dynamics and community structure, they can provide important information about the general biodiversity of the ecosystem. Recent data on the possible use of anisakid nematodes are presented in this section as biological indicators of (i) the definition of fish stocks within a multidisciplinary approach, (ii) integrity and stability of trophic webs and (iii) habitat disturbance.
8.1. Anisakis spp. larvae as biological tags of fish stocks One of the major goals of the last decade in the study of the parasite fauna of aquatic organisms has been, among the others, the assessment of fish stocks, their movement and recruitment. Indeed, the use of parasites to discriminate fish host populations has been one of the most useful approaches in a multidisciplinary study of fish stock detection and characterization. Nowadays, the modern concept of ‘fish stock’ integrates all the information gathered from a broad spectrum of techniques, ranging from morphology (morphometrics, meristic, otolith microchemistry) to biology (life-history characteristics, mark-recapture, parasites) and genetic structure (allozymes, microsatellites, DNA sequences) of the fish hosts throughout their geographical range. In this ‘holistic approach’ to the definition of fish stock (Begg and Waldman, 1999), among the biological approaches, the use of parasites as ‘biological tags’ has become a useful tool, mainly concerning species with a high commercial value in fisheries. Trends in the use of parasites as biological tags in population studies of marine fish species was recently reviewed (MacKenzie, 2002). Among the parasite species that have been used in fish stock definition, the larval anisakid nematodes of the genus Anisakis represent one of the best biological tags (MacKenzie, 2002). Allozyme markers were first used to differentiate populations of herring, Clupea harengus L., in the North Atlantic, on the basis of different allele frequencies observed at one enzymatic locus in Anisakis larvae (Beverly-Burton et al., 1977). The application of diagnostic genetic markers in the identification of larval Anisakis to species level, and the biogeographical aspects of Anisakis spp., have, in recent years, allowed the fish stock identification of several demersal and pelagic species (Mattiucci et al., 2007a). A parasitological survey carried out on several fish specimens of the European hake, Merluccius merluccius, from 14 localities in the North East Atlantic and
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
121
Mediterranean Sea included in the range of distribution of the fish species, has allowed detection, by allozyme markers, of the occurrence of different biological species of larval Anisakis (n = 1950). Indeed, seven species of Anisakis were genetically recognized [A. pegreffii, A. simplex (s.s.), A. typica, A ziphidarum, A. physeteris, A. brevispiculata and A. paggiae] showing statistically significant differences in their distribution in the hake sampled from Mediterranean and Atlantic waters (Mattiucci et al., 2004) (Fig. 2.9). This indicated that (i) two stocks of M. merluccius from Mediterranean and Atlantic waters exist; (ii) in the North Atlantic area, at least two different population units of European hake are present; one northern and another from off the Atlantic coast of Morocco; and (iii) some sub-structuring in the Mediterranean population was also recognized (Mattiucci et al., 2004). These findings are in accordance with genetic results obtained from allozyme markers on the same fish host samples, demonstrating that Atlantic and Mediterranean populations of M. merluccius belong to two different stocks. These two populations are separated at the Almeria-Oran front (Cimmaruta et al., 2005), as this front has been proved to be the boundary between Atlantic and Mediterranean stocks of several marine organisms (Pannacciulli et al., 1997; Rios et al., 2002). In this area a strong inflow transports surface Atlantic waters into the Albora´n Sea, creating an Atlantic domain within the Mediterranean, and a partial isolation of the Albora´n with respect to the other Mediterranean water masses. It was also demonstrated that salinity and temperature are responsible for maintaining the genetic differentiation between the two fish population groups through selective processes (Cimmaruta et al., 2005). In the case of horse mackerel, Trachurus trachurus, different populations can be distinguished according to the spatial distribution of the two most dominant parasite species, genetically recognized as A. simplex (s.s.) and A. pegreffii. Indeed, the significant differences indicated by their relative proportions between the samples from the Mediterranean Sea (excluding the Albora´n Sea) in comparison with the Atlantic ones seem to indicate the discrete sub-structuring of populations of T. trachurus in Mediterranean and Atlantic waters. Moreover, in Atlantic waters, the differences found in the proportion of A. pegreffii between the southern horse mackerel samples and those remaining seem to suggest a possible existence of a ‘southern’ sub-population of T. trachurus readily distinguished from a ‘northern’ one. The samples of T. trachurus from the Albora´n Sea were closer to the southern populations of the Atlantic than to the other samples from the Mediterranean Sea. This finding, and particularly the presence of A. simplex (s.s.) in some individual hosts from the Albora´n Sea, suggests that these populations could be the result of a migration of this small pelagic fish species from Atlantic waters into this extreme western area of the Mediterranean Sea, and also possibly
122
Simonetta Mattiucci and Giuseppe Nascetti
A
Merluccius merluccius
B
Trachurus trachurus
C
Xiphias gladius
A. simplex (s.s.) A. pegreffii A. brevispiculata A. physeteris A. ziphidarum A. typica A. paggiae Anisakis sp.
50 45 40 35 –50 –45 –40 –35 –30 –25 –20 –15
50 45 40 35 –50 –45 –40 –35 –30 –25 –20 –15 –10 15 10 5 0 –30
–25
–20
–15
–10
–5
FIGURE 2.9 Relative proportions of larval specimens of Anisakis spp. from different fish hosts: (A) Merluccius merluccius; (B) Trachurus trachurus; (C) Xiphias gladius throughout their range of distribution. It shows the use of biogeographical aspects of Anisakis spp., genetically identified to species level, as biological tags for their fish stocks definition in European waters.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
123
mixing with Mediterranean populations. This, in turn, is related to the possible Almeria–Albora´n boundary in Mediterranean waters. Furthermore, the high percentage of fish exhibiting infection by A. simplex (s.s.) from this area was in accordance with the high dispersal capacity of this small pelagic fish (Mattiucci et al., 2008a) (Fig. 2.9). This is in agreement with the high genetic variability values shown for T. trachurus, at the nuclear level (allozymes), in the same fish samples (Cimmaruta et al., 2008). However, in contrast with the parasitological findings, a substantial genetic homogeneity, inferred from allozyme markers on the same fish samples, was found between the Atlantic and Mediterranean populations of horse-mackerel; a partial sub-structuring of T. trachurus observed between the far eastern and western Mediterranean Sea has been reported (Cimmaruta et al., 2008). As for the swordfish, Xiphias gladius, it has been recently demonstrated, using different molecular genetic markers and biological data, that the Mediterranean population is separated from the Atlantic one (Cimmaruta et al., 2006; Kotoulas et al., 2006; Lu et al., 2006; Reeb and Block, 2006). The existence of a further separation between northern and southern populations of swordfish in the Atlantic has been also suggested (Alvarado-Bremer et al., 2006). The occurrence of different species of Anisakis spp. larvae in Mediterranean and Atlantic swordfish populations was recently shown (Mattiucci et al., 2007a). In particular, the absence of A. simplex (s.s.) in the Mediterranean swordfish samples seems to support the idea that the Mediterranean population of X. gladius is separated from the Atlantic one. Moreover, in Atlantic waters, the possible existence of two populations, one northern and one southern, has been suggested (Mattiucci et al., 2007a) (Fig. 2.9). This is also supported by population genetics data obtained from the swordfish sampled in this area, indicating a possible boundary of the two stocks between 0 N and 5 S in Atlantic waters (Alvarado-Bremer et al., 2006). Recently, it has been shown that the polymorphism observed in mtDNA cox1 gene sequences, analysed for several samples of A. simplex (s.s.) collected between and within spawning seasons of its fish host (the herring, Clupea harengus), exhibited a high degree of temporal stability (Cross et al., 2007). This underlines the potential use of this species as a biological tag, and mtDNA cox1 as a suitable genetic marker for future investigation in fish host stock structure definition (Cross et al., 2007).
8.2. Anisakids as indicators of trophic web stability and habitat disturbance of marine ecosystems Food webs are networks of trophic relationships which map the location of energy flow in a community. Despite the increasing details that have been incorporated into marine food webs, some functional groups remain
124
Simonetta Mattiucci and Giuseppe Nascetti
neglected. Among these are the parasites and host–parasite interactions that are known to affect, among others, community structure and trophic relationships (Marcogliese, 2001, 2005; Poulin and Morand, 2004). The transmission pathways of parasites with indirect life cycles are fully included in food webs of aquatic ecosystems. In other words, just as food webs have exerted strong selective pressure on the evolution of parasite transmission strategies, parasites are now shaping some of the ecological properties of existing food webs. The transmission routes of anisakid nematodes follow closely the trophic relationships among their successive hosts, and thus they are parasites embedded in food webs. As a consequence, the completion of life cycles as complicated as those of anisakid nematodes requires a stable trophic web. As a result, the life cycle of anisakid nematodes in marine ecosystems with various degrees of habitat disturbance could be affected by changes in host population size. Indeed, when the population size of the hosts participating in the life cycle of these parasites is reduced due to different causes (pollution, by-catch of marine mammals, viral diseases of marine mammals, overfishing, etc.) (Fig. 2.10), the population size of their anisakid endo-parasites could also be reduced. This would result in a higher probability of genetic drift in the parasite gene pools and, consequently, a decrease in their genetic variability values. It has recently been shown (Mattiucci and Nascetti, 2007) that the distribution of the genetic variability of anisakid nematode populations in geographical areas with different levels of environmental stress is likely to reflect the influence of a range of factors that could promote their genetic diversity. These include a large effective parasite population size, the wide range, availability, and population size of their hosts, and the stability of marine trophic webs. The values of the genetic variability [estimated at the parameters of percentage of polymorphic loci (P), mean number of alleles per locus (A) and expected heterozygosity per locus (He)], obtained at 19 allozyme loci, were compared among 53 populations of anisakid nematodes belonging to 20 species of Anisakis, Pseudoterranova and Contracaecum from several hosts in the Boreal and Austral regions (Mattiucci and Nascetti, 2007). Austral populations of species belonging to these three genera exhibited significantly higher genetic variability values than those from the Boreal regions [expected mean of heterozygosity per locus, He = 0.19 (in Austral populations) and He = 0.09 (in Boreal populations) (P < 0.01)] (Fig. 2.11). A more remarkable difference in their genetic variability values was observed when only Antarctic and subAntarctic populations were compared directly with Arctic and sub-Arctic populations [He = 0.23 and He = 0.07 (P < 0.001), respectively] (Mattiucci and Nascetti, 2007). One conclusion is that the observed values of genetic variability could be related to extreme latitudes, a parameter often considered as relevant (Nevo et al., 1984). However, the data suggested that a
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
Host
Definitive
Intermediate/ paratenic
First intermediate
125
Habitat disturbance By-catch and hunting, viral diseases
Overfishing, contaminants
Global warming, sea water acidification, pollution
FIGURE 2.10 Relationship between habitat disturbance and host population size. It depicts causes of habitat disturbance (right) that can affect the population size of definitive and intermediate hosts (at different trophic levels of a marine food web) involved in speculative life-cycle pathways of anisakid nematodes of the genera Anisakis, Pseudoterranova and Contracaecum (includes only species from pinnipeds). (Modified from Mattiucci and Nascetti, 2007, with permission from Elsevier.)
significantly higher level of genetic variability found in the Antarctic members considered (i.e. C. osculatum D, C. osculatum E, C. radiatum, P. decipiens E, A. simplex C and A. pegreffii populations from sub-Antarctic regions) coincide with a lower degree of habitat disturbance (e.g. overfishing, by-catch of cetaceans, hunting and diseases mortality of seals, sea water pollution and acidification). This would allow host species to reach higher population sizes, resulting in higher anisakid population sizes, with a reduced probability of genetic drift phenomena in the parasite gene pools. Consequently, a higher level of genetic diversity in the Antarctic populations of these nematodes was observed. Likewise, a much higher abundance and intensity of infection was observed in the Antarctic and sub-Antarctic populations and species of anisakid nematodes, where more than 105 individual worms were typically collected from a single host, compared to <102 from Arctic and sub-Arctic hosts (Beverly-Burton, 1971; Brattey and Stenson, 1993;
60⬚
ircle olar C Arctic P
He = 0.07
A. ziphidarum Mediterranean Sea
P. azarasi NW Pacific
C. osculatum A NW Atlantic
C. osculatum B NE Atlantic
A. physeteris CE Atlantic
Boreal populations
A. brevispiculata CE Atlantic
A. pegreffii Mediterranan Sea
C. osculatum A NE Atlantic
A. typica CE Atlantic
A. physeteris Mediterranean Sea
P. bulbosa NW Pacific
C. osculatum (s.s.) Off Iceland
C. osculatum (s.s.) Baltic Sea
A. simplex (s.s.) W Mediterranean Sea
A. simplex (s.s.) Spanish Atlantic Coast
C. margolisi NE Pacific
P. bulbosa NW Atlantic P. bulbosa Barents Sea
P. decipiens NE Atlantic
He 0.02–0.03 0.04–0.05
P. decipiens NE Pacific
C. osculatum B Barents Sea
A. simplex (s.s.) Baltic Sea
C. osculatum B NW Atlantic
A. simplex (s.s.) NW Atlantic
P. decipiens (s.s.) NW Atlantic
P. krabbei NE Atlantic
A. pegreffii Spanish Atlantic Coast
C. osculatum A Barents Sea
A. simplex (s.s.) NE Atlantic
0.06–0.07
0.08–0.09
0.10–0.11
A. ziphidarum Off South Africa
A. brevispiculata Off South Africa
30⬚
A. simplex C NE Pacific
0.12–0.13
0.15–0.16
0.17–0.18
0.20–0.21
0.23–0.24
0.29–0.30
0.32–0.33
A. pegreffii Falklands Islands
C. ogmorhini (s.s.) Off New Zealand
C. ogmorhini (s.s.) Off Argentina
C. osculatum D Weddell Sea
A. simplex C New Zealand
C. osculatum E Ross Sea
C. radiatum Weddell Sea
C. osculatum D Ross Sea
C. radiatum Ross Sea
A. pegreffii New Zealand
P. decipiens E Antarctic
A. simplex C Antarctic
A. typica Brazilian Coast
C. ogmorhini (s.s.) Off South Africa
C. mirounga Antarctic
A. pegreffii Off South Africa
C. osculatum D Ross Sea A. simplex C Off South Africa
Austral populations
He = 0.19
60⬚
Antarcti c
120⬚
60⬚
0⬚
60⬚
Polar C
ircle
120⬚
180⬚
FIGURE 2.11 Distribution of the genetic diversity (expected mean heterozygosity per locus, He) (in white coloured cells) among anisakid populations from Boreal (in light grey coloured cells) and Austral regions (in dark grey coloured cells). The arrows represent the mean values of He in Boreal and Austral populations. The genetic variability of anisakid populations is higher in the Antarctic region than in other geographical areas, whether they be similar or ecologically differentiated. Such difference can be explained by the lower habitat disturbance of the Antarctic region, which permits the maintenance of more stable trophic webs in this ecosystem (redrawn from Mattiucci and Nascetti, 2007).
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
127
Klo¨ser et al., 1992; Mattiucci and Nascetti, 2007; McClelland et al., 1983) (Tables 2.12 and 2.13). The data are consistent with biotic factors, such as host density of those suitable definitive and intermediate hosts for the anisakid nematodes in the Antarctic and sub-Antarctic areas, maintaining the high genetic diversity in the anisakid gene pools. In support of this hypothesis, a high density of marine mammal populations has been documented in these geographical areas (Wickens, 1995), relative to populations from the Arctic-Boreal regions (Read et al., 2006). For example, in Antarctic waters, the Weddell seal (main definitive host of C. osculatum D and C. osculatum E, see Section 3.3) has not suffered a reduction in its population numbers in recent years (Kendall et al., 2003). This is contrary to the phocids of the Arctic–Boreal region, whose population size has decreased due to different causes of habitat disturbance (hunting, by-catch, viral diseases, etc.) (Andersen et al., 2006; Barron et al., 2003; Di Guardo et al., 2005; Pastor et al., 2004). Similarly, the Antarctic fish species examined here and utilized as intermediate hosts by these nematodes have not been subjected to the population reduction from over-fishing that has occurred in the host fish species from the Boreal region (i.e. Gadus morhua) (Anon, 1997; Cook et al., 1997). Various causes of habitat disturbance (pollution, capture, overfishing, global climate change) possibly place undue stress on the trophic webs involving different species of marine mammals, fish and invertebrates (suitable as definitive and intermediate hosts of anisakid nematodes). This in turn could adversely affect the population size of their endoparasitic anisakid nematodes and, consequently, their genetic variability. Small anisakid nematode burdens tend to have a lower genetic diversity, as smaller populations lose diversity more quickly than large populations, due to genetic drift. Large populations of anisakid nematodes, such as those from the Antarctic, show higher levels of genetic diversity. These data fit well into the pattern which has recently been demonstrated for a range of other parasitic nematode species, showing that low levels of genetic diversity are often positively associated with low levels of infection (Webster et al., 2007). It is likely that elevated host density in the Antarctic and sub-Antarctic areas will lead to an increase in anisakid parasite prevalence and abundance in both suitable definitive and intermediate hosts. The high levels of parasitic infection reported for the C. osculatum (s.l.) complex in Antarctic fish species, which are prey for the Weddell seal, are consistent with the high integrity and stability of the food webs in this marine ecosystem. This, in turn, facilitates the completion of the life cycles of Antarctic and sub-Antarctic anisakid nematodes (Mattiucci and Nascetti, 2007). These results support the hypothesis that parasite infections give indications about the hosts that are available to them (Hudson et al., 2006; Poulin, 2006).
128
Simonetta Mattiucci and Giuseppe Nascetti
TABLE 2.12 Prevalence (P, %), Intensity (I) and Abundance (A) of the infection with the species Contracaecum osculatum (s.l.) in intermediate/paratenic (fish) and definitive (pinnipeds) hosts from the Boreal and Antarctic regions Host species/ geographic origin
Intermediate host Boreal region Gadus morhua (NE Atlantic) Hippoglossoides platessoides (NE Atlantic) Antarctic region Gymnodraco acuticeps (Ross Sea) Cygnodraco mawsoni (Ross Sea) Cryodraco antarcticus (Ross Sea) Chionodraco hamatus (Ross Sea) Trematomus bernacchii (Ross Sea) Definitive host Boreal region Pagophilus groenlandicus (NE Atlantic) Halichoerus grypus (NE Atlantic) Phoca vitulina (NE Atlantic) Erignathus barbatus (NE Atlantic) Antarctic region Leptonychotes weddellii (Weddell Sea) Leptonychotes weddellii (Ross sea)
Parasite
P
I (min–max)
A
C. osculatum B
50
(4–20)
4.0
C. osculatum A
26.0
(2–10)
1.2
C. osculatum D
100.0
(53–554)
308.7
C. osculatum D
100.0
(60–520)
210.0
C. osculatum D
100.0
(86–619)
342.8
C. osculatum D
100.0
(30–209)
105.1
C. osculatum E
100.0
(20–350)
110.2
C. osculatum B
97.7
(13–2351)
321.9
C. osculatum B
80.7
(44–2783)
528.5
C. osculatum B
100.0
(1–543)
76.8
C. osculatum A
100.0
(48–1223)
313.7
C. osculatum (s.l.)
100.0
(20671– 130200)
42270.2
C. osculatum (s.l.)
100.0
(6300– 32500)
25310.1
Modified from Mattiucci and Nascetti (2007, with permission from Elsevier).
129
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
TABLE 2.13 Prevalence (P,%), intensity (I) and abundance (A) of parasitic infection levels by Anisakis pegreffii in different definitive and intermediate hosts from the Boreal and Austral regions Host species/geographic origin
Definitive host Boreal region Tursiops truncatus Stenella coeruleoalba Delphinus delphis Austral region Caperea marginata Cephalorhynchus haevinsedi Tursiops truncatus Intermediate host Boreal region Trachurus trachurus Merluccius merluccius Lepidopus caudatus Engraulis encrasicolus Micromesistius poutassou Austral region Merluccius capensis Lepidopus caudatus
P
I (min–max)
A
Mediterranean Sea
10.0
(1–15)
10.5
Mediterranean Sea
2.0
(1–30)
15.0
Mediterranean Sea
2.0
(1–22)
20.0
SE Atlantic Ocean (South Africa coast) SE Atlantic Ocean (South Africa coast) SE Atlantic Ocean (South Africa coast)
–
–
100.5
–
120.4
–
–
250.9
Mediterranean Sea
44.4
(1–50)
10.5
Mediterranean Sea
85.0
(1–130)
25.0
Mediterranean Sea
80.0
(10–120)
20.0
Mediterranean Sea
17.7
(1–10)
1.8
Mediterranean Sea
48.5
(1–12)
5.1
SE Atlantic Ocean (South Africa coast) SE Atlantic Ocean (South Africa coast)
98.0
(10–130)
90.5
100
(30–250)
212.0
(continued)
130
Simonetta Mattiucci and Giuseppe Nascetti
TABLE 2.13
(continued)
Host species/geographic origin
P
I (min–max)
A
Etrumeus whiteheadi
100
(20–120)
80.8
100
(20–360)
205.1
95.0
(30–170)
90.5
Thyrsites atun
Merluccius hubbsi
SE Atlantic Ocean (South Africa coast) SE Atlantic Ocean (South Africa coast) SW Atlantic Ocean (Falklands Islands)
Modified from Mattiucci and Nascetti (2007, with permission from Elsevier).
The Antarctic ecosystem is generally considered ‘pristine’ (Battaglia et al., 1997). Existing results on the genetic variability of anisakid nematodes have shown that, despite the ‘extreme’ ecological conditions of marine Antarctic ecosystems, the genetic diversity of the host–parasite system formed by ascaridoid worms in fish and marine mammals is higher in the Antarctic region than in other geographical areas whether they be similar or ecologically differentiated. Such difference can be explained by the lower habitat disturbance of the Antarctic region, which permits the maintenance of more stable trophic webs in this ecosystem (Mattiucci and Nascetti, 2007).
9. CONCLUSIONS AND IDENTIFICATION OF GAPS IN OUR KNOWLEDGE OF ANISAKIDS TO BE FILLED BY FUTURE RESEARCH The objective of this review was to compile current knowledge on the anisakid species of Anisakis, Pseudoterranova and Contracaecum acquired since the introduction of molecular markers as an aid to their systematics. Thus, we have attempted to detail the advances in their taxonomy, epidemiology, geographical distribution, population genetics and phylogenetic relationships, highlighting ecological implications and co-evolutionary processes. New insights into our knowledge of anisakids have been developed, but several questions still need to be addressed. This section thus highlights some of the main gaps which, in our opinion, will present new perspectives and opportunities for future research on anisakid nematodes.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
131
9.1. Molecular systematics From a taxonomic point of view, we have collected examples of, and evidence for, criteria useful for the species diagnosis of anisakid nematodes. Describing reproductive isolation is crucial for speciation and the use of allozyme markers, and (by MAE) is particularly useful for this aim because (1) they are co-dominant markers (homozygotes are distinguishable from heterozygotes), (2) the polymorphisms which they can reveal are representative of the genetic variation of an anisakid population, and (3) they allow a more complete understanding of Mendelian inheritance of these polymorphisms. Starting from these landmarks we have seen that, in the case of anisakid species, MAE could be the best choice, as a first genetic approach, to define the taxonomic status of a species following the BSC, because they are able to demonstrate the absence of gene flow and, thus, the reproductive isolation between taxa. They are also able to distinguish possible F1 hybrids between related taxa and/or introgressed individuals, where other molecular markers (for instance, RAPDs and/or other not co-dominant markers) may fail to demonstrate them. Moreover, it has been shown that once anisakid species have been designated by allozymes, phylogenetic systematics inferred from other molecular markers (both nuclear and mitochondrial) were congruent and corroborated those results. On the other hand, phylogenetic systematics of anisakid nematodes based on DNA gene sequence analysis has been shown, in most cases and with few exceptions, to be congruent with allozyme data sets establishing their taxonomic status and depicting phylogenetic relationships between related taxa of anisakids. It has also been useful for comparison with phylogenies postulated for their definitive hosts. Among the PCR-DNA methods, the mitochondrial gene of the cytochromoxidase 2 (mtDNA cox2) has been demonstrated to be useful both for species identification distinguishing all the anisakid taxa so far detected, and also for their phylogenetic study. This also underlines the utility of being able to infer phylogenetic hypotheses for anisakid nematodes based on data concerning large and varied characters. Thus, allozymes from MAE and/or DNA-based markers (barcodes) are recommended as rapid and precise tools for the identification of anisakid nematodes and for their cataloguing (barcoding). In the near future, this will allow the comparison of new samples and taxa with deposited DNA material and sequences (obtained from well-established taxa of anisakids) as well as from type-material in museums, to avoid the future synonymy of anisakid taxa. It is also expected to have, as much as possible, an integrated approach in the study of anisakid nematode systematics, with a reconciliation of genetics with morphology, after the genetic recognition of a biological species. Indeed, in some of the anisakids detected genetically, the formal
132
Simonetta Mattiucci and Giuseppe Nascetti
description of the species is still lacking (i.e. for the C. osculatum species complex). Molecular systematics will in the future clarify the taxonomic status of some species still waiting to be genetically characterized among those considered as inquirendae and will help to establish their genetic relationships with related taxa.
9.2. Identification of human infections The precise and easy identification to species level of anisakid nematodes has a significant implication for the diagnosis of human anisakid infections (anisakidosis) and, therefore, it is an important component of disease surveillance. To date, PCR-DNA methods have shown that human infections in Italy are attributable to A. pegreffii (see D’Amelio et al., 1999; Mattiucci et al., 2007b), while in Japan infections are mainly due to A. simplex (s.s.), despite the reported occurrence of A. pegreffii in fish from the southern Sea of Japan (Umehara et al., 2007). The DNA-based diagnostic techniques have an advantage when an anisakid larva is extracted by endoscopy, because only very small portion of the larva is required for its specific identification and anisakid larvae in any case afford scant features for identification to species level. PCR-DNA markers to recognize larval anisakids in eosinophilic granulomas, after their surgical removal, could be also applied in future for diagnosis of human cases. The collection of basic information on human infections by anisakids is important in the assessment of their relative epidemiological role, especially where different species are sympatric and even syntopic in the same fish species. This becomes more relevant when the fish are of high commercial value and are used for human consumption. This will in future allow us to show distinct differences between the diseases caused by different anisakid species. For instance, the clinical syndrome in human infections with Pseudoterranova larvae is different in Japan from that reported in the USA (Oshima, 1987). While the nature of the pathogenic role is not yet fully understood, Desowitz (1986) has suggested that various clinical aspects of human infections might be caused by different ‘strains’ of P. decipiens in the two different geographical areas. As different sibling species of P. decipiens are distributed in fish from these areas (P. azarasi and P. bulbosa in Japan, but P. decipiens (s.s.) and P. bulbosa in USA), the correct identification in the future of human infections caused by these anisakid species may be of particular importance.
9.3. Molecular ecology and life cycle Genetic markers have proved to be an essential tool for basic ecological studies where the ranges of anisakid species overlap. This allows us to investigate patterns of their differential distribution in both definitive and
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
133
intermediate/paratenic hosts in order to clarify whether they are the result of ecological processes such as competition and adaptation. In fact, despite the numerous observations on the definitive and intermediate hosts so far collected for the different genetically identified anisakid species, further studies are required to confirm the host range and the geographical distribution of several anisakid species. In addition, studies need to be extended to other areas where anisakids of uncertain taxonomic designation are reported but have not been fully investigated. Anisakid specimens have been recorded from these areas but data concerning their definitive and intermediate hosts are still patchy (for instance, for the species indicated in this review as Anisakis sp., Anisakis sp.1 and Anisakis sp. 2). Other aspects related to possible life-history pathways speculated for different anisakid species are still scanty and need to be fully demonstrated. This may include, for instance, proof that A. physeteris has a life cycle with squid rather than fish as suitable intermediate hosts. This was suggested by Mattiucci et al. (2001, 2004), taking into account the finding that other cephalopods have been reported to be infected by this anisakid species (but not genetically identified) in Pacific waters. These included Onychoteuthis borealis japonica (see Nagasawa and Moravec, 2002) and other squid species (i.e. Todarodes japonicus, Doryteuthis bleekeri and Ommastrephes bartrami) (see Smith and Wootten, 1978). Similarly, according to the high rates of prevalence found in several pelagic and demersal fish species of A. simplex (s.s.) and A. pegreffii, it seems that these parasites use pelagic and mesopelagic food chains to complete their life cycles. However, although this was anticipated by several authors (Klimpel et al., 2007; Mattiucci et al., 1997, 2004; Pontes et al., 2005), further investigations are required to confirm these hypotheses. These would include the molecular identification of anisakid larval stages from invertebrate species adapted to benthic or pelagic food chains and involved as first intermediate hosts in these anisakid life cycles. Indeed, despite several invertebrate species having been recorded as first intermediate hosts for Anisakis simplex (s.l.) and Pseudoterranova decipiens (s.l.) (Hurst, 1984; Marcogliese, 1993; Smith and Mooney Snyder, 2005), no data are so far available on the specific identification of the larval anisakids recovered from those hosts.
9.4. Host–parasite co-evolutionary aspects The long history of host–anisakid associations could have led to reciprocal adaptations in the hosts and their parasites (classical co-adaptation) as well as contemporaneous events in the two lineages (co-speciation). The available host historical biogeography mapped onto the phylogenetic trees postulated for the anisakid nematodes of the genera Anisakis and
134
Simonetta Mattiucci and Giuseppe Nascetti
Contracaecum have made possible reconstructions of co-evolutionary scenarios. However, other genetically unstudied taxa of anisakid nematodes belonging to Anisakis, Pseudoterranova and Contracaecum from cetaceans and pinnipeds should be investigated, and their phylogeny mapped onto that of their definitive hosts as inferred from other molecular markers. Comparative phylogenetics should also be studied to address the identification of temporal links between host and parasite phylogenies, and thus provide an internal time calibration for comparative studies of rates of evolution in the two groups. Moreover, in a parasite clade that shows evidence for host-switching, further studies will determine whether there are geographical, morphological or ecological correlates of host-switching. Indeed reconstruction of the biogeography history of host-switching events may reveal whether colonization of new hosts by anisakid nematodes was simply ‘opportunistic’ or whether the parasite is acquiring a particular resource in that host taxon which is not itself correlated with host phylogeny. Co-adaptation and co-evolution in anisakid nematodes will also be addressed in the future by further examination of the host–parasite assemblages already attempted in the group of Contracaecum spp. from fish-eating birds (Mattiucci et al., 2008c).
9.5. Genetic variability of anisakids as an indicator of habitat disturbance It has recently been shown in several organisms that natural and anthropogenic environmental changes could affect the genetic structure of animal populations causing ‘genetic erosion’ (loss of number of alleles per locus, decrease in heterozygosity and polymorphism rates) in their gene pools. Different mechanisms such as population fragmentation and size reduction, gene flow interruption and genetic drift can be invoked for this. Thus, we may assume that habitat disturbance of marine ecosystems and food webs can have considerable long-term effects on the genetic and demographic viability of the parasite populations due to a reduction of host population numbers. Indeed, assuming that habitat disturbance and fragmentation reduce the local effective population size (N) or migration rate (m) or both, increased genetic drift should redistribute genetic diversity, such that variation within populations is reduced and variation between populations is increased. In a small and isolated population, the average heterozygosity decreases at a constant rate per generation, so that small populations have a higher probability of losing genetic diversity than larger and continuously distributed populations. The extent to which a species could be affected by habitat disturbance is determined by its degree of specialization and dispersal potential.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
135
Two of the main factors influencing the genetic variability of a population are the amount of gene flow between populations and the population size. In nematode parasites, gene flow is largely determined by the dispersal of host animals (Blouin et al., 1995) and parasite abundance (as a proxy for parasite population size) which are strongly correlated with host density (Arneberg et al., 1998). As a consequence, variation in host dispersal and density could lead to differences in the patterns of genetic diversity distributed among parasite populations (Criscione and Blouin, 2005; May, 1993). Two complementary strategies have generally been used to examine the genetic effects of habitat disturbance (anthropogenic impact) on the genetic diversity (variability) of a population: (1) comparison of genetic variability of different sets of populations inhabiting disrupted ecosystems (space dimension) and/or (2) comparison of the genetic variability of a particular data set of populations from the same geographical area at different times (time dimension). As for the first point, it has been shown that the existence of a correlation between genetic variability values and parasitic infection levels of anisakid populations, from different geographical areas of the Boreal and Austral hemispheres, have different levels of habitat disturbance (i.e. Arctic vs Antarctic ecosystems). Could anthropogenic changes such as habitat disturbance provoke the reduction of host population size? In turn this would mean a reduction in habitats for their anisakid nematodes that could lead to a reduction in the genetic variability of the parasites’ gene pools, resulting in a loss of genetic polymorphism in some genes. Future efforts could be directed towards testing this hypothesis by monitoring genetic diversity (variability) of anisakid populations over time. Nowadays, the ‘genetic monitoring’ definition encompasses the quantification of temporal changes in population genetic data generated using suitable molecular genetic markers such as allozymes, microsatellites, single nucleotide polymorphisms (SNPs), amplified fragment length polymorphism (AFLP) and single-strand polymorphisms (SSCPs) (Schwartz et al., 2006). Preliminary data on the ‘genetic monitoring’ of anisakid populations through time seem to indicate that the possible phenomenon of the loss of polymorphism could have affected their gene pools. This has been observed in populations of Anisakis pegreffii, a sibling species of the A. simplex complex widespread in the Mediterranean Sea but also occurring in the Austral region. Loss of observed heterozygosity at some polymorphic allozyme loci has been found in the Mediterranean populations of A. pegreffii collected from fish hosts in recent years, as compared with that estimated in samples collected earlier in the genetic studies on these parasites (Fig. 2.12). In contrast, the Austral populations of A. pegreffii have maintained the same level of heterozygosity at the same
136
Simonetta Mattiucci and Giuseppe Nascetti
35 30 25
6Pgdh
20 Aat-2
15
Mpi
10 5 Mpi Aat-2 6Pgdh
2006 (South Pacific)
1994 (South Pacific)
2006 (Mediterranean Sea)
1998 (Mediterranean Sea)
1982 (Mediterranean Sea)
0
FIGURE 2.12 Temporal variation in the distribution of the genetic diversity (expected heterozygosity, he) at three allozyme loci (6Pgdh, Aat-2 and Gpi) in populations of Anisakis pegreffii from the Mediterranean Sea and South Pacific Ocean. Loss of observed heterozygosity (genetic erosion) at some polymorphic allozyme loci has been found in recent Mediterranean populations of A. pegreffii with respect to those estimated from the same populations in previous years and from the Austral region.
gene loci through time, despite the high level of gene flow observed. It is possible that this finding in Mediterranean waters could be related to a reduced population size in A. pegreffii. This in turn could correlate to habitat disturbance-mediated reductions in population sizes of their definitive and intermediate hosts. Clearly, further comparative genetic analyses of these parasite populations and their levels of abundance over time need to be undertaken to investigate this possibility. Molecular genetic markers, other than allozymes, could provide estimates of intraspecific genetic variation of these populations over time. Among these, those which have been demonstrated to show high genetic nucleotide variation in nematode populations (such as mitochondrial genes) (Blouin, 2002; Blouin et al., 1998; Hu and Gasser, 2006; Hu et al, 2004; Kim et al., 2006) could be promising for gathering data from earlier anisakid populations in museum collections. Thus, monitoring of the genetic diversity of anisakid nematodes and their abundance levels in hosts from different geographical areas, over time and space, could be used in the future as a tool for monitoring trophic web stability and for assessing the general biodiversity of marine ecosystems. Given that many of the anisakid nematodes are host specific, an ecosystem rich in their hosts should also be the one rich in anisakid parasites.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
137
We could thus conclude with a ‘paradox of anisakid’: A healthy marine ecosystem is one with high level of infections by anisakid nematodes!
ACKNOWLEDGEMENTS We apologize for not citing several authors and their valuable contributes on anisakid nematodes knowledge due to space constraint. We are very grateful to David I. Gibson for his helpful advice on the paper, to Steve C. Webb for reading the manuscript and suggestions, and to Robin Overstreet for comments on a first draft of the manuscript. Constructive suggestions of an anonymous referee improved the paper. We wish to thank Michela Paoletti and Carlo Taccari for their help in preparing tables and figures. Part of these studies was supported by European Community, Italian Ministry for Agriculture and Fisheries (DG Fisheries and Aquaculture), I Faculty of Medicine ‘Sapienza— University of Rome’, and ‘Tuscia University’ in Viterbo.
REFERENCES Abe, N., Ohya, N., and Yanagiguchi, R. (2005). Molecular characterization of Anisakis pegreffii larvae in Pacific cod in Japan. J. Helminthol. 79, 303–306. Abe, N., Tominaga, K., and Kimata, I. (2006). Usefulness of PCR-restriction fragments length polymorphism analysis of the internal transcribed spacer region of rDNA for identification of Anisakis simplex complex. Jpn. J. Infect. Dis. 59, 60–62. Abollo, E., Gestal, C., and Pascal, S. (2001). Anisakis infestation in marine fish and cephalopods from Galician waters: An updated perspective. Parasitol. Res. 87, 492–499. Adams, B. J. (1998). Species concepts and the evolutionary paradigm in modern nematology. J. Nematol. 30, 1–21. Adams, A. M., Murrell, K. D., and Cross, J. H. (1997). Parasites of fish and risks to public health. Rev. Sci. Technol. 16, 652–660. Alvarado-Bremer, J. R., Mejuto, J., Go´mez-Ma´rquez, J., Pla-Zanuy, C., Vin˜as, J., Marques, C., Hazin, F., Griffiths, M., Ely, B., Sa´nchez, B., and Greig, T. W. (2006). Genetic population structure of Atlantic swordfish: Current status, controversies, and future directions. Report of the 2006 ICCAT Swordfish Stock Structure Workshop, Crete (Greece), 13–14 March 2006.’’ SCRS/2006/041. Andersen, M., Gwynn, J. P., Dowdall, M., Kovacs, K. M., and Lydersen, C. (2006). Radiocaesium (137Cs) in marine mammals from Svalbard, the Barents Sea and the North Greenland Sea. Sci. Total Environ. 363, 87–94. Anderson, T. J. C., Blouin, M. S., and Beech, R. N. (1998). Population biology of parasitic nematodes: Application of genetic markers. Adv. Parasitol. 41, 219–283. Anon, R. N. (1997). Fishing by numbers reveals its limits. Nature 387, 110. Arduino, P., Nascetti, G., Cianchi, R., Plo¨tz, J., Mattiucci, S., D’Amelio, S., Paggi, L., Orecchia, P., and Bullini, L. (1995). Isozyme variation and taxonomic rank of Contracaecum radiatum (v. Linstow, 1907) from the Antarctic Ocean (Nematoda, Ascaridoidea). Syst. Parasitol. 30, 1–9. Arnason, U., Bodin, K., Gullberg, A., Ledje, C., and Mouchaty, S. (1995). A molecular view of pinniped relationships with particular emphasis on the true seals. J. Mol. Evol. 40, 78–85. Arnason, U., Gullberg, A., and Janke, A. (2004). Mitogenomic analyses provide new insight into cetacean origin and evolution. Gene 333, 27–34. Arneberg, P., Skorping, A., Grenfell, B., and Read, A. F. (1998). Host densities as determinants of abundance in parasite communities. Proc. R. Soc. Lond., B: Biol. Sci. 265, 1283–1289.
138
Simonetta Mattiucci and Giuseppe Nascetti
Audicana, M. T., Ansotegui, I. J., de Corres, L. F., and Kennedy, M. W. (2002). Anisakis simplex: Dangerous—Dead and alive? Trends Parasitol. 18, 20–25. Ayala, F. J. (1975). Genetic differentiation during the speciation process. Evol. Biol. 8, 1–78. Barron, M. G., Heintz, R., and Krahn, M. M. (2003). Contaminant exposure and effects in pinnipeds: Implications for sea lion declines in Alaska. Sci. Total Environ. 311, 111–133. Battaglia, B., Valencia, J., and Walton, D. W. H. (1997). In ‘‘Antarctic Communities: Species, Structure and Survival,’’ p. 464. Cambridge University Press. Baylis, H. A. (1916). Some ascarids in the British Museum (Nat. Hist.). Parasitology 8, 360–378. Baylis, H. A. (1937). On the ascarids parasitic in seals, with special reference to the genus Contracaecum. Parasitology 24, 121–130. Begg, G. A., and Waldman, J. R. (1999). An holistic approach to fish stock identification. Fish. Res. 43, 35–44. Berland, B. (1961). Nematodes from some Norwegian marine fishes. Sarsia 2, 1–50. Berland, B. (1964). Phocascaris cystophorae sp. nov. (Nematoda) from the hooded seal, with ˚ rbok for Universitetet i Bergen, Mat.-naturv., Series 1963 17, an emendation of the genus. A 1–21. Beverly-Burton, M. (1971). Helminths from the Weddell Sea, Leptonychotes weddellii (Lesson, 1826) in the Antarctic. Can. J. Zool. 49, 75–83. Beverly-Burton, M., Nyman, O. L., and Pippy, J. H. C. (1977). The morphology, and some observation on the population genetics of Anisakis simplex larvae (Nematoda: Ascaridoidea) from fishes of the North Atlantic. J. Fish. Res. Board Can. 34, 105–112. Blouin, M. S. (2002). Molecular prospecting for cryptic species of nematodes: Mitochondrial DNA versus internal transcribed spacer. Int. J. Parasitol. 32, 527–531. Blouin, M. S., Yowell, C. A., Courtney, C. H., and Dame, J. B. (1995). Host movement and the genetic structure of populations of parasitic nematodes. Genetics 141, 1007–1014. Blouin, M. S., Yowell, C. A., Courtney, C. H., and Dame, J. B. (1998). Substitution bias, rapid saturation, and the use of mtDNA for nematode systematics. Mol. Biol. Evol. 15, 1719–1727. Brattey, J., and Davidson, W. S. (1996). Genetic variation within Pseudoterranova decipiens (Nematoda: Ascaridoidea) from Canadian Atlantic marine fishes and seals: Characterization by RFLP analysis of genomic DNA. Can. J. Fish. Aquat. Sci. 53, 333–341. Brattey, J., and Stenson, G. B. (1993). Host specificity and abundance of parasitic nematodes (Ascaridoidea) from the stomachs of five phocid species from Newfoundland and Labrador. Can. J. Zool. 71, 2156–2166. Bristow, G. A., and Berland, B. (1992). On the ecology and distribution of Pseudoterranova decipiens (Nematoda: Anisakidae) in an intermediate host, Hippoglossoides platessoides, in northern Norwegian waters. Int. J. Parasitol. 22, 203–208. Brooks, D. R. (1990). Parsimony analysis in historical biogeography and coevolution: Methodological and theoretical update. Syst. Zool. 39, 14–30. Brooks, D. R., and Hoberg, E. P. (1981). Hennig’s parasitological method: A proposed solution. Syst. Zool. 30, 229–249. Brooks, D. R., and Hoberg, E. P. (2007). How will global climate change affect parasite-host assemblages? Trends Parasitol. 23, 571–574. Bruce, N. L., and Cannon, L. R. G. (1990). Ascaridoid nematodes from sharks from Australia and the Solomon Islands, Southwestern Pacific Ocean. Invertebr. Taxon. 4, 763–783. Bullini, L., and Sbordoni, V. (1980). Electrophoretic studies of gene-enzyme systems: Microevolutionary processes and phylogenetic inference. Boll. Zool. 47, 95–112. Bullini, L., Nascetti, G., Paggi, L., Orecchia, P., Mattiucci, S., and Berland, B. (1986). Genetic variation of ascaridoid worms with different life cycles. Evolution 40, 437–440. Bullini, L., Arduino, P., Cianchi, R., Nascetti, G., D’Amelio, S., Mattiucci, S., Paggi, L., and Orecchia, P. (1994). Genetic and ecological studies on nematode endoparasites of the genera Contracaecum and Pseudoterranova in the Antarctic and Arctic-Boreal regions.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
139
In ‘‘Proceedings of the 2nd Meeting ‘Biology in Antarctica’, Padova’’ (A. Martello, ed.), pp. 131–146. Edizioni Universitarie Patavine, Padova. Bullini, L., Arduino, P., Cianchi, R., Nascetti, G., D’Amelio, S., Mattiucci, S., Paggi, L., Orecchia, P., Plotz, J., Smith, J. H., and Brattey, J. (1997). Genetic and ecological research on anisakid endoparasites of fish and marine mammals in the Antarctic and Arctic-Boreal regions. In ‘‘Antarctic Communities: Species, Structure and Survival’’ (B. Battaglia, J. Valencia, and D. W. H. Walton, eds.), pp. 39–44. Cambridge University Press, Cambridge. Burt, M. D. B. (1994). The sealworm situation. In ‘‘Parasitic and Infectious Diseases’’ (M. E. Scott and G. Smith, eds.), pp. 347–362. Academic Press, New York. Cassens, I., Vicario, S., Waddell, V. G., Balchowsky, H., Van Belle, D., Ding, W., Chen, F., Mohan, R. S. L., Simoes-Lopes, P. C., Bastida, R., Meyer, A., Stanhope, M. J., et al. (2000). Independent adaptation to riverine habitats allowed survival of ancient cetacean lineages. Proc. Natl. Acad. Sci. USA 97, 11343–11347. Cavalli-Sforza, L. L., and Edwards, A. W. F. (1967). Phylogenetic analysis: Models and estimation procedures. Am. J. Hum. Genet. 19, 233–257. Chai, Y. Y., Murrell, D. K., and Lymbery, A. (2005). Fish-borne parasitic zoonoses: Status and issues. Int. J. Parasitol. 35, 1233–1254. Charleston, M. A. (2003). Recent results in cophylogeny mapping. Adv. Parasitol. 54, 303–330. Cimmaruta, R., Bondanelli, P., and Nascetti, G. (2005). Genetic structure and environmental heterogeneity in the European hake (Merluccius merluccius). Mol. Ecol. 14, 2577–2591. Cimmaruta, R., Paoletti, M., Bondanelli, P., Santos, M. N., Garcia, A., and Nascetti, G. (2006). Genetic structure of Mediterranean and Atlantic swordfish investigated using allozymes and RFLPs. Report of the 2006 ICCAT swordfish stock structure workshop Crete (Greece) 13–14 March 2006. Cimmaruta, R., Bondanelli, P., Ruggi, A., and Nascetti, G. (2008). Genetic structure and temporal stability in horse mackerel (Trachurus trachurus). Fish. Res. 89, 114–121. Cook, R. M., Sinclair, A., and Stefansson, G. (1997). Potential collapse of North Sea cod stocks. Nature 385, 521–522. Criscione, C. D., and Blouin, M. S. (2005). Effective size of macroparasite populations: A conceptual model. Trends Parasitol. 21, 212–217. Criscione, C. D., Poulin, R., and Blouin, M. S. (2005). Molecular ecology of parasites: Elucidating ecological and microevolutionary processes. Mol. Ecol. 14, 2247–2257. Cross, M. A., Collins, C., Campbell, N., Watts, P. C., Chubb, J. C., Cunningham, C. O., Hatfield, E. M. C., and MacKenzie, K. (2007). Levels of intra-host and temporal sequence variation in a large CO1 sub-units from Anisakis simplex sensu stricto (Rudolphi 1809) (Nematoda: Anisakidae): Implications for fisheries management. Mar. Biol. 151, 695–702. Crow, J. F., and Aoki, K. (1984). Group selection for a polygenic behavioural trait: Estimating the degree of population subdivision. Proc. Natl. Acad. Sci. USA 81, 6073–6077. D’Amelio, S., Nascetti, G., Mattiucci, S., Cianchi, R., Orecchia, P., Paggi, L., Berland, B., and Bullini, L. (1990). Ricerche elettroforetiche su alcune species del genere Contracaecum, parassiti di uccelli ittiofagi (Ascaridida: Anisakidae). Parassitologia 32(suppl. 1), 77. D’Amelio, S., Mattiucci, S., Paggi, L., Koie, M., Podvyaznaya, I., Pugachev, O., Rusinek, O., Timoskhkin, O., and Nascetti, G. (1995). Taxonomic rank and origin of Contracaecum osculatum baicalensis Mozgovoi and Ryjikov, 1950, parasite of Phoca sibirica from Lake Baikal, with data on its occurrence in fish hosts. ‘‘Abstracts, IVth International Symposium on Fish Parasitology, Munich, October 3–7 2005,’’ 27. D’Amelio, S., Mathiopoulos, K. D., Brandonisio, O., Lucarelli, G., Doronzo, F., and Paggi, L. (1999). Diagnosis of a case of gastric anisakidosis by PCR-based restriction fragment length polymorphism analysis. Parassitologia 41, 591–593. D’Amelio, S., Mathiopoulos, K., Santos, C. P., Pugachev, O. N., Webb, S. C., Picanco, M., and Paggi, L. (2000). Genetic markers in ribosomal DNA for the identification of members
140
Simonetta Mattiucci and Giuseppe Nascetti
of the genus Anisakis (Nematoda: Ascaridoidea) defined by polymerase chain reactionbased restriction fragment length polymorphism. Int. J. Parasitol. 30, 223–226. D’Amelio, S., Barros, N. B., Ingrosso, S., Fauquier, D. A., Russo, R., and Paggi, L. (2007). Genetic characterization of members of the genus Contracaecum (Nematoda: Anisakidae) from fish-eating birds from west-central Florida, USA, with evidence of new species. Parasitology 134, 1041–1051. Dailey, M. D. (1975). The distribution and intraspecific variation of helminth parasites in pinnipeds. Rapport et Proces-Verbaux des Reunions-Conseil International pour l’Exploration de la Mer 169, 338–352. Daniels, R. (1982). Feeding ecology of some fishes of the Antarctic Peninsula. Fish. Bull. 80, 575–588. Davey, J. T. (1971). A revision of the genus Anisakis Dujardin, 1845 (Nematoda: Ascaridata). J. Helminthol. 45, 51–72. Deardoff, T. L., and Overstreet, R. M. (1979). Contracaecum multipapillatum (=C. robustum) from fishes and birds in the northern Gulf of Mexico. J. Parasitol. 66, 853–856. Deardoff, T. L., and Overstreet, R. M. (1980). Review of Hysterothylacium and Iheringascaris (both previously = Thynnascaris) (Nematoda: Anisakidae) from the northern Gulf of Mexico. Proc. Biol. Soc. Wash. 93, 1035–1079. Deardorff, T. L., and Overstreet, R. M. (1981). Terranova ceticola n. sp. (Nematoda: Anisakidae) from the dwarf sperm whale; Kogia simus (Owen), in the Gulf of Mexico. Syst. Parasitol. 3, 25–28. Delyamure, S. L., Kurochkin, Y. V., and Skryabin, A. S. (1964). Contributions to the helminth fauna of the Caspian seal (Phoca caspica Gm.). Trudy Astrakanskogo Zapovednika 9, 105–118 (In Russian). Deme´re´, T. A., Berta, A., and Adam, P. J. (2003). Pinnipedimorphs evolutionary biogeography. Bull. Am. Mus. Nat. Hist. 279, 32–76 (Chapter 3). Desdevises, Y. (2007). Cophylogeny: Insights from fish-parasite systems. Parassitologia 49, 125–128. Desowitz, R. S. (1986). Human and experimental anisakiasis in the United States. Hokkaido J. Med. Sci. 61, 358–351. Di Guardo, G., Marruchella, G., Agrini, U., and Kennedy, S. (2005). Morbillivirus infections in aquatic mammals: A brief overview. J. Vet. Sci. 52, 88–93. Dollfus, R. (1966). Helminthofauna de Kogia breviceps (de Blainville, 1838) cetacea odeontocete. Annls Soc. Sci. Nat. Charente-Marit. 4, 3–6. Eichler, W. (1948). Some rules in ectoparasitism. Ann. Mag. Natl. Hist. 12, 588–598. Fagerholm, H. P. (1989). Intra-specific variability of the morphology in a single population of the seal parasite Contracaecum osculatum (Rudolphi) (Nematoda: Ascaridoidea), with a redescription of the species. Zool. Scripta 18, 33–41. Fagerholm, H. P. (1991). Systematic implications of male caudal morphology in ascaridoid nematode parasites. Syst. Parasitol. 19, 215–228. Fagerholm, H. P., and Gibson, D. I. (1987). A redescription of the pinniped parasite Contracaecum ogmorhini (Nematoda, Ascaridoidea), with an assessment of its antiBoreal circumpolar distribution. Zool. Scripta 16, 19–24. Fahrenholz, H. (1913). Ectoparasitism und Abstammungslehre. Zool. Anz. Leipzig 41, 371–374. Farjallah, S., Slimane, B. B., Busi, M., Paggi, L., Amor, N., Blel, H., Said, K., and D’Amelio, S. (2008). Occurrence and molecular identification of Anisakis spp. from the North African coasts of Mediterranean Sea. Parasitol. Res. 102, 371–379. Felsenstein, J. (1985). Confidence limits on phylogenies: An approach using the bootstrap. Evolution 39, 783–791. Foster, B. A., and Montgomery, C. (1993). Planktivory in benthic nototheniid fish in McMurdo Sound, Antarctica. Environ. Biol. Fish. 36, 313–318.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
141
George-Nascimento, M., and Llanos, A. (1995). Micro-evolutionary implications of allozymic and morphometric variations in sealworms Pseudoterranova sp. (Ascaridoidea: Anisakidae) among sympatric hosts from the southeastern Pacific Ocean. Int. J. Parasitol. 25, 1163–1171. George-Nascimento, M., and Urrutia, X. (2000). Pseudoterranova cattani sp. nov. (Ascaridoidea: Anisakidae), a parasite of the South American sea lion Otaria byronia De Blainville from Chile. Revta Chil. Hist. Nat. 73, 93–98. Gibson, D. I. (1983). The systematics of ascaridoid nematodes: A current assessment. In ‘‘Concepts in Nematode Systematics’’ (A. R. Stone, H. M. Platt, and L. F. Khalil, eds.) pp. 321–338. Academic Press, New York. Gibson, D. I., and Colin, J. A. (1982). The Terranova enigma. Parasitology 85, XXXVI–XXXVII. Guttman, L. A. (1968). A general nonmetric technique for finding the smallest coordinate space for a configuration of points. Psychometric 33, 469–506. Hartwich, G. (1957). Zur Systematik der Nematoden-Superfamilie Ascaridoidea. Zool. Jb., Abteilung fu¨r Systematik, Jena 85, 211–252. Hartwich, G. (1964). Revision der Vogelparasitischen Nematoden Mitteleuropas II. Die Gattung Contracaecum Railliet and Henry, 1912 (Ascaridoidea). Mitt. Zool. Mus. Berl. 40, 15–53. Hartwich, G. (1974). Keys to genera of the Ascaridoidea. In ‘‘CIH Keys to the Nematode Parasites of Vertebrates’’ (R. C. Anderson, A. G. Chabaud, and S. Wilmott, eds.), p. 153. Farnham Royal, Commonwealth Agriculture Bureau, Richmond. Hartwich, G. (1975). Rhabditida und Ascaridida. Tierwelt Dtl. 62, 1–256. Hoberg, E. P. (1992). Congruent and synchronic patterns in biogeography and speciation among seabirds, pinnipeds, and cestodes. J. Parasitol. 78, 601–615. Hoberg, E. P. (2005). Coevolution in marine systems. In ‘‘Marine Parasitology’’ (K. Rodhe, ed.), pp. 327–339. CBI Publishing, Oxon. Ho¨st, P. (1932). Phocascaris phocae n.gen.n.sp., eine neue Askaridenart aus Phoca groenlandica. Fabr. Zbl. Bakt. 1Ab. Orig. 125, 335–340. Hu, M., and Gasser, R. B. (2006). Mitochondrial genomes of parasitic nematodes: Progress and perspectives. Trends Parasitol. 22, 78–84. Hu, M., D’Amelio, S., Zhu, X. Q., Paggi, L., and Gasser, R. (2001). Mutation scanning for sequence variation in three mitochondrial DNA regions for members of the Contracaecum osculatum (Nematoda: Ascaridoidea) complex. Electrophoresis 22, 1069–1075. Hu, M., Chilton, N. B., and Gasser, R. B. (2004). The mitochondrial genomics of parasitic nematodes of socio-economic importance: Recent progress and implications for population genetics and systematic. Adv. Parasitol. 56, 133–212. Hudson, P. J., Dobson, A. P., and Lafferty, K. D. (2006). Is a healthy ecosystem one that is rich in parasites? Trends Ecol. Evol. 21, 381–385. Huizinga, H. W. (1967). Studies on the life cycle and development of Contracaecum spiculigerum (Rudolphi, 1809) (Ascaridoidea: Heterocheilidae) from marine piscivorous birds. J. Elisa Mitchell Sci. Soc. 82, 180–195. Hurst, R. J. (1984). Marine invertebrate hosts of New Zealand Anisakidae (Nematoda). N. Z. J. Mar. Freshwater Res. 18, 187–196. Johnston, T. H., and Mawson, P. M. (1941). Nematodes from Australian marine mammals. Rec. S. Aust. Mus. 6, 429–434. Johnston, T. H., and Mawson, P. M. (1945). Parasitic nematodes. Rep. Bri. NZ. Austral. Antarct. Res. Exped. Ser. B 5, 74–159. Karokhin, V. I. (1946). Two new species of Porrocaecum from Siberian birds of prey. In ‘‘Helminthological collection, Nauka, Moscow’’ (V. P. Pod’yanolskaya, ed.), pp. 132–141. In Russian.
142
Simonetta Mattiucci and Giuseppe Nascetti
Kendall, K. A., Ruhl, H. A., and Wilson, R. C. (2003). Distribution and abundance of marine bird and pinniped populations within Port Foster, Deception Islands, Antarctica. DeepSea Res. Part II: Top. Stud. Oceanogr. 50, 1873–1888. Kijewska, A., Rokicki, J., Sitko, J., and Wegrzyn, G. (2002). Ascaridoidea: A simple DNA assay for identification of 11 species infecting marine and freshwater fish, mammals and fish-eating birds. Exp. Parasitol. 101, 35–39. Kim, K. H., Eom, K. S., and Park, J. K. (2006). The complete mitochondrial genome of Anisakis simplex (Ascaridida: Nematoda) and phylogenetic implications. Int. J. Parasitol. 36, 319–328. Kimura, M. (1980). A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J. Mol. Evol. 16, 111–120. King, J. E. (1983). ‘‘Seals of the World’’ (British Museum Natural History) 2nd edn. p. 240. London and Oxford University Press, Oxford. Klimpel, S., Kellermanns, E., Palm, H. W., and Moravec, F. (2007). Zoogeography of fish parasites of the pearlside (Maurolicus muelleri), with genetic evidence of Anisakis simplex (s.s.) from the Mid-Atlantic Ridge. Mar. Biol. 152, 725–732. Klo¨ser, H., Plo¨tz, J., Palm, H., Bartsch, A., and Hubold, G. (1992). Adjustment of anisakid nematode life cycles to the high Antarctic food web as shown by Contracaecum radiatum and C. osculatum in the Weddell sea. Antarct. Sci. 4, 171–178. Kie, M., and Fagerholm, H. P. (1995). The life cycle of Contracaecum osculatum (Rudolphi, 1802) sensu stricto (Nematoda, Ascaridoidea, Anisakidae) in view of experimental infections. Parasitol. Res. 81, 481–489. Kotoulas, G., Mejuto, J., Antoniou, A., Tserpes, G., Piccinetti, C., Peristeraki, P., Kasapidis, P., Garcia-Cortes, B., Oikonomaki, K., De la Serna, J. M., and Magoulas, A. (2006). Genetic structure of the swordfish Xiphias gladius at a global scale using microsatellite markers. ‘‘Report of the 2006 ICCAT swordfish stock structure workshop, Crete (Greece), 13–15 March, 2006.’’ SCRS/2006/035. La Rosa, G., Marucci, G., Zarlenga, D. S., and Pozio, E. (2001). Trichinella pseudospiralis populations of the PaleArctic region and their relationship with populations of the NeArctic and Australian regions. Int. J. Parasitol. 31, 297–305. Legendre, P., Desdevises, Y., and Bazin, E. (2002). A statistical test for host-parasite coevolution. Syst. Biol. 51, 217–234. Li, A., D’Amelio, S., Paggi, L., He, F., Gasser, R. B., Lun, Z., Abollo, E., Turchetto, M., and Zhu, X. (2005). Genetic evidence for the existence of sibling species within Contracaecum rudolphii (Hartwich, 1964) and the validity of Contracaecum septentrionale (Kreis, 1955) (Nematoda: Anisakidae). Parasitol. Res. 96, 361–366. Lu, C. P., Chen, C. A., Hui, C. F., Tzeng, T. D., and Yeh, S. Y. (2006). Population genetic structure of the swordfish, Xiphias gladius (Linnaeus, 1758), in the Indian Ocean and West Pacific inferred from the complete DNA sequence of the mitochondrial control region. Zool. Stud. 45, 269–279. MacKenzie, K. (2002). Parasites as biological tags in population studies of marine organisms: An update. Parasitology 124, 153–163. Marcogliese, D. J. (1993). Larval nematodes infecting Amphiporeia virginiana (Amphipoda: Pontoporeioidea) on Sable Island, Nova Scotia. J. Parasitol. 79, 959–962. Marcogliese, D. J. (1997). Fecundity of sealworm (Pseudoterranova decipiens) infecting grey seals (Halichoerus grypus) in the Gulf of St. Lawrence, Canada: Lack of density-dependent effects. Int. J. Parasitol. 27, 1401–1409. Marcogliese, D. J. (2001). Implications of climate change for parasitism of animals in the aquatic environment. Can. J. Zool. 79, 1331–1352. Marcogliese, D. J. (2005). Parasites of the superorganism: Are they indicators of ecosystem health? Int. J. Parasitol. 35, 705–716.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
143
Margolis, L. (1956). Parasitic helminths and arthropods from pinnipedia of the Canadian pacific coast. J. Fish. Res. Board Can. 13, 489–505. Marques, J. F., Cabral, H. N., Busi, M., and D’Amelio, S. (2006). Molecular identification of Anisakis species from Pleuronectiformes off the Portuguese coast. J. Helminthol. 80, 47–51. Mattiucci, S., and Nascetti, G. (2006). Molecular systematics, phylogeny and ecology of anisakid nematodes of the genus Anisakis Dujardin, 1845: An update. Parasite´ 13, 99–113. Mattiucci, S., and Nascetti, G. (2007). Genetic diversity and infection levels of anisakid nematodes parasitic in fish and marine mammals from Boreal and Austral hemispheres. Vet. Parasitol. 148, 43–57. Mattiucci, S., Nascetti, G., Bullini, L., Orecchia, P., and Paggi, L. (1986). Genetic stucture of Anisakis physeteris and its differentiation from the Anisakis simplex complex (Ascaridida: Anisakidae). Parasitology 93, 383–387. Mattiucci, S., Nascetti, G., Cianchi, R., Paggi, L., Arduino, P., Margolis, L., Brattey, J., Webb, S. C., D’Amelio, S., Orecchia, P., and Bullini, L. (1997). Genetic and ecological data on the Anisakis simplex complex with evidence for a new species (Nematoda, Ascaridoidea, Anisakidae). J. Parasitol. 83, 401–416. Mattiucci, S., Paggi, L., Nascetti, G., Ishikura, H., Kikuchi, K., Sato, N., Cianchi, R., and Bullini, L. (1998). Allozyme and morphological identification of Anisakis, Contracaecum and Pseudoterranova from Japanese waters (Nematoda: Ascaridoidea). Syst. Parasitol. 40, 81–92. Mattiucci, S., Paggi, L., Nascetti, G., Abollo, E., Webb, S. C., Pascual, S., Cianchi, R., and Bullini, L. (2001). Genetic divergence and reproductive isolation between Anisakis brevispiculata and Anisakis physeteris (Nematoda: Anisakidae). Int. J. Parasitol. 31, 9–14. Mattiucci, S., Paggi, L., Nascetti, G., Portes Santos, C., Costa, G., Di Benedetto, A. P., Ramos, R., Argyrou, M., Cianchi, R., and Bullini, L. (2002a). Genetic markers in the study of Anisakis typica (Diesing, 1860): Larval identification and genetic relationships with other species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae). Syst. Parasitol. 51, 159–170. Mattiucci, S., Turchetto, M., Brigantini, F., and Nascetti, G. (2002b). On the occurrence of the sibling species of Contracaecum rudolphii complex (Nematoda: Anisakidae) in cormorants (Phalacrocorax carbo sinensis) from Venice and Caorle lagoons: Genetic markers and ecological studies. Parassitologia 44, 105. Mattiucci, S., Cianchi, R., Nascetti, G., Paggi, L., Sardella, N., Timi, J., Webb, S. C., Bastida, R., Rodriguez, D., and Bullini, L. (2003). Genetic evidence for two sibling species within Contracaecum ogmorhini Johnston and Mawson (1941) (Nematoda: Anisakidae) from otariid seals of Boreal and Austral regions. Syst. Parasitol. 54, 13–23. Mattiucci, S., Abaunza, P., Ramadori, L., and Nascetti, G. (2004). Genetic identification of Anisakis larvae in European hake from Atlantic and Mediterranean waters for stock recognition. J. Fish Biol. 65, 495–510. Mattiucci, S., Nascetti, G., Dailey, M., Webb, S. C., Barros, N., Cianchi, R., and Bullini, L. (2005). Evidence for a new species of Anisakis Dujardin, 1845: Morphological description and genetic relationships between congeners (Nematoda: Anisakidae). Syst. Parasitol. 61, 157–171. Mattiucci, S., Contadini, R., Paoletti, M., and Nascetti, G. (2006). Relazioni genetiche tra specie del genere Contracaecum (Railliet et Henry, 1912), parassiti di pinnipedi, ed aspetti coevolutivi ospite-parassita. Abstracts XVI Congresso Societa` Italiana di Ecologia 98. Mattiucci, S., Abaunza, P., Damiano, S., Garcia, A., Santos, M. N., and Nascetti, G. (2007a). Distribution of Anisakis larvae identified by genetic markers and their use for stock characterization of demersal and pelagic fish from European waters: An update. J. Helminthol. 81, 117–127. Mattiucci, S., Paoletti, M., De Angelis, M., Sereno, S., and Cancrini, G. (2007b). Human Anisakidosis in Italy: Molecular and histological identification of two new cases. Abstracts of the VII International Symposium on Fish Parasites: Parassitologia 49(suppl. 2), 226.
144
Simonetta Mattiucci and Giuseppe Nascetti
Mattiucci, S., Farina, V., Campbell, N., Mackenzie, K., Ramos, P., Pinto, A. L., Abaunza, P., and Nascetti, G. (2008a). Anisakis spp. larvae (Nematoda: Anisakidae) from Atlantic horse mackerel: Their genetic identification and use as biological tags for host stock identification. Fish. Res. 89, 146–171. Mattiucci, S., Paoletti, M., Webb, S. C., Sardella, N., Timi, J. T., Berland, B., and Nascetti, G. (2008b). Genetic relationships among species of Contracaecum Railliet and Henry, 1912 and Phocascaris Ho¨st, 1932 (Nematoda: Anisakidae) from pinnipeds based on mitochondrial cox2 sequences, and congruence with allozyme data. Parasite 15(3). Mattiucci, S., Paoletti, M., Olivero-Verbel, J., Baldiris, R., Arroyo-Salgado, B., Garbin, L., Navone, G., and Nascetti, G. (2008c). Contracaecum bioccai n. sp. from the brown pelican Pelecanus occidentalis (L.) in Colombia (Nematoda: Anisakidae): Morphology, molecular evidence and its genetic relationship with congeners from fish-eating birds. Syst. Parasitol. 69, 101–121. May, R. M. (1993). Agriculture: Resisting resistance. Nature 361, 593–594. Mayr, E. (1963). ‘‘Animal Species and Evolution.’’ Belknap Press, Harvard University Press, Cambridge, Massachusetts. Mayr, E. (1976). ‘‘Evolution and the Diversity of Life.’’ Belknap Press, Harvard University Press, Cambridge, Massachusetts. McClelland, G. (2002). The trouble with sealworms (Pseudoterranova decipiens species complex, Nematoda): A review. Parasitology 124, 183–203. McClelland, G., Misra, R. K., and Marcogliese, D. J. (1983). Variation in abundance of larval anisakines, sealworm (Phocanema decipiens) and related species in cod and flatfish from the Southern Gulf of St. Laurence (4T) and the Breton shelf (4Vn). Can. Tech. Rep. Fish. Aquat. Sci. No. 27, p. 1201. McClelland, G., Misra, R. K., and Martell, D. J. (1985). Variations in abundance of larval anisakines, sealworm (Pseudoterranova decipiens) and related species, in eastern Canadian cod and flatfish. Can. Tech. Rep. Fish. Aquat. Sci. No. 1392, p. 57. McClelland, G., Misra, G. R. K., and Martell, D. J. (1990). Larval anisakine nematodes in various fish species from Sable Island Bank and vicinity. Can. Bull. Fish. Aquat. Sci. 222, 83–113. Milinkovitch, M. C. (1995). Molecular phylogeny of cetaceans prompts revision of morphological transformations. Trends Ecol. Evol. 10, 328–334. Mozgovoi, A. A. (1951). Ascarids of mammals of the U.S.S.R. (Anisakidae). Trudy gel’mint. Lab., Akademiya Nauk SSR 5, 12–22. Mozgovoi, A. A. (1953). Ascaridata of animals and man, and the diseases caused by them. Osn. Nemat. II. Izdatielstvo AN SSSR, Moskva, 616. Myers, B. J. (1959). Phocanema, a new genus for the anisakid nematode of seals. Can. J. Zool. 37, 459–465. Nadler, S. A. (1992). Phylogeny of some ascaridoid nematodes, inferred from comparison of 18s and 28s rRNA sequences. Mol. Biol. Evol. 9, 932–944. Nadler, S. A. (1995). Microevolution and genetic structure of parasite populations. J. Parasitol. 81, 395–403. Nadler, S. A. (2002). Species delimitation and nematode biodiversity: Phylogenies rule. Nematology 4, 615–625. Nadler, S. A. (2005). Speciation and species delimitation. In ‘‘Marine Parasitology’’ (K. Rohde, ed.), pp. 339–345. CABI Publishing, Oxon. Nadler, S. A., and Hudspeth, D. S. S. (1998). Ribosomal DNA and phylogeny of the Ascaridoidea (Nematoda: Secernentea): Implications for morphological evolution and classification. Mol. Phylogenet. Evol. 10, 221–236. Nadler, S. A., D’Amelio, S., Fagerholm, H. P., Berland, B., and Paggi, L. (2000). Phylogenetic relationships among species of Contracaecum Railliet and Henry, 1912 and Phocascaris
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
145
Ho¨st, 1932 (Nematoda: Ascaridoidea) based on nuclear rDNA sequence data. Parasitology 121, 455–463. Nadler, S. A., D’Amelio, S., Dailey, M. D., Paggi, L., Siu, S., and Sakanari, J. A. (2005). Molecular phylogenetics and diagnosis of Anisakis, Pseudoterranova, and Contracaecum from Northern Pacific marine mammals. J. Parasitol. 91, 1413–1429. Nagasawa, K., and Moravec, F. (2002). Larval anisakid nematodes from four species of squid (Cephalopoda: Teuthoidea) from the central and western North Pacific Ocean. J. Nat. Hist. 36, 883–891. Nascetti, G. (1992). Studio e riconoscimento di specie gemelle in nematodi endoparassiti: Analisi della variazione genetica, dell’isolamento riproduttivo, della nicchia ecologica e della competizione. Atti della Societa` Italiana di Ecologia 15, 287–312. Nascetti, G., Paggi, L., Orecchia, P., Smith, J. W., Mattiucci, S., and Bullini, L. (1986). Electrophoretic studies on Anisakis simplex complex (Ascaridida: Anisakidae) from the Mediterranean and North East Atlantic. Int. J. Parasitol. 16, 633–640. Nascetti, G., Bullini, L., Cianchi, R., Paggi, L., Orecchia, P., Mattiucci, S., D’Amelio, S., and Berland, B. (1990). Genetic relationships among anisakid species belonging to the genera Contracaecum and Phocascaris. Bull. Soc. fr. Parasitol. 8, 261. Nascetti, G., Cianchi, R., Mattiucci, S., D’Amelio, S., Orecchia, P., Paggi, L., Brattey, J., Berland, B., Smith, J. W., and Bullini, L. (1993). Three sibling species within Contracaecum osculatum (Nematoda, Ascaridida, Ascaridoidea) from the Atlantic Arctic-Boreal region: Reproductive isolation and host preferences. Int. J. Parasitol. 23, 105–120. Nei, M. (1972). Genetic distance between populations. Am. Nat. 106, 283–292. Nevo, E., Beiles, A., and Ben-Shlomo, R. (1984). The evolutionary significance of genetic diversity: Ecological, demographic and life history correlates. Evol. Dyn. Genet. Divers. 53, 14–213. Nikaido, M., Matsumo, F., Hamilton, H., Brownell, R. L., Cao, Y., Ding, W., Zuoyan, Z., Shedlock, A. M., Fordyce, R. E., Hasegawa, M., and Okada, N. (2001). Retroposon analysis of major cetaceans lineages: The monophyly of toothed whales and the paraphyly of river dolphins. Proc. Natl. Acad. Sci. USA 98, 7384–7389. Orecchia, P., Paggi, L., Mattiucci, S., Nascetti, G., Smith, J. W., and Bullini, L. (1986a). Electrophoretic identification of larvae and adults of Anisakis (Ascaridida: Anisakidae). J. Helminthol. 60, 331–339. Orecchia, P., Berland, B., Paggi, L., Nascetti, G., Mattiucci, S., and Bullini, L. (1986b). Divergenza genetica di specie dei generi Contracaecum e Phocascaris (Ascaridida, Anisakidae) con diverso ciclo biologico. Ann. Ist. sup. Sanita` 22, 345–348. Orecchia, P., Mattiucci, S., D’Amelio, S., Paggi, L., Plotz, J., Cianchi, R., Nascetti, G., Arduino, P., and Bullini, L. (1994). Two new members in the Contracaecum osculatum complex (Nematoda, Ascaridoidea) from the Antarctic. Int. J. Parasitol. 24, 367–377. Osche, G. (1963). Morphological, biological, and ecological considerations in the phylogeny of parasitic nematodes. In ‘‘The Lower Metazoa, Comparative Biology and Phylogeny’’ (E. C. Dougherty, ed.), pp. 283–302. California University Press, Berkeley. Oshima, T. (1987). Anisakiasis is the Sushi bar guilty? Parasitol. Today 3, 44–48. Paggi, L., and Bullini, L. (1994). Molecular taxonomy in anisakids. Bull. Scand. Soc. Parasitol. 4, 25–39. Paggi, L., Nascetti, G., Cianchi, R., Orecchia, P., Mattiucci, S., D’Amelio, S., Berland, B., Brattey, J., Smith, J. W., and Bullini, L. (1991). Genetic evidence for three species within Pseudoterranova decipiens (Nematoda, Ascaridida, Ascaridoidea) in the North Atlantic and Norwegian and Barents Seas. Int. J. Parasitol. 21, 195–212. Paggi, L., Mattiucci, S., D’Amelio, S., and Nascetti, G. (1998a). Nematodi del genere Anisakis in pesci, cefalopodi e cetacei del Mar Mediterraneo e dell’Oceano Atlantico e Pacifico. Biol. Mar. Mediterr. 5, 1585–1592.
146
Simonetta Mattiucci and Giuseppe Nascetti
Paggi, L., Nascetti, G., Webb, S. C., Mattiucci, S., Cianchi, R., and Bullini, L. (1998b). A new species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae) from beaked whale (Ziphiidae): Allozyme and morphological evidence. Syst. Parasitol. 40, 161–174. Paggi, L., Mattiucci, S., Ishikura, H., Kikuchi, K., Sato, N., Nascetti, G., Cianchi, R., and Bullini, L. (1998c). Molecular genetics in anisakid nematodes from the Pacific Boreal Region. In ‘‘Host Response to International Parasitic Zoonoses’’ (H. Ishikura, M. Aikawa, H. Itakura, and K. Kikuchi, eds.), pp. 83–107. Springer-Verlag, Tokyo. Paggi, L., Mattiucci, S., Gibson, D. I., Berland, B., Nascetti, G., Cianchi, R., and Bullini, L. (2000). Pseudoterranova decipiens species A and B (Nematoda: Ascaridoidea): Nomenclatural designation, morphological diagnostic characters and genetic markers. Syst. Parasitol. 45, 185–197. Paggi, L., Mattiucci, S., and D’Amelio, S. (2001). Allozyme and PCR-RFLP markers in anisakid nematodes, aethiological agents of human anisakidosis. Parassitologia 43, 21–27. Palm, H., Anderson, K., Klo¨ser, H., and Plo¨tz, J. (1994). Occurrence of Pseudoterranova decipiens (Nematoda) in fish from the southeastern Weddell Sea (Antarctic). Polar Biol. 14, 539–544. Pa`lsson, J., and Beverly-Burton, M. (1984). Helminth parasites of capelin, Mallotus villosus (Pisces: Osmeridae) of the north Atlantic. Proc. Helminthol. Soc. Wash. 51, 248–254. Pannacciulli, F., Bishop, J. D. D., and Hawkins, S. J. (1997). Genetic structure of populations of two species of Chthamalus (Crustacea: Cirripedia) in the north-east Atlantic and Mediterranean. Mar. Biol. 128, 73–82. Pastor, T., Garza, J. C., Allen, P., Amos, W., and Aguilar, A. (2004). Low genetic variability in the highly endagered Mediterranean monk seal. J. Hered. 95, 291–300. Paterson, A. M., Gray, R. D., and Wallis, G. P. (1993). Parasites, petrels and penguins: Does louse phylogeny reflect seabird phylogeny? Int. J. Parasitol. 23, 515–526. Petter, A. J. (1972). Redescription of Anisakis insignis Diesing (Ascaridoidea), parasite of the Amazon river dolphin, Inia geoffrensis. In ‘‘Investigation on Cetacea’’ (G. Pilleri, ed.), Vol. IV, pp. 93–99. Brain Anatomy Institute, University of Berne, Switzerland. Pontes, T., D’Amelio, S., Costa, G., and Paggi, L. (2005). Molecular characterization of larval anisakid nematodes from marine fishes of Madeira by a PCR-based approach, with evidence for a new species. J. Parasitol. 91, 1430–1434. Poulin, R. (2006). Variation in infection parameters among populations within parasite species: Intrinsic properties versus local factors. Int. J. Parasitol. 36, 877–885. Poulin, R., and Morand, S. (2004). ‘‘The Parasite Biodiversity.’’ Smithsonian Institution Press, S. Smithsonian Books, Washington. Pozio, E., and Murrel, K. D. (2006). Systematics and epidemiology of Trichinella. Adv. Parasitol. 63, 367–439. Railliet, A., and Henry, A. (1912). Quelques nematodes parasites des reptiles. Bull. Soc. Path. exot. 5, 251–259. Read, A. J., Drinker, P., and Northridge, S. (2006). By-catch of marine mammals in U.S. and global fisheries. Conserv. Biol. 20(1), 163–169. Reeb, C. A., and Block, B. (2006). Ten microsatellite loci show that Mediterranean swordfish are genetically unique from other populations worldwide. ‘‘Report of the 2006 ICCAT swordfish stock structure workshop, Crete (Greece), 13–15 March 2006.’’, SCRS/2006/02. Rios, C., Sanz, S., Saavedra, C., and Pena, J. B. (2002). Allozyme variation in populations of scallops, Pecten jacobaeus (L.) and P. maximus (L.) (Bivalvia: Pectinidae), across the Almeria-Oran front. J. Exp. Mar. Biol. Ecol. 267, 223–244. Saraiva, A., Faranda, A., Damiano, S., Hermida, M., Santos, M. J., Ventura, C. H., Soares, J. P., Cruz, C., Eiras, J. C., and Mattiucci, S. (2007). Six species of Anisakis (Nematoda: Anisakidae) parasites of the black scabbardfish, Aphanopus carbo from NE Atlantic waters: Genetic markers and fish biology. Abstracts of VII International Symposium on Fish Parasites, Parassitologia 49(suppl. 2), 229.
Anisakid Nematodes and Host–Parasite Co-evolutionary Processes
147
Schwartz, M. K., Luikart, G., and Waples, R. S. (2006). Genetic monitoring as a promising tool for conservation and management. Trends Ecol. Evol. 22, 25–33. Shiraki, T. (1974). Larval nematodes of family Anisakidae (Nematoda) in the Northern Sea of Japan—As a causative agent of eosinophilic phlegmone or granuloma in the human gastro-intestinal tract. Acta Med. Biol. 22, 57–98. Slatkin, M., and Barton, N. H. (1989). A comparison of three indirect methods for estimating average levels of gene flow. Evolution 43, 1349–1368. Smith, J. W., and Mooney Snyder, J. (2005). New locality for third-stage larvae of Anisaks simplex (sensu lato) (Nematoda: Ascaridoidea) in euphausiids Euphasia pacifica and Thysanoessa raschii from Prince William Sound, Alaska. Parasitol. Res. 97, 539–542. Smith, J. W., and Wootten, R. (1978). Anisakis and anisakiasis. Adv. Parasitol. 16, 93–163. Sprent, J. F. A. (1977). Ascaridoid nematodes of amphibians and reptiles: Dujardinascaris. J. Helminthol. 51, 253–287. Sprent, J. F. A. (1978). Ascaridoid nematodes of amphibians and reptiles: Gedoelstascaris n. g. and Ortleppascaris n. g. J. Helminthol. 52, 261–282. Sprent, J. F. A. (1979). Ascaridoid nematodes of amphibians and reptiles: Multicaecum and Brevimulticaecum. J. Helminthol. 53, 91–116. Sprent, J. F. A. (1983). Ascaridoid nematodes of amphibians and reptiles: Typhlophorus, Hartwichia and Trispiculascaris. J. Helminthol. 57, 179–189. Sures, B. (2004). Environmental parasitology: Relevancy of parasites in monitoring environmental pollution. Trends Parasitol. 20, 170–177. Thompson, R. M., Mouritsen, K. N., and Poulin, R. (2005). Importance of parasites and their life cycle characteristics in determining the structure of a large marine food web. J. Anim. Ecol. 74, 77–85. Timi, J. T., Sardella, N. H., and Mattiucci, S. (2003). Contracaecum ogmorhini s.s. Johnston and Mawson, 1941 (Nematoda: Anisakidae), parasite of Arctocephalus australis (Zimmermann, 1783) off the Argentinian coast. Helminthologia 40, 27–31. Umehara, A., Kawakami, Y., Matsui, T., Araki, J., and Uchida, A. (2006). Molecular identification of Anisakis simplex sensu stricto and Anisakis pegreffii (Nematoda: Anisakidae) from fish and cetacean in Japanese waters. Parasitol. Int. 55, 267–271. Umehara, A., Kawakami, Y., Araki, J., and Uchida, A. (2007). Molecular identification of the etiological agent of the human anisakiasis in Japan. Parasitol. Int. 56, 211–215. Umehara, A., Kawakami, Y., Araki, J., and Uchida, A. (2008). Multiplex PCR for the identification of Anisakis simplex sensu stricto, Anisakis pegreffii and the other anisakid nematodes. Parasitol. Int. 57, 49–53. Vacchi, M., La Mesa, M., and Castelli, A. (1994). Diet of two coastal nototheniid fish from Terra Nova Bay, Ross Sea. Antarct. Sci. 6, 61–65. Valentini, A., Mattiucci, S., Bondanelli, P., Webb, S. C., Mignucci-Giannone, A., ColomLlavina, M. M., and Nascetti, G. (2006). Genetic relationships among Anisakis species (Nematoda: Anisakidae) inferred from mitochondrial cox2 sequences, and comparison with allozyme data. J. Parasitol. 92, 156–166. Van Thiel, P. H. (1960). Anisakiasis. Parasitology 52, 16–17. Vidal-Martinez, V. M., Osorio-Saraiba, D., and Overstreet, R. M. (1994). Experimental infection of Contracaecum multipapillatum (Nematoda: Anisakinae) from Mexico in the domestic cat. J. Parasitol. 80, 576–579. Webster, L. M. I., Johnson, P. C. D., Adam, A., Mable, B. K., and Keller, L. F. (2007). Macrogeographic population structure in parasitic nematode with avian host. Vet. Parasitol. 144, 156–166. Wickens, P. A. (1995). A review of operational interactions between pinnipeds and fisheries. FAO Fisheries Technical Paper 346. Yamaguti, S. (1961). ‘‘Systema Helminthum: The Nematodes of Vertebrates,’’ 679 pp. Interscience Publishers.
148
Simonetta Mattiucci and Giuseppe Nascetti
Yamaguti, S., and Arima, S. (1942). Porrocaecum azarasi n. sp. (Nematoda) from the Japanese seal. Trans. Sapporo Nat. Hist. Soc. 17, 113–116. Zarlenga, D. S., Hoberg, E. P., Stringfellow, F., and Lichtenfield, J. R. (1998). Comparison of two polymorphic species of Ostertagia and phylogenetic relationships with the Ostertaginiiae (Nematoda: Tricostrongiloidea) inferred from DNA repeat and mitochondrial DNA sequences. J. Parasitol. 84, 806–812. Zhu, X. Q., Gasser, R. B., Podolska, M., and Chilton, N. B. (1998). Characterisation of anisakid nematodes with zoonotic potential by nuclear ribosomal DNA sequences. Int. J. Parasitol. 28, 1911–1921. Zhu, X. Q., Gasser, R. B., Jacobs, D. E., Hung, G. C., and Chilton, N. B. (2000a). Relationships among some ascaridoid nematodes based on ribosomal DNA sequence data. Parasitol. Res. 86, 738–744. Zhu, X. Q., D’Amelio, S., Paggi, L., and Gasser, R. B. (2000b). Assessing sequence variation in the internal transcribed spacers of ribosomal DNA within and among members of the Contracaecum osculatum complex (Nematoda: Ascaridoidea: Anisakidae). Parasitol. Res. 86, 677–683. Zhu, X., D’Amelio, S., Hu, M., Paggi, L., and Gasser, R. B. (2001). Electrophoretic detection of population variation within Contracaecum ogmorhini (Nematoda: Ascaridoidea: Anisakidae). Electrophoresis 22, 1930–1934. Zhu, X. Q., D’Amelio, S., Palm, H. W., Paggi, L., George-Nascimento, M., and Gasser, R. B. (2002). SSCP-based identification of members within the Pseudoterranova decipiens complex (Nematoda: Ascaridoidea: Anisakidae) using genetic markers in the internal transcribed spacers of ribosomal DNA. Parasitology 124, 615–623. Zhu, X. Q., Podolska, M., Liu, J. S., Yu, H. Q., Chen, H. H., Lin, Z. X., Luo, C. B., Song, H. Q., and Lin, R. Q. (2007). Identification of anisakid nematodes with zoonotic potential from Europe and China by single-strand conformation polymorphism analysis of nuclear ribosomal DNA. Parasitol. Res. 101, 1703–1707.
CHAPTER
3 Atopic Disorders and Parasitic Infections Aditya Reddy* and Bernard Fried*
Contents
Abstract
150 151 154
1. 2. 3. 4.
Introduction Atopic Disorders Relationship of Parasites to Atopic Disorders Laboratory Studies on Atopy Using Selected Parasites and Rodent Models 5. Concluding Remarks References
175 182 184
This chapter examines the relationship between atopic disorders and parasitic infections. Atopy is an exaggerated IgE-mediated Type-1 immune response in predisposed individuals. Conflicting information exists in regard to the relationship of parasitic infections and the classic allergic diseases, that is, atopic dermatitis, allergic rhinitis and asthma. Attention is paid to the explanations for these discrepancies in the literature found within both human and animal studies on atopy with particular emphasis on helminthic infections. The factors that cause only a proportion of atopic individuals to develop clinical disease have not been defined although helminths confer protection in many studies examined. Early childhood infections help induce a Th1-biased immunity and prevent the induction of the Th2 system that causes atopy. Acute parasitic infections may increase manifestations of allergy, whereas chronic infections with parasites decrease atopic predisposition. Nonetheless, a causal association between geohelminth infection and atopic disorders has not been established. Some helminthic
* Department of Biology, Lafayette College, Easton, Pennsylvania 18042, USA
Advances in Parasitology, Volume 66 ISSN 0065-308X, DOI: 10.1016/S0065-308X(08)00203-0
#
2008 Elsevier Ltd. All rights reserved.
149
150
Aditya Reddy and Bernard Fried
substances, especially the cytokines, have respiratory and antiallergic effects, and may therefore become useful as therapeutic modalities for many atopic and allergic disorders.
1. INTRODUCTION The purpose of this chapter is to examine the relationship between atopic disorders and parasitic infections. Conflicting information exists in regard to the relationship of parasitic infections and certain atopic disorders. An understanding of atopy is a prerequisite for the analysis of this complex relationship. Atopy or the atopic syndrome is an allergic hypersensitivity affecting parts of the body not in direct contact with an allergen. Atopy is often defined as the genetic tendency to develop classic allergic diseases, that is, atopic dermatitis, allergic rhinitis (hay fever) and asthma. An allergen is a substance that is foreign to the body and can cause an allergic reaction in certain individuals. The main allergens involved are house-dust mites (HDM), house dust, human dander, feathers, mould and pollen from trees, weeds and grasses. Dust mites are medically important acarines and are included in this chapter because allergic reactions to arthropods are involved in the development of asthma and the provocation of attacks (Burgess, 1993). Atopic diseases are strongly correlated at a population level (International Study of Asthma and Allergies in Childhood (ISAAC) Steering Committee, 1998), and individuals who develop early atopic dermatitis are often at high risk of subsequently developing respiratory allergic disease (Spergel and Paller, 2003). Current literature on human atopic disorders and parasitic infection is scattered among various epidemiological and biomedical journals and presented in diverse ways. In this chapter, attention is paid to the explanations for these discrepancies in the literature found within both human and rodent studies on atopy with particular emphasis on helminthic infections. Most studies on atopy and parasites involve the helminths. Smits et al. (2005) noted that various population studies have provided a strong case for the involvement of helminth infections in this regard. A popular explanation known as the ‘hygiene hypothesis’ proposes that the increased prevalence in allergy is due to a diminished or altered exposure to gut-dwelling microbes, resulting in a disordered immunoregulation. Zaccone et al. (2006) noted that the ‘hygiene hypothesis’ was initially postulated to explain the inverse correlation between the incidence of infections and the rise of allergic diseases, particularly in the developed world. The ‘hygiene hypothesis’ has lately been extended to also incorporate autoimmune diseases in general (Zaccone et al., 2006). Epidemiological and experimental data both suggest that infections or the exposure to non-pathogenic bacteria protect individuals from developing
Atopic Disorders and Parasitic Infections
151
some autoimmune and atopic disorders (Kamradt et al., 2005). Numerous population studies have provided a strong case for the involvement of helminth infections in this respect. Analysis of helminth-induced immune responses showed that helminths not only prime for polarized Th2 responses but also potently induce T-cell hyporesponsiveness (Smits et al., 2005). Recently, it has been demonstrated that helminths induce suppressed host immune responses by priming for regulatory T cells. A spectrum of CD4+ T cells, including Th3 cells, TR cells, CD4+, CD25+ cells and NKT cells play a critical role in regulating these allergic diseases (Akbari et al., 2003). It is proposed that this regulatory T cell-inducing activity accounts for the protection observed in the development of allergic disorders. This chapter also provides an understanding of the relevant protective mechanisms against atopic disorders associated with parasites and summarizes a wide body of both clinical and experimental literature which may help clarify this complex interaction; we also highlight the need for further research in certain areas. The focus of this chapter is on those helminths, protozoans and medically important arthropods that are related to atopic disorders. This chapter excludes coverage of atopic disorders that are associated with bacterial, viral, fungal and metabolic disorders. This chapter is concerned mainly with humans as hosts, but where other hosts, particularly rodents, are used in experimental work, those studies are covered in our chapter. In contrast to microbial exposures which can promote Th2-supressing Th1 responses, and may skew Th2 infections, helminths appear to suppress atopy, which suggests an alternative explanation for these findings. In addition, the production of interleukin (IL)-4, IL-5, IL-9 and/or IL-13 by Th2 cells in response to helminthic infections mediates a range of responses that can be protective or pathogenic in atopic disorders. Although it has been well established that during these infections there is a stimulation of IgE against their own antigens as well as a strong induction of non-specific TH2/IL-4 polyclonal IgE, similar to the allergic processes itself, many authors debate if the presence of these infections correlates inversely or not with the rate prevalence of atopy or respiratory allergy (Mingomataj et al., 2006). The stimulation of a robust anti-inflammatory regulatory network by persistent immune challenge offers a unifying explanation for the observed inverse association of many infections with allergic disorders (Yazdanbakhsh et al., 2002).
2. ATOPIC DISORDERS The atopic disorders considered herein are atopic asthma, allergic rhinitis and atopic dermatitis because these types are often associated with parasitic infections. The atopic disorders considered herein are mainly
152
Aditya Reddy and Bernard Fried
concerned with various genera of acarines, helminths and protozoans. The sections used for most of our coverage will discuss atopic influences upon the host and, conversely, host influences upon the atopic disorder. The epidemiology of certain atopic disorders is described. The chapter also covers current research on the topic of pathophysiological pathways relating atopy and parasites and key pathogenic features of atopic disorders such as elevated IgE levels and a characteristic T helper cell cytokine pattern. Atopic asthma is the most common chronic disease of children, and the continued rising trends in asthma and atopy prevalence worldwide still increase irrespective of improved knowledge of many aspects of the diseases (von Hertzen and Haahtela, 2004). Atopic asthma refers to the onset of wheezing, cough and shortness of breath upon contact of an allergen. Importantly, there is a temporal relationship between the onset of allergic rhinitis and asthma, with rhinitis often preceding the development of asthma (Cruz et al., 2007a). A multi-centre study by Flo¨istrup et al. (2006), involving 6630 children aged 5–13 years in five European countries, suggested that restrictive use of antibiotics and antipyretics are associated with a reduced risk of allergic disease in children. The immune response to helminth infections has long been known to share key features with the allergic response; in particular, both are typified by enhanced T helper 2 (Th2) responses with high levels of IL-4, IL-5 and IL-13, accompanied by eosinophilia and significant IgE production (Yazdanbakhsh et al., 2001). Allergic rhinitis is seasonal or perennial itching, sneezing, rhinorrhea, nasal congestion and sometimes conjunctivitis, caused by exposure to pollens or other allergens. The seasonal type of allergic rhinitis is commonly known as ‘hay fever’ because it is most prevalent during haying season. Diagnosis is by history and skin testing. Treatment is with a combination of antihistamines, decongestants, nasal corticosteroids, and, for severe, refractory cases, desensitization. Treatment of rhinitis with intranasal glucocorticosteroids, antihistamines, leukotriene antagonists or immunotherapy may reduce asthma associated morbidity (Cruz et al., 2007a). Treatment of seasonal and perennial allergic rhinitis is generally the same, although attempts at environmental control (e.g. eliminating dust mites and cockroaches) are recommended for perennial rhinitis. Allergic conjunctivitis is most commonly a short-term (acute) problem but may uncommonly be a long-term (chronic) condition. Perennial rhinitis is caused by year-round exposure to indoor inhaled allergens (e.g. dust mites, cockroaches) or by strong reactivity to plant pollens in sequential seasons. Acute allergic conjunctivitis occurs with hay fever and other seasonal allergy. Atopic dermatitis is often referred to as ‘eczema’, which is a general term for many types of dermatitis. It is the most common of the many
Atopic Disorders and Parasitic Infections
153
types of eczema. Atopic dermatitis is a non-contagious inflammatory chronic disease that affects the skin. Atopic dermatitis is definitively the most common childhood skin disorder in developed countries (Williams et al., 1999). The prevalence of this condition has actually increased two- to threefold in the last three decades, affecting 15–20% of young children (Larsen and Hanifin, 2002). In atopic dermatitis, the skin becomes extremely itchy and inflamed, causing redness, swelling, cracking, weeping, crusting and scaling. Atopic dermatitis most often affects infants and young children, but it can continue into adulthood or first show up later in life. The relationship between atopic disorders and helminth infections is not only of interest to epidemiologists but also has intrigued immunologists, because of the close parallels between the allergic inflammation caused by the host immune responses to many environmental allergens and that caused by parasite antigens (Cooper, 2004). Both are associated with high levels of IgE, tissue eosinophilia and mastocytosis, mucus hypersecretion and T cells that preferentially secrete Th2 cytokines (IL-4, IL-5 and IL-13) (Holgate, 1999; Maizels and Yazdanbakhsh, 2003). Complex genetic, environmental and site-specific factors contribute to development of allergies. A role for genetic factors is suggested by familial inheritance of disease, association between atopy and specific human leukocyte antigen system (HLA; the name of the human major histocompatibility complex) loci and polymorphisms of genes for the high-affinity IgE receptor b chain, IL-4 and CD14. The various disorders are clearly interrelated; new evidence supports previous Allergic Rhinitis and its Impact on Asthma (ARIA) consensus statements, such as: (i) allergic rhinitis is a risk factor for asthma; (ii) patients with persistent rhinitis should be evaluated for asthma; (iii) most patients with asthma have rhinitis; (iv) a combined strategy should be used to treat the airways and (v) in low- to middle-income countries, a different strategy may be needed (Cruz et al., 2007a). Previous ARIA publications have stated that most patients with allergic and non-allergic asthma have rhinitis, that many patients with allergic rhinitis have increased bronchial hyper-reactivity, and that there is a temporal relationship between the initial onset of the allergic rhinitis and asthma, with rhinitis often preceding the development of asthma (Bachert et al., 2002). Type I hypersensitivity reactions underlie all atopic and many allergic disorders. The terms atopy and allergy are often used interchangeably but are different. Atopy is an exaggerated IgE-mediated immune response; all atopic disorders are type I hypersensitivity disorders. Vasodilation, edema and bronchial constriction secondary to mediators released by mast cells after interaction with IgE-linked allergen comprise the early events of immediate hypersensitivity (Carvalho et al., 2006). Allergy is any exaggerated immune response to a foreign antigen regardless of
154
Aditya Reddy and Bernard Fried
mechanism. Thus, all atopic disorders are considered allergic, but many allergic disorders (e.g. hypersensitivity pneumonitis) are not atopic; allergic disorders are the most common disorders among people. Atopic disorders most commonly affect the nose, eyes, skin and lungs. Allergic atopic disorders, such as rhinitis, asthma and atopic dermatitis, are the result of a systemic inflammatory reaction triggered by type 2 T helper (Th2) cell-mediated immune responses against antigens (allergens) of complex genetic and environmental origin (Romagnani, 2004). These disorders include atopic dermatitis and contact dermatitis (which may be primary skin disorders or symptoms of systemic disorders), latex allergy, allergic lung disorders (e.g. asthma, allergic bronchopulmonary aspergillosis, hypersensitivity, pneumonitis) and allergic reactions to venomous stings. Many studies demonstrate parallel increasing prevalence of asthma and rhinitis, but in regions of highest prevalence, it may be reaching a plateau (Cruz et al., 2007a).
3. RELATIONSHIP OF PARASITES TO ATOPIC DISORDERS Both helminth infection and allergy are common diseases and it has been noted that helminth infection is negatively associated with atopy, prevalence of allergic diseases and the severity of asthma (Carvalho et al., 2006). It is estimated that about one-quarter of the world’s population, 1.5 billion, are infected with one or more of the major soil-transmitted helminths, including hookworms, ascarids and whipworms (Kamal and Sayed, 2006). The prevalence of asthma and rhinitis is also high in countries where helminths are endemic (Asher and Weiland, 1998), and studies that evaluate the immunological and clinical consequences of this association are examined in this chapter. Interestingly, the prevalence of asthma is increasing mainly in the developed world where 130 million people worldwide are estimated to suffer from the condition (Sears, 1997, 1998). Irrespective of improved knowledge of many aspects of atopic diseases, the unfavourable trends in their prevalence particularly among children are significant; a growing body of evidence suggests that something may lack from our societal affluence that has the capacity to provide protection against the development of atopic diseases (von Hertzen and Haahtela, 2004). A number of epidemiological studies have suggested that the increase in the prevalence of allergic disorders that has occurred over the past few decades is attributable to a reduced microbial and parasitic burden during childhood, as a consequence of westernized lifestyle (Romagnani, 2004). However, specific mechanisms by which the reduced exposure of children to pathogenic and non-pathogenic microbes results in enhanced responses of Th2 cells are still controversial (Yazdanbakhsh et al., 2002).
Atopic Disorders and Parasitic Infections
155
Atopy, measured by the presence of allergen-specific IgE in sera or skin test reactivity to allergen extracts, has been found to be a strong risk factor for asthma in developed countries (Beasley et al., 2001), but the proportion of atopics with clinical asthma (or the proportion of asthmatics with atopy) appears to be much smaller in some rural areas of Europe (Priftanji et al., 2001) and in many developing nations (Cooper et al., 2003; Davey et al., 2005). Additionally, epidemiologic studies have pointed to a protective role of helminthic infections in the development of allergy and asthma; however, evidence for this inverse association has not been consistently established (Arruda and Santos, 2005). Importantly, not only intestinal helminth parasites but also several other pathogens and apparently also many commensals are able to elicit the anti-inflammatory (regulatory) network including the regulatory T cells that secrete IL-10 and TGFb, the anti-inflammatory cytokines, that inhibit deleterious immunopathologic responses (Maloy and Powrie, 2001). The secretion of these antiinflammatory cytokines, particularly IL-10, by regulatory T cells appears now to hold the key to counter-regulation of both atopic and autoimmune diseases (Gale, 2002). Helminths, similar to allergens also induce a type 2 immune response and though pathology can be documented in some patients infected with worms, the majority of helminth-infected individuals have an asymptomatic form of infection or present mild clinical manifestations (Carvalho et al., 2006). Reddy and Fried (2007) noted that long-lived helminthic parasites are remarkable in their ability to down-regulate host immunity, protecting themselves from elimination, and also minimize severe pathological host changes in inflammatory diseases such as Crohn’s. As an example of mild or asymptomatic infection, the larvae (juvenile forms) of Ascaris lumbricoides may induce eosinophilia and high total IgE levels ( Joubert et al., 1979; Li et al., 2000) that are either of no clinical consequence or are able to elicit severe pulmonary reactions resembling type I hypersensitivity during their migration through the lungs (Lo¨ffler’s syndrome) (Kennedy, 2000). Parasite-specific IgE antibodies act to exclude parasites from the host, whereas non-specific IgE antibody production during parasite invasion is involved in the host evasion of those parasites; it is thus important to clarify whether antibody production is as a result of the infection of parasites and in the immune-evasive mechanisms system of those parasites (Muto et al., 2001). Th2 responses induced by allergens or helminths share many common features; however, though allergen-specific IgE can almost always be detected in atopic patients, helminth-specific IgE is often not detectable and anaphylaxis often occurs in atopy but not helminth infections (Erb, 2007). Erb (2007) suggested that this may be due to T regulatory responses induced by the helminths or the lack of helminth-specific IgE. Alternatively,
156
Aditya Reddy and Bernard Fried
non-specific IgE induced by the helminths may protect from mast cell or basophil degranulation by saturating IgE binding sites; both of these mechanisms have been found to be involved in helminth-induced protection from allergic responses (Erb, 2007). Additionally, Redecke et al. (2004) indicated that Toll-like receptor stimulation during the initial phase of immune activation determines the polarization of the adaptive immune response and may play a role in the initiation of Th2-mediated immune disorders, and induces experimental asthma. Westernized countries are suffering from an epidemic rise in immunologic disorders, such as childhood allergy; a popular explanation is that the increased prevalence in allergy is due to a diminished or altered exposure to gut-dwelling microbes and helminths, resulting in abnormal immunoregulation (Smits et al., 2005). Renewed interest in microorganisms, not as initiators or inciters of asthma, but as protectors against atopic diseases, began finally after the publication of the three pioneer studies on the inverse association between childhood infections and the subsequent development of asthma and atopy (Matricardi et al., 1997; Shaheen et al., 1996; Shirakawa et al., 1997). In allergic individuals antibodies are largely responsible for initiating hypersensitivity reactions such as asthma, whereas in helminth infections, the IgE production is thought to be responsible for a protective immune response to the parasite, as well as for immune-mediated pathology (Weiss, 2000). Increasingly, there is evidence of important effects on other innate cell types, particularly mast cells, dendritic cells and eosinophils (Maizels et al., 2004). The sum effect of these immune changes to host reactivity is to create an anti-inflammatory environment, which is most favourable to parasite survival (Maizels et al., 2004). Helminths have demonstrated the ability to down-regulate the type 1 inflammatory response and attenuate autoimmune disease in experimental models (La Flamme et al., 2003; Sabin et al., 1996). Studies have shown a strong inverse relationship between atopy and geohelminth infection, indicating that atopy may protect against geohelminth infection. Cooper et al. (2004a) found that resistance to ascariasis in atopic individuals may occur through greater immunoglobulin E-mediated responses and expression of T helper cell type 2 (Th2) cytokines to parasite antigens. To investigate the effect that atopy has on the immune response to Ascaris antigens, Cooper et al. (1993) recruited school-age children from rural schools in Ecuador. In the study, immunologic variables were compared between children stratified by atopic and/or A. lumbricoides infection status; the variables included cytokine expression by peripheral blood mononuclear cells (PBMCs) and histamine release in response to A. lumbricoides antigens. Atopic children had both greater frequencies of PBMCs expressing IL-4 and IL-5 and enhanced histamine release, compared with the same parameters in non-atopic children. Stratification by
Atopic Disorders and Parasitic Infections
157
atopic and A. lumbricoides infection status found the greatest histamine and Th2 cytokine responses in the epidermis of atopic, non-infected children. The authors used multivariate regression analyses to show significant effects for atopic status but not for infection status on Th2 cytokine expression and histamine release (Cooper et al., 2004a). Infection with Schistosoma mansoni or exposure to eggs from this helminth inhibits the development of type 1 diabetes in non-obese diabetic (NOD) mice when soluble extract injections are begun at 4 weeks; NOD mice are deficient in natural killer T cells (Zaccone et al., 2003). Hosts deploy a set of defence mechanisms against these parasites that together control infection by most members of that class, even though a specific defence mechanism may not be required to defend against a particular parasite and may even damage a host infected with that parasite (Finkelman et al., 1997). In the La Flamme et al. (2003) study, analysis of the cellular composition of murine spinal cords and brains revealed that a pre-established infection with S. mansoni decreased central nervous system inflammation, particularly of macrophages and CD4 T cells. The authors suggested that S. mansoni may negatively regulate the onset of experimental autoimmune encephalomyelitis by down-regulating the production of pro-inflammatory cytokines and altering CNS inflammation. In humans it is known that S. mansoni infection is inversely associated with a positive skin test (PST) response to aeroallergens (Araujo et al., 2000; van den Biggelaar et al., 2000). In the Araujo et al. (2000) study subjects infected by S. mansoni with more than 200 eggs/g of faeces (n = 42) and uninfected subjects (n = 133) were selected from an endemic area of schistosomiasis. The history of allergy and results of the immediate hypersensitivity prick tests with inhalant allergen extracts were registered and total IgE and IgE specific to S. mansoni and aeroallergens were measured in serum by ELISA. There was a tendency for higher total and S. mansoni-specific IgE levels in infected patients, an opposite trend, that is higher aeroallergen-specific IgE, was observed in uninfected subjects. A strong and statistically significant inverse association was found between the immediate skin test response to common aeroallergens and infection by S. mansoni; this indicates that immediate hypersensitivity reactions may be suppressed in S. mansoni-infected individuals (Araujo et al., 2000). van den Biggelaar et al. (2000) tested 520 Gabonese schoolchildren for skin reaction to house-dust mites and other allergens, for Schistosoma haematobium eggs in urine, and for microfilariae in blood samples. There has been nearly an epidemic rise in allergic disease throughout the world; however, this significant increase in the prevalence of allergic diseases has not been reported on the African continent (Ndiaye and Bousquet, 2004). In the van den Biggelaar et al. (2000) study, both total and mite-specific IgE antibodies were measured; a subsample selected on the basis of their skin test to house-dust mite received detailed
158
Aditya Reddy and Bernard Fried
immunological investigations. In the study, children with S. haematobium had a lower prevalence of a positive skin reaction to house-dust mite than those free of this infection; the degree of sensitization to house-dust mites could not explain this difference in skin prick positivity. In addition, schistosome antigen-specific concentrations of IL-10 were significantly higher in infected children, and higher specific concentrations of this anti-inflammatory cytokine were negatively associated with the outcome of skin test reactivity to mites. The authors concluded that the antiinflammatory cytokine, IL-10, induced in chronic S. haematobium infection, appears central to suppressing atopy in African children (van den Biggelaar et al., 2000). Further, worms may attenuate the clinical manifestations of asthma (Medeiros et al., 2003). Medeiros et al. (2003) subjects were from 3 low-socioeconomic areas: a rural area endemic for schistosomiasis (group 1) in addition to a rural area (group 2) and a slum area (group 3), both of which were not endemic for schistosomiasis. Symptom frequency, use of anti-asthma drugs and pulmonary abnormal findings at physical examination were less in group 1 subjects than in group 2 and 3 subjects (p = 0.0001). Initial data that helminth infections were inversely associated with atopy were published in the 1970s and, more recently, several studies have shown that helminths can protect against allergy (Araujo et al., 2000; Carswell et al., 1976; Cooper et al., 2004b; Jarrett and Kerr, 1973; Lynch et al., 1993; van den Biggelaar et al., 2002). Jarrett and Kerr (1973) demonstrated a high incidence of infection with the pinworm Enterobius vermicularis with allergic asthma, but also in a non-allergic group; the study in which there is no evidence of infection with any other helminth parasites. Presence or absence of pinworm infection was not clearly correlated with differences in total serum IgE level in either allergic or non-allergic children although levels of this immunoglobulin were raised in the allergic group ( Jarrett and Kerr, 1973). The authors therefore suggested that hypersensitivity to E. vermicularis allergen absorbed from the bowel might contribute to the allergic symptoms. Carswell et al. (1976) studied two rural Tanzanian primary schools to test the hypothesis that parasitic infection prevents the development of asthma; 242 pupils were interviewed to determine the prevalence of pupils with recurrent episodes of wheezing. The authors found no difference in immediate cutaneous hypersensitivity to 22 allergens between the asthmatics and controls and therefore suggested that parasitic infection does not prevent the development of asthma. The study by Cooper et al. (2004b) was more conclusive: a total of 1002 children from seven rural schools were recruited in an area in which the prevalence of geohelminth infections was high (70.1% were infected with at least one geohelminth parasite) and the prevalence of allergic sensitization was high (20.0% had evidence of aeroallergen sensitization). Factors associated with significant
Atopic Disorders and Parasitic Infections
159
protection against atopy in multivariate analyses were the presence of overcrowding in the child’s home, low socio-economic level and infection with the geohelminth parasites (A. lumbricoides and Trichuris trichiura), and the protective effects of the three factors were all statistically independent (Cooper et al., 2003). The authors concluded that low socioeconomic level, overcrowding and geohelminth infection, are all independently protective against atopy among school-age children living in a rural area of Latin America. Parasites have a built-in anti-inflammatory defence, and there is increasing evidence that these defence mechanisms include effects on regulatory T cells, of which there are three populations, some secreting anti-inflammatory cytokines such as transforming growth factor-b (TGF-b) and IL-10, and many others that seem to depend on contact with activated T cells to survive (Leonardi-Bee et al., 2006). Leonardi-Bee et al. (2006) suggested that despite mounting evidence that this population of cells is expanded during parasite infection, the pathologic impact of this expansion is not yet clear. High levels of IL-4, IL-5, IL-13, eosinophils and production of high IgE levels suggest that worm infection predisposes individuals to allergic reactions (Carvalho et al., 2006). This statement is supported by evidence that helminth proteases exhibit significant homology with known allergens, and that allergic manifestations in the skin and the respiratory tract during acute helminth infection can be documented (Donnelly et al., 2006). The cysteine protease of dust mite, Der p1, the aspartic protease of cockroach, Bla g 2, the serine protease of Aspergillus fumigatus and the bacterial subtilisins are all major allergenic molecules responsible for the increase in asthma and atopic conditions worldwide (Donnelly et al., 2006). These proteases induce Th2-driven inflammatory responses in the airways by disrupting the epithelial cell junctions so that these, and other molecules, gain access to, and alter the function of, underlying cells of the innate immune system (dendritic cells, mast cells, basophils and macrophages) as well as B and T cells (Donnelly et al., 2006). The authors state that the anti-inflammatory responses observed in chronic helminthiases, involving IL-10 and TGF-b, that are primarily responsible for controlling immune-mediated damage to the host that is initiated by secreted proteases, actually protect against similar inflammatory damage by allergens. Helminth parasites secrete proteases to gain entry into their hosts, and to feed on and migrate through tissues; the helminth activity leads to tissue damage and the activation of inflammatory responses dominated by elevated IgE, eosinophilia and Th2 cells, much like allergenic responses. Studies conducted in Caracas, Venezuela, showed a low prevalence of PST to aeroallergens in individuals living in an endemic area of Ascaris lumbricoides infection, but after anthelmintic treatment, there was an increase in the number of positive skin prick test (SPT) responses (Lynch et al., 1993).
160
Aditya Reddy and Bernard Fried
It is well known that helminthic infection can cause a polyclonal stimulation of the synthesis of IgE, which is dependent on IL-4 production, and it has been suggested that this can modulate the expression of allergic reactivity in tropical areas (Lynch et al., 1993). Lynch et al. (1993) evaluated the effect of regular anthelmintic treatment, for a period of 22 months, on certain aspects of the allergic reactivity of children in a slum area of Caracas where helminths are endemic and found that inhibition of allergic reactivity is reversible by anthelmintic treatment. Human ascariasis is associated with a polarized Th2-type immune response, with elevated levels across a range of serum immunoglobulins. While some authors propose that parasite-specific antibody responses may confer a degree of protection from re-infection, others suggest that such responses merely reflect infection intensity without involving protective mechanisms. Another opinion suggests that such antibody responses do protect, but only indirectly as a consequence of inflammatory processes (King et al., 2005). A recent study suggested that immune response to A. lumbricoides (Ascaris-sIgE) may be a risk factor of atopic disease in populations exposed to mild A. lumbricoides infection and that Mycobacterium tuberculosis infection may be protective against this risk, probably by stimulation of anti-inflammatory networks (Obihara et al., 2006). Geohelminths, that is, ascarids, hookworms and whipworms, are prevalent infections of childhood and might contribute to the low prevalence of allergic disease reported from rural areas of the tropics; Cooper et al. (2003) sought to establish whether geohelminth infections protect against atopy and to explore whether this protection is dependent on infection chronicity in a study of a total of 2865 children aged 5–19 years from 55 schools. The authors found that active infections with geohelminth parasites and the presence of serologic markers of chronic infections (high levels of total serum IgE or anti-A. lumbricoides IgG4) are independent protective factors against allergen skin test reactivity among school-age children living in an endemic region of the rural tropics (Cooper et al., 2003). Cooper (2002) stated that in the case of infections such as A. lumbricoides and Trichuris trichiura that are acquired at an early age, and constant exposure occurs throughout childhood, school-age children with the heaviest parasite burdens would be expected to have the lowest rates of atopy and allergic disease. In the study, active infection with any geohelminth and infections with A. lumbricoides or Ancylostoma duodenale was also associated with significant protective effects against allergen skin test reactivity. Children with the highest levels of total IgE or with anti-A. lumbricoides IgG4 antibodies were protected against skin test reactivity, and the protective effects of high IgE or anti-A. lumbricoides IgG4 and or active geohelminth infections were statistically independent (Cooper et al., 2003).
Atopic Disorders and Parasitic Infections
161
Chronic blood and tissue parasite infections that are capable of modulating immune responses in the host are negatively associated with skin test reactivity in a sensitized population; a low frequency of positive SPT response to dust mites was documented in a pivotal study among school-age children residing in Gabon, Africa (van den Biggelaar et al., 2001). The study took place in an area of Gabon endemic for blood and tissue parasites; 520 schoolchildren were parasitologically examined and skin prick tested for a set of common environmental aeroallergens. In schoolchildren, schistosome and filarial infections increased with age, whereas malaria was more prevalent in younger children; the authors found that, in contrast to allergen sensitization that increased with age, skin test reactivity tended to decline. The number of children with mitespecific IgE antibodies (47%) far exceeded the number responding to skin prick testing (11%) and mite sensitization was found to be the highest in children infected with schistosomes and/or filariae whereas skin test reactivity was lowest (van den Biggelaar et al., 2001). Although the majority of studies in this field have shown an inverse association between helminth infection and atopy, there are a few reports associating geohelminths with an increased risk of asthma (Guemont, 1973; Kayhan et al., 1978; Palmer et al., 2002). Kayhan et al. (1978) studied 50 patients with bronchial asthma and 50 normal controls of whom 40% of the patients with bronchial asthma and 14% of the controls had ascarids in the stool examinations. The authors noted that the significant difference in these two groups indicated that intestinal parasites should be looked for in patients with bronchial asthma. Palmer et al. (2002) found that independently of the other factors assessed, infection with A. lumbricoides was associated with increased risk of asthma (p < 0.001), an increased number of skin tests positive to aeroallergens (p < 0.001). In the study, the interrelationships of current and past infection with A. lumbricoides and asthma and atopy were investigated in a cross-sectional sample of 2164 children between the ages of 8 and 18 years from Anqing Province, China. The association of sensitization to common aeroallergens with increased asthma risk was enhanced in children infected with A. lumbricoides, and the infection was associated with an increased risk of asthma independent of sensitization to aeroallergens in this selected population (Palmer et al. 2002). These authors suggested that the complex relationship between ascariasis and susceptibility to childhood asthma among predisposed children may involve an interaction with the immune response to inhaled aeroallergens. A possible explanation for these conflicting data includes age of the population, parasite burden, the time of the exposure to the worm (acute or chronic) and also the species of helminth (Carvalho et al., 2006). Hosts may have evolved the ability to recognize features that characterize parasitic gastrointestinal nematodes as a class of triggers for a stereotypic cytokine response, but may not have the ability to
162
Aditya Reddy and Bernard Fried
distinguish features of individual parasites as stimuli for more specific protective cytokine responses (Finkelman et al., 1997). Individuals infected with a low parasite burden of A. lumbricoides did not differ on the frequency of positive SPT to dust mites from those living without infection in the same area (Ponte et al., 2006). The cross-sectional study of 113 patients with asthma or rhinitis was conducted in a region endemic for geohelminths. Stool examinations and SPTs to aeroallergens were performed in all the patients; Der p-specific IL-10 production was measured in the supernatant of peripheral blood mononuclear cell (PBMC) cultures of a subsample of 53 patients. Ponte et al. (2006) concluded that in patients with asthma or rhinitis living in an urban area endemic for geohelminths, there was no association between Ascaris infection and skin reactivity to aeroallergens and no difference in Der p-specific IL-10 production by PBMCs. The authors stated that these negative findings indicate that in contrast to what was observed in Schistosoma infection, A. lumbricoides infection in individuals living in an urban area does not induce strong regulatory responses (Ponte et al., 2006). Similarly, among African children with bronchial asthma evaluated for A. lumbricoides skin-specific IgE, 27% of 270 children were shown to have detectable specific IgE to the A. lumbricoides antigen, compared to 8% of controls (Aderele and Oduwole, 1982). In a study of about 2300 East German children, those who were Ascaris-IgE seropositive had tenfold higher levels of total IgE and higher prevalence rates of allergen-specific IgE seropositivity, allergic rhinitis and asthma. The study concluded that contact with low doses of helmintic antigen is associated with an increase of total and specific IgE production and that helmintic infections in East German children are not causative factors for a low prevalence of allergies in the former East Germany (Dold et al., 1998). In an early South African study utilizing skin-specific IgE determination to evaluate the incidence of allergic asthma in A. lumbricoides infected patients, 17% of the nonallergic controls and 51% of the allergic asthmatics had a clinically detectable immunogenic response to the parasite. The predicted incidence of asthma was significantly higher than the observed incidence in the subjects in whom skin-specific IgE had been found. Inhalation of A. lumbricoides antigen-induced asthmatic reactions in 7 of 8 patients who were Ascaris-positive on skin testing, but not in the negative controls ( Joubert et al., 1979). Scha¨fer et al. (2005) conducted a study to investigate the association between worm infestation and eczema in a proper temporal sequence and under consideration of allergic sensitization. Their data support the concept that a lack of immune-stimulation by parasitic infections contributes to the development of allergies. Two surveys were performed in East German school children for which questionnaire data included the history of eczema and worm infestation and their time of onset. Specific IgE
Atopic Disorders and Parasitic Infections
163
antibodies to five common aeroallergens were measured and used to define both non-atopic and atopic eczema. Logistic regression analyses were performed to control for relevant confounding variables (age, sex, parental school education and history of allergies). In order to confirm the findings the authors applied a corresponding conditional regression analysis on cases and controls matched by age and sex. A total of 4169 children participated who were on average 9.2 years old (47% girls). Overall 17.0% reported a prior worm infestation (Ascaris 44%, E. vermicularis 33%) and 18.1% of the children had a history of eczema. Eczema occurred significantly less frequently in children who had a worm infection (prior to the onset of eczema) compared with children without a history of infection. The authors found that atopic eczema was affected more by a prior worm infestation than the non-atopic eczema (Scha¨fer et al. 2005). A total of 29.1% of the children exhibited specific IgE antibodies to at least one aeroallergen. Sensitized children gave significantly less history of personal worm infection. In this study, worm infection was associated with a reduced frequency of subsequent eczema, especially atopic eczema. The authors concluded that allergic sensitization, especially to HDM, and worm infection are negatively associated (Scha¨fer et al., 2005). Lynch et al. (1987) found that the prevalence of positive SPT to allergens was lower in individuals with high parasite burden of geoheminths compared to those with a lower parasite burden in Caracas, Venezuela. The study was conducted on the premise that as some factors associated with the tropical environment can modify the expression of atopic disease, various indicators of allergic reactivity merit comparison between allergic and non-allergic subjects of different socioeconomic levels (Lynch et al., 1987). Furthermore, Araujo et al. (2000) demonstrated lower prevalence of SPT response to aeroallergens in individuals infected with a high parasite burden of S. mansoni living in a rural area, compared to noninfected individuals living in the same area. In the study, subjects infected by S. mansoni with more than 200 eggs/g of faeces (n = 42) and uninfected subjects (n = 133) were selected from an endemic area of schistosomiasis. Interestingly, the proportion of individuals with a PST to allergens was higher in the uninfected group (24.3%) than in the group with more than 200 eggs/g of faeces (4.8%) (Lynch et al., 1987). The history of allergy and results of the immediate hypersensitivity prick tests with inhalant allergen extracts were documented; total IgE and IgE specific to S. mansoni and aeroallergens were measured in serum by ELISA. The study indicates that immediate hypersensitivity reactions may be suppressed in S. mansoniinfected individuals (Lynch et al., 1987). Araujo and de Carvalho (2006) noted that cells from asthmatic individuals infected with S. mansoni produced lower levels of IL-5 than asthmatics free of infections. The authors discussed the association between S. mansoni infection, atopy and severity of asthma and possible
164
Aditya Reddy and Bernard Fried
mechanisms by which individuals living in helminth endemic areas are protected against the development of allergies. In contrast, the authors noted that IL-10 is more readily produced by allergen-stimulated cells from asthmatics who are infected and is detected only at low levels by cells from helminth-free asthmatics (Araujo and de Carvalho, 2006). It is well known that Th2 cytokines are involved in the pathogenesis of allergies and asthma, and some studies indicate that IL-10 is the key cytokine that inhibits the Th2-inflammatory response in allergy (Donnelly et al., 2006). Medeiros et al. (2004) evaluated the association between exposure to dust mites and skin reactivity to mite allergens in subjects with a history of wheezing in the last 12 months selected from a rural endemic area for schistosomiasis (group I, n = 21), and two non-S. mansoni endemic locale, a rural area (group II, n = 21) and a urban slum area (group III, n = 21). Although skin reactivity to indoor allergens is decreased in subjects from helminthic endemic areas, the degree of exposure to mite allergens had not been investigated in these areas prior to this pivotal study. In the study, all subjects were evaluated by SPTs with mite allergens, and for total and specific immunoglobulin E (IgE) against dust mites, antibodies for S. mansoni, and for intestinal parasites. Dust samples from subjects’ homes were quantified for mite allergens and the species of the mite also identified. The authors found that, except for S. mansoni infection which was more prevalent in group I than in groups II and III (p < 0.0001), the prevalence of intestinal parasites, and total and specific IgE levels were similar for all groups. Despite the levels of mite allergens and specifically to Der p 1 detected in dust samples of subjects homes from all three areas, the frequency of positive skin reactivity to mite antigens was significantly lower (19.0%) in subjects from group I relative to group II (76.2%) and group III (57.1%; p < 0.001). These results suggested that S. mansoni infection could modulate the immediate hypersensitivity of skin response to mite allergens in highly exposed individuals (Medeiros et al., 2004) Leonardi-Bee et al. (2006) conducted a systematic review and meta-analysis of epidemiologic studies to determine whether total or species-specific current parasite infection is associated with a reduced risk of asthma or wheeze. Leonardi-Bee et al. (2006) searched MEDLINE, EMBASE and CINAHL (up to January 2006); reviews; and reference lists from publications and included studies that reported asthma or wheeze as an outcome measure and ascertained parasite infection by faecal examination. Pooled odds ratios (OR) and 95% confidence intervals (CI) were estimated by Leonardi-Bee et al. (2006) using data obtained from published papers, or where available, original data provided by authors, using random effect models. In the 33 studies which met the inclusion criteria, the authors found that infection with any parasite was associated with a small, non-significant increase in asthma risk (OR, 1.24; 95% CI,
Atopic Disorders and Parasitic Infections
165
0.98–1.57; 29 studies). In species-specific analysis, Ascaris lumbricoides was associated with significantly increased odds of asthma (OR, 1.34; 95% CI, 1.05–1.71; 20 studies), while hookworm infection was associated with a significantly strong reduction (OR, 0.50; 95% CI, 0.28–0.90; 9 studies) that was directly and significantly related to the severity of infection (p < 0.001). Other helminth species had no significant effects on asthma; infection effects on wheeze were derived from smaller numbers, but revealed a broadly similar pattern of results (Leonardi-Bee et al., 2006). Leonardi-Bee et al. (2006) suggested that an alternative explanation is that the associations observed in the study rise from reverse causation, and that, especially, allergic individuals are less likely to acquire hookworm infection. The cross-sectional nature of the data available to the authors precluded any further insight into this possibility. Overall, the Leonardi-Bee et al. (2006) study indicated that different species of parasite infection may have important effects on the pathogenesis of asthma, and that in particular, the potential individual or public health benefits of hookworm infection merit further investigation. Another factor that Carvalho et al. (2006) mentioned as an explanation for the conflicting results regarding the role of helminth infections on the development of allergies is that most published data did not consider the fact that exposure to allergens is required to develop a SPT response. Despite difficulties demonstrating in vivo that worms may be killed by components of the type 2 immune response, there is evidence from experimental models that IL-4 and IL-13 contribute to parasite elimination in stool by stimulating peristalsis and increasing intestinal fluid (Finkelman et al., 1997). Studies with rodents infected with Trichinella spiralis, Heligmosomoides polygyrus, Nippostrongylus brasiliensis and Trichuris muris have provided considerable information about immune mechanisms that protect against parasitic gastrointestinal nematodes (Finkelman et al., 1997). The focus of a review by Arruda and Santos (2005) was to discuss the potential role of shared antigens between parasites and environmental allergens in modulating allergic immune responses, specifically tropomyosin. It has been postulated that Ascaris lumbricoides and other helminths could share antigens capable of inducing cross-reactive IgE antibody responses after exposure via the inhalation route, such as the antigens in mites and cockroaches; one example is tropomyosin, a protein associated with strong IgE antibody responses, which is highly conserved in invertebrates (Arruda, 2005). Interestingly, cross-reactivity was reported to IgE-binding proteins from Anisakis simplex, Blattella germanica (German cockroach) and unidentified chironomids (midge-like larvae); there was also a strong association among Pandalus borealis (Atlantic shrimp), A. lumbricoides and Daphnia magna (the water flea), but this was not confirmed in laboratory studies; the common allergen in these diverse species may have been tropomyosin (Pascual et al., 1997).
166
Aditya Reddy and Bernard Fried
The relationship of asthma and allergy to parasitic infection could vary according to the particular kinds of parasites; however, the molecular basis of these effects is largely unknown. As the biological cycle of A. lumbricoides results in pulmonary passage of larvae, and other nematodes, including hookworms and Strongyloides stercoralis, also undergo pulmonary passage of larvae, it may be that A. lumbricoides allergen, on the evidence of the reactions it causes, is the most potent of all allergens of parasitic origin (Arruda, 2005). It may be that exposure to cross-reactive allergens such as tropomyosin at the initial A. lumbricoides infection could facilitate subsequent development of cross-reactive IgE antibody responses upon exposure to mite or cockroach, and that this could lead to airway inflammation and asthma. It may be that infection with A. lumbricoides would have an adjuvant effect on the development of asthma in a subset of A. lumbricoides infected children who become sensitized to the tropomyosin allergen. Experimental work by Kennedy (2000) has demonstrated a dramatic alteration in the parasite’s antigens, which occurs during its development. A. suum larvae have been shown to change the composition of both their secreted and surface antigens during the tissue-invasive stage of infection, thus expressing quite different antigens than those provoking pulmonary hypersensitivity responses (Kennedy, 2000). This may be similar for A. lumbricoides, and likely direct inflammatory responses in the lungs caused by parasite antigens can account for wheezing and other respiratory symptoms. Children potentially immune to A. lumbricoides were identified in an area of Nigeria where infection is hyperendemic in a study by McSharry et al. (1999). Among those individuals who produced IgG antibody to recombinant ABA-1 allergen, a major allergen of both A. lumbricoides and A. suum, the naturally immune group had significantly more IgE antibody to the allergen than did those susceptible to the infection. IgE antibody responses in conjunction with innate inflammatory processes, therefore, appear to associate with natural immunity to ascariasis in this community (McSharry et al., 1999). Despite exceptionally high total IgE levels in the study, there was no evidence that atopic responses to local common allergens were associated with natural immunity to Ascaris spp. A number of studies have reported sensitization and subsequent allergic symptoms to Ascaris spp., including a report of 41 patients with eosinophilic asthma-associated A. lumbricoides infection (Li et al., 2000). Ascaris spp. material is known to provoke allergic reactions in laboratory workers (Ogilvie and de Savigny, 1982). Additionally, A. lumbrioides has been associated with an increased risk of wheezing, asthma and allergic sensitization in certain populations (Palmer et al., 2002; Sales et al., 2002). It is also possible that the response of the induction of large quantities of IgE antibody is in part directed against the helminths’ own antigens, but that a polyclonal stimulation also occurs that may increase the allergic
Atopic Disorders and Parasitic Infections
167
reactivity towards environmental allergens. A study demonstrated that regular anthelmintic treatment resulted in significant improvement in all clinical indicators of asthma, not only during the period of anthelmintic administration, but also for the following year (Lynch et al., 1997). However, after 2 years without treatment, the severity of asthma reverted to its initial level. The authors concluded that intestinal helminthic infections can contribute to the clinical symptoms of asthma in an endemic situation (Lynch et al., 1997). However, as mentioned, a more recent study found that anthelmintic treatment of chronically infected children resulted in increased atopic reactivity, arguing that helminths directly suppress allergic reactions (van den Biggelaar et al., 2004). Children in whom atopic and allergic responses are controlled by peripheral tolerance to cross-reactive antigens may continue to suppress allergic inflammation under circumstances of continued exposure to geohelminth infection, but loss of exposure caused by migration or sustained anthelmintic treatment may cause peripheral tolerance to decrease with activation of allergic responses (Lynch et al., 1993; van den Biggelaar et al., 2004). In contrast, children born to uninfected mothers and who live in circumstances of low-level transmission to geohelminth parasites may react with enhanced allergic responses to early geohelminth exposures (Gelpi and Mustafa, 1967), and suppression of these infections by anthelmintic treatment may actually cause an improvement in allergic symptoms (Lynch et al., 1997). The effect of geohelminth infection on wheeze and allergen sensitization is inconsistent across various studies. Davey et al. (2005) investigated the association between self-reported wheeze, self-reported asthma, allergic sensitization and geohelminth infection in urban and rural areas of Butajira, southern Ethiopia. Questionnaire data on wheeze, asthma and a range of confounding variables was gathered in a cross-sectional study of 7649 people aged 5 years or older. Allergic skin sensitization to the dust mite Dermatophagoides pteronyssinus and cockroach was measured, and a stool sample collected for qualitative and quantitative geohelminth analysis. The authors found a weak association between allergic sensitization and wheeze. Pereira et al. (2007) hypothesized that helminths attenuate allergic asthma, whereas other factors are related to the expression of a non-atopic wheeze/asthma phenotype. Although asthma is common in urban centers in Latin America, atopic asthma may not be the predominant phenotype among children (Pereira et al., 2007). In the study, a total of 1982 children from Southern Brazil with a mean SD age of 10.1 0.76 years completed asthma questionnaires, and 1011 were evaluated for intestinal parasites and atopy using SPTs. Wheeze in the previous 12 months was reported by 25.6% and 9.3% showed current asthma; 13% were SPT-positive and 19.1% were positive for any helminth infection. Most children with either
168
Aditya Reddy and Bernard Fried
wheeze or asthma were SPT-negative; however, the study found that severe wheeze was more prevalent among the atopic minority; helminth infections were inversely associated with positive SPT results. Bronchiolitis, inflammation of the arterioles, before the age of 2 years was the major independent risk factor for asthma at age 10 years; high-load A. lumbricoides infection, a family history of asthma and positive SPT results were also asthma risk factors. In a typical case, an infant under 12 months of age develops cough, wheeze and shortness of breath over 1 or 2 days. Pereira et al. (2007) concluded that most asthma and wheeze are of the non-atopic phenotype, suggesting that some helminths may exert an attenuating effect on the expression of the atopic portion of the disease, whereas viral bronchiolitis predisposes more specifically to recurrent airway symptoms. Another study by Karadag et al. (2006) focused on 613 children 6–13 years of age from rural areas of Austria, Germany and Switzerland, who took part in the Allergy and Endotoxin study. Allergic diseases and farming characteristics were assessed by a standardized questionnaire and as a crude measure of possible exposure to helminths, IgG antibodies to A. lumbricoides were studied by the authors. Results of the study show that exposure to ascarids, as determined by the levels of antibody to A. lumbricoides, was more frequent among farmers’ children than nonfarmers’ children (39.8% vs 31.1%, p = 0.03). This positive serology was found to be significantly associated with high total IgE levels and eosinophilia. However, there was no association between anti-nematode serology and the prevalences of asthma, wheeze, hay fever or atopy; a weak association for atopy was observed after adjustment for total IgE (Karadag et al., 2006). Karadag et al. (2006) concluded that immunoglobulin G antibodies to A. lumbricoides, as a crude measure of possible exposure to helminths, did not indicate any protective effect against allergic diseases in this population, though farmers’ children had increased antibody levels reactive to helminth parasites indicating exposure; this could not explain the protective effect of farming against atopic diseases. Conversely, other epidemiological studies in developing countries suggest that intestinal parasite infection may reduce the risk of asthma (Cooper et al., 2003). In the Cooper et al. (2003) study, the presence of geohelminth infections was protective against allergen skin test reactivity (p < 0.001) and symptoms of exercise-induced wheeze (p = 0.008) but not against other wheeze symptoms or symptoms of allergic rhinitis or atopic eczema. The study found infection intensity with A. lumbricoides or Trichuris trichiura was associated with a reduction in the prevalence of allergen skin test reactivity but not with allergic symptoms. The authors concluded that geohelminth infections did not explain the low prevalence of allergic symptoms in the study population. In a survey of 7155 children aged 1–4 years living in urban and rural areas of Jimma,
Atopic Disorders and Parasitic Infections
169
Ethiopia, the conclusion was reached that Ascaris and possibly hookworm infection protected against wheeze in young children (Dagoye et al., 2003). Infection with parasites in the study population was common, predominantly with T. trichuris (54%), A. lumbricoides (38%) and hookworm (10%). Wheezing in the past year was significantly more prevalent in urban (4.4%) than rural children (2.0%), and was less prevalent in those infected with A. lumbricoides (age, sex and urban/rural adjusted odds ratio, 0.5; 95% confidence interval, 0.3–0.9), particularly in relation to high-intensity infection. The effects of protection against wheeze were not mediated by inhibition of allergen sensitization (Dagoye et al., 2003). A. lumbricoides may also cause chronic urticaria (or hives) as a result of its close association with the parasite larva Anisakis simplex; this has been suggested as the cause of acute urticaria and anaphylaxis in a clinical study by Lopez-Saez et al. (2003). Urticaria is generally caused by direct contact with an allergenic substance, or an immune response to food or some other allergen, but can also appear for other reasons, notably emotional stress. In 101 patients with chronic urticaria, 35% were found to have skin-specific IgE to A. simplex and serum-specific IgE to A. simplex was positive in 55%. A total of 22% of all the patients had detectable serum-specific IgE to A. lumbricoides, and of these 91% had serum-specific IgE to A. simplex. The authors concluded that sensitization to A. simplex is higher among patients with chronic urticaria and that sensitization to other parasites occurs because of cross-reactivity, but that a causal relationship between the presence of specific IgE to A. simplex and chronic urticaria had not been fully established (Lopez-Saez et al., 2003). Perzanowski et al. (2002) evaluated differences in the relationship between asthma and immune responses to allergens in children living in rural and urban areas of Kenya. Children (mean age, 11 years) from a rural village (n = 136) and a small town (n = 129) were studied by skin testing and IgE and IgG antibody measurement. In the study, asthma was evaluated by symptoms, as well as spirometry before and after vigorous exercise to test for exercise-induced bronchospasm. School children from a prior study performed in Atlanta, Georgia, were used for comparison of anthropometric and immunologic results. Compared with the urban area of Kenya, children living in the rural area had a lower percentage of body fat, smaller and fewer skin test responses to allergens, a higher prevalence of IgE antibodies to A. lumbricoides (67% vs 26%) and tenfold higher total IgE. In the urban area of Kenya, the authors found a strong correlation between bronchospasm and atopy determined both by IgE antibodies (p = 0.02) and skin tests (p = 0.002). By contrast, in the rural area, none of the 13 children with bronchospasm were skin-test positive (vs 13/109 of children without bronchospasm). The study found that among the rural children, there was no association between immune responses to allergens and airway-related symptoms or reactivity. The authors
170
Aditya Reddy and Bernard Fried
conclude that the association between asthma and atopy seen in the town may represent an important step in the increase in asthma seen both in urban Africa and in the West (Perzanowski et al., 2002). Ferreira et al. (2007) investigated the prevalence and risk factors for wheezing and asthma in young Amazonian children. They found an increased risk of asthma was independently associated with early introduction of bottle feeding, defined as <4 months of exclusive breastfeeding (p = 0.033), and seropositivity to the zoonotic nematode Toxocara canis (p = 0.016), suggesting two potential targets for public health interventions (Ferreira et al. 2007). A population-based cross-sectional survey of 606 children aged 6–59 months was performed in two small towns in Acre State, Northwestern Brazil. Information on outcome variables (recent wheezing and medical diagnosis of asthma) and demographic, socioeconomic, environmental, maternal and nutritional variables was obtained by interviewing children’s mothers or immediate family. In the study, infections with intestinal parasites and antibodies to T. canis were diagnosed using standard laboratory techniques. Multiple unconditional logistic regression models were used to describe associations between both independent variables and outcomes. The prevalence of recent wheezing (defined as one or more reported episodes in the past 12 months) was 42.6%, but only 19.8% of wheezing children were also reported to have a medical diagnosis of asthma (prevalence, 8.5%); 21.5% of the children in the Ferreira et al. (2007) study had antibodies to T. canis. The study by Ogorodova et al. (2007) aimed to estimate the relationship between the prevalence of allergic disease and helminth invasion by the liver fluke Opisthorchis felineus in rural and urban populations in West Siberia, Russia. Two hundred and one people from a village and 196 from the city were screened for the presence of atopy and O. felineus invasion. Opisthorchiasis was found in 66 participants (32.8%) from the village and in 22 of the city subjects (11.2%). It was found that atopic diseases were more common in the urban population than in the rural: 52.8% and 31.4%, respectively. Positive skin prick tests were significantly higher in the urban population than in rural people, 83.2% versus 24.4%, respectively. It was found that in the city, the presence of antibodies to O. felineus negatively correlates with the atopic sensitization as determined by SPT. However, in the village setting, opisthorchiasis was positively associated with atopic diseases. The authors noted that the study data confirmed the negative association of rural lifestyle and atopic diseases prevalence and also indicated that O. felineus invasion might be a modifying factor of this relationship. Nyan et al. (2001) conducted a study to determine the prevalence of atopy and intestinal helminth infection and to relate these to wheeze history and serum total IgE in a community sample of adults from an urban and a rural area of the Gambia. The low prevalence of atopy in
Atopic Disorders and Parasitic Infections
171
traditional societies has been attributed to high parasite-driven blocking IgE concentrations (Nyan et al., 2001). Six hundred and ninety-three adults were interviewed about respiratory symptoms using a modified version of the International Union Against Tuberculosis and Lung Disease questionnaire, and had skin prick testing using four allergens. Stools were examined after formol-ether concentration. Total serum IgE concentration was measured in a subset of participants. The prevalence of atopy (mean weal diameter 3 mm) in the urban and rural area was 35.3% and 22.5% (p = 0.05); the dust mite D. pteronyssinus and mould were identified as the most common sensitizing allergens. Prevalence of wheeze history in the previous 12 months was 4.4% and 3.5% for the urban and rural areas, respectively; wheezing was not found to be significantly associated with atopy. Interestingly, there was an inverse association between atopy and intestinal helminth infection; 7% of atopic subjects had helminths, compared to 13% of non-atopic subjects (p = 0.03). Non-atopic individuals had total serum IgE concentrations about 2.5 times the upper limit of the reference range in non-atopic Western populations. The authors concluded that IgE concentration was not associated with the presence of helminth infection and suggest that atopy might protect against helminth infection (Nyan et al., 2001). Scrivener et al. (2001) tested the hypothesis that the risk of asthma is reduced by intestinal parasites or hepatitis A infection, and increased by exposure to dust-mite allergen or organophosphorus insecticides in urban and rural areas of Jimma, Ethiopia. The study of asthma and atopy included 12,876 individuals; those who reported wheeze in the previous 12 months were recorded, and a random subsample of controls was also included. Parasites in faecal samples were assessed, Der p 1 levels in bedding, hepatitis A antibodies, serum cholinesterase (a marker of organophosphorus exposure), total and specific serum IgE and skin sensitization to D. pteronyssinus in 205 cases and 399 controls aged over 16 years. The authors analyzed the effects of parasitosis, Der p 1 level, hepatitis A seropositivity and cholinesterase concentration on risk of wheeze, and the role of IgE and skin sensitization in these associations using multiple logistic regressions. The risk of wheeze was found to be independently reduced by hookworm infection by an odds ratio of 0.48 (p = 0.03), increased in relation to Der p 1 level (p = 0.05) and was unrelated to hepatitis A seropositivity or cholinesterase concentration in the rural population. In the urban population, D. pteronyssinus skin sensitization was more strongly related to wheeze than in the rural areas (p for interaction = 0.017), where D. pteronyssinus sensitization was common, but unrelated to wheeze in the presence of high-intensity parasite infection. The authors concluded that high levels of parasite infection might prevent asthma symptoms in atopic individuals (Scrivener et al., 2001).
172
Aditya Reddy and Bernard Fried
IgG4 has been proposed to act as a ‘blocking antibody’ because of its ability to compete for the same epitopes as IgE thus preventing IgEdependent allergic responses; IgG4 and IgE are both elevated in helminth infections (Turner et al., 2005; Yazdanbakhsh and Wahyuni, 2005). In a study aimed at determining the relationship between anti-parasite IgG4 and IgE antibodies and A. lumbricoides infection status, anti-parasite responses, including antibody levels to recombinant Ascaris allergen-1A (rABA-1A), a target of serum IgE antibodies in endemic populations were examined (Turner et al., 2005). Individuals who had detectable levels of IgE but not IgG4 antibodies to rABA-1A (11%) had lower average levels of infection compared with individuals who produced anti-rABA-1A IgG4 (40%) and sero-negative individuals (49%). The ratio of IgG4/IgE in rABA-1A responders positively correlated with intensity of infection. IgG4 levels positively correlated with infection level in younger children (age 4–11) where average levels of infection were increasing, whereas allergen-specific IgE emerged as a correlate of immunity in older children and adults (age 12–36) where infection levels were decreasing. The study demonstrated that in a gastrointestinal helminth infection, differential regulation of anti-allergen antibody isotypes relate to infection level, and concluded that the results were consistent with the concept that IgG4 antibody can block IgE-mediated immunity and, therefore, allergic processes in humans (Turner et al., 2005). Erb (2007) described the generation of an anti-N. brasiliensis-specific IgE antibody which was used to identify a novel N. brasiliensis antigen (Nb-Ag1). The Nb-Ag1-specific IgE could only be detected for a short period of time during infection, and that these levels were sufficient to prime mast cells thereby leading to active cutaneous anaphylaxis after the application of Nb-Ag1. It is the first report clearly showing that a low level of helminth-specific IgE, transiently produced, was able to induce mast cell degranulation in the presence of large amounts of polyclonal IgE (Erb, 2007). Huang et al. (2002) examined the association between E. vermicularis infection and allergy in primary school children. The authors identified a negative association between pinworm infection and allergic airway diseases, which could partially be attributed to the protective effect of pinworm infection on development of allergic symptoms; other mechanisms of association could not be ruled out (Huang et al. 2002). The peri-anal tape test for E. vermicularis is routinely performed in Taipei primary schools; Huang et al. (2002) collected data from school records and questionnaires distributed to all children in four primary schools grades 1 through 6 (n = 3107). In the study, the prevalence of physician-diagnosed asthma (9.3% vs 14.1%, p = 0.007) and allergic rhinitis (27.4% vs 38.3%, p = 0.001) was lower in E. vermicularis positive compared to uninfected children. E. vermicularis was not correlated with atopic dermatitis or
Atopic Disorders and Parasitic Infections
173
parent allergy; with logistic regression controlling for sex, parent allergy and lower respiratory infection, current asthma and rhinitis were negatively associated with pinworm. The authors found that, in children in grades 3–6 who had no asthma or rhinitis before age 7, those with early infection (pinworm diagnosed at or before grade 1) had a lower risk of having a diagnosis of rhinitis during school years, compared to the uninfected group (Huang et al., 2002). There are no published data available on how maternal geohelminth infections and early infant infections may affect the occurrence of food allergy or the development and progression of the atopic disorders (Cooper et al., 2004b). Early exposure to geohelminth parasites could affect the early expression of allergic disease, prevent the development of atopic dermatitis in infancy or contribute to milder clinical forms of disease, and reduce the risk of severe asthma and allergic rhinitis or asthma symptoms later in life (Cooper et al., 2004b). Geiger et al. (2007) described hookworm interactions with their human hosts by comparing lymphocyte phenotyping, proliferative responses, and cytokine and chemokine secretion patterns in adults who are either mono-infected with Necator americanus or egg-negative controls resident in an area of high transmission in Brazil. The authors propose that the longevity of hookworms in their human hosts results from a stagespecific, down-modulation of the immune response. Cellular immune responses against crude hookworm antigen extracts from various developmental stages were evaluated simultaneously. In the study, principal component analysis was used to reduce the standardized immune responses; random effects multivariate regression was then used to investigate whether principal components differ between the two groups once potential confounders and effect modifiers had been accounted for. The authors found that although hookworm patients had reduced percentages of T and B cells, they had higher levels of activated CD4+ T and CD19+ B cells (Geiger et al., 2007). This state of host ‘immune activation’ coincided with lower proliferative responses, especially to third-stage larval antigen; cytokine levels in mono-infected adults were also lower and characterized by a mixed Th1/Th2-type profile. Excretory/secretory antigen obtained from adult worms was found to be a potent modulator of the immune response, resulting in diminished TNF-a and IL-10 secretion in PBMCs from hookworm-infected patients (Geiger et al., 2007). Signal transducer and activator of transcription 6 (Stat6) is vital for Th2-mediated responses during allergic airway disease. Alonso-Trujillo et al. (2007) used susceptible BALB/c mice and resistant STAT6-/- BALB/ c mice to analyze the role of nitric oxide (NO) in determining the outcome of murine cysticercosis caused by the cestode Taenia crassiceps. The level of exhaled NO is increased in patients with allergic asthma and seasonal rhinitis (Gratziou et al., 1999). NO is one of the few gaseous signalling
174
Aditya Reddy and Bernard Fried
molecules known. It is a key biological messenger, playing a role in a variety of biological processes.NO, known as the ‘endothelium-derived relaxing factor’, or ‘EDRF’, is biosynthesized from arginine and oxygen by various nitric oxide synthase (NOS) enzymes and by reduction of inorganic nitrate. The endothelium of blood vessels uses NO to signal the surrounding smooth muscle to relax, thus dilating the artery and increasing blood flow. Cruz et al. (2007b) conducted an exploratory study to investigate the safety of omalizumab (anti-IgE monoclonal antibody) in subjects with allergic asthma and/or perennial allergic rhinitis at high risk of intestinal helminth infection. The primary safety outcome in the study was risk of infections with intestinal helminths during anti-IgE therapy. A randomized, double-blind, placebo-controlled trial was conducted in 137 subjects (12–30 years) at high risk for geohelminthic infections. All subjects received pre-study anthelmintic treatment, followed by 52 weeks’ treatment with omalizumab or placebo. Of the omalizumab subjects 50% (34/68) experienced at least one intestinal geohelminth infection compared with 41% (28/69) of placebo subjects (p = 0.14), providing some evidence for a potential increased incidence of geohelminth infection in subjects receiving omalizumab. Omalizumab therapy was well tolerated, and did not appear to be associated with increased morbidity attributable to intestinal helminths as assessed by clinical and laboratory adverse events, maximal helminth infection intensities and additional anthelmintic requirements. Time to first infection was similar between the two treatment groups. Cruz et al. (2007b) concluded that omalizumab therapy is safe and well tolerated, but may be associated with a modest increase in the incidence of geohelminth infection. The study by Alonso-Trujillo et al. (2007) was conducted to clarify immune mechanisms that underlie resistance and susceptibility to cysticercosis, which are not completely understood. After T. crassiceps infection, wild-type BALB/c mice developed a strong Th2-like response, produced high levels of IgG1, IgE, IL-5, IL-4 and discrete levels of NO, yet remained susceptible to T. crassiceps infection. Interestingly, similarly infected BALB/c mice treated with No-nitro-L-arginine methyl ester (L-NAME, an inhibitor of NO synthase) mounted a similar immune response but with lower levels of NO and harboured nearly 100% more parasites than No-nitro-D-arginine methyl ester (D-NAME, inactive enantiomer)-treated mice. To further clarify the role of NO in murine cysticercosis, the authors treated STAT6-/- male mice (known to be highly resistant to T. crassiceps) with L-NAME during 8 weeks of infection. As expected, STAT6-/- mice mounted a strong Th1-like response, synthesized high levels of IgG2a, IFN-g and IL-17, while their macrophages displayed increased transcription of tumour necrosis factor (TNF)-a as well as inducible nitric oxide synthase (iNOS) and efficiently controlled
Atopic Disorders and Parasitic Infections
175
T. crassiceps infection. However, STAT6-/- male mice receiving L-NAME mounted a similar immune response but with lower iNOS transcripts concomitantly with decreased levels of NO in sera and, as mentioned, displayed significantly higher parasite burdens. The authors suggested that macrophage activation and NO production are effector mechanisms that importantly contribute in host resistance to T. crassiceps infection (Alonso-Trujillo et al., 2007). The aim of a study by Gratziou et al. (1999) was to investigate the significance of atopy on NO production in the lower airways. Measurements of exhaled NO were performed in 131 stable asthmatic patients with chronic mild asthma (95 atopics and 36 non-atopics), 72 patients with perennial rhinitis (57 atopics and 15 non-atopics) and in 100 healthy controls (20 atopics and 80 non-atopics). Patients with either asthma or rhinitis had higher exhaled NO values (13.3 1.2 parts per billion and 11.7 1.1 ppb) than controlled subjects (p < 0.01). Exhaled NO levels were significantly higher in atopic asthmatics (19 3.6 ppb) compared with non-atopic patients (5.6 0.8 ppb, p < 0.001). Similar findings were observed in patients with rhinitis (13.3 1.3 ppb in atopics and 5.8 1.2 ppb in non-atopics, p < 0.001). No difference was found in NO levels between atopic and non-atopic control subjects (4.8 0.8 ppb and 4.5 0.3 ppb). The authors have shown that increased exhaled NO levels are detected only in atopic patients with asthma and/or rhinitis and not in non-atopic patients. These findings may suggest that the allergic nature of airway inflammation is largely responsible for the higher NO production in the lower airways (Gratziou et al., 1999).
4. LABORATORY STUDIES ON ATOPY USING SELECTED PARASITES AND RODENT MODELS Murine and other animal models have been useful in understanding the mechanisms of the development of atopic disorders. Martin (1976) found that experimental injection of atopic serum and the serum of patients infected with Ascaris into the peritoneal cavity of rats after 24 h produced an increase in mesenteric mast cells along with vascular congestion. The increase of mast cells was shown to have statistical significance in atopic serum (hay fever and atopic dermatitis) as well as in serum of patients infected with Ascaris. Martin (1976) stated that although it is known that the serum from atopic patients and those infected with Ascaris has a high level of IgE, it is unknown if this increase in mast cells in the rat is produced by human IgE. Studies regarding the role of T cells in allergic asthma have been facilitated by the development of mouse models of this disease, though these models do not accurately reflect human asthma they mimic cardinal asthma manifestations (Motta and van Oosterhout, 2006).
176
Aditya Reddy and Bernard Fried
Though T helper type 2 lymphocytes are considered as the main effector cells in the pathology of atopic asthma, new T cells subtypes have been characterized during the past decade, which have greatly extended our vision on the immunoregulation of this disease (Motta and van Oosterhout, 2006). Several studies have shown an inverse association between exposures to Toxoplasma gondii or harbouring of Schistosoma sp. or intestinal helminths and allergy or mentioned allergic symptoms (van der Kleij and Yazdanbakhsh, 2003). It is known that migration of helminth larvae through the tissues of their mammalian hosts can cause considerable pathology through the IgE response or eosinophilic inflammation, yet the evolutionary factors responsible for this migratory behaviour are poorly understood (Mingomataj et al., 2006). Navigation and survival in an array of different habitants requires costly biochemical and morphological adaptations (Read and Skorping, 1995). Living in an exposed extracellular niche, in confrontation with a potentially hostile environment, the persistent, chronic lifestyle of helminths is persuasive evidence in itself for their profound ability to modulate their hosts’ immune response (Else, 2005). The most successful human helminth of the western world is the pinworm E. vermicularis, and some 50% of young children in Europe and North America may have been infected by the organism around the middle of the twentieth century (Gale, 2002). Helminths may inhibit the development of atopic disease via induction of regulatory T cells and secretion of Il-10, and Gale (2002) noted that pinworms inhibit diabetes development in the NOD mouse. Pinworms usually are benign, often asymptomatic and may have immunomodulatory properties that protect against the development of immune-mediated disorders including diabetes and asthma; the author concluded that their decline in response to improved living conditions may explain a number of features of the epidemiology of childhood atopy and diabetes (Gale, 2002). Reece et al. (2006) utilized the transient pulmonary phase of N. brasiliensis development to study the innate immune responses induced by this helminth parasite in wild-type (WT) and severe-combined immune deficient (SCID) BALB/c mice. While it is well established that infection with the rodent hookworm N. brasiliensis induces a strongly polarized Th2 immune response, little was known about the innate host-parasite interactions that lead to the development of this robust Th2 immunity (Reece et al., 2006). In the study, histological analysis demonstrated that the cellular infiltrates caused by N. brasiliensis transit through the lungs were quickly resolved in WT mice but not in SCID mice. The authors used microarray-based gene expression analysis to demonstrate that there was a rapid induction of genes encoding molecules that participate in innate immunity and in repair/remodelling during days 2–4
Atopic Disorders and Parasitic Infections
177
post-infection in the lungs of WT and SCID mice. Of particular note was the rapid up-regulation in both WT and SCID mice of the genes encoding YM1, FIZZ1 and Arg1, indicating a role for alternatively activated macrophages in pulmonary innate immunity. Immunohistochemistry demonstrated that nearly all alveolar macrophages became YM1-producing macrophages as early as day 2 post-infection. The authors found that while the innate responses induced during the lung phase of N. brasiliensis infection were similar in complexity and magnitude in WT and SCID mice, only mice with functional T cells were capable of maintaining elevated levels of gene expression beyond the innate window of host reactivity. The authors concluded that induction of alternatively activated alveolar macrophages could be important for dampening the level of inflammation in the lungs and contribute to the long-term decrease in pulmonary inflammation that has been associated with many helminth infections (Reece et al, 2006). Brazis et al. (1998) studied the potential effect of an increase in serum IgE on mast cell activity by analyzing the histamine releasability of mature mast cells isolated from the skin of atopic A. lumbricoides sensitive and in healthy dogs. No histamine release was detected upon the immunological stimulation of cells not previously sensitized with atopic or A. lumbricoides sensitive dog serum; however, when passively sensitized, skin mast cells were challenged with either Asc SI antigen or anti-IgE, the mast cell histamine release increased in a stimulus concentrationdependent manner (Brazis et al., 1998). The authors suggested that, since there is either little or no increase in the density of IgE receptors in atopic or Ascaris hypersensitive dogs versus controls, the digestive process might affect cell behaviour in vitro or that an underlying increase of receptors affects the release of granule-stored mediators but influences mast cell activity in a differential manner (Brazis et al., 1998). Trujillo-Vargas et al. (2007) conducted a study to evaluate if N. brasiliensis excretory–secretory (NES) products prevent the development of asthma. In the study, mice were immunized with ovalbumin/alum intraperitoneally in the absence or presence of helminthic products and then challenged intranasally with ovalbumin; 6 days later mice were analyzed to see if they had developed Th2 responses in the lung. Their results showed that mice immunized with NES and ovalbumin generated NESbut not ovalbumin-specific IgE clearly indicating that N. brasiliensis produce substances which drive a helminth-specific IgE response under limiting conditions in which an allergen-specific IgE response cannot be generated. The authors found that application of the helminthic products together with ovalbumin/alum during the sensitization period totally inhibited the development of eosinophilia and goblet cell metaplasia in the airways and also strongly reduced the development of airway hyperreactivity (Trujillo-Vargas et al., 2007). Allergen-specific IgG1 and IgE
178
Aditya Reddy and Bernard Fried
serum levels were also significantly reduced; these findings correlated with decreased levels of IL-4 and IL-5 in the airways in NES-treated animals. The study found suppressive effects on the development of allergic responses that were independent of the presence of Toll-like receptors 2 and 4, IFN-g, and most important, IL-10 which has been studied extensively. Interestingly, Trujillo-Vargas et al. (2007) found that suppression was still observed when the helminthic products were heated or treated with proteinase K; paradoxically, they also found that strong helminth product-specific Th2 responses were induced in parallel with the inhibition of ovalbumin-specific responses. In a recent review, Knox (2007) stated that there is now substantial evidence to show that nematode parasites utilize these proteinase inhibitors to protect themselves from degradation by host proteinases, to facilitate feeding and to manipulate the host response to the parasite. Infections with N. brasiliensis can also induce dominant type 2 responses from antigen-specific T helper cells; the potency of the Th2 bias can also drive Th2 responses to bystander antigens introduced at the same time as infection (Holland et al., 2000). Holland et al. (2000) found that the Th2-promoting effect of infection can be reproduced with soluble NES released by adult parasites in vitro. Immunization of BALB/c mice with NES results in the production of IL-4 with elevated total serum IgE and specific IgG1 antibodies; NES is also able to stimulate IL-4 and polyclonal IgE production in other mouse strains (C57BL/6, B10.D2, CBA). In the study, these findings occurred whether NES is administered without adjuvant as soluble protein in phosphate-buffered saline or with complete Freund’s adjuvant which normally favours Th1 responses. Adjuvants are agents which modify the effect of other agents while having few if any direct effects when given by themselves. In this sense, they are very roughly analogous with chemical catalysts. NES therefore possesses intrinsic adjuvanticity in the Holland et al. study. Moreover, co-administration of hen’s egg lysozyme (HEL) with NES in the absence of other adjuvants resulted in generation of HEL-specific lymphocyte proliferation, IL-4 release and IgG1 antibody responses. The authors documented that NES can act as an adjuvant for third-party antigens (Holland et al., 2000). Interestingly, proteinase K digestion or heat treatment of NES before immunization abolished the IL-4-stimulating activity, indicating that the factors acting to promote Th2 induction are proteins secreted by the adult parasite. Wang et al. (2001) conducted a study to determine if a pre-existing helminth infection would increase or decrease subsequent allergic responses to an unrelated allergen in the lungs. In the study, BALB/ cByJ mice were infected with the nematode S. stercoralis prior to ovalbumin (OVA) immunization and to an intratracheal challenge. Bronchoalveolar lavage (BAL) and Bronchoalveolar lavage fluid (BALF) were
Atopic Disorders and Parasitic Infections
179
collected 3 days post-challenge and cellular and humoral immune responses were quantified. Intracellular cytokine staining demonstrated increased IL-4 and IL-5 producing cells in BAL from mice infected with S. stercoralis before OVA sensitization; increased IL-5 protein levels and decreased IFN-g protein levels were also observed in the BALF (Wang et al., 2001). In the study there was, however, no increase in airway eosinophil accumulation in mice infected with parasites prior to sensitization with OVA as compared to mice exposed to the OVA alone. Furthermore, levels of the small cytokine eotaxin in the lungs induced by OVA were suppressed in mice infected with the parasite before OVA sensitization. In response to the presence of allergens, eotaxin directly promotes the accumulation of eosinophils, a prominent feature of allergic inflammatory reactions. The development of OVA-specific IgE responses in BALF was likewise impaired in mice infected with the parasite before sensitization with OVA. The authors concluded that a pre-existing helminth infection may potentiate a systemic Type 2 response yet simultaneously suppress in the lungs allergen-specific IgE responses and eotaxin levels in response to subsequent exposure to allergens (Wang et al., 2001). Lima et al. (2002) investigated the influence of extracts from Ascaris suum (ASC) on the development of pulmonary eosinophilic inflammation in a murine model of asthma. Adult worm extracts suppress the IgE antibody production against unrelated antigens. In the study, heatcoagulated egg white alone (EWI) or mixed with ASC (EWI + ASC) was implanted subcutaneously in B10.A or C57BL/6 mice, and 14 days later they were challenged intratracheally with OVA or exposed to aerosolized OVA for 4 days. The suppressive effect of ASC was demonstrated on the accumulation of cells into airways, with subsequent reduction of eosinophil numbers and of eosinophil peroxidase activity in EWI + ASC-immunized mice. In addition, this effect correlated with a marked reduction of IL-5 and IL-4 levels in the BAL from C57BL/6 and B10. A mice, respectively, and of eotaxin in BAL and lung tissue from both strains. OVA-specific IgG1 and IgE levels were also impaired in serum and BAL from these mice; airway hyper-reactivity to methacholine was obtained in B10.A mice sensitized with EWI, but the respiratory mechanical parameters returned to normal levels in both EWI + ASC-immunized mice. These authors concluded that ASC has a profound inhibitory effect on lung inflammation and hyper-responsiveness and that suppression of IL-5 or IL-4 and of eotaxin contributes to this effect (Lima et al., 2002). Pinto et al. (2006) found that Angiostrongylus costaricensis infection in mice decreased pulmonary inflammatory response to OVA. A. costaricensis is a nematode of wild rodents with widespread occurrence in Central and South America; transmission to humans may occur by ingesting infected slugs (intermediate host) or contaminated food or water (Morera, 1988). Geiger et al. (2001) demonstrated a predominantly Th1 cytokine profile in
180
Aditya Reddy and Bernard Fried
A. costaricensis infected BALB/c mice, with the gut as the primary site of infection. This nematode was therefore seen as useful as an experimental model by Pinto et al. (2006) for the study of the association between asthma and intestinal parasitic infection. In their study, Pinto et al. (2006) infected seven BALB/c mice with A. costaricensis by orogastric gavage (10 larvae/mouse) on day (D) 0. The mice were immunized against OVA by intraperitoneal injection on D 5 and D 12 and received an intranasal OVA challenge (40 ml) on D 15 and D 17. On D 19 BAL was performed. In the study, six BALB/c mice (control group) were immunized with OVA using the same protocol, but were not infected with A. costaricensis. Interleukin (IL)-1b and IL-6 levels were measured in the BAL fluid using commercial ELISA assays; total cell counts and differential cell counts were performed in the BAL fluid samples. The group infected with A. costaricensis had lower total cell count in the BAL fluid when compared with the control group. BAL fluid IL-1b levels from the infected group were significantly lower than those of the control group. IL-6 levels in BAL fluid were not different between the groups studied (Pinto et al., 2006). Promising results were obtained from murine asthma models in which IFN-g, given either during the primary sensitization or during the secondary immune response, decreased both IgE production and airway inflammation and also normalized airway function (Lack et al., 1994, 1996). In addition, systemic as well as pulmonary IFN-g gene delivery was proven to be effective not only for preventing but also for suppressing established allergen-induced airway hyper-responsiveness (AHR) in a mouse model (Dow et al., 1999; Li et al., 1996). However, no beneficial effects were observed in clinical trials on asthma patients with subcutaneous or aerosolized IFN-g as was observed in the murine models (Boguniewicz et al., 1995; Martin et al., 1993). Doligalska et al. (2007) evaluated levels of apoptosis and the immune response in BALB/c mice infected with H. polygyrus. H. polygyrus infection in mice is associated with reduced local and systemic immune responses, thus providing an ideal model to study the mechanisms of immune regulation (Rzepecka et al., 2007). In the study, cell proliferation, apoptosis and cytokine production were measured in mesenteric lymph nodes (MLN) without exposure to H. polygyrus antigens in culture. The authors concluded that inhibited apoptosis and cytokine production reported might reflect a state of cell hyporesponsiveness in the prepatent phase of infection. These changes were accompanied by changes in the percentage of CD4+ cells in MLN and popliteal lymph nodes; lymph node changes are noted in several allergic and atopic disorders (Doligalska et al., 2007). In another study by Rzepecka et al. (2007) the process by which H. polygyrus infection modulates the influx of eosinophils into the airways of asthmatic mice was investigated. The authors observed
Atopic Disorders and Parasitic Infections
181
a reduction in the total number and percentage of lung eosinophils that coincided with decreased levels of eotaxin in BALF, lower expression of the CCR3 receptor on eosinophils and impaired chemotaxis of these cells towards eotaxin. Allergen-induced immune response were therefore down-regulated as production of Th1 (IFN-g)-, Th2 (IL-4, IL-5)- and Treg (IL-10)-related cytokines as well as IL-6 and TNF-a levels which diminished upon nematode infection. On the basis of the study, the authors concluded that the attenuation of allergic inflammation during H. polygyrus infection is a consequence of the dichotomy of the immune response in the face of concurrent antigenic challenge. Data from Wilson et al. (2005) support the contention that helminth infections elicit a regulatory T cell population able to down-regulate allergen-induced lung pathology in vivo. Allergic diseases mediated by Th2 cell immune responses are rising dramatically in most developed countries and exaggerated Th2 cell reactivity could result, for example, from diminished exposure to Th1 cell inducing microbial infections (Wilson et al., 2005). Go´mez-Garcı´a et al. (2006) studied the role of intact carbohydrate structures on T. crassiceps compounds in the induction of biased type 2 and anti-inflammatory immune responses on peptidestimulated T cells by using DO11.10 transgenic (OVA Tg) mice. The authors findings suggest that carbohydrate components in T. crassiceps soluble antigens are involved in modulating immune responses to bystander antigens and that do not signal via TLR4 (Go´mez-Garcı´a et al., 2006). On the basis that Th2 cell stimulating helminth parasites may also counteract allergies, possibly by generating regulatory T cells which suppress both Th1 and Th2 arms of immunity, Wilson et al. (2005) tested the ability of the Th2 cell inducing gastrointestinal nematode H. polygyrus to influence experimentally induced airway allergy to ovalbumin and the HDM allergen Der p 1. In the study, inflammatory cell infiltrates in the lung were suppressed in infected mice compared with uninfected controls; suppression was reversed in mice treated with antibodies to CD25. Interestingly, the authors found that suppression was transferable with mesenteric lymph node cells (MLNC) from infected animals to uninfected sensitized mice, demonstrating that the effector phase was targeted. MLNC from infected mice contained elevated numbers of CD4+ CD25+ Foxp3+ T cells, higher TGF-b expression and produced strong IL-10 responses to the parasite antigen. However the authors found that MLNC from IL-10-deficient animals transferred suppression to sensitized hosts, indicating that IL-10 is not the primary modulator of the allergic response. Murine host suppression was associated with CD4+ T cells from MLNC, with the CD4+ CD25+ surface marker defining the most active cell population. Jenkins et al. (2005) explored the various lines of evidence that excretory/ secretory molecules from S. mansoni cercariae down-regulate the host’s
182
Aditya Reddy and Bernard Fried
immune response. They highlighted the immunological factors that were produced and may be involved in regulating the immune system (e.g. IL-10 and eicosanoids), as well as speculating on possible mechanisms of immune modulation (e.g. mast-cell activation, T-cell apoptosis and/or the skewed activation of antigen-presenting cells). The authors also drew attention to several previously mentioned molecules of S. mansoni origin that have the potential to stimulate the regulatory response (e.g. glycans) and link these to potential host receptors (e.g. TLRs and C-type lectins) (Jenkins et al., 2005).
5. CONCLUDING REMARKS It is probable that down-regulatory immune mechanisms, which dampen the anti-parasite response, might benefit the host by blocking progression to atopic reactions (Yazdanbakhsh et al., 2001). The protective effect of helminths is probably mediated by regulatory T lymphocytes; co-evolutionary partners might have contributed to the development of this form of response, and parasites and the indigenous biota of the gut are plausible candidates (Gale, 2002). Cooper et al. (2004a) states that a causal association between geohelminth infections and allergy remains to be proven. The strongest epidemiological evidence for a causal association between geohelminth infection and allergy is provided by intervention studies that demonstrate evidence for an effect of anthelmintic treatment on atopy or asthma risk (Cooper et al., 2004a). The vast majority of helminth infections do not lead to associated clinical morbidity even in those areas where such infections are a significant public health concern (Bundy et al., 2004). The early development and maturation of ‘the immunoregulatory network’ (Maizels and Yazdanbakhsh, 2003), in which regulatory T cells truncate strong inflammatory responses generated in response to infectious exposures such as geohelminths, could be an important mechanism for the avoidance of potentially damaging allergic inflammatory responses to both environmental allergens and parasite antigens (Cooper, 2002). The suspected key players for helminth infection-mediated immune suppression of autoimmunity and atopy are T regulatory cells and dendritic cells, which produce immunosuppressive cytokines, such as IL-10, IL-5 and transforming growth factor-b (Kamradt et al. 2005). Araujo and de Carvalho (2006) explain the inverse correlation between helminth infections and atopy as factors including competition between helminth-induced polyclonal IgE and aeroallergen-specific IgE for high-affinity receptors present on mast cells, increased number of regulatory T cells and high levels of regulatory cytokines produced during helminthic infections. Cooper et al. (2006) also note that the timing of exposure to geohelminth infections
Atopic Disorders and Parasitic Infections
183
may be of particular importance in determining the effect of geohelminths on allergy and atopic disorders. Early exposures (in utero, if the mother is infected or in early infancy) to parasites could have important qualitative or quantitative effects on immune maturation and the development of appropriate immune regulation (Cooper et al., 2006). While many microparasites such as viruses, bacteria and protozoans escape immune attack by antigenic variation or sequestration in specialized niches, helminths appear to flourish in exposed extracellular locations, such as the lymphatics, bloodstream or gastrointestinal tract (Maizels et al., 2004). The proposed role of helminth parasities could be one of immunomodulation rather than disease induction, possibly mediated by interaction with other influences upon the development of the mucosal immune system; this hypothesis could be tested in casecontrol studies by the development of serological markers or skin testing (Gale, 2002). Cooper et al. (2004a) suggested that both prospective and intervention studies are required which investigate the development of allergy in early life at a time when children are first exposed to geohelminth parasites and their antigens. Chronic helminth infections, but not acute infections, may be associated with the expression of regulatory networks necessary for down-modulating allergic immune responses to harmless antigens (Smits and Yazdanbakhsh, 2007). The lack of consensus in the literature could be explained by design variability in individual studies (e.g. cross-sectional and uncontrolled confounding); or it may reflect real differences, if the modulatory effects of geohelminth infections on allergic reactivity markedly differ for different geohelminths, are modified by prevalence of geohelminth infections or are due to timing of exposure in relation to immune maturation or sensitization (Cooper et al., 2006). In a comprehensive review, Flohr (2003) found no clear evidence for a direct relationship between atopic dermatitis and endoparasites. The authors noted that this may be due to the overall small number of studies and insufficient methodological rigor in the existing body of research (Flohr, 2003). Cooper et al. (2006) stated that an explanation for the conflicting findings of epidemiological studies is that geohelminths decrease the risk of allergy in areas of high infection prevalence and increase the risk of allergy in areas of low prevalence. They found that chronic geohelminth infections are inversely associated with allergy, and anthelmintic treatment may increase the prevalence of allergy (Cooper et al., 2006). Their review focused on the relationship between geohelminth parasites and allergy and referred to other helminth infections (e.g. schistosomiasis) where relevant. The authors cautiously stated that observations on geohelminths are not necessarily generalizable to other helminth parasites, particularly helminth parasites that parasitize the vascular (e.g. schistosomes)
184
Aditya Reddy and Bernard Fried
and lymphatic (e.g. some filarial parasites), vessels and that may have differential modulatory effects on host immunity (Cooper et al., 2006). Some helminthic substances, especially the cytokines, could have respiratory and anti-allergic effects, and may therefore become useful as therapeutic modalities for many atopic and allergic disorders. If studied further, identification of these underlying mechanisms could open the way to new forms of immune intervention.
REFERENCES Aderele, W. I., and Oduwole, O. (1982). Ascaris and bronchial asthma in children. Afr. J. Med. Med. Sci. 11, 161–166. Alonso-Trujillo, J., Rivera-Montoya, I., Rodrı´guez-Sosa, M., and Terrazas, L. I. (2007). Nitric oxide contributes to host resistance against experimental Taenia crassiceps cysticercosis. Parasitol. Res. 100, 1341–1350. Akbari, O., Stock, P., De Kruyff, R. H., and Umetsu, D. (2003). Role of regulatory T cells in allergy and asthma. Curr. Opin. Immunol. 15, 627–633. Araujo, M. I., and de Carvalho, E. M. (2006). Human schistosomiasis decreases immune responses to allergens and clinical manifestations of asthma. Chem. Immunol. Allergy 90, 29–44. Araujo, M. I., Lopes, A. A., Medeiros, M., Cruz, A. A., Sousa-Atta, L., Sole, D., and Carvalho, E. M. (2000). Inverse association between skin response to aeroallergens and Schistosoma mansoni infection. Int. Arch. Allergy Immunol. 123, 145–148. Arruda, L. K. (2005). Tropomyosin in parasites—A crossreactive IgE-binding protein? Allergy Clin. Immunol. Int. 17, 243–245. Arruda, L. K., and Santos, A. B. (2005). Immunologic responses to common antigens in helminthic infections and allergic disease. Curr. Opin. Allergy Clin. Immunol. 5, 399–402. Asher, M. I., and Weiland, S. K. (1998). The International study of asthma and allergies in childhood (ISAAC). ISAAC steering committee. Clin. Exp. Allergy 28, 52–66. Bachert, C., van Cauwenberge, P., and Khaltaev, N. (2002). Allergic rhinitis and its impact on asthma (in collaboration with the World Health Organization. Executive summary of the workshop report. 7–10 December 1999, Geneva, Switzerland). Allergy 57, 841–855. Beasley, R., Pekkanen, J., and Pearce, N. (2001). Has the role of atopy in the development of asthma been over-emphasized? Pediatr. Pulmonol. 23, S149–S150. Boguniewicz, M., Martin, R. J., Martin, D., Gibson, U., Celniker, A., Williams, M., and Leung, D. Y. M. (1995). The effects of nebulized interferon-gamma in asthmatic airways. J. Allergy Clin. Immunol. 95, 133–135. Brazis, P., Queralt, M., de Mora, F., Ferrer, L., and Puigdemont, A. (1998). Comparative study of histamine release from skin mast cells dispersed from atopic, Ascaris-sensitive and healthy dogs. Vet. Immunol. Immunopathol. 66, 43–51. Bundy, D. A. P., Chan, M. S., Medley, G. F., and Jamison, D. (2004). Intestinal nematode infections. In ‘‘The Global Epidemiology of Infectious Diseases’’ (C. J. L. Murray, A. D. Lopez, and C. D. Athers, eds.), pp. 243–300. World Health Organization, Geneva. Burgess, I. (1993). Allergic reactions to arthropods. Indoor Built Environ. 2, 64–70. Carswell, F., Meakins, R. H., and Harland, P. S. (1976). Parasites and asthma in Tanzanian children. Lancet 2, 706–707. Carvalho, E. M., Bastos, L. S., and Araujo, M. I. (2006). Worms and allergy. Parasite Immunol. 28, 525–534.
Atopic Disorders and Parasitic Infections
185
Cooper, P. J. (2002). Can intestinal helminth infections (geohelminths) affect the development and expression of asthma and allergic disease? Clin. Exp. Immunol. 128, 398–404. Cooper, P. J. (2004). Intestinal worms and human allergy. Parasite Immunol. 26, 455–467. Cooper, P. J., Chico, M. E., Rodrigues, L. C., Strachan, D. P., Anderson, H. R., Rodriguez, E. A., and Griften, G. E. (1993). Reduced risk of atopy among school-age children infected with geohelminth parasites in a rural area of the tropics. J. Allergy Clin. Immunol. 111, 995–1000. Cooper, P. J., Chico, M. E., Bland, M., Griffin, G. E., and Nutman, T. B. (2003). Allergic symptoms, atopy, and geohelminth infections in a rural area of Ecuador. Am. J. Respir. Crit. Care Med. 168, 313–317. Cooper, P. J., Chico, M. E., Sandoval, C., and Nutman, T. B. (2004a). Atopic phenotype is an important determinant of immunoglobulin E-mediated inflammation and expression of T helper cell type 2 cytokines to Ascaris antigens in children exposed to ascariasis. J. Infect. Dis. 190, 1338–1346. Cooper, P. J., Chico, M. E., Rodrigues, L. C., Strachan, D. P., Anderson, H. R., Rodriguez, E. A., Gaus, D. P., and Griffin, G. E. (2004b). Risk factors for atopy among school children in a rural area of Latin America. Clin. Exp. Allergy 34, 845–852. Cooper, P. J., Barreto, M. L., and Rodrigues, L. C. (2006). Human allergy and geohelminth infections: A review of the literature and a proposed conceptual model to guide the investigation of possible causal associations. Br. Med. Bull. 79–80, 203–218. Cruz, A. A., Popov, T., Pawankar, R., Annesi-Maesano, I., Fokkens, W., Kemp, J., Ohta, K., Price, D., and Bousquet, J. (2007a). Common characteristics of upper and lower airways in rhinitis and asthma: ARIA update, in collaboration with GA(2)LEN. Allergy 62, 1–41. Cruz, A. A., Lima, F., Sarinho, E., Ayre, G., Martin, C., Fox, H., and Cooper, J. P. (2007b). Safety of anti-immunoglobulin E therapy with omalizumab in allergic patients at risk of geohelminth infection. Clin. Exp. Allergy 37, 197–207. Dagoye, D., Bekele, Z., Woldemichael, K., Nida, H., Yimam, M., Hall, A., Venn, A. J., Britton, J. R., Hubbard, R., and Lewis, S. A. (2003). Wheezing, allergy, and parasite infection in children in urban and rural Ethiopia. Am. J. Respir. Crit. Care Med. 15, 1369–1373. Davey, G., Venn, A. J., Belete, H., Berhane, Y., and Britton, J. R. (2005). Wheeze, allergic sensitisation and geohelminth infection in Butajira, Ethiopia. Clin. Exp. Allergy 35, 301–307. Dold, S., Heinrich, J., Wichmann, H. E., and Wjst, M. (1998). Ascaris-specific IgE and allergic sensitization in a cohort of school children in the former East Germany. J. Allergy Clin. Immunol. 102, 414–420. Doligalska, M., Donskow-Schmelter, K., Rzepecka, J., and Drela, N. (2007). Reduced apoptosis in BALB/c mice infected with Heligmosomoides polygyrus. Parasite Immunol. 29, 283–291. Donnelly, S., Dalton, J. P., and Loukas, A. (2006). Proteases in helminth-and allergen-induced inflammatory responses. Chem. Immunol. Allergy 90, 45–64. Dow, S. W., Schwartze, J., Heath, T. D., Potter, T. A., and Gelfand, E. W. (1999). Systemic and local interferon gamma gene delivery to the lungs for treatment of allergen-induced airway hyperresponsiveness in mice. Hum. Gene Ther. 10, 1905–1914. Else, K. J. (2005). Have gastrointestinal nematodes outwitted the immune system? Parasite Immunol. 27, 407–415. Erb, K. L. (2007). Helminths, allergic disorders and IgE-mediated immune responses: Where do we stand? Eur. J. Immunol. 37, 1170–1173. Ferreira, M. U., Rubinsky-Elefant, G., de Castro, T. G., Hoffmann, E. H., da Silva-Nunes, M., Cardoso, M. A., and Muniz, P. T. (2007). Bottle feeding and exposure to Toxocara as risk factors for wheezing illness among under-five Amazonian children: A population-based cross-sectional study. J. Trop. Pediatr. 53, 119–124.
186
Aditya Reddy and Bernard Fried
Finkelman, F. D., Shea-Donohue, T., Goldhill, J., Sullivan, C. A., Morris, S. C., Madden, K. B., Gause, W. C., and Urban, J. F., Jr. (1997). Cytokine regulation of host defense against parasitic gastrointestinal nematodes: Lessons from studies with rodent models. Annu. Rev. Immunol. 15, 505–533. Flohr, C. (2003). Dirt, worms and atopic dermatitis. Br. J. Dermatol. 148, 871–877. Flo¨istrup, H., Swartz, J., Bergstro¨m, A., Alm, J. S., Scheynius, A., van Hage, M., Waser, M., Braun-Fahrla¨nder, C., Schram-Bijkerk, D., Huber, M., Zutavern, A., von Mutius, E., et al. The PARSIFAL Study Group (2006). Allergic disease and sensitization in Steiner school children. J. Allergy Clin. Immunol. 117, 59–66. Gale, E. A. M. (2002). A missing link in the hygiene hypothesis? Diabetologia 45, 588–594. Geiger, S. M., Abrahams-Sandi, E., Soboslay, P. T., Hoffmann, W. H., Pfaff, A. W., GraeffTeixeira, C., and Schulz-Key, H. (2001). Cellular immune responses and cytokine production in BALB/c and C57BL/6 mice during the acute phase of Angiostrongylus costaricensis infection. Acta Trop. 80, 59–68. Geiger, S. M., Caldas, I. R., Glone, B. E., Campi-Azevedo, A. C., De Oliveira, L. M., Brooker, S., Diemert, D., Correa-Oliveira, R., and Bethony, J. M. (2007). Stage-specific immune responses in human Necator americanus infection. Parasite Immunol. 29, 347–358. Gelpi, A. P., and Mustafa, A. (1967). Seasonal pneumonitis with eosinophilia: A study of larval ascariasis in Saudi Arabs. Am. J. Trop. Med. Hyg. 16, 646–657. Go´mez-Garcı´a, L., Rivera-Montoya, I., Rodrı´guez-Sosa, M., and Terrazas, L. I. (2006). Carbohydrate components of Taenia crassiceps metacestodes display Th2-adjuvant and antiinflammatory properties when co-injected with bystander antigen. Parasitol. Res. 99, 440–448. Gratziou, C., Lignos, M., Dassiou, M., and Roussos, C. (1999). Influence of atopy on exhaled nitric oxide in patients with stable asthma and rhinitis. Eur. Respir. J. 14, 897–901. Guement, J. M. (1973). Bronchial asthma, spasmodic coryza and parasitic diseases. Nouv. Presse Med 2, 2125. Holgate, S. T. (1999). The epidemic of allergy and asthma. Nature 402, B2–B4. Holland, M. J., Harcus, Y. M., Riches, P. L., and Maizels, R. M. (2000). Proteins secreted by the parasitic nematode Nippostrongylus brasiliensis act as adjuvants for Th2 responses. Eur. J. Immunol. 30, 1977–1987. Huang, S. L., Tsai, P. F., and Yeh, Y. F. (2002). Negative association of Enterobius infestation with asthma and rhinitis in primary school children in Taipei. Clin. Exp. Allergy 32, 1029–1032. International Study of Asthma and Allergies in Childhood (ISAAC) Steering Committee (1998). Worldwide variation in prevalence of symptoms of asthma, allergic rhinoconjunctivitis, and atopic eczema: ISAAC. Lancet 351, 1225–1232. Jarrett, E. E., and Kerr, J. W. (1973). Threadworms and IgE in allergic asthma. Clin. Allergy 3, 203–207. Jenkins, S. J., Hewitson, J. P., Jenkins, G. R., and Mountford, A. P. (2005). Modulation of the host’s immune response by schistosome larvae. Parasite Immunol. 27, 385–393. Joubert, J. R., de Klerk, H. C., and Malan, C. (1979). Ascaris lumbricoides and allergic asthma: A new perspective. S. Afr. Med. J. 56, 599–602. Kamal, S. M., and Sayed, K. E. (2006). Immune modulation by helminthic infections: Worms and viral infections. Parasite Immunol. 28, 483–496. Kamradt, T., Goggel, R., and Erb, K. J. (2005). Induction, exacerbation and inhibition of allergic and autoimmune diseases by infection. Trends Immunol. 26, 260–267. Karadag, B., Ege, M., Bradley, J. E., Braun-Fahrlander, C., Riedler, J., Nowak, D., and von Mutius, E. (2006). The role of parasitic infections in atopic diseases in rural schoolchildren. Allergy 61, 996–1001.
Atopic Disorders and Parasitic Infections
187
Kayhan, B., Telatar, H., and Karacadag, S. (1978). Bronchial asthma associated with intestinal parasites. Am. J. Gastroenterol. 69, 605–606. Kennedy, M. W. (2000). Immune response to Anisakis simplex and other ascarid nematodes. Allergy 55, 7–13. King, E. M., Kim, H. T., Dang, N. T., Michael, E., Drake, L., Needham, C., Haque, R., Bundy, D. A., and Webster, J. P. (2005). Immuno-epidemiology of Ascaris lumbricoides infection in a high transmission community: Antibody responses and their impact on current and future infection intensity. Parasite Immunol. 27, 89–96. Knox, D. P. (2007). Proteinase inhibitors and helminth parasite infection. Parasite Immunol. 29, 57–71. La Flamme, A. C., Ruddenklau, K., and Backstrom, B. T. (2003). Schistosomiasis decreases central nervous system inflammation and alters the progression of experimental autoimmune encephalomyelitis. Infect. Immun. 71, 4996–5004. Lack, G., Renz, H., Saloga, J., Bradley, K. L., Loaden, J., Leung, D. Y. M., Larsen, G., and Gelfand, E. W. (1994). Nebulized but not parenteral IFN-gamma decreases IgE production and normalizes airway function in a murine model of allergen sensitization. J. Immunol. 152, 2546–2554. Lack, G., Bradley, K. L., Hamelmann, E., Renz, H., Loader, J., Leung, D. Y. M., Larsen, G., and Gelfand, E. W. (1996). Nebulized IFN-gamma inhibits the development of secondary allergen responses in mice. J. Immunol. 157, 1432–1439. Larsen, F. S., and Hanifin, J. M. (2002). Epidemiology of atopic dermatitis. Immunol. Allergy Clin. North Am. 22, 1–24. Leonardi-Bee, J., Pritchard, D., and Britton, J. The Parasites in Asthma Collaboration (2006). Asthma and current intestinal parasite infection: Systematic review and meta-analysis. Am. J. Respir. Crit. Care Med. 174, 514–523. Li, X. M., Chopra, R. K., Chou, T. Y., Schofield, B. H., Wills-Karp, M., and Huang, S. K. (1996). Mucosal IFN-gamma gene transfer inhibits pulmonary allergic responses in mice. J. Immunol. 157, 3216–3219. Li, C. Y., Li, F., and Qiu, H. L. (2000). Clinical analysis of 41 cases with eosinophil asthma associated Ascaris infection. [Chinese] Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi 18, 223. Lima, C., Perini, A., Garcia, M. L. B., Martins, M. A., Teixeira, M. M., and Macedo, M. S. (2002). Eosinophilic inflammation and airway hyper-responsiveness are profoundly inhibited by a helminth (Ascaris suum) extract in a murine model of asthma. Clin. Exp. Allergy 32, 1659–1666. Lopez-Saez, M. P., Zubeldia, J. M., Caloto, M., Olalde, S., Pelta, R., Rubio, M., and Baeza, M. L. (2003). Is Anisakis simplex responsible for chronic urticaria? Allergy Asthma Proc. 24, 339–345. Lynch, N. R., Lopez, R. I., Di Prisco-Fuenmayor, M. C., Hagel, I., Medouze, L., Viana, G., Ortega, C., and Prato, G. (1987). Allergic reactivity and socio-economic level in a tropical environment. Clin. Allergy 17, 199–207. Lynch, N. R., Hagel, I., Perez, M., Di Prisco, M. C., Lopez, R., and Alvarez, N. (1993). Effect of antihelminthic treatment on the allergic reactivity of children in a tropical slum. J. Allergy Clin. Immunol. 92, 404–411. Lynch, N. R., Palenque, M., Hagel, I., and Di Prisco, M. C. (1997). Clinical improvement of asthma after anthelminthic treatment in a tropical situation. Am. J. Respir. Crit. Care Med. 156, 50–54. Maizels, R. M., and Yazdanbakhsh, M. (2003). Immune regulation by helminth parasites: Cellular and molecular mechanisms. Nat. Rev. Immunol. 3, 733–744.
188
Aditya Reddy and Bernard Fried
Maizels, R. M., Balic, A., Gomez-Escobar, N., Nair, M., Taylor, M. D., and Allen, J. E. (2004). Helminth parasites—Masters of regulation. Immunol. Rev. 201, 89–116. Maloy, K. J., and Powrie, F. (2001). Regulatory T cells in the control of immune pathology. Nat. Immunol. 2, 816–822. Martin, E. S. (1976). Increase of mast cells in the peritoneal cavity of rats treated atopic patient’s serum. Allergol. Immunopathol. 4, 131–138. Martin, R. J., Boguniewicz, M., Henson, J. E., Celniker, A. C., Williams, M., Giorno, R. C., and Leung, D. Y. M. (1993). The effects of inhaled interferon-gamma in normal human airways. Am. Rev. Respir. Dis. 148, 1677–1682. Matricardi, P. M., Rosmini, F., Ferrigno, L., Nisini, R., Rapicetta, M., Chionne, P., Stroffolini, T., Pasquini, P., and D’Amelio, R. (1997). Cross-sectional retrospective study of prevalence of atopy among Italian military students with antibodies against hepatitis A virus. Br. Med. J. 314, 999–1003. McSharry, C., Xia, Y., Holland, C. V., and Kennedy, M. W. (1999). Natural immunity to Ascaris lumbricoides associated with immunoglobulin E antibody to ABA-1 allergen and inflammation indicators in children. Infect. Immun. 67, 484–489. Medeiros, M., Jr., Figueiredo, J. P., Almeida, M. C., Matos, M. A., Araujo, M. I., Cruz, A. A., Atta, A. M., Rego, M. A. V., de Jesus, A. R., and Taketomi, E. (2003). Schistosoma mansoni infection is associated with a reduced course of asthma. J. Allergy Clin. Immunol. 111, 947–951. Medeiros, M., Jr., Almeida, M. C., Figueiredo, J. P., Atta, A. M., Mendes, C. M., Arau´jo, M. I., Taketomi, E. A., Terra, S. A., Silva, D. A., and Carvalho, E. M. (2004). Low frequency of positive skin tests in asthmatic patients infected with Schistosoma mansoni exposed to high levels of mite allergens. Pediatr. Allergy Immunol. 15, 142–147. Mingomataj, E.C ¸ ., Xhixha, F., and Gjata, E. (2006). Helminths can protect themselves against rejection inhibiting hostile respiratory allergy symptoms. Allergy 61, 400–406. Morera, P. (1988). Abdominal angiostrongyliasis. A public health problem? Rev. Soc. Bras. Med. Trop. 21, 81–83. Motta, A. C., and van Oosterhout, A. J. M. (2006). T cells in asthma: Lessons from mouse models. Drug Discov. Today Dis. Models 3, 199–204. Muto, R., Imai, S., Tezuka, H., Furuhashi, Y., and Fujita, K. (2001). The biological activity of ABA-1-like protein from Ascaris lumbricoides. J. Med. Dent. Sci. 48, 95–104. Ndiaye, M., and Bousquet, J. (2004). Allergies and parasitoses in sub-Saharan Africa. Clin. Rev. Allergy Immunol. 26, 105–114. Nyan, O. A., Walraven, G. E. L., Banya, W. A. S., Milligan, P., Van Der Sande, M., Ceesay, S. M., Del Prete, G., and McAdam, K. P. (2001). Atopy, intestinal helminth infection and total serum IgE in rural and urban adult Gambian communities. Clin. Exp. Allergy 31, 1672–1678. Obihara, C. C., Beyers, N., Gie, R. P., Hoekstra, M. O., Fincham, J. E., Marais, B. J., Lombard, C. J., Dini, L. A., and Kimpen, J. L. (2006). Respiratory atopic disease, Ascarisimmunoglobulin E and tuberculin testing in urban South African children. Clin. Exp. Allergy 36, 640–648. Ogilvie, B. M., and de Savigny, D. (1982). Immune response to nematodes.. In ‘‘Immunology of Parasitic Infections’’ (S. Cohen and K. S. Warren, eds.), pp. 715–757. Blackwell Scientific Publications, Oxford, England. Ogorodova, L. M., Freidin, M. B., Sazonov, A. E., Fedorova, O. S., Gerbek, I. E., Cherevko, N. A., and Yu Lebedeva, N. (2007). A pilot screening of prevalence of atopic states and opisthorchosis and their relationship in people of Tomsk Oblast. Parasitol. Res. 101, 1165–1168. Palmer, L. J., Celedon, J. C., Weiss, S. T., Wang, B., Fang, Z., and Xu, X. (2002). Ascaris lumbricoides infection is associated with increased risk of childhood asthma and atopy in rural China. Am. J. Respir. Crit. Care Med. 165, 1489–1493.
Atopic Disorders and Parasitic Infections
189
Pascual, C. Y., Crespo, J. F., San Martin, S., Ornia, N., Ortega, T., Caballero, M., Mun˜ozPereira, M., and Martin-Esteban, M. (1997). Cross-reactivity between IgE-binding proteins from Anisakis, German cockroach, and chironomids. Allergy 52, 514–520. Pereira, M. U., Sly, P. D., Pitrez, P. M., Jones, M. H., Escouto, D., Dias, A. C. O., Weiland, S. K., and Stein, R. T. (2007). Nonatopic asthma is associated with helminth infections and bronchiolitis in poor children. Eur. Respir. J. 29, 1154–1160. Perzanowski, M. S., Nganga, L. W., Carter, M. C., Odhiambo, J., Ngan, P., Vaughan, J. W., and Chapman, M. P. (2002). Atopy, asthma, and antibodies to Ascaris among rural and urban children in Kenya. J. Pediatr. 140, 582–588. Pinto, L. A., Dias, A. C. O., Rymer, B. L., Fernandes, F. F., Barbosa, G. L., Jones, M. H., Teixeira, C. G., Stein, R. T., and Pitrez, P. M. C. (2006). Effect of Angiostrongylus costaricensis extract on eosinophilic pulmonary response in BALB/c mice. Parasitol. Res. 98, 295–298. ´ .A. Ponte, E., Lima, F., Araujo, M. I., Maria, I., Oliveira, R. R., Cardoso, L. S., and Cruz, A (2006). Skin test reactivity and Der p1-induced IL-10 production in patients with asthma or rhinitis infected with Ascaris. Ann. Allergy Asthma Immunol. 96, 713–718. Priftanji, A., Strachan, D., Burr, M., Sinamati, J., Shkurti, A., Grabocka, E., Kaur, B., and Fitzpatrick, S. (2001). Asthma and allergy in Albania and the UK. Lancet 358, 1426–1427. Read, A. F., and Skorping, A. (1995). The evolution of tissue migration by parasitic nematode larvae. Parasitology 111, 359–371. Redecke, V., Hacker, H., Datta, S. K., Fermin, A., Pitha, P. M., Broide, D. H., and Razet, E. (2004). Activation of Toll-like receptor 2 induces a Th2 immune response and promotes experimental asthma. J. Immunol. 172, 2729–2743. Reddy, A., and Fried, B. (2007). The use of Trichuris suis and other helminth therapies to treat Crohn’s disease. Parasitol. Res. 100, 921–927. Reece, J. J., Siracusa, M. C., and Scott, A. L. (2006). Innate immune responses to lung-stage helminth infection induce alternatively activated alveolar macrophages. Infect. Immunol. 74, 4970–4981. Romagnani, S. (2004). The increased prevalence of allergy and the hygiene hypothesis: Missing immune deviation, reduced immune suppression, or both? Immunology 112, 352–363. Rzepecka, J., Donskow-Schmelter, K., and Doligalska, M. (2007). Heligmosomoides polygyrus infection down-regulates eotaxin concentration and CCR3 expression on lung eosinophils in murine allergic pulmonary inflammation. Parasite Immunol. 29, 405–413. Sabin, E. A., Araujo, M. I., Carvalho, E. M., and Pearce, E. J. (1996). Impairment of tetanus toxoid-specific Th1-like immune responses in humans infected with Schistosoma mansoni. J. Infect. Dis. 173, 269–272. Sales, V. S., Rodrigues, C. E., Trombone, A. P. F., Lima, R. C., Santos, A. B. R., Oliver, C., Ferriani, V. P. L., Chapman, M. D., and Arruda, L. K. (2002). Infection with Ascaris lumbricoides in preschool children: Role in wheezing and IgE responses to inhalant allergens. J. Allergy Clin. Immunol. 109, S27. Scha¨fer, T., Meyer, T., Ring, J., Wichmann, H.-E., and Heinrich, J. (2005). Worm infestation and the negative association with eczema (atopic/nonatopic) and allergic sensitization. Allergy 60, 1014–1020. Scrivener, S., Yemaneberhan, H., Zebenigus, M., Tilahun, D., Girma, S., Ali, S., McElroy, P., Custovic, A., Woodcock, A., and Pritchard, D. (2001). Independent effects of intestinal parasite infection and domestic allergen exposure on the risk of wheeze in Ethiopia: A nested case-control study. Lancet 358, 1493–1499. Sears, M. R. (1997). Descriptive epidemiology of asthma. Lancet 350, SII1–SII4. Sears, M. R. (1998). Evolution of asthma through childhood. Clin. Exp. Allergy 28, 82–91. Shaheen, S. O., Aaby, P., Hall, A. J., Barker, D. J. P., Heyes, C. B., Shiella, A. W., and Goudiaby, A. (1996). Measles and atopy in Guinea-Bissau. Lancet 347, 1792–1796.
190
Aditya Reddy and Bernard Fried
Shirakawa, T., Enomoto, T., Shimazu, S., and Hopkin, J. M. (1997). The inverse association between tuberculin responses and atopic disorder. Science 275, 77–79. Smits, H. H., and Yazdanbakhsh, M. (2007). Chronic helminth infections modulate allergenspecific immune responses: Protection against development of allergic disorders? Ann. Int. Med. 39, 428–439. Smits, H. H., Hartgers, F. C., and Yazdanbakhsh, M. (2005). Helminth infections: Protection from atopic disorders. Curr. Allergy Asthma Rep. 5, 42–50. Spergel, J. M., and Paller, A. S. (2003). Atopic dermatitis and the atopic march. J. Allergy Clin. Immunol. 112, S118–S127. Turner, J. D., Faulkner, H., Kamgno, J., Kennedy, M. W., Behnke, J., Boussinesq, M., and Bradley, J. E. (2005). Allergen-specific IgE and IgG4 are markers of resistance and susceptibility in a human intestinal nematode infection. Microbes Infect. 7, 990–996. Trujillo-Vargas, C. M., Werner-Klein, M., Wohlleben, G., Polte, T., Hansen, G., Ehlers, S., and Erb, K. J. (2007). Helminth-derived products inhibit the development of allergic responses in mice. Am. J. Respir. Crit. Care Med. 175, 336–344. van den Biggelaar, A. H., van Ree, R., Rodrigues, L. C., Lell, B., Deelder, A., Kremsner, P., and Yazdanbakhsh, M. (2000). Decreased atopy in children infected with: A role for parasite-induced interleukin-10. Lancet 356, 1723–1727. van den Biggelaar, A. H., Lopuhaa, C., van Ree, R., van der Zee, J. S., Jans, J., Hoek, A., Migombet, B., Borrmann, S., Luckner, D., Kremsner, P. G., and Yazdanbakhsh, M. (2001). The prevalence of parasite infestation and house dust mite sensitization in Gabonese schoolchildren. Int. Arch. Allergy Immunol. 126, 231–238. van den Biggelaar, A. H., Boorman, S., Kremsner, P., and Yazdanbakhsh, M. (2002). Immune responses induced by repeated treatment do not result in protective immunity to Schistosoma haematobium: Interleukin (IL)-5 and Il-10 responses. J. Infect. Dis. 186, 1474–1482. van den Biggelaar, A. H., Rodrigues, L. C., van Ree, R., Zee, J. S., van der HoeksmaKruize, Y. C. M., Souverijn, J. H. M., Missinou, M. A., Borrmann, S., Kremsner, P. G., and Yazdanbakhsh, M. (2004). Long-term treatment of intestinal helminths increases mite skin-test reactivity in Gabonese schoolchildren. J. Infect. Dis. 189, 892–900. van der Kleij, D., and Yazdanbakhsh, M. (2003). Control of inflammatory diseases by pathogens: Lipids and the immune system. Eur. J. Immunol. 33, 2953–2963. von Hertzen, L. C., and Haahtela, T. (2004). Asthma and atopy—The price of affluence? Allergy 59, 124–137. Wang, C. C., Nolan, T. J., Schad, G. A., and Abraham, D. (2001). Infection in mice with the helminth Strongyloides stercoralis suppresses pulmonary allergic responses to ovalbumin. Clin. Exp. Allergy 31, 495–503. Weiss, S. (2000). Parasites and asthma/allergy: What is the relationship? J. Allergy Clin. Immunol. 105, 205–210. Williams, H., Robertson, C., Stewart, A., Aı¨t-Khaled, N., Anabwani, G., Anderson, R., Asher, I., Beasley, R., Bjo¨rkste´n, B., Burr, M., Clayton, T., Crane, J., et al. (1999). Worldwide variations in the prevalence of symptoms of atopic eczema in the International Study of Asthma and Allergies in Childhood. J. Allergy Clin. Immunol. 103, 125–138. Wilson, M. S., Taylor, M. D., Balic, A., Finney, C. A., Lamb, J. R., and Maizels, R. M. (2005). Suppression of allergic airway inflammation by helminth-induced regulatory T cells. J. Exp. Med. 202, 1199–1212. Yazdanbakhsh, M., and Wahyuni, S. (2005). The role of helminth infections in protection from atopic disorders. Curr. Opin. Allergy Clin. Immunol. 5, 386–391. Yazdanbakhsh, M., van den Biggelaar, A., and Maizels, R. M. (2001). Th2 responses without atopy: Immunoregulation in chronic helminth infections and reduced allergic disease. Trends Immunol. 22, 372–377. Yazdanbakhsh, M., Kremsner, P. G., and van Ree, R. (2002). Allergy, parasites, and the hygiene hypothesis. Science 296, 490–494.
Atopic Disorders and Parasitic Infections
191
Zaccone, P., Fehervari, Z., Jones, F. M., Sidobre, S., Kronenberg, M., Dunne, D. W., and Cooke, A. (2003). Schistosoma mansoni antigens modulate the activity of the innate immune response and prevent onset of type 1 diabetes. Eur. J. Immunol. 33, 1439–1449. Zaccone, P., Fehervari, Z., Phillips, J. M., Dunne, D. W., and Cooke, A. (2006). Parasitic worms and inflammatory diseases. Parasite Immunol. 28, 515–523.
This page intentionally left blank
CHAPTER
4 Heartworm Disease in Animals and Humans John W. McCall,* Claudio Genchi,† Laura H. Kramer,‡ Jorge Guerrero,} and Luigi Venco}
Contents
1. Introduction (Biology and Life Cycle) 1.1. Development in mosquitoes 1.2. Development in animals 1.3. Experimental infections in animal models 2. Epidemiology in Domestic and Wild Hosts 2.1. D. immitis prevalence and distribution in domestic animals 2.2. Heartworm in wild carnivores 2.3. Molecular surveys of D. immitis in mosquito vectors 3. Pathogenesis, Immunology and Wolbachia Endosymbiosis 3.1. Pathogenesis 3.2. Immunology 3.3. Wolbachia endosymbiosis 4. Canine Heartworm Disease 4.1. Clinical presentation 4.2. Diagnosis 4.3. Chemoprophylaxis 4.4. Therapy against adult worms and microfilariae
195 195 196 197 199 202 208 208 209 209 213 214 222 222 223 227 230
* Department of Infectious Diseases, College of Veterinary Medicine, University of Georgia, Athens, {
{ }
}
Georgia, 30602, USA DIPAV, Sezione di patologia Generale e Parassitologia, Universita` degli Studi di Milano, Via Celoria 10, 20133 Milano, Italy Dipartimento di Produzione Animale, Universita` di Parma, via del Taglio 8, 43100 Parma, Italy Department of Pathobiology, School of Veterinary Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104, USA Clinica Veterinaria Citta` di Pavia, Viale Cremona 179, 27100 Pavia, Italy
Advances in Parasitology, Volume 66 ISSN 0065-308X, DOI: 10.1016/S0065-308X(08)00204-2
#
2008 Elsevier Ltd. All rights reserved.
193
194
John W. McCall et al.
4.5. Surgical extraction of adult worms 5. Feline Heartworm Disease 5.1. Clinical presentation 5.2. Diagnosis 5.3. Chemoprophylaxis 5.4. Symptomatic treatment 5.5. Adulticide therapy 5.6. Surgical extraction of adult worms 6. Heartworm Disease in Ferrets 6.1. Clinical presentation 6.2. Diagnosis 6.3. Chemoprophylaxis 6.4. Therapy against adult heartworms 7. Human Dirofilariosis 7.1. Clinical aspects of human dirofilariasis by D. immitis 7.2. Epidemiology of human D. immitis infection 7.3. Conclusions 8. Emerging Strategies in Heartworm Treatment and Control 8.1. Arsenical therapy 8.2. ‘Safety-Net’ and adulticidal properties of prolonged monthly prophylactic doses of macrocyclic lactones 8.3. Arsenical plus prophylactic doses of macrocyclic lactones for adulticidal therapy 8.4. Anti-filarial effects of tetracyclines and doxycycline plus ivermectin Acknowledgments References
Abstract
232 233 233 239 251 252 253 253 253 253 254 255 256 258 258 260 262 263 263
263 264 264 265 265
Heartworm disease due to Dirofilaria immitis continues to cause severe disease and even death in dogs and other animals in many parts of the world, even though safe, highly effective and convenient preventatives have been available for the past two decades. Moreover, the parasite and vector mosquitoes continue to spread into areas where they have not been reported previously. Heartworm societies have been established in the USA and Japan and the First European Dirofilaria Days (FEDD) Conference was held in Zagreb, Croatia, in February of 2007. These organizations promote awareness, encourage research and provide updated guidelines for the diagnosis, treatment and prevention of heartworm disease. The chapter begins with a review of the biology and life cycle of the parasite. It continues with the prevalence and distribution of the disease in domestic and wild animals, with emphasis on more recent data on the spreading of the disease and the use of
Heartworm Disease in Animals and Humans
195
molecular biology techniques in vector studies. The section on pathogenesis and immunology also includes a discussion of the current knowledge of the potential role of the Wolbachia endosymbiont in inflammatory and immune responses to D. immitis infection, diagnostic use of specific immune responses to the bacteria, immunomodulatory activity and antibiotic treatment of infected animals. Canine, feline and ferret heartworm disease are updated with regard to the clinical presentation, diagnosis, prevention, therapy and management of the disease, with special emphasis on the recently described Heartworm Associated Respiratory Disease (HARD) Syndrome in cats. The section devoted to heartworm infection in humans also includes notes on other epizootic filariae, particularly D. repens in humans in Europe. The chapter concludes with a discussion on emerging strategies in heartworm treatment and control, highlighting the potential role of tetracycline antibiotics in adulticidal therapy.
1. INTRODUCTION (BIOLOGY AND LIFE CYCLE) The domestic dog and some wild canids are the normal definitive hosts for the heartworm Dirofilaria immitis and thus serve as the main reservoir of infection. However, even less suitable hosts such as cats and ferrets occasionally have low-level, transient microfilaraemias and probably serve as a source of infection for mosquitoes during these short periods, as it has been demonstrated that mosquitoes fed on microfilaraemic blood from a cat produced infective, third-stage larvae (L3) that developed to fully reproductive adult worms in a dog (Donahoe, 1975).
1.1. Development in mosquitoes The life cycle of D. immitis is relatively long (usually 7–9 months) compared with most parasitic nematodes. The susceptible mosquito becomes infected when taking a blood meal needed for egg development from a microfilaraemic host. The microfilariae (270–365 mm long and 6–8 mm wide) remain in the mosquito midgut for approximately 24 h before migrating into the large cells of the malpighian tubules. The larvae then become shorter and stouter as they develop into the ‘sausage’ stage. By the fifth day, a gut may be differentiated, consisting of an oesophagus, intestine and rectum. On day 6 or 7, the larvae leave the malpighian tubule cells and enter the lumen of the tubules. The larvae molt to the second stage 8–10 days after infection and again to the third stage 2–3 days later. One to two days later, these L3 perforate the distal ends of the tubules and migrate via the haemocoel (body cavity) to the head and mouthparts where they become infective (1100–1300 mm long) (see Abraham, 1988; Cancrini and Kramer, 2001; Taylor, 1960). The time required for the development of
196
John W. McCall et al.
microfilariae to the infective, third stage is temperature-dependent. At 27 C and 80% relative humidity, development takes about 10–14 days (McCall, 1981; Orihel, 1961).
1.2. Development in animals Infective L3 are transmitted to the definite host while the mosquito is taking a blood meal. During the blood meal, the tip of the labellum (fleshy part of the mouthparts) ruptures and the L3 are deposited on the skin of the host in a drop of the mosquito’s blood (haemolymph). After the mosquito feeds, these sexually differentiated larvae enter the animal’s body via the puncture wound made by the blade-like stylets in the mouthparts (McGreevy et al., 1974). Three days after infection of the dog, most of the larvae are found in the subcutaneous (SC) tissues near their entry site. By day 21, most of them have migrated to the abdomen of the dog, and by day 41, they may be recovered from either the abdomen or the thorax. Worms reach the heart as early as day 70 and all have arrived at the heart by day 90–120. Apparently, L3 and L4 travel between muscle fibres during migration, whereas juveniles (immature adults) penetrate muscle and eventually the jugular, or other veins, directing them towards the heart (Kotani and Powers, 1982; Kume and Itagaki, 1955; Orihel, 1961). The molt from L3 to L4 begins as early as day 3 (Kotani and Powers, 1982; Lichtenfels et al., 1985) and as late as day 9–12 (Orihel, 1961). L4 molt to the final stage at day 50–70; the first worms entering the heart on day 70–85 are 2–4 cm in length. After reaching the heart, the female worms will increase in length by almost tenfold. They become sexually mature about day 120 post-infection. Dogs develop patent infections (i.e., have microfilariae circulating in their blood) as early as 6 months (J. W. McCall, 2007, unpublished data) but usually by 7–9 months post-infection (Kotani and Powers, 1982; Orihel, 1961). Early migration and development of D. immitis in ferrets is similar to that in dogs (Supakorndej et al., 1994) but this has not been studied in cats. When juvenile heartworms first reach the heart and lungs, the pressure of venous blood forces them into the small pulmonary arteries (Rawlings et al., 1978). As they grow and increase in size, they progressively migrate upstream into larger and larger arteries until the worms become fully mature. The eventual location of the mature adult worms appears to depend mainly on the size of the dog and the worm burden. A medium-sized dog (e.g., Beagle) with a low worm burden (i.e., 10) usually has worms mainly in the lobar arteries and main pulmonary artery. As the worm burden increases, worms are also located in the right ventricle. Dogs with more than 40 worms are more likely to have caval syndrome, where most of the worms migrate into the right ventricle, right atrium and the caudal vena cava, thus interfering with valvular function and/or blood flow (Atwell and Buoro, 1988; Ishihara et al., 1978; Jackson, 1975).
Heartworm Disease in Animals and Humans
197
1.3. Experimental infections in animal models Much useful research can be done on animals with naturally acquired infections, and indeed such infections are required for certain studies. However, more specific and definitive results often can be obtained using fewer animals with experimentally induced infections. The nature of the research, required characteristics of the infection in the host, host response and economics usually dictate the specific model selected for a research project. Numerous attempts have been made to establish heartworm infections in various species and strains of mice and rats and jirds (Meriones unquiculatus) and the varying degrees of success with these models has been reviewed by Abraham (1988). More recently, McCall (2001) reviewed the development of the parasite in the intermediate and definitive hosts and described the various procedures now routinely used in the laboratory for experimentally inducing heartworm infections in the most commonly used animal hosts (i.e., dogs, cats and ferrets) and characterized these infections in terms of infection rate, worm burden, microfilaraemia pattern, antigen (and antibody for cats) response and duration of infection in the host. Selected parasitological data on these models are presented in Table 4.1. Essentially all dogs (Dzimianski et al., 1989) and ferrets (Supakorndej et al., 1992) and 60–73% of cats can be infected by SC injection of L3 or the bite of infected mosquitoes (Mansour et al., 1995; McCall et al., 1992). Male cats are more susceptible than female cats in terms of infection rate, the average number of worms per cat, the number of worms in individual cats and the survival rate of infected cats (McCall, 1992), but this sex difference has not been noted in dogs or ferrets and is not evident in all surveys of cats with naturally acquired infections. Intravenous (IV) transplantation of adult heartworms, usually but not always from donor dogs, in dogs and cats produces ‘instant’ infections with known numbers of male and/or female worms of known age in all recipient animals, with predictably high worm survival. This also allows for use of fewer animals to obtain statistically valid data. Also, light, moderate, heavy and single-sex or dual-sex infections can be established to simulate many of the diverse conditions encountered in naturally acquired infections. SC transplantation of juvenile (immature adult) heartworms in dogs, cats and ferrets also provides highly uniform and predictable infections for some studies. Although there are some obvious similarities, there are some other very important differences in infections induced by SC injection of L3 or the bite of infected mosquitoes in dogs, cats and ferrets. As seen in Table 4.1, the average percent recovery of adult worms in dogs (56%) and ferrets (40%, range 34–54%) is considerably higher than in cats (6%). The life span of adult worms in dogs is reported to be 5–7 years (Newton, 1968). As cats are considered to be abnormal hosts for heartworms, it has
TABLE 4.1
Selected parasitological data on some animal modelsa
Host
n
Parasite stage
DOGb
47 39
L3 Adult
18 24
Adult Immature (113–116 days old) L3 Adult
CATc
FERRETc
26 10 7 25 8
10
a b c
Juvenile L3 Immature (97–107 days old) Immature (117–137 days old)
See McCall (2001) for further details and references. Microfilaraemias high and persistent. Microfilaraemias low and transient.
No.
Route
Overall average (%), Recovery (range)
50 5–15 pairs (M&F) 1–3 F; 5–10 M 1–7 F; 13–24 M
SC IV transplantation
56 (26–84) 91 (86–94)
100 100
IV transplantation IV transplantation
100 92 (0–100)
100 96
SC IV transplantation
6 (1–31) 96 (83–100)
73 100
IV transplantation SC SC transplantation
61 (40–100) 40 (7–70) 83 (50–100)
100 100 100
SC transplantation
30 (0–67)
90
100 3–4 pairs (M&F) 7–10 15–60 3 pairs (M&F) 3 pairs (M&F)
Infection rate (%)
Heartworm Disease in Animals and Humans
199
been estimated that the worms live only 2–3 years in this host. This has lead to the proposed assumption that early death of juvenile worms about 4–9 months after infection in cats, but not dogs, is largely responsible for the Heartworm Associated Respiratory Disease (HARD) syndrome often misdiagnosed as feline asthma (Dillon et al., 2007; Nelson et al., 2007). Although this explanation seems plausible, this early death of worms in non-treated cats has not been confirmed with experimentally induced infections. Moreover, it is quite likely that some worms live considerably longer than 2–3 years in cats. There are no reports of the life span of adult worms in ferrets, but considering the relatively high susceptibility and mortality in infected ferrets, it seems reasonable to assume that most infected ferrets do not outlive the parasite. Adult worm size also varies among hosts. In dogs, females measure 250–310 mm long and 1.0–1.3 mm wide and males measure 120–200 mm in length and 0.7–0.9 mm in width (Lok, 1988; Manfredi et al., 2001) and male worms are only about one-fifth the mass of females ( J. W. McCall, 1992, unpublished data). Fully developed worms in cats do not reach the size of those seen in dogs (average length of females in cats, 210 mm) (see McCall et al., 1992). Similarly, heartworms in ferrets are smaller than those of the same age in dogs (see Supakorndej et al., 2004). The size of the worms in the host probably is influenced more by the size of the host than by the rate of development, with smaller hosts having smaller worms (McCall, 1992).
2. EPIDEMIOLOGY IN DOMESTIC AND WILD HOSTS Filarial infections are worldwide diseases caused by different nematode genera and species. Although D. immitis is the most important and a potentially life-threatening species, several filarial worms can infect domestic and wild carnivores. Their importance is mainly due to the need to differentiate circulating microfilariae for a correct diagnosis and treatment of heartworm disease in dogs and cats (see Section 4.2) and to study the risk of zoonotic infections (see Section 7). For this reason, although this section is specifically dedicated to D. immitis epidemiology, some information is given on other species able to infect domestic and wild carnivores. Even though D. immitis occurs worldwide, in North and South America, Acanthocheilonema (previously Dipetalonema) reconditum is present with prevalences up to 22% in Parana´, Brazil (Patton and Faulkner, 1992; Reifur et al., 2004; Theis et al., 2001a), while in Europe, Africa and Middle and Far East, Dirofilaria (Nochtiella) repens, A. (previously Dipetalonema) reconditum, Acantocheilonema (previously Dipetalonema) dracunculoides and Cercopithifilaria (previously Dipetalonema) grassii are not infrequently found in both dogs and cats and wild carnivores (Genchi et al., 2001a,
200
John W. McCall et al.
2007a; Hargis et al., 1999; Schwan and Schro¨ter, 2006) (for further information see Sections 2.2 and 4.2). Both Dirofilaria species are transmitted by different genera and species of mosquitoes (Culicidae) that act as intermediate hosts and vectors, while Acantocheilonema species and C. grassii are transmitted by ticks, lice, fleas and flies (Nelson, 1962; Olmeda-Garcia et al., 1993) (Table 4.2). Although the biology and the ecology of the arthropod vectors are different, some factors such as the abundance of both domestic and wild reservoirs (microfilaraemic hosts), socioeconomical changes that have increased the number of pets and ‘travelling’ dogs (cats are often amicrofilaraemic and are not an efficient source of infection for mosquitoes; McCall et al., 1992) due to the availability of easier and faster means of transportation favour the spreading of dirofilarial infections. However, in a non-arid environment, the main factor is temperature that influences mosquito abundance in the environment and expands the mosquito activity season mainly in areas characterized by a subtropical and temperate climate. For instance, predictive models have demonstrated an actual risk of spreading filarial infections into areas previously unaffected in the northern and southern hemispheres (Genchi et al., 2005; Medlock et al., 2007; Vezzani and Carbajo, 2006) and some empirical findings have already confirmed such a trend (Fok, 2007; Svobodova et al., 2006). Nowadays, Dirofilaria infections are spread from Canada and the northern States of the USA (e.g., Washington) (43 –48 N parallels) (Klotins et al., 2000; Theis et al., 2001a; Zimmerman et al., 1992) to 34 550 S parallel in South America (La Plata, Argentina; Vezzani et al., 2006), although models based on the temperature threshold below which filarial development will not proceed in the mosquito have shown that the risk of infection is extended to the 45 S parallel (Vezzani and Carbajo, 2006). However, it is of interest to note that while the highest endemic area in America is between the 24 N and 34 N parallels, in Europe Dirofilaria infections are mostly confined between the 43.5 N and 45.5 N parallels, demonstrating that although the temperature is critical, many other factors are involved in the epidemiology of the disease that have an affect on geographical distributions. Development of D. immitis to the infective stage (L3) in mosquitoes occurs at a rate that is dependent on ambient temperature, and development may not occur at a threshold temperature below 14 C (Fortin and Slocombe, 1981). Development is assumed to be cumulative and may be calculated in terms of degree-days above the developmental threshold, or heartworm development units (HDU). The model of heartworm seasonality assumes a requirement of 130 heartworm development units ( C) for complete development and a maximum life expectancy of 30 days for common vector mosquitoes (Slocombe et al., 1989). Using this laboratoryderived model that requires numerous inherent assumptions and was
TABLE 4.2
Filarial species infecting dogs and cats
Causative agent
Vector
Dirofilaria immitis
Culicidae
Dirofilaria (Nochtiella) repens Acanthocheilonema (Dipetalonema) reconditum
Culicidae
Acanthocheilonema dracunculoides Cercopithifilaria (Acanthocheilonema) grassii a b c d
Mf: microfilariae. M: male. F: female. Microfilariae in the skin.
Fleas, lice and ticks
Flies and ticks (R. sanguineus) Ticks (R. sanguineus)
Prepatent period
120–180 days 189–259 days 427–476 days
Mfa, in blood
Adults Mb/Fc
Localization of adult worms
290–330 mm
12–18 cm, 25–30 cm 5–7 cm, 10–17 cm 9–17 mm, 21–25 mm
Pulmonary arteries/ right heart Subcutaneous tissue/ muscular fasciae Subcutaneous tissue/ muscular fasciae
15–31 mm, 33–55 mm Unknown, 23–24 mm
Peritoneal cavity
320–370 mm 269–283 mm
?
195–230 mm
?
570 mmd
Subcutaneous tissue/ muscular fasciae
202
John W. McCall et al.
designed to study only the influence of macro-environmental temperature on the heartworm development period, investigators have predicted the seasonal limits of transmission in Canada (Slocombe et al., 1995), the USA (Knight and Lok, 1995) and Europe (Genchi et al., 2005; Medlock et al., 2007) and formulated recommendations for timing of heartworm chemoprophylaxis and scheduling of diagnostic testing. Furthermore, predictive risk maps have been produced coupling this basic model with a Geographic Information Systems (GIS) based on a thermal regime (Genchi et al., 2005; Medlock et al., 2007) and information about mosquito vectors (Vezzani and Carbajo, 2006). While these model-based predictions are academically appealing, they do not yet consider several potentially important factors, such as influence of microclimate and the unique biological habits and adaptations of the numerous mosquito vectors on larval development. More than 70 species of mosquitoes have been shown to be capable of developing microfilariae to the L3, but fewer than a dozen of these species are believed to be major vectors (Otto and Jachowski, 1981). However, the introduction of Aedes (Stegomyia) albopictus in western countries has added one more competent vector (Gratz, 2004; Tatem et al., 2006a,b) to previously described mosquito species for the transmission of heartworm infection (Cancrini et al., 2006; Genchi et al., 1992a). Although the susceptibility of different geographical strains may vary, the likelihood of the presence of at least one susceptible vector species in a geographic area that is conducive to the propagation of mosquitoes is high, and once the ubiquitous heartworm parasite is introduced into an area, its transmission is virtually insured.
2.1. D. immitis prevalence and distribution in domestic animals Although D. immitis has been found in over 30 mammals species, such as domestic and wild canids, domestic and wild felids, mustelids, monkeys, marine mammals, rodents and ungulates (Otto, 1975), dogs are the most frequently infected, with the heaviest worm burden (Genchi et al., 1988) and the most competent reservoirs. In spite of efforts aimed at prevention and control, particularly in dogs, infection appears to be spreading into areas previously considered to be free of the disease (Genchi et al., 2005) and many countries are now endemic for heartworm infection (Genchi et al., 2007). Heartworm prevalence and distribution are better known for dogs. Generally, when dogs are living in high endemic areas, no differences in prevalence are found between sexes, breed, length of hair coat and activity (working dogs vs pet dogs), while the size, the age and the inside/outside habitation are critical, as the risk of infection is significantly higher for large-sized dogs (which are more attractive to mosquitoes), for dogs older than 3 years and for dogs living outdoors (Bolio-Gonzalez et al., 2007;
Heartworm Disease in Animals and Humans
203
Genchi et al., 1988, 1993). In natural infections, the number of adult parasites increases with the dog’s age (up to more than 150 worms per dog in heavily endemic areas) with an overdispersion pattern within the dog population, as most dogs harbour less than 15 adult worms (Genchi et al., 1988). Information on the frequency of diagnosis of infection in cats is gradually becoming available. It is now generally accepted that heartworm disease may occur in cats in any area where dogs are infected (Genchi et al., 1992b, 1998a; Guerrero et al., 1992a; Kramer and Genchi, 2002; McCall et al., 1992) but the geographical distribution and level of infection are less predictable in cats than in dogs. In highly endemic areas, with sufficient rainfall, essentially every unprotected dog becomes infected (McTier et al., 1992a). In contrast to dogs, about 75% of cats can be infected experimentally with D. immitis L3 (McCall et al., 1992). However, the prevalence rate of naturally acquired infections in cats is between 5% and 20% of that for dogs in the same geographical area (Ryan and Newcomb, 1995). Furthermore, in some regions, wild carnivores are infected and act as competent (microfilaemic) reservoirs.
2.1.1. North america Heartworm infection in dogs has now been diagnosed in all of the 50 states of the USA. Although transmission of infection has not been clearly documented for Alaska, the disease is considered to be endemic in all of the remaining 49 states. Heartworm is enzootic along the Atlantic Seaboard and Gulf Coast areas, with the southeastern states generally showing the highest prevalence values. Infection continues to be diagnosed at a high frequency in the Mississippi River basin and in states along the Ohio and Missouri Rivers. New foci have been detected in northern California and Oregon, and autochthonous infections in dogs in the states of Wyoming, Utah, Idaho and Washington have been documented in recent years (Zimmerman et al., 1992). There is a high probability that the introduction of the tree-hole breeding mosquito, Aedes sierrensis, in Salt Lake City, Utah, during the last decade or so is associated with the establishment of enzootic heartworm transmission in the area (Scoles and Dickson, 1995). The tiger mosquito, Aedes albopictus, a known vector of heartworm in Japan, has spread rapidly throughout much of the USA since it was introduced from Asia in 1985. It breeds mainly in piles of discarded tires and is spread within the country by movement of tires from place to place (Tatem et al., 2006a,b). This mosquito readily feeds on dogs and other mammals and is able to host and transmit D. immitis. Two recent surveys conducted by Merial in cooperation with the American Heartworm Society showed that the number of canine heartworm cases diagnosed in the USA were close to a quarter of a million. The first survey conducted in 2002 requesting diagnostic data for 2001 had the
204
John W. McCall et al.
participation of 15,366 clinics that reported diagnosing heartworm infection in 244,291 dogs. The second survey conducted in 2005 reviewing data for 2004 had the participation of 12,173 clinics (out of a total of 25,000) that reported 250,000 cases of canine heartworm infection diagnosed that year (Guerrero et al., 2006). Interestingly, 8800 of the responders in the second survey had also responded in the first survey. Analysing the data obtained in the clinics that participated in both surveys, the number of canine heartworm cases decreased in 17 states, in 3 states the numbers were the same and in 30 states plus Washington, DC, the number of cases increased. Nationally, the reported positive cases of heartworm infection in dogs increased slightly in those clinics reporting in both 2002 and 2005. In a parallel study, the national prevalence of heartworm infection in dogs was performed by evaluating medical records of more than 500 Banfield Pet Hospitals that see approximately 80,000 pets on a weekly basis. Data collected from January 2002 to December 2005 was evaluated. Results of this study showed that 1.46% of the 871,839 dogs examined tested positive for circulating antigen of D. immitis. Based on this data, the estimate of pet dogs in the USA and the proportion of them probably on heartworm prevention, the investigators (Apotheker et al., 2006, unpublished data, personal communication) estimated that 509,932 dogs in the USA had heartworm infection, a figure that is almost exactly double the number of cases reported in the Merial-AHS surveys of 2002 and 2005. Moreover, in the 2005 Merial-AHS survey, close to half of the total number of Companion animal clinics in the USA responded to the survey. It is noteworthy that groups of dogs exported from the New Orleans area and other areas near Louisiana, USA, in the aftermath of Hurricane Katrina in August 2005 to northern states and Canada had very high prevalences of heartworm infection (34–51%) (Levy et al., 2007). Heartworm is endemic in all the states in the USA (Nelson et al., 2005a) and although infected dogs exported from the disaster area are unlikely to have a perceptible impact on the overall prevalence of dirofilariosis, it may increase the risk of establishing new foci at least at the regional level. In Canada, the overall canine prevalence rate is 0.24% (Slocombe, 1992). Prevalence is higher (8.4%) in endemic areas of southwestern Ontario (Klotins et al., 2000). The most significant reports of heartworm infections in British Columbia are from the Okanagan Valley area, which represents a classic instance of recent introduction of the parasite and a resulting local pocket of infection (Zimmerman et al., 1992). Hunters, in previous years, had transported hunting dogs from Texas to this area, setting up new foci of infection. In Mexico, Guerrero et al. (1992b) in a survey involving 2599 dogs from 15 different cities reported an overall prevalence of 7.5%. The highest rates of infection were among dogs from cities located along the coast of the Gulf of Mexico. A more recent study in Meridia, Yucatan, confirmed the
Heartworm Disease in Animals and Humans
205
national prevalence (8.3%) (Bolio-Gonzalez et al., 2007), though in coastal areas the prevalence is higher (19.6%) (Labarthe and Guerrero, 2005).
2.1.2. South america The prevalence and spread of heartworm infection in South America has been recently reviewed by Labarthe and Guerrero (2005) and Vezzani et al. (2006). Surveys indicate that heartworm is endemic in several countries of South America. In Brazil, the overall prevalence of canine heartworm infection in the state of Rio de Janeiro was 21.3% (Guerrero et al., 1992b), with the highest rate for dogs in the northern beaches (49%) followed by dogs from the mountain towns near the cities of Rio de Janeiro (27.4%) and Niteroi (26.4%). The rate in the suburbs of Rio de Janeiro was around 33% (Labarthe et al., 1992). Labarthe et al. (1997a,b) confirmed and extended these findings, and also reported heartworm infection in random-source cats in the city. Alves et al. (1999) found a prevalence rate, as determined by necropsy of dogs in the city of Recife in northeastern Brazil of 2.3%. In the same state (Pernambuco), prevalence in dogs on Itamaraca´ Island was found to be as high as 43%. Labarthe (1997) reviewed the literature on the prevalence in Brazil and found reports of prevalence rates as high as 45% in the state of Sa˜o Paulo. The distribution and prevalence of heartworm disease in Argentina has been recently reviewed by Vezzani et al. (2006). Endemic areas were identified, with the prevalence in dogs ranging from 0% to 23.5% in the Greater Buenos Aires, and depending on environmental and urban characteristics (Rosa et al., 2002), up to 74% in the rural areas of the northeastern province of Formosa (Mancebo et al., 1992). A rate of 10.9% was recorded for Corrientes. More recently, Peteta et al. (1998) reported a prevalence, determined by microfilarial and antigen testing, of 13.6% for dogs in Villa La Na`ta, which is located near the Parana Delta and surrounded by the Lujan River and Villanueva and LaRioja Channels in the Tigre district of the province of Buenos Aires. In Venezuela, examination of canine blood samples submitted to the School of Veterinary Medicine, Central University of Venezuela in Aragua, revealed that 2.3% were positive for microfilariae of D. immitis (Perez and Arlett, 1998). Recent studies performed in Lima, Peru, reported that 4.35% of the blood samples from 140 randomly selected dogs were positive for circulating antigen of D. immitis (Gonzales, 2002).
2.1.3. Central america and the caribbean
Kozek et al. (1995) reviewed the prevalence of canine filariae in the Caribbean islands and conducted a thorough epidemiological survey in Puerto Rico. Prevalence values for heartworm infection in Puerto Rico ranged from 3.1% to 20.4%, with the highest rate recorded for the city of Ponce on the southern coast. In Cuba, prevalence for Havana ranged from
206
John W. McCall et al.
7% to 19% and from 37% to 63% on the Isla de Juventud. For Curaca˜o, it ranged from 9% to 11% and was 53% for the Grand Bahamas and 18% for the Dominican Republic.
2.1.4. Australia Heartworm infection is enzootic along the northern coastal areas of Western Australia and the eastern states of Queensland, New South Wales and Victoria, where prevalence rates generally mimic those for the southeastern states of the USA, with north to south prevalences reversed for the southern hemisphere. The prevalence of heartworm infection in dogs in Sydney was as high as 30% in the late 1980s, and cats were also found to be infected (Kendall et al., 1991). More recently, the rate for dogs in this area was reported to be only 11.4% (Bidgood and Collins, 1996).
2.1.5. Asia and the south pacific Heartworm disease is well established in most of the Islands of the Pacific and in many countries of Asia but survey results are not readily available for every country. A prevalence of 11.3% for dogs from the Fars province of Iran has been reported ( Jafari et al., 1996). Heartworm is enzootic on the islands of Japan, where the disease is well known and prevalence is well documented. A survey conducted on stray dogs and cats in the Kanto region of Japan in 1985 revealed a prevalence rate of 59% for dogs and 2% for cats (Tanaka et al., 1985). More recently, Roncalli et al. (1998) reported that prevalence rates for feline heartworm infection in Japan range from 0.5% to 9.5% in stray cats and from 3.0% to 5.2% in house cats. An antigentest survey of German shepherds in five areas of South Korea revealed an overall prevalence of 28.3% (Lee et al., 1996). Prevalence was highest in Hoengsong-gun (84.4%), while Yechon-gun and Chungwon-gun areas had rates of 20.0% and 14.3%, respectively. None of the dogs in the Kimhae-shi and Kwanju areas was positive. More recently, prevalences rates of 48% in outdoor dogs and 2.6% in stray cats were reported by Lee (2003) and Liu et al. (2005). Kuo et al. (1995) reported a 53.8% prevalence rate for dogs in the Taipei province of Taiwan, and Wu and Fan (2003) recorded an overall prevalence of 57% in stray dogs in Taiwan.
2.1.6. Europe The prevalence and spread of heartworm infection in Europe has been comprehensively reviewed by Genchi et al. (2005). The disease is diagnosed mainly in the southern European countries of Italy, Spain, Portugal and France, with scattered reports from Greece, Turkey and some Eastern European countries such as Croatia, Serbia, Bulgaria, Romania and the Czech Republic (prevalence ranging from 2% to 48%; FEDD, 2007), but not in Hungary where until now only D. repens has been found in dogs (Fok, 2007). An increasing number of cases are now being diagnosed in
Heartworm Disease in Animals and Humans
207
northern countries such as Austria, Germany and The Netherlands in dogs that were either imported from the Mediterranean area or had accompanied their owners to the area. One possible exception is a heartworm-positive dog from the Canton of Tessin (Switzerland), which appears to have acquired an autochthonous infection. The area of highest prevalence values for dogs and cats is along the Po River Valley in northern Italy, from where the first observation of the worm was made at necropsy of an hunting dog in 1626 (Birago, 1626). The prevalence rate for cats in this area is high (up to 21%), and the rate for dogs not treated with preventive drugs ranges from 35% to 80%. The disease has recently spread northward into the provinces of Friuli-Venezia-Giulia and in central areas of the peninsula, though no infection or very low prevalence has been found in southern areas where temperatures are more favourable for larval development in mosquitoes. For instance, D. immitis prevalence (microfilaraemic dogs) is 0.01% in Sicily (Giannetto et al., 1997, 2007) and 0.6% in Campania (Cringoli et al., 2001), two regions located between the 40.5 N and 37 N parallels. However, the spread of Aedes albopictus in Italy and the evidence that this mosquito species can act as a natural vector for D. immitis could enhance the risk of transmission from animals to humans, considering the aggressive anthropophilic behaviour of the species (30–48 bites/h) (Cancrini et al., 2003). The highest rates for dogs in Spain are reported for the southern provinces of Huelva (37%), Cadiz (12%) and Badajoz (8%) (Guerrero et al., 1992b). During the last few years, D. immitis infection appears to be spreading into other regions of Catalun˜a. A recent survey of Barcelona showed that 12.8% of the dogs were infected. The Canary Islands of Tenerife (21%; Montoya et al., 2006) and Las Palmas (36.0%) are highly endemic and about 59% of the dogs on Gran Canaria Island are infected with heartworms (Montoya et al., 1998). In Portugal, infection is diagnosed mainly in the southern regions, with prevalence values ranging from 12% (Algarve) to 30% (Island of Madeira). Although limited survey data are available, prevalence values for dogs range from 2% to 17% for Slovenia, Bulgaria, Greece and Turkey and up to 65% for Romania, and some of these areas are considered to be endemic. In Croatia, canine heartworm infection has been reported from the Istrian peninsula (about 16% prevalence) (Zˇivicˇnjak et al., 2007).
2.1.7. Africa Heartworm is found in dogs from various regions of Africa, but no information is available regarding infections in cats. Infection in dogs appears to be common throughout western Africa and in eastern parts, extending from the Republic of South Africa and Mozambique (Schwan and Durand, 2002) to the Republic of Sudan. Matola (1991) reported a prevalence of 10.2% for dogs in Tanzania.
208
John W. McCall et al.
2.2. Heartworm in wild carnivores D. immitis infections has been reported in wild felids, such as the ocelot (Leopardus pardalis), mountain lion (Felis concolor), clouded leopard (Neofelis neburosa), snow leopard (Uncia uncia), Bengal tiger (Panthera tigris), lion (Panthera leo) and black bear (Ursus americanus) in their natural habitat or caged in urban zoos and free ranging in ‘safari’ parks worldwide. However, wild felids and ursids are infrequently microfilaemic (Crum et al., 1978; Kennedy and Patton, 1981; Murata et al., 2003; Okada et al., 1983; Paul-Murphy et al., 1994; Pence et al., 2003; Ruiz de Yba´n˜e´z et al., 2006) and are not thought to be reservoirs of the infection. As with domestic cats, generally they are not involved in the transmission of this parasite. This is the case of animals born in urban zoos located in heavily endemic areas, showing that heartworm transmission is active also in heavily urbanized areas (C. Genchi, 1982, unpublished data). In wild canids (Canis lupus, C. latrans, C. aureus, Vulpes vulpes, Urocyon spp. and Nyctereutes procionides), D. immitis has been frequently observed. In the USA, 21–42% of coyotes (C. latrans), depending on the capture area (Nakagaki et al., 2000; Nelson et al., 2003; Pappas and Lunzman, 1985; Sacks and Caswell-Chen, 2003) and 58–100% of Island foxes from California Channel Island (Roemer et al., 2000) were found infected. The demographic data suggests that heartworm is not a major factor influencing the population dynamics. In Europe, heartworm prevalence in red foxes (V. vulpes) from Spain, Italy and Bulgaria ranges from 0.4% to12% (C. Genchi, 2005, unpublished data; Gorta´zar et al., 1994, 1998; Magi et al., 2007; Man˜as et al., 2005; Segovia et al., 2001) and is 2.1% in the wolf (C. lupus) (Segovia et al., 2001). In Bulgaria, heartworm infection was also found in jackals (C. aureus) with a prevalence of 8.9% (Kirkova et al., 2007). Information on microfilaraemia is generally scant in such hosts because most of the surveys were carried out serologically or post-mortem. In general, foxes harbour few parasites, often of the same sex, and the risk of their acting as a competent reservoir of infection is low. On the contrary, in California, USA, coyotes have heavy worm burdens and microfilariae in their peripheral blood and can serve as an active reservoir (Garcia and Voigt, 1989).
2.3. Molecular surveys of D. immitis in mosquito vectors The development of molecular methods to identify filarial larvae (Favia et al., 1996, 2000) has allowed a more accurate identification of their mosquito vectors, opening the way for some molecular surveys of D. immitis in their vectors. Recently, wild Culex pipiens, Anopheles albopictus, A. maculipennis sensu lato and Coquillettidia richiardii have been found to be able to transmit D. immitis in Italy (Cancrini et al., 2003, 2006); Aedes aegyptii and C. pipiens in Argentina (Vezzani et al., 2006) and Anopheles sinensis sensu
Heartworm Disease in Animals and Humans
209
lato, A. sineroides, Aedes vexans nipponii, Armigeres subalbatus and C. pipiens in Korea (Lee et al., 2007). Such results will contribute to the improvement of risk maps, which until now have relied on a thermal regimen.
3. PATHOGENESIS, IMMUNOLOGY AND WOLBACHIA ENDOSYMBIOSIS Canine and feline heartworm disease is characterized by both acute and chronic inflammatory lesions in the lungs and other organs because of the presence of adult worms and circulating microfilariae. Furthermore, mild-to-severe side effects are an inevitable consequence of successful adulticide therapy. D. immitis, the causative agent of canine and feline heartworm disease, harbours intracellular bacteria named Wolbachia pipientis. Indeed, most filarial species studied, with very few exceptions, contain these micro-organisms that are thought to play an essential role in the biology and reproductive functions of their filarial hosts. Sironi et al. (1995) demonstrated that D. immitis harbours Wolbachia. Indeed, as a Gram-negative bacteria, Wolbachia have the potential to play an important role in the pathogenesis and immunoresponse to filarial infection. The immunopathology of filarial disease is extremely complex and the clinical manifestations of infection are strongly dependent on the type of immune response elicited by the parasite. Furthermore, the fact that adult parasites can survive for years in otherwise immunocompetent hosts is likely due to the parasite’s ability to avoid/modulate the immune response. It is therefore extremely important to identify which components of the parasite interact with the host’s immune system, including Wolbachia. This section will briefly review the pathogenesis and immune response during heartworm disease and will then illustrate what is currently known about the relationship between Wolbachia and filarial worms, the interaction between Wolbachia and dogs and cats infected with D. immitis, and how the understanding of what Wolbachia actually does to/for the worm may aid researchers in finding interesting new targets for control and prevention of heartworm disease.
3.1. Pathogenesis The pathophysiological response to heartworm infection is mainly due to the presence of adult worms of D. immitis in the pulmonary arteries. The primary lesions in this disease occur in the pulmonary arteries and lung parenchyma. They are mostly attributable to the intravascular adult parasites; they cause pulmonary hypertension, that if not treated progresses inevitably to congestive heart failure (CHF). Other syndromes are related to the disturbance of blood flow due to the location of heartworms in the
210
John W. McCall et al.
right atrium at the level of the tricuspidal valve. This event causes massive haemolysis and related haemoglobinuria, responsible for the vena caval syndrome (Ishihara et al., 1978; Kitagawa et al., 1987). Microfilariae appear to play a relatively minor pathogenic role but may cause clinically significant pneumonitis and glomerulonephritis. Some individuals develop a hypersensitivity to microfilariae which then disappear from the blood. Occasionally, aberrant migration results in parasites becoming trapped in ectopic locations, such as the anterior chamber of the eye (Weiner et al., 1980) or systemic arteries (Liu et al., 1966; Slonka et al., 1977). Data regarding immunopathogenesis and in particular the role of cytokines, pro-inflammatory mediators as well as cellular components of the immune system in the development of heartworm-related lesions has been recently reviewed (Grandi et al., 2005). Although the spectrum of pathologies related to chronic heartworm disease is broad, the most important clinical manifestation in dogs is CHF (cor pulmonale; a change in structure and function of the right ventricle of the heart as a result of a respiratory disorder). Heartworms are, despite what their physical presence could suggest, primary agents of vascular disease, rather than simply the cause of obstruction and/or blood flow disturbances. Intimal proliferation occurs in arteries occupied by living worms and embolic worm fragments trigger thrombosis, both of which may completely obstruct segments of the pulmonary arteries (Fig. 4.1). These effects, both leading to pulmonary hypertension, are
FIGURE 4.1 Lung from a dog with heartworm infection. Note the obstruction of vascular lumen due to endothelial proliferation and worm fragments (hematoxylin/ eosin, 40).
Heartworm Disease in Animals and Humans
211
strongly correlated to worm burden, which in turn is related to the degree of their distribution through lung parenchyma (Knight, 1987). The juvenile heartworms are only about 2.5 cm long when they reach the systemic venous circulation. They passively embolize the pulmonary arteries and are disbursed in proportion to the lobar blood flow. Generally, the larger and more readily accessible right caudal lobar artery accumulates more worms than the left (Atwell and Rezakhani, 1986a). Contact between the parasite and the intima of the pulmonary arteries is an important, if not essential, initial step in the development of the endovascular lesions. The earliest lesions are limited to the small peripheral branches where the worms first come to rest. As the parasite grows, lesions occur in more proximal segments. Intimal thickening and narrowing of the vessel lumen in small peripheral branches of the pulmonary arteries are the major cause of obstructed blood flow and pulmonary hypertension. The intimal proliferation is caused by migration of medial smooth muscle cells through the internal elastic laminae (Munnel et al., 1980; Schaub et al., 1981). The pathogenesis of the arteritis caused by heartworms remains a matter of speculation. Disruption of the endothelial cell junction and denuding of the intimal surface are characteristics of the first lesions that occur only a few days after the worms occupy the vessels. The evidence suggests that injury to the endothelium occurs immediately upon arrival of the parasite, too soon for the components of an acquired immune response to fall into place without prior sensitization. Furthermore, the disappearance of endothelial cells occurs without evidence of degeneration and is followed by an aggressive buildup of cells and structural elements. This suggests that cells have been dislodged rather than destroyed in situ. Macrophages, granulocytes and platelets are attracted to the site of endothelial damage and adhere to the exposed subendothelium. Shortly after their arrival, vascular smooth muscle cells migrate into the intima and a very active process of myointimal proliferation produces rapid growth of the lesions. The prominence of platelets in the acute lesions and their documented ability to stimulate growth of the vascular smooth muscle, through the release of platelet-derived growth factors (Ross, 1986), are hypothesized to be a likely mechanism for triggering and sustaining the growth of these lesions (Schaub and Rawlings, 1980; Schaub et al., 1981). Although the lesions thicken the wall of these large elastic vessels and produce a rough texture on the intimal surface, they do not obstruct blood flow by narrowing the lumen. On the contrary, the large distributing arteries actually dilate as pulmonary hypertension becomes increasingly severe. Pulmonary blood flow is impeded primarily by the reduction in cross-sectional area of the arterial vascular bed, caused by obliterative endarteritis of small peripheral branches. Recently, in heartworm infected dogs, it has been demonstrated that there is a markedly
212
John W. McCall et al.
increased plasma level of endothelin-1, a mediator that induces acute vasoconstriction and chronic vascular remodelling. It is probable that both of these events contribute in turn to the development of pulmonary hypertension (Uchide and Saida, 2005). Thrombosis and thromboembolism may compromise the pulmonary circulation further. As worms accumulate, lesions also develop in the large distributing arteries, which dilate and become stiffer, and the pulmonary blood pressure rises. The decreased distension of the large vessels significantly increases cardiac work by coupling the right ventricle directly to the high vascular resistance in the obstructed peripheral vasculature. Right ventricular hypertrophy is a compensatory response to the increased pressure load. As heartworm disease impedes flow in an increasing number of branches, the pulmonary vascular reserve diminishes. For a time, normal pulmonary blood pressure is preserved at rest and rises only modestly during exercise as patent arteries reach full distension. Eventually, the pulmonary arterial tree is restricted to the point that it assumes the characteristics of a system of rigid tubes and pulmonary vascular resistance becomes fixed. At this stage, pressure rises in direct proportion to further increase in flow. Consequently, the more severe the disease and active the patient, the more cardiac work must be performed. In advanced cases of heartworm disease, low-output CHF develops as a result of the right ventricle’s inability to generate and sustain the high perfusion pressures required to move blood through the lung. Recently, a decrease in extracellular collagen matrix has been observed in the myocardium of heartworm-infected dogs that may contribute to ventricle dilatation, thereby markedly affecting the systolic and diastolic functions of the heart (Wang et al., 2005). Frequently, dogs at this stage experience syncope (a temporary loss of consciousness and posture) when attempting to suddenly increase cardiac output. Right-sided congestive heart failure (R-CHF) with ascites, hepatomegaly and cachexia is a late sequela and may be precipitated by an acute episode of pulmonary thromboembolism (Fig. 4.1).
3.1.1. Pathogenesis of feline heartworm disease Although live adult worms in the pulmonary arteries cause a local arteritis, some cats never manifest clinical signs. When signs are evident, they usually develop during two stages of the disease: (1) arrival of juvenile heartworms in the pulmonary vasculature and (2) death of adult heartworms. The first stage coincides with the arrival of immature adult worms in the pulmonary arteries and arterioles approximately 3–4 months postinfection. These early signs are due to an acute vascular and parenchymal inflammatory response to the newly arriving worms and the subsequent death of some, if not most, of these same worms. This initial phase is often misdiagnosed as asthma or allergic bronchitis but in actuality is part of a syndrome now known as Heartworm Associated Respiratory Disease
Heartworm Disease in Animals and Humans
213
(HARD, see www.heartwormsociety.org). Clinical signs associated with this acute phase subside as the worms mature but demonstrable histopathological lesions are evident even in those cats that clear the infection. The most notable microscopic lesion is occlusive medial hypertrophy of the small pulmonary arterioles, but other changes are also noted in the bronchi, bronchioles, alveoli and pulmonary arteries. Browne et al. (2005) recently reported that cats with serological evidence of exposure to heartworms, including those without adult heartworms in the lungs and heart, have a greater prevalence of pulmonary arterial lesions than heartwormnegative cats without serological evidence of exposure. The cat’s response to the parasite is intense and the role of inflammation is very important (Furlanello et al., 1998) following the death of adult worms. The cause of the acute and often fatal crisis in the cat is lung injury resulting in respiratory distress. Frequently, this event is associated with the death of as few as one adult heartworm. The lung can become acutely oedematous and respiratory failure, rather than heart failure, becomes the life-threatening event. Because pulmonary hypertension is only an occasional event, R-CHF and severe cor pulmonale is uncommon in feline heartworm infection (Dillon, 1999; Genchi et al., 1995).
3.2. Immunology There are several features of heartworm disease that suggest that a protective immune response eventually develops against one or more developmental stages of D. immitis. First, following experimental infection, not all inoculated infective larvae are able to complete migration to the lungs and become adults (Yoshida et al., 1997). This would imply that the host immune system is able to eliminate migrating larvae and attempts to immunize dogs with irradiated L3 or with chemically abbreviated infections have met with variable success (Grieve et al., 1988; Mejia and Carlow, 1994; Yoshida et al., 1997). Mejia and Carlow (1994) reported that vaccinated dogs preferentially recognized several larval (14, 20, 30, 34, 39 kDa), adult worm (20 kDa) and microfilarial (36, 38, 71, 84 kDa) antigens. Frank and Grieve (1996) further identified and characterized two proteins that appear to be larval specific (L3–L4 molt) and, more importantly, are specifically recognized by sera from dogs immunized by chemically abbreviating their infections. The implications for vaccine development are obvious, but still in experimental phases. The identification of immunodominant antigens also has diagnostic potential. Synthetic peptides based on the sequences of Frank and Grieve (1996) have been used more recently to study antibody responses in experimentally and naturally infected cats (Prieto et al., 2002) with promising results. Another feature of heartworm disease is the so-called ‘occult infection’, defined as the presence of adult worms in the lung/heart, but with the
214
John W. McCall et al.
absence of circulating microfilariae. There are several, well-defined types of occult infection, including pre-patent infections, infections of only one sex, amicrofilaraemic infections induced by drugs and immune-mediated occult infections. Immune-mediated occult infections are due to a specific, humoral response towards microfilarial antigens (Rawlings et al., 1982). Light and transmission electron microscopy of lungs from dogs with immune-mediated occult infections have shown that microfilariae undergo massive attack by eosinophils, neutrophils and lymphocytes, suggesting that cell-mediated hypersensitivity may also play a role in eliminating microfilariae. More recent in vitro studies have shown that microfilarial agglutination, a feature of in vitro exposure of microfilariae to sera from dogs with immune-mediated occult infection, depends on the formation of immune complexes between anti-microfilarial antibodies and escretory/secretory products (Hayasaki, 2001). As with other pathogenic filariae, the immunology of D. immitis infection is complex and there is still much to be done to shed light on the interaction between parasite and host. Interestingly, heartworm infection may behave in much the same way as human filarial infections. Recently, Morcho´n et al. (2007a) reported that dogs with circulating microfilariae have higher expression of circulating interlukin (IL)-4, IL-10 and inducible nitric oxide synthetase (iNOS) mRNA than those with occult infection, suggesting that the presence of circulating microfilariae is associated with immune tolerance towards infection. In human lymphatic filariasis (Brugia malayi and Wuchereria bancrofti), both microfilariae and adults are long-lived and studies suggest that the immune response elicited by each stage probably has its own distinct features (O’Connor et al., 2003). In particular, the role of microfilariae in the immunology of these infections has been widely studied. Microfilariaemic individuals tend to show immunosuppression or ‘unresponsiveness’ to parasite antigens, as reflected by the inability of peripheral blood mononuclear cells to secrete interferon-g (Luder et al., 1996, Sartono et al., 1999). Recent studies of the cytokine milieu in microfilaria-positive infections have shown that several cytokines predominate: namely, the immunosuppressive IL-10 and the ‘Th2-type’ IL-4 [see O’Connor et al. (2003) for review].
3.3. Wolbachia endosymbiosis Following the discovery of the endosymbiont bacteria Wolbachia in pathogenic filarial worms, not only has its biological role in the reproduction and survival of the nematode host been the subject of intense research, but the possible role of contact between the filarial infected host and these Gram-negative bacteria in the pathogenesis of disease is also being widely studied [see Taylor et al. (2005a) for review].
Heartworm Disease in Animals and Humans
215
W. pipientis, the only species thus far identified in the genus, are Gramnegative bacteria belonging to the order Rickettsiales. They are closely related to other bacteria belonging to the same group, such as Ehrlichia spp. and Anaplasma spp. (Bandi et al., 2001). Electron microscopy, histology and immunohistochemistry have offered a clear description of the distribution of Wolbachia in D. immitis (Bandi et al., 1999; Kozek, 2005; Kramer et al., 2003; Sacchi et al., 2002). They are found throughout all the stages of the life cycle of the nematode although they occur in varying proportions between individual worms and different developmental stages. In adult D. immitis, Wolbachia is predominantly found throughout the hypodermal cells of the lateral cords (Fig. 4.2). In females, Wolbachia is also present in the ovaries, oocytes and developing embryonic stages within the uteri (Fig. 4.3). They have not been demonstrated in the male reproductive system (Sacchi et al., 2002), suggesting that the bacterium is maternally transmitted through the cytoplasm of the egg and not through the sperm. All current evidence suggests that Wolbachia is a symbiont in filarial worms (see Taylor et al., 2005a), that is, the presence of the bacteria is essential for the filarial worm’s survival. The phenomenon of bacterial endosymbiosis is well known in arthropods, but less so in nematodes. There are, however, several features of the relationship between Wolbachia and filarial worms (including D. immitis) that suggest its symbiotic nature: (1) in those species of filarial worms that have been identified as harbouring Wolbachia, all of the individuals are infected (100% prevalence); (2) the
FIGURE 4.2 Anti-Wolbachia surface protein (WSP) immunohistochemistry. D. immitis female with positive staining within a lateral hypodermal chord. (ABC-HRP, 100).
216
John W. McCall et al.
FIGURE 4.3 Anti-Wolbachia immunohistochemistry. D. immitis female with positive-staining microfilariae within the uterus. (ABC/HRP, 100).
evolution of the bacteria match that of the filarial worms, as shown by phylogenic studies; (3) the bacteria are transmitted from female to offspring, thus the symbiont guarantees its own future by increasing the fitness of the host that is involved in its transmission and (4) removal of Wolbachia (antibiotics/radiation) leads to sterility of females and eventual death of adults. It is still unclear, however, exactly what Wolbachia does to make it so important for its filarial host. Several hypotheses have been suggested. Studies on population dynamics in B. malayi (Fenn and Blaxter, 2004; McGarry et al., 2004) have shown that the numbers of bacteria remain stable in microfilariae and in the larval stages L2 and L3 within the mosquito. After approximately a week following infection of the mammalian host, bacteria numbers increase rapidly throughout L4 development. This means that the major period of bacterial population growth occurs within the first month of infection of the definitive host, suggesting a role in evasion of mammalian immunity, the long-term survival of adult worms and possibly in molting. Interestingly, Morcho´n et al. (2004) have recently reported that the development of a strong anti-Wolbachia antibody response against experimental infection of cats with D. immitis occurs after 1–2 months of infection: Is it possible that this intense immune response to Wolbachia is characteristic of resistance to infection in this host? Perhaps the most important recent discovery concerning Wolbachia in filarial nematodes is the completion of the entire genome sequencing and
Heartworm Disease in Animals and Humans
217
annotation of the metabolic pathways of the Wolbachia from B. malayi (Comb et al., 2005; Foster et al., 2005). This will certainly lead to a clearer understanding of the endosymbiotic relationship. To summarize, Wolbachia is essential for its filarial host, including D. immitis. Evolutionary studies confirm that filarial worms have been harbouring Wolbachia for tens of millions of years. During this time, the bacteria have learned to supply needed metabolites to the worm during molting and embryonal development and the worm, in turn, has guaranteed the bacteria’s survival and transmission. It is indeed a ‘one hand washing the other’ situation that may, however, be the key to novel strategies for the control/treatment of filarial infection, including heartworm disease in canines, felines and ferrets.
3.3.1. Wolbachia-derived molecules as pathogen-associated molecular patterns
In hosts that are infected with filarial nematodes harbouring Wolbachia, the bacteria are released following worm death through larval molt, natural attrition, microfilarial turnover and pharmacological intervention (Taylor et al., 2001; Fig. 4.4). In human and murine models of infection, the release of bacteria has been shown to be associated with the up-regulation of pro-inflammatory cytokines, neutrophil recruitment and an increase in specific immunoglobulins. Thus, the role of Wolbachia in the host response to filarial infection may include interaction between bacterial molecules and the innate and adaptive immune system. The innate immune system
FIGURE 4.4 Anti-Wolbachia immunohistochemistry. Kidney from a dog with heartworm infection. Note positive staining for the WSP within microfilariae (ABC/HRP, 60).
218
John W. McCall et al.
represents a defence mechanism against molecular structures that are conserved among a wide range of organisms. It consists of the recognition of specific ‘markers’ (pathogen-associated molecular patterns, PAMPs) that signal the presence of ‘generic’ pathogens. The consequential recognition of these PAMPs by Toll-like receptors (TLRs) on the surface of antigen-presenting cells leads to the production of reactive oxygen species and pro-inflammatory cytokines and to the up-regulation of co-stimulatory molecules that assist in the development of an adaptive immune response. Potential PAMP candidates for Wolbachia include the Wolbachia surface protein (WSP) and GroEL. Reports of a possible role for Wolbachia in the immunopathogenesis of filarial infection have come from both in vitro and in vivo studies. Studies on the human filarial nematodes Onchocerca volvulus and B. malayi/W. bancrofti have demonstrated that (1) adverse reactions to filaricidal therapy (ivermectin, DEC) are associated with the presence of Wolbachia and/or its DNA in the bloodstream and peak levels of Wolbachia correlate with levels of proinflammatory cytokines such as TNFa (Cross et al., 2001); (2) O. volvulusinduced skin nodules feature neutrophil infiltration around adults and microfilariae; this inflammation subsequently subsides following antibiotic-mediated removal of Wolbachia (Brattig et al., 2001). Interestingly, a major surface protein of Wolbachia from D. immitis has been shown to provoke chemiokinesis and IL-8 production in canine neutrophils in vitro (Bazzocchi et al., 2003); (3) filarial worm extracts stimulate cells in vitro to produce pro-inflammatory cytokines in a TLR-dependent manner and this effect is abolished with antibiotic-mediated removal of Wolbachia (Brattig et al., 2004). Furthermore, this effect is not present with extracts of filarial worms that do not harbour Wolbachia; (4) chronic pathology in lymphatic filariasis (elephantiasis, hydrocoele) is correlated with a strong specific humoral response to the WSP (Hise et al., 2004). The hypothesis that D. immitis-infected dogs also come into contact with Wolbachia either through microfilarial turnover or natural death of adult worms has been tested recently (Kramer et al., 2005). In this study, intense staining for the WSP was observed in various tissues from dogs who had died from natural heartworm disease. Bacteria were observed in the lungs and particularly in organs such as the kidney and liver, where microfilariae normally circulate (Fig. 4.4). Interestingly, immunocomplex glomerulonephritis is a frequent complication of heartworm disease and the localization of WSP in glomeruli is suggestive of a role for Wolbachia in renal pathology. It has been reported that infection in dogs with Ehrlichia canis, a bacteria closely related to Wolbachia, features immune-complex formation that may be responsible for renal lesions. Furthermore, when specific antibody responses to Wolbachia were examined, a stronger humoral response in dogs with circulating microfilariae compared with dogs with occult infection was observed, supporting the hypothesis that
Heartworm Disease in Animals and Humans
219
microfilarial turnover is an important source of Wolbachia in dogs with heartworm disease. Interestingly, studies in naturally infected dogs indicate that the presence of both pro-inflammatory mediators (IL-2, iNOS) and the immunomodulatory cytokine IL-10 are characteristic of heartworm disease in dogs. However, as cited above, dogs with patent infection (i.e., circulating microfilariae) have higher expression of circulating IL-4, IL-10 and iNOS mRNA than those with occult infection (Morcho´n et al., 2007a). Is it possible that the fine balance of inflammatory pathology/ long-term survival of adult worms may in some way be dependent on continuous exposure to Wolbachia? Brattig et al. (2004) reported that blood cells from patients with O. volvulus, when incubated in vitro with the WSP, produced high levels of IL-10 and the authors suggest that Wolbachia may contribute to the down-regulation of pro-inflammatory mediators, thus establishing the necessary homeostasis for chronic infection. O’Connor et al. (2003) reviewed the role of NO in filarial disease, reporting evidence that several filarial antigens (microfilarial extracts, filarial cystatins) are capable of inducing NO production in vivo and in vitro. The authors suggest that this, in turn, may induce peripheral tolerance through NO-mediated apoptosis of antigen-specific T lymphocytes. They also cite the potential immunoregulatory influence of Wolbachia in NO production during filarial infection. Bazzocchi et al. (2007) have recently reported that D. immitis Wolbachia-derived WSP prevents apoptosis in human neutrophils, further complicating the possible role of this endosymbiont during filarial infection. Interaction between Wolbachia and the humoral immune system has also been reported by several authors in different hosts infected with different species of filariae, including dogs and cats with D. immitis (Bazzocchi et al., 2000; Brattig et al., 2004; Kramer et al., 2005; Morcho´n et al., 2004; Punkosdy et al., 2003; Simo´n et al., 2003). Naturally infected cats produce antibodies that recognize WSP in Western blot analysis. In a more recent study, the antibody response against specific molecules of D. immitis and Wolbachia endosymbionts in both naturally and experimentally infected cats with and without preventive (ivermectin) treatment was evaluated. Increased antibody production against filarial antigens and WSP was observed in experimentally infected cats without treatment. However, in experimentally infected cats treated with a preventive drug, there was a transient increase in anti-D. immitis IgG that decreased dramatically in association with the death of the larvae, while the anti-WSP IgG response increased constantly until the end of the experiment (6 months). The immune response to Wolbachia antigens was detected as early as 2 months after infection, before detection of specific antibodies against D. immitis antigens. These findings suggest that Wolbachia also plays an important role in the immune response to heartworm infection in cats that may also have diagnostic value (Morcho´n et al., 2004).
220
John W. McCall et al.
Specific immune responses against WSP have also been studied in dogs with naturally acquired heartworm infection. As mentioned above, higher anti-WSP total IgG titres were observed in dogs with circulating microfilariae (mfþ) compared with dogs with occult infection (mf). There was also a predominance of IgG2 antibodies, indicating a bias towards cell-mediated immunity against Wolbachia (Kramer et al., 2005). Perhaps, one of the most interesting results seen so far with infection by D. immitis concerns human dirofilariaisis. Simo´n et al. (2003) reported specific humoral recognition of WSP in patients with pulmonary nodules due to D. immitis and have suggested the use of this antibody response in the differential diagnosis of the disease. Little data is currently available for the potential pro-inflammatory/ immunomodulatory effect of GroEL/HSP60 of Wolbachia. The protein from the Wolbachia of D. immitis has been produced in recombinant form and used in inoculation trials in mice. When inoculated alone, Wolbachia GroEl/ HSP60 does not appear to stimulate pro-inflammatory responses; however, when inoculated in combination with WSP, there is a stronger innate inflammatory response compared with WSP alone (Morcho´n et al., 2007b). Recently, it was reported that Wolbachia-derived HSP60 from W. bancrofti generates antibody responses in infected or exposed individuals and an elevated IgG and IgG1 reactivity is observed in people with filarial pathology, suggesting that another Wolbachia-associated molecule may indeed play a role in filarial disease (Suba et al., 2007). In conclusion, there is increasing evidence that Wolbachia participates in the inflammatory and immune response to D. immitis infection. Areas of future research should include the possible diagnostic use of specific immune responses to Wolabchia, its potential immunomodulatory activity (prevention) and the effects of antibiotic treatment in infected animals.
3.3.2. What are the effects of antibiotic treatment on D. immitis?
Wolbachia can be eliminated from filarial worms through antibiotic therapy of the infected host. Numerous studies on several filarial worms have shown that various treatment protocols/dosages (tetracycline and synthetic derivatives appear to be the most effective) are able to drastically reduce, if not completely remove, the endosymbiont from the worm host. Such depletion of Wolbachia is then followed by clear anti-filarial effects, including inhibition of larval development, female worm sterility and adulticide effects (Gilbert et al., 2005; Langworthy et al., 2000; Taylor et al., 2005b). We know from Bandi et al. (1999) that doxycycline treatment of naturally infected dogs drastically reduces Wolbachia loads in D. immitis. The effects of different treatment regimens in dogs experimentally infected with adult D. immitis on microfilariaemia, antigenemia, worm vitality and
Heartworm Disease in Animals and Humans
221
Wolbachia content has been evaluated recently. Long-term, intermittent doxycyline therapy resulted in a slow yet constant decrease in circulating microfilariae. Furthermore, quantitative polymerase chain reaction (PCR) analysis of Wolbachia/nematode DNA ratios showed a significant decrease in Wolbachia content in worms recovered from treated dogs that was not, however, associated with worm death (C. Bazzocchi et al., in press). If we consider Wolbachia as a potential cause of inflammation in the course of filarial disease, depletion of the bacteria may still be beneficial, independently of its effect on the worm. Indeed, preliminary results from our study indicate that efficient depletion of Wolbachia from D. immitis can alleviate pulmonary pathology following specific adulticide therapy (L. M. Kramer, 2007, unpublished data). Given the recent and very promising developments in the use of tetracyclines for micro-macrofilaricidal therapy in human filarial infections, it is hoped that similar attention will be given to canine, feline and ferret heartworm disease, all of which could greatly benefit from alternative therapeutic strategies.
3.3.3. Immunization The first successful attempt to protect dogs from heartworm infection was carried out by Wong (1974) more than 30 years ago. In that study, D. immitis-irradiated larvae were used to immunize eight dogs, three of which were completely protected from challenge infection. Since this study, several experiments have been carried out demonstrating varying levels of protection using irradiated larvae (Mejia and Carlow, 1994) or chemically abbreviated infections in dogs (Grieve et al., 1988; Yoshida et al., 1997). Overall, the efficacy of such studies did not exceed 70%. In experimental models, studies carried out to identify protective antigens focused mainly on surface antigens, excretory/secretory, molecules that may be produced to counteract the host immune response and antigens recognised by sera from naturally infected dogs or putatively immune individuals (Abraham and Grieve, 1991; Grieve et al., 1992; Mejia and Carlow, 1994), but the level of protection was not sufficient to develop an effective vaccine. In the same way, D. immitis gut-associated antigens were found to react only with sera from heavily- or long-term, infected dogs and were not recognized by ‘normally’ infected hosts (McGonigle et al., 2001), even if partial protection was observed (reduction of 24.5–61% of live worms). Frank et al. (1998a) compared antigen-driven proliferative responses and cytokine production profiles in popliteal lymph node cells collected from both a heartworm-naive dog and an immune dog that had previously received six chemically abbreviated D. immitis infections. Stimulation indices after 4 days in culture indicated that cells collected
222
John W. McCall et al.
from a putatively immune dog proliferated more vigorously than did cells collected from a naive dog in response to soluble or either excretory/secretory larval antigens or to bvDi20/22 (Frank and Grieve, 1996; Frank et al., 1996, 1998a). In cats, the use of bvDi20/22 antigen for 6 days was found to stimulate peripheral blood mononuclear cells in vitro (Frank et al., 1998b). In spite of these results and the number of natural and recombinant molecules studied until now as potential vaccine candidates, including proteins associated with the timing of the moult from L3 to L4 (Frank and Grieve, 1991), the results remain quite poor when compared with the efficacy observed by Wong (1974).
4. CANINE HEARTWORM DISEASE 4.1. Clinical presentation Heartworm infection is a severe and potentially fatal disease caused mainly by the adult stages of D. immitis. Despite the name ‘heartworm’, which suggests primarily cardiac involvement, worms are located mainly in the pulmonary arteries and initial damage is to the lung. Heartworm disease should be considered a pulmonary disease that involves the right cardiac chambers only in the last stages of the disease and in infections with heavy worm burdens. The pathological mechanisms that are responsible for clinical disease are reviewed in Section 3. The clinical presentation of canine heartworm disease is usually chronic. Most infected dogs do not show any symptoms for months or years, depending on the worm burden, individual reactivity and exercise, as arterial damage is more severe in dogs with intensive exercise than in dogs at rest (Dillon et al., 1995a). Signs of the disease develop gradually and may begin with a chronic cough and coughing may be followed by moderate to severe dyspnoea, weakness and sometimes lipothymia after exercise or excitement. At this time, abnormal pulmonary sounds (crackles) over the caudal lung lobes and splitting of the second heart sound can often be heard. Later, when R-CHF is developing, swelling of the abdomen and sometimes legs from fluid accumulation, anorexia, weight loss and dehydration are usually noted. At this stage, cardiac murmurs over the right side of the thorax due to tricuspid valve insufficiency and abnormal cardiac rhythm due to atrial fibrillation are common findings. Sudden death rarely occurs and dogs usually die following respiratory distress or cachexia. Occasionally, acute episodes can also be observed during the chronic course of the disease. After severe, spontaneous thromboembolism due to the natural death of many worms, dogs may show acute life-threatening
Heartworm Disease in Animals and Humans
223
dyspnoea and haemoptysis. Furthermore, in small-sized dogs, the displacement of adult worms from the pulmonary arteries to the right cardiac chambers due to pulmonary hypertension and the sudden fall in right cardiac output is a common event. In this case, affected dogs present the so-called ‘caval syndrome’. Caval syndrome is characterized by acute haemodynamic changes and presents as an acute clinical syndrome. Dyspnoea, tricuspidal cardiac murmur, acute intravascular haemolysis and haemoglobinuria, which is considered pathognomonic for caval syndrome, are the most typical signs and without immediate surgery to remove the worms, a fatal outcome is usual (Atwell and Buoro, 1988; Kitagawa et al., 1986, 1987; Venco, 1993). Because erythrocytes from dogs with caval syndrome have increased osmotic and mechanical fragility and poikilocytosis, haemolysis is mainly a consequence of the mechanical shearing and physical collision of erythrocytes with heartworms (Atwell and Buoro, 1988; Rawlings, 1986).
4.2. Diagnosis Adult heartworm infection in dogs can be diagnosed with blood tests that detect circulating microfilariae or adult antigens, but further diagnostic procedures are usually required to determine the severity of disease and treatment options (Knight, 1995). Microfilariae must be morphologically differentiated from those of other filarial species that release microfilariae into the blood (see below and Table 4.3), especially in European and far East countries where several filarial species can infect dogs (see Section 2). Microfilariae of these genera are found occasionally during blood examinations in dogs, and less frequently in cats, with no clinical signs of disease. Adult worms of these species are localized in SC tissues. However, it has been reported that dogs with D. repens can present cutaneous disorders of varying severity, such as pruritus, dermal swelling and subcutaneous nodules containing the parasites (Beneth et al., 2002; Bredal et al., 1998; Hargis et al., 1999). Severe infections with allergic reactions likely due to microfilarial sensitization have also been reported (Hargis et al., 1999; Mandelli and Mantovani, 1966; Restani et al., 1962). Microfilariae can be differentiated through morphology, histochemical staining or molecular methods.
4.2.1. Blood test for microfilariae Blood samples are examined for the presence of microfilariae after conÒ centration [modified Knott test (Knott, 1939) or filtration (Difil Test)]. If microfilariae are seen and identified as D. immitis based on morphology (Table 4.3), this is considered definitive proof of infection (specificity 100%). However, up to 30% of infected dogs do not have circulating microfilariae even though they harbour adult worms. This may be due
224
John W. McCall et al.
TABLE 4.3 Morphological features of microfilariaea from the blood of dogs infected with filariae
a b
Species
Length (mm)
Width (mm)
Dirofilaria immitis
290–330
5–7
Dirofilaria repens
300–360
6–8
Acanthocheilonema reconditum
260–283
4
Acantocheilonema dracunculoidesb
190–247
4–6.5
Features
No sheath, cephalic end pointed, tail straight with the end pointed No sheath, cephalic end obtuse, tail sharp and filiform often ending as an umbrella handle No sheath, cephalic end obtuse with a prominent cephalic hook, tail button hooked and curved Sheath, cephalic end obtuse, caudal end sharp and extended
Ò
Microfilariae measured by Knott test; when measured by Difil Test lengths are shorter. Microfilariae from the uterus.
to the age of adult worms (young adults in pre-patent infections or older female worms with decreased fecundity) or to the host immune response against adult nematodes or microfilariae. Other factors that can affect the presence of microfilariae in the blood stream are the administration of microfilaricidal drugs or macrocyclic lactones at off-label dosages or, infrequently in dogs, the presence of only worms of the same sex. Therefore, the sensitivity of testing for microfilariae is not sufficient to rule out infection in the case of a negative result. It is noteworthy that the intensity of microfilaraemia is not correlated with the adult worm burden: in general, highly microfilaraemic dogs harbour only a few worms.
4.2.2. Histochemical stains Histochemical staining for microfilariae has been extensively described by Chalifoux and Hunt (1971) and more recently by Periba`n˜ez et al. (2001). D. immitis microfilariae show two acid phosphatase activity spots located around the anal and the excretory pores, respectively; D. repens microfilariae show only one acid phosphatase activity spot located around the anal pore; A. dracunculoides microfilariae show three acid phosphatase
Heartworm Disease in Animals and Humans
225
activity spots localized around the anal, excretory and medium poles. A. reconditum microfilariae show acid phosphatase activity throughout the body.
4.2.3. Blood test for adult female circulating antigens Tests designed to detect adult heartworm antigens based on ELISA and immunochromatography/lateral flow staining techniques are considered highly specific, as cross-reactivity with other canine parasites (i.e., D. repens, Acanthocheilonema spp.) does not occur. These tests allow for detection of adult heartworm antigens produced only by female worms and may provide information about worm burden (Knight, 1995; Venco et al., 2004). The sensitivity is actually very high, but false negative results may occur in prepatent (infections of less than 5 months duration) or very light infections or when only male worms are present (McCall, 1992). They are currently available as in-clinic tests as well as through many veterinary reference laboratories.
4.2.4. Molecular methods PCR is a sensitive and accurate tool to discriminate microfilariae from the different filarial worms that infect dogs. Several PCR methods using different primers have been developed (Casiraghi et al., 2006; Favia et al., 1996; Mar et al., 2002; Rishniw et al., 2006). Its use is advisable in case of morphological abnormalities of microfilariae that is not infrequent in dogs treated incorrectly with preventive drugs or when multiple infections with more than one species of filarial worm makes morphological differentiation difficult. Furthermore, PCR is a useful tool for the molecular survey of filarial infections in mosquito vectors (Cancrini et al., 2006; Sang-Eun et al., 2007) and for identifying filarial larvae in mosquitoes (Cancrini et al., 2003).
4.2.5. Radiography Thoracic radiographs may show, in the advanced stage of disease, enlargement of pulmonary arteries, abnormal pulmonary patterns and, in severe cases, right-sided cardiomegaly. If R-CHF is present, peritoneal and pleural effusion can be noted (Calvert and Rawlings, 1988; Rawlings, 1986). Radiography is useful to assess the severity of the pulmonary lesions but not for evaluating the worm burden (Venco et al., 2004). Because radiographic signs of advanced pulmonary vascular disease may persist long after the infection has run its course, some of the most severely diseased dogs may have disproportionately light worm burdens (Fig. 4.5). On the contrary, some inactive dogs may have heavy worm burdens and be clinically asymptomatic with no or negligible radiographic lesions (Fig. 4.6).
226
John W. McCall et al.
FIGURE 4.5 Thoracic radiograph showing dramatic enlargement of pulmonary arteries, right-sided cardiomegaly and diffuse bronchointerstitial pattern in a 10-year-old heartworm infected dog with right-sided congestive heart failure. Only two adult worms were found at necropsy.
FIGURE 4.6 Thoracic radiograph showing a focal patchy bronchointerstitial pattern (white arrow) in a 4-year-old dog harbouring a heavy worm burden; 38 adult worms were surgically removed from this dog via a jugular vein. Pulmonary arteries and cardiac silhouette appear normal.
Heartworm Disease in Animals and Humans
227
4.2.6. Electrocardiography As electrocardiograms display the electrical activity of the heart, abnormalities (electrical axis right deviation, atrial fibrillation) are usually found only in the last stage of the disease, when right cardiac chambers are severely damaged.
4.2.7. Echocardiography Echocardiography allows a direct visualization of cardiac chambers and associated vessels (Moise, 1988). It also allows the visualization of parasites in the right cardiac chambers, caudal vena cava, main pulmonary artery and the proximal tract of both caudal pulmonary arteries (see Section 5.2). Live adult heartworms are visualized as short, double, linear parallel objects floating in the right cardiac chambers or within the lumen of vessels (Badertscher et al., 1988; Moise, 1988). Echocardiography is performed in dogs mainly in cases where clinical and radiographic findings suggest severe disease. Cardiac ultrasound can increase the accuracy in staging the disease and estimating the worm burden, both of which affect the treatment programme and the prognosis (Venco et al., 2003).
4.3. Chemoprophylaxis Canine heartworm infection can be effectively prevented by treating dogs with drugs. These drugs fall into two basic classes, the piperazine derivative diethylcarbamazine citrate (DEC) and macrocyclic lactones or macrolides (avermectins and milbemycins). The required daily administration of DEC is inconvenient, more than one missed dose can result in a breakdown in protection and the overall prevention programme is difficult to manage. Furthermore, some dogs develop diarrhoea and vomiting while on the drug, which may necessitate discontinuation. More important, DEC administration can cause severe anaphylactoid reactions in microfilaraemic dogs (Plumb, 1995). The use of DEC is fairly limited in the USA and Canada, and the drug is no longer available in Europe. Chemoprophylaxis against D. immitis with macrocyclic lactones is based on the property of these compounds to kill the third and fourth larval stages of Dirofilaria worms. Thus, although the terms ‘prophylaxis’ or ‘prevention’ are currently used, the administration of these drugs actually interrupts the development of larvae transmitted by mosquitoes to the animal during the previous 30–60 days. Therefore, monthly oral administration throughout the transmission season of ivermectin at 6 mcg/kg, milbemycin oxime at 500 mcg/kg or moxidectin at 3 mcg/kg provides effective protection against heartworm infection in dogs (Table 4.4). Selamectin is also 100% effective in preventing the development of heartworm infection in dogs when administered monthly at
TABLE 4.4 Minimum effective dosages of macrocyclic lactones for the prevention of Dirofilaria immitis in dogs and cats and indications against other parasites ML
Presentation
Species
ML dose
Indications
Minimum age for treatment
IVM IVM/PYR
Tablets/chewables Chewables Chewables Flavour tablets Tablets Tablets Tablets Tablets Injectable Topical Tablets Topical Topical
Dog Cat Dog Dog Dog Cat Dog Dog Dog Dog Cat Dog Cat
6 mcg/kg 24 mcg/kg 6 mcg/kg 0.5 mg/kg 0.5 mg/kg 2 mg/kg 0.5 mg/kg 3 mcg/kg 0.17 mg/kg 2.5 mg/kg 1 mg/kg 6 mg/kg 6 mg/kg
Di, Dr Di, At, Ab Di, Dr, Tc, Tl, Ac, Us Di, Tc, Tl, Ac, Tv Di, Tc, Tl, Ac, Tv, Dc, Tae, Eg, Ms Di, At, Tc, Dc, Tae, Em Di, Tc, Ac, Tv, Cf Di Di, Dr, Ac, Us Di, Tc, Ac, Tv, Cf, Oc, Ss, Dx Di, Tc, At, Cf, Oc Di, Dr, Tc, Cf, Ss, Oc, Trc Di, Tct, At, Cf, Oc, Fs
6 weeks 6 weeks 6 weeks 2 weeks/0.5 kg 2 weeks 6 weeks/0.5 kg 2 weeks 6 weeks 12 weeks 7 weeks/1 kg 9 weeks 6 weeks 6 weeks
MBO MBO/PZQ MBO/LFN MOX MOX/IMCP SLM
ML, macrocyclic lactone; IVM, ivermectin; PYR, pyrantel pamoate; MBO, milbemycine oxime; PZQ, praziquantel; LFN, lufenuron; MOX, moxidectin; IMCP, imidacloprid; SLM, selamectin. Di, Dirofilaria immitis; Dr, D. repens; Ac, Ancylostoma caninum; At, A. tubeforme; Ab, A. braziliense, Tc, Toxocara canis; Tct, T. cati; Us, Uncinaria stenocephala; Tv, Trichuris vulpis; Dc, Dipylidium caninum; Ms, Mesocestoides sp; Tae, Taenia spp, Eg, Echnococcus granulosus; Em, Echinococcus multilocularis; Cf, Ctenocephalides felis; Trc, Trichodectes canis; Fs, Felicola subrostrata; Oc, Otodectes cynotis; Ss, Sarcoptes scabiei; Dx: Demodex canis.
Heartworm Disease in Animals and Humans
229
6 mg/kg and given throughout the transmission season. There are several important characteristics unique to selamectin: the drug is given as a topical formulation, thereby avoiding problems associated with oral administration. At the dosage recommended for heartworm prevention in dogs (and cats), selamectin is also effective in preventing and controlling flea infestations (Ctenocephalides felis) and for treating and controlling ear mites (Otodectes cynotis), biting lice (Trichodectes canis), sarcoptic mange (Sarcoptes scabiei), hookworm (Ancylostoma caninum) and roundworm (Toxocara canis) infection. Milbemycin oxime at the recommended dose for heartworm prophylaxis is also effective against Toxocara canis, T. leonina, A. caninum and Trichuris vulpis [for a review see Guerrero et al. (2002)]. Furthermore, several combinations of macrolides with other active compounds against endo- and ectoparasites are also available that include treatment and control of certain gastrointestinal and external parasites (Table 4.4). Ivermectin, selamectin and moxidectin at approved doses for heartworm prevention in dogs have also been found to be effective in preventing D. repens infection (Genchi et al., 2002a; Marconcini et al., 1993; Rossi et al., 2002, 2004). All of these macrolide drugs have a wide range of efficacy that allows them to be administered every 30 days. Treatment with any one of them should begin within a month after the beginning of the transmission season and the final dose should be administered within 1 month after the end of mosquito activity. At the present time, however, the most accepted recommendation is to treat year-round in endemic areas (Nelson et al., 2005a). This provides a safeguard in the case of omission or delay of a monthly treatment, or when the chemoprophylactic history cannot be verified. Ivermectin, milbemycin oxime and selamectin have both been found to provide a high degree of protection when administered on a regular basis, beginning 3 months after infection. Furthermore, monthly treatment with ivermectin over a 1-year period has been shown to be more than 95% effective in preventing development of D. immitis larvae that were 4 months old; however, under the same conditions, milbemycin oxime was only 41.5–49.3% effective as a clinical prophylactic agent (McCall, 2005). This retroactive or ‘reachback’ effect has not been reported for moxidectin, although products with this compound do have a label claim for efficacy of 2 months duration. This so-called ‘safety net’ or ‘reachback’ effect of macrocyclic lactones is very useful to compensate for missed or delayed treatments (e.g., no information about prevention in dogs with possible prepatent infection coming from/living in endemic areas), but should not be considered as justification to modify the recommended monthly interval for treatment (McCall, 2005). In heartwormendemic areas, puppies that are born during the transmission season should be given their first dose of macrocyclic lactones between 2–8 weeks of age.
230
John W. McCall et al.
Injectable formulations of moxidectin to be administered solely by licensed veterinarians are available in Italy, Spain and Australia for prevention of heartworm infection in dogs. The commercial formulations (moxidectin sustained release injectable for dogs) have been approved for use in dogs 12 weeks of age and older. It is able to protect dogs for an entire heartworm transmission season (Genchi et al., 2002b; Lok et al., 2005). For dogs over 6 months of age, pre-treatment testing utilizing both an antigen test and Knott test for circulating microfilariae to verify the absence of D. immitis or D. repens infections is compulsory in dogs. Retesting should always be carried out after the first season of preventive treatment and must include testing for circulating adult antigens as well as for microfilariae. The reason for this is that regular chemoprophylactic use of the macrolides will usually clear any microfilariae from the blood, so that even if an infection has become patent during therapy, it might not be detected by screening for microfilariae. However, in some cases, dogs can be microfilaraemic, mainly when one or more treatments are inadvertently omitted towards the end of the transmission season. Testing should be performed at a minimum of 6 months after the last administration of one of the macrolide compounds. It is also advisable to repeat testing at the beginning of each transmission season before the start of treatment in order to exclude the possibility of infection due to poor owner compliance during the preceding season, or to verify that there was no pre-existing infection.
4.4. Therapy against adult worms and microfilariae 4.4.1. Pre-treatment assessment of the patient It is generally agreed that the treatment of heartworm infection is problematic and there are several approaches that can be used, including the option of not treating at all. In any case, it is important to understand that treating for heartworm infection is neither simple nor safe in itself. Prior to therapy, heartworm-infected dogs must be assessed and rated for risk of post-adulticide thromboembolism. Previously, four classes were described in order to develop the correct prognosis (from classes one to four; Di Sacco and Vezzoni, 1992), but currently a more simple classification is preferred that classifies the patient into one of two categories (low and high risk). Important factors include how many worms are thought to be present based on antigen testing and ultrasound examination (Venco et al., 2004), the size of the dog, the age of the dog (dogs ranging from 5 to 7 years are at high risk of harbouring the heaviest worm burden; Venco et al., 2004), concurrent health factors, severity of pulmonary disease and the degree to which exercise can be restricted during the recovery period.
Heartworm Disease in Animals and Humans
231
The categories into which patients are grouped are as follows: 1. Low risk of thromboembolic complications: light worm burden and no parenchymal and/or pulmonary vascular lesions. Dogs included in this group must satisfy all of the following conditions: a. no symptoms, b. normal thoracic radiographs, c. low level of circulating antigens or a negative antigen test with circulating microfilariae, d. no worms visualized by echocardiography, e. no concurrent diseases and f. possibility of restricting exercise (owner compliance). 2. High risk of thromboembolic complications: in this group, all dogs that show one or more of these conditions should be included: a. symptoms related to the disease (coughing, lipothymia, swelling of the abdomen), b. abnormal thoracic radiographs, c. high level of circulating antigens, d. worms visualized by echocardiography, e. concurrent diseases and f. no possibility of restricting exercise (lack of owner compliance).
4.4.2. Anti-inflammatory and antithrombotic therapy Symptomatic therapy includes drugs and measures that can improve cardiopulmonary circulation and lung inflation in order to relieve symptoms in dogs that cannot undergo causal therapy or to prepare them for an adulticide or surgical therapy. Restriction of exercise and, in selected cases, cage rest seem to be the most important measures to improve cardiopulmonary circulation and reduce pulmonary hypertension (Dillon et al., 1995a). Anti-inflammatory doses of glucocorticosteroids (prednisolone, two mg/kg s.i.d. for 4 or 5 days) given at a diminishing rate can control pulmonary inflammation and thromboembolism. Diuretics (furosemide, one mg/kg b.i.d.) can reduce fluid effusion when R-CHF is present. Digoxin may be administered but only to control atrial fibrillation. The use of aspirin is debatable and no sure evidence of a beneficial antithrombotic effect has been reported. The empiric use of aspirin is therefore not advised (Knight, 1995). In selected cases of respiratory distress, oppioids and oxygen may be used to treat life-threatening situations.
4.4.3. Adulticidal and microfilaricidal treatment The organic arsenical compound melarsomine dihydrochloride is the only approved and effective heartworm adulticide. Two intramuscular injections of 2.5 mg/kg b.w. 24 h apart is the standard regimen. However, a more gradual two-phase treatment regimen is strongly advised to reduce
232
John W. McCall et al.
the risk of pulmonary thromboembolism. This protocol includes a single injection of melarsomine followed by two injections 24 h apart after an interval of at least 30 days (Keister et al., 1992; Vezzoni et al., 1992). In fact, one administration of melarsomine at the dose of 2.5 mg/kg b.w. kills about 90% of the male worms and 10% of the female worms, resulting therefore in a 50% reduction of the total worm burden (which is safer in terms of embolism and shock). Because it is safer and more efficacious, the three-injection alternative protocol is the treatment of choice of the American Heartworm Society (2005) and several university teaching hospitals, regardless of the stage of disease or of the risk category (Nelson et al., 2005a). Pulmonary thromboembolism is an inevitable consequence of successful adulticide therapy. If several worms die, widespread pulmonary thrombosis frequently develops. Mild thromboembolism may be clinically inapparent, but in severe cases life-threatening respiratory distress can occur. These complications can be alleviated by restricting exercise (no walks, no running around; the dog must stay indoors and, in selected cases, be confined to cage rest) during the 30–40 days following treatment. Calcium heparin and anti-inflammatory doses of glucocorticosteroid should be given to control clinical signs of thromboembolism (Di Sacco and Vezzoni, 1992; Vezzoni et al., 1992). It is now known that certain macrolides have adulticidal properties (McCall, 2005; McCall et al., 2001a). Experimental studies have shown that ivermectin has partial adulticidal properties when used continuously for 16 months or more at preventive doses (6–12 mcg/kg/month) and greater than 95% adulticidal efficacy if administered continuously for about 30 months (McCall et al., 2001a). While there may be a role for this therapeutic strategy in a few selected cases (e.g., client-owned dogs in which age or concurrent medical problems advise against using melarsomine), the current recommendations are that this extra-label use of ivermectin should not to be adopted as the primary adulticidal approach, and that this kind of therapy should be used carefully (Nelson et al., 2005a). In fact, the adulticidal effect of ivermectin generally requires a prolonged period before heartworms are completely eliminated. Furthermore, the older the worms are when first exposed to ivermectin, the longer they take to die. In the meantime, the infection persists and continues to cause disease (Rawlings et al., 2001). Clinical observations suggest that some heartworm-positive, active dogs under long-term ivermectin treatment worsen if ivermectin is given monthly for 2 years (Venco et al., 2004).
4.5. Surgical extraction of adult worms Surgical therapy is advised when worms have become displaced in the right cardiac chambers and/or the caudal vena cava, causing the sudden onset of severe symptoms (caval syndrome). Surgical extraction is carried
Heartworm Disease in Animals and Humans
233
out under general anaesthesia with Flexible Alligator Forceps (Fuji Photo Optical LTD, Japan) introduced via the jugular vein. The caudal vena cava, right cardiac chambers and the main pulmonary artery and lobar branches can be accessed with flexible alligator forceps, aided by fluoroscopic guidance (Ishihara et al., 1990). Intra-operative mortality with this technique is very low. Overall survival and rate of recovery of dogs at high risk of pulmonary thromboembolism is improved significantly by physically removing as many worms as possible. When the facilities are available, worm extraction is the procedure of choice for the most heavily infected and high-risk dogs. Before electing this method of treatment, echocardiographic visualization of the pulmonary arteries should be performed to determine that a sufficient number of worms are in accessible locations. Surgical removal of heartworms avoids pulmonary thromboembolism, as compared to pharmacological adulticide drugs such as melarsomine (Morini et al., 1998). This procedure, however, requires specialized training and instrumentation, including fluoroscopic imaging capabilities. Nevertheless, it remains a very good and a safe alternative for the management of high-risk patients and the best choice in dogs harbouring a heavy worm burden.
5. FELINE HEARTWORM DISEASE 5.1. Clinical presentation Heartworm infection in cats is basically pulmonary in nature. Infected cats can be asymptomatic carriers of the parasite, or can present nonspecific clinical signs or clinical signs suggestive of a respiratory affliction. Clinically, the infection can be asymptomatic, chronic or acute. The most frequent clinical signs are of respiratory or digestive origin. Among the respiratory signs, the most frequent is dyspnoea, although coughing, tachypnea and, less frequently, sneezing can be seen. Sometimes it is also possible to detect an increase in respiratory sounds (Atkins et al., 1998a). Vomiting not associated with eating is the most frequent gastrointestinal sign, but other symptoms such as diarrhoea are occasionally described. (Dillon et al., 1996, 1997a,b). Some affected cats also present prostrated, anorectic and showing weight loss (Atkins et al., 1998a). In those very rare cases where worms are located in the right heart, a systolic cardiac murmur due to tricuspid valve insufficiency and galloping cardiac rhythm are common findings at auscultation (Atkins et al., 1998a). Neurological signs such as ataxia, syncope, blindness or vestibular alterations can be seen depending on the ectopic localization of the migrating worms (Atkins et al., 1998a; Dillon et al., 1996, 1997a,b, 1998; McCall et al., 1994). Abdominal oedema, pneumothorax or chylothorax
234
John W. McCall et al.
have also been described although rarely observed in the clinic (Atkins et al., 1998b; Dillon et al., 1997b; Glaus et al., 1995; Treadwell et al., 1998). Clinical signs are usually more evident in two stages of the cat infection: when young adult parasites arrive in the pulmonary arteries (4–6 months after infection) and die, and at a later time when adult D. immitis die (Atkins et al., 1998a; Dillon et al., 1995b). Feline heartworm disease is now recognized as a significant pulmonary syndrome defined as Heartworm Associated Respiratory Disease (HARD, which is discussed in Section 3 of this chapter. Clinical signs associated with HARD are described in Table 4.5. Results of a study by Dillon et al. (2007) demonstrated that arterial disease occurs, along with airway disease, even in cats in which no adult parasites developed. In this study, 60 laboratory-born and -raised cats were each injected SC with 100 D. immitis L3 and allocated to three groups of 20 cats that were given topical selamectin (6 mg/kg) monthly to prevent infection, oral ivermectin (150 mcg/kg) every 2 weeks starting at 84 days PI to simulate early death of immature worms in the pulmonary arteries or served as non-treated controls, respectively. Half of the cats were necropsied at 8 months PI; the remaining cats are scheduled for necropsy at 16 months PI. No worms were recovered from the 10 selamectin-treated cats, only one ivermectin-treated cat had worms and was excluded, and 9 of the 10 control cats had adult heartworms. At approximately 4, 6 and 9 months PI, control and ivermectin cats had abnormal radiographic interstitial and bronchial scores compared to selamectin cats. At necropsy, lungs were grossly abnormal in control and ivermectin cats. Lung histopathological scores for the control and ivermectin cats were significantly different (p < 0.05) from selamectin cats for pulmonary arterioles (Figs. 4.7–4.9), capillaries, bronchioles (Figs. 4.10–4.12) and alveoli (Figs. 4.13–4.15), and lesions were more severe in the arterial segments of control and ivermectin cats (Figs. 4.16 and 4.17). Ivermectin cats had only immature heartworms, but significant parenchymal and airway
TABLE 4.5 Signs described in cats with Heartworm Associated Respiratory Disease (HARD)
Anorexia Difficulty breathing Collapse Coughing Convulsions Diarrhoea Blindness
Rapid heart rate Lethargy Vomiting Fainting Sudden death Weight loss
Heartworm Disease in Animals and Humans
235
FIGURE 4.7 Small pulmonary arteriole of a heartworm-challenged cat on monthly chemoprophylaxis. (Used with permission of the American Heartworm Society.)
disease were manifested radiographically and histologically. Thus, unlike in dogs, heartworm disease in cats should not be linked only with adult worm infection, and airway and parenchymal disease should be associated with the arrival and early death of immature worms soon after they reach the pulmonary arteries. Levy et al. (2007) recently confirmed this finding in cats with naturally acquired heartworm disease. They evaluated pulmonary pathology in cats that were free of adult heartworms but were positive for heartworm antibodies, indicating either very early heartworm infection or previous aborted infection. Lung lesions characterized by pulmonary artery occlusive hypertrophy were common in cats with adult heartworms (80%) but not in cats free of heartworm and heartworm antibodies (10%). They also report that 50% of cats with heartworm antibodies, but not adult heartworms, had the same type of pulmonary lesions, suggesting that even transient infection with heartworm leaves cats with long-lasting pulmonary pathology.
236
John W. McCall et al.
FIGURE 4.8 Small pulmonary arteriole of a cat with a chemically abbreviated heartworm infection. (Used with permission of the American Heartworm Society.)
In acute cases, cats can die so quickly that pet owners are usually not able to report any clinical signs before sudden death (Dillon, 1988; Dillon et al., 1995b, 1997a,b). When the worms reach the pulmonary arteries, acute inflammatory responses at the vascular and parenchymal level follow. There is activation of pulmonary intravascular macrophages, normally numerous in comparison with dogs, and hyperplasia of pulmonary type II cells, which are producers of surfactant (Dillon et al., 1995b). The activation of pulmonary intravascular macrophages is a unique and likely the most important feature of heartworm infection in cats and is responsible for the exacerbated pulmonary reaction (Atkins et al., 1998a; Dillon et al., 1995b). The patient may present with respiratory signs that can be confused with asthma, and may be associated with vomiting, anorexia and prostration (Atkins et al., 1998c). At this stage, pulmonary radiographic findings include an interstitial inflammatory pattern related to the increased production of cytokines by macrophages (Dillon et al., 1995b; Holmes et al., 1992). If this acute stage is not fatal, the cat may tolerate the infection and become asymptomatic, or else becomes chronic. Chronic infection features one or more clinical signs that may or may not be present, including dyspnoea, sporadic vomiting, tachypnea, respiratory difficulty, weight
Heartworm Disease in Animals and Humans
237
FIGURE 4.9 Small pulmonary arteriole of a non-treated cat with adult heartworms. (Used with permission of the American Heartworm Society.)
loss and, rarely, cardiac murmurs (Dillon et al., 1987). Signs that are common in dogs like ascitis, exercise intolerance and CHF are not frequently noted in cats (Dillon et al., 1998). Some infected cats recover from the infection spontaneously, with or without symptomatic treatments (Atkins et al., 1995). Recently, Genchi et al. (in press) reported the results of a study performed to assess the duration, outcome (self-cure or death) and life expectancy of felineheartworm infected cats. Of 43 heartworm-infected cats in this trial, 9 (21%) died during the study and 34 self-cured (79%). Throughout the study, 27 cats showed no symptoms (63%), 3 died suddenly after 38–40 months from the diagnosis and 6 died during the follow-up period 8–41 months after the diagnosis (Table 4.6). Symptomatic cats, mainly showing signs of dyspnoea and vomiting, were treated at the onset of symptoms. Necropsy was performed on three sudden-death cats whose owners agreed to the necropsy. Severe thromboembolic processes were found in the pulmonary arteries with two to three worms trapped in the thrombi.
238
John W. McCall et al.
FIGURE 4.10 Bronchiole of a heartworm-challenged cat on monthly chemoprophylaxis. (Used with permission of the American Heartworm Society.)
The worms were still barely alive. According to the authors, the probability of death increased with the age at diagnosis. Litster et al. (in press), studying the pathogenesis of acute death syndrome in cats infected with D. immitis, determined that the most severe lung reactions occurred in hypersensitized cats exposed to dead worms or antigens collected from adult parasites. The authors described a marked increase in dyspnoea and a dramatic decrease in oxygen blood saturation. Clinical signs described were dyspnoea, retching, defecation or flatulence, urination, less frequently haemorrhage from the nostrils or the anus and sometimes facial swelling. The authors concluded that the acute death syndrome resembles an acute systemic anaphylactic reaction. Even in cases of chronic infection, there is always the possibility of reversion to the dangerous acute phase when adult parasites die. Adult worm death is associated with an intense pulmonary inflammatory reaction in response to thromboembolism that in turn is responsible for pulmonary infarction and haemorrhage. Circulatory collapse and respiratory failure usually follow. Clinical signs at this stage can include dyspnoea, cyanosis, hypothermia, ataxia, haemoptysis and syncope (Rogers, 1998).
Heartworm Disease in Animals and Humans
239
FIGURE 4.11 Bronchiole of a cat with a chemically abbreviated heartworm infection. (Used with permission of the American Heartworm Society.)
Recently bacteriae of the genus Wolbachia have been studied by several investigators as potential contributors to the pathology caused by D. immitis (Dingman et al., 2007; Genchi et al., 2001b; see Section 3.3). The general pulmonary pathology in the cat is similar to that in the dog and is covered in Section 3.1 of this chapter.
5.2. Diagnosis Diagnosis of feline heartworm disease is more elusive than for dogs. A willingness to pursue a high index of suspicion is critical and often involves the use of multiple procedures and tests, some of which
240
John W. McCall et al.
FIGURE 4.12 Bronchiole of a non-treated cat with adult heartworms. (Used with permission of the American Heartworm Society.)
may need to be repeated on several occasions (see http://www. heartwormsociety.com). Furthermore, clinical diagnosis of the disease is challenging due to the fact that many cats are asymptomatic. When symptoms are present, they are often non-specific and transitory or complicated by localization of the parasite in ectopic sites (Dillon et al., 1998; Henckler, 1998; Robertson-Plouch et al., 1998). This makes it difficult to diagnosis in a patient or to establish the true prevalence of infection. Moreover, the occurrence of sudden death complicates diagnosis even further (Dillon, 1988, 1998). In spite of these problems, several tests and procedures are available to facilitate the ante mortem diagnosis of infection (Dillon, 1984, 1998; Dillon et al., 1998; Labarthe et al., 1997b). Serological tests to detect antibodies or adult parasite antigens, thoracic radiography and echocardiography are
Heartworm Disease in Animals and Humans
241
FIGURE 4.13 Alveoli of a heartworm-challenged cat on monthly chemoprophylaxis. (Used with permission of the American Heartworm Society.)
FIGURE 4.14 Alveoli of a cat with a chemically abbreviated heartworm infection. (Used with permission of the American Heartworm Society.)
currently the most widely used diagnostic tools (Atkins et al., 1998a; Snyder et al., 2000; Venco et al., 1998a,b). The American Heartworm Society (Nelson et al., 2005b) has published a guide to aid in the interpretation of the results of these procedures and tests (Table 4.7).
242
John W. McCall et al.
FIGURE 4.15 Alveoli of a non-treated cat with adult heartworms. (Used with permission of the American Heartworm Society.)
FIGURE 4.16 Pulmonary artery of a heartworm-challenged cat on monthly chemoprophylaxis. (Used with permission of the American Heartworm Society.)
5.2.1. Detection of microfilariae Microfilariaemia is rare in infected cats. However, when circulating microfilariae are present, there is assurance that the cat is infected with adult D. immitis (Gomes et al., 2001).
Heartworm Disease in Animals and Humans
243
FIGURE 4.17 Pulmonary artery of a non-treated cat with adult heartworms. (Used with permission of the American Heartworm Society.)
TABLE 4.6
Duration (in months) of heartworm (HW) infection in naturally infected cats
Sudden death Death through HW followup Self-cured with symptoms Self-cured without symptoms a
No. of cats
AMa
Median
Range
3 6
39.0 24.2
39 21
38–40 8–41
23 11
35.3 32.7
40 33
18–49 21–48
AM: arithmetic mean.
5.2.2. Radiological examination Thoracic radiography is an important tool for the diagnosis of cardiopulmonary diseases. However, in heartworm infected cats thoracic abnormalities are often transient and highly variable or absent. In cases of confirmed diagnosis, radiography can be useful to monitor progression or resolution of the disease (Brawner et al., 1998). Findings such as enlarged peripheral branches of the pulmonary arteries (mainly in caudal lobes) associated with low definition of their margins (better visualized in
244
TABLE 4.7
Interpretation of heartworm diagnostic procedures/tests in the cat
Test
Brief description
Result
Interpretation
Limitations
Antibody test
Detects antibodies produced by the cat in response to presence of heartworm larvae. May detect infections as early as 8 weeks post-transmission by mosquito.
Negative
Antibodies confirm infection with heartworm larvae, but do not confirm disease causality.
Detects antigen produced by the adult female heartworm or from the dying male (>5) or female heartworms. Detects vascular enlargement (inflammation caused by young L5 and, later, hypertrophy), pulmonary parenchymal inflammation and oedema (the latter only in ARDSlike syndrome).
Negative
Lower index of suspicion Increasing index of suspicion; 50% or more cats will have pulmonary arterial disease; confirms cat is at risk Lower index of suspicion Confirms presence of heartworms Lower index of suspicion Enlarged arteries greatly increases index of suspicion
Radiographic signs subjective and affected by clinical interpretation.
Antigen test
Thoracic radiography
Positive
Positive
Normal Signs consistent with FHD
Immature or maleonly worm infections are rarely detected.
Echocardiography
Detects echogenic walls of the immature or mature heartworm residing in the lumen of the pulmonary arterial tree, if within the visual window of the ultrasound.
No worms seen Worms seen
No change to index of suspicion Confirms presence of heartworms in the structure
Ultrasonographer experience with heartworm detection appears to influence accuracy rate.
Note: In the cat, no single test will detect all heartworm cases. While antigen tests are highly specific for detecting adult heartworm antigen, they will not detect infections with only live male worms. The clinician must use a combination of test results to determine the likelihood of heartworm disease as the aetiology of the cat’s symptomatology. Reprinted by permission of the American Heartworm Society.
245
246
John W. McCall et al.
a ventrodorsal position) (Donahoe et al., 1976a,b; Suter and Lord, 1974), accompanied by varying degrees of pulmonary parenchymal disease and hyperinflation (Schafer and Berry, 1995), are considered the most typical features consistent with infection. Other suggestive findings include diffuse interstitial infiltrate, atelectasia and perivascular dense zones (Brawner et al., 1998). Recently, Venco et al. (in press) evaluated the clinical evolution, radiographic findings and outcome of naturally acquired feline heartworm infection in a field study performed on privately owned cats. Thirtyfour asymptomatic cats, positively diagnosed as infected with heartworm based on positive antibody and antigen tests together with an echocardiogram were used. These cats were examined every 3 months from the time of heartworm diagnosis to the final outcome (self-cure or death). The owners were instructed to take their cat to the Veterinary Hospital as soon as any clinical sign was observed. Self-cure was defined as a negative antigen test and no worms visualized by echocardiography. Twenty-eight of the 34 cats (82%) self-cured, 21 (62%) of them never showed clinical signs and 6 cats (18%) died suddenly. In the majority of cats (21 of 34, 62%) enrolled in the study, the infection lasted more than 3 years. Findings on thoracic radiographs were variable. Focal pulmonary parenchymal pattern and pulmonary artery enlargement in the caudal lung lobes were the most frequently observed abnormalities (Figs. 4.18 and 4.19). Litster et al. (2005) recently demonstrated a tendency for the cardiac silhouette to increase in size during infection. However, the differences in the parameters evaluated compared with normal cats appear very small and suggest that their use as a clinical-diagnostic instrument is extremely limited because it is not possible to establish a reliable cutoff value to discriminate between infected and healthy cats.
5.2.3. Antibody testing The most commonly utilized antibody detection tests are based on immunochromatographic (later flow) or ELISA technology. All commercially available tests have proven to be highly sensitive and specific, particularly in cats with relatively heavy, experimentally induced infections (Bestul et al., 1998; Frank et al., 1998a,b; McCall et al., 1995, 1998, 2001b; Watkins et al., 1998). Thus, antibody test were originally recommended first in those cases where heartworm was suspected in a cat (Genchi et al., 1998b; Knight et al., 2001; McCall et al., 2001b). However, more variable results have been obtained in some studies on cats with naturally acquired infections (Levy et al., 2003; Nelson, 1998; Snyder et al., 2000). According to Levy (2007), the specificity of the antibody detection tests on sera from cats with naturally acquired infections ranges from 78% to 99% and the sensitivity from 32% to 90%. It is important to keep in mind that a cat becomes sensitized as he/she repeatedly becomes infected. It is also
Heartworm Disease in Animals and Humans
247
FIGURE 4.18 Thoracic radiograph showing a focal bronchointerstitial pattern (white circle) in a Domestic Short Hair cat infected with heartworms.
FIGURE 4.19 Thoracic radiograph clearly showing enlargement of the cranial pulmonary artery (white arrow) in a heartworm infected Domestic Short Hair cat with an infection lasting more than 3 years.
known that infections interrupted either naturally or due to chemoprophylaxis are still immunogenic enough to stimulate a persistent antibody response even in the absence of adult parasites (Dillon et al., 1996, 1997a,b;
248
John W. McCall et al.
Donoghue et al., 1998; McCall et al., 2001b). Levy et al. (2003), in a serological survey of a group of naturally infected cats, found adult heartworms in the heart/lungs during necropsy in 4.9% of cats, while 14.8% had at least two positive results in the antibody tests. They concluded that approximately 10% of the cats were antibody-positive without evidence of the presence of the parasites. Therefore, the interpretation of a positive heartworm antibody test indicates that the cat may (1) be infected with an adult worm(s), (2) have an ectopic infection, (3) be infected with only L4 or L5 or (4) be cleared naturally or chemically of a recently acquired infection. Levy (2007) recently reported that in areas heavily endemic for heartworm infection in dogs, seropositivity in cats may reach 40% or more and that 3–15% of the cats are infected with heartworms. Interestingly, many (25%) antibody-positive cats live exclusively indoors (Atkins et al., 1998b; Miller et al., 1998; Robertson-Plouch et al., 1998). Lorentzen and Caola (in press) recently reported that the percentage of cats tested for feline heartworm disease in cats between 2000 and 2007 in the USA was only 0.4%, as compared with 38% for dogs.
5.2.4. Antigen testing The ELISA- or immunochromatography-based antigen tests for detection of D. immitis circulating antigens are heavily dependent on the number of mature female parasites present in the animal (Fox et al., 1998; McCall et al., 1995, 1998). In earlier studies, all of them were considered to be highly specific and sensitive for experimentally induced infections in cats with relatively heavy, female worm burdens, but sensitivity was relatively low in cats with more typical, light worm burdens (Genchi et al., 1998b; McCall et al., 1995, 1998). In general, it appears that the sensitivity of these tests has improved over the last several years (McCall et al., 2001c). Thus, for cats with naturally acquired infections, the results have been more variable over time. Levy (2007) recently reported a specificity of 78–99% and a sensitivity of 68–86% for cats with naturally acquired infections. These authors reported that only one of nine cats confirmed positive by necropsy had a negative antigen test result. A positive antigen detection test result is indicative of active adult infection; however, a negative test does not rule out that the animal may be infected with only male worms, a single female parasite, young (prepatent period) adult worms or very old adult worms, most of which are common for cats (McCall et al., 1995; Piche´ et al., 1998). Thus, their ability to consistently detect heartworm infection in cats generally requires the presence of two or more mature female worms (McCall et al., 1998). The accuracy of diagnosis is increased significantly by the concomitant use of both the antibody and antigen tests (Snyder et al., 2000) and this combination is now recommended by the American Heartworm Society (Nelson et al., 2007).
Heartworm Disease in Animals and Humans
249
Based on an analysis of data collected from 2000 to 2007 on antigen testing of cats in different regions of the USA, Lorentzen and Caola (in press) show that in certain regions the prevalence is close to half of that for the canine heartworm-infected population.
5.2.5. Ecocardiography This technique is a useful diagnostic tool with high sensitivity, but relies strongly on the expertise of the individual performing the test. (DeFrancesco and Atkins, 1998; DeFrancesco et al., 2001; Venco et al., 1998a,c). The adult parasite is echogenic, producing images of two, short parallel lines (0.5–1 cm in length) (Figs. 4.20 and 4.21). The pulmonary arteries (PA), right ventricle (RV) or rarely the right atria (RA) must be examined carefully because infections with one or only a few parasites could be overlooked. Recently, Atkins et al. (in press) evaluated 80 adult cats, each inoculated SC with 60 D. immitis L3 on test day 1, to test the potential quantification of the parasite burden by echocardiography. The cats were examined at 8 months and again at approximately 12 months
FIGURE 4.20 Echocardiogram (right parasternal short-axis view) of a heartworm infected cat. The heartworm is visualized as a double, linear parallel object (white arrow) floating in the lumen of the right atrium.
250
John W. McCall et al.
FIGURE 4.21 Echocardiogram (right parasternal long-axis view) of the same cat as in Fig. 4.20. The worm is visualized in the right ventricle (white arrow).
post-infection to confirm and enumerate the number of adult D. immitis residing in the cardiovascular system. Worm burdens were confirmed at necropsy. Worm burden by echocardiography correlated well, but not precisely, with post-mortem counts and under-, over- and exactly estimated the actual post-mortem heartworm burden 53%, 27% and 22% of the time, respectively. Both false-negative and false-positive results were obtained. In any case, echocardiography, combining both high sensitivity and specificity, must be considered a fundamental tool for the diagnosis of feline heartworm infection.
5.2.6. Tracheal wash cytology The findings are generally not specific. Eosinophilia is a consistent finding; however, other parasitic infections could also be responsible (Rogers, 1998). Eosinophilia in tracheal washes may also be indicative of heartworm infections but we have to keep in mind that Paragonimus kellicotti or the larvae of Aelurostrongylus abstrusus could also produce this cytological picture.
Heartworm Disease in Animals and Humans
251
5.2.7. Necropsy Necropsy has long been considered a gold standard for post-mortem diagnosis, but dissection of the heart and lungs is insensitive for immature infections as well as ectopic infections (Atkins et al., 1998c; Genchi et al., 1998b; Levy et al., 2007; McCall et al., 1992, 1995). For that reason, it is recommended that the vena cava, right heart and pulmonary arteries be dissected and examined up to their finest and most distal branches for parasite fragments or immature parasites (Miller et al., 1998). In cases of sudden death, the necropsy is imperative. If neurological signs were seen before death, a more complete necropsy involving the brain and spinal cord, as well as the heart and lungs and body cavities, should be done, keeping in mind the type of signs observed (Atkins et al., 1995).
5.2.8. Differential diagnosis Dirofilariosis must be a part of the differential diagnosis when a cat is presented with respiratory signs, such as dyspnoea and coughing, as well as vomiting not associated with feeding or sudden death (Berndt and Ware, 1987; Dillon, 1988, 1998). Keep in mind that HARD can be presented even before the parasite reaches the adult stage. Pneumothorax, chylothorax, pulmonary abscesses, pleural effusion, bronchitis, asthma or hypertension also can be caused by D. immitis (Rogers, 1998). Other parasites, mainly A. abstrusus, can also cause respiratory signs in cats (Calvert et al., 1994; Dillon, 1984).
5.3. Chemoprophylaxis Monthly chemoprophylaxis is the only safe and effective option for cats living in areas where heartworm infection is considered endemic in dogs and exposure to infective mosquitoes is possible (Blagburn, 2001). Even cats that are considered to be ‘indoor-only’ by their owners may be at risk. Several heartworm chemoprophylactics are available for use by the veterinarian. Monthly doses of either ivermectin or milbemycin oxime orally, topical selamectin or topical moxidectin plus imidacloprid are highly effective and safe for routine use (Arther et al., 2005; Clark et al., 1992; Guerrero et al., 2002; Longhofer et al., 1995; McTier et al., 1992b, 1998; Paul et al., 1992; Stewart et al., 1992a,b). Preventives should be started in kittens at 8 weeks of age and be administered to all cats in heartworm endemic areas during the heartworm transmission season. The individual monthly prophylactic doses for these preventatives are 24 mg/kg of body weight for ivermectin, 2.0 mg/kg for milbemycin oxime, 1 mg/kg for moxidectin and 6–12 mg/kg for selamectin. Administration of these drugs in cats is not
252
John W. McCall et al.
precluded by antibody or antigen seropositivity. The efficacy of DEC for heartworm chemoprophylaxis in cats has not been evaluated. Administering a preventive to cats year-round has merit due to several reasons: complementary activity against some common intestinal parasites and, in the case of topical imidacloprid and moxidectin and topical selmectin, activity against some ectoparasites. Year-round preventatives have greater owner compliance and retroactive efficacy as a safeguard for inadvertently missed doses. In view of the difficulty in diagnosis and adulticide treatment of feline heartworm disease, the only reasonable choice for control of heartworm disease is the monthly administration of heartworm preventive medication to cats in any area where dogs receive the heartworm preventive treatment.
5.4. Symptomatic treatment If a cat does not display overt clinical signs despite radiographic evidence of pulmonary vascular/interstitial lung disease consistent with HARD, it may be advised to allow time for self-cure. These subclinical cases can be monitored periodically (6- to 12-month intervals) by antibody and antigen re-testing and thoracic radiography. In those cats destined to recover, regression of radiographic signs and especially seroconversion of a positive antigen test to negative status provide evidence that the period of risk probably has passed (Nelson et al., 2007). Prednisone in diminishing doses is often an effective medical support for infected cats with radiographic evidence of lung disease, whether or not they are symptomatic. Also, this should be initiated whenever antibody- and/or antigen-positive cats display clinical signs. An empirical oral regimen is 2 mg/kg body weight/day, declining gradually to 0.5 mg/kg every other day by 2 weeks and then discontinued after an additional 2 weeks. At that time, the effects of treatment should be reassessed based on the clinical response and/or thoracic radiography. This treatment may be repeated in cats with recurrent clinical signs (Atkins et al., 1998a). Cats that become acutely ill need to be stabilized promptly with supportive therapy appropriate for treating shock. Depending on the circumstances, this may include IV corticosteroids, balanced electrolyte solutions, bronchodilators and oxygen via intranasal catheter or closed cage. Diuretics are inappropriate, even for infected cats with severe interstitial or patchy alveolar lung patterns. Aspirin and other non-steroidal anti-inflammatory drugs have failed to produce demonstrable benefit and actually may exacerbate the parenchymal pulmonary disease (Atkins et al., 1998a).
Heartworm Disease in Animals and Humans
253
5.5. Adulticide therapy Adulticidal treatment is considered the last-resort treatment for cats in stable condition but which continue to manifest clinical signs that are not controlled by empirical corticosteroid therapy. There is insufficient experience with melarsomine dihydrochloride at this time, thus melarsomine is not recommended for use in cats. Preliminary data suggests that melarsomine is toxic to cats at doses as low as 3.5 mg/kg (see Goodman, 1996; McLeroy, 1998). Ivermectin given orally at 24 mg/kg monthly for 2 years has been reported to reduce worm burdens by 65% as compared with untreated cats (Guerrero et al., 2002). Since most cats have light worm burdens, it is not worm mass alone that is problematic, but the ‘anaphylactic’ type reaction that results when the worms die. This will likely also occur when the ivermectin-treated worms die but the extent of the reaction is unknown and unpredictable in both scenarios.
5.6. Surgical extraction of adult worms In principle, it is preferable to remove heartworms rather than destroy them in situ. This can be accomplished successfully by introducing brush strings (Venco et al., 1998c), catheters or loop snares via right jugular venotomy, or alligator forceps can be inserted through a right ventricular purse string incision after left thoracotomy (Glaus et al., 1995; Rawlings et al., 1994; Venco et al., 1998c). Before attempting either approach, heartworms should be identified ultrasonographically (Borgarelli et al., 1997) in locations that can be reached with these inflexible instruments. Although it may not be possible to retrieve every worm, the surgical option may be a reasonable alternative to symptomatic support or adulticide treatment of cats that are heavily infected and/or are in critical condition (Rawlings et al., 1994). This procedure has to be performed carefully, as the accidental dissection of a worm can lead to a shock-like reaction and to death of the cat (Venco et al., 1998a).
6. HEARTWORM DISEASE IN FERRETS 6.1. Clinical presentation Although ferrets (Mustela putorius furo) are highly susceptible to heartworm infection and worm burdens are relatively light compared with dogs, even a few adult worms can cause severe disease and even death of ferrets (Kemmerer, 1998). Many cases of unexplained sudden death of ferrets in heartworm endemic areas are heartworm related (Kemmerer, 1998). Due to the large size of the adult heartworms and the small
254
John W. McCall et al.
pulmonary arteries and right heart chambers, most live worms inhabit the cranial and caudal venae cavae of ferrets (Supakorndej et al., 1992, 1995). Clinical signs of heartworm disease in ferrets are similar to those seen in dogs, but often progress much more rapidly and thus require more urgency for early diagnosis and management of the patient. Clinical signs upon presentation include lethargy, inappetance, exercise intolerance, pleural effusion, cyanosis and dyspnoea. In one study on ferrets with naturally acquired infections, the following clinical signs were scored as mild, moderate or severe: coughing (92%, moderate to severe), dyspnoea (84%, severe), exercise intolerance (84%, moderate to severe), cyanosis (84%, moderate), pleural effusion (92%, severe), heart murmur (84%), decreased appetite/weight loss (84%, mild to severe), posterior paresis (23%, severe) and vomiting (7%, mild) (Antinoff, 2001). In this same study, haematological abnormalities consistently present included monocytosis and anaemia (43%). The most consistent biochemical abnormality was mild hyperchloremia (43%). By urinalysis, bilirubinuria was detected in 83% and trace amounts of blood were detected in 67% of the ferrets. One of seven (14.3%) had caval syndrome. Elevations in liver enzymes were only sporadically detected. In a study with 10 male and female ferrets experimentally infected with heartworm L3 and 10 male and female non-infected ferrets, eosinophil counts in infected ferrets were generally higher than those in non-infected ferrets beginning at week 22 and significantly higher beginning at week 30, with peak values at week 34. There were no dramatic changes in other haematological, serum biochemical or urinalysis values (Supakorndej et al., 1992). In regard to pathology, the cause of death in cases of sudden death of ferrets with naturally acquired infections appears to be due to pulmonary embolism, with entire lung lobes blackened and necrotic. In patients who show ante mortem signs, microscopic pathology very often shows a severe pulmonary arteritis and an eosinophilic or granulomatous pneumonitis (Kemmerer, 1998).
6.2. Diagnosis The same diagnostic tests and procedures used in dogs and cats generally can be used in ferrets, with varying degrees of success (McCall, 1998). Histochemical stains and molecular methods generally are not needed in detecting heartworm infection in ferrets, but they should be as effective in this species as in dogs and cats.
6.2.1. Blood tests for microfilariae Infections in ferrets that have circulating microfilariae are usually patent (i.e., microfilaraemic) by the seventh month after infection (Supakorndej et al., 1992). However, concentration tests (modified Knott test or filtration
Heartworm Disease in Animals and Humans
255
tests) are generally unreliable screening tests due to the low level and transient microfilaraemia in ferrets (McCall, 1998).
6.2.2. Blood test for adult female circulating antigen Adult heartworm antigen detection tests available for use in dogs and cats have been used with much success in diagnosing infection in ferrets (Kemmerer, 1998; Supakorndej et al., 1992). In two studies with experimentally induced infections, 40–80% of the infected ferrets were antigenpositive 4 months after inoculation and all were positive at 6 months. This is at least 1 month earlier than in dogs and cats, probably due to a greater concentration of the antigen in the smaller blood volume of the ferret (Supakorndej et al., 1992, 1994).
6.2.3. Radiography, angiography and echocardiography Radiographic changes seen in heartworm-infected ferrets differ from those seen in dogs and cats (McCall, 1998). Heartworm-infected dogs manifest disease radiographically by the enlargement of the pulmonary arteries, prominent pulmonary trunk and an enlarged right heart. Signature radiographic features of heartworm disease in cats most commonly include enlargement of the pulmonary arteries with ill-defined margins (mainly caudal lobes) and pulmonary infiltrate (Donahoe et al, 1976b). Classic radiographic changes in infected ferrets are cardiomegaly and severe pleural effusion (Kemmerer, 1998; Supakorndej et al., 1995). Adult heartworms are readily visualized as filling defects in the cranial (and sometimes caudal) venae cavae of infected ferrets (Supakorndej et al., 1995) and the right heart and pulmonary arteries of dogs (Thrall et al., 1979) and cats (L. Neuwirth and J. W. McCall, 1997, unpublished data) by non-selective angiography (Fig. 4.22). Adult heartworms are detected sonographically as early as 5 months after experimentally induced infection in ferrets (L. Neuwirth and J. W. McCall, 1997, unpublished data) and 6.5 months PI in cats (Selcer et al., 1996).
6.3. Chemoprophylaxis No drug is approved for use in preventing heartworm disease by suppressing the development of the migrating larval parasites in the host tissue of ferrets. Fortunately, there are several reports of macrocyclic lactone preventatives approved for dogs and cats being highly effective in preventing the disease in ferrets. Single oral doses of ivermectin (premarket formulation) given at 50 or 200 mcg/kg to ferrets 1 month after inoculation of infective, L3 were 100% protective and a dose of 12.5 mcg/kg was 84% protective (Blair and Campbell, 1978; Campbell and Blair, 1978). Supakorndej et al. (1992) reported that ivermectin given as a single dose of
256
John W. McCall et al.
FIGURE 4.22 Lateral thoracic angiogram of an infected male ferret taken at 40 weeks PI showing enlargement of the right atrium and cranial vena cava (white arrow). Worms inside the enlarged cranial vena cava can be seen as dark, filling defects (large black arrow). A worm can be seen in the azygous vein (two black arrows). (Used with permission of the American Heartworm Society.)
3.0 or 6.0 mcg/kg was 100% protective and a dose of 0.5 mcg/kg was partially effective. Other macrocyclic lactones are effective in preventing heartworm infection. In a recent study, topical application of a product containing imidacloprid (10%) plus moxidectin (1%) at a dosage range of 1.9– 3.33 mcg/kg was completely effective in preventing heartworm disease in ferrets (Shaper et al., 2007). When given topically at the dosage recommended for dogs and cats (minimum, 6 mg/kg) 1 month after infection, selamectin was highly effective (99.5%) in preventing the disease ( J. W. McCall, 2000, unpublished data).
6.4. Therapy against adult heartworms Adulticidal therapy for heartworm-infected ferrets has proved to be only marginally successful, at best (Kemmerer, 1998). Both thiacetarsamide sodium, which is no longer available, and melarsomine dihydrochloride have been used and the survival rate with both drugs has been
Heartworm Disease in Animals and Humans
257
disappointing. Ferrets are at high risk of sudden death due to thromboembolism following treatment. Although intramuscular administration of melarsomine makes it a more attractive option, compared with IV injection of thiacetarsamide, survival with melarsomine has not been superior to thiacetarsamide. In one clinic study, 20 of 34 (59%) ferrets treated with thiacetarsamide at the dosage recommended for dogs survived while only 2 of 5 (40%) treated with melarsomine survived(Kemmerer, 1998). In another clinic, only 2 of 6 (33.3%) ferrets survived treatment with melarsomine (one IM injection of 2.5 mg/kg followed 1 month later by two injections 24 h apart plus ivermectin administered orally once a month at 50 mcg/kg) (Antinoff, 2001). The average survival time of the two remaining ferrets was 435 days. In the same clinic, only 2 of 4 (50%) ferrets survived treatment with ivermectin, with a mean survival time of at least 230 days, when observations were discontinued. In both groups, the ferrets also received prednisolone (1 mg/kg PO every 24 h), furosemide (2–4 mg/kg every 8–12 h), diltiazem (7.5 mg/kg every 24 h) and other cardiac medications whenever indicated by the patient’s clinical signs. A laboratory study with 28 ferrets experimentally infected by SC transplantation of six (three males, three females) juvenile heartworms and treated when the worms were 7 months old indicated that ferrets generally tolerated a high dose of 3.25 mg/kg (i.e., 30% higher than the dosage recommended for dogs) given IM once, twice (24 h apart) or thrice (one injection followed 1 month later by two injections 24 h apart). However, at necropsy for worm counts 3 months after the first or only treatment, efficacy in the surviving ferrets was only 35.3% for the single injection and ranged from 63.2% to 80.3% for the two- and three-injection protocols (J. W. McCall and P. Supakorndej, 2001, unpublished data). Moreover, two of seven (14.3%) ferrets given either a single injection or two injections and three of seven (42.9%) given three injections died during the study. Most died due to thromboembolism and pulmonary infarction of pulmonary arteries. An earlier study with experimentally infected ferrets given two IM injections of the dosage used for dogs (2.5 mg/kg) 24 h apart was only about 70% effective and the mortality rate was unacceptable ( J. W. McCall and P. Supakorndej, 1999, unpublished data). Thus, it appears that there may not be a significant advantage to melarsomine therapy in ferrets. It has been suggested that where heartworms are diagnosed on routine examination in a non-symptomatic or mildly symptomatic patient, use of prednisone and heartworm prevention, that is, monthly prophylactic doses of ivermectin, may be found to produce a higher long-term survival rate, but further study in this area is needed before this approach can be widely recommended (Kemmerer, 1998).
258
John W. McCall et al.
7. HUMAN DIROFILARIOSIS Several filarial nematodes of mammals have been reported to occasionally infect humans, including D. repens, D. ursi, D. tenuis and D. immitis. The public health significance of the different species depends largely on their geographic distribution. For instance, in Europe, the main concern is about D. repens. The number of zoonotic infections has dramatically increased in the last few decades [for a comprehensive review, see Pampiglione and Rivasi (2000)] and the infection now can be included in the list of emerging zoonoses. According to Raccurt (1999) and Pampiglione et al. (2001), 48% of the human cases in Italy and France (the second European country in prevalence for human infections) have been diagnosed in only the last 10 years, and the infection is spreading in many southern and eastern European countries (Fok, 2007; Kramer et al., 2007) probably as a consequence of the movement of infected dogs and global warming, both of which increase the transmission season. The infective larvae of zoonotic filariae invade a variety of human tissues and elicit little or no evident response from the host during the course of their development. However, when these parasites die in the tissues, the host mounts an inflammatory response to their presence. It is unclear in these cases whether the parasite becomes moribund and the host responds to the dying worm or whether the host ultimately mounts a response that kills the worm (Orihel and Eberhard, 1998). Humans are considered accidental hosts and these parasitic infections rarely reach patency. This section reviews what is currently known about human infection with D. immitis.
7.1. Clinical aspects of human dirofilariasis by D. immitis Humans living in areas endemic for canine heartworm infection are at risk of being bitten by an infected mosquito, which may then transmit the parasite during the blood meal. It is thought that many infective larvae are eliminated by the human host’s immune system (Simo´n et al., 2005). However, one or more worms may continue to migrate through the connective tissue and into the venous circulation and right heart, eventually reaching the pulmonary arteries. Here, development may continue within the small- to medium-sized branches of the pulmonary arterial tree. The nematode likely induces vasculitis and is ultimately killed or dies as a result of the inflammatory response (Theis, 2005). The dead/ dying worm is then incorporated into a granuloma, recognized as the typical ‘coin’ lesion on a chest radiogram or CT scan (Fig. 4.23). Radiological characteristics of the coin lesion show a solitary pulmonary nodule that has well-defined and smooth edges, is spherical or oval
Heartworm Disease in Animals and Humans
259
FIGURE 4.23 Typical ‘coin lesion’ containing a dead/dying heartworm in a granuloma in a lower lung lobe on a chest radiogram of an infected human. (Courtesy of Fernando Simo´n, University of Salamanca, Spain).
in shape and is homogeneously dense (Muro and Cordero, 2001). Most cases are asymptomatic at the time lesions are observed (Bielawski et al., 2001; Milanez de Campos et al., 1997; Stephen, 2001). However, in a recent review, Theis (2005) reported that patients may present coughing or some symptom of pneumonitis prior to diagnosis. The discovery of a coin lesion requires an intense diagnostic work-up to exclude other infectious agents (fungi, bacteria) and neoplasia. As there is yet no adequate or reliable diagnostic test available for pulmonary dirofilariasis, thoracotomy and surgical resection of the lung tissue containing the nodule(s) is the only option. It has been estimated that that the cost of evaluating a coin lesion is approximately $80,000 (Theis, 2005). Fortunately, surgery is also curative. Miyoshi et al. (2006) reported that video-assisted thoracic surgery is a less invasive alternative for surgical resection of coin lesions, including those due to D. immitis. Serology has been used in epidemiological surveys and for risk assessment studies (see below), but is not diagnostic. Miyoshi et al. (2006), for
260
John W. McCall et al.
example, reported that of seven cases of confirmed pulmonary dirofilariosis, only four showed a positive antibody response to Dirofilaria antigens by ELISA. On the other hand, a positive antibody response to D. immitis antigens, whether crude (Simo´n et al., 1991; Vieira et al., 1998) or purified (Perera et al., 1994, 1998; Sun and Sugane, 1992), does not necessarily indicate the presence of a pulmonary lesion (Simo´n et al., 2005) but may only reflect previous exposure to the parasite. Simo´n et al. (2003) reported a significantly higher anti-WSP seroprevalence in patients with confirmed pulmonary dirofilariasis when compared both to healthy individuals living in D. immitis-endemic areas and to humans from nonendemic zones. Indeed, the release of Wolbachia from a dead worm and its subsequent contact with the host’s immune system may aid in the serological diagnosis of disease. More recently, Morcho´n et al. (2006) reported that the same patients had a higher serum level of the eiconasoid thromboxane B2 than the two other groups. Despite these recent advances, however, the serological diagnosis of human pulmonary dirofilariasis is still unreliable. The vast majority of cases of D. immitis infection in humans report the detection of lesions in the lung. Even though this is likely due to the natural predisposition of this parasite for the pulmonary vasculature, it could also be due to the fact that the lung is an organ system that is readily surveyed by indirect, non-invasive techniques (radiography, computerized tomography and so on) (Theis, 2005). Indeed, ‘aberrant’ localization of D. immitis is a feature of canine and feline infection and has also been reported in humans. Theis et al. (2001b) reported the finding of an immature male D. immitis in a branch of the left testicular artery, the identity of which was confirmed by PCR. Kim et al. (2002) reported an immature male D. immitis within a hepatic nodule. These examples indicate that many different arteries may permit the localization and development of D. immitis and that an extrapulmonary location does not exclude D. immitis as the causative agent.
7.2. Epidemiology of human D. immitis infection Human infection with D. immitis has been reported in many countries of the world and is possible wherever the parasite is endemic in dogs. The large number of mosquito species that can transmit the parasite, together with the natural sharing of environmental habitats by humans and dogs, likely accounts for the high risk of transmission (Genchi et al., 2001a). It has been suggested that due to the asymptomatic nature of the infection and the lack of a reliable diagnostic test, the number of cases of human pulmonary dirofilariasis is likely underestimated (Theis, 2005). Several surveys and isolated case reports, however, allow a glimpse of what Simo´n et al. (2005) have termed ‘the tip of the iceberg’.
Heartworm Disease in Animals and Humans
261
Animal dirofilariosis is endemic in several Mediterranean countries, and many human cases have been found in Spain, France, Italy and Greece (Muro et al., 1999). In Spain, cases of pulmonary dirofilariosis have been reported especially in the western part of the country (Cordero et al., 1990, 1992; Ruiz-Moreno et al., 1998). Most human cases from France have been reported in areas near the Mediterranean coast (Languedoc-Roussillon, Provence-Coˆte d’Azur) and Corse (Raccurt, 1999). In Italy, human dirofilariosis is distributed throughout the country, with varying incidence. The majority of human cases reported have been attributed to D. repens (Pampiglione and Rivasi, 2000; Pampiglione et al., 2001) in spite of the fact that D. immitis predominates in the northern part of Italy where D. immitis is predominant (Genchi et al., 2001a). In Greece, the first case of pulmonary dirofilaiosis was published only recently, suggesting a change in epidemiology (Pampiglione and Rivasi, 2000). In central and northern European countries, which are currently considered non-endemic for canine heartworm infection, isolated cases of human dirofilariasis are being detected with increasing frequency. Jelinek et al. (1996), for example, reported two cases of pulmonary dirofilariosis in a German hospital. In Austria, Auer (2003) reported 14 cases from 1981 to 2003 plus 20 serologically positive individuals with clinical symptoms and/or geographic anamnesis associated with SC and pulmonary dirofilariosis; however, it is uncertain if the infections were caused by D. immitis or D. repens. Recently, Fok (2007) reviewed human dirofilariosis in Hungary and concluded that all the autochtonous infections were caused by D. repens, including the first described by Babes in 1879 and one unusual case of spermatic cord localization reported by Pampiglione et al. (1999). Due to the difficulty of diagnosing pulmonary dirofilariasis, published case reports likely do not reflect the true risk that D. immitis poses for human health. Serological studies, on the other hand, can offer useful data on the epidemiological situation of a given population. In an endemic zone of western Spain, for example, where D. immitis infects 33.3% of the resident dogs, 5.6% of the population had circulating IgG, 2.6% had IgM and 12.6% had IgE against D. immitis antigens (Espinoza et al., 1993; Muro et al., 1991; Simo´n et al., 1991), for a total seroprevalence of approximately 21%. Moreover, in a radiological follow-up of the same population, eight cases of pulmonary dirofilariosis were found over a 2-year period. This suggests that the parasitosis is likely under-diagnosed (Muro et al., 1999). In another study, conducted in a highly endemic area of Italy (Pavia), seroprevalence in humans for D. immitis was approximately 33% (Prieto et al., 2000). Uge et al. (1990) reported that approximately 50–60% of the dogs in Japan that do not receive preventive treatment for heartworm disease are infected with D. immitis. Recently, Miyoshi et al. (2006) reported a case of human pulmonary dirofilariosis and reviewed the literature concerning
262
John W. McCall et al.
the infection in Japan. They reported that a further 24 cases of human pulmonary dirofilariosis had been reported in Japan from 1998 to 2004. Of these 24 patients with human pulmonary dirofilariosis, 12 (50%) were men. Their average age was 54 years and ranged from 29 to 80 years. Of the patients reviewed, 67% were asymptomatic. In the symptomatic cases, the most common symptom was cough (25%) and other symptoms resembling the common cold. No serological studies have yet been carried out in Japan. Theis (2005) recently published a retrospective study of human pulmonary dirofilariosis in the USA. This study clearly shows that the vast majority of these cases were in patients living in the highly endemic, south-eastern areas of the country. A total of 110 cases have been diagnosed since 1941. Many important features of the disease were also reviewed, including salient information during diagnostic work-ups. Coin lesions due to D. immitis ranged from 1 to 4.5 cm in diameter. The vast majority of patients were asymptomatic at the time of diagnosis (this is true in most cases of coin lesion). The vast majority of coin lesions due to D. immitis were located in the peripheral areas of the lungs and most of the worms were identified as immature adults recently molted from L4. Recently, Vezzani et al. (2006) reviewed human D. immitis infections in Central and South America. Since the first case (De Magalha˜es, 1887) in Brazil, 50 new cases have been documented, mainly from Rio de Janeiro, Sa˜o Paulo and Floriano´polis. In a retrospective study of 24 human cases (1982–1996) from Sa˜o Paulo, Brazil, Milanez de Campos et al. (1997) found a higher prevalence in male individuals (70%) than in female (30%). Fiftyfour per cent of the patients were asymptomatic and 75% had a wellcircumscribed, non-calcified, subpleural, pulmonary nodule. The authors concluded that a subpleural, non-calcified pulmonary nodule in the appropriate clinical and epidemiological setting should alert the clinician to the possibility of Dirofilaria infection and that human pulmonary dirofilariois should be considered in the differential diagnosis of pulmonary nodules. Sporadic cases have been reported from Venezuela, Colombia and Argentina. In addition, Vieira et al. (1998) detected serological evidence for human dirofilariosis in inhabitants of the Colombian Amazon.
7.3. Conclusions Many vector-borne diseases are currently considered emerging and/or increasing in prevalence (Pherez, 2007), including human dirofilariosis. The particular severity of the pulmonary localization of D. immitis in the human host makes this infection a diagnostic challenge and a therapeutic risk. There is most definitely a need for a reliable serological test to aid physicians in their decision whether to resect a lung nodule immediately
Heartworm Disease in Animals and Humans
263
or to ‘watch and wait’, thus saving patients the risk, pain, expense and psychological strain of surgery. However, and this cannot be repeated enough, the only sure way to protect human health from this zoonotic parasite is to administer preventive treatment to dogs living in endemic areas. There are many safe and highly effective drugs that can prevent heartworm disease in animals and only through their widespread use, human infection will no longer pose a risk.
8. EMERGING STRATEGIES IN HEARTWORM TREATMENT AND CONTROL 8.1. Arsenical therapy Arsenical drugs have been the mainstay for heartworm adulticidal therapy for the last four to five decades. However, treatment strategies have changed and indeed continue to evolve since melarsomine dihydrochloride replaced thiacetsamide sodium in the early 1990s. The manufacturer provides two treatment protocols for melarsomine: the ‘standard’ two-injection protocol for Class I (mild clinical signs) and Class II (moderate clinical signs) dogs and the ‘alternative’ threeinjection protocol for Class III (severe clinical signs) disease (see Section 4.4). Generally, this was followed 3–4 weeks later by administration of a microfilaricide that often had to be repeated once or twice to clear the dog of circulating microfilariae.
8.2. ‘Safety-Net’ and adulticidal properties of prolonged monthly prophylactic doses of macrocyclic lactones Starting in 1995, several studies have demonstrated that prolonged administration of monthly prophylactic doses of a macrocyclic lactone, particularly ivermectin, kills older larvae, immatures (juveniles), young adults and mature adults (see McCall, 2005). Moreover, a high percentage of the dogs become amicrofilaraemic within 6–9 months after dosing is started. The rate-of-kill with this slow-kill treatment is dependent on the age of the heartworms when treatment is started, with 3-month-old larvae requiring up to 1 year and mature adults needing 2.5 years to provide efficacy of at least 95%. Although monthly ivermectin is not an approved alternative to melarsomine therapy and it is particularly risky in very active and symptomatic dogs (Venco et al., 2004; see Section 4.4), it clearly provides potent ‘safety-net’ activity against older larvae, ‘juveniles’, and young adults in cases of owner compliance failure, even when the owner and veterinarian are not aware that the animal is infected.
264
John W. McCall et al.
8.3. Arsenical plus prophylactic doses of macrocyclic lactones for adulticidal therapy The American Heartworm Society (Nelson et al., 2005a) recognizes the safety-net and adulticidal properties of some of the macrocyclic lactones, particularly IVM. Their 2005 canine guidelines state that administration of a chemoprophylactic dose of a macrocyclic lactone should begin as soon as the dog is diagnosed with a heartworm infection. While controversial due to the theoretical risk of inducing resistance to macrocyclic lactones, it may be beneficial to administer a macrocyclic lactone for up to 6 months prior to administration of melarsomine, when the presentation does not demand immediate intervention. The reasoning for this approach is to greatly reduce, if not completely eliminate, circulating microfilariae and kill migrating D. immitis larvae, and in the case of ivermectin, stunt immature D. immitis and reduce female worm mass by inhibiting the reproductive system. Moreover, administration for 3 months also will allow immature worms to reach an age at which they are known to be susceptible to killing by melarsomine (Atkins and Miller, 2003; Keister et al., 1992) and administration for longer than 3 months should result in reduced antigenic mass that in turn may reduce the risk of thromboembolism.
8.4. Anti-filarial effects of tetracyclines and doxycycline plus ivermectin By eliminating the Wolbachia endosymbionts, tetracyclines have been shown to prevent development of the larval stages, inhibit embryogenesis in adult worms and eventually kill adult filariae, usually one and one-half to two years after treatment is started, of several species of filariae in laboratory animals, large animals and humans (see Kramer et al., 2007; Taylor et al., 2005a). In view of the above, a recent laboratory trial with 30 dogs experimentally infected with adult heartworms tested the efficacy of weekly prophylactic doses of ivermectin (6 mcg/kg) and intermittent 2- to 6-week regimens of doxycycline (10 mg/kg/day) alone and together for 9 months (McCall, 2007). Also, one group was given ivermectin and doxycycline plus melarsomine and one received only melarsomine. Ivermectin or doxycycline alone gradually reduced microfilaraemia levels to negative or near-negative values, while no microfilariae were seen in the ivermectin plus doxycycline group after week 9. Compared with non-treated controls, ivermectin plus doxycyxline reduced the adult worm burden by 78.3%, while these drugs administered individually were only 8.7% (doxycycline) and 20.3% (ivermectin) effective. Interestingly, Wolbachia were still detected at necropsy in worms from dogs treated with doxycycline only,
Heartworm Disease in Animals and Humans
265
but were virtually eliminated in those receiving both doxycycline and ivermectin (Bazzocchi et al., in press). Gross pathology and histopathological observations of the lungs revealed the most severe pathology in the dogs receiving only melarsomine and the least evidence of pathology in those receiving doxycycline, ivermectin and melarsomine. These latter dogs also showed significantly less severe arterial lesions and a virtual absence of thrombi. In the same study (McCall, 2007), L3 collected from mosquitoes fed on microfilaraemic blood from dogs treated with doxycycline were normal in appearance and motility but were not able to develop in dogs, thus preventing further transmission of the disease even when microfilariae were present. These and other observations strongly suggest that administration of both doxycycline and ivermectin for several months prior to melarsomine or without melarsomine will eliminate adult heartworms with less potential for severe thromboembolism than melarsomine alone and will block transmission. Thus, it is quite likely that doxycycline eventually will be included in heartworm adulticide therapy for dogs. Furthermore, the potentially life-threatening infections and high risk associated with melarsomine treatment strongly encourage the testing of doxycycline plus ivermectin as an alternative adulticidal therapy for heartworm-infected cats and ferrets.
ACKNOWLEDGMENTS The authors thank Drs. Byron Blagburn and Ray Dillon, College of Veterinary Medicine, Auburn University for figures 4.7, 4.8, 4.9, 4.10, 4.11, 4.12, 4.13, 4.14, 4.15, 4.16 and 4.17
REFERENCES Abraham, D. (1988). Biology of Dirofilaria immitis. In ‘‘Dirofilariasis’’ (P. F. L. Boreham and R. B. Atwell, eds.), pp. 29–46. CRC Press, Inc., Boca Raton, Florida. Abraham, D., and Grieve, R. B. (1991). Passive transfer of protective immunity to larval Dirofilaria immitis from dogs to Balb/c mice. J. Parasitol. 77, 254–257. Alves, L. C., Silva, L. V. A., Faustino, M. A. G., McCall, J. W., Supakorndej, P., Labarthe, N. W., Sanchez, M., and Caires, O. (1999). Survey of canine heartworm in the city of Recife, Pernambuco, Brazil. Memo´rias do Instituto Oswaldo Cruz 94, 587–590. American Heartworm Society (2005). 2005 guidelines for the diagnosis, prevention and management of heartworm (Dirofilaria immitis) infection in dogs. URL: www. heartwormsociety.org. Antinoff, N. (2001). Clinical observations in ferrets with naturally occurring heartworm disease and preliminary evaluation of treatment, with ivermectin and with and without melarsomine. In ‘‘Recent Advances in Heartworm Disease, Symposium ’01’’ (R. L. Seward, ed.), pp. 45–47. American Heartworm Society, Batavia, Illinois.
266
John W. McCall et al.
Arther, R. G., Charles, S., Ciszewski, D. K., Davis, W. L., and Settji, T. S. (2005). Imidacloprid/ moxidectin topical solution for the prevention of heartworm disease and the treatment and control of flea and intestinal nematodes of cats. Vet. Parasitol. 133, 219–225. Atkins, C. E., and Miller, M. W. (2003). Is there a better way to administer heartworm adulticidal therapy? Vet. Med. 98(4), 310–317. Atkins, C. E., Atwell, R. B., Dillon, R., Genchi, C., Hayasaki, M., Knight, D. H., Lukof, D. K., McCall, J. W., and Slocombe, J. O. D. (1995). Guidelines for the diagnosis, treatment and prevention of heartworm (Dirofilaria immitis) infection in cats. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Kinght, eds.), pp. 309–312. American Heartworm Society, Batavia, Illinois. Atkins, C. E., Atwell, R. B., Courtney, C. H., Dillon, R., Genchi, C., Hagio, M., Holmes, R. A., Knight, D. H., Lukof, D. K., McCall, J. W., and Venco, L. (1998a). Guidelines for the diagnosis, treatment and prevention of heartworm (Dirofilaria immitis) infection in cats. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 265–271. American Heartworm Society, Batavia, Illinois. Atkins, C. E., DeFrancesco, T. C., Coats, J. R., and Keene, B. W. (1998b). Feline heartworm disease: The North Carolina experience. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 123–125. American Heartworm Society, Batavia, Illinois. Atkins, C. E., DeFrancesco, T. C., Miller, M. W., Meurs, K. M., and Keene, B. (1998c). Prevalence of heartworm infection in cats with signs of cardiorespiratory abnormalities. J. Am. Vet. Med. Assoc. 212, 517–520. Atkins, C. E., Arther, R. G., Ciszewski, D. K., Davis, W. L., Ensley, S. M., Guity, P. S., Chopade, H., Hoss, H., and Settje, T. L. Echocardiographic quantification of Dirofilaria immitis in experimentally-infected cats. Vet. Parasitol. In press. Atwell, R. B., and Buoro, I. B. J. (1988). Caval syndrome. In ‘‘Dirofilariasis’’ (P. F. L. Boreman and R. B. Atwell, eds.), pp. 191–203. CRC Press, Inc., Boca Raton, Florida. Atwell, R. B., and Rezakhani, A. (1986a). Inoculation of dogs with artificial larvae similar to those of Dirofilaria immitis: Distribution within the pulmonary arteries. Am. J. Vet. Res. 47, 1044–1045. Auer, H. (2003). Cases of human dirofilariosis in Austria. In ‘‘Helmintologische Fachgespra¨che Helmintological Colloquium. Wein 14th November 2003’’ (H. Auer, C. Ho¨rweg, and H. Sattmann, eds.), Naturhistorische Museum Wien www.vu-wien.ac.at/i116/ oegtp/downloads_oegtp/Fachgespraeche _2003.pdf. p. 15. Babes, V. 1879 cited by Kotla´n, 1951: Kotla´n, A. (1951). On a new case of human filarioidosis in Hungary. Acta Vet. Acad. Sci. Hung. 1, 69–79. Badertscher, R. R., Losonsky, J. M., Paul, A. J., and Kneller, S. K. (1988). Two dimensional echocardiography for diagnosis of dirofilariasis in nine dogs. J. Am. Vet. Med. Assoc. 7, 843–846. Bandi, C., McCall, J. W., Genchi, C., Corona, S., Venco, L., and Sacchi, L. (1999). Effects of tetracycline on the filarial worms Brugia pahangi and Dirofilaria immitis and their bacterial endosymbiont Wolbachia. Int. J. Parasitol. 29, 357–364. Bandi, C., Trees, A. J., and Brattig, N. W. (2001). Wolbachia in filarial nematodes: Evolutionary aspects and implications for the pathogenesis and treatment of filarial diseases. Vet. Parasitol. 98, 215–238. Bazzocchi, C., Ceciliani, F., McCall, J. W., Ricci, I., Genchi, C., and Bandi, C. (2000). Antigenic role of the endosymbionts of filarial nematodes: IgG response against the Wolbachia surface protein in cats infected with Dirofilaria immitis. Proc. R. Soc. Lond. B Biol. Sci. 267, 1–6. Bazzocchi, C., Genchi, C., Paltrinieri, S., Lecchi, C., Mortarino, M., and Bandi, C. (2003). Immunological role of the endosymbionts of Dirofilaria immitis: The Wolbachia surface protein activates canine neutrophils with production of IL-8. Vet. Parasitol. 117, 73–83. Bazzocchi, C., Comazzi, S., Santoni, R., Bandi, C., Genchi, C., and Mortarino, M. (2007). Wolbachia surface protein (WSP) inhibits apoptosis in human neutrophils. Parasite Immunol. 29, 73–79.
Heartworm Disease in Animals and Humans
267
Bazzocchi, C., Mortarino, M., Grandi, G., Kramer, L. H., Genchi, C., Bandi, C., Genchi, M., Sacchi, L., and McCall, J. W. Combined ivermectin-doxycycline treatment has microfilaricidal and adulticide activity against D. immitis in experimentally infected dogs. Intern. J. Parasitol., (in press). Beneth, G., Volansky, Z., Anug, Y., Favia, G., Bain, O., Golstein, R. E., and Harrus, S. (2002). Dirofilaria repens infection in a dog: Diagnosis and treatment with melarsomine and doramectin. Vet. Parasitol. 105, 173–178. Berndt, J. S., and Ware, W. (1987). An overview of Feline heartworm disease. Iowa State Univ. Vet. 49, 91–93. Bestul, K. J., McCall, J. W., Supakorndej, N., Ma, D., and Carlow, C. K. S. (1998). Evaluation of the AssureÒ /FH antibody assay for the detection of Feline heartworm infection. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 179–185. American Heartworm Society, Batavia, Illinois. Bidgood, A., and Collins, G. H. (1996). The prevalence of Dirofilaria immitis in dogs in Sydney. Aust. Vet. J. 73, 103–104. Bielawski, B. C., Harrington, D., and Joseph, E. (2001). A solitary pulmonary nodule with zoonotic implications. Chest 119, 1250–1252. Birago, F. (1626). ‘‘Trattato Cinegetico, Ovvero Della Caccia.’’ p. 77. V. Sfondato, Milano, Italy. Blagburn, B. L. (2001). New trends and technologies in heartworm prevention. URL:http: www.dvmnewsmagazine.com. Blair, L. S., and Campbell, W. C. (1978). Trial of avermectin B1a, mebendazole and melarsoprol against pre-cardiac Dirofilaria immitis in the ferret Mustela putorius furo. J. Parasitol. 64, 1032–1034. Bolio-Gonzalez, M. E., Rodriguez-Vivas, R. I., Sauri-Arceo, C. H., Gutierrez-Blanco, E., Ortega-Pacheco, A., and Colin-Flores, R. F. (2007). Prevalence of Dirofilaria immitis infection in dogs from Meridia, Yucatan, Mexico. Vet. Parasitol. 148, 166–169. Borgarelli, M., Venco, L., Piga, P. M., Bonino, F., and Ryan, W. G. (1997). Surgical removal of heartworms from the right atrium of a cat. J. Am. Vet. Med. Assoc. 211, 68–69. Brattig, N. W., Buttner, D. W., and Hoerauf, A. (2001). Neutrophil accumulation around Onchocerca worms and chemotaxis of neutrophils are dependent on Wolbachia endobacteria. Microbes Infect. 3, 439–446. Brattig, N. W., Bazzocchi, C., Kirschning, C. J., Reiling, N., Buttner, D. W., Ceciliani, F., Geisinger, F., Hochrein, H., Ernst, M., Wagner, H., Bandi, C., and Hoerauf, A. (2004). The major surface protein of Wolbachia endosymbionts in filarial nematodes elicits immune responses through TLR2 and TLR4. J. Immunol 73, 437–445. Brawner, W. R., Jr., Dillon, A. R., Robertson-Plouch, C. K., and Guerrero, J. (1998). Radiographic diagnosis of feline heartworm disease and correlation to other clinical criteria: Results of a multicenter clinical case study. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 91–95. American Heartworm Society, Batavia, Illinois. Bredal, W. P., Gjerde, B., Eberhard, M. L., Aleksandersen, M., Wilhelmsen, D. K., and Mansfield, L. S. (1998). Adult Dirofilaria repens in a subcutaneous granuloma on the chest of a dog. J. Small Anim. Pract. 39, 595–597. Browne, L. E., Carter, T. D., Levy, J. K., Snyder, P. S., and Johnson, C. M. (2005). Pulmonary arterial disease in cats seropositive for Dirofilaria immitis but lacking adult heartworms in the heart and lungs. Am. J. Vet. Res. 65, 1544–1549. Calvert, C. A., and Rawlings, C. A. (1988). Canine heartworm disease. In ‘‘Canine and Feline Cardiology’’ (P. R. Fox, ed.), pp. 519–549. Churchill Livingstone, Inc, New York. Calvert, C. A., Rawlings, C. A., and McCall, J. W. (1994). Feline heartworm disease. In ‘‘The Cat: Diseases and Clinical Management’’ (R. G. Sherding, ed.), 2nd edn., 2nd edn. Vol. 1, pp. 623–645. Churchill Livingstone, New York.
268
John W. McCall et al.
Campbell, W. C., and Blair, L. S. (1978). Dirofilaria immitis: Experimental infections in the ferret (Mustela putorius furo). J. Parasitol. 64, 119–122. Cancrini, G., and Kramer, L. H. (2001). Insect vectors of Dirofilaria spp. In ‘‘Heartworm Infection in Humans and Animals’’ (F. Simon and C. Genchi, eds.), pp. 63–82. University of Salamanca, Salamanca, Spain. Cancrini, G., Frangipane di Regalbono, A., Ricci, I., Tessarin, C., Gabrielli, S., and Pietrobelli, M. (2003). Aedes albopictus is a natural vector for Dirofilaria immitis in Italy. Vet. Parasitol. 118, 195–202. Cancrini, G., Magi, M. C., Gabrielli, S., Aruspici, M., Tolari, M., and Dell’Omodarme Prati, M. C. (2006). Natural vectors of dirofilariasis in rural and urban areas of the Tuscany region, central Italy. J. Med. Entomol. 43, 574–579. Casiraghi, M., Mazzocchi, C., Mortarino, M., Ottina, E., and Genchi, C. (2006). A simple molecular method for discrimination common filarial nematodes of dogs (Canis familiaris). Vet. Parasitol. 141, 368–372. Chalifoux, L., and Hunt, R. D. (1971). Histochemical differentiation of Dirofilaria immitis and Dipetalonema reconditum. J. Am. Vet. Med. Assoc. 158, 602–605. Clark, J. N., Pulliam, J. D., Alva, R., and Daurio, C. P. (1992). Safety of orally administered ivermectin in cats. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 103–109. American Heartworm Society, Batavia, IL. Comb, D., Koonin, E., and Slatko, B. (2005). The Wolbachia genome of Brugia malayi: Endosymbiont evolution within a human pathogenic nematode. Public Library of Science Biology 3, e121. Cordero, M., Mun˜oz, M. R., Muro, A., and Simo´n, F. (1990). Transient solitary pulmonary nodule caused by Dirofilaria immitis. Eur. Respir. J. 3, 1070–1071. Cordero, M., Mun˜oz, M. R., Muro, A., Simo´n, F., and Perera, M. L. (1992). Small calcified nodule: An undescribed radiologic manifestation of human pulmonary dirofilariasis. J. Infect. Dis. 165, 398–399. Cringoli, G., Rinald, L., Veneziano, V., and Capelli, G. (2001). Prevalence survey and risk analysis of filariosis in dogs from Mt. Vesuvio area of southern Italy. Vet. Parasitol. 102, 243–252. Cross, H. F., Haarbrink, M., Egerton, G., Yazdanbakhsh, M., and Taylor, M. J. (2001). Severe reactions to filarial chemotherapy and release of Wolbachia endosymbionts into blood. Lancet 358, 1873–1875. Crum, J. M., Nettles, V. F., and Davidson, W. R. (1978). Studies on endoparasites of the black bear (Ursus americanus) in the southeastern United States. J. Wildl. Dis. 14, 178–186. DeFrancesco, T. C., and Atkins, C. E. (1998). The utility of echocardiography in the diagnosis of feline heartworm disease: A review of published reports. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 103–106. American Heartworm Society, Batavia, Illinois. DeFrancesco, T. C., Atkins, C. E., Miller, M. W., Meurs, K. M., and Keene, B. W. (2001). Use of echocardiography for the diagnosis of heartworm disease in cats: 43 cases (1985–1997). J. Am. Vet. Med. Assoc. 218, 66–69. De Magalha˜es, P. S. (1887). Descric¸a˜o de una espe´cie de filarias encontradas no corac¸a˜o humano, precedida de una contribuic¸a˜o para o estudio de filariose de Wucherer e do respectivo parasito adulto, a Filo`aria brancofti (Cobbold) on Filaria sanguinis hominis (Lewis). Revista dos Cursos Pra´ticos e Teo´ricos da Facultad Medicina do Rio de Janeiro 3, 4193. Dillon, R. (1984). Feline dirofilariasis. Vet. Clin. North Am. Small Anim. Pract. 14, 1185–1199. Dillon, R. (1988). Feline heartworm disease. In ‘‘Dirofilariasis’’ (P. F. L. Boreham and R. B. Atwell, eds.), pp. 205–215. CRC Press, Boca Raton, FL. Dillon, R. (1998). Clinical significance of feline heartworm disease. Vet. Clin. North Am. Small Anim. Pract. 28, 1547–1565.
Heartworm Disease in Animals and Humans
269
Dillon, R. (1999). Feline heartworm disease. In ‘‘Textbook of Canine and Feline Cardiology’’ (P. R. Fox, D. Sisson, and N. S. Moı¨se, eds.), 2nd edn. pp. 727–735. W.B. Saunders, Philadelphia, PE. Dillon, A. R., Brawner, W. R., Grieve, R. B., Buxton-Smith, B., and Schultz, R. D. (1987). The chronic effects of experimental Dirofilaria immitis infection in cats. Semin. Vet. Med. Surg. (Small Anim.) 2, 72–77. Dillon, R., Brawner, W. R., and Hanrahan, L. (1995a). Influence of number of parasites and exercise on the severity of heartworm disease in dogs. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), p. 113. American Heartworm Society, Batavia, Illinois. Dillon, A. R., Warner, A. E., and Molina, R. M. (1995b). Pulmonary parenchymal changes in dogs and cats after experimental transplantation of dead Dirofilaria immits. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 97–101. American Heartworm Society, Batavia, Illinois. Dillon, A. R., Atkins, C., Clekis, T., Genchi, C., Knight, D. H., McCall, J. W., and Miller, M. W. (1996). Feline heartworm disease, Part 1. Feline Pract. 24, 12–16. Dillon, A. R., Atkins, C., Clekis, T., Genchi, C., Knight, D. H., McCall, J. W., and Miller, M. W. (1997a). Feline heartworm disease, Part 2. Feline Pract. 25, 26–30. Dillon, A. R., Atkins, C., Clekis, T., Genchi, C., Knight, D. H., McCall, J. W., and Miller, M. W. (1997b). Feline heartworm disease, Part 3. Feline Pract. 25, 12–21. Dillon, A. R., Brawner, W. R., Jr., Robertson-Plouch, C. K., and Guerrero, J. (1998). Feline heartworm disease: Correlations of clinical signs, serology, and other diagnostics— results of a multicenter study. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 153–157. American Heartworm Society, Batavia, Illinois. Dillon, A. R., Blagburn, B. L., Tillson, D. M., Brawner, W. R., Welles, B., Johnson, C., Spencer, J., Kaltenboeck, R., and Rynders, P. E. (2007). Immature heartworm infection produces pulmonary parenchymal, airway and vascular disease in cats. In ‘‘State of the Heartworm ’07 Symposium.’’ 12th Triennial Symposium of the American Heartworm Society held 13–14 July 2007 in Washington, DC, p. 24. American Heartworm Society, Batavia, Illinois. Dingman, P., Levy, J., Kramer, L., Lappin, M., and Johnson, C. (2007). Association of Wolbachia with heartworm disease in dogs and cats. In ‘‘State of the Heartworm ’07 Symposium.’’ 12th Triennial Symposium of the American Heartworm Society held 13–14 July 2007 in Washington, DC, p. 15. American Heartworm Society, Batavia, Illinois. Di Sacco, B., and Vezzoni, A. (1992). Clinical classification of heartworm disease for the purpose of adding objectivity to the assessment of therapeutic efficacy of adulticidal drugs in the field. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 209–214. American Heartworm Society, Batavia, Illinois. Donahoe, J. M. (1975). Experimental infection of cats with Dirofilaria immitis. J. Parasitol. 61, 599–605. Donahoe, J. M., Kneller, S. K., and Lewis, R. E. (1976a). In vivo pulmonary arteriography in cats infected with Dirofilaria immitis. J. Am. Vet. Radiol. Soc. 17, 147–151. Donahoe, J. M. R., Kneller, S. K., and Lewis, R. E. (1976b). Hematologic and radiographic changes in cats after inoculation with infective larvae of Dirofilaria immitis. J. Am. Vet. Med. Assoc. 168, 413–417. Donoghue, A. R., Piche´ Radecki, S. V., Schaad, L. E., and Frank, G. R. (1998). Effect of prophylaxis on antiheartworm antibody levels in cats receiving trickle experimental infections of Dirofilaria immitis. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 135–138. American Heartworm Society, Batavia, Illinois. Dzimianski, M. T., McTier, T. L., McCall, J. W., and Raynaud, J. P. (1989). Assessment of filaricidal activity of a new filaricide (RM 340) against immature and adult heartworms
270
John W. McCall et al.
using experimental canine models. In ‘‘Proceedings of the Heartworm Symposium ’89’’ (G. H. Otto, ed.), pp. 147–153. American Heartworm Society, Batavia, Illinois. Espinoza, E., Cordero, M., Muro, A., Lorente, F., and Simo´n, F. (1993). Anti-Dirofilaria immitis IgE: Seroepidemiology and seasonal variation in an exposed human population. Trop. Med. Parasitol. 44, 172–176. Favia, G., Lanfrancotti, A., Della Torre, A., Cancrini, G., and Coluzzi, M. (1996). Polymerase chain reaction-identification of Dirofilaria repens and Dirofilaria immitis. Parasitology 113, 567–571. Favia, G., Cancrini, G., Ricci, I., Mazzocchi, C., Magi, M., Pietrobelli, M., Genchi, C., and Bandi, C. (2000). 5S ribosomal spacer sequences of some filarial parasites: Comparative analysis and diagnostic implications. Mol. Cell. Probes 14, 285–290. FEDD (2007). ‘‘First European Dirofilaria Days.’’ Abstract Book, p. 38. Zagreb, Croatia. Fenn, K., and Blaxter, M. (2004). Quantification of Wolbachia bacteria in Brugia malayi through the nematode lifecycle. Mol. Biochem. Parasitol. 137, 361–364. Fok, E. (2007). The importance of dirofilariosis in carnivores and humans in Hungary, past and present. In ‘‘Dirofilaria immitis and Dirofilaria repens in dog and cat and human infections’’ (C. Genchi, L. Rinaldi, and G. Cringoli, eds.), pp. 181–188. Rolando Editore, Napoli, Italy. Fortin, J. F., and Slocombe, J. O. D. (1981). Temperature requirements for the development of Dirofilaria immitis in Aedes triseriatus and Ae. vexans. Mosq. News 41, 625–633. Foster, J., Ganatra, M., Kamal, I., Ware, J., Makarova, K., Ivanova, N., Bhattacharyya, A., Kapatral, V., Kumar, S., Posfai, J., Vincze, T., Ingram, J., et al. (2005). The Wolbachia genome of Brugia malayi: Endosymbiont evolution within a human pathogenic nematode. Public Library of Science Biology 3, e121. Fox, J. C., Wagner, R., and Crotty, C. (1998). Development of a PCR test and an IFA test for Dirofilaria immits in cats and their use along with an ELISA heartworm antigen test to determine the prevalence of infections in the Tulsa, Oklahoma Area. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 167–171. American Heartworm Society, Batavia, Illinois. Frank, G. R., and Grieve, R. B. (1991). Metabolic labelling of Dirofilaria immitis third- and fourth-stage larvae and their excretory-secretory products. J. Parasitol. 77, 950–956. Frank, G. R., and Grieve, R. B. (1996). Purification and characterization of three larval excretory-secretory proteins of Dirofilaria immitis. Mol. Biochem. Parasitol. 75, 221–229. Frank, G. R., Tripp, C. A., and Grieve, R. B. (1996). Molecular cloning of a developmentally regulated protein isolated from excretory-secretory products of larval Dirofilaria immitis. Mol. Biochem. Parasitol. 75, 231–240. Frank, G. R., Sabin, E. A., and Chandrashekar, R. (1998a). Heartworm vaccine and immunology research. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 247–256. American Heartworm Society, Batavia, Illinois. Frank, G. R., Mondesire, R. R., Brandt, K. S., and Wisnewski, N. (1998b). Antibody to the Dirofilaria immitis aspartyl protease inhibitor homologue is a diagnostic marker for feline heartworm infections. J. Parasitol. 84, 1231–1236. Furlanello, T., Caldin, A., Vezzoni, A., Venco, L., and Kitagawa, H. (1998). Patogenesi. In ‘‘La Filariosi cardiopolmonare del cane e del gatto’’ (C. Genchi, L. Venco, and A. Vezzoni, eds.), pp. 31–46. Societa´ Culturale Italiana Veterinari per Animali da Campagnia, Cremona, Italy. Garcia, R., and Voigt, W. G. (1989). Coyotes as a reservoir for canine heartworm infection in California. In ‘‘Proceedings of Heartworm Symposium ’89’’ (G. F. Otto, ed.), pp. 7–12. American Heartworm Society, Washington, DC. Genchi, C., Traldi, G., Di Sacco, B., and Benedetti, M. C. (1988). Epidemiological aspects of canine heartworm disease in Italy. In ‘‘Atti del IV Seminario: Filariosi,’’ pp. 53–64. Societa´ Culturale Italiana Veterinari per Animali da Campagnia, Cremona, Italy.
Heartworm Disease in Animals and Humans
271
Genchi, C., Di Sacco, B., and Cancrini, G. (1992a). Epizootiology of canine and feline heartworm infection in Northern Italy: Possible mosquito vectors. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 39–46. American Heartworm Society, Batavia, Illinois. Genchi, C., Guerrero, J., Di Sacco, B., and Formaggini, L. (1992b). Prevalence of Dirofilaria immitis infection in Italian cats. In ‘‘Proceedings of Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 97–102. American Heartworm Society, Batavia, Illinois. Genchi, C., Venco, L., Magnino, S., Di Sacco, B., Perera, L., Bandi, C., Pignatelli, P., Formaggini, L., and Mazzucchelli, M. (1993). Aggiornamento epidemiologico sulla filariosi del cane e del gatto. Veterinaria 7, 5–11. Genchi, C., Solari, B. F., Bandi, C., Di Sacco, B., Venco, L., and Vezzoni, A. (1995). Factors influencing the spread of heartworm in Italy. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 65–71. American Heartworm Society, Batavia, Illinois. Genchi, C., Solari Basano, F., Marrone, R. V., and Petruschke, G. (1998a). Canine and feline heartworm in Europe with special emphasis on Italy. In ‘‘Proceedings of the Heartworm Symposium ’98’’ (R. L. Seward and D. H. Knight, eds.), pp. 75–82. American Heartworm Society, Batavia IL. Genchi, C., Kramer, L., Venco, L., Prieto, G., and Simo´n, F. (1998b). Comparison of antibody and antigen testing with echocardiography for the detection of heartworm (Dirofilaria immitis) in cats. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 173–177. American Heartworm Society, Batavia, Illinois. Genchi, C., Kramer, L. H., and Prieto, G. (2001a). Epidemiology of canine and feline dirofilariosis: A global view. In ‘‘Heartworm Infection in Humans and Animals’’ (F. Simo´n and C. Genchi, eds.), pp. 121–133. Ediciones Universidad de Salamanca, Salamanca, Spain. Genchi, C., Simoncini, L., McCall, J. W., Venco, L., Lo, N., Bazzocchi, C., Casiraghi, M., Kramer, L., and Bandi, C. (2001b). The Wolbachia endosymbionts of Dirofilaria immitis: Implications for pathology, immunology and control. In ‘‘Recent Advances in Heartworm Disease: Symposium ’01’’ (R. L. Seward, ed.), pp. 27–33. American Heartworm Society, Batavia, Illinois. Genchi, C., Poglayen, G., and Kramer, L. (2002a). Efficacia di selamectin nella profilassi delle infestazioni da Dirofilaria repens nel cane. Veterinaria 16, 69–71. Genchi, C., Rossi, L., Cardini, G., Kramer, L. H., Venco, L., Casiraghi, M., Genchi, M., and Agostini, A. (2002b). Full season efficacy of moxidectin microsphere sustained release formulation for the prevention of heartworm (Dirofilaria immitis) infection in dogs. Vet. Parasitol. 110, 85–91. Genchi, C., Rinaldi, L., Cascone, C., Mortarino, M., and Cringoli, G. (2005). Is heartworm really spreading in Europe? Vet. Parasitol. 133, 137–148. Genchi, C., Guerrero, J., McCall, J. W., and Venco, L. (2007). Epidemiology and prevention of Dirofilaria infections in dogs and cats. In ‘‘Dirofilaria immitis and D. repens in Dog and Cat and Human Infections’’ (C. Genchi, L. Rinaldi, and G. Cringoli, eds.) pp. 147–161. Rinaldo Editore, Napoli, Italy. Genchi, C., Venco, L., Ferrari, N., Mortarino, M., and Genchi, M. Feline heartworm (Dirofilaria immitis) infection: A statistical elaboration of the duration of the infection and life expectancy in asymptomatic cats. Vet. Parasitol. (in press). Giannetto, S., Pampiglione, S., Santoro, V., and Virga, A. (1997). Research of canine filariosis in Trapani province (Western Sicily). Parassitologa 39, 403–405. Giannetto, S., Poglayen, G., Gaglio, G., and Brianti, E. (2007). Prevalence and epidemiological aspects of microfilaraemia in dogs in Sicily. In ‘‘FEDD, First European Dirofilaria Days.’’ p. 5. Zagreb, Croatia. Gilbert, J., Nfon, C. K., Makepeace, B. L., Njongmeta, L. M., Hastings, I. M., Pfarr, K. M., Renz, A., Tanya, V. N., and Trees, A. J. (2005). Antibiotic chemotherapy of onchocerciasis:
272
John W. McCall et al.
In a bovine model, killing of adult parasites requires a sustained depletion of endosymbiotic bacteria (Wolbachia species). J. Infect. Dis. 192, 1483–1493. Glaus, T. M., Jacobs, G. J., Rawlings, C. A., Watson, E. D., and Calvert, C. A. (1995). Surgical removal of heartworms from a cat with caval syndrome. J. Am. Vet. Med. Assoc. 206, 663–666. Gomes, L. A. M., Serra˜o, M. L., Dantas, C., and Labarthe, N. (2001). Estudo da dirofilariose em gatos no Engenho do Mato, regia˜o oceaˆnica de Nitero´i, R.J., Brasil. Revista Brasileira de Cieˆncia Veterina´ria 8, 160–162. Gonzales, J. A. (2002). Seroprevalencia de la dirofilariosis y ehrlichiosis canina en los distritos de Chorrillos, La Molina y San Juan de Miraflores. Thesis. Universidad Nacional Mayor de San Marcos p. 98. Lima, Peru. Goodman, D. A. (1996). Evaluation of a single dose of melarsomine dihydrochloride for adulticidal activity against Dirofilaria immitis in cats. In ‘‘Thesis for Master of Science Degree.’’ p. 95. University of Georgia, Athens, Georgia. Gorta´zar, C., Castillo, J. A., Lucientes, J., Blanco, J. C., Arriolabengoa, A., and Calvete, C. (1994). Factors affecting Dirofilaria immitis prevalence in red foxes in Northeastern Spain. J. Wildl. Dis. 30, 545–547. Gorta´zar, C., Villafuente, R., Lucientes, J., and Ferna´ndez-de-Luco, D. (1998). Habitat related differences in helminth parasites of red foxes in the Ebro valley. Vet. Parasitol. 80, 75–81. Grandi, G., Kramer, L. H., Bazzocchi, C., Mortarino, M., Venco, L., and Genchi, C. (2005). Dirofilaria immitis and the endosymbiont Wolbachia: Inflammation and immune response in heartworm disease. Jpn. J. Vet. Parasitol. 4, 11–16. Gratz, N. G. (2004). Critical review of the vector status of Aedes albopictus. Med. Vet. Entomol. 18, 215–237. Grieve, R. B., Abraham, D., Mika-Grieve, M., and Seibert, B. P. (1988). Induction of protective immunity in dogs to infection with Dirofilaria immitis using chemically abbreviated infections. Am. J. Trop. Med. Hyg. 39, 373–379. Grieve, R. B., Frank, G. R., Mika-Grieve, M., Culpepper, J. A., and Mok, M. (1992). Identification of Dirofilaria immitis larval antigens with immunoprophylactic potential using sera from immune dogs. J. Immunol. 148, 2511–2515. Guerrero, J., McCall, J. W., Dzimianski, M. T., McTier, T. L., Holmes, R. A., and Newcomb, K. M. (1992a). Prevalence of D. immitis infection in cats from the southeastern United States. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 91–95. American Heartworm Society, Batavia, Illinois. Guerrero, J., Ducos de la Hitte, J., Genchi, C., Rojo, F., Gomez-Bautista, M., Carvalho Valera, M., Labarthe, N., Bordin, E., Gonzales, G., Mancebo, O., Patino, F., Uribe, L. F., et al. (1992b). Update on the distribution of Dirofilaria immitis in dogs from southern Europe and Latin America. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 31–37. American Heartworm Society, Batavia, Illinois. Guerrero, J., McCall, J. W., and Genchi, C. (2002). The use of macrocyclic lactones in the control and prevention of heartworm and other parasites in dogs and cats. In ‘‘Macrocyclic Lactones in Antiparasitic Therapy’’ ( J. Vercruysse and R. S. Rew, eds.), pp. 353–369. CABI Publishing, Oxon, UK. Guerrero, J., Nelson, C. T., and Carithers, D. (2006). Results and realistic implications of the 2004 AHS-Merial heartworm survey. In ‘‘Proceedings of the 51st Annual Meeting of the American Association of Veterinary Parasitologists.’’ pp. 62–63. Honolulu, Hawaii. Hargis, A. M., Lewis, T. P., Duclos, L. D., Loeffler, D. G., and Rausch, R. L. (1999). Dermatitis associated with microfilariae (Filarioidea) in 10 dogs. Vet. Dermatol. 10, 95–107. Hayasaki, M. (2001). Immunological analysis of agglutination in Dirofilaria immitis microfilariae. Jpn. Vet. Med. Sci. 63, 903–907. Henckler, S. (1998). Feline heartworm awareness: The first step toward diagnosis. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 107–108. American Heartworm Society, Batavia, Illinois.
Heartworm Disease in Animals and Humans
273
Hise, A. G., Gillette-Ferguson, I., and Pearlman, E. (2004). The role of endosymbiotic Wolbachia bacteria in filarial disease. Cell Microbiol. 6, 97–104. Holmes, R. A., Clark, J. N., Casey, H. W., Henk, W., and Plue, R. E. (1992). Histopathologic and radiographic studies of the development of heartworm pulmonary vascular disease in experimentally infected cats. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 81–89. American Heartworm Society, Batavia, Illinois. Ishihara, K., Kitagawa, H., Ojima, M., Yagata, Y., and Suganuma, Y. (1978). Clinicopathological studies on canine dirofilarial hemoglobinuria. J. Vet. Sci. 40, 525–530. Ishihara, K., Kitagawa, H., and Sasaky, Y. (1990). Efficacy of heartworm removal in dogs with dirofilarial hemoglobinuria using Flexible Alligator Forceps. Jpn. J. Vet. Sci. 53, 591–599. Jackson, R. F. (1975). The venae cavae syndrome. In ‘‘Proceedings of the Heartworm Symposium ’74’’ (H. C. Morgan, ed.), pp. 48–50. Veterinary Medicine Publishing, Bonner Springs, Kansas. Jafari, S., Gaur, S. N. S., and Khaksar, Z. (1996). Prevalence of Dirofilaria immitis in dogs of Fars province of Iran. J. Appl. Anim. Res. 9, 27–31. Jelinek, T., Schulte-Hillen, J., and Loscher, T. (1996). Human dirofilariasis. Int. J. Dermatol. 35, 872–875. Keister, D. M., Dzimianski, M. T., McTier, T. L., McCall, J. W., and Brown, J. (1992). Dose selection and confirmation of RM 340, a new filaricide for the treatment of dogs with immature and mature Dirofilaria immitis. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 225–229. American Heartworm Society, Batavia, Illinois. Kemmerer, D. W. (1998). Heartworm disease in the domestic ferret. In ‘‘Recent Advances in Heartworm Disease, Symposium ’98’’ (R. L. Seward, ed.), pp. 87–89. American Heartworm Society, Batavia, Illinois. Kennedy, S., and Patton, S. (1981). Heartworms in a Bengal tiger (Panthera tigris). J. Zoo Anim. Med. 12, 20–22. Kendall, K., Collins, G. H., and Pope, S. E. (1991). Dirofilaria immitis in cats from inner Sydney. Aust. Vet. J. 68, 356–357. Kim, M. K., Kim, C. H., Yeom, B. W., Park, S. H., Choi, S. Y., and Choi, J. S. (2002). The first human case of hepatic dirofilariasis. J. Korean Med. Sci. 17, 686–690. Kirkova, Z., Ivanov, A., and Georgieva, D. (2007). Dirofilariosis in dogs and wild carnivores in Bulgaria. In ‘‘FEDD, First European Dirofilaria Days.’’ Abstract Book, P. S. Zagret, Crcatia . Kitagawa, H., Sasaki, Y., and Ishihara, K. (1986). Clinical studies on canine dirofilarial hemoglobinuria: Relationship between the heartworm mass at the tricuspid valve orifice and the plasma hemoglobin concentration. Jpn. J. Vet. Sci. 48, 99–103. Kitagawa, H., Sasaky, Y., and Ishiara, K. (1987). Canine dirofilaria hemoglobinuria: Changes in right heart hemodynamics and heartworm migration from pulmonary artery toward right atrium following beta-blocker administration. Nippon Juigaku Zasshi 49, 1081–1086. Klotins, K. C., Wayne Martin, S., Bonnett, B. N., and Peregrine, A. S. (2000). Canine heartworm testing in Canada: Are we being effective. Can. Vet. J. 41, 929–937. Knight, D. H. (1987). Heartworm infection. Vet. Clin. North Am. Small Anim. Pract. 17, 1463–1518. Knight, D. H. (1995). Guidelines for diagnosis and management of heartworm (Dirofilaria immitis) infection. In ‘‘Kirk’s Current Veterinary Therapy XII, Small Animal Practice’’ ( J. D. Bonagura, ed.), pp. 879–887. WB Saunders Co, Philadelphia, Pennsylvania. Knight, D. H., and Lok, J. B. (1995). Seasonal timing of heartworm chemoprophylaxis in the United States. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 37–42. American Heartworm Society, Batavia, Illinois. Knight, D. H., Doiron, D. W., Longhofer, S. L., Rubin, S. B., Downing, R., Nelson, C. T., McCall, J. W., and Seward, R. L. (2001). 2002 Guidelines for the diagnosis, prevention and management of heartworm (Dirofilaria immitis) infection in cats. In ‘‘Recent Advances
274
John W. McCall et al.
in Heartworm Disease: Symposium ’01’’ (R. L. Seward, ed.), pp. 267–273. American Heartworm Society, Batavia, Illinois. Knott, J. (1939). A method for making microfilarial survey on day blood. Trans. R. Soc. Med. Hyg. 33, 191–196. Kotani, T., and Powers, K. G. (1982). Developmental stages of Dirofilaria immitis in the dog. Am. J. Vet. Res. 43, 2199–2206. Kozek, W. J. (2005). What is new in the Wolbachia/Dirofilaria interactions? Vet. Parasitol. 133, 127–132. Kozek, W. J., Vazquez, A. E., Gonzalez, C., Iguina, J., Sanchez, E., de Jesus, F., Cardona, C. J., Gomez, C., Seneriz, R., and Diaz-Umpierre, J. (1995). Prevalence of canine filariae in Puerto Rico and the Caribbean. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 55–64. American Heartworm Society, Batavia, Illinois. Kramer, L., and Genchi, C. (2002). Feline heartworm infection: Serological survey of asymptomatic cats living in northern Italy. Vet. Parasitol. 104, 43–50. Kramer, L. H., Passeri, B., Corona, S., Simoncini, L., and Casiraghi, M. (2003). Immunohistochemical/immunogold detection and distribution of the endosymbiont Wolbachia of D. immitis and Brugia pahangi using a polyclonal antiserum raised against WSP (Wolbachia surface protein). Parasitol. Res. 89, 381–386. Kramer, L. H., Tamarozzi, F., Morchon, R., Lopez-Belmonte, J., Marcos- Atxutegi, C., MartinPacho, R., and Simon, F. (2005). Immune response to and tissue localization of the Wolbachia surface protein (WSP) in dogs with natural heartworm (Dirofilaria immitis) infection. Vet. Immunol. Immunopathol. 106, 303–308. Kramer, L. H., McCall, J. W., Grandi, G., and Genchi, C. (2007). Wolbachia and its importance in veterinary filariasis. Issues Infect. Dis. 5, 124–132. Kramer, L., Grandi, G., Leoni, M., Passeri, B., McCall, J., Genchi, C., Mortarino, M., and Bazzocchi, C. Wolbachia and its influence on the pathology and immunology of Dirofilaria immitis infection. Vet. Parasitol. (in press). Kume, S., and Itagaki, S. (1955). On the life-cycle of Dirofilaria immitis in the dog as the final host. Br. Vet. J. 111, 16–24. Kuo, T. F., Yang, C. Y., and Yau, C. F. (1995). Study on the incidence of pound dogs infested with heartworms and a comparison of different methods of detection during the summer season in Taipei. J. Chinese Soc. Vet. Sci. 21, 97–104. Labarthe, N. V. (1997). Dirofilariose canina: Diagno´stico, prevenc¸a˜o e tratamento adulticida: Revisao de literatura. Clin. Vet. 10, 10–16. Labarthe, N., and Guerrero, J. (2005). Epidemiology of heartworm: What is happening in South America and Mexico. Vet. Parasitol. 133, 149–156. Labarthe, N. V., Almosny, N. R., Soares, A. M., and Souza-Silva, L. C. (1992). Update on the distribution of Dirofilaria immitis in the State of Rio de Janeiro, Brazil. In ‘‘Proceedings of the XVII World Small Animal Veterinary Association World Congress.’’ pp. 291–293. Rome, Italy. Labarthe, N., Almosny, N., Guerrero, J., and Duque-Arau´jo, A. M. (1997a). Description of the occurrence of canine dirofilariasis in the State of Rio de Janeiro, Brazil. Memo´rias do Instituto Oswaldo Cruz 92, 47–51. Labarthe, N., Ferreira, A. M. R., Guerrero, J., Newcomb, K., and Paes-de-Almeida, E. (1997b). Survey of Dirofilaria immitis (Leidy, 1856) in random source cats in metropolitan Rio de Janeiro, Brazil, with descriptions of lesions. Vet. Parasitol. 71, 301–306. Langworthy, N., Renz, A., Meckenstedr, U., Henkle-Duhrsen, K., Bronvoort, M., Tanya, V., Donnelly, M., and Trees, A. J. (2000). Macrofilaricidal activity of tetracycline against the filarial nematode Onchocerca ochengi: Elimination of Wolbachia precedes worm death and suggests a dependent relationship. Proc. R. Soc. Lond. B Biol. Sci. 267, 1063–1069. Lee, J. C., Lee, C. Y., Shin, S. S., and Lee, C. G. (1996). A survey of canine heartworm infections among German shepherds in South Korea. Korean J. Parasitol. 34, 225–231.
Heartworm Disease in Animals and Humans
275
Lee, S. E. (2003). The survey of canine dirofilariasis and differential diagnosis of microfilaria. In ‘‘Thesis for Degree of Master of Veterinary Medicine’’ p. 27. Chungnam National University, South Korea. Lee, S.-E., Heung-Chul, K., Sung-Tae, C., Klein, T. A., and Wong-Ja, L. (2007). Molecular survey of Dirofilaria immitis and Dirofilaria repens in the Republic of Korea. Vet. Parasitol. 148, 149–155. Levy, J. (2007). Diagnostic changes in feline heartworm disease. In ‘‘State of the Heartworm ’07 Symposium.’’ 12th Triennial Symposium of the American Heartworm Society held 13–14 July 2007 in Washington, DC, Abstract Book, pp. 30. American Heartworm Society, Batavia, Illinois. Levy, J. K., Snyder, P. S., Tavares, L. M., Hooks, J. L., Pegelow, M. J., Slater, M. R., Hughes, K. L., and Salute, M. E. (2003). Prevalence and risk factors for heartworm infection in cats from Northern Florida. J. Am. Anim. Hosp. Assoc. 39, 533–537. Levy, J. K., Edinboro, C. H., Glotfelty, C.-S., Dingman, P. A., West, A. L., and KirklandCady, K. D. (2007). Seroprevalence of Dirofilaria immitis, feline leukemia virus, and feline immunodeficiency virus infection among dogs and cats exported from the 2005 Gulf Coast hurricane disaster area. J. Am. Vet. Med. Assoc. 231, 218–225. Lichtenfels, J. R., Pilitt, P. A., Kotani, T., and Powers, K. G. (1985). Morphogenesis of developmental stages of Dirofilaria immitis (Nematoda) in dogs. Proceedings of the Helminthological Society of Washington 52, 98–102. Litster, A., Atkins, C., Atwell, R., and Buchanan, J. (2005). Radiographic cardiac size in cats and dogs with heartworm disease compared with the reference values using the vertebral heart scale methol: 53 cases. J. Vet. Cardiol. 7, 33–40. Litster, A., Atkins, C., and Atwell, R. Acute death in heartworm-infected cats: Unraveling the puzzle. Vet. Parasitol. (in press). Liu, S. K., Das, K. M., and Tashjian, R. J. (1966). Adult Dirofilaria immitis in the arterial system of a dog. J. Am. Vet. Med. Assoc. 148, 1501–1507. Liu, J., Song, K. H., Lee, S. E., Lee, J. I., Haysaki, M., You, M. J., and Kim, D. H. (2005). Serological and molecular survey of Dirofilaria immitis infection in cats in Gyunggi province, South Korea. Vet. Parasitol. 130, 125–129. Lok, J. B. (1988). Dirofilaria spp.: Taxonomy and distribution. In ‘‘Dirofilariasis’’ (P. F. L. Boreham and R. B. Atwell, eds.), pp. 1–28. CRC Press, Inc., Boca Raton, Florida. Lok, J. B., Knight, D. H., Nolan, T. J., Grubbs, S. T., Cleale, R. M., and Heaney, K. (2005). Efficacy of an injectable, sustained release formulation of moxidectin in preventing experimental heartworm infection in mongrel dogs challenged 12 months after administration. Vet. Parasitol. 128, 129–135. Longhofer, S. L., Daurio, C. P., Plue, R. E., Alva, R., Wallace, D. H., and Roncalli, R. A. (1995). Ivermectin for the prevention of feline heartworm disease. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Kinght, eds.), pp. 177–182. American Heartworm Society, Batavia, Illinois. Lorentzen, L., and Caola, A. Incidence of positive heartworm antibody and antigen tests at IDEXX Laboratories: Trends and potential impact on feline heartworm awareness and prevention. Vet. Parasitol. (in press). Luder, C. G., Schulz-Key, H., Banla, M., Pritze, S., and Sobaslay, P. T. (1996). Dirofilaria immitis infection in dogs. Vet. Parasitol. 133, 255–266. Magi, M., Calderini, P., Gabrielli, S., Dell’Omodarme, M., Macchioni, F., Prati, M. C., and Cancrini, G. (2007). An update of filarial parasites in Vulpes vulpes of Tuscany (Central Italy). In ‘‘FEDD, First European Dirofilaria Days.’’ Abstract Book, pp. 9. Zagreb, Croatia. Man˜as, S., Ferrer, D., Castella, J., and Lopez-Martin, J. M. (2005). Cardiopulmonary helminth parasites of red fox (Vulpes vulpes) in Catalonia, northeastern Spain. Vet. J. 169, 118–120.
276
John W. McCall et al.
Mancebo, O., Russo, A., Bulman, G., Cabajal, L., and Villavicencio de Mancebo, V. (1992). Dirofilaria immitis: Caracteristicas, prevalencia y diagno´stic de la dirofilariasis en la poblacio´n canina en a´reas urbanas, suburbanas, y rurales de la Prov. De Formosa (Argentina) y descriptio´n de la enfermedade en el coati comun (Nasua solitaria). Pet’s 8, 95–117. Mandelli, G., and Mantovani, A. (1966). Su di un caso di infestazione massiva da Dirofilaria repens nel cane. Parassitologia 8, 21–28. Manfredi, M. T., Vieira, C., Bandi, C., Casiraghi, M., and Simon, F. (2001). Phylogeny, systematic and structural aspects. In ‘‘Heartworm Infection in Humans and Animals’’ (F. Simon and C. Genchi, eds.), pp. 19–40. University of Salamanca, Salamanca, Spain. Mansour, A. E., McCall, J. W., McTier, T. L., Supakorndej, N., and Ricketts, R. R. (1995). Epidemiology of feline dirofilariasis: Infections induced by simulated natural exposure to Aedes aegypti experimentally infected with heartworms. In ‘‘Proceedings of Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 87–95. American Heartworm Society, Batavia, Illinois. Mar, P. H., Yang, I. C., Chang, G. N., and Dei, A. C. (2002). Specific polymerase chain reaction for differential diagnosis of Dirofilaria immitis and Dipetalonema reconditum using primers derived from internal transcribed spacer region 2 (ITS2). Vet. Parasitol. 106, 243–252. Marconcini, G., Magi, M., and Hecht Contin, B. (1993). Sulia validita` dell’inevrmectina nella profilassi dell’infestazione da con Dirofilaria repens in cani naturalmente esposti al contagio. Parassitologia 35, 67–71. Matola, Y. G. (1991). Periodicity of Dirofilaria immitis microfilariae in a dog from Muheza district. Tanzania J. Helminthol. 65, 76–78. McCall, J. W. (1981). The role of arthropods in the development of animal models for filariasis. J. Ga. Entomol. Soc. 16, 283–293. McCall, J. W. (1992). A parallel between experimentally induced canine and feline heartworm disease. In ‘‘Proceedings of XVII World Small Animal Veterinary Association World Congress, 24th–27th September 1992.’’ pp. 255–261. Delfino Publisher, Rome. McCall, J. W. (1998). Dirofilarisis in the domestic ferret. Clin. Tech. Small Anim. Pract. 13, 109–112. McCall, J. W. (2001). Experimental infections in animal models. In ‘‘Heartworm Infection in Humans and Animals’’ (F. Simon and C. Genchi, eds.), pp. 147–159. University of Salamanca, Salamanca, Spain. McCall, J. W. (2005). The safety-net story about macrocyclic lactone heartworm preventatives: A review, an update and recommendations. Vet. Parasitol. 133, 197–206. McCall, J. W., Genchi, C., Kramer, L., Guerrero, J., Dzimianski, M. T., Supakorndej, P., Mansour, A. M., McCall, S. D., Supakorndej, N., Grandi, G., and Carson, B. Heartworm and Wolbachia: Therapeutic implications. Vet. Parasitol. (in press). McCall, J. W., Dzimianski, M. T., McTier, T. L., Jernigan, A. D., Jun, J. J., Mansour, A. E., Supakorndej, P., Plue, R. E., Clark, J. N., Wallace, D. H., and Lewis, R. E. (1992). Biology of experimental heartworm infections in cats. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 71–79. American Heartworm Society, Batavia, Illinois. McCall, J. W., Calvert, C. A., and Rawlings, C. A. (1994). Heartworm infection in cats: A life-threatening disease. Vet. Med. 89, 639–648. McCall, J. W., Supakorndej, N., Ryan, W., and Soll, M. D. (1995). Utility of an ELISA-based antibody test for detection of heartworm infection in cats. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 127–133. American Heartworm Society, Batavia, Illinois. McCall, J. W., Guerrero, J., Supakorndej, P., Mansour, A. E., and Lumpkin, M. (1998). Evaluation of the accuracy of heartworm antigen and antibody tests for cats. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 127–133. American Heartworm Society, Batavia, Illinois.
Heartworm Disease in Animals and Humans
277
McCall, J. W., Guerrero, J., Roberts, R. E., Supakorndej, N., Mansour, A. E., Dzimianski, M. T., and McCall, S. D. (2001a). Further evidence of clinical prophylactic (reach-back) and adulticidal activity of monthly administration of ivermectin and pyrantel pamoate (Heartgard Plus) in dogs experimentally infected with heartworms. In ‘‘Recent Advances in Heartworm Disease, Symposium ’01’’ (R. L. Seward, ed.), pp. 189–200. American Heartworm Society, Batavia, Illinois. McCall, J. W., Supakorndej, N., McCall, S. D., and Mansour, A. E. (2001b). Evaluation of feline heartworm antibody test kits and diagnostic laboratory tests. In ‘‘Recent Advances in Heartworm Disease: Symposium ’01’’ (R. L. Seward, ed.), pp. 125–133. American Heartworm Society, Batavia, Illinois. McCall, J. W., Supakorndej, N., Donoghue, A. R., Turnbull, R. K., and Radecki, S. V. (2001c). Evaluation of the performance of canine heartworm antigen test kits licensed for use by veterinarians and canine heartworm antigen tests conducted by diagnostic laboratories. In ‘‘Recent Advances in Heartworm Disease: Symposium ’01’’ (R. L. Seward, ed.), pp. 97–104. American Heartworm Society, Batavia, Illinois. McGarry, H. F., Egerton, G., and Taylor, M. J. (2004). Population dynamics of Wolbachia bacterial endosymbionts in Brugia malayi. Mol. Biochem. Parasitol. 135, 57–67. McGonigle, S., Yoho, E. R., and James, E. R. (2001). Immunization of mice with fractions derived from intestine of Dirofilaria immitis. Int. J. Parasitol. 31, 1459–1466. McGreevy, P. B., Theis, J. H., Lavoipierre, M. M. J., and Clark, J. (1974). Studies on filariasis. III. Dirofilaria immitis: Emergence of infective larvae from the mouthparts of Aedes aegypti. J. Helminthol. 48, 221–224. McLeroy, L. W. (1998). Evaluation of melarsomine dihydrochloride for adulticidal activity against Dirofilaria immitis in cats with intravenously transplanted adult heartworms. In ‘‘Thesis for Master of Science Degree.’’ p. 95. University of Georgia, Athens, Georgia. McTier, T. L., McCall, J. W., Dzimianski, M. T., Raynaud, J. P., Holmes, R. A., and Keister, D. M. (1992a). Epidemiology of heartworm infection in beagles naturally exposed to infection in three southeastern states. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 47–57. American Heartworm Society, Batavia, Illinois. McTier, T. L., McCall, J. W., Dzimianski, M. T., Mansour, A. E., Jernigan, A., Clark, J. N., Plue, R. E., and Daurio, C. P. (1992b). Prevention of heartworm infection in cats by treatment with ivermectin at one month post-infection. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. S. Soll, ed.), pp. 111–116. American Heartworm Society, Batavia, Illinois. McTier, T. L., McCall, J. W., Jenigan, A. D., rowan, T. G., Giles, C. J., Bishop, B. F., Evans, N. A., and Bruce, C. I. (1998). UK-124,114, a novel avermectin for the prevention of heartworms in dogs and cats. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 187–192. American Heartworm Society, Batavia, Illinois. Medlock, J. M., Barrass, I., Kerrod, E., Taylor, M. A., and Leach, S. (2007). Analysis of climatic predictions for extrinsic incubation of Dirofilaria in the United Kingdom. Vector-borne Zoonotic Dis. 7, 4–14. Mejia, J. S., and Carlow, C. K. (1994). An analysis of the humoral immune response of dogs following vaccination with irradiated infective larvae of Dirofilaria immitis. Parasite Immunol. 16, 157–164. Milanez de Campos, J. R., Barbas, C. S., Filomeno, L. T., Minamoto, H., Filho, J. V., and Jatene, F. B. (1997). Human pulmonary dirofilariasis: Analysis of 24 cases from Sao Paulo, Brazil. Chest 112, 729–733. Miller, M. W., Atkins, C. E., Stemme, K., Robertson-Plouch, C., and Guerrero, J. (1998). Prevalence of exposure to Dirofilaria immitis in cats in multiple areas of the United States. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 161–166. American Heartworm Society, Batavia, Illinois.
278
John W. McCall et al.
Miyoshi, T., Tsubouchi, H., Iwasaki, A., Shiraishi, T., Nabeshima, K., and Shirakusa, T. (2006). Human pulmonary dirofilariasis: A case report and review of the recent Japanese literature. Respirology 11, 343–347. Moise, N. S. (1988). Echocardiography. In ‘‘Canine and Feline Cardiology’’ (P. R. Fox, ed.), pp. 113–156. Churchchill Livingstone, Inc., New York. Montoya, J. A., Morales, M., Ferrer, O., Molina, J. M., and Corbera, J. A. (1998). The prevalence of Dirofilaria immitis in Gran Canaria, Canary Islands, Spain (1994–1996). Vet. Parasitol. 75, 221–226. Montoya, J. A., Morales, M., Juste, M. C., Ban˜ares, A., Simon, F., and Genchi, C. (2006). Seroprevalence of canine heartworm disease (Dirofilaria immitis) on Tenerife Island: An epidemiological update. Parasitol. Res. 100, 103–105. Morcho´n, R., Ferreira, A. C., Martin-Pacho, J., Montoya, A., Mortarino, M., Genchi, C., and Simo´n, F. (2004). Specific IgG antibody response against antigens of Dirofilaria immitis and its Wolbachia endosymbiont bacterium in cats with natural and experimental infections. Vet. Parasitol. 125, 313–321. Morcho´n, R., Lo´pez-Belmonte, J., Rodriguez-Barbero, A., and Simo´n, F. (2006). High levels of serum thromboxane B2 are generating human pulmonary dirofilariosis. Clin. Vaccine Immunol. 13, 1175–1176. Morcho´n, R., Lopez-Belmonte, J., Bazzocchi, C., Grandi, G., Kramer, L., and Simo´n, F. (2007a). Dogs with patent Dirofilaria immitis infection have higher expression of circulating IL-4, IL-10 and iNOS mRNA than those with occult infection. Vet. Immunol. Immunopathol. 115, 184–188. Morcho´n, R., Bazzocchi, C., Lopez-Belmonte, J., Martin-Pacho, J. R., Kramer, L. H., Grandi, G., and Simo´n, F. (2007b). iNOs expression is stimulated by the major surface protein (rWSP) from Wolbachia bacterial endosymbiont of Dirofilaria immitis following subcutaneous injection in mice. Parasitol. Int. 56, 71–75. Morini, S., Venco, L., Fagioli, P., and Genchi, C. (1998). Surgical removal of heartworms versus melarsomine treatment of naturally infected dogs with risk of thromboembolism. In ‘‘Proceedings of the American Heartworm Symposium ’98’’ (R. L. Seward, ed.), pp. 235–340. American Heartworm Society, Batavia, Illinois. Munnel, J. F., Weldon, J. S., Lewis, R. E., Thrall, D. E., and McCall, J. W. (1980). Intimal lesions of the pulmonary artery in dogs with experimental dirofilariasis. Am. J. Vet. Res. 41, 1108–1112. Murata, K., Yanai, T., Agatsuma, T., and Uni, S. (2003). Dirofilaria immitis infection of a snow leopard (Uncia uncia) in a Japanese zoo with mitochondrial DNA analysis. J. Vet. Med. Sci. 65, 945–947. Muro, A., and Cordero, M. (2001). Clinical aspects and diagnosis of human pulmonary dirofilariosis. In ‘‘Heartworm Infection in Humans and Animals’’ (F. Simo´n and C. Genchi, eds.), pp. 191–202. Ediciones Universidad de Salamanca, Salamanca, Spain. Muro, A., Cordero, M., Ramos, A., and Simon, F. (1991). Seasonal changes in the levels of anti-Dirofilaria immitis in an exposed human population. Trop. Med. Parasitol. 42, 371–374. Muro, A., Genchi, C., Cordero, M., and Simo´n, F. (1999). Human dirofilariasis in the European Union. Parasitol. Today 15, 386–389. Nakagaki, K., Suzuki, T., Hayama, S. I., and Kanda, E. (2000). Prevalence of filarial infection in racoon dogs in Japan. Parasitol. Int. 49, 253–256. Nelson, G. S. (1962). Dipetalonema reconditum (Grassi, 1889) from the dog with a note on its development in the flea, Ctenocephalides canis and the louse, Heterodoxus spiniger. J. Helminthol. 36, 297–308. Nelson, C. T. (1998). Incidence of Dirofilaria immitis in shelter cats from Southeast Texas. In ‘‘Proceedings of the American Heartworm Symposium ’98’’ (R. L. Seward, ed.), pp. 63–66. American Heartworm Society, Batavia, Illinois.
Heartworm Disease in Animals and Humans
279
Nelson, T. A., Gregory, D. G., and Laursen, J. R. (2003). Canine heartworms in coyotes in Illinois. J. Wildl. Dis. 39, 593–599. Nelson, C. T., McCall, J. W., Rubin, S. B., Buzhardt, L. F., Doiron, D. W., Graham, W., Longhofer, S. L., Guerrero, J., Robertson-Plough, C., and Paul, A. (2005a). 2005 Guidelines for the diagnosis, prevention and management of heartworm (Dirofilaria immitis) infection in dogs. Vet. Parasitol. 133, 255–266. Nelson, C. T., McCall, J. W., Rubin, S. B., Buzhardt, L. F., Doiron, D. W., Graham, W., Longhofer, S. L., Guerrero, J., Robertson-Plough, C., and Paul, A. (2005b). 2005 Guidelines for the diagnosis, prevention and management of heartworm (Dirofilaria immitis) infection in cats. Vet. Parasitol. 133, 267–275. Nelson, C. T., Seward, R. L., McCall, J. W., Rubin, S. B., Buzhardt, L. F., Graham, W., Longhofer, S. L., Guerrero, J., Robertson-Plough, C., Paul, A., and Carithers, D. (2007). 2007 Guidelines for the diagnosis, treatment and prevention of heartworm (Dirofilaria immitis) in cats. URL:http://www.heartwormsociety.org. Newton, W. L. (1968). Longevity of an experimental infection with Dirofilaria immitis in a dog. J. Parasitol. 54, 187–188. O’Connor, R. A., Jenson, J. S., Osborne, J., and Devaney, E. (2003). An enduring association? Microfilariae and immunosupression in lymphatic filariasis. Trends Parasitol. 19, 565–570. Okada, R., Imal, S., and Ishii, T. (1983). Clouded leopard, Neofelis neburosa, new host for Dirofilaria immitis. Jpn. J. Vet. Sci. 45, 849–852. Olmeda-Garcia, A. S., Rodriguez-Rodriguez, J. A., and Rojo Vazquez, F. A. (1993). Experimental transmission of Dipetalonema dracunculoides (Cobbod, 1970) by Rhipicephalus sanguineus (Latrille, 1806). Vet. Parasitol. 47, 339–342. Orihel, T. C. (1961). Morphology of the larval stages of Dirofilaria immitis in the dog. J. Parasitol. 47, 251–262. Orihel, T. A., and Eberhard, M. L. (1998). Zoonotic filariasis. Clin. Microbiol. Rev. 11, 366–381. Otto, G. F. (1975). Occurrence of the heartworm in unusual locations and unusual hosts. In ‘‘Proceedings of the Heartworm Symposium ’74’’ (H. C. Morgan, ed.), pp. 6–13. VM Publishing Inc., Bonner Springs, Kansas. Otto, G. F., and Jachowski, L. A., Jr. (1981). Mosquitoes and canine heartworm disease. In ‘‘Proceedings of the Heartworm Symposium ’80’’ (G. F. Otto, ed.), pp. 17–32. Veterinary Medicine Publishing Co, Edwardsville, Kansas. Pampiglione, S., and Rivasi, F. (2000). Human dirofilariasis due to Dirofilaria (Nochtiella) repens: An update of world literature from 1995 to 2000. Parassitologia 42, 235–254. Pampiglione, S., Elek, G., Pa´lfi, P., and Varga, I. (1999). Human Dirofilaria repens infection in Hungary: A case in the spermatic cord and a review of the literature. Acta Vet. Acad. Sci. Hung. 47, 77–83. Pampiglione, S., Rivasi, F., Angeli, G., Boldorini, R., Incensati, R. M., Pastormerlo, M., Pavesa, M., and Ramponi, A. (2001). Dirofilariasis due to Dirofilaria repens in Italy, an emergent zoonosis: Report of 60 new cases. Histopathology 38, 344–354. Pappas, L. G., and Lunzman, A. T. (1985). Canine heartworm in the domestic and wild canids of southeastern Nebraska. J. Parasitol. 71, 828–830. Patton, S., and Faulkner, C. T. (1992). Prevalence of Dirofilaria immitis and Dipetalonema reconditum infection in dogs: 805 cases (1980–1989). J. Am. Vet. Med. Assoc. 200, 1533–1534. Paul, A. J., Acre, K. E., Todd, K. S., Wallace, D. H., Jernigan, A. D., and Wallig, M. A. (1992). Efficacy of ivermectin against Dirofilaria immitis in cats 30 and 45 days post-infection. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 117–119. American Heartworm Society, Batavia, Illinois. Paul-Murphy, J., Work, T., Hunter, D., McFie, E., and Fjelline, D. (1994). Serologic survey and serum biological reference ranges of the free ranging mountain lion (Felis concolor) in California. J. Wildl. Dis. 30, 205–215.
280
John W. McCall et al.
Pence, D. B., Tewes, M. E., and Leack, L. L. (2003). Helminths of the ocelot from southern Texas. J. Wildl. Dis. 39, 683–689. Perera, L., Muro, A., Cordero, M., Villar, E., and Simo´n, F. (1994). Evaluation of a 22 kDa Dirofilaria immitis antigen for the immunodiagnosis of human pulmonary dirofilariasis. Trop. Med. Parasitol. 45, 249–252. Perera, L., Perez-Arellano, J. L., Corsero, M., Simo´n, F., and Muro, A. (1998). Utility of antibodies against a 22 kDa molecule of Dirofilaria immitis in the diagnosis of human pulmonary dirofilariasis. Trop. Med. Int. Health 3, 151–155. Perez, M., and Arlett, M. (1998). Parasitological survey at the small animal hospital of the School of Veterinary Science, Central University of Venezuela. In ‘‘Proceedings of XXIII World Small Animal Veterinary Association World Congress.’’ p. 801. Buenos Aires, Argentina. Periba`n˜ez, M. A., Lucientes, J., Arce, S., Morales, M., Castello, J. A., and Gracia, M. J. (2001). Histochemical differentiation of Dirofilaria immitis, Dirofilaria repens and Achantocheilonema dracunculoides microfilariae by staining with a commercial kit, Leucognost-SP. Vet. Parasitol. 102, 173–175. Peteta, L., Sigal, G., Ribicich, M., Rosa, A., and Perez Tort, G. (1998). Canine dirofilariasis in Villa la Nata, Province of Buenos Aires, Argentina. In ‘‘Proceedings of XXIII World Small Animal Veterinary Association World Congress.’’ p. 815. Buenos Aires, Argentina. Pherez, F. M. (2007). Factors affecting the emergence and prevalence of vector borne infections (VBI) and role of vertical transmission (VT). J. Vector Borne Dis. 44, 157–163. Piche´, C. A., Cavanaugh, M. T., Donoghue, A. R., and Radecki, S. V. (1998). Results of antibody and antigen testing for feline heartworm infection at HeskaÒ Veterinary Diagnostic Laboratories. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 139–143. American Heartworm Society, Batavia, Illinois. Plumb, D. C. (1995). ‘‘Veterinary Drug Handbook.’’ pp. 206–208. Pharma Publishing, White Bear Lake, Minnesota. Prieto, G., Cancrini, G., Muro, A., Genchi, C., and Simo´n, F. (2000). Seroepidemiology of Dirofilaria immitis and Dirofilaria repens in humans from three areas of Southern Europe. Res. Rev. Parasitol. 60, 95–98. Prieto, G., Ceciliani, F., Venco, L., Morchon, R., and Simon, F. (2002). Feline dirofilariosis: Antibody response to antigenic fractions containing specific 20 to 30 kDa polypeptides from the adult Dirofilaria immitis somatic antigen. Vet. Parasitol. 103, 341–353. Punkosdy, G. A., Addiss, D. G., and Lammie, P. J. (2003). Characterization of antibody responses to Wolbachia surface protein in humans with lymphatic filariasis. Infect. Immun. 71, 5104–5114. Raccurt, C. P. (1999). La dirofilariose, zoonose e´mergente et me´connue en France. Me´d. Trop. 59, 389–400. Rawlings, C. A. (1986). ‘‘Heartworm Disease in Dogs and Cats.’’ p. 329. WB Saunders Co, Philadelphia, USA. Rawlings, C. A., McCall, J. W., and Lewis, R. E. (1978). The response of the canine’s heart and lungs to Dirofilaria immitis. J. Am. Anim. Hosp. Assoc. 14, 17–32. Rawlings, C. A., Dawe, D. L., and McCall, J. W. (1982). Four types of occult Dirofilaria immitis infections in dogs. J. Am. Vet. Med. Assoc. 180, 1323–1326. Rawlings, C. A., Calvert, C. A., Glaus, T. M., and Jacobs, G. J. (1994). Surgical removal of heartworms. Semin. Vet. Med. Surg. (Small Anim.) 9, 200–205. Rawlings, C. A., Bowman, D. D., Howerth, E. W., Stansfield, D. G., Legg, W., and Luempert, L. G., III (2001). Response of dogs treated with ivermectin or milbemycin starting at various intervals after Dirofilaria immitis infection. Vet. Ther. 2, 193–207. Reifur, L., Thomaz-Soccol, V., and Montinari-Ferreira, F. (2004). Epidemiological aspects of filariosis on the coast of Parana´, Brazil: With emphasis on Dirofilaria immitis. Vet. Parasitol. 122, 273–286.
Heartworm Disease in Animals and Humans
281
Restani, R., Rossi, G., and Semproni, G. (1962). Due interessanti reperti clinici in cani portatori di Dirofilaria repens. Atti della Societa` Italiana delle Scienze Veterinarie 16, 406–412. Rishniw, M., Barr, S. C., Simpson, K. W., Frongillo, M. F., Franz, M., and Dominguez Alpizar, J. L. (2006). Discrimination between six species of canine microfilariae by a single polymerase chain reaction. Vet. Parasitol. 135, 303–314. Robertson-Plouch, C. K., Dillon, A. R., Brawner, W. R., and Guerrero, J. (1998). Prevalence of feline heartworm infections among cats with respiratory and gastrointestinal signs: Results of a multicenter study. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 57–62. American Heartworm Society, Batavia, Illinois. Roemer, G. W., Coonan, T. J., Garcelon, D. K., Starbird, C. H., and McCall, J. W. (2000). Spatial and temporal variation in the seroprevalence of canine heartworm antigen in the Island fox. J. Wildl. Dis. 36, 723–728. Rogers, A. H. (1998). Feline heartworm disease in northeast Florida: A case report. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 117–122. American Heartworm Society, Batavia, Illinois. Roncalli, R. A., Yamane, Y., and Nagata, T. (1998). Prevalence of Dirofilaria immitis in cats in Japan. Vet. Parasitol. 75, 81–89. Rosa, A., Ribicich, M., Bett, A., Kistermann, J., Cardillo, N., Basso, N., and Hallu, R. (2002). Prevalence of canine dirofilariosis in the City of Buenos Aires and its outskirts (Argentina). Vet. Parasitol. 109, 261–264. Ross, R. (1986). The pathogenesis of atherosclerosis: An update. N. Engl. J. Med. 314, 488–500. Rossi, L., Ferroglio, E., and Agostini, A. (2002). Use of moxidectin tablets in the control of canine subcutaneous dirofilariosis. Vet. Rec. 150, 383. Rossi, L., Ferroglio, E., and Agostini, A. (2004). Use of an injectable, sustained-release formulation of moxidectin to prevent canine subcutaneous dirofilariosis. Vet. Rec. 154, 26–27. Ruiz-Moreno, J. M., Bornay-Llynares, F. J., Prieto Maza, G., Medrano, M., Simo´n, F., and Eberhard, M. (1998). Subconjunctival infection with Dirofilaria repens. Serological confirmation of cure following surgery. Arch. Ophthalmol. 116, 1370–1372. Ruiz de Yba´n˜e´z, M. R., Martı´nez-Carrasco, C., Ortiz, J. M., Atout, T., and Bain, O. (2006). Dirofilaria immitis in an African lion (Panthera leo). Vet. Rec. 158, 240–242. Ryan, G. R., and Newcomb, K. M. (1995). Prevalence of feline heartworm disease: A global review. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 79–86. American Heartworm Society, Batavia, Illinois. Sacchi, L., Corona, S., Casiraghi, M., and Bandi, C. (2002). Does fertilization in the filarial nematode Dirofilaria immitis occur through endocytosis of spermatozoa? Parasitology 124, 87–95. Sacks, B. N., and Caswell-Chen, E. P. (2003). Reconstructing the spread of Dirofilaria immitis in California coyotes. J. Parasitol. 89, 319–323. Sang-Eun, L., Heung-Chul, K., Sung-Tae, C., Terry, A. K., and Won-Ja, L. (2007). Molecular survey of Dirofilaria immitis and Dirofilaria repens by direct PCR for wild caught mosquitoes in the Republic of Korea. Vet. Parasitol. 148, 149–155. Sartono, E., Lopriore, C., Kruize, Y. C., Kurniawan-Atmadja, A., Maizels, R. M., and Yazdanbakhsh, M. (1999). Reversal in microfilarial density and T cell responses in human lymphatic filariasis. Parasite Immunol. 21, 565–571. Schafer, M., and Berry, C. R. (1995). Cardiac and pulmonary artery mensuration in feline heartworm disease. Vet. Radiol. Ultrasound 36, 499–505. Schaub, R. G., and Rawlings, C. A. (1980). Pulmonary vascular response during phases of canine heartworm disease: Scanning electron microscopy study. Am. J. Vet. Res. 41, 1082–1089. Schaub, R. G., Rawlings, C. A., and Keith, J. C. (1981). Platelet adhesion and myointimal proliferation in canine pulmonary arteries. Am. J. Pathol. 104, 13–22.
282
John W. McCall et al.
Schwan, E. V., and Durand, D. T. (2002). Canine filariosis caused by Dirofilaria immitis in Mozambique: A small survey on the identification of microfilarie. J. S. Afr. Vet. Assoc. 73, 124–126. Schwan, E. V., and Schro¨ter, F. G. (2006). First record of Acanthocheilonema dracunculoides from domestic dogs in Namibia. J. S. Afr. Vet. Assoc. 77, 220–221. Scoles, G. A., and Dickson, S. L. (1995). New foci of canine heartworm associated with introductions of new vector species. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 27–35. American Heartworm Society, Batavia, Illinois. Segovia, J. M., Torres, J., Miquel, J., Llaneza, L., and Feliu, C. (2001). Helminths in the wolf, Canis lupus, from north-western Spain. J. Helminthol. 75, 183–192. Selcer, B. A., Newell, S. M., Mansour, A. E., and McCall, J. W. (1996). Radiographic and 2-D echocardiographic findings in eighteen cats experimentally exposed to D. immitis via mosquito bites. Vet. Radiol. Ultrasound 37, 37–44. Shaper, R., Heine, J., Arther, R. G., Charles, S. D., and McCall, J. W. (2007). Imidacloprid plus moxidectin to prevent heartworm infection (Dirofilaria immitis) in ferrets. Parasitol. Res. 101, S57–S62. Simo´n, F., Muro, A., Cordero, M., and Martı´n, J. (1991). A seroepidemiologic survey of human dirofilariasis in Western Spain. Trop. Med. Parasitol. 42, 106–108. Simo´n, F., Prieto, G., Morcho´n, R., Bazzocchi, C., Bandi, C., and Genchi, C. (2003). Immunoglobulin G antibodies against the endosymbionts of filarial nematodos (Wolbachia) in patients with pulmonary dirofilariasis. Clin. Diagn. Lab. Immunol. 10, 180–181. Simo´n, F., Lopez-Belmonte, J., Marcos-Atxutegi, C., Morchon, R., and Martın-Pacho, J. R. (2005). What is happening outside North America regarding human dirofilariasis? Vet. Parasitol. 133, 181–189. Sironi, M., Bandi, C., Sacchi, L., Di Sacco, B., Damiani, G., and Genchi, C. (1995). A close relative of the arthropods endosymbiont Wolbachia in a filarial worm. Mol. Biochem. Parasitol. 74, 223–227. Slocombe, J. O. D. (1992). Reflections on heartworm surveys in Canada over 15 years. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 21–30. American Heartworm Society, Batavia, Illinois. Slocombe, J. O. D., Surgeoner, G. A., and Srivastava, B. (1989). Determination of the heartworm transmission period and its use in diagnosis and control. In ‘‘Proceedings of the Heartworm Symposium ’89’’ (G. F. Otto, ed.), pp. 19–26. American Heartworm Society, Washington, DC. Slocombe, J. O. D., Srivastava, B., and Surgeoner, G. A. (1995). The transmission period for heartworm in Canada. In ‘‘Proceedings of the Heartworm Symposium ’95’’ (M. D. Soll and D. H. Knight, eds.), pp. 43–48. American Heartworm Society, Batavia, Illinois. Slonka, G. F., Castleman, W., and Krum, S. (1977). Adult heartworms in arteries and veins of a dog. J. Am. Vet. Med. Assoc. 170, 717–719. Snyder, P. S., Levy, J. K., Salute, M. E., Gorman, S. P., Kubilis, P. S., Smail, P. W., and George, L. L. (2000). Performance of serologic tests used to detect heartworm infection in cats. J. Am. Vet. Med. Assoc. 216, 693–700. Stephen, H. G. (2001). Migrating worms. In ‘‘Principle and Practice of Clinical Parasitology’’ (H. G. Stephen and D. P. Richard, eds.), pp. 535–551. John Wiley and Sons, Chichester, UK. Stewart, V. A., Blagburn, B. J., Hendrix, C. M., Hepler, D. I., and Grieve, R. B. (1992a). Milbemycin oxime as an effective preventative of heartworm (Dirofilaria immitis) infection in cats. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 127–131. American Heartworm Society, Batavia, Illinois. Stewart, V. A., Hepler, D. I., and Grieve, R. B. (1992b). Efficacy of milbemycin oxime in chemoprophylaxis of dirofilariasis in cats. Am. J. Vet. Res. 53, 2274–2277.
Heartworm Disease in Animals and Humans
283
Suba, N., Shiny, C., Taylor, M. J., and Narayanan, R. B. (2007). Brugia malayi Wolbachia hsp60 IgG antibody and isotype reactivity in different clinical groups infected or exposed to human bancroftian lymphatic filariasis. Exp. Parasitol. 116, 291–295. Sun, S., and Sugane, K. (1992). Immunodiagnosis of human dirofilariasis by enzyme linked immunosorbent assay using recombinant DNA-derived fusion protein. J. Helminthol. 66, 220–226. Supakorndej, P., McCall, J. W., Lewis, R. E., Rowan, S. J., Mansour, A. E., and Holmes, R. A. (1992). Biology, diagnosis and prevention of heartworm infection in ferrets. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 60–69. American Heartworm Society, Batavia, Illinois. Supakorndej, P., McCall, J. W., and Jun, J. J. (1994). Early migration and development of Dirofilaria immitis in the ferret, Mustela putorius furo. J. Parasitol. 80, 237–244. Supakorndej, P., Lewis, R. E., McCall, J. W., Dzimianski, M. T., and Holmes, R. A. (1995). Radiographic and angiographic evaluations of ferrets experimentally infected with Dirofilaria immitis. Vet. Radiol. Ultrasound 36, 23–29. Supakorndej, P., McCall, J. W., and Jun, J. J. (2004). Early migration and development of Dirofilaria immitis in the ferret, Mustela putorius furo. J. Parasitol. 80, 237–244. Suter, P. F., and Lord, P. F. (1974). Radiographic differentiation of disseminated pulmonary parenchymal diseases in dogs and cats. Vet. Clin. North Am. 4, 687–710. Svobodova, Z., Svobodova, V., Genchi, C., and Forejtek, P. (2006). The first report of authoctonous dirofilariosis in dogs in the Czech Republic. Helminthology 43, 242–245. Tanaka, H., Watanabe, M., and Ogawa, Y. (1985). Parasites of stray dogs and cats in the Kanto region, Honshu, Japan. J. Vet. Med. Japan 771, 657–661. Tatem, A. J., Hay, S. I., and Rogers, D. J. (2006a). Global traffic and disease vector dispersal. Proc. Natl. Acad. Sci. USA 103, 6242–6247. Tatem, A. J., Rogers, D. J., and Hay, S. I. (2006b). Global transport networks and infectious disease spread. Adv. Parasitol. 62, 293–343. Taylor, A. E. R. (1960). The development of Dirofilaria immitis in the mosquitoes Aedes aegypti. J. Helminthol. 34, 27–38. Taylor, M. J., Cross, H. F., Ford, L., Makunde, W. H., Prasad, G. B. K. S., and Bilo, K. (2001). Wolbachia bacteria in filarial immunity and disease. Parasite Immunol. 23, 401–409. Taylor, M. J., Bandi, C., and Hoerauf, A. (2005a). Wolbachia bacterial endosymbionts of filarial nematodes. Adv. Parasitol. 60, 248–286. Taylor, M. J., Makunde, W. H., McGarry, H. F., Turner, J. D., Mand, S., and Hoerauf, A. (2005b). Macrofilaricidal activity after doxycycline treatment of Wuchereria bancrofti: A double-blind, randomised placebo-controlled trial. Lancet 365, 2116–2121. Theis, J. H. (2005). Public health aspects of dirofilariasis in the United States. Vet. Parasitol. 133, 157–180. Theis, J. H., Stevens, F., and Law, M. (2001a). Distribution, prevalence, and relative risk of filariosis in dogs from the State of Washington (1997–1999). J. Am. Anim. Hosp. Assoc. 37, 339–347. Theis, J. H., Gilson, A., Simon, G. E., Bradshaw, B., and Clark, D. (2001b). Case report: Unusual location of Dirofilaria immitis in a 28-yearold man necessitates orchiectomy. Am. J. Trop. Med. Hyg. 64, 317–322. Thrall, D. E., Badertscher, R. R., and McCall, J. W. (1979). The pulmonary arterial circulation in dogs experimentally infected with Dirofilaria immitis: Its angiographic evaluation. J. Vet. Radiol. 20, 74–79. Treadwell, N. G., Scott-Moncrieff, J. C., Smith, J., and Rivers, B. J. (1998). Pneumothorax as a presenting sign of Dirofilaria immitis infection in cats. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 113–116. American Heartworm Society, Batavia, Illinois.
284
John W. McCall et al.
Uchide, T., and Saida, K. (2005). Elevated endothelin-1 expression in dogs with heartworm disease. J. Vet. Med. Sci. 67, 1155–1161. Uge, S., Matsumura, T., Ishibashi, K., Yatomi, K., Yoda, Y., and Kataoka, N. (1990). Occult infection of Dirofilaria immitis in stray dogs captured in Hyogo prefecture, Japan. Jpn. J. Parasitol. 39, 425–430. Venco, L. A. (1993). Diagnosis of vena cava syndrome. Veterinaria 7, 11–18. Venco, L., Borgarelli, M., Ferrari, E., Morini, S., and Genchi, C. (1998a). Surgical removal of heartworms from naturally-infected cats. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 241–246. American Heartworm Society, Batavia, Illinois. Venco, L., Calzolari, D., Mazzocchi, D., and Morini, S. (1998b). The use of echocardiography as a diagnostic tool for detection of feline heartworm (Dirofilaria immitis) infections. Feline Pract. 26, 6–9. Venco, L., Morini, S., Ferrari, E., and Genchi, C. (1998c). Technique for identifying heartworms in cats by 2-D echocardiography. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 97–102. American Heartworm Society, Batavia, Illinois. Venco, L., Genchi, C., Vigevani Colson, P., and Kramer, L. (2003). Relative utility of echocardiography, radiography, serologic testing and microfilariae counts to predict adult worm burden in dogs naturally infected with heartworms. In ‘‘Recent Advances in Heartworm Disease; Symposium ’01’’ (R. L. Seward and KnightD. H. , eds.), pp. 111–124. American Heartworm Society, Batavia, Illinois. Venco, L., McCall, J. W., Guerrero, J., and Genchi, C. (2004). Efficacy of long-term monthly administration of ivermectin on the progress of naturally acquired heartworm infection in dogs. Vet. Parasitol. 124, 259–268. Venco, L., Genchi, C., Genchi, M., Grandi, G., and Kramer, L. H. Clinical evolution and radiogrphic findings of feline heartworm infection in asymptomatic cats. Vet. Parasitol. (in press). Vezzani, D., and Carbajo, A. E. (2006). Spatial and temporal transmission risk of Dirofilaria immitis in Argentina. Int. J. Parasitol. 36, 1463–1472. Vezzani, D., Eiras, D. F., and Wisnivesky, C. (2006). Dirofilariasis in Argentina: Historical review and first report of Dirofilaria immitis in a natural mosquito population. Vet. Parasitol. 136, 259–273. Vezzoni, A., Genchi, C., and Raynaud, J. P. (1992). Adulticide efficacy of RM 340 in dogs with mild and severe natural infections. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 231–240. American Heartworm Society, Batavia Illinois. Vieira, C., Velez, I. D., Montoya, M. N., Agudelo, S., Alvarez, M. I., Genchi, C., and Simon, F. (1998). Dirofilaria immitis in Tikuna Indians and their dogs in the Colombian Amazon. Ann. Trop. Med. Parasitol. 92, 123–125. Wang, J. S., Tung, K. C., Huang, C. C., and Lai, C. H. (2005). Alteration of extracellular collagen matrix in the myocardium of canines infected with Dirofilaria immitis. Vet. Parasitol. 131, 261–265. Watkins, B. F., Toro, M., and Toro, G. (1998). Prevalence of heartworm antibody and antigenpositive sera among submissions to a commercial laboratory in the USA. In ‘‘Recent Advances in Heartworm Disease: Symposium ’98’’ (R. L. Seward, ed.), pp. 145–152. American Heartworm Society, Batavia, Illinois. Weiner, D. J., Aguirre, G., and Dubielzig, R. (1980). Ectopic-site filariid infection with immunologic follow-up of the host. In ‘‘Proceedings of the Heartworm Symposium ’80’’ (G. F. Otto, ed.), pp. 51–54. American Heartworm Society, Bonner Springs, Kansas. Wu, C. C., and Fan, P. C. (2003). Prevalence of canine dirofilariasis in Taiwan. J. Helminthol. 77, 83–88.
Heartworm Disease in Animals and Humans
285
Wong, M. M. (1974). Dirofilaria immitis: Fate and immunogenicity of irradiated infective stage larvae in Beagles. Exp. Parasitol. 35, 465–474. Yoshida, M., Nakagaki, K., Nogami, S., Harasawa, R., Maeda, R., Katae, H., and Hayashi, Y. (1997). Immunologic protection against canine heartworm infection. J. Vet. Med. Sci. 59, 1115–1121. Zimmerman, G. I., Knapp, S. E., Foreyt, W. J., Erekson, N. T., and Mackenzie, G. (1992). Heartworm infections in dogs in the northwestern United States and British Columbia, Canada. In ‘‘Proceedings of the Heartworm Symposium ’92’’ (M. D. Soll, ed.), pp. 15–20. American Heartworm Society, Batavia, Illinois. ˇ ivicˇnjak, T., Martinkovic´, F., and Beck, R. (2007). Canine dirofilariosis in Croatia: Let’s face Z it. In ‘‘FEDD, First European Dirofilaria Days.’’ Abstract Book, pp. 35. Zagreb, Croatia.
This page intentionally left blank
INDEX A Allergen(s) aeroallergens, 157 in allergic rhinitis, 152 asthma and immune responses, 169 definition of, 150 hypersensitivity, 153–154 Th2 responses, 155 tropomyosin, 166 Allergic conjunctivitis, 152 Allergic rhinitis, 152 Allergic Rhinitis and its Impact on Asthma (ARIA), 153 Anisakid nematodes average genetic distance, 100 among Anisakis spp., 101 among Contracaecum spp., 103 among P. decipiens complex, 102 at intraspecific level, 106–107 as biological indicators of habitat disturbance, 124–127 of Merluccius merluccius, 121–122 of Trachurus trachurus, 121–123 of trophic web stability, 124–127 of Xiphias gladius, 122–123 competitive exclusion, 119 genetic variability factors influencing, 135 genetic erosion, 134 genetic monitoring, 135–136 habitat disturbance, 134–136 host-anisakid co-evolutionary processes, 133–134 host-parasite phylogeny of Anisakis spp. and cetacean hosts, 109–111 biogeographical method, 109 Contracaecum spp. and pinnipeds host, 112–114 cophylogeny mapping, 108–109 mode of speciation, 114 host preferences among, 115
identification tools in human infections, 132 for speciation, 131–132 interspecific differentiation allozyme level, 101–102 mitochondrial level, 105 single strand conformation polymorphism (SSCP) analysis, 104–105 intra- and inter-specific competition, 118–119 intraspecific differentiation in Anisakis spp., 106, 108 in C. osculatum complex, 107–108 in P. decipiens complex, 106–107 intraspecific gene flow, 107–108 life cycle, 49, 124–125, 133 molecular systematics allozyme markers, 53–54 morphospecies, 52 PCR-DNA markers, 54–55 niche subdivision in C. osculatum complex, 116–118 in Pseudoterranova spp., 115–117 peripatric model, 100, 114 phylogenetic systematics allozyme data, 91–92 in Anisakis spp., 92–95 in Contracaecum spp., 96–99 in Pseudoterranova spp., 95–96 species detection of, 50–51 Anisakidosis, 49–50 Anisakis spp. average genetic distance, 101, 106–107 as biological tags of Merluccius merluccius, 121–122 of Trachurus trachurus, 121–123 of Xiphias gladius, 122–123 in clade I A. pegreffii in, 72 A. simplex in, 62 A. typica in, 72–73
287
288
Index
Anisakis spp. (cont.) A. ziphidarum in, 73–74 in clade II A. brevispiculata in, 75 A. paggiae in, 75 A. physeteris in, 74 distribution areas of, 63 genetic heterogeneity of, 76 genetic monitoring of, 135–136 host-parasite phylogeny of biogeographical method, 109 and cetacean hosts, 109–111 cophylogeny mapping, 108–109 host preferences, 115 identification of, 56–57 life-cycle pathways of, 124–125 mode of speciation, 114 phylogenetic studies of MP and NJ in, 93–96 UPGMA phenograms in, 92–93 taxonomy of adult morphology in, 55, 58 definitive hosts in, 64–65 intermediate hosts in, 66–71 phylogenetic analysis in, 61 Anopheles albimanus, 14 Anopheles atroparvus, 13 Anopheles freeborni, 14 Anopheles gambiae, 13 Anopheles maculopennis, 13 Anopheles punctipennis, 13 Anopheles quadrimaculatus, 13–14 Anti-malarial drugs chloroquine, 10–11 proguanil, 10 pyrimethamine, 10–11, 15 Anti-Wolbachia surface protein (WSP), 215 Ascaris lumbricoides atopic asthma studies, 162, 166 chronic urticaria, 169 in histamine release, 177 IgG4 and IgE antibodies, 172 parasite burden of, 162 Ascaris suum (ASC), 179 Atopic asthma allergic rhinitis impacts, 153 Ascaris lumbricoides, 162, 166 non-atopic phenotype, 167–168 parasitic infection in, 158–159 prevalence of, 154 protease effect in, 159 risk factors for, 160, 170
Atopic dermatitis, 152–153 Atopic disorders vs. allergic disorders, 154 allergic rhinitis in, 152–153 asthma in, 152 dermatitis in, 152–153 and helminthic infections aeroallergens responses, 162 anthelmintic treatment, 167 anti-inflammatory defence, 159 Ascaris-IgE seropositive, 162 asthma risk, 164–165 childhood allergy studies, 156 chronic blood and tissue parasite infections, 161 chronic urticaria, 169 cysteine protease, 159 geohelminth infection, 159–160 hepatitis A infection, 171 hookworm interactions, 173 house-dust mites (HDM) responses, 157–158 human ascariasis, 160 IgE concentration, 171 IgG4 and IgE antibodies, 172 IL-10 in, 164 immunologic variables, 156–157 mite allergens level, 164 nitric oxide (NO) levels, 174–175 omalizumab safety, 175 opisthorchiasis, 170 parasite burden, 163 shared antigens, 165–166 skin prick test (SPT) responses, 159–160 Th2 responses in, 151, 155–156 type 2 immune responses, 155 in vivo allergens exposure, 165 wheeze and allergen sensitization, 167–168 wheezing and asthma risk factors, 170 worm infestation and eczema, 162–163 laboratory studies, using parasites and rodent models allergic responses in, 178–179 Angiostrongylus costaricensis in, 179–180 asthma murine model, 179 BALB/c mice, 180–181 helminth larvae, 176 mesenteric lymph nodes (MLN), 181–182
Index
Nippostrongylus brasiliensis excretory–secretory (NES) products, 177–178 T helper type 2 lymphocytes in, 176 Average genetic distance see DNei B Biological species concept (BSC), 50, 53 Brooks’ parsimony analysis, 109 C Canine heartworm disease adulticidal and microfilaricidal treatment in melarsomine dihydrochloride administration in, 231–232 pulmonary thromboembolism and macrolides dosage in, 232 adult worms surgical extraction, 232–233 caval syndrome in, 223 chemoprophylaxis reaction of Knott test in, 230 macrocyclic lactones supplement in, 227–228 macrolide drugs treatment in, 229–230 selamectin administration and characteristics, 227–229 clinical presentation, 222–223 diagnosis of, 223 electrocardiography and echocardiography in, 227 ELISA and PCR methods in, 225 histochemical stain studies in, 224–225 microfilariae blood tests in, 223–224 thoracic radiography method in, 225–226 therapy vs. adult worms and microfilariae anti-inflammatory and antithrombotic, 231 patients’ pre-treatment assessment, 230–231 Caval syndrome, 223 Chloroquine, 10–11 Coin lesion, 258–259 Contracaecum spp. average genetic distance, 103 co-evolutionary processes, 133–134 definitive and intermediate hosts of, 86–88 distribution areas of, 63 host-parasite phylogeny of
289
biogeographical method, 109 cophylogeny mapping, 108–109 and pinnipeds hosts, 112–114 host preferences, 115 identification of, 59–60 infection in hosts, 127–128 life-cycle pathways of, 124–125 mode of speciation, 114 phylogenetic studies of NJ tree in, 98–99 UPGMA phenogram in, 97 single strand conformation polymorphism (SSCP) analysis, 104–105 taxonomy of allozyme markers in, 84–85 C. mirounga in, 90–91 C. ogmorhini complex in, 89 C. osculatum baicalensis in, 89–90 C. osculatum complex in, 85, 89 C. radiatum in, 90 morphological character in, 83–84 Cophylogeny mapping, 108–109 D Dirofilaria immitis antigen testing method, 248–249 blood tests morphology of, 223–224 canine and feline heartworm disease in, 209 human dirofilariosis in clinical aspects of, 258–260 epidemiology of, 260–262 immunology occult infection, 213–214 vaccine development in, 213 infection and pathogenesis congestive heart failure (CHF) in, 210 of endovascular lesions, 211–212 of feline heartworm disease, 212–213 in pulmonary arteries, 209–210 right ventricular hypertrophy, 212 infections in characterized areas, 200 heartworm development units (HDU), 200 stage (L3) development, 200–202 lifecycle in animals, 196 experimental infections in animal models, 197–199 in mosquitoes, 195–196
290
Index
Dirofilaria immitis (cont.) macrocyclic lactones administration, prevention by, 227–228 prevalence and distribution in Africa, 207 in Australia, Asia and South Pacific, 206 in Central America and Caribbean, 205–206 in Europe, 206–207 in North America, 203–204 in South America, 205 in wild carnivores, 208 Wolbachia endosymbiosis GroEL/HSP60, effect of, 220 human filarial nematodes, studies on, 218–219 humoral immune system, 219–220 NO production, role of, 219 DNei, 100 among Anisakis spp., 101 among Contracaecum spp., 103 among P. decipiens complex, 102 at intraspecific level, 106–107 E Egg white alone (EWI), heat-coagulated, 179 Enterobius vermicularis, 158, 172, 176 F Feline heartworm disease adulticide therapy and adult worm extraction in, 253 chemoprophylaxis in, 251–252 clinical presentation acute and chronic stages in, 236–237 arterial and airway disease, 234–235 heartworm associated respiratory disease (HARD), 234 symptoms in, 237–239 diagnosis of antibody testing method, 246–248 antigen testing method, 248–249 ecocardiography in, 249–250 microfilariae detection, 242–243 necropsy and difilariosis in, 251 serological tests, 240–242 thoracic radiography, 243–247 tracheal wash cytology in, 250 pathogenesis of, 212–213 symptomatic treatment in, 252
Ferret heartworm disease vs. adulticidal therapy, 256–257 chemoprophylaxis, 255–256 clinical presentation, 253–254 diagnosis of, 254–255 G Genetic erosion, 134 Genetic monitoring, 135 Geohelminth infections asthma risk in, 161–162 maternal, 173 omalizumab in, 174 parasites in, 160 prevalence of, 158–159 on wheeze and allergen sensitization, 167–168 H Habitat disturbance, marine ecosystems anisakids genetic variability factors influencing, 135 genetic erosion, 134 population size reductions, 136 of Antarctic region, 125–126 causes of, 127 and host population size, 124–125 Hardy–Weinberg equilibrium, 54, 77 Hay fever, 152 Heartworm adulticidal therapy arsenic drugs, 263 ivermectin and doxycycline dosage in, 264–265 macrocyclic lactones prophylactic doses ivermectin administration in, 263 melarsomine (MEL) administration in, 264 Heartworm associated respiratory disease (HARD) clinical signs, 213, 234 symptomatic treatment, 252 Helminthic infections, 150–151 House-dust mites (HDM) allergen, 163, 181 Human dirofilariosis clinical aspects by D. immitis coin lesion in, 258–259 serological diagnosis, 260 epidemiology of, 260–261 pulmonary dirofilariasis epidemiology, 261–262
Index
I Internal transcribed spacers of ribosomal DNA (ITS-rDNA), 104 L Lesseptian migration, 73 M Malaria epidemics, 25 infection fever types, 4 mode of transmission, 4–5 parasite types, 3–4 Plasmodium, 4–5 Malaria strain theory see also Plasmodium falciparum; Plasmodium vivax antigenic properties genetic diversity, P. falciparum, 19 heterologous immunity, 17 homologous immunity, 16 superinfection, 18, 23–27 clinical and parasitological response anti-malarial immunity, molecular basis, 23 heterologous tolerance, 21 immunity criteria, 20 inoculations, heterologous vs. homologous, 22 resistance to reinfection, 19–20 clinical virulence definition, 9 strain severity, 7–8 symptomatic differences, 6 transmission, 8 drug resistance chloroquine, 10–11 molecular-genetic basis for, 11–12 proguanil and pyrimethamine, 10 infectivity Anopheles, 12–13 clone vs. isolate, 15 gametocyte production, 14–15 vector-parasite transmission tests, 13 latency and relapse cross-inoculation responses, 32 host response, 29 hypnozoite formation and reactivation, 32–33 incubation period, 28 vs. P. vivax, 28–29
291
relapse frequencies, 29–30 strain diversity, 31–32 strain characteristics of antigenic properties, 16–18 clinical virulence, 6–9 drug resistance, 9–12 infectivity, 12–15 latency and relapse, 28–33 Maximum parsimony analysis in Anisakis spp., 93, 95 in Contracaecum spp., 99 in Pseudoterranova spp., 96 Merozoite surface protein 1 (MSP-1), 23 Mesenteric lymph node cells (MLNC), 181 MLEE see Multilocus allozyme electrophoresis Moxidectin canine heartworm diseases, 227–230 feline heartworm diseases, 251–252 ferret heartworm diseases, 256 injectable and commercial formulations of, 230 MP analysis see Maximum parsimony analysis Multilocus allozyme electrophoresis, 52, 131 Mustela putorius furo, 253 N Neighbour joining (NJ) method in Anisakis spp., 93–95 in Contracaecum and Phocascaris spp., 98–99 in Pseudoterranova spp., 95 Nippostrongylus brasiliensis, 165, 176–178 Nippostrongylus brasiliensis excretory–secretory (NES) products, 177–178 Nitric oxide (NO), 173–175 O Occult infections definition, 213 immune-mediated, 214 Opisthorchiasis, 170 Opisthorchis felineus, 170 Ovalbumin (OVA), 178–180 P Parasite strain, 35 PCR see Polymerase chain reaction Perennial allergic rhinitis, 152
292
Index
Peripheral blood mononuclear cells (PBMCs), 156, 162 Phocascaris spp. distribution of, 63 genetic divergence, 105 identification of, 59–60 phylogenetic studies, 96–99 taxonomy of, 84–85 Plasmodium falciparum antigenic properties, 17 chloroquine resistance, 10–11 clinical virulence, 6–7 gametocytemia, 14–15 genetic diversity, 19 latency and relapse, 28–33 vs. P. vivax, 28–29 pyrimethamine resistance in, 10 sterilizing effect on, 10 strain characteristics of, 36 strain severity of, 7 vector-parasite transmission test, 13 Plasmodium malariae, 4 Plasmodium ovale, 4 Plasmodium vivax Anopheles in, 13–14 clinical virulence of fever periodicity in, 6 strain severity in, 7–8 drug resistance in, 10 gametocyte production in, 12 heterologous immunity in, 17 latency in, 28–29 relapse and frequency, 29–30 relapse pattern, 32 superinfection in, 18 Polymerase chain reaction in anisakid nematodes in clade I, 72–73 identification of, 53–55 Positive skin test (PST), 157, 159, 163 Proguanil, 10 Pseudoterranova spp. average genetic distance, 102 distribution areas of, 63 host preferences, 115 in human infections, 132 identification of, 58 intraspecific differentiation, 106–107 life-cycle pathways of, 124–125 mode of speciation, 114 niche subdivision, 115–117 phylogenetic studies of, 95–96
taxonomy of definitive and intermediate hosts in, 80–82 P. azarasi in, 83 P. krabbei and P. bulbosa in, 79 population genetic analysis in, 77–78 Terranova in, 76–77 Pyrimethamine, 10–11, 15 R Reproductive isolating mechanisms (RIMs), 100 S Schistosoma haematobium, 157–158 Schistosoma mansoni, 157, 163–164 Schistosomiasis, 157–158, 163–164 Seasonal allergic rhinitis, 152 Selamectin canine heartworm diseases, 227–229 feline heartworm diseases, 234 ferret heartworm diseases, 251 Severe-combined immune deficient (SCID), 176–177 Single strand conformation polymorphism (SSCP), 104–105 Superinfection multiple strain immunity, 25–27 parasite density effects, 27–28 and reinfection, 23 U Unweighted pair group method using arithmetic averages (UPGMA phenograms) in Anisakis spp., 92–93 in Contracaecum spp., 97 in Pseudoterranova spp., 95 V Virulence definition of, 9 of P. falciparam, 6–7 of P. vivax fever periodicity in, 6 strain severity in, 6 W Wolbachia endosymbiosis antibiotic treatment on D. immitis
Index
doxycycline treatment, 220–221 immunization methods in, 221–222 use of tetracyclins, 221 with Brugia malayi, 216–217 with D. immitis, 215–216 effect of GroEL/HSP60, 220
293
humoral immune system, 219–220 role of NO production, 219 studies on human filarial nematodes, 218–219 Wolbachia surface protein (WSP) and GroEL role, 218
This page intentionally left blank
CONTENTS OF VOLUMES IN THIS SERIES Volume 41 Drug Resistance in Malaria Parasites of Animals and Man W. Peters Molecular Pathobiology and Antigenic Variation of Pneumocystis carinii Y. Nakamura and M. Wada Ascariasis in China P. Weidono, Z. Xianmin and D.W.T. Crompton The Generation and Expression of Immunity to Trichinella spiralis in Laboratory Rodents R.G. Bell Population Biology of Parasitic Nematodes: Application of Genetic Markers T.J.C. Anderson, M.S. Blouin and R.M. Brech Schistosomiasis in Cattle J. De Bont and J. Vercruysse
Volume 42 The Southern Cone Initiative Against Chagas Disease C.J. Schofield and J.C.P. Dias Phytomonas and Other Trypanosomatid Parasites of Plants and Fruit E.P. Camargo Paragonimiasis and the Genus Paragonimus D. Blair, Z.-B. Xu, and T. Agatsuma Immunology and Biochemistry of Hymenolepis diminuta J. Anreassen, E.M. Bennet-Jenkins, and C. Bryant Control Strategies for Human Intestinal Nematode Infections
M. Albonico, D.W.T. Cromption, and L. Savioli DNA Vaocines: Technology and Applications as Anti-parasite and Anti-microbial Agents J.B. Alarcon, G.W. Wainem and D.P. McManus
Volume 43 Genetic Exchange in the Trypanosomatidae W. Gibson and J. Stevens The Host-Parasite Relationship in Neosporosis A. Hemphill Proteases of Protozoan Parasites P.J. Rosenthal Proteinases and Associated Genes of Parasitic Helminths J. Tort, P.J. Brindley, D. Knox, K.H. Wolfe, and J.P. Dalton Parasitic Fungi and their Interaction with the Insect Immune System A. Vilcinskas and P. Go¨tz
Volume 44 Cell Biology of Leishmania B. Handman Immunity and Vaccine Development in the Bovine Theilerioses N. Boulter and R. Hall The Distribution of Schistosoma bovis Sonaino, 1876 in Relation to Intermediate Host Mollusc-Parasite Relationships H. Mone´, G. Mouahid, and S. Morand
295
296
Contents of Volumes in This Series
The Larvae of Monogenea (Platyhelminthes) I.D. Whittington, L.A. Chisholm, and K. Rohde Sealice on Salmonids: Their Biology and Control A.W. Pike and S.L. Wadsworth
Volume 45 The Biology of some Intraerythrocytic Parasites of Fishes, Amphibia and Reptiles A.J. Davies and M.R.L. Johnston The Range and Biological Activity of FMR Famide-related Peptides and Classical Neurotransmitters in Nematodes D. Brownlee, L. Holden-Dye, and R. Walker The Immunobiology of Gastrointestinal Nematode Infections in Ruminants A. Balic, V.M. Bowles, and E.N.T. Meeusen
Satellites, Space, Time and the African Trypanosomiases D.J. Rogers Earth Observation, Geographic Information Systems and Plasmodium falciparum Malaria in Sub-Saharan Africa S.I. Hay, J. Omumbo, M. Craig, and R.W. Snow Ticks and Tick-borne Disease Systems in Space and from Space S.E. Randolph The Potential of Geographical Information Systems (GIS) and Remote Sensing in the Epidemiology and Control of Human Helminth Infections S. Brooker and E. Michael Advances in Satellite Remote Sensing of Environmental Variables for Epidemiological Applications S.J. Goetz, S.D. Prince, and J. Small
Volume 46
Forecasting Diseases Risk for Increased Epidemic Preparedness in Public Health M.F. Myers, D.J. Rogers, J. Cox, A. Flauhalt, and S.I. Hay
Host-Parasite Interactions in Acanthocephala: A Morphological Approach H. Taraschewski
Education, Outreach and the Future of Remote Sensing in Human Health B.L. Woods, L.R. Beck, B.M. Lobitz, and M.R. Bobo
Eicosanoids in Parasites and Parasitic Infections A. Daugschies and A. Joachim
Volume 48
Volume 47 An Overview of Remote Sensing and Geodesy for Epidemiology and Public Health Application S.I. Hay Linking Remote Sensing, Land Cover and Disease P.J. Curran, P.M. Atkinson, G.M. Foody, and E.J. Milton Spatial Statistics and Geographic Information Systems in Epidemiology and Public Health T.P. Robinson
The Molecular Evolution of Trypanosomatidae J.R. Stevens, H.A. Noyes, C.J. Schofield, and W. Gibson Transovarial Transmission in the Microsporidia A.M. Dunn, R.S. Terry, and J.E. Smith Adhesive Secretions in the Platyhelminthes I.D. Whittington and B.W. Cribb The Use of Ultrasound in Schistosomiasis C.F.R. Hatz Ascaris and Ascariasis D.W.T. Crompton
Contents of Volumes in This Series
Volume 49
Volume 52
Antigenic Variation in Trypanosomes: Enhanced Phenotypic Variation in a Eukaryotic Parasite H.D. Barry and R. McCulloch
The Ecology of Fish Parasites with Particular Reference to Helminth Parasites and their Salmonid Fish Hosts in Welsh Rivers: A Review of Some of the Central Questions J.D. Thomas
The Epidemiology and Control of Human African Trypanosomiasis J. Pe´pin and H.A. Me´da Apoptosis and Parasitism: from the Parasite to the Host Immune Response G.A. DosReis and M.A. Barcinski Biology of Echinostomes Except Echinostoma B. Fried
297
Biology of the Schistosome Genus Trichobilharzia P. Hora´k, L. Kola´rova´, and C.M. Adema The Consequences of Reducing Transmission of Plasmodium falciparum in Africa R.W. Snow and K. Marsh
The Malaria-Infected Red Blood Cell: Structural and Functional Changes B.M. Cooke, N. Mohandas, and R.L. Coppel
Cytokine-Mediated Host Responses during Schistosome Infections: Walking the Fine Line Between Immunological Control and Immunopathology K.F. Hoffmann, T.A. Wynn, and D.W. Dunne
Schistosomiasis in the Mekong Region: Epidemiology and Phytogeography S.W. Attwood
Volume 53
Volume 50
Molecular Aspects of Sexual Development and Reproduction in Nematodes and Schistosomes P.R. Boag, S.E. Newton, and R.B. Gasser Antiparasitic Properties of Medicinal Plants and Other Naturally Occurring Products S. Tagboto and S. Townson
Interactions between Tsetse and Trypanosomes with Implications for the Control of Trypanosomiasis S. Aksoy, W.C. Gibson, and M.J. Lehane Enzymes Involved in the Biogenesis of the Nematode Cuticle A.P. Page and A.D. Winter
Volume 51
Diagnosis of Human Filariases (Except Onchocerciasis) M. Walther and R. Muller
Aspects of Human Parasites in which Surgical Intervention May Be Important D.A. Meyer and B. Fried
Volume 54
Electron-transfer Complexes in Ascaris Mitochondria K. Kita and S. Takamiya Cestode Parasites: Application of In Vivo and In Vitro Models for Studies of the Host-Parasite Relationship M. Siles-Lucas and A. Hemphill
Introduction – Phylogenies, Phylogenetics, Parasites and the Evolution of Parasitism D.T.J. Littlewood Cryptic Organelles in Parasitic Protists and Fungi B.A.P. Williams and P.J. Keeling
298
Contents of Volumes in This Series
Phylogenetic Insights into the Evolution of Parasitism in Hymenoptera J.B. Whitfield Nematoda: Genes, Genomes and the Evolution of Parasitism M.L. Blaxter Life Cycle Evolution in the Digenea: A New Perspective from Phylogeny T.H. Cribb, R.A. Bray, P.D. Olson, and D.T.J. Littlewood Progress in Malaria Research: The Case for Phylogenetics S.M. Rich and F.J. Ayala Phylogenies, the Comparative Method and Parasite Evolutionary Ecology S. Morand and R. Poulin Recent Results in Cophylogeny Mapping M.A. Charleston Inference of Viral Evolutionary Rates from Molecular Sequences A. Drummond, O.G. Pybus, and A. Rambaut Detecting Adaptive Molecular Evolution: Additional Tools for the Parasitologist J.O. McInerney, D.T.J. Littlewood, and C.J. Creevey
Volume 55 Contents of Volumes 28–52 Cumulative Subject Indexes for Volumes 28–52 Contributors to Volumes 28–52
Volume 56 Glycoinositolphospholipid from Trypanosoma cruzi: Structure, Biosynthesis and Immunobiology J.O. Previato, R. Wait, C. Jones, G.A. DosReis, A.R. Todeschini, N. Heise and L.M. Previata Biodiversity and Evolution of the Myxozoa E.U. Canning and B. Okamura
The Mitochondrial Genomics of Parasitic Nematodes of Socio-Economic Importance: Recent Progress, and Implications for Population Genetics and Systematics M. Hu, N.B. Chilton, and R.B. Gasser The Cytoskeleton and Motility in Apicomplexan Invasion R.E. Fowler, G. Margos, and G.H. Mitchell
Volume 57 Canine Leishmaniasis J. Alvar, C. Can˜avate, R. Molina, J. Moreno, and J. Nieto Sexual Biology of Schistosomes H. Mone´ and J. Boissier Review of the Trematode Genus Ribeiroia (Psilostomidae): Ecology, Life History, and Pathogenesis with Special Emphasis on the Amphibian Malformation Problem P.T.J. Johnson, D.R. Sutherland, J.M. Kinsella and K.B. Lunde The Trichuris muris System: A Paradigm of Resistance and Susceptibility to Intestinal Nematode Infection L.J. Cliffe and R.K. Grencis Scabies: New Future for a Neglected Disease S.F. Walton, D.C. Holt, B.J. Currie, and D.J. Kemp
Volume 58 Leishmania spp.: On the Interactions they Establish with Antigen-Presenting Cells of their Mammalian Hosts J.-C. Antoine, E. Prina, N. Courret, and T. Lang Variation in Giardia: Implications for Taxonomy and Epidemiology R.C.A. Thompson and P.T. Monis Recent Advances in the Biology of Echinostoma species in the ‘‘revolutum’’ Group B. Fried and T.K. Graczyk
Contents of Volumes in This Series
Human Hookworm Infection in the 21st Century S. Brooker, J. Bethony, and P.J. Hotez The Curious Life-Style of the Parasitic Stages of Gnathiid Isopods N.J. Smit and A.J. Davies
Volume 59 Genes and Susceptibility to Leishmaniasis Emanuela Handman, Colleen Elso, and Simon Foote Cryptosporidium and Cryptosporidiosis R.C.A. Thompson, M.E. Olson, G. Zhu, S. Enomoto, Mitchell S. Abrahamsen and N.S. Hijjawi Ichthyophthirius multifiliis Fouquet and Ichthyophthiriosis in Freshwater Teleosts R.A. Matthews Biology of the Phylum Nematomorpha B. Hanelt, F. Thomas, and A. SchmidtRhaesa
Volume 60 Sulfur-Containing Amino Acid Metabolism in Parasitic Protozoa Tomoyoshi Nozaki, Vahab Ali, and Masaharu Tokoro The Use and Implications of Ribosomal DNA Sequencing for the Discrimination of Digenean Species Matthew J. Nolan and Thomas H. Cribb
299
Volume 61 Control of Human Parasitic Diseases: Context and Overview David H. Molyneux Malaria Chemotherapy Peter Winstanley and Stephen Ward Insecticide-Treated Nets Jenny Hill, Jo Lines, and Mark Rowland Control of Chagas Disease Yoichi Yamagata and Jun Nakagawa Human African Trypanosomiasis: Epidemiology and Control E.M. Fe`vre, K. Picozzi, J. Jannin, S.C. Welburn and I. Maudlin Chemotherapy in the Treatment and Control of Leishmaniasis Jorge Alvar, Simon Croft, and Piero Olliaro Dracunculiasis (Guinea Worm Disease) Eradication Ernesto Ruiz-Tiben and Donald R. Hopkins Intervention for the Control of SoilTransmitted Helminthiasis in the Community Marco Albonico, Antonio Montresor, D.W. T. Crompton, and Lorenzo Savioli Control of Onchocerciasis Boakye A. Boatin and Frank O. Richards, Jr. Lymphatic Filariasis: Treatment, Control and Elimination Eric A. Ottesen
Advances and Trends in the Molecular Systematics of the Parasitic Platyhelminthes Peter D. Olson and Vasyl V. Tkach
Control of Cystic Echinococcosis/ Hydatidosis: 1863–2002 P.S. Craig and E. Larrieu
Wolbachia Bacterial Endosymbionts of Filarial Nematodes Mark J. Taylor, Claudio Bandi, and Achim Hoerauf
Control of Taenia solium Cysticercosis/ Taeniosis Arve Lee Willingham III and Dirk Engels
The Biology of Avian Eimeria with an Emphasis on their Control by Vaccination Martin W. Shirley, Adrian L. Smith, and Fiona M. Tomley
Implementation of Human Schistosomiasis Control: Challenges and Prospects Alan Fenwick, David Rollinson, and Vaughan Southgate
300
Contents of Volumes in This Series
Volume 62 Models for Vectors and Vector-Borne Diseases D.J. Rogers Global Environmental Data for Mapping Infectious Disease Distribution S.I. Hay, A.J. Tatem, A.J. Graham, S.J. Goetz, and D.J. Rogers Issues of Scale and Uncertainty in the Global Remote Sensing of Disease P.M. Atkinson and A.J. Graham Determining Global Population Distribution: Methods, Applications and Data D.L. Balk, U. Deichmann, G. Yetman, F. Pozzi, S.I. Hay, and A. Nelson Defining the Global Spatial Limits of Malaria Transmission in 2005 C.A. Guerra, R.W. Snow and S.I. Hay The Global Distribution of Yellow Fever and Dengue D.J. Rogers, A.J. Wilson, S.I. Hay, and A.J. Graham Global Epidemiology, Ecology and Control of Soil-Transmitted Helminth Infections S. Brooker, A.C.A. Clements and D.A.P. Bundy Tick-borne Disease Systems: Mapping Geographic and Phylogenetic Space S.E. Randolph and D.J. Rogers Global Transport Networks and Infectious Disease Spread A.J. Tatem, D.J. Rogers and S.I. Hay Climate Change and Vector-Borne Diseases D.J. Rogers and S.E. Randolph
Volume 63 Phylogenetic Analyses of Parasites in the New Millennium David A. Morrison
Targeting of Toxic Compounds to the Trypanosome’s Interior Michael P. Barrett and Ian H. Gilbert Making Sense of the Schistosome Surface Patrick J. Skelly and R. Alan Wilson Immunology and Pathology of Intestinal Trematodes in Their Definitive Hosts Rafael Toledo, Jose´-Guillermo Esteban, and Bernard Fried Systematics and Epidemiology of Trichinella Edoardo Pozio and K. Darwin Murrell
Volume 64 Leishmania and the Leishmaniases: A Parasite Genetic Update and Advances in Taxonomy, Epidemiology and Pathogenicity in Humans Anne-Laure Ban˜uls, Mallorie Hide and Franck Prugnolle Human Waterborne Trematode and Protozoan Infections Thaddeus K. Graczyk and Bernard Fried The Biology of Gyrodctylid Monogeneans: The ‘‘Russian-Doll Killers’’ T.A. Bakke, J. Cable, and P.D. Harris Human Genetic Diversity and the Epidemiology of Parasitic and Other Transmissible Diseases Michel Tibayrenc
Volume 65 ABO Blood Group Phenotypes and Plasmodium falciparum Malaria: Unlocking a Pivotal Mechanism Marı´a-Paz Loscertales, Stephen Owens, James O’Donnell, James Bunn, Xavier Bosch-Capblanch, and Bernard J. Brabin Structure and Content of the Entamoeba histolytica Genome C. G. Clark, U. C. M. Alsmark, M. Tazreiter, Y. Saito-Nakano, V. Ali,
Contents of Volumes in This Series
S. Marion, C. Weber, C. Mukherjee, I. Bruchhaus, E. Tannich, M. Leippe, T. Sicheritz-Ponten, P. G. Foster, J. Samuelson, C. J. Noe¨l, R. P. Hirt, T. M. Embley, C. A. Gilchrist, B. J. Mann, U. Singh, J. P. Ackers, S. Bhattacharya, A. Bhattacharya, A. Lohia, N. Guille´n, M. Ducheˆne, T. Nozaki, and N. Hall Epidemiological Modelling for Monitoring and Evaluation of Lymphatic Filariasis Control
301
Edwin Michael, Mwele N. MalecelaLazaro, and James W. Kazura The Role of Helminth Infections in Carcinogenesis David A. Mayer and Bernard Fried A Review of the Biology of the Parasitic Copepod Lernaeocera branchialis (L., 1767)(Copepoda: Pennellidae Adam J. Brooker, Andrew P. Shinn, and James E. Bron
60⬚
i olar C Ar tic P
rcle
30⬚
0⬚
30⬚
Antarc
60⬚
tic Po
60⬚
120⬚ A. pegreffii
A. simplex (s.s. )
C. osculatum (s.s.) P. decipiens (s.s.) Ph. phocae
A. simplex C
C. osculatum A P. krabbei
A. typica
C. osculatum B
P. bulbosa
P. azarasi
A. physeteris
C. o. baicalensis P. decipiens E
0⬚ A. brevispiculata C. osculatum D
lar Cir
cle
60⬚ A. ziphidarum C. osculatum E
120⬚ A. paggiae C. radiatum
180⬚
Anisakis sp. C. mirounga
C. ogmorhini (s.s)
C. margolisi
P. cattani
Ph. cystophorae
Plate 2.1 World map showing the so far known distribution areas of anisakid species of Anisakis (▭), Pseudoterranova (△), Contracaecum (○) and Phocascaris (?). The geographical areas indicated are related to the sampling localities for their definitive and intermediate hosts.
A (allozyme data)
B (mtDNA cox2) A. simplex (s.s.)
A. simplex (s.s.) 71
A. pegreffii
100 96
A. pegreffii
A. simplex C
99
100
A. simplex C
A. typica
A. typica
A. ziphidarum
73
A. ziphidarum
71
76 Anisakis sp.
Anisakis sp.
A. physeteris
A. paggiae 50
A. brevispiculata
A. brevispiculata A. physeteris
A. paggiae P. decipiens
70
(outgroup) P. decipiens Clade I Clade II
Plate 2.2 Genetic relationships among Anisakis spp. depicted by (A) neighbour-joining (NJ) tree inferred from chord-distance values (Dc, Cavalli-Sforza and Edwards, 1967) from allozyme data; (B) NJ inferred from K2P distance values obtained by mtDNA cox2 sequences analysis (data from Valentini et al., 2006). Bootstrap values 60 are shown at the internal nodes. Pseudoterranova ceticola as outgroup. The two data sets are congruent in depicting the existence of two main clades: (I) includes the A. simplex species complex [A. pegreffii, A. simplex (s.s.), A. simplex C], A. typica, A. ziphidarum and Anisakis sp.; (II) comprises the species A. physeteris, A. brevispiculata and A. paggiae. The species included in the two main clusters show a different larval morphology: type I (sensu Berland, 1961) in the first group and type II in the second one.
A (allozyme data)
B (mtDNA cox2)
98
C. osculatum A
C. osculatum B
C. osculatum E
C. osculatum (s.s.)
C. osculatum D
P. cystophorae C. osculatum A
C. o. baicalensis
C. osculatum D C. osculatum (s.s.)
90
96
62 94
C. o. baicalensis
C. osculatum B
90
95
96 99
C. osculatum E
85
P. cystophorae C. radiatum
88
C. radiatum
95 C. mirounga
C. mirounga
C. ogmorhini (s.s. )
C. ogmorhini (s.s. )
100 C. margolisi
C. margolisi P. ceticola
100
(outgroup)
P. ceticola
Clade I Clade II
Plate 2.4 Genetic relationships between species of Contracaecum and Phocascaris from pinnipeds depicted by (A) NJ tree inferred from chord-distance values (Dc, Cavalli-Sforza and Edwards, 1967) from allozyme data and (B) NJ inferred from mtDNA cox2 sequences analysis. Bootstrap values 60 are shown at the internal nodes. Pseudoterranova ceticola as outgroup. The two data sets are congruent in depicting the existence of two main clades: (I) includes the species of C. osculatum complex, Phocascaris cystophorae, C. radiatum, C. mirounga; (II) comprises C. ogmorhini (s.s.) and C. margolisi. The two phylogenetic trees show P. cystophorae nested in the subclade formed by the species of C. osculatum complex.
Host Bottlenose dolphin
Parasite A. pegreffii 71
F
Short-finned pilot whale
Mage19
E
A. simplex (s.s.)
Isi14 Isi36 Isi36 Mago24 Mago26 Mago32
G
Narwhal
Amz13 Bando1
D
Amazon river dolphin H
60
A. simplex C
Amz11
La plata dolphin
Mago8 Mago13
Yangtze river dolphin
I
C
Tuti24 Tuti35
Mago21 Mago22
A. typica 73
Baird’s beaked whale Mesoplodon sp.
B
A. ziphidarum
76
Anisakis sp.
Sperm8 Sperm28 Sperm47
Ganges river dolphin J
A Bando1 SP316
100
Dall’s porpoise
K Sp9
Sp2
Pygmy sperm whale
Pm72 Pm52 M11
A. paggiae A. brevispiculata
Sperm whale L Hump20 Hump203 aaa792
70
A. physeteris Humpback whale Fin whale
(outgroup) P. ceticola Dolphins
Minke whale
Hosts of Anisakis spp.
Ziphiids Physeterids
Plate 2.5 Tanglegram of phylogenies of Anisakis spp. and their cetacean hosts. It shows the phylogeny of a group of extant cetaceans (adapted from Nikaido et al., 2001) mapped into the phylogeny of Anisakis spp. inferred from mtDNA cox2 sequence analysis. Lines depict the observed host–parasite co-speciation events; the dotted line indicates possible host-switching events (redrawn from Mattiucci and Nascetti, 2006).
Host P. vitulina vitulina P. vitulina richardsi
97 65
P. largha H. grypus
Parasite C. osculatum B C. osculatum (s.s.)
95
Ph. cystophorae C. osculatum A
P. hispida C. o. baicalensis
90 87
62 94
89
P. fasciata
C. osculatum D
96 99
P. groenlandicus E. barbatus
74
80
L. weddellii
85
C. osculatum E C. radiatum
95
M. schauinslandi H. leptonyx
97
M. leonina
C. mirounga
C. ursinus Z. californianus
100 100
88
91
A. gazella
C. margolisi
100
C. ogmorhini (s.s.)
A. forsteri (outgroup) P. ceticola
97
93 100
O. rosmarus Brown bear Polar bear
American black bear Domestic cat Lion
Hosts of Contracaecum spp.
Phocinae Monachinae Otariidae
Tiger
Plate 2.6 Tanglegram of phylogenies of Contracaecum spp. and their pinnipeds hosts. It shows the phylogeny of a group of extant pinnipeds (adapted from Arnason et al., 1995) mapped into the phylogeny of Contracaecum spp. inferred from mtDNA cox2 sequence analysis (adapted from Mattiucci et al., 2008b). Lines depict possible host–parasite co-speciation events; the dotted line indicates possible host-switching events.
North Atlantic Ocean
Halichoerus grypus
Erignathus barbatus
Pagophilus groenlandicus
Pseudoterranova bulbosa
Contracaecum osculatum A
Pseudoterranova decipiens (s.s.)
Contracaecum osculatum B
Pseudoterranova krabbei
Contracaecum osculatum (s.s.)
Phoca vitulina
Plate 2.7 Relative proportions of Pseudoterranova decipiens and Contracaecum osculatum species complexes in definitive hosts (seals) from the North Atlantic Ocean. It shows differences in their definitive host preferences.
A
Merluccius merluccius
B
Trachurus trachurus
C
Xiphias gladius
A. simplex (s.s.) A. pegreffii A. brevispiculata A. physeteris A. ziphidarum A. typica A. paggiae Anisakis sp.
50 45 40 35 –50 –45 –40 –35 –30 –25 –20 –15
50 45 40 35 –50 –45 –40 –35 –30 –25 –20 –15 –10 15 10 5 0 –30
–25
–20
–15
–10
–5
Plate 2.9 Relative proportions of larval specimens of Anisakis spp. from different fish hosts: (A) Merluccius merluccius; (B) Trachurus trachurus; (C) Xiphias gladius throughout their range of distribution. It shows the use of biogeographical aspects of Anisakis spp., genetically identified to species level, as biological tags for their fish stocks definition in European waters.
Host
Definitive
Intermediate/ paratenic
First intermediate
Habitat disturbance By-catch and hunting, viral diseases
Overfishing, contaminants
Global warming, sea water acidification, pollution
Plate 2.10 Relationship between habitat disturbance and host population size. It depicts causes of habitat disturbance (right) that can affect the population size of definitive and intermediate hosts (at different trophic levels of a marine food web) involved in speculative life-cycle pathways of anisakid nematodes of the genera Anisakis, Pseudoterranova and Contracaecum (includes only species from pinnipeds). (Modified from Mattiucci and Nascetti, 2007, with permission from Elsevier.)
Plate 4.1 Lung from a dog with heartworm infection. Note the obstruction of vascular lumen due to endothelial proliferation and worm fragments (hematoxylin/eosin, 40).
Plate 4.2 Anti-Wolbachia surface protein (WSP) immunohistochemistry. D. immitis female with positive staining within a lateral hypodermal chord. (ABC-HRP, 100).
Plate 4.3 Anti-Wolbachia immunohistochemistry. D. immitis female with positivestaining microfilariae within the uterus. (ABC/HRP, 100).
Plate 4.4 Anti-Wolbachia immunohistochemistry. Kidney from a dog with heartworm infection. Note positive staining for the WSP within microfilariae (ABC/HRP, 60).
Plate 4.7 Small pulmonary arteriole of a heartworm-challenged cat on monthly chemoprophylaxis. (Used with permission of the American Heartworm Society.)
Plate 4.8 Small pulmonary arteriole of a cat with a chemically abbreviated heartworm infection. (Used with permission of the American Heartworm Society.)
Plate 4.9 Small pulmonary arteriole of a non-treated cat with adult heartworms. (Used with permission of the American Heartworm Society.)
Plate 4.10 Bronchiole of a heartworm-challenged cat on monthly chemoprophylaxis. (Used with permission of the American Heartworm Society.)
Plate 4.11 Bronchiole of a cat with a chemically abbreviated heartworm infection. (Used with permission of the American Heartworm Society.)
Plate 4.12 Bronchiole of a non-treated cat with adult heartworms. (Used with permission of the American Heartworm Society.)
Plate 4.13 Alveoli of a heartworm-challenged cat on monthly chemoprophylaxis. (Used with permission of the American Heartworm Society.)
Plate 4.14 Alveoli of a cat with a chemically abbreviated heartworm infection. (Used with permission of the American Heartworm Society.)
Plate 4.15 Alveoli of a non-treated cat with adult heartworms. (Used with permission of the American Heartworm Society.)
Plate 4.16 Pulmonary artery of a heartworm-challenged cat on monthly chemoprophylaxis. (Used with permission of the American Heartworm Society.)
Plate 4.17 Pulmonary artery of a non-treated cat with adult heartworms. (Used with permission of the American Heartworm Society.)
