Foundations of Wildlife Diseases 9780520958951

Table of contents :
Contents
Preface
1. Introduction
2. Introduction to Immunity
3. Nematodes, Acanthocephala, Pentastomes, and Leeches
4. Flatworms: Trematodes And Cestodes
5. The Parasitic Insects, Mites, And Ticks
6. Kingdom Protista
7. Kingdom Fungi
8. Introduction to Non-eukaryotic Agents
9. Eubacteria
10. Viruses
11. Special Topics
12. Summary and Future Directions
Appendix One. Systematics of Major Parasites Groups
Appendix Two. Serological and Molecular Diagnostic Tests
Glossary
Index

Citation preview

Foundations of  Wildlife DiseaseS

Foundations of Wildlife DiseaseS

Richard G. Botzler and Richard N. Brown

U ni v er si t y o f C a l ifo r ni a P r e s s

University of California Press, one of the most distinguished university presses in the United States, enriches lives around the world by advancing scholarship in the humanities, social sciences, and natural sciences. Its activities are supported by the UC Press Foundation and by philanthropic contributions from individuals and institutions. For more information, visit www.ucpress.edu. University of California Press Oakland, California © 2014 by The Regents of the University of California Library of Congress Cataloging-in-Publication Data Botzler, Richard George, 1942–   Foundations of wildlife diseases / Richard G. Botzler and Richard N. Brown.   pages  cm   Includes bibliographical references and index.   ISBN 978-0-520-27609-3 (cloth : alk. paper)—ISBN 978-0-520-95895-1 (e-book)   1.  Wildlife diseases.  I. Brown, Richard N. (Richard Neal), 1958– II. Title.  SF996.4.B68 2014  639.9’6—dc23

2013048057

22 21 20 19 18 17 16 15 14 13 10 9 8 7 6 5 4  3 2 1 The paper used in this publication meets the minimum requirements of ANSI/NISO Z39.48-1992 (R 2002) (Permanence of Paper).8

contents

Preface / vii

• • •

1 Introduction / 1 2 Introduction to Immunity  /  27 3 Nematodes, Acanthocephala, Pentastomes, and Leeches  /  45

•

4 Flatworms: Trematodes and Cestodes  /  85

•

8 Introduction to Non-Eukaryotic Agents  /  241

• • • •

9 Eubacteria / 259 10

viruses / 315

11 Special Topics  /  353 12 Summary and Future Directions  /  377

Appendix One / 381

•

Appendix Two / 401

• •

Index / 429

5 The Parasitic Insects, Mites, and Ticks  /  125 6 Kingdom Protista  /  167 7 Kingdom Fungi  /  205

Glossary / 411

preface

The discipline of wildlife diseases is a dynamic field of increasing importance to a conservationconscious society. Beginning as a secondary interest among a group of wildlife managers in the early 1900s, it became a formal field of study with the formation of the Wildlife Disease Association (WDA) in 1951; at that time the WDA primarily comprised wildlife professionals trained in traditional wildlife management, as well as some veterinarians, but all sharing an interest in better understanding the role of diseases in wildlife populations. In the intervening years, our increased understanding of wildlife diseases has led to clarifying significant conservation issues; we recognize that the relationships among vertebrate wildlife, infectious agents, ecological disturbances and loss, pollutants, climate change, invasive species, and other factors are very complex. Making sense of the multitude of specific relationships can be overwhelming. Most of these problems cannot be successfully addressed by any one professional, and require networking of specialists in diverse fields to obtain better understandings. Trained conservation professionals, as well as veterinarians with a wildlife background, are essential for successfully addressing these issues. For undergraduate and graduate students interested in

wildlife diseases, we present a foundation for thinking about infective agents and their interactions with wildlife. This book is intended as a first formal introduction to the field of wildlife diseases written for upper-division and graduate students who have a good foundational grounding in biology and zoology; these include students studying wildlife and natural resources, as well as the natural and biological sciences, and veterinary students who are extending their interests to diseases of wildlife. We do not expect that students necessarily have extensive foundations in microbiology, parasitology, or other disease issues. With the ideas of this text, we hope students will come to understand the basic life history strategies used by infective agents for being successful among vertebrate wildlife, as well as understanding the mechanisms by which wildlife can defend themselves from these agents. Hosts and infective agents each constantly evolve new tools and strategies to gain an advantage over the other. We envision host–infective agent relations as a dynamic “evolutionary dance” in which each vertebrate host and each infective agent has an array of tools and strategies for gaining an advantage over the other, and for countering the tools and strategies of the other. Current wildlife disease relationships

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are the result of a perpetual natural selection for successful tools and strategies among both wildlife hosts and the infective agents. Understanding these relationships is key to more fully understanding the ecology and evolution of wildlife diseases and to laying the foundation to better understand the important contemporary and leading-edge work being conducted in the field. We use a natural history approach to provide a general introduction to wildlife diseases, pulling together information that is distributed over a wide variety of texts and professional publications into one concise source for the reader. We first give an overview and some basic definitions needed for understanding wildlife diseases. Because of its importance and applicability for all discussions of the infective agents, we next outline basic methods of defense employed by vertebrate hosts. This is followed by outlining the major life history strategies found among infective agents. We also cover some important noninfectious diseases of wildlife, and a few special problems and emerging diseases. Our primary focus among wildlife is on mammals and birds, with limited coverage of amphibians and reptiles. Arthropods and other invertebrates are addressed as contributors to important disease processes among these wildlife hosts. We believe that our emphasis on broad ecological comparisons among pathogens using virulence as a transmission strategy, those using chronic carriers as a strategy, and those relying on indirect transmission is unique to our text. For taxonomic purposes, we generally follow traditional lines of parasite classification, but we also include recent changes reflecting new evolutionary perspectives. We do not attempt comprehensive coverage of each taxonomic group, but emphasize an understanding of the variation in life history strategies among

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preface

those members within each group causing diseases among wildlife. We then present one or two examples of wildlife pathogens as illustrations of the main life history strategies in the group. For the wildlife pathogens presented, we provide a consistent organizational approach to allow readers to more easily compare key features among infective agents. The text is not intended as a descriptive survey of the most common or even the most serious wildlife diseases, a strategy often used in parasitology or medical microbiology texts. It will not compete with the specialized infectious and parasitic disease texts that play an important role in summarizing the major diseases affecting vertebrate animals. But by providing a clear foundation for understanding how wildlife and infective agents interact, our text provides an important foundation for readers to use these more specialized texts as they continue developing as wildlife disease professionals. We acknowledge, but do not develop, such contemporary and important wildlife disease topics as invasion biology and impacts on faunal structure, co-evolutionary relations, the roles of climate change and ecological disturbances as contributors to the emergence of new diseases, or ecosystem approaches to understanding the complex interactions of pathogens and the resulting patterns of disease. Rather, the text is intended to provide the foundation needed by students to thoughtfully and intelligently address these and other critical topics. Where possible, we also identify early sources of significant ideas in our discipline’s history; giving credit for the ideas allows us to appreciate the early innovative researchers, and better understand the evolution of thinking on these topics. We hope you as readers enjoy learning and reflecting on this fascinating, dynamic field of study!

ONE

Introduction

CONTENTS Why Study Wildlife Diseases Human Health Domestic Animal Health Wildlife Health

Density and Disease Disease Models

1 2 3 4

Select Definitions and Concepts 6 Health and Disease 6 Parasitism6 Diseases in Populations 9

The study of wildlife diseases encompasses the health, disease, and fitness of wildlife, and the broad range of factors that can potentially affect their well-being. These factors include a wide array of infectious organisms such as helminths, arthropods, and microorganisms, as well as toxins, traumas, metabolic dysfunctions, genetic problems, and habitat fragmentation. Such factors not only act individually, but also may interact synergistically in complex ways to affect wildlife health. In this first chapter we give a brief summary of the basis for the emergence of wildlife diseases as a discipline and provide some key general concepts used throughout this text.

10 11

Causes of Disease

12

Role of Diseases in Wildlife Populations

13

Can Diseases Regulate Wild Populations?

15

Overview and Summary

17

Literature Cited

18

Why Study Wildlife Diseases A focus on the health and well-being of wildlife themselves is of relatively recent origin. Historically, much of the initial interest in the health and diseases of wildlife stemmed from other concerns, particularly human health and the health of domestic animals (Friend 1976, Gulland 1995, Simpson 2002). In time, more direct interest emerged in the wildlife themselves, with an effort to gain a more complete biological and ecological understanding of how diseases interact with host populations; this is exemplified by the emergence of wildlife health and conservation medicine as distinctive disciplines.

1

More broadly, because parasitism is such a common mode of life, with parasites comprising the majority of species on earth (Zimmer 2000), there is considerable interest in understanding the basic biological relations parasitism entails, so as to better understand evolutionary history and ecological principles. In conventional Darwinian theory, natural selection adapts creatures to their immediate local environments through a process of specialization, operating to produce features that reduce organismal flexibility for future evolution; for example, specialization through simplification and loss of structures among helminth parasites often has been extreme (Gould 2002). Wildlife provide important models to better understand key host–parasite relationships. In recent years, an increased sense of human responsibility toward the natural world, including a greater concern for wildlife, has emerged. Two examples of these concerns are the growing interest among the public in wildlife rehabilitation, as well as concerns over environmental issues such as oil spills and other forms of pollution. In recent years, increased tensions in global political relationships and a desire for greater national and international biosecurity (Dudley 2004) have further stimulated interest in a number of wildlife diseases.

Human Health Public health risks were an early concern, and they remain a significant source of interest in the study of wildlife diseases (Beran and Steele 1994, Ashford and Crewe 2003). These interests have been focused primarily on wildlife diseases to which humans are susceptible, such as rabies, bubonic plague, avian influenza, Lyme disease, hantavirus pulmonary syndrome, and West Nile fever in North America. Such diseases transmitted between humans and other vertebrate animals are termed zoonoses (sing. zoonosis) (Soulsby 1974). We apply the term zoonosis as including (a) diseases common to both humans and nonhumans and which may be transmitted from nonhumans 2

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to humans or from humans to nonhumans, (b) diseases that are transmitted to humans and for which humans typically are a dead-end host (zooanthroponoses), and (c) diseases transmitted from humans to nonhumans and for which nonhumans typically are a dead-end host (anthropozoonoses). Interestingly, in Russian literature the opposite definitions are given for each term (Bender 2007). Estimates for the number of zoonotic diseases range from as low as 100 (Benenson 1990) to as high as 3,000 if all 2,000 serotypes of Salmonella spp. are counted as separate species (Beran and Steele 1994, Ashford and Crewe 2003), with new zoonotic pathogens being described every year. In one detailed count, an estimate of 816 zoonotic pathogens was made after omitting the numerous Salmonella spp. serotypes (Woolhouse and Gowtage-Sequeria 2005). Along with their health impacts, some zoonoses have had particularly significant sociological impacts. Bubonic plague, caused by the bacterium Yersinia pestis, is considered to have given rise to at least three major world-wide epidemics (Stenseth et al. 2008) and to be one of the greatest natural disasters in human history (Fig. 1.1). In the fourteenth century alone, it is estimated that plague caused or contributed to the death of one-fourth of the western European population and of 2 million persons in England alone (Poland et al. 1994). Equally important was the nearly complete social disruption that accompanied many plague epidemics, with consequent shifts in world politics and power (McNeill 1977). On a historical note, the old children’s nursery rhyme “Ring-around-the-Rosie” sometimes has been described as a poem stemming from plague epidemics. However, this appears to be fallacious, as the poem was not recognized until the eighteenth century, long after the most significant plague epidemics in humans; also, the poem has many variations that contain little apparent connection to plague (Munro 1996) and include one proposed to depict a smallpox epidemic (Glickman 1987). Rabies is another disease that has had a significant impact on human cultures.

human diseases have been identified as zoonotic (Lederberg et al. 1992); one estimate is that about three-fourths of the emerging diseases in humans are zoonoses (Taylor and Woodhouse 2000). Many emergent diseases in humans have followed changes in habitat or populations of wild hosts, with rodent species frequently being of special concern (Mills et al. 1994, Poss et al. 2002). Hantavirus infections and avian influenza both can produce significant human mortality and loom as potential sources of serious zoonoses. Besides giving insights into the sources and risks of zoonotic diseases to humans, the study of diseases in wildlife also may provide models for better understanding similar human diseases. Alternatively, the understanding of human diseases can provide important insights into diseases transmitted to nonhumans (anthropozoonoses), including other primates (Cranfield et al. 2002).

Figure 1.1  Plague cemetery in Nürnberg, Germany. Headstones often were laid on top of graves to prevent wild pigs and other animals from scavenging the dead (photo by R. Botzler).

The “big bad wolf” from the Little Red Riding Hood fairy tale may play on the fear many of European ancestry held for wolves (Canis lupus); this fear has been traced to a severe wolf rabies outbreak in Europe during the 1760s (Clarke 1971). However, in a more extensive study, Linnell et al. (2002) considered most wolf attacks on humans in France during the 1760s to be predatory rather than rabies-induced. The authors cite a number of reported wolf attacks on humans, especially in Finland, France, and Estonia, and more recently India, Russia, and additional regions of Asia; but they also assert that attacks by normal, healthy wolves are quite rare and unusual, and have not been reported in North America (Linnell et al. 2002). Public health concerns have played an important role in the study of many other wildlife diseases. For example, most emerging

Domestic Animal Health In addition to disease agents shared with humans, wildlife also share many disease agents with domestic animals. Rinderpest (literally “cattle pestilence”), caused by a morbillivirus closely related to the human measles virus, was considered a great scourge among domestic cattle and wildlife in Africa (Branagan and Hammond 1965, Plowright 1982) before its eradication (Anonymous 2011). The impacts of this disease on imported cattle were so severe that strenuous efforts were made to contain infections; these efforts included establishing belts of immune cattle in key habitats, and even constructing a 265-km barrier fence between Lake Tanganyika and Lake Nyassa in Africa, with a 40-km game-free strip maintained on each side of the fence by professional shooters (Plowright 1982, McCallum and Dobson 1995). This led to considerable wildlife mortality during the ongoing slaughter of animals in this game-free strip; the fence further disrupted the natural daily and seasonal migration patterns for wildlife between key habitats.

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In addition to the concern for humans, rabies also may have a considerable impact on wildlife and domestic animal populations (Beran 1994). Rabies in vampire bats (Desmodus rotundus) occurs from tropical Mexico to northern Argentina and Chile, and the disease has led to direct losses of millions of dollars annually among cattle, as well as additional losses through reduction in production and secondary infections among livestock (Kverno and Mitchell 1976). Rabid bats also have contributed considerably to public health costs for rabies control and postbite prophylaxis of humans. The environmental and ecological effects of killing vampire bats as a means of control have not been assessed. Avian cholera, caused by the bacterium Pasteurella multocida, is a serious disease among both domestic (Heddleston 1972) and many wild birds, including migratory wildfowl (Botzler 1991, Samuel et al. 2007). In North America, the first known epizootics among wildfowl occurred in Texas (Gordus 1993) and California (Rosen and Bischoff 1949, 1950) and were associated with exposure to carcasses of domestic chickens that had died from avian cholera. However, despite these historical connections and an ongoing concern that wild birds may be a source of infection for domestic birds, there is little direct evidence for consistent transmission of Pasteurella multocida between wild and domestic birds (Snipes et al. 1988, 1989; Christiansen et al. 1992). Many helminth parasites and protozoa also are shared between domestic animals and wildlife (Fig. 1.2). These include lungworms and intestinal nematodes, tapeworms and flukes among mammals (Longhurst et al. 1952, Soulsby 1968, Dunn 1969, Fraser and Mays 1986), as well as a number of intestinal parasites among domestic (Soulsby 1968) and wild (Wehr 1971) birds. Although interest in shared diseases between wildlife and domestic animals initially stemmed from veterinary concerns about the role of wildlife as sources of diseases for domestic animals, it also is important to note that wildlife can be adversely affected by diseases acquired from domestic animals. Recent epizootics in 4

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Figure 1.2  A sheep dying from effects of liver flukes (Fasciola hepatica) in habitat typical of the snail intermediate host (Courtesy of W. Frank, Universität Hohenheim, Germany).

African mammals of both canine distemper (Harder et al. 1995, Roelke-Parker et al. 1996, Carpenter et al. 1998) and rabies (Cleaveland and Dye 1995) followed transmission of these viruses from domestic dog reservoirs in conjunction with increasing encroachment of human populations on wildlife reserves (Poss et al. 2002). As an added concern, some of the emerging infectious diseases in both wildlife and domestic animals also are zoonotic (Mahy and Brown 2000, Friend et al. 2001, Daszak and Cunningham 2002, Kahn 2006). Some diseases intersect with public health, domestic animals, and wildlife. Inf luenza, caused by a myxovirus, is an example. Wild ducks, geese, shorebirds, and domestic pigs are important reservoirs for these viruses and can transmit them to domestic fowl (Slemons and Brugh 1994, Acha and Szyfres 2003, Fouchier et al. 2005). Human cases usually occur after exposure to infected domestic animals such as pigs and fowl, in which viral strains have undergone a genetic recombination (Slemons and Brugh 1994, Alexander and Brown 2000); human cases of the highly pathogenic avian influenza H5N1 have been tied to domestic fowl.

Wildlife Health While research addressing the role of wildlife diseases among humans and domestic animals has occurred over many years, a focus

Figure 1.3  An American coot (Fulica americana) dying from avian cholera (Pasteurella multocida infection). Note the convulsions and torticollis (“twisted neck”).

on wildlife diseases with a specific concern for the wildlife themselves has developed more recently, emerging first as a formal discipline about 1951, with the founding of the Wildlife Disease Association. Initially, most available information among interested professionals was transmitted through the Wildlife Disease Association Newsletter, which was replaced with the Bulletin of the Wildlife Disease Association in 1965, which, in turn, was continued as the Journal of Wildlife Diseases after 1970. Several books emerged during and after the 1960s that also helped establish wildlife disease as a distinct discipline (McDiarmid 1962, 1969; Davis et al. 1970, 1971; Davis and Anderson 1971; Page 1976; von Braunschweig 1979; Davis et al. 1981; Fowler 1981; Wobeser 1981; Edwards and McDonnell 1982; Hoff and Davis 1982; Fairbrother et al. 1996; Samuel et al. 2001; Williams and Barker 2001; Majumdar et al. 2005; Wobeser 2006; Thomas et al. 2007; Atkinson et al. 2008). Most early concerns about wildlife diseases among biologists and managers were focused on major mortality events or mortality factors affecting the management of economically important game species (e.g., waterfowl, ungulates, upland game) (Fig. 1.3). In recent years, greater emphasis has been placed on better assessing the role of parasites and diseases

on general fitness (survival, fecundity, mate selection) for all wildlife species. Concern for diseases in threatened and endangered species, and in relocation and translocation programs also has increased. Many disease investigations in conservation programs continue to be focused on high-profile species that have undergone a sudden demographic crash (Munson and Karesh 2002). Recent emerging infectious diseases of wildlife include the chytrid fungus (Batrachochytrium dendrobatidis) (Daszak et al. 2004), white nose syndrome (Pseudogymnoascus [Geomyces] destructans) (Blehert et al. 2009), and devil facial tumor disease (Hawkins et al. 2006). Emerging wildlife diseases can have a particularly severe impact on small, fragmented populations (Daszak and Cunningham 2002) and have been responsible for some local and regional extinctions (McCallum and Dobson 1995, Daszak and Cunningham 1999, Woodroffe 1999). Reasons proposed for the intensified transmission and better detection of emerging diseases include removal of geographic barriers to human and animal transport, as well as ecosystem disruption, climate change, and habitat fragmentation (Graczyk 2002). In a broader sense, devising conservation strategies that are practical in the current understanding of the “state of the Earth” will require models that address disease risks

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(Munson and Karesh 2002), including risks to humans, domestic animals and plants, wildlife, and whole ecosystems, and that further recognize that all of these groups have shared risks and cannot be viewed in isolation from the others. Such strategies call for a better understanding of the role of wildlife in these larger disease models, as well as call for policies and approaches to integrate human, agriculture, and wildlife disease studies, as addressed in the One Health concept (www.onehealth.com).

Select Definitions and Concepts Prior to detailed discussions of wildlife disease topics, it is useful to consider some basic definitions.

Health and Disease Two approaches commonly have been used to provide a conceptual base for health and disease in wildlife (Wylie 1970). In one, health is viewed as a concept analogous to temperature. As such, while there may be lower limits (death or absolute zero, respectively), there is no true upper limit for either health or temperature. In such an analogy, health is the concept to be understood and measured, and one seeks to assess how far from death an organism may be. Among humans, features used to assess health can entail physical, mental, emotional, and spiritual components. Considering just physical features, states of human health have been assessed by body mass index, body fat, erythrocyte and serum enzyme measures, and urine metabolites (Berkow and Fletcher 1992). For wildlife and other nonhuman animals, far less is understood regarding what normal physiological values constitute physical health. Measures used also may include body weights, fat indices, as well as a variety of red blood cell, serum enzyme, and urine metabolite values (Malpas 1977, Warren and Kirkpatrick 1978, Seal et al. 1981, DelGuidice et al. 1990, Harder and Kirkpatrick 1994). In contrast to viewing health on a purported linear scale, health also may be assessed as 6

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adaptability to change (Wylie 1970). Less important here is the number of parasites or problems an animal or population encounters than its response and adaptability to these stresses. One working definition of human health is the capacity to achieve socially determined goals (mental, physical, and social well-being, vigor, resilience, productivity, flourishing) within a set of socioecological constraints, only one of which is disease (Murray et al. 2002). Munson and Karesh (2002) view disease as any disorder of body functions, systems, or organs; such a disorder is not necessarily confined to one caused by an extrinsic factor such as viral or bacterial infection, infestation with parasites, or exposure to toxins (Munson and Karesh 2002). In a very broad outlook, disease has been defined simply as a reduction in fitness (Clayton and Moore 1997). For purposes of this text, we use the definition that wildlife disease is “any impairment that interferes with or modifies the performance of normal functions, including responses to environmental factors such as nutrition, toxicants, and climate; infectious agents; inherent or congenital defects, or combinations of these factors” (Wobeser 1981). Such a definition can be applied to both individuals and populations. Our primary emphasis in this text is on infectious and, secondarily, noninfectious causes of disease.

Parasitism Parasitism is a term used to describe one set of interspecific relationships along a broad continuum of relationships between species (Dindal 1975). Broadly, the term symbiology (literally, “living together”) has been used to describe the study of any persistent relationship (symbiosis) between two different species; parasitism is one part of that larger set of relationships (Read 1970). It sometimes is argued that while parasites may elude clear definition, they generally are “known primarily when we see them” (Moore 2002). Broadly, parasites are organisms living partly or completely at the expense of another

organism (its host). Parasitic lifestyles are regularly represented among four of the major kingdoms: Animalia, Fungi, Protista, and Monera (with a much less common occurrence among Plantae). Viruses raise an interesting problem; they have some features of living systems (e.g., genome, replication, evolution), but are not functionally active outside their host cells and generally are not considered to be living microorganisms (van Regenmortel and Mahy 2004); however, they commonly are treated as highly specialized parasites and included with other microorganisms in this text. Prions, infectious proteins lacking nucleic acids, also raise interesting problems and are addressed as a special topic in Chapter 11. Parasitic relationships can overlap with some forms of predation (e.g., parasitoid wasps and some fly maggots that kill their hosts) as well as certain mutualistic relationships (e.g., many intestinal bacteria). Generally, parasites live in relatively long contact with the host for part of their life cycle; in contrast, predators and prey generally have a relatively short period of contact that ends in death and consumption of the prey by the predators. Further, parasites typically relinquish the role of regulating their relationship with the external environment to the host during part of their life cycle. Endoparasites live within a host’s body during at least part of their life cycle and depend completely on the host to regulate their relationship with the external environment during that time; examples of endoparasites include some helminths, fungi, protozoa, and many infectious microorganisms. Parasites living on the exterior of the host are termed ectoparasites; these have a partial dependence on the host to regulate their relationship with the external environment when they are present on the host; examples include parasitic arthropods such as ticks, fleas, lice, and mites. Finally, parasites generally are characterized as competing with the host for its resources, and often reduce host fitness (e.g., survival, fecundity) (Clayton and Moore 1997). Many parasites are transmitted directly between susceptible hosts; these species have

direct life cycles and are termed “monoxenous” (Gr. mono 5 one, xenous 5 host). However, others, including many parasitic helminths and protozoa, require two or more hosts to complete their life cycle; these have indirect life cycles and are termed “heteroxenous.” For parasites with an indirect life cycle, the host in which the sexually mature stage of the parasite occurs is called the definitive host. An intermediate host is an additional required host for those parasites to complete their life cycle. In the intermediate host, the parasites undergo some developmental changes and may multiply, but do not reach their sexually mature stage; the intermediate host typically is a different taxonomic group from the definitive host. A less common type of host is a paratenic (transport) host, an organism which serves to transfer a larval stage or stages from one host to another but in which little or no development takes place (Anderson 1992). A paratenic host is not required for completion of the life cycle, but often is a prey species of the definitive host and facilitates completion of the life cycle; a paratenic host often is considered an “optional” host. The term vector is defined by some as any host that transmits parasites, including intermediate, definitive, and paratenic hosts (Clayton and Moore 1997); in contrast, other scholars tend to restrict the term primarily or exclusively to arthropods (Wobeser 2006). The persistent presence of a parasite in a host is termed an infection (Pratt 1963); a related term, infestation, is used to describe the persistent presence of ectoparasites. Prevalence is the number of animals infected by a parasite divided by the number of animals in the population examined, and commonly is reported as a percent value (Margolis et al. 1982, Bush et al. 1997). In contrast, incidence is the number of new hosts that become infected with a particular parasite during a specified time interval, divided by the number of uninfected hosts present at the start of that time interval (Margolis et al. 1982, Bush et al. 1997); incidence often is reported as number per 1,000 in the population. Intensity is the number

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of parasites of a particular species in or on a single infected animal; mean intensity is the total number of parasites of a particular species found in each host, divided by the number of hosts infected with that parasite (Margolis et al. 1982, Bush et al. 1997). Mortality refers to the death of a host and morbidity is the condition of having an illness, weakness, or other disability. In contrast, necrosis refers to a localized area of death in a tissue or organ rather than the death of a whole organism. Virulence is a measure of the impact of parasites upon their host. Broadly, virulence can be defined as any collective effects on host fitness, including mortality, morbidity, and reduced fecundity (Clayton and Moore 1997). Virulence also has been described more precisely as a complex property comprising three characteristics: infectivity, invasiveness, and pathogenicity (Frobisher 1962, Pratt 1963). This latter definition has been applied primarily to microorganisms and is considered here in further detail. Infectivity is defined as the ability to initiate and maintain an infection in the host (Pratt 1963). This trait is dependent on the capacity of a parasite to establish a persistent presence by evading or overcoming local defense mechanisms of the host. For example, bacteria causing plague (Yersinia pestis), typhoid fever (Salmonella typhi), or shigellosis (Shigella spp.) do not have high infectivity to laboratory workers under normal circumstances (Pratt 1963). In contrast, Francisella tularensis, the cause of tularemia, is a highly infectious bacterium readily transferred to workers within the laboratory (Pratt 1963, Hopla and Hopla 1994). Invasiveness is defined as the ability to progress further into the host from the initial site of infection (Pratt 1963). For example, many bacteria, including Pasteurella multocida, the cause of avian cholera, commonly invade the blood stream, causing a septicemia (presence of pathogenic bacteria or their toxins in the blood) among infected wildfowl. Pathogenicity is the ability to injure a host (Pratt 1963) by damaging host tissues. For larger parasites such as helminths and arthropods, 8

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Figure 1.4  Sigmoid curve typical of an LD50 test (Courtesy of National Library of Medicine. Based on original from http://aquaticpath.umd.edu/appliedtox/images=toxtutor/ chart-5.gif. Drawn by Patient Education Institute).

physical damage and blood loss are common causes of morbidity and mortality. Among bacteria, toxins affecting the nervous system, heart, or kidney are more typical causes of pathogenicity (Pratt 1963). In the laboratory, pathogenicity in living animals commonly has been measured by an LD50 test; an LD50 is the dose of an infective agent or toxin lethal for 50% of a test population (Fig. 1.4) (Reed and Muench 1938). The LD50 test has been highly regarded as a measure of pathogenicity because of its consistency and its requirement for relatively few test animals. However, the LD50 test also is limited to assessing mortality and cannot assess more subtle influences such as morbidity, reduced reproductive success, or increased susceptibility to predation; further, it also is useful only for single, isolated mortality factors and is not readily adapted to assessing synergystic relationships between two or more factors influencing mortality. In recent years, the LD50 (and the consequent killing of laboratory animals) increasingly has been replaced by alternative tests to measure pathogenicity, such as assessment of cellular pathology through the use of tissue cultures. However, it still is used in toxicology, where every registered pesticide must have at least three avian LD50 tests (A. Fairbrother, pers. comm.). In summary, for a microorganism to be considered virulent, it must simultaneously be infective, invasive, and pathogenic for a given host

(Pratt 1963). For example, Mycobacterium tuberculosis, a cause of human tuberculosis, is very invasive and highly pathogenic to guinea pigs (Cavia porcellus) in the laboratory (Wämoscher and Stöcklin 1927, Dörr and Gold 1932, Wilson and Miles 1964); yet there are no evident reports of its occurrence among guinea pigs in their natural habitats (Shope 1927, Wilson and Miles 1964, Williams 2001). Thus, while very invasive and pathogenic, an apparent absence of infectivity would result in the bacteria not being considered virulent for guinea pigs. Likewise, although Pasteurella multocida, the cause of avian cholera, is virulent to at least 180 species of birds (Samuel et al. 2007), there is no evidence that P. multocida can invade the blood stream (cause a septicemia) or cause a clinical disease among turkey vultures (Cathartes aura) (Botzler 1991, Samuel et al. 2007); thus by definition P. multocida is not invasive, and consequently not virulent, for turkey vultures. Historically, microparasite virulence was viewed as a sign of recent association between a host and parasite, and it was argued that subsequent host–parasite co-evolution would lead to a reduction of virulence and even the development of commensalism or mutualism (Burnet and White 1972). A more recent hypothesis is that virulence also can be maintained by natural selection and may increase or decrease in evolutionary response to environmental conditions or the density and behavior of hosts (Levin 1996). Thus the level of virulence expressed by parasites may result from the strategy developed by the infective agent for optimal transmission and survival (Ewald 1994). Alternatively, it also has been proposed that the virulence of microparasites is coincidental to parasite-expressed characters that evolved for other functions, or emerged as the product of short-sighted evolution in infected hosts (Levin 1996). All of these factors may play a role in different circumstances.

Diseases in Populations Several key terms are used in describing diseases in populations. The term epidemic (epi:

upon; dem: people) refers to a disease affecting many people within an area at one time, at a significantly greater occurrence than expected. Examples include outbreaks of bubonic plague and human inf luenza. The term epizootic has been used to refer to epidemics within nonhuman animals and the term epornitic occasionally is used to refer to epidemics among avian populations. The term pandemic refers to a worldwide epidemic (among humans, since most other species don’t have the same broad distribution). For example, human influenza is estimated to have caused the death of >20 million humans worldwide during the pandemic of 1918–19 (Slemons and Brugh 1994). Bubonic plague also has caused numerous pandemics among humans (McNeill 1977). In contrast, the term endemic refers to a parasite or disease with a low incidence, but one that is regularly present in a host population. Enzootic is a similar term that has been used in reference to diseases characteristic of nonhumans. For example, Yersinia pestis, the cause of bubonic plague, is maintained among some rodent populations such as the California vole (Microtus californicus) and the deer mouse (Peromyscus maniculatus) in western North America; Y. pestis is considered enzootic in these species (Poland et al. 1994). One potentially confusing aspect of the term endemic is that in ecological literature it commonly refers to a species that evolved solely in a limited area or region, as on certain islands (Van Dyke 2003); use of the alternative term “enzootic” in disease literature can help reduce that potential confusion. The terms endemic and enzootic also have been used to signify a parasite or disease characteristic of a geographic region (rather than a particular host species). Thus, plague also can be characterized as enzootic to dry grasslands, mountain meadows, and some deserts of western North American and other regions of the world. As another example, avian cholera, caused by Pasteurella multocida, regularly causes epornitics among wildfowl of North America; while found in all North American flyways, it generally is considered enzootic to

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northern California, Nebraska’s Rainwater Basin, and the Texas playa lakes (Friend 1999). The term reservoir has been used by authors in several ways. Simply stated, it can be defined as the sum of all sources of infection—the natural habitat of the parasite (Pratt 1963). More specifically, a reservoir of infection has been defined as an ecologic system in which an infectious agent survives indefinitely (Ashford 1997); such an ecologic system would encompass all of the vertebrate and invertebrate host populations and encompass the pertinent environmental factors as well as any quantitative factors, such as critical community size, needed to maintain an infectious agent indefinitely (Ashford 2003). For our use, we generally refer to Ashford’s modified 2003 definition of a reservoir. Examples of reservoirs range from red foxes (Vulpes vulpes) for rabies viruses in Europe (Rupprecht et al. 2001), to soil, mud, or water as reservoirs for the bacterium Listeria monocytogenes (Bille et al. 1999). However, the notion of reservoir for a parasite or disease may vary with geographic scale, such as that occurring within a specific watershed versus a more general assessment of the reservoir on a worldwide basis. Further, in describing reservoirs, researchers only address a limited number of aspects of the environment and thus may miss key criteria in their descriptions. The term reservoir species has been used in a specialized sense to refer to an introduced host that has artificially raised the size of the collective host populations or densities, consequently allowing pathogen transmission even when the endemic host population had been reduced below the density at which a pathogen is able to maintain effective transmission (Daszak and Cunningham 2002). Time scale is another important factor. Arthropod-borne agents of vertebrates that survive in temperate regions often survive during a period of time (e.g., cold, dry) when arthropod survival is low. In some cases the infective agent survives primarily in the vertebrate population and the vertebrates could be considered the primary reservoir. However, our use of 10

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High

A

Proportion dying or risk of infection

Low

B

Host density

High

Figure 1.5  Contrast of density dependent (A) and density-independent (B) disease impacts.

reservoir (e.g., in Chapters 9 and 10) will generally take an annual or multi-year perspective, and thus we will refer to arthropod–vertebrate reservoirs in these chapters, even where some members (e.g., arthropods) play a smaller role during some parts of the year.

Density and Disease Parasites and diseases whose risk of infection or impact varies consistently with the density (number per unit area) of their host populations are termed density-dependent, whereas those whose risk or impact do not change in response to differing densities are called densityindependent. Thus, for a density-dependent disease, the rate of transmission of a disease through a population, or risk of infection for the susceptible individual, varies consistently with the density of the host population (Fig. 1.5). Generally, parasites transmitted directly between hosts of a susceptible species, without the need for intermediate hosts or vectors, cause densitydependent diseases (Scott 1988). Rabies often is cited as a classical densitydependent disease (Macdonald 1980, Bacon 1985). One would expect that a rabid fox would have a higher probability of encountering an uninfected, susceptible fox in a high-density fox population, compared to a low-density fox population. Thus the risk of rabies transmission would be higher in a high-density than in a low-density population of susceptible animals.

In contrast, among density-independent diseases, the rate of transmission of the causative agent through a population, or risk of infection for the host, is independent of the density of the susceptible host population. Pesticides and environmental toxins, infectious diseases with reservoirs in soil or water, severe weather, and accidents tend to have density-independent impacts. For example, the risk to an individual bird of flying into a power line during a migration would not be expected to change in a consistent fashion if the flock size increased or decreased. Also, one of the devastating impacts of some pesticides is that their lethal effects are unabated even as the host populations reach very low levels (Hickey and Anderson 1968, Risebrough 1978, Peek 1986). It is important to note that the significant distinguishing feature between densitydependent and density-independent diseases is the proportion (rather than the actual number) of a susceptible population affected. For example, if the hosts in a susceptible population have a 5% risk of mortality (in a density-independent situation), one would expect approximately 5 hosts in a population 100, or approximately 50 hosts in a population 1,000, to die. Thus, while the total numbers of animals dying increases in larger populations, the actual proportion of animals affected (5%) is unchanged when transmission is density independent. Density dependence becomes more complicated with parasites undergoing indirect life cycles. For example, among parasitic helminths, the intestinal tapeworm Echinococcus granulosus involves large canids (e.g., wolves, Canis lupus; coyotes, Canis latrans) as definitive hosts and ungulates (e.g., moose, Alces alces; deer, Odocoileus spp.) as intermediate hosts. For a given season, the risk of infection to wolves by ingesting the tapeworm infective stage (hydatid cysts) is more directly dependent on the density of hydatid-infected moose than on the density of the other infected wolves (who are shedding eggs infective to moose). Likewise, in a given season the risk of moose becoming parasitized by ingesting tapeworm eggs is more directly

affected by the density of wolves shedding eggs than by the density of other hydatid-infected moose. Thus, over a shorter term, such as a single season, risks of acquiring indirect life cycle parasites tend to be density independent. However, over several seasons, one would expect that increases in wolves (shedding eggs) would lead to increases in infected moose, which, in turn, would lead to increases in the prevalence of infection among the wolves (eating infected moose); thus, over a longer term, risks of acquiring indirect life cycle parasites can become density dependent. There is some similarity for arthropodborne parasites (e.g., West Nile virus) to the relationship described above for indirect life cycle parasites. However, arthropod-borne parasite life cycles are complicated by additional factors such as the repeated feeding by some arthropods in a season, with their consequent increased likelihood of acquiring parasites. In such cases, the risk of infection to a susceptible vertebrate host is potentially dependent on both the density of infected vectors and the density of the other infected vertebrates. Likewise, the risk of infection to a vector is certainly influenced by the density of infected vertebrates, but also can be influenced by the density of other infected vectors that may infect vertebrates and make them available later in a season to the uninfected vectors. It is more difficult to break the cycle in this case as both vertebrate hosts and vectors can contribute to a condition more closely representing that of density dependence. While over a shorter term, such as a single season, arthropod-borne diseases often tend to be density independent, the influences of multiple feeding each season may lead to an additional density-dependent influence as well, over several seasons.

Disease Models We present two models to use as foundations for conceptualizing key relationships in the field of wildlife diseases. One entails

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Parasite

Environment Host Figure 1.6  Parasites, hosts, and their environments are closely intertwined—each affecting the other two in varying degrees.

an understanding of three general components: parasites and other causative agents of diseases, the affected vertebrate and invertebrate hosts, and the environment in which these occur. In this model each component interacts with the other two, and must be described and understood in relation to the other two (Fig. 1.6). Any change in one can drastically alter the balance of a resulting disease. An earlier and more complex disease model, proposed in 1939 by Soviet biologist Evgeny N. Pavlovsky, first was used to describe arthropod-borne diseases (Pavlovski 1966). It variously has been termed the Natural Nidality Doctrine of Transmissible Diseases, the Landscape Theory of Epidemiology, the Landscape Theory of Zoonotic Diseases (Pollitzer and Meyer 1961, Pavlovski 1966), and the Natural Nidality Theory (Nelson 1980). Pavlovski believed that most transmissible diseases exist in nature as discrete foci or nidi (sing. nidus, “hearth,” “home”). A nidus is defined as that portion of a region with a definite geographic character, and would be similar in usage to that of the terms biotope, ecosystem, or habitat type. A nidus can be a small local area or a broad geographic region (Pavlovski 1966). Thus, rabies in the range of infected red foxes (Vulpes vulpes) in Europe would be an example of a broad nidus; likewise, plague would be seen as having nidi among dry grasslands (steppes), mountain meadows, and some desert habitats in western North America and Asia. A nidus also can be dynamic, as in the cases of a rabies nidus among red foxes moving across Europe (Macdonald 1980), or a plague nidus shifting in North America (Barnes 1982). However, most 12

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nidi are more permanently associated with specific regions and habitat types. Within each nidus is an ecological association termed a biocoenosis, composed of the infective agent, the wild vertebrate reservoir species, intermediate hosts or vectors (typically arthropods or other invertebrates), and any wild vertebrate amplifying hosts. All of these organisms are limited in their geographic and ecological distribution by the environmental determinants of the habitat. These biocoenoses allow circulation of the parasites indefinitely and can be viewed as integrated wholes (Pavlovski 1966). Microscale disease foci are influenced by qualities of the entire ecosystem. Focal diseases in discrete sites may spread out from the nidi to cause epizootics among susceptible hosts. Susceptible wildlife, humans, or other domestic animals might become involved if they invade an active nidus. This holistic approach to the study of disease ecology has been used as a foundation for the study of plague in North America (Nelson 1980) as well as a number of other diseases (Pavlovski 1966).

Causes of Disease Diseases can be caused both by infective agents and by noninfectious causes including toxins, tumors, nutritional and metabolic problems traumas, and many others. Most diseases we address are caused by infective agents. Infective agents most commonly are living organisms and typically are classified among five major kingdoms (Whittaker 1969). Viruses and prions are additional agents that, while not considered living entities, also are infective agents (Büchen-Osmond 2003). Living organisms with a parasitic lifestyle comprise the majority of species in the world; by one estimate parasites outnumber free-living species in a four to one ratio (Zimmer 2000). Among living organisms, an initial division typically is made between eukaryotic and prokaryotic organisms. Eukaryotic organisms all have double-stranded DNA enclosed within a nuclear membrane, an endoplasmic reticulum,

mitochondria, Golgi apparatuses, lysosomes, and other cellular organelles; these organisms divide by mitosis or meiosis. Traditionally they have been distributed among the Kingdoms Protista (single-celled eukaryotes), Plantae, Animalia, and Fungi (Whittaker 1969) (Appendix 1). In contrast, prokaryotic organisms lack true nuclei and have single-stranded DNA; they also lack such organelles as mitochondria, endoplasmic reticula, Golgi apparatuses, and lysosomes (Murray et al. 1999). Prokaryotes divide by fission rather than by mitosis or meiosis. Prokaryotes include both the Archaebacteria and Eubacteria (“true bacteria”) (Murray et al. 1999) and are classified in the Kingdom Monera (Whittaker 1969) (Appendix 1). Viruses lie at the boundary between life and inert matter and are not typically included in classifications of living organisms, even though they regularly replicate, mutate, evolve, and serve as significant influences on the evolution of their hosts (Villarreal 2004). Classification schemes have emerged for viruses based on their proposed evolutionary relationships (Büchen-Osmond 2003, van Regenmortel and Mahy 2004). Recently, considerable interest has emerged in the role of infectious proteins (“prions”), which have no nucleic acids (Büchen-Osmond 2003) but are important contributors to some wildlife diseases. Views about the evolutionary relationships within and between various groups have been changing (Doolittle 1999). We note some of the recent changes in proposed classifications (e.g., Adl et al. 2005); however, because of its long and well-established history, we base our discussions on the traditional five-kingdom system (Monera, Protista, Fungi, Animalia, and Plantae) (Whittaker 1969) in this text (Appendix 1). Viruses and prions are treated as addenda to the five-kingdom system. For each major group we provide a general definition of the group, a brief description of some of its distinctive features, and a summary of any recent taxonomic changes. Although our main focus is on diseases caused by infective agents, there also is a wide

variety of noninfectious diseases to which wildlife are subject (Fairbrother et al. 1996). Two that we will address include cancers and toxins.

Role of Diseases in Wildlife Populations Disease agents function by reducing the fitness of their hosts in a variety of ways (Scott 1988). In wildlife management, factors that directly reduce wildlife numbers have been termed decimating factors, and diseases are one of many different decimating factors (Leopold 1933). In contrast, welfare factors are non-lethal factors such as a shortage of food, water, or cover that reduce wildlife reproductive success (Leopold 1933) or make wildlife more susceptible to other mortality factors such as predation, accidents, and so on; diseases also can function as welfare factors. Historically, a major focus of wildlife managers was on the role of diseases as decimating factors, especially among economically important wildlife such as ungulates, waterfowl, and upland game. These kinds of diseases often are exemplified by microparasites that undergo multiplication within their hosts. Such diseases commonly produce epizootics where waves of infection pass through populations, alternating with periods in which the pathogen disappears following a loss of susceptible hosts as they die or survive and become immune. Examples include avian cholera in wildfowl (Botzler 1991, Samuel et al. 2007), hemorrhagic diseases of deer and other ungulates (Howerth et al. 2001), or tularemia in rabbits (Mörner and Addison 2001). The 1988 epizootic of phocine distemper virus in the North Sea population of harbor seals (Phoca vitulina) is a particularly well-documented example of a decimating disease (Hudson et al. 2003). Here the parasite appeared in a series of harbor seal populations around the coasts of northern Europe, and then disappeared following a lack of new susceptible animals (Hudson et al. 2003). Disease also can serve as a welfare factor by reducing the reproductive success of susceptible animals (Gulland 1995). Among bacteria,

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Salmonella pullorum reduces the egg-laying capacity of ring-necked pheasants (Phasianus colchicus) by 75% or more, and hatched birds often are stunted and less fit (Biester and Schwarte 1965). Brucella abortus infects and causes abortions in bison (Bison bison), elk (Cervus elaphus), and other ungulates (Thorne 2001). Infections by many viruses, including members of the families Parvoviridae, Herpesviridae, Paramyxoviridae, and Orbiviridae, can result in abortion or neonatal death. Because neonatal mortality or reproductive failure resulting from infectious agents may be difficult to discern, host population size may be modulated by virus infection in the absence of measurable adult mortality (Poss et al. 2002). Diseases also can reduce the energy resources available for host immunity and lead to greater susceptibility to other parasites. Such parasites benefit when poor nutrition or other environmental conditions reduce the efficiency of the immune system, making their hosts more vulnerable (Chandra and Newberne 1977, Gershwin et al. 1985). For example, normally quiescent but opportunistic bacteria carried in the intestinal tract (e.g., Salmonella spp.) or respiratory tract (e.g., Pasteurella spp.) can cause overt disease in the presence of a compromised immune system. Also, some species experiencing diseases are more susceptible to other stresses such as cold or food shortage (Sheppe and Adams 1957), thus contributing to diminished well-being of individuals and populations. There also are interactions with malnourishment, infections, and environmental chemicals on growth and reproduction (Porter et al. 1984) Also, macroparasites commonly occur as enzootic infections, more commonly causing host morbidity than mortality. Sick animals may be less cautious and have slower reflexes than healthy animals (Poulin 1994). Such behavioral changes in animals may lead to greater susceptibility to predation or accidents. Likewise, lead poisoning (plumbism) and botulism intoxication may make waterfowl more susceptible to predation. Neurological 14

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diseases, such as canine distemper or rabies, may enhance the likelihood of some terrestrial mammals dying from highway mortality or other accidents. Sexual selection also may be influenced by parasites and diseases. For example, secondary sexual characteristics of male birds, including brightness or color and vocalizations, may signal a male’s overall well-being and freedom from parasites. Males resistant to parasites within a species may be more attractive to breeding females due to their brighter plumage, more vigorous songs, or other superior mating behaviors, compared to infected males (Hamilton and Zuk 1982, Loye and Zuk 1991, MØller 1991, Zuk 1991). Linked to these findings is evidence that parasitism may be more common among individual animals affected by developmental asymmetry of secondary sexual characteristics (MØller 1996, MØller and Swaddle 1997, Thornhill and MØller 1997). Conversely, higher parasite levels may contribute to greater asymmetry of secondary characteristics. For example, parasite-infected reindeer (Rangifer tarandus) have less symmetrical antlers (Folstad et al. 1996), and miteinfected barn swallows (Hirundo rustica) have higher levels of asymmetry in wing length and tail feathers compared to uninfected members of their respective species (MØller 1992). In turn, such levels of asymmetry could influence females seeking males for mating (MØller and Swaddle 1997) and thereby affect mating success and fitness. The cost in fitness from an infectious bacterium or virus that kills an animal or weakens it to the point where it is susceptible to predation or starvation is self-evident. The fitness costs from arthropods, intestinal nematodes, and some microparasites often are far more subtle (Hart 1997). For example, a light parasite load that may not noticeably impact a healthy, well-fed adult bird may severely affect it in times of nutritional or socially related stress, or in conjunction with the physiological demands of laying and incubating eggs, provisioning nestlings, escaping from a predator, or fighting with conspecifics

(Hart 1997). In cases where male offspring grow larger and more quickly than female offspring, parasitism can impede the ability of avian mothers to raise males, shifting the sex ratio and affecting population viability; removing their parasites allows the mothers to forage longer and rear more sons (Reed et al. 2008). Also, among polygynous species, pathogens are dispersed by infected females after the resident male dies, and the effects of pathogen-mediated dispersal increases as the harem size (number of females) increases (Nunn et al. 2008).

Can Diseases Regulate Wild Populations? Although the mortality from a disease can be dramatic, there often is little relationship between observed mortality and the effectiveness of a disease in limiting or regulating a host population. For example, avian cholera can be an explosive local disease, killing thousands of birds on a given site (Friend 1999). Approximately 37,000 birds died in one California epornitic, yet it was estimated that even such severe mortality affected only about 0.5% of the California waterfowl populations and that these losses could be recovered readily on the breeding grounds (Rosen 1972). Among waterfowl, avian cholera generally is less important than habitat destruction or hunting in limiting populations. However, there are cases where diseases can substantially influence wildlife populations, especially on initial introduction to a population. Some microorganisms can suppress wild host populations through reduced survival, reduced fecundity, or both (Scott 1988, Tompkins and Begon 1999, Hudson et al. 2003). In a classic case, myxoma virus, a poxvirus, has caused a long-term depression of European rabbit (Oryctolagus cuniculus) populations in Australia (Fenner and Ratcliffe 1965) and Europe (Ross 1982). Rabies also can temporarily suppress affected host populations (Bacon 1985). Canine distemper, a viral disease, has caused severe declines of some African lion (Panthera leo) populations (Morell 1994) as

well as near extinction of black-footed ferrets (Mustela nigripes) (Thorne and Williams 1988). Among macroparasites, there are a number of studies with evidence for helminth and arthropod parasites effectively controlling wild animal populations through reduced survival or fecundity of the hosts (Tompkins and Begon 1999). The parasites involved included two species of fleas, four species of mites, two species of bugs, one species of fly, and three species of nematodes; the affected hosts included eight species of birds and three of mammals. One of the best documented cases involves Trichostrongylus tenuis, an intestinal nematode, that helps drive population cycles of red grouse (Lagopus lagopus) in Scotland (Potts et al. 1984; Hudson et al. 1985; Hudson and Dobson 1989; Dobson and Hudson 1992; Hudson et al. 1992, 1998, 2003). Among toxins, there is strong evidence that during their regular use, dichlorodiphenyltrichloroethane (DDT) and other environmental toxins suppressed populations of raptors and fish-eating birds (Hickey and Anderson 1968). For example, use of DDT depressed peregrine falcon (Falco peregrinus) populations by reducing eggshell thickness, interfering with calcium carbonate deposition in eggshells, and altering reproductive behaviors (Enderson and Berger 1970). Significant recovery of several raptorial and other bird species occurred after banning many persistent and bioaccumulative pesticides in the United States (Anderson et al. 1975, Spitzer et al. 1978, Grier 1982, Grue et al. 1983, Bolen and Robinson 2003). Pathogens infecting a broad range of host species can cause serious problems for endangered populations (McCallum and Dobson 1995), and species-wide extinctions have been linked to diseases. For example, there is good evidence that avian malaria (Plasmodium relictum capistranoae) and avian pox (Poxviridae) have caused some population suppressions, local extirpations, and even species extinctions among native Hawaiian birds. These losses involved some complex interactions among the native hosts, introduced species of hosts, parasites, and vectors, as well as well as habitat

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(Warner 1968, van Riper et al. 1986). Interestingly, there also is recent evidence for limited species recovery among some native Hawaiian birds that did not become extinct (Woodworth et al. 2005). While it is highly likely that disease caused at least some of these extinctions, the evidence still is indirect. The first known definitive report of a parasite causing species extinction is the loss of a land snail (Pardula turgida) brought about by a microsporidian parasite (Steinhausia spp.) (Cunningham and Daszak 1998). Bighorn sheep (Ovis canadensis) introduced into Lava Beds National Monument, California, were locally extirpated from effects of Pasteurella multocida pneumonia in July 1980 following their apparent contact with domestic sheep on adjacent grazing leases (Foreyt and Jessup 1982). The response of managers to prevent this loss was complicated by political conflicts among the several federal and state agencies and ranchers with responsibilities for the animals or land. There also is evidence that local populations of prairie dogs (Cynomys spp.) can be extirpated by bubonic plague in short-grass prairies (Kartman et al. 1962, Barnes 1982). Rinderpest, a morbillivirus infection, historically caused substantial reductions among wild ungulate populations in Africa, including local extirpations of some species and significant changes in the species composition of African ungulates in many regions (Talbot and Talbot 1963, Holmes 1982, Plowright 1982, McCallum and Dobson 1995). This introduced pathogen swept through southern Africa between 1890 and 1899 and killed up to 90% of the populations of some native wild species (Plowright 1982). Rinderpest is benign in its ancient cattle host (McCallum and Dobson 1995), but highly virulent to the wildebeest (Connochaetes taurinus) and cape buffalo (Syncerus caffer), as well as introduced cattle recently exposed to this morbillivirus (Plowright 1982). Rinderpest exemplifies a disease in populations lacking past exposure or innate immunity; the causative virus infected a large proportion of the susceptible populations and mortality was high. Wild ungulates were blamed as reservoir hosts for susceptible breeds 16

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of European cattle and were slaughtered in areas around cattle ranches. However, control of rinderpest in Tanganyika wildlife through use of a vaccine in cattle in the 1950s provided evidence that cattle, rather than wildlife, played a central role as rinderpest reservoirs (Branagan and Hammond 1965). Plowright later concluded that even large populations, in excess of 100,000, of susceptible wild African ungulates were unable to sustain rinderpest infections in the absence of cattle (Plowright 1982). Following a worldwide cattle vaccination campaign to combat the disease, rinderpest was declared to be only the second disease to be eradicated on a worldwide basis, following smallpox (Anonymous 2011). Mathematical models of microparasitic diseases were developed to assess expected impacts of these diseases on their hosts (McCallum and Dobson 1995). Some generalizations that emerged are that most pathogens do not depress host population equilibria far below their disease-free carrying capacity (Anderson 1979), and that parasites highly pathogenic for individuals usually have only a minor effect on host populations. Often, if a disease is detectable at high prevalence, it probably is mild and unlikely to be a major problem to an endangered species. Also some parasites highly pathogenic in the laboratory are unlikely to cause problems in low-density populations because infected animals die before the disease can be spread. These conclusions are subject to two major qualifications. First, they apply to single-host species models, and many pathogens implicated in extinctions of one host have other reservoir hosts in which they are relatively benign (van Riper et al. 1986, Thorne and Williams 1988). Thus if a pathogen is a generalist and an endangered species is susceptible, the pathogen can cause the endangered species to decline if it has a sympatric host species (reservoir species). Second, the mathematical models assume the disease primarily increases host mortality. If the disease decreases fecundity, then diseases at high prevalence may have a significant impact on host populations without causing increased deaths (McCallum 1994); DDT had

such effects (Enderson and Berger 1970). Similar generalizations have been drawn from models of helminth and other macroparasitic infections (Anderson 1980). Where diseases affect hosts differentially, occurrence of sympatric populations of vertebrate hosts with a shared parasite can result in one host benefiting by a greater impact of the parasite on the other (Hudson and Greenman 1998). Parasite-mediated competition can act when an invading species introduces a parasite to a vulnerable resident species. One example is the likely significant impact on the native red squirrel (Sciurus vulgaris) of the introduction of a parapox virus by the introduced eastern gray squirrel (S. carolinensis) (Tompkins et al. 2002). Similarly, diseases introduced with domestic dogs have exerted significant impact on rarer and endangered indigenous species as Ethiopian wolves (Canis simensis) (Laurenson et al. 1998) and wild dogs (Lycaon picta) (Kat et al. 1995). Likewise, bighorn sheep are more susceptible to the effects of Pasteurella multocida than are domestic sheep, and the pasteurellae caused a likely extirpation of bighorns from Lava Beds National Monument (Foreyt and Jessup 1982, Foreyt 1989). Parasitemediated competition also has been proposed as a mode of action among white-tailed deer (Odocoileus virginianus) in gaining competitive advantage over moose (Alces alces) in areas of the eastern United States and Canada (Kearney and Gilbert 1976). The parasite Parelaphostrongylus tenuis is a meningeal nematode with little or no impact on white-tailed deer; in contrast, other ungulate species are far more susceptible (Lankester 2001). However, this purported role of regulating moose populations by the parasite has been controversial (Nudds 1990). In this context, it has been proposed that where a parasite species infects more than one host species, the pathogen will be least pathogenic to the host with the larger range and more pathogenic to the species with limited range; such a relationship has potentially serious impacts for rare and endangered species with limited distributions (Price et al. 1988). However, it must be recognized that parasite infections or toxins are only one of several

elements affecting host population numbers over time (Scott 1988). It often is difficult to clearly distinguish the specific role of diseases as decimating factors. It is even more difficult to document their roles as welfare factors in interactions with nutrition, stress, genetic problems, predator–prey interactions, accidents, climate, or other ancillary factors. Diseases may exert selective pressures on various social behaviors. For example, mating behaviors, social avoidance, group size, and group isolation may have been affected by selection pressures to reduce transmission of pathogens (Loehle 1995). A final, positive note is that while parasites can be detrimental to host fitness in one environment, they can be beneficial to it in another. There is some evidence that parasitized individuals may enjoy a selective advantage over unparasitized conspecific hosts in some circumstances (Thomas et al. 2000).

Overview and Summary Overall, wildlife diseases can be serious decimating factors to affected host populations. They can suppress and regulate these populations, cause local extirpations, and have been associated with species extinction. Most of the emphasis in this book will focus on the role of diseases as decimating factors. Additionally, diseases can serve as welfare factors and may reduce reproductive success or increase the likelihood of death from other causes. Diseases can influence sexual selection among hosts. Further, diseases may interact with other extrinsic factors such as nutrition and stress in their hosts, and even have been proposed as a means by which their hosts can gain an advantage over a competing host species. With this brief introduction, we are ready to begin a more detailed look into the fascinating world of wildlife diseases. We begin (Chapter 2) with a summary of the tools and strategies wildlife hosts can use in protecting themselves from the effects of various diseases they encounter, so as to be able to clearly address these defense mechanisms in

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later discussion of these diseases. In many ways, the relationships between hosts and parasites can be described as an evolutionary “dance,” a constant competition with each side seeking to respond appropriately to the moves of the other to optimize their own success and avoid a loss of fitness. Considering disease in an evolutionary context, there never are ideal phenotypes (Ewald 1994). Hosts and parasites each are full of compromises, and each is under considerable selective pressure as it evolves an optimal level of success. Many of the discomforts felt by organisms experiencing a disease (e.g., fever, diarrhea, allergies, anxiety) are connected with contemporary defense mechanisms (Ewald 1994). We then assess the major macroparasites by taxonomic group in Chapters 3 through 5. For purposes of this text, we provide the taxonomic information appropriate to a level that is most practical for an introduction to wildlife diseases; the actual taxonomic levels addressed among various parasite and host groups are not consistent. Next we address eukaryotic singlecelled organisms by basic taxonomic group in Chapters 6 and 7. Then our emphasis is upon prokaryotic and other microparasites, including bacteria and viruses, in Chapters 8 through 10. Understanding the basic life cycles and life history strategies of each parasite group as their selective pressures work to optimize their biological success, the problems they encounter, and how they overcome them, underlie an understanding of how these diseases ultimately can be managed. In Chapter 11 we address a few special topics such as noninfectious diseases, including toxins, cancers, prion diseases, and the global amphibian decline. While of more recent interest, these issues increasingly are recognized as having considerable importance for wildlife. Finally, in Chapter 12 we address specific applications and special topics, including emergent diseases, special problems, and a look at future wildlife disease studies, management, and conservation. Our goal is to provide a broader understanding of wildlife diseases from an ecological 18

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and evolutionary approach of key taxonomic groups, rather than emphasizing a clinical and pathological perspective. Hosts and parasites are constantly interacting with each other in a dynamic fashion. Each species tries to optimize its own success, and the degree of sophistication that has evolved in that process is remarkable. Methods of transmission and types of host defenses are among the more striking examples. It is a sense of this larger picture we wish to emphasize. In approaching a study of wildlife diseases, we remind readers that we work from the perspective of contemporary Western science. This approach has led to enormous advancements in human understanding of the natural world, including significant insights into the understanding of the health and diseases of wildlife. It also is a system with some limitations and often is characterized as being hierarchical, elitist, and dualistic in its approach; many argue that no one cultural worldview is privileged (Klukhohn and Leighton 1946/1962, Feyerabend 1987, Abram 1996, Nakashima 1998, Berkes 1999), and we acknowledge that Western science is only one of a number a ways of viewing an understanding of wildlife health and diseases. Of course, we also recognize the many benefits and values of using Western science as a foundation for understanding the world. Finally, the major emphasis is on disease in wildlife populations rather than a focus on individuals. The effects of wildlife diseases on individual animals is covered well in veterinary references focusing on captive wildlife (Fraser and Mays 1986, Fowler et al. 2003) as well as more general veterinary texts (Fraser and Mays 1986). There also are some valuable references covering more specialized topics (Murphy et al. 1999, Mullen and Durden 2002, Stockham and Scott 2002).

Literature Cited Abram, D. 1996. A more-than-human world. In A. Weston (editor), An invitation to environmental philosophy. Oxford University Press, New York.

Acha, P. N., and B. Szyfres. 2003. Influenza. Pp. 155–172 in Zoonoses and communicable disease common to man and animals. Pan American Health Organization Scientific and Technical Publication No. 580, Washington, DC. Adl, S. M., A. G. B. Simpson, M. A. Farmer, R. A. Andersen, O. R. Andersen, J. R. Barta, S. S. Bowser, G. Brugerolle, R. A. Fensome, S. Fredericq, T. Y. James, S. Karpov, P. Kugrens, J. Krug, C. E. Lane, L. A. Lewis, J. Lodge, D. H. Lynn, D. G. Mann, R. M. McCourt, L. Mendoza, Ø. Moestrup, S. E. MozleyStandridge, T. A. Nerad, C. A. Shearer, A. V. Smirnov, F. W. Spiegel, and M. F. J. R. Taylor. 2005. The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. Journal of Eukaryotic Microbiology 52:399–451. Alexander, D. J., and I. H. Brown. 2000. Recent zoonoses caused by influenza A viruses. Revue Scientifique et Technique Office International des Epizooties 19:197–225. Anderson, D. W., J. J. R. Jehl, W. Risebrough, L. A. Woods, L. R. Deweese, and W. G. Edgecomb. 1975. Brown pelicans: Improved reproduction off the southern California coast. Science 190:806–808. Anderson, R. C. 1992. Nematode parasites of vertebrates: Their development and transmission. CAB International, Wallingford, UK. Anderson, R. M. 1979. Parasite pathogenicity and the depression of host population equilibria. Nature 279:150–152. Anderson, R. M. 1980. Depression of host population abundance by direct life cycle macroparasites. Journal of Theoretical Biology 82:283–311. Anonymous. 2011. Joint FAO/OIE Committee on Global Rinderpest Eradication final report. Food and Agriculture Organisation of the United Nations (FAO), Rome, Italy; World Organisation for Animal Health (OIE), Paris, France. Ashford, R. W. 1997. What it takes to be a reservoir host. Belgian Journal of Zoology 127:85–90. Ashford, R. W. 2003. When is a reservoir not a reservoir? Emerging Infectious Diseases 9:3. Ashford, R. W., and W. Crewe. 2003. The parasites of Homo sapiens: An annotated checklist of the protozoa, helminths, and arthropods for which we are home. Taylor and Francis, London, UK. Atkinson, C. T., N. J. Thomas, and D. B. Hunter. 2008. Parasitic diseases of wild birds. Blackwell Publishing, Ames, IA. Bacon, P. J. (editor). 1985. Population dynamics of rabies in wildlife. Academic Press, London, UK.

Barnes, A. M. 1982. Surveillance and control of bubonic plague in the United States. Symposia of the Zoological Society of London 50:237–270. Bender, J. 2007. Chimpanzees, antibiotic resistance— Uganda (02). International Society for Infectious Diseases. . Benenson, A. S. (editor). 1990. Control of communicable diseases in man. 15th ed. American Public Health Association, Washington, DC. Beran, G. W. 1994. Rabies and infections by rabiesrelated viruses. Pp. 307–357 in G. W. Beran and J. H. Steele (editors), Handbook of zoonoses, Section B: Viral. CRC Press, Boca Raton, FL. Beran, G. W., and J. H. Steele (editors). 1994. Handbook of zoonoses, Section A: Bacterial, rickettsial, chlamydial, and mycotic. 2nd ed. CRC Press, Boca Raton, FL. Berkes, F. 1999. Sacred ecology: Traditional ecological knowledge and resource management. Taylor & Francis, Philadelphia, PA. Berkow, R., and A. J. Fletcher. 1992. The Merck manual of diagnosis and therapy. 16th ed. Merck Research Laboratories, Rahway, NJ. Biester, H. E., and L. H. Schwarte. 1965. Diseases of poultry. Iowa State University Press, Ames, IA. Bille, J., J. Rocourt, and B. Swaminathan. 1999. Listeria, Erysipelothrix, and Kurthia. Pp. 346–356 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Blehert, D. S., A. C. Hicks, M. Behr, C. U. Meteyer, B. M. Berlowski-Zier, E. L. Buckles, J. T. H. Coleman, S. R. Darling, A. Gargas, R. Niver, J. C. Okoniewski, R. J. Rudd, and W. B. Stone. 2009. Bat white-nose syndrome: An emerging fungal pathogen? Science 323:227. Bolen, E. G., and W. L. Robinson. 2003. Wildlife ecology and management. 5th ed. Prentice Hall, Upper Saddle River, NJ. Botzler, R. G. 1991. Epizootiology of avian cholera in wildfowl. Journal of Wildlife Diseases 27:367–395. Branagan, D., and J. A. Hammond. 1965. Rinderpest in Tanganyika: A review. Bulletin of Epizootic Diseases of Africa 13:225–246. Büchen-Osmond, C. 2003. Taxonomy and classification of viruses. Pp. 1217–1226 in Manual of clinical microbiology. ASM Press, Washington, DC. Burnet, F. M., and D. O. White. 1972. Natural history of infectious diseases. Cambridge University Press, Cambridge, UK. Bush, A. O., K. D. Lafferty, J. M. Lotz, and A. W. Shostak. 1997. Parasitology meets ecology on its own terms: Margolis et al. revisited. Journal of Parasitology 83:575–583.

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Carpenter, M. A., M. J. Appel, M. E. Roelke-Parker, L. Munson, H. Hofer, M. East, and S. J. O’Brien. 1998. Genetic characterization of canine distemper virus in Serengeti carnivores. Veterinary Immunology and Immunopathology 65:259–266. Chandra, R. K., and P. M. Newberne. 1977. Nutrition, immunity, and infection. Plenum Press, New York. Christiansen, K. H., T. E. Carpenter, K. P. Snipes, D. W. Hird, and G. Y. Ghazikhanian. 1992. Restriction endonuclease analysis of Pasteurella multocida isolates from three California turkey premises. Avian Diseases 36:272–281. Clarke, C. H. D. 1971. The beast of Gévaudan. Natural History 80:44–51, 66–73. Clayton, D. H., and J. Moore (editors). 1997. Host– parasite evolution: General principles and avian models. Oxford University Press, Oxford, UK. Cleaveland, S., and C. Dye. 1995. Maintenance of a microparasite infecting several host species: Rabies in the Serengeti. Parasitology 111:S33–S47. Cranfield, M., L. Gaffikin, J. Sleeman, and M. Rooney. 2002. The mountain gorilla and conservation medicine. Pp. 282–296 in A. A. Aguirre, R. S. Ostfeld, G. M. Tabor, C. House, and M. C. Pearl (editors), Conservation medicine: Ecological health in practice. Oxford University Press, Oxford, UK.  Cunningham, A. A., and P. Daszak. 1998. Extinction of a species of land snail due to infection with a microsporidian parasite. Conservation Biology 12:1139–1141. Daszak, P., and A. A. Cunningham. 1999. Extinction by infection. Trends in Ecology and Evolution 14:279. Daszak, P., and A. A. Cunningham. 2002. Emerging infectious diseases: A key role for conservation medicine. Pp. 40–61 in A. A. Aguirre, R. S. Ostfeld, G. M. Tabor, C. House, and M. C. Pearl (editors). Conservation medicine: Ecological health in practice. Oxford University Press, Oxford, UK. Daszak, P., A. Strieby, A. A. Cunningham, J. E. Longcore, C. C. Brown, and D. Porter. 2004. Experimental evidence that the bullfrog (Rana catesbeiana) is a potential carrier of chytridiomycosis, an emerging fungal disease of amphibians. Herpetological Journal 14:201–207. Davis, J. W., and R. C. Anderson (editors). 1971. Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Davis, J. W., R. C. Anderson, L. Karstad, and D. O. Trainer. 1971. Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA.

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introduction

Davis, J. W., L. H. Karstad, and D. O. Trainer (editors). 1970. Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Davis, J. W., L. H. Karstad, and D. O. Trainer (editors). 1981. Infectious diseases of wild mammals. 2nd ed. Iowa State University Press, Ames, IA. DelGuidice, G. D., L. D. Mech, and U. S. Seal. 1990. Effects of winter undernutrition on body composition and physiological profiles of white-tailed deer. Journal of Wildlife Management 54:539–550. Dindal, D. L. 1975. Symbiosis: Nomenclature and proposed classification. The Biologist 57:129–142. Dobson, A. P., and P. J. Hudson. 1992. Regulation and stability of a free-living host–parasite system: Trichostrongylus tenuis in red grouse. II: Population models. Journal of Animal Ecology 61:487–498. Doolittle, W. F. 1999. Phylogenetic classification and the universal tree. Science 284:2124–2128. Dörr, R., and E. Gold. 1932. Zur Frage der “Virulenz” der Tuberkelbazillen. Zeitschrift für Immunoforschung 74:7–24. Dudley, J. P. 2004. Global zoonotic disease surveillance: An emerging public health and biosecurity imperative. BioScience 54:982–983. Dunn, A. M. 1969. The wild ruminant as reservoir host of helminth infection. In A. McDiarmid (editor), Diseases in free-living wild animals. Symposia of the Zoological Society of London No. 24. Academic Press, London, UK. Edwards, M. A., and U. McDonnell (editors). 1982. Animal disease in relation to animal conservation. Symposia of the Zoological Society of London No. 50. Academic Press, London, UK. Enderson, J. H., and D. D. Berger. 1970. Pesticides: Eggshell thinning and lowered production of young in prairie falcons. BioScience 20:355–356. Ewald, P. W. 1994. Evolution of infectious disease. Oxford University Press, Oxford, UK. Fairbrother, A., L. N. Locke, and G. L. Hoff. 1996. Noninfectious diseases of wildlife. 2nd ed. Iowa State University Press, Ames, IA. Fenner, F., and F. N. Ratcliffe. 1965. Myxomatosis. Cambridge University Press, New York. Feyerabend, P. 1987. Farewell to reason. Verso, London, UK. Folstad, I., P. Arneberg, and A. J. Karter. 1996. Antlers and parasites. Oecologia 105:556–558. Foreyt, W. J. 1989. Fatal Pasteurella haemolytica pneumonia in bighorn sheep after direct contact with clinically normal domestic sheep. American Journal of Veterinary Research 50:341–344. Foreyt, W. J., and D. A. Jessup. 1982. Fatal pneumonia of bighorn sheep following association with domestic sheep. Journal of Wildlife Diseases 18:163–168.

Fouchier, R. A. M., V. Munster, A. Wallensten, T. M. Bestebroer, S. Herfst, D. Smith, G. F. Rimmelzwaan, B. Olsen, and A. D. M. E. Osterhaus. 2005. Characterization of a novel Influenza A virus hemagglutinin subtype (H16) obtained from black-headed gulls. Journal of Virology 79:2814–2822. Fowler, M. E. (editor). 1981. Wildlife diseases of the Pacific Basin and other countries. Fruitridge Printing, Sacramento, CA. Fowler, M. E., R. E. Miller, and Morris Animal Foundation. 2003. Zoo and wild animal medicine. 5th ed. Saunders, St. Louis, MO. Fraser, C. M., and A. Mays. 1986. The Merck veterinary manual. 6th ed. Merck & Co., Inc., Rahway, NJ. Friend, M. 1976. Philosophical considerations. Pp. 7–18 in L. A. Page (editor), Wildlife diseases. Plenum Press, New York Friend, M. 1999. Avian cholera. Pp. 75–92 in M. Friend and J. C. Franson (editors), Field manual of wildlife diseases: General field procedures and diseases of birds. U.S. Geological Survey, Biological Resources Division, Washington, DC. Friend, M., R. G. McLean, and F. J. Dein. 2001. Disease emergence in birds: Challenges for the twenty-first century. The Auk 118:290–303. Frobisher, M. 1962. Fundamentals of microbiology. W. B. Saunders Company, Philadelphia, PA. Gershwin, M. E., R. S. Beach, and L. S. Hurley. 1985. Nutrition and immunity. Academic Press, Orlando, FL. Glickman, F. S. 1987. “A ring around the rosie” (the rash that was). Journal of the American Academy of Dermatology 16:1282–1282. Gordus, A. G. 1993. Notes on the first known avian cholera epizootic in wildfowl in North America. Journal of Wildlife Diseases 29:367. Gould, S. J. 2002. The structure of evolutionary theory. Belknap Press of Harvard University Press, Cambridge, MA. Graczyk, T. K. 2002. Zoonotic infections and conservation. Pp. 220–228 in R. S. O. A. A. Aguirre, G. M. Tabor, C. House, and M. C. Pearl (editors), Conservation medicine: Ecological health in practice. Oxford University Press, New York. Grier, J. W. 1982. Ban of DDT and subsequent of reproduction in bald eagles. Science 218:1232–1235. Grue, C. E., W. J. Fleming, D. G. Busby, and E. F. Hill. 1983. Assessing hazards of organophosphate pesticides to wildlife. Transactions of the North American Wildlife and Natural Resources Conference 48:200–220. Gulland, F. M. D. 1995. Impact of infectious diseases on wild animal populations: A review. Pp. 20–51 in B. T. Grenfell and A. P. Dobson (editors),

Ecology of infectious diseases in natural populations. Cambridge University Press, Cambridge, UK. Hamilton, W. D., and M. Zuk. 1982. Heritable true fitness and bright birds: A role for parasites? Science 218:384–387. Harder, J. D., and R. L. Kirkpatrick. 1994. Physiological methods in wildlife research. Pp. 275–306 in T. A. Bookhout (editor), Research and management techniques for wildlife and habitats. The Wildlife Society, Bethesda, MD. Harder, T. C., M. Kenter, M. J. Appel, M. E. RoelkeParker, T. Barrett, and A. D. Osterhaus. 1995. Phylogenetic evidence of canine distemper virus in Serengeti’s lions. Vaccine 13:521–523. Hart, B. L. 1997. Behavioural defence. Pp. 59–77 in D. H. Clayton and J. Moore (editors), Host– parasite evolution: General principles and avian models. Oxford University Press, Oxford, UK. Hawkins, C. E., C. Baars, H. Hesterman, G. J. Hocking, M. E. Jones, B. Lazenby, D. Mann, N. Mooney, D. Pemberton, S. Pyecroft, M. Restani, and J. Wiersma. 2006. Emerging disease and population decline of an island endemic, the Tasmanian devil Sarcophilus harrisii. Biological Conservation 131:307–324. Heddleston, K. L. 1972. Avian pasteurellosis. Pp. 219–251 in M. S. Hofstad, B. W. Calnek, C. F. Helmboldt, W. M. Reid, and J. N. W. Yoder (editors), Diseases of poultry. Iowa State University Press, Ames, IA. Hickey, J. J., and D. W. Anderson. 1968. Chlorinated hydrocarbons and eggshell changes in raptorial and fish-eating birds. Science 162:271–272. Hoff, G. L., and J. W. Davis (editors). 1982. Noninfectious diseases of wildlife. Iowa State University Press, Ames, IA. Holmes, J. C. 1982. Impact of infectious disease agents on the population growth and geographical distribution of animals. Pp. 37–51 in R. M. Anderson and R. M. May (editors), Population biology of infectious diseases. Springer-Verlag, New York. Hopla, C. E., and A. K. Hopla. 1994. Tularemia. Pp. 113–126 in G. W. Beran (editor), Handbook of zoonoses. CRC Press, Boca Raton, FL. Howerth, E. W., D. E. Stallknecht, and P. D. Kirkland. 2001. Bluetongue, epizootic hemorrhagic disease, and other orbivirus-related diseases. Pp. 77–97 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Hudson, P. J., and A. P. Dobson. 1989. Population biology of Trichostrongylus tenuis, a parasite of economic importance for red grouse management. Parasitology Today 5:283–291.

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Hudson, P. J., A. P. Dobson, and D. Newborn. 1985. Cyclic and non-cyclic populations of red grouse: A role for parasitism. In D. Rollinson and R. M. Anderson (editors), Ecology and genetics of host-parasite interactions. Linnean Society Symposium Series, Vol. 11. Hudson, P. J., A. P. Dobson, and D. Newborn. 1998. Prevention of population cycles by parasite removal. Science 282:2256–2258. Hudson, P. J., and J. Greenman. 1998. Competition mediated by parasites: Biological and theoretical progress. Trends in Ecology and Evolution 13:387–390. Hudson, P. J., D. Newborn, and A. P. Dobson. 1992. Regulation and stability of a free-living host– parasite system: Trichostrongylus tenuis in red grouse. I: Monitoring and parasite reduction experiments. Journal of Animal Ecology 61:477–486. Hudson, P. J., A. Rizzoli, B. T. Grenfell, H. Heesterbeek, and A. P. Dobson (editors). 2003. The ecology of wildlife diseases. Oxford University Press, Oxford, UK. Kahn, L. H. 2006. Confronting zoonoses, linking human and veterinary medicine. Emerging Infectious Diseases 12:556–561. Kartman, L., S. F. Quan, and R. R. Lechleitner. 1962. Die-off of a Gunnison’s prairie dog colony in central Colorado. II: Retrospective determination of plague infection in flea vectors, rodents, and man. Zoonoses Research 1:201–224. Kat, P. W., K. A. Alexander, J. S. Smith, and L. Munson. 1995. Rabies and African wild dogs in Kenya. Proceedings of the Royal Society B: Biological Sciences 262:229–233. Kearney, S. R., and F. F. Gilbert. 1976. Habitat use by white-tailed deer and moose on sympatric range. Journal of Wildlife Management 40:645–657. Klukhohn, C., and D. Leighton. 1946/1962. The Navaho. Doubleday and Co., Garden City, NY. Kverno, N. B., and G. C. Mitchell. 1976. Vampire bats and their effect on cattle production in Latin America. World Animal Review 17:1–7. Lankester, M. W. 2001. Extrapulmonary lungworms of cervids. Pp. 228–278 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors). Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Laurenson, K., C. Sillero-Zubiri, H. Thompson, F. Shiferaw, S. Thirgood, and J. Malcolm. 1998. Disease as a threat to endangered species: Ethiopian wolves, domestic dogs and canine pathogens. Animal Conservation 1:273–280. Lederberg, J., R. E. Shope, and J. S. C. Oakes. 1992. Emerging infections: Microbial threats to health

22

introduction

in the United States. Institute of Medicine, National Academies Press, Washington, DC. Leopold, A. S. 1933. Game management. Charles Scribner’s Sons, New York. Levin, B. R. 1996. The evolution and maintenance of virulence in microparasites. Emerging Infectious Diseases 2:93–102. Linnell, J. D., R. A. C., Z. Andersone, J. C. B. L. Balciauskas, L. Boitani, S. Brainerd, U. Bretienmoser, I. Kojola, O. Liberg, J. Loe, H. Okarma, H. C. Pedersen, C. Promberger, H. Sand, E. J. Solberg, H. Valdmann, and P. Wabakken. 2002. The fear of wolves: A review of wolf attacks on humans. Norsk Institut for Naturforskning (NINA), Trondheim, Norway. Loehle, C. 1995. Social barriers to pathogen transmission in wild animal populations. Ecology 76:326–335. Longhurst, W. M., A. S. Leopold, and R. F. Dasmann. 1952. A survey of California deer herds: Their ranges and management problems. California Department of Fish and Game, Bureau of Game Conservation Game Bulletin No. 6. Loye, J. E., and M. Zuk (editors). 1991. Bird–parasite interactions. Oxford University Press, Oxford, UK. Macdonald, D. W. 1980. Rabies and wildlife: A biologist’s perspective. Oxford University Press, Oxford, UK. Mahy, B. W. J., and C. C. Brown. 2000. Emerging zoonoses: Crossing the species barrier. Revue Scientifique et Technique Office International des Epizooties 19:33–40. Majumdar, S. K., J. E. Huffman, F. J. Brenner, and A. I. Panah (editors). 2005. Wildlife diseases: Landscape epidemiology, spatial distribution and utilization of remote sensing technology. Pennsylvania Academy of Science, Easton, PA. Malpas, R. C. 1977. Diet and the condition and growth of elephants in Uganda. Journal of Applied Ecology 14:489–504. Margolis, L., G. W. Esch, J. C. Holmes, A. M. Kuris, and G. A. Schad. 1982. The use of ecological terms in parasitology (report of an ad hoc committee of the American Society of Parasitologists). Journal of Parasitology 68:131–133. McCallum, H. 1994. Quantifying the impact of disease on threatened species. Pacific Conservation Biology 1:107–117. McCallum, H., and A. Dobson. 1995. Detecting disease and parasite threats to endangered species and ecosystems. Trends in Ecology and Evolution 10:190–194. McDiarmid, A. 1962. Diseases of free-living wild animals. Food and Agriculture Organization of the United Nations, Rome, Italy.

McDiarmid, A. (editor). 1969. Diseases in free-living wild animals. Symposia of the Zoological Society of London No. 24. Academic Press, London, UK. McNeill, W. H. 1977. Plagues and peoples. Anchor Press, Doubleday, Garden City, NY. Mills, J. N., B. A. Ellis, J. E. Childs, J. K. T. McKee, J. I. Maiztegui, C. J. Peters, T. G. Ksiazek, and P. B. Jahrling. 1994. Prevalence of infection with Junin virus in rodent populations in the epidemic area of Argentine hemorrhagic fever. American journal of Tropical Medicine and Hygiene 51:554–562. MØller, A. P. 1991. Parasites, sexual ornaments, and mate choice in the barn swallow. Pp. 328–343 in J. E. Loye and M. Zuk (editors), Bird–parasite Interactions. Oxford University Press, Oxford, UK. MØller, A. P. 1992. Parasites differentially increase the degree of fluctuating symmetry in secondary sexual characters. Journal of Evolutionary Biology 5:691–699. MØller, A. P. 1996. Parasitism and developmental instability of hosts: A review. Oikos 77:189–196. MØller, A. P., and J. P. Swaddle. 1997. Asymmetry, developmental stability, and evolution. Oxford University Press, Oxford, UK. Moore, J. 2002. Parasites and the behavior of animals. Oxford University Press, Oxford, UK. Morell, V. 1994. Serengeti’s big cats going to the dogs. Science 264:1664. Mörner, T., and E. Addison. 2001. Tularemia. Pp. 303–312 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Mullen, G., and L. Durden (editors). 2002. Medical and veterinary entomology. Academic Press, Boston, MA. Munro, I. 1996. “Ring around the rosie” mini-FAQ. . Munson, L., and W. B. Karesh. 2002. Disease monitoring for the conservation of terrestrial animals. Pp. 95–117 in A. A. Aguirre, R. S. Ostfeld, G. M. Tabor, C. House, and M. C. Pearl (editors), Conservation medicine: Ecological health in practice. Oxford University Press, New York. Murphy, F. A., E. P. J. Gibbs, M. C. Horzinek, and M. J. Studdert. 1999. Veterinary virology. 3rd ed. Academic Press, San Diego, CA. Murray, P. R., E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken. 1999. Manual of clinical microbiology. 7th ed. American Society for Microbiology, Washington, DC. Murray, T. P., J. J. Kay, D. Waltner-Toews, and E. F. Raez-Luna. 2002. Linking human and ecosystem health on the Amazon frontier. Pp. 297–309 in A. A. Aguirre, R. S. Ostfeld,

G. M. Tabor, C. House, and M. C. Pearl (editors), Conservation medicine: Ecological health in practice. Oxford University Press Oxford, UK.  Nakashima, D. J. 1998. Conceptualizing nature: The cultural context of resource management. Nature and Resources 34:8–22. Nelson, B. C. 1980. Plague studies in California: The roles of various species of sylvatic rodents in plague ecology in California. Proceedings of the Vertebrate Pest Conference 9:89–96. Nudds, T. 1990. Retroductive logic in retrospect: The ecological effects of meningeal worms. Journal of Wildlife Management 54:396–402. Nunn, C. L., P. H. Thrall, K. Stewart, and A. H. Harcourt. 2008. Emerging infectious diseases and animal social systems. Evolutionary Ecology 22:519–543. Page, L. A. 1976. Wildlife diseases. Plenum Press, New York. Pavlovski, E. N. 1966. Natural nidality of transmissible diseases. University of Illinois Press, Urbana, IL. Peek, J. M. 1986. Pesticides and herbicides. Pp. 205–223 in A review of wildlife management. Prentice-Hall, Englewood Cliffs, NJ. Plowright, W. 1982. The effects of rinderpest and rinderpest control on wildlife in Africa. Pp. 1–28 in M. A. Edwards and U. McDonnell (editors), Symposia of the Zoological Society of London No. 50. Academic Press, London, UK. Poland, J., T. J. Quan, and A. M. Barnes. 1994. Plague. Pp. 93–112 in G. W. Beran and J. H. Steele (editors), Handbook of zoonoses, Section A: Bacterial, rickettsial, chlamydial, and mycotic. CRC Press, Boca Raton, FL. Pollitzer, R., and K. F. Meyer. 1961. The ecology of plague. Pp. 433–501 in J. M. May (editor), Studies in disease ecology. Hafner Publishing Company, New York. Porter, W. P., R. Hinsdill, A. Fairbrother, L. J. Olson, J. Jaeger, T. Yuill, S. Bisgaard, W. G. Hunter, and K. Nolan. 1984. Toxicant-disease-environment interactions associated with suppression of immune system, growth, and reproduction. Science 224:1014–1017. Poss, M., R. Biek, and A. Rodrigo. 2002. Viruses as evolutionary tools to monitor population dynamics. Pp. 118–129 in A. A. Aguirre, R. S. Ostfeld, G. M. Tabor, C. House, and M. C. Pearl (editors), Conservation medicine: Ecological health in practice. Oxford University Press, New York. Potts, G. R., S. C. Tapper, and P. J. Hudson. 1984. Population fluctuations in red grouse: Analysis of bag records and a simulation model. Journal of Animal Ecology 53:21–36.

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Poulin, R. 1994. Meta-analysis of parasite-induced behavioural changes. Animal Behaviour 48:137–146. Pratt, H. D. 1963. Epidemiology and control of vector-borne diseases. U.S. Public Health Service, Communicable Disease Center, Atlanta, GA. Price, P. W., M. Westoby, and B. Rice. 1988. Parasite-mediated competition: Some predictions and tests. American Naturalist 131:544–555. Read, C. P. 1970. Parasitism and symbiology. Ronald Press, New York. Reed, L. J., and H. A. Muench. 1938. A simple method of estimating fifty percent endpoints. American Journal of Hygiene 27:493–497. Reed, T. E., F. Daunt, M. E. Hall, R. A. Phillips, S. Wanless, and E. J. A. Cunningham. 2008. Parasite treatment affects maternal investment in sons. Science 321:1681–1682. Risebrough, R. W. 1978. Pesticides and other toxicants. In H. P. Brokaw (editor), Wildlife and America. Council on Environmental Quality, Washington, DC. Roelke-Parker, L. M. M. E., C. Packer, R. Kock, S. Cleaveland, M. Carpenter, S. J. O’Brien, A. Pospischil, R. Hofmann-Lehmann, J. Lutz, G. L. M. Mwamengele, M. N. Mgasa, G. A. Machange, B. A. Summers, and M. J. G. Appel. 1996. A canine distemper virus epidemic in Serengeti lions (Panthera leo). Nature 379:441–445. Rosen, M. N. 1972. The 1970–71 avian cholera epornitic’s impact on certain species. Journal of Wildlife Diseases 8:75–78. Rosen, M. N., and A. I. Bischoff. 1949. The 1948–49 outbreak of fowl cholera in birds in the San Francisco Bay area and surrounding counties. California Fish and Game 35:185–192. Rosen, M. N., and A. I. Bischoff. 1950. The epidemiology of fowl cholera as it occurs in the wild. Transactions of the North American Wildlife Conference 15:147–154. Ross, J. 1982. Myxomatosis: The natural evolution of the disease. Pp. 77–95 in Symposium of the Zoological Society of London No. 50. Academic Press, London, UK. Rupprecht, C. E., K. Stöhr, and C. Meredith. 2001. Rabies. Pp. 3–36 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Samuel, M. D., R. G. Botzler, and G. A. Wobeser. 2007. Avian cholera. Pp. 239–269 in N. J. Thomas, D. B. Hunter, and C. T. Atkinson (editors), Infectious diseases of wild birds. Blackwell Publishing Professional, Ames, IA.

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introduction

Samuel, W. M., M. J. Pybus, and A. A. Kocan. 2001. Parasitic diseases of wild mammals. 2nd ed. Iowa State University Press, Ames, IA. Scott, M. E. 1988. The impact of infection and disease on animal populations: Implications for conservation biology. Conservation Biology 2:40–56. Seal, U. S., L. J. Verme, and J. J. Ozoga. 1981. Physiologic values. Pp. 17–33 in W. R. Davidson, F. A. Hayes, V. F. Nettles, and F. E. Kellogg (editors), Diseases and parasites of white-tailed deer. Miscellaneous Publication No. 7. Tall Timbers Research Station, Tallahassee, FL. Sheppe, W. A., and J. R. Adams. 1957. The pathogenic effect of Trypanosoma duttoni in hosts under stress conditions. Journal of Parasitology 43:55–59. Shope, R. E. 1927. The quantity of cholesterol in the blood serum of the guinea pig as an inherited character: Its relation to natural resistance to tuberculosis, and to tuberculosis infection. Journal of Experimental Medicine 45:59–68. Simpson, V. R. 2002. Wild animals as reservoirs of infectious diseases in the UK. The Veterinary Journal 163:128–146. Slemons, R. D., and M. Brugh. 1994. Influenza. Pp. 385–395 in G. W. Beran (editor), Handbook of zoonoses. CRC Press, Boca Raton, FL. Snipes, K. P., T. E. Carpenter, J. L. Corn, R. W. Kasten, D. W. Hirsh, D. W. Hird, and R. H. McCapes. 1988. Pasteurella multocida in wild mammals and birds in California: Prevalence and virulence for turkeys. Avian Diseases 32:9–15. Snipes, K. P., D. W. Hirsh, R. W. Kasten, L. M. Hansen, D. W. Hird, T. E. Carpenter, and R. H. McCapes. 1989. Use of an rRNA probe and restriction endonuclease analysis to fingerprint Pasteurella multocida isolated from turkeys and wildlife. Journal of Clinical Microbiology 27:1847–1853. Soulsby, E. J. L. 1968. Helminths, arthropods and protozoa of domesticated animals (Mönnig). Williams and Wilkins Company, Baltimore, MD. Soulsby, E. J. L. 1974. Parasitic zoonoses, clinical and experimental studies. Academic Press, New York. Spitzer, P. R., R. W. Risebrough, W. Walker, R. Hernandez, A. Poole, D. Puleston, and I. C. Nisbet. 1978. Productivity of ospreys in Connecticut–Long Island increases as DDE residues decline. Science 202:333–335. Stenseth, N. C., B. B. Atshabar, M. Begon, S. R. Belmain, E. Bertherat, E. Carniel, K. L. Gage, H. Leirs, and L. Rahalison. 2008. Plague: Past, present, and future. Public Library of Science Medicine 5:0009–0013 (e0003). Stockham, S. L., and M. A. Scott. 2002. Fundamentals of veterinary clinical pathology. Iowa State University Press, Ames, IA.

Talbot, L. M., and M. H. Talbot. 1963. The wildebeest in western Masailand, East Africa. Wildlife Monographs 12:1–88. Taylor, L. H., and M. E. J. Woodhouse. 2000. Zoonoses and the risk of disease emergence. Poster 122 in International Conference on Emerging Infectious Diseases. Centers for Disease Control, Atlanta, GA. Thomas, F., R. Poulin, J.-F. Guégan, Y. Michalakis, and F. Renaud. 2000. Are there pros as well as cons to being parasitized? Parasitology Today 16:533–536. Thomas, N. J., D. B. Hunter, and C. T. Atkinson. 2007. Infectious diseases of wild birds. Blackwell Publishing Professional, Ames, IA. Thorne, E. T. 2001. Brucellosis. Pp. 372–395 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Thorne, E. T., and E. S. Williams. 1988. Disease and endangered species: The black-footed ferret as a recent example. Conservation Biology 2:66–74. Thornhill, R., and A. P. MØller. 1997. Developmental stability, disease and medicine. Biological Reviews (Cambridge) 72:497–548. Tompkins, D. M., and M. Begon. 1999. Parasites can regulate wildlife populations. Parasitology Today 15:311–313. Tompkins, D. M., A. W. Sainsbury, P. Nettleton, D. Buxton, and J. Gurnell. 2002. Parapoxvirus causes a deleterious disease in red squirrels associated with UK population declines. Proceedings of the Royal Society B: Biological Sciences 269:529–533. Van Dyke, F. 2003. Conservation biology: Foundations, concepts, applications. McGraw-Hill, Boston, MA. van Regenmortel, M. H. V., and B. W. J. Mahy. 2004. Emerging issues in virus taxonomy. Emerging Infectious Diseases 10:8–13. van Riper, C. H., III, S. G. van Riper, M. L. Goff, and M. Laird. 1986. The epizootiology and ecological significance of malaria in Hawaiian land birds. Ecological Monographs 56:327–344. Villarreal, L. P. 2004. Are viruses alive? Scientific American 291:101–105. von Braunschweig, A. 1979. Wildkrankheiten. Landbuch-Verlag, Gmbh, Hannover, West Germany. Wämoscher, L., and H. Stöcklin. 1927. Infektionsversuche mit enzelnen Tuberkelbazillen, durchgeführt mit der mikrurgischen Methode. Centralblatt für Bakteriologie, Parasitenkunde und Infektionskrankheiten. 1 Abteilung. Originale 104:86–90. Warner, R. E. 1968. The role of introduced diseases in the extinction of the endemic Hawaiian avifauna. The Condor 70:101–120.

Warren, R. J., and R. L. Kirkpatrick. 1978. Indices of nutritional status in cottontail rabbits fed controlled diets. Journal of Wildlife Management 42:154–158. Wehr, E. E. 1971. Nematodes. Pp. 185–233 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Whittaker, R. H. 1969. New concepts of kingdoms of organisms. Science 163:150–160. Williams, E. S. 2001. Paratuberculosis and other mycobacterial diseases. Pp. 361–371 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Williams, E. S., and I. K. Barker (editors). 2001. Infectious diseases of wild mammals. 3rd ed. Iowa State University Press, Ames, IA. Wilson, G. S., and A. A. Miles. 1964. Topley and Wilson’s Principles of bacteriology and immunity. 5th ed. Williams and Wilkins Company, Baltimore, MD. Wobeser, G. A. 1981. Diseases of wild waterfowl. Plenum Press, New York. Wobeser, G. A. 2006. Essentials of disease in wild animals. Blackwell Publishing Professional, Ames, IA. Woodroffe, R. 1999. Managing disease threats to wild mammals. Animal Conservation 2:185–193. Woodworth, B. L., C. T. Atkinson, D. A. LaPointe, P. J. Hart, C. S. Spiegel, E. J. Tweed, C. Henneman, J. LeBrun, T. Denette, R. DeMots, K. L. Kozar, D. Triglia, D. Lease, A. Gregor, T. Smith, and D. Duffy. 2005. Host population persistence in the face of introduced vectorborne diseases: Hawaii amakihi and avian malaria. Proceedings of The National Academy of Sciences of the United States of America 102:1531–1536. Woolhouse, M. E. J., and S. Gowtage-Sequeria. 2005. Host range and emerging and reemerging pathogens. Emerging Infectious Diseases 11:1842–1847. Wylie, C. M. 1970. The definition and measurement of health and disease. Public Health Reports 85:100–104. Zimmer, C. 2000. Parasite rex: Inside the bizarre world of nature’s most dangerous creatures. Free Press, New York. Zuk, M. 1991. Parasites and bright birds: New data and a new prediction. Pp. 317–327 in J. E. Loye and M. Zuk (editors), Bird–parasite interactions. Oxford University Press, Oxford, UK 

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TWO

Introduction to Immunity

CONTENTS Introduction27 Leukocytes Involved

Basis of Lymphocyte Heterogeneity Basis of Immunologic Memory

28

Innate Immunity 29 Passive Barriers 30 Antimicrobial Defenses 30 Inflammation32 Summary33 Acquired Immunity Unique Qualities of the Acquired Immune System Humoral Immunity Cell-mediated Immunity

33 33 34 35

Immunity among Different Animal Groups Comparing Acquired Immunity in Mammals and Birds Acquired Immunity in Reptiles Immunity in Amphibians Immunity in Arthropods

38 39 39 39

Behavioral Defenses in Vertebrates Future Directions Literature Cited

40 40 41

37

disease or foreign agent. The study of immunity is termed immunology. Some of the most complex and dense ideas of this book will occur in this chapter; yet an introduction to them is essential for understanding the range and complexity of vertebrate host response to protect against the effects of diseases. The functions of a host immune system are complex and often overlap. One function, of course, is host defense—to protect the body

Introduction All organisms, including humans, live in a sea of infectious or toxic agents inhabiting the air we breathe, the food we eat, and the water we drink. Yet most of us and other organisms are healthy for the majority of our lives because we can resist these agents. This resistance is called immunity, and is further defined as the condition of being protected against an infectious



37 37



27

from infectious agents. Another function is surveillance; cells of the immune system constantly find and destroy mutant (cancer) cells that continually arise in hosts. A third function is homeostasis, in which damaged or worn out parts of the body are removed to allow for replacement by new tissues. To protect hosts from infectious diseases and cancers, the immune system must be able to rapidly distinguish between cells and molecules associated with the host’s own body (termed “self”) from cells and molecules not considered part of the host body (“nonself”) (Clough and Roth 1998). The resistance by a host against infectious and toxic agents generally has been classified into two broad but overlapping physiological systems: the innate system, also called the nonspecific or natural system, and the acquired, or specific, immune system. The innate system gives a general protection and is continually in operation, or at least ready at all times to respond against foreign threats. The innate system has a fairly low level of discrimination, and essentially distinguishes “self” from “nonself” (Wakelin and Apanius 1997). The acquired immune system is activated by, and responds to, specific foreign chemicals of invading parasites or toxins, or to unusual cells such as cancer cells or parasite-infected cells. The term antigen is used to denote any chemical recognized as foreign (nonself) to the host and that stimulates an immune response or reacts with products of the immune response. The acquired immune system is capable of sophisticated discrimination and can distinguish among an almost infinite number of foreign antigens (Wakelin and Apanius 1997). This acquired system also is slow to respond and initially may require one to two weeks for an effective response the first time it encounters an antigen; however, it typically responds more quickly on subsequent exposure to the same antigen. A brief description of these major systems is provided in this chapter. Greater depth on these topics is available from a number of resources 28   introduction to immunity

(Sell 1996; Janeway and Travers 1997; Wakelin and Apanius 1997; Clough and Roth 1998; Tizard 2002, 2004; Davison et al. 2008; Parnham 2009; Tizard 2013).

Leukocytes Involved Several basic types of leukocytes (leuko: white, cyte: cell) work together to carry out the primary functions of the immune system. Among mammals these include neutrophils, eosinophils, basophils, lymphocytes, and macrophages (and their immature form, monocytes) (Fig. 2.1) (Clough and Roth 1998); birds lack neutrophils, but have heterophils instead (Davison et al. 2008). Stem cells in the host continually replicate, and their progeny gradually differentiate through several series of divisions to produce these various leukocytes, as well as erythrocytes and platelets. Leukocytes can be distinguished both by their structures and by their functions. For example, some leukocytes have irregular, lobulated nuclei with several nuclear segments connected by strands of chromatin; these cells are called polymorphonuclear (poly: many, morpho: shaped, nucleus) leukocytes and include basophils, eosinophils, as well as neutrophils in mammals and heterophils in birds. All of these polymorphonuclear leukocytes also have distinct granules in their cytoplasms and collectively may be called granulocytes. The color of these granules after staining with hematoxylin and eosin dyes has given rise to their names. Basophil granules are stained a deep blue by the basic hematoxylin dye, eosinophils granules are stained a red color by the acidic eosin dyes, and neutrophils granules are stained weakly by both the basic and acidic dyes. In contrast, lymphocytes, monocytes, and macrophages lack the distinct intracellular granules and have round or kidney-shaped nuclei (Fig. 2.1). Leukocytes also can be classified by their func­ tions. Phagocytes (phago: eating, cyto: cell) are leukocytes active in ingesting foreign material. Phagocytes include monocytes, macrophages, neutrophils, heterophils, and eosinophils.

figure 2.1  Relationships of various types of blood cells among mammals (Copyright 2009, from The Immune System, by Parham. Reproduced by permission of Garland Science/Taylor & Francis Books, LLC).

Among mammals, neutrophils, monocytes, eosinophils, and basophils, along with heterophils in birds, generally are included as part of the innate immune system. In contrast, most lymphocytes are part of the acquired immune system and circulate between the blood and lymphoid tissues in search of the specific antigens and/or target cells such as cancer cells or host cells containing intracellular parasites (Clough and Roth 1998); these are the B-lymphocytes and T-lymphocytes. However, one additional lymphocyte, the natural killer (NK) lymphocyte, is part of the innate immune system rather than the acquired immune system. It recognizes and kills abnormal cells such as tumoror virus-infected cells without any previous

exposure to them (Clough and Roth 1998, Tizard 2004).

Innate Immunity Innate immunity is a protective system involving a variety of mechanisms that are always in place and do not depend on previous exposure to infectious agents, toxins, or other foreign chemicals (Clough and Roth 1998). Elements of the innate immune system appear to have arisen early in evolutionary history and some elements are found among all multicellular organisms examined to date (O’Neill 2004). Some aspects of the innate immune system act simply as passive physical barriers introduction to immunity   29

and impediments to infection. These include intact skin and mucous membranes, the normal microbial flora living on host tissues, many fatty acids of the skin, a low stomach pH, and the cilia of the respiratory and urinary tracts (Wilson and Miles 1964). Other aspects of the innate immune system involve specific antimicrobial components. This more active aspect involves the complement system, interferon, antimicrobial peptides, phagocytic cells, and natural killer (NK) lymphocytes (Clough and Roth 1998). For example, inflammation is a specific process using many of the innate immune system tools, and in which many host cell types work cooperatively in response to infection or injury, including release of histamine. Inflammation typically results in pain, heat, redness, swelling of affected tissues, and loss of function.

Passive Barriers The skin and mucous membranes provide a very effective physical and chemical barricade to foreign agents (Wilson and Miles 1964, Janeway and Travers 1997). Very few invading organisms can penetrate intact skin, despite the important exceptions of infective stages (cercariae) of some trematodes (Bush et al. 2001) and bacteria such as Francisella tularensis, the cause of tularemia (Hopla and Hopla 1994). Hair, feathers, and scales can further reduce success of parasites penetrating host skin. If parasites do reach the skin, they encounter a variety of excreted salts, organic acids, and fatty acids that have antiparasite effects. The cilia in the mucous membrane of the respiratory and genitourinary tracts constantly work to move foreign objects out of the body. The acid pH of the stomach is very effective at killing many invading bacteria. Further, enzymes in the intestine, saliva, and tears have antibacterial effects. The normal bacterial flora of the skin, digestive tract, and other mucous membranes essentially fill those sites with microorganisms to which hosts are well adapted, and which inhibit the successful 30   introduction to immunity

colonization of those sites by invading, foreign organisms (Wilson and Miles 1964).

Antimicrobial Defenses Cytokines are a very extensive group of small proteins produced by leukocytes that act as cellular growth and differentiation factors, and also influence the immune responses of the host. Cytokines affect those cells that have specific receptor molecules for them; in host defense they often destroy target cells such as microbeinfected cells or tumor cells (Bellanti 1978, Buisseret 1982). Examples of cytokines include histamine, serotonin, prostaglandins, at least 30 types of interleukins, several types of interferons, as well as tumor necrosis factors and some growth factors; there are many others. Interleukins are cytokines that regulate the interactions between lymphocytes and other leukocytes (Tizard 2004). Many individual cytokines affect a variety of cells and organs, and many cytokines appear to be redundant in their biological activities; this complexity has given rise to the concept of a cytokine network, a web of signals among all the cell types of the immune system mediated by complex mixtures of cytokines (Tizard 2004). Cells producing cytokines include basophils, platelets, mast cells, neutrophils, macrophages, monocytes, and lymphocytes; cytokines produced by lymphocytes sometimes are called lymphokines. Macrophages secrete at least five different types of cytokines and most leukocytes secrete more than one type. The complement system is a multipleenzyme system present in the blood that rapidly responds to and attacks some infectious agents, as well as attracting other host defense mechanisms to the site of infection (Clough and Roth 1998). Most vertebrates, including amphibians, reptiles, and birds, have some type of complement system similar to that of mammals (Tizard 2004). A number of different types of complement have been identified that often operate in a sequential, cascading system (Swierkosz and Hodinka 1999). A complement system can be triggered by some microbial

surface chemicals and by some antigen– antibody complexes (Clough and Roth 1998). In the event of a bacterial infection, members of a complement system may damage bacterial membranes, attract neutrophils, increase blood flow to the site of infection (vasodilation), and also facilitate leakage of protective plasma proteins from blood vessels into the site of infection. Complement chemicals combine with antibodies already attached on the surface of the invading bacteria to make those bacteria more susceptible to phagocytosis; this process is called opsonization (Hayden 1995, Swierkosz and Hodinka 1999). Complement sometimes interacts with disease agents in such a way as to cause harm to the host as well (Leslie 2012). Interferon is a group of chemicals among the cytokines tied to antiviral and anticancer defense. Some interferons are produced by virus-infected macrophages, dendritic cells, or fibroblasts, and bind to uninfected host cells in a way that prevents virus replication from occurring in those cells (Clough and Roth 1998). Other interferons are produced by lymphocytes and regulate the immune response by stimulating T-lymphocytes, macrophages, and neutrophils, as well as influencing the classes of antibodies that B-lymphocytes secrete (Clough and Roth 1998). Basophils release histamine and other vasoactive substances, and are active in moderating the process of inflammation (Wedemeyer et al. 2000). Mast cells sometimes are considered “chemical factories” that produce a multitude of compounds; these include histamines to make the blood vessels leaky, protein-slicing enzymes such as chymase and tryptase, and other cytokines to incite inflammation and attract other leukocytes. However, mast cells also are responsible for many allergic reactions, and thus can have adverse effects on the host (Leslie 2007). Though they are derived from similar progenitor cells, basophils and mast cells undergo a different development history; basophils (like other granulocytes) circulate in peripheral blood, whereas mast cells do not circulate in blood but complete

their differentiation in vascularized tissues (Wedemeyer et al. 2000). Although not addressed in this summary, there are many animal antimicrobial peptides having protective effects against disease agents. These gene-encoded antimicrobial peptides have significant roles in animal defense systems at all taxonomic levels (Andreu and Rivas 1999). Another important component of the innate immune system is phagocytosis, the process whereby some leukocytes ingest foreign particles. The major types of phagocytic cells among mammals are neutrophils, eosinophils, monocytes, and macrophages (Tizard 2004); in birds, heterophils function in place of neutrophils (Andreasen et al. 1991). Neutrophils contain several types of chemicals important for killing microorganisms and breaking down complex chemicals; for example, lysosomes are intracellular granules of neutrophils containing lysozyme, a very effective chemical for rupturing bacterial cell walls. Neutrophils are attracted by a number of cytokines and complement to injured and infected sites of the host; in turn, they also produce some cytokines that influence host responses. Eosinophils are phagocytic, but less efficient than neutrophils or mononuclear cells. Their functions are not fully clear, but they appear to ingest antigen–antibody complexes, remove products of mast cell degranulation, and attack multicelled parasites, (Meeusen and Balic 2000, Klion and Nutman 2004) They also limit inflammatory reactions by inactivating histamine and inhibiting edema. Eosinophils have receptors on their membranes for a type of antibody called immunoglobulin E, and there is evidence that eosinophils kill some helminth parasites after binding to them; eosinophils also detoxify some inflammationinducing substances released by the mast cells and basophils (Clough and Roth 1998). Eosinophils are attracted to a site by a variety of cytokines, including histamine, as well as antigen–antibody complexes. There are two closely related mononuclear cell types: monocytes and macrophages. After introduction to immunity   31

their production from stem cells, monocytes circulate in the blood for a few days and then migrate to the tissues to develop into macrophages. Both cell types remove and destroy invading organisms, including those that multiply intracellularly, and also attack damaged and worn out cells and tumor cells; their role in removing tumor cells is considered very important (Schultz et al. 1977). Like neutrophils, monocytes and macrophages also are attracted to an infected site by cytokines and complement, and they are further stimulated to phagocytize when an object is coated with antibody, complement, or with a chemical combination of antibody and complement (Tizard 2004). Phagocytic cells such as neutrophils and monocytes are important for providing a rapid response to reduce the impact of bacterial infections while the slower humoral and cell-mediated immune systems of the host develop their responses (Clough and Roth 1998); they are particularly effective against extracellular bacteria (Baxt et al. 2013). Neutrophils are attracted to a number of chemicals produced by infecting bacteria. After ingesting these invading bacteria, neutrophils kill the bacteria with lysozyme and other chemicals they carry. However, as the neutrophils themselves often die at a site of infection, the lysozyme and other toxic chemicals they produce may spill out and exacerbate the damage to local host cells. Bacterial cells also have several mechanisms to evade phagocytosis (Baxt et al. 2013). Natural killer (NK) cells, another defensive cell, are a type of small lymphocyte that can recognize and kill some virally infected cells and cancer cells; they also play an important role in tumor rejection. These NK cells are so named because they are active naturally without previous exposure to antigen (Clough and Roth 1998). Natural killer cells also have receptors for antibody molecules and thus can bind to and kill antibody-coated cells in a process called antibody-dependent cell-mediated toxicity; when NK cells are in the process of actively detecting and killing antibody-coated 32   introduction to immunity

cells they also have been called killer cells (Clough and Roth 1998).

Inflammation Inflammation is the host response to tissue damage or invading microorganisms, and is a means by which defensive cells and molecules are concentrated rapidly at injured or infected host sites (Tizard 2004). The innate immune system recognizes generic classes of foreign molecules produced by a variety of pathogens. When such foreign molecules are detected by a host, the innate system triggers an inflammatory response in which various cells of the immune system attempt to wall off the invader and halt its spread (O’Neill 2005). An important role of inflammation is facilitating the movement of defensive cells from the blood stream into sites of infection or injury (Tizard 2004). Inf lammation also involves the process of returning an injured site to a condition of homeostasis and may include scar formation, the replacement of dead cells with connective tissue, and regeneration of some cells such as skin. Inflammation is a very complex array of responses coordinated primarily by the innate immune system. Its overt physical features traditionally are described as redness, swelling, heat, pain, and loss of function. Much work of the innate immune system contributes to both enhancing and eventually reducing the inflammatory process. In addition to the benefits provided to the host, excess activity by complement, and leukocytes such as neutrophils and mast cells, also may cause tissue damage and exacerbate problems for the host (Tizard 2004). The inflammatory response is initiated by Toll-like receptors (TLRs), an ancient family of proteins that mediate innate immunity in a wide variety of vertebrates and invertebrates; these TLRs are made by many leukocytes of the innate system (O’Neill 2005). Most TLRs recognize molecules important to the survival of bacteria, viruses, fungi, and other parasites, such as bacterial lipopolysaccharide, the lipoteichoic

acid found in bacterial cell walls, a protein of bacterial flagella, and the genetic material of viruses (O’Neill 2004). Approximately 10 different TLRs appear to protect humans from virtually every know pathogen (O’Neill 2004). Besides the specialized immune cells, there also is the capacity of most cells to defend themselves against infections. This host protection is termed cell-autonomous immunity and operates across all domains of life (Randow et al. 2013).

Summary Collectively, the various cells and processes of innate immunity are part of a host’s first line of defense against infective agents and play a powerful protective role for hosts. Innate immunity also interacts extensively with the acquired immune system, helping to determine the antigens to which the acquired immune system responds and the nature of that response (Fearon and Locksley 1996).

Acquired Immunity The acquired immune system is mediated by lymphocytes; it is most developed in vertebrates and best studied in mammals. Among mammals, common lymphoid progenitors (Fig. 2.1) are produced in the bone marrow and circulate through the blood stream as undifferentiated lymphocytes. Some migrate to the thymus gland, where they undergo a rigorous but incompletely understood “maturation” process and emerge as T (thymus-derived)-lymphocytes. Others migrate to the bursa in birds, or to bone marrow or Peyer’s patches in mammals (Tizard 2004), where they undergo a comparable maturation process and emerge as mature B (bursa/ bone marrow-derived)-lymphocytes. Both Band T-lymphocytes have distinctive sites in the lymph nodes which they inhabit. Acquired immunity has two key components: humoral immunity and cell-mediated immunity (Bartl et al. 1994). Humoral immunity is based on B-lymphocytes that can respond

to foreign proteins (antigens) by producing specific serum proteins (antibodies) against the antigens. When these specific antibodies chemically combine with a specific antigen, this combination often results in the host being protected from adverse effects of that antigen or the parasite on which it occurs. Antigens often are large and complex chemicals. The specific chemical moieties of an antigen that stimulate antibody production are called epitopes; each antigen can have multiple epitopes. Antibodies are defined as immunoglobulins. In general, mammals carry three chemical forms of globulins: a-globulins, b-globulins, and g-globulins. Immunoglobulins all are g-globulins. Cell-mediated immunity is based on T-lymphocytes and is particularly effective in destroying parasite-infected cells and cancer cells. The T-lymphocytes can act by directly destroying target cells or by releasing cytokines which, in turn, destroy the target cells or influence other host cells to enhance immune responses. Activities of the cell-mediated immune system overlap with those of the humoral immune system.

Unique Qualities of the Acquired Immune System There are some important and distinctive characteristics of the acquired immune system. One is a very high level of specificity. For example, a single B- or T-lymphocyte will respond to only one specific epitope. Because there are likely millions of different types of antigens a host can encounter, there must be millions of different types of antibody-producing B-lymphocytes in mammalian hosts, and a comparable number of T-cells. Consequently there is considerable heterogeneity among both B- and T-cells. Thus, although the immune response to antigens is highly specific, the immune system also has the potential to respond to a wide diversity of different antigens and their epitopes and, consequently, to their associated parasites and toxins. introduction to immunity   33

en ite tig g s n a in nd bi

bi an nd tig in en g si te

variable constant

light chain

heavy chain

figure 2.2  Diagram of an antibody (redrawn with permission of N. Anderson, University of Arizona).

A third important feature of the acquired immune system is an immunological memory. After the first exposure to an antigen, called a primary response, there is a 12- to 14-day lag before antibodies are detectable. For the hosts surviving this first exposure, a second, later exposure to that same antigen results in a shorter time span, often about 2 days, before specific antibodies to that antigen are detected. This is called a secondary response. Also, far greater numbers of those specific antibodies are produced in a short period in the secondary response. Thus, once a host has been exposed to a specific antigen, it develops an immunological memory for that antigen and responds much more quickly and aggressively to its presence. The features of specificity, heterogeneity, and memory are characteristic for both the humoral and cell-mediated systems of acquired immunity.

Humoral Immunity In their basic form, antibodies consist of four polypeptides—two heavy chains and two light chains that are joined to form a “Y”-shaped molecule (Fig. 2.2). The tips of each “Y” are highly variable and consist of 110–130 amino acids, which vary to give the antibody its specificity for binding an antigen. This variable region includes the ends of both light and heavy chains. 34   introduction to immunity

Each B-lymphocyte has about 200,000 to 500,000 antigen receptor sites on its surface for a specific antigen (Tizard 2004). These surface receptor sites resemble antibodies. As foreign materials invade a host, some make their way through the blood to the lymph nodes, where they come in contact with many leukocytes, including monocytes, macrophages, and lymphocytes. The B-lymphocyte receptors can bind to free, soluble antigen. However, to be fully activated, the B-cell also requires co-stimulation by a specialized T-lymphocyte called a helper T-cell that has been presented with that same antigen (Tizard 2004). When an antigen (or a specific epitope) attaches to a receptor site on the surface of the specific types of B-lymphocyte with which it can bind, that clone of B-lymphocytes is stimulated to multiply; each of the progeny also produces quantities of that specific antibody. Thus, humoral immunity is a result of clonal selection, the selection for the specific B-lymphocyte type that will produce antibodies against a particular epitope (antigen) among the many clones of B-lymphocytes in the host. B-lymphocytes actively producing antibody also are called plasma cells. Of course, parasitic organisms are not composed of a single antigen; rather, their surfaces are a collection of many different macromolecular chemical groups. Thus, a host will recognize a number of these different chemical groups (epitopes) and will produce a polyclonal antibody response to each parasite. Once antigen recognition has occurred, B-lymphocytes proliferate to produce clones of B-cells (plasma cells) bearing identical receptor sites, each actively engaged in antibody production. Memory cells are a subset of B-lymphocytes that may remain sequestered in the host for months to years after an earlier response to an antigen and provide a quick and enhanced response upon later exposure to the same antigen (Wakelin and Apanius 1997). Hosts also can produce a number of different immunoglobulin classes having the same antigen specificity (Bush et al. 2001). Mammalian immunoglobulins (Ig) are divided into five major

classes, IgM, IgG, IgA, IgD, and IgE, based on their chemical structure and immune function. Immunoglobulin M first evolved in cartilaginous fishes and is found in amphibians, reptiles, birds, and mammals; it is often the first antibody to respond to an antigen (Tizard 2004). In contrast to most other antibodies, its shape is a pentamere with five “Y” structures fused at the base of the “Y”; IgM stimulates macrophages to engulf those objects it coats. This antibody also damages bacterial membranes; in combination with complement molecules, IgM produces holes in bacterial cell membranes and walls. However, the plasma cells soon convert from IgM to IgG production in mammals. Immunoglobulin G, found only in mammals, is the most abundant antibody present in the mammalian circulatory system (Tizard 2004). It is very effective against blooddisseminated microorganisms (septicemia) and is also able to cross the placental barrier in mammals. The avian equivalent of this immunoglobulin is IgY (Tizard 2004). Immunoglobulin A is reported in birds and mammals and provides a barrier against pathogenic organisms entering the host body; IgA-producing B-lymphocytes are found on the linings of intestines, as well as in the respiratory and genitourinary tracts, saliva, tears, and perspiration (Tizard 2004). This immunoglobulin occurs in high concentrations in the colostrum of mammals; colostrum is the first milk produced immediately after birth. Thus, nursing females transmit a rich supply of protective antibodies and a quick, but temporary, immunity to offspring. Immunoglobulin A is locally synthesized in mammary tissues, although many IgA-producing cells in the mammary gland are derived from precursors originating in the intestine; these intestinal cells are a source of antibodies against intestinal pathogens (Tizard 2004). Immunoglobulin E is known to occur only in mammals; IgE, like IgA, is made by plasma cells located beneath body surfaces (Tizard 2004). While its function is not entirely clear, IgE may help initiate the inflammatory response. Immunoglobulin E binds to the surface of mast

cells and basophils, leading to histamine release when a specific antigen combines with the IgE; its presence also is associated with allergic reactions such as hay fever and asthma in humans. It may function as a defense against helminths or arthropods (Tizard 2004). Immunoglobulin D is known to occur in humans, some other mammals, certain fish, and some amphibians and reptiles, but not in birds (Tizard 2004, Davison et al. 2008). Immunoglobulin D may be the form of antibody attached directly to B-lymphocytes as the antigen receptor; it rarely is found free in the blood (Tizard 2004).

Cell-Mediated Immunity The cell-mediated immune system is based on T-lymphocytes and is particularly effective in destroying parasite-infected cells and cancer cells. The T-lymphocytes can act by directly destroying target cells or by releasing cytokines which, in turn, destroy the target cells or influence other host cells in ways to enhance immune responses. There are several types of T-lymphocytes, distinguished by the type of antigen receptors they carry, their accessory molecules, and their functions (Tizard 2004). These include two types of helper T-cells, regulatory T-cells, cytotoxic T-cells, and natural killer (NK) cells. Helper T-lymphocytes respond to antigens by secreting a variety of cytokines that influence the responses and activities of all other leukocyte types (Clough and Roth 1998). One type of helper T-cell collaborates with B-lymphocytes to enhance antibody formation; another induces a cell-mediated immune response and assists the direct cell-to-cell contacts associated with cellmediated immunity (Clough and Roth 1998). Regulatory T-cells can suppress some immune responses. For example, they can suppress the proliferation of helper T-cells in response to antigens and prevent inappropriate T-cell activation in the absence of antigens (Tizard 2004). Cytotoxic T-lymphocytes respond to foreign antigens presented on cell surfaces by killing the cells that produced the antigens (Clough and Roth 1998). Cytotoxic T-lymphocytes differ introduction to immunity   35

T cell TCRɑ chain

TCR β chain

peptide from infectious agent

MHC protein infected cell

figure 2.3  Model of an infected antigen-processing cell with a Major Histocompatibility Complex (MHC) protein presenting a foreign peptide to a T cell receptor (TCR) (From Bartl et al. Molecular evolution of the vertebrate immune system. Proceedings of the National Academy of Sciences 91: 10769-10770. Copyright 1994, National Academy of Sciences, USA).

from the natural killer (NK) cells discussed in the innate immune system in that the cytotoxic T-cells respond to specific antigens, whereas NK cells do not require previous exposure to a specific antigen to kill virus-infected or cancerous cells. Cancerous host cells or host cells infected with intracellular microorganisms typically will have foreign peptides on their surface that attract cytotoxic T-lymphocytes (Clough and Roth 1998). Cytotoxic T-lymphocytes use specialized T-cell receptors on their surfaces to attach to peptides from infectious agents that have been bound to proteins on the surface of infected cells or tumor cells (Fig. 2.3) (Bartl et al. 1994). The T-cells can kill these infected or tumor cells by binding molecules on the surface of these cells; this binding often initiates dissolution of the nucleus or disrupts the fluid equilibrium of the cell (Clough and Roth 1998). In the process, the T-lymphocytes also may cooperate with phagocytes or B-cells in destroying these target cells. Thus, T-lymphocytes, B-lymphocytes, and the innate immune system all can work in a cooperative fashion. Some T-lymphocyte cytokines may regulate the activity of other T-lymphocytes, as well as B-lymphocytes. Depending on the specific cytokines released, T-lymphocytes can inhibit the response of helper T-lymphocytes involved in the antibody response and inhibit proliferation of the cytotoxic T-lymphocytes (Bush et al. 2001). They also may induce and influence the inflammatory responses by regulating the production, migration, and functional activation 36   introduction to immunity

of granulocytes and monocytes (Wakelin and Apanius 1997). Different parasites selectively stimulate different helper T-lymphocytes subsets; each T-lymphocyte subset secretes a characteristic set of cytokines, referred to as a T-lymphocyte cytokine profile (Bush et al. 2001). During chemical breakdown and processing of antigens, the resulting peptides become bound to major histocompatibility complex (MHC) receptors on the surface of the antigenprocessing cells (Tizard 2004). The MHC proteins occur on the membranes of host cells and are highly pleomorphic between individuals in a species; they are required in the presentation of processed peptide antigens to T-lymphocytes (Clough and Roth 1998). T-lymphocytes use their T-cell receptors to respond to antigen in the form of processed peptides bound to the cell surface proteins encoded in the MHC. The importance of MHC molecules in immune responses against pathogens led to the proposal that the high diversity of MHC molecules evolved as a result of parasite interactions (Snell 1968, Doherty and Zinkernagel 1975). Further, there is evidence that parasites with a long history of co-evolution with their hosts have had the greatest impact on this MCH polymorphism (Klein and O’Huigin 1997). For T-lymphocytes, two types of receptor molecules are required for recognition of antigen. First, the MHC molecule on the antigen-processing cell (e.g., dendritic cell) binds the processed antigenic peptide; second, the T-cell receptor binds the MHC molecule–antigenic peptide complex (Tizard 2004). The MHC receptor molecules appear on the surface of dendritic or other antigen-presenting cells (Fig. 2.3). It is only when antigenic fragments are presented by an MHC receptor in this way that they can be recognized by specific T-lymphocyte receptors or initiate specific B- or T-lymphocyte immune responses (Doherty and Zinkernagel 1975, Wakelin and Apanius 1997). This dependence on correct MHC-mediated presentation is called MHC restriction and constitutes a potential bottleneck for recognition of parasite antigens (Zinkernagel and Doherty 1979, 1997). While processing of complex antigens such as those derived from

parasites may produce many epitopes, most subsequent immune response is directed against a relatively small number of “immunodominant epitopes” (Wakelin and Apanius 1997). Both B- and T-cells have similar types of specificity, heterogeneity, and memory. Thus, each T-lymphocyte also responds to a single epitope and, following combination with a specific antigen, there is a clonal proliferation of these cells. Cellular immunity involves the presentation of antigenic epitopes in combination with specific macrophage surface molecules (the major histocompatibility proteins) to the T-lymphocytes.

Basis of lymphocyte heterogeneity A mammalian acquired immune system requires antigen receptors (immunoglobulins and T-cell receptors), antigen-presenting molecules (MHC), and gene-rearranging proteins (Fig. 2.3) (Bartl et al. 1994). It is estimated that mammals can produce up to 1015 different antigen receptors to be expressed on B- and T-lymphocytes, but that in order to produce this enormous diversity they use fewer than 500 genes (Tizard 2004). Initially, stem-cell lymphocyte precursors all are alike in genetic composition within an individual host. In mammals these precursor cells undergo a maturation process in the bone marrow, Peyer’s patches, or thymus whereby their genetic compositions are altered so that they encode unique receptor proteins and each becomes able to recognize and respond to a unique epitope. The key to generating receptor diversity of gene-rearranging proteins lies in the fact that multiple genes are required to code for each receptor peptide chain. Several genes code for each variable region of the B-cell or T-cell receptors, whereas one gene codes for the constant region. As a result, a single constant-region gene can be combined with any one of a large number of different variable-region genes to make a complete receptor peptide chain. Thus, instead of having to store information about all possible receptor chains, it is only necessary to store the information (genes) for all the variable domains and to match these, when required,

with the appropriate constant-region gene to produce a complete range of receptors. Following this, light and heavy chains may be paired in different combinations, a process called combinatorial association (Tizard 2004). The development of lymphocyte heterogeneity can be seen as analogous to shuffling a deck of several thousand cards and then randomly distributing a limited number of cards (genes) among many players (lymphocytes). There are a huge number of possible combinations of genes, with little likelihood of identical combinations. And in the case of immunity, there are three separate gene segments from which one is selecting and distributing genes (Bartl et al. 1994).

Basis of immunological memory Immunological memory is based on differential responses by B- or T-lymphocytes of a clone encountering a specific antigen. Most B- or T-lymphocytes that respond to a specific antigen multiply, produce antibodies (B-cells), or attack target cells (T-cells), and die. However, a few cells in each of these clones remain quiescent; these long-lived “memory cells” remain ready to respond to a subsequent exposure to the same antigen. Length of immunological memory is greatly influenced by the nature of the antigen. Although some antigens, such as mumps and smallpox in humans, may elicit a lifelong immunity, many others (e.g., rabies) have an effective immunity of a few years. In contrast, botulism toxin results in a short memory or none whatsoever, based on recovered birds that were banded and released, and succumbed to the disease the following year. It is possible that a sublethal dose of toxin may have too little antigenic mass to stimulate an antibody response.

Immunity among Different Animal Groups All animals, both invertebrates and vertebrates, possess innate immune defenses triggered by tissue damage or microbial invasion. introduction to immunity   37

The acquired immune system evolved only after the emergence of jawless fishes; thus acquired immune mechanisms evolved only in the more recently evolved vertebrates (Tizard 2004).

Comparing Acquired Immunity in Mammals and Birds Although both mammals and birds have complex immune systems, there are some important differences between them (Wakelin and Apanius 1997, Davison et al. 2008). Like mammals, birds are hosts for virtually every group of parasites, from viruses to helminths and arthropods (Wakelin and Apanius 1997). And like mammals, avian immune systems are characterized by a true two-component (B- and T-lymphocyte) system, including specific immunoglobulin antibody production, highly developed specific cellular immunity, and specific memory. However, birds do not have lymph nodes, but have a central sinus that is the main lumen of a lymphatic vessel, which contains germinal centers functionally equivalent to mammalian lymph nodes (Tizard 2004). The thymus is the source of T-lymphocytes for both mammals and birds; it lies along the jugular vein in birds but overlies the heart in mammals. Rather than relying on bone marrow for differentiation of B-cells, birds have a discrete organ, the bursa of Fabricius, located near the cloaca, that provides a microenvironment for the differentiation and expansion of the B-cell compartment (Wakelin and Apanius 1997). The bursa also is the source of erythrocyte production in birds. Based on the immunoglobulins of domestic galliform birds, there are three principal immunoglobulin classes in birds: IgY (similar to IgG of mammals), IgM, and IgA (Tizard 2004). Thus far, IgE and IgD have not been reported in birds (Davison et al. 2008) Avian IgM is homologous to the IgM from other vertebrate classes (Wakelin and Apanius 1997). Birds produce primary and secondary responses similar to mammals, although the predominance of IgM in the primary 38   introduction to immunity

response and of IgY in the secondary response is less marked in birds than in mammals. Immunoglobulin Y is the functional equivalent of IgG in birds, reptiles, and amphibians, and it is viewed as the ancestor to the uniquely mammalian antibodies IgG and IgE (Warr et al. 1995). Avian IgY differs from mammalian IgG in possessing an additional polypeptide domain (Wakelin and Apanius 1997). Because of the structural differences from mammalian IgG, many authors prefer to use the term IgY in place of avian IgG (Wakelin and Apanius 1997, Tizard 2004). Mammalian IgG has several subclasses, but this is not clearly established for avian IgY (Benedict and Yamaga 1976, Tizard 2004). A functional analogue of mammalian IgA has been identified in galliform and columbiform birds (Porter and Parry 1976, Goudswaard et al. 1977). However, the evolutionary relationships between IgA of birds and those of mammals are not clear (Hädge and Ambrosius 1983, 1984). Ducks, geese, and swans (Anseriformes) have some features distinct from other avian groups. The morphology of the bursa differs from other avian orders (von Rautenfield and Budras 1982). Some species of ducks have multiple copies of the gene for antigen-binding sites of immunoglobulins, in contrast to the single copy found in other orders (McCormack et al. 1989). Anseriforms have a very uncommon form of immunoglobulin isotype called IgN that has only two of the four polypeptide domains that compose the IgY heavy chain (Magor et al. 1992, Tizard 2004). Thus, IgN has a lower molecular weight than IgY and reduced effectiveness for antibody activities such as agglutination and complement activation (Higgins 1989). Anseriforms secrete an IgM isotype into bile for protection of mucosal surfaces in the intestine (Ng and Higgins 1986), whereas galliforms and columbiforms secrete an IgA isotype into bile, tracheal mucus, and tears (Wakelin and Apanius 1997). Thus, while their functional capabilities of immune responses have not been systematically compared, anseriforms

appear immunologically distinct from other birds (Wakelin and Apanius 1997). Among galliform birds, newly hatched chicks have limited but detectable innate and acquired immune responses (Seto and Henderson 1967). Immunoglobulin Y antibodies are transported into the yolk and provide protection for chicks for the first two weeks of life until the immunoglobulins are catabolized (Rose et al. 1974). The qualitative and quantitative expressions of acquired immune responses increase with the growth of galliform chicks and plateaus between 6 and 12 weeks (Rose 1967, Solomon 1968, Lawrence et al. 1981, Suresh et al. 1993).

Acquired Immunity in Reptiles Immunoglobulins first appeared at the level of jawless fishes. Distinctive T- and B-cells first are seen among amphibians (Sell 1996). Reptiles have regulatory T-cells, cells with surface immunoglobulin and lymphoid organs that resemble those of mammals (Sell 1996). Primitive lymph nodules surround the aorta, vena cava, and jugular veins. Lymphocytes and plasma cells are found in the nodules in the intestinal wall, and some lymphocytes are found in the kidneys of reptiles (Tizard 2004). In most reptiles, a bursa-like organ is present (Black 2002). In most reptiles also, the thymus develops from pharyngeal pouches and is structurally similar to the thymus of other vertebrate classes (Tizard 2004); however, no thymus has been described for alligators and crocodiles (Black 2002). Among turtles, the IgM is comparable to mammalian IgM in size, chain structure, and carbohydrate content (Tizard 2002). Turtles and lizards can mount both primary and secondary antibody responses, with IgM produced in the primary response and IgY produced in the secondary response (Tizard 2004). There is evidence of cell-mediated immune responses such as mixed lymphocyte reactions and delayed hypersensitivity reactions among reptiles (Tizard 2004). Both humoral and cellular immune processes decline

with reduced ambient temperatures (Liu and Walford 1972, Tizard 2002).

Immunity in Amphibians Resistance to pathogens involves both an innate immune system and an acquired immune system (Carey et al. 1999). A first defense against pathogens found in the skin and digestive tract of amphibians consists of small, basic antimicrobial peptides active against bacteria, yeasts, and fungi (Jacob and Zasloff 1994, Nicolas and Mor 1995, Rollins-Smith et al. 2002). Like other vertebrates, amphibians have phagocytic cells such as macrophages and neutrophils, a complement system, and natural killer (NK) cells (Carey et al. 1999). Innate immunity is perceived to be an important part of amphibian defenses (Carey et al. 1999, Richmond et al. 2009). Acquired immunity has not been studied in many amphibians. Based on the African clawed frog (Xenopus laevis) and the axolotl (Ambystoma mexicanum), the acquired immune system of amphibians appears to have many similarities to that of mammals (Carey et al. 1999). These include B- and T-lymphocytes (Bleicher and Cohen 1981), several types of immunoglobulins (Carey et al. 1999), cytokines (Haynes and Cohen 1993), and major histocompatibility complex (MHC) class I and class II genes (Flajnik and DuPasquier 1990). Xenopus laevis lacks lymph nodes and lymphopoietic bone marrow, but does have a thymus and spleen (Carey et al. 1999). Interestingly, salamanders have many of the same immune features described for frogs and toads, but often have a weaker acquired immune response; the reasons are not clear (Carey et al. 1999).

Immunity in Arthropods Because of their role as disease vectors to vertebrates, there is also some interest in arthropod immunity. Among their initial defenses, arthropods have a hard chitinous covering that is shed periodically; many also have a high (alkaline) pH in their guts as well as a number introduction to immunity   39

of antibacterial substances that inhibit infection by bacteria (Heimpel and Harshbarger 1965). Arthropods generally protect themselves against invasion by processes of phagocytosis, some antibody-like molecules, and by physical barriers (Tizard 2002). Several types of phagocytic cells occur in invertebrates, including hemocytes and coelomocytes (Heimpel and Harshbarger 1965). These cells function similarly to mammalian phagocytes and undergo chemotaxis, adherence, and ingestion, as well as digestion of foreign materials; they also contain proteases (Tizard 2002). Toll-like receptors have been found in all multicellular organisms examined to date (O’Neill 2004). Arthropods possess a family of enzymes that, when activated, can generate a cascade of proteases leading to the production of phenoloxidase, an enzyme that binds to foreign surfaces and generates melanin around sites where immune defense reactions occur (Tizard 2002). Phenoloxidase enhances phagocytosis and is bactericidal and fungicidal (Tizard 2002). Some insects produce proteins that slow bacterial growth or lyse bacteria. Some insects also induce proteins that mimic antibody molecules. However, little is known about the mechanisms of these antibacterial proteins (Tizard 2002). Although plague bacteria (Yersinia pestis) can remain in fleas for well over a year in some circumstances, there also is evidence that bacteriophages in the intestines of fleas lyse the microbes and can free 60–70% of the infected fleas from plague bacilli in one to three days (Pavlovski 1966). Almost all bacteria are cleared from an insect by phagocytosis and melanization, a unique immune defense mechanism primarily of insects involving the production of melanin, a brown-black pigment that accompanies innate immune responses against several microorganisms (Haine et al. 2008, Schneider and Chambers 2008). Antimicrobial proteins likely remove the bacteria remaining after these more general defensive mechanisms have finished, and this may help explain why 40   introduction to immunity

bacteria generally have not been successful in developing resistance to arthropod peptides.

Behavioral Defenses of Vertebrates While not often addressed, behavioral defenses on the part of the vertebrate host can have considerable influence on the success of parasites attempting to infect a host (Hart 1997). It often is the collective, subtle effects of parasitism that appear to account for behavioral patterns hosts use to reduce parasite loads (Hart 1990). Behaviors for which there is some evidence of control over parasites may be thought of as a first line of defense against parasites, with physiological and immunological forms of resistance serving as a second barrier (Hart 1997). Hosts use a number of strategies for parasite avoidance (Hart 1997). Nest-borne ectoparasites can be avoided or controlled by avoiding old nest sites or abandoning severely infested nests; in some cases ectoparasites may be controlled by the use of secondary plant compounds. Ectoparasites can be removed from the body and plumage by self-grooming, allopreening, heterospecific cleaning, and rubbing ants or other insects containing formic acids on their feathers (anting). Hosts also attempt to avoid flying insects by using fly-repelling behavior, defensive sleeping or resting, grouping away from parasitic flies, as well as seeking arthropod-free microhabitats. Finally, hosts can avoid and control many microparasites by nest sanitation, territoriality, and specific mating behaviors (Hart 1997).

Future Directions There are a number of interesting but rarely addressed ideas worthy of future study. For example, disease resistance mechanisms are energetically costly but rarely measured (Groater and Holmes 1997); in one case, the fever accompanying malaria can increase the human basal metabolic rate by 40% (Hall 1985, Holmes and Zohar 1990). Mounting immune responses may be particularly challenging for growing and developing hosts (Fair et al. 1999). There

also is evidence that suppressing an immune response when infected, under stress, or malnourished may have survival value under some circumstances (Hanssen et al. 2004). Disease tolerance has been identified as a defense strategy (Medzhitov et al. 2012). Resistance in parasites may carry a cost in reduced productivity (Groater and Holmes 1997). For example, mice resistant to Nematospiroides dubius had a larger body size, but decreased litter size (Brindley and Dobson 1981). Much of the damage ascribed to parasites may in fact be due to host responses to the parasites, so-called “immunopathology” (Castro 1990). Specific mechanisms that confer resistance to one parasite may increase susceptibility to another parasite (Groater and Holmes 1997). Elevated production of new red blood cells in mice confers resistance to one isolate of malaria (Plasmodium yoelii) but susceptibility to another (Sayles and Wassom 1988). One also sees this interplay among cattle in resistance to helminths versus ticks (McKinnon 1990), as well as between Eimeria tenella and other coccidia in chickens (Bumstead et al. 1991). Literature Cited Andreasen, C. B., K. S. Latimer, B. G. Harmon, J. R. Glisson, J. M. Golden, and J. Brown. 1991. Heterophil function in healthy chickens and in chickens with experimentally induced staphylococcal tenosynovitis. Veterinary Pathology 28:419–427. Andreu, D., and L. Rivas. 1999. Animal antimicrobial peptides: An overview. Peptide Science 47:415–533. Bartl, S., D. Baltimore, and I. L. Weissman. 1994. Molecular evolution of the vertebrate immune system. Proceedings of the National Academy of Sciences of the United States of America 91:10769–10770. Baxt, L. A., A. C. Garza-Mayers, and M. B. Goldberg. 2013. Bacterial subversion of host innate immune pathways. Science 340:697–701. Bellanti, J. A. 1978. Immunology II. W. B. Saunders Company, Philadelphia, PA. Benedict, A. A., and K. Yamaga. 1976. Immunoglobulins and antibody production in avian species. Pp. 335–375 in J. J. Marchalonis (editor), Comparative immunology. Blackwell Scientific, Oxford, UK. Black, S. J. 2002. Introduction to infection, resistance, and immunity. Pp. 1–11 in J. P. Kreier (editor),



Infection, resistance, and immunity. Taylor & Francis, New York. Bleicher, P. A., and N. Cohen. 1981. Monoclonal anti-IgM can separate T-cell from B-cell proliferative responses in the frog Xenopus laevis. Journal of Immunology 127:1549–1555. Brindley, P. J., and C. Dobson. 1981. Genetic control of liability to infection with Nematospiroides dubius in mice: Selection of refractory and liable populations of mice. Parasitology 83:51–65. Buisseret, P. D. 1982. Allergy. Scientific American 247:86–95. Bumstead, N., B. M. Millard, P. Barrow, and J. K. A. Cook. 1991. Genetic basis of disease resistance in chickens. Pp. 10–23 in J. B. Owen and R. F. E. Axford (editors), Breeding for disease resistance in farm animals. CAB International, Wallingford, UK. Bush, A. O., J. C. Fernández, G. W. Esch, and J. R. Seed. 2001. Parasitism: The diversity and ecology of animal parasites. Cambridge University Press, Cambridge, UK. Carey, C., N. Cohen, and L. Rollins-Smith. 1999. Amphibian declines: An immunological perspective. Developmental and Comparative Immunology 23:459–472. Castro, G. A. 1990. Intestinal pathology. Pp. 283–316 in J. M. Behnke (editor), Parasites: Immunity and pathology. The consequences of parasitic infection in mammals. Taylor & Francis, London, UK. Clough, N. C., and J. A. Roth. 1998. Understanding immunology. Mosby-Year Book, Inc., St. Louis, MO. Davison, F., B. Kaspers, and K. A. Schat. 2008. Avian immunology. Elsevier, Burlington, MA. Doherty, P. C., and R. Zinkernagel. 1975. Enhanced immunologic surveillance in mice heterozygous at the H-2 gene complex. Nature (London) 256:50–52. Fair, J. M., E. W. Hansen, and R. E. Ricklefs. 1999. Growth, developmental stability and immune response in juvenile Japanese quails (Coturnix coturnix japonica). Proceedings of the Royal Society B: Biological Sciences 266:1735–1742. Fearon, D. T., and R. M. Locksley. 1996. The instructive role of innate immunity in the acquired immune response. Science 272:50–54. Flajnik, M. F., and L. DuPasquier. 1990. The major histocompatibility complex of frogs. Immunological Reviews 150:47–63. Goudswaard, J., J.-P. Vaerman, and J. F. Heremans. 1977. Three immunoglobulin classes in the pigeon (Columba livia). International Archives of Allergy and Applied Immunology 53:409–419. Groater, C. P., and J. C. Holmes. 1997. Parasitemediated natural selection. Pp. 9–29 in

introduction to immunity   41

D. H. Clayton and J. Moore (editors), Hostparasite evolution: General principles and avian models. Oxford University Press, Oxford, UK. Hädge, D., and H. Ambrosius. 1983. Evolution of low molecular weight immunoglobulins. III: The immunoglobulin of chicken bile—not an IgA. Molecular Immunology 20:597–606. Hädge, D., and H. Ambrosius. 1984. Evolution of low molecular weight immunoglobulins. IV: IgY-like immunoglobulins of birds, reptiles and amphibians, precursors of mammalian IgA. Molecular Immunology 21:699–707. Haine, E. R., Y. Moret, M. T. Siva-Jothy, and J. Rolff. 2008. Antimicrobial defense and persistent infection in insects. Science 322:1257–1259. Hall, A. 1985. Nutritional aspects of parasitic infection. Progress in Food and Nutrition Science 9:227–256. Hanssen, S. A., D. Hasselquist, I. Folstad, and K. E. Erikstad. 2004. Costs of immunity: Immune responsiveness reduces survival in a vertebrate. Proceedings of the Royal Society B: Biological Sciences 271:925–930. Hart, B. L. 1990. Behavioral adaptations to pathogens and parasites: Five strategies. Neuroscience and Biobehavioral Reviews 14:273–294. Hart, B. L. 1997. Behavioural defence. Pp. 59–77 in D. H. Clayton and J. Moore (editors), Host– parasite evolution: General principles and avian models. Oxford University Press, Oxford, UK. Hayden, F. G. 1995. Antiviral agents. Pp. 411–450 in G. L. Mandell, J. E. Bennett, and R. Dolin (editors), Mandell, Douglas and Bennett’s principles and practice of infectious disease. Churchill Livingstone, New York. Haynes, L., and N. Cohen. 1993. Further characterization of an interleukin-2-like cytokine produced by Xenopus laevis T lymphocytes. Developmental Immunology 23:1–23. Heimpel, A. M., and J. C. Harshbarger. 1965. Symposium on microbial insecticides. V: Immunity in insects. Bacteriological Reviews 29:397–405. Higgins, D. A. 1989. Precipitating antibodies of the duck (Anas platyrhynchos). Comparative Biochemistry and Physiology 93B:135–144. Holmes, J. C., and S. Zohar. 1990. Pathology and host behavior. Pp. 34–63 in C. J. Barnard and J. M. Behnke (editors), Parasitism and host behaviour. Taylor & Francis, London, UK. Hopla, C. E., and A. K. Hopla. 1994. Tularemia. Pp. 113–126 in G. W. Beran (editor), Handbook of zoonoses. CRC Press, Boca Raton, FL. Jacob, L., and M. Zasloff. 1994. Potential therapeutic applications of magainins and other antimicrobial agents of animal origin. Pp. 197–223 in J. Marsh

42   introduction to immunity

and J. A. Goode (editors), Antimicrobial peptides. Ciba Foundation Symposiuim, Vol. 186. Janeway, C. A., and P. Travers. 1997. Immunobiology: The immune system in health and disease. 3rd ed. Current Biology Ltd., London, UK. Klein, J., and C. O’Huigin. 1997. MHC polymorphism and parasites. Pp. 81–88 in W. D. Hamilton and J. C. Howard (editors), Infection, polymorphism and evolution. Chapman & Hall, London, UK. Klion, A. D., and T. B. Nutman. 2004. The role of eosinophils in host defense against helminth parasites. Journal of Allergy and Clinical Immunology 113:30–37. Lawrence, E. C., F. Arnaud-Battandier, J. Grayson, I. R. Koski, N. J. Dooley, A. V. Muchmore, and R. M. Blaese. 1981. Ontogeny of humoral immune function in normal chickens: A comparison of immunoglobulin secreting cells in bone marrow, spleen, lungs and intestine. Clinical and Experimental Immunology 43:450–457. Leslie, M. 2007. Mast cells show their might. Science 317:614–616. Leslie, M. 2012. The new view of complement. Science 337:1034–1037. Liu, R. K., and R. L. Walford. 1972. The effect of lowered body temperature on lifespan and immune and non-immune processes. Gerontologia 18:363–388. Magor, K. E., G. W. Warr, D. Middleton, M. R. Wilson, and D. A. Higgins. 1992. Structural relationship between the two IgY of the duck, Anas platyrhynchos: Molecular genetic evidence. Journal of Immunology 149:2627–2633. McCormack, W. T., L. M. Carlson, L. W. Tjoelker, and C. B. Thompson. 1989. Evolutionary comparison of the avian IgL locus: Combinatorial diversity plays a role in the generation of the antibody repertoire of some avian species. International Immunology 1:332–341. McKinnon, M. J. 1990. Genetic relationships between parasite resistance, growth and fertility in tropical beef cattle. Australian Association of Animal Breeding and Genetics 8:163–171. Medzhitov, R., D. S. Schneider, and M. P. Soares. 2012. Disease tolerance as a defense strategy. Science 335:936–941. Meeusen, E. N. T., and A. Balic. 2000. Do eosinophils have a role in the killing of helminth parasites? Parasitology Today 16:95–101. Ng, P. L. K., and D. A. Higgins. 1986. Bile immunoglobulin of the duck (Anas platyrhynchos). I: Preliminary characterization and ontogeny. Immunology 58:323–327.

Nicolas, P., and A. Mor. 1995. Peptides as weapons against microorganisms in the chemical defense system of vertebrates. Annual Review of Microbiology 49:277–304. O’Neill, L. A. J. 2004. TLRs: Professor Mechnikov, sit on your hat. Trends in Immunology 25:687–693. O’Neill, L. A. J. 2005. Immunity’s early-warning system. Scientific American 252:38–45. Parnham, P. 2009. The immune system. 3rd ed. Garland Science, New York. Pavlovski, E. N. 1966. Natural nidality of transmissible diseases. University of Illinois Press, Urbana, IL. Porter, P., and S. H. Parry. 1976. Further characterization of chicken IgA in chicken serum and secretions with evidence of a possible homologue of mammalian secretory component. Immunology 31:407–415. Randow, F., J. D. MacMicking, and L. C. James. 2013. Cellular self-defense: How cell-autonomous immunity protects against pathogens. Science 340:701–705. Richmond, J. Q., A. E. Savage, K. R. Zamudio, and E. B. Rosenblum. 2009. Toward immunogenetic studies of amphibian chytridiomycosis: Linking innate and acquired immunity. BioScience 59:311–320. Rollins-Smith, L. A., J. K. Doersam, J. E. Longcore, S. K. Taylor, J. C. Shamblin, C. Carey, and M. A. Zasloff. 2002. Antimicrobial peptide defenses against pathogens associated with global amphibian declines. Developmental and Comparative Immunology 26:63–72. Rose, M. E. 1967. The influence of age of host on infection with Eimeria tenella. Journal of Parasitology 53:924–929. Rose, M. E., E. Orlans, and N. Buttress. 1974. Immunoglobulin classes in hen’s egg: Their segregation in yolk and white. European Journal of Immunology 4:521–523. Sayles, P. C., and D. L. Wassom. 1988. Immunoregulation in murine malaria: Susceptibility of inbred mice to infection with Plasmodium yoelii depends on the dynamic interplay of host and parasite genes. Journal of Immunology 141:241–248. Schneider, D. S., and M. C. Chambers. 2008. Rogue insect immunity. Science 322:1199–1200. Schultz, R. M., J. D. Papamatheakis, and M. A. Chirigos. 1977. Interferon: an inducer of macrophage activation by polyanions. Science 197:674–676. Sell, S. 1996. Immunology, immunopathology, and immunity. 5th ed. Appleton & Lange, Stamford, CT. Seto, F., and W. G. Henderson. 1967. Natural and immune hemagglutinin forming capacity of immature chickens. Journal of Experimental Zoology 169:501–512.



Snell, G. D. 1968. The H-2 locus of the mouse: Observations and speculations concerning its comparative genetics and its polymorphism. Folia Biologica 14:335–358. Solomon, J. D. 1968. Immunity to Salmonella gallinarum during ontogeny of the chicken. Immunology 15:197–236. Suresh, M., J. M. Sharma, and S. W. Belzer. 1993. Studies on lymphocyte subpopulations and the effect of age on immune competence in turkeys. Developmental and Comparative Immunology 17:525–535. Swierkosz, E. M., and R. L. Hodinka. 1999. Antiviral agents and susceptibility tests. Pp. 1624–1639 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Tizard, I. 2002. Comparative immunology. Pp. 247–264 in J. P. Kreier (editor), Infection, resistance, and immunity. Taylor & Francis, New York. Tizard, I. R. 2004. Veterinary immunology: An introduction. 7th ed. Saunders (Elsevier), Philadelphia, PA. Tizard, I. R. 2013. Veterinary immunology. 9th ed. Elsevier, Philadelphia, PA. von Rautenfield, B. D., and K.-D. Budras. 1982. The bursa cloacae (Fabricii) of struthioniforms in comparison with the bursa of other birds. Journal of Morphology 172:123–138. Wakelin, D., and V. Apanius. 1997. Immune defence: Genetic control. Pp. 30–58 in D. H. Clayton and J. Moore (editors), Host– parasite evolution: General principles and avian models. Oxford University Press, Oxford, UK. Warr, G. W., K. E. Magor, and D. A. Higgins. 1995. IgY: Clues to the origins of modern antibodies. Immunology Today 16:392–398. Wedemeyer, J., M. Tsai, and S. J. Galli. 2000. Roles of mast cells and basophils in innate and acquired immunity. Current Opinion in Immunology 12:624–631. Wilson, G. S., and A. A. Miles. 1964. Topley and Wilson’s Principles of bacteriology and immunity. 5th ed. The Williams and Wilkins Company, Baltimore, MD. Zinkernagel, R. M., and P. C. Doherty. 1979. MHC-restricted cytotoxic T cells: Studies on the biological role of polymorphic major transplantation antigens determining T-cell restrictionspecificity, function, and responsiveness. Advances in Immunology 27:51–177. Zinkernagel, R. M., and P. C. Doherty. 1997. The discovery of MHC restriction. Immunology Today 18:14–17.

introduction to immunity   43

THREE

Nematodes, Acanthocephala, Pentastomes, and Leeches

CONTENTS Phylum Nematoda 45 Introduction45 Direct Life Cycle Nematodes 48 Intestinal Nematodes 48 Amidostomum anseris48 Abomasal parasites 49 Lungworms52 Dictyocaulus viviparus52 Direct Life Cycle Nematodes with Unusual Features 54 Baylisascaris procyonis54 Trichinella spp. 58

Indirect Life Cycle Nematodes 62 Parelaphostrongylus tenuis62 Filarial Nematodes 65 Elaeophora schneideri65 Phylum Acanthocephala 68 Introduction68 Polymorphus/Profilicollis spp. 69 Other Acanthocephala 72 73

Leeches (Annelida: Hirudinea)

73

Literature Cited

74

While lacking a body cavity, nematodes have a fluid-filled false cavity without a peritoneal lining (pseudocoel) between the somatic musculature and digestive tract that serves as a primary component of the hydrostatic skeleton, and thus is critical for movement; it also serves to transport nutrients between tissues (Bush et al. 2001). Nematodes are covered by a non-cellular cuticle that allows selective permeability and helps maintain their shape by hydrostatic

Phylum Nematoda Introduction Members of the Phylum Nematoda (Greek “nema” 5 thread) are bilaterally symmetrical worm-like metazoans that are round in cross section. The overall body shape of a cylinder tapered at both ends tends to be uniform among most nematode species (Bush et al. 2001) (Fig. 3.1).



Phylum Pentastoma



45

Buccal cavity Vestibule Muscular esophagus (pharynx) Nerve ring Cervical papilla Glandular esophagus Intestine

Male Posterior Spicule Gubernaculum Preanal caudal papilla Cloacal opening (anus) Caudal ala Postanal caudal papilla Female Posterior Uterus Vulva Ovejector Anus

figure 3.1  Generalized anatomy of male and female nematodes (McDonald, 1974; Courtesy of Malcolm McDonald and the U. S. Fish and Wildlife Service).

pressure. With rare exceptions they have separate sexes (dioecious). Nematodes are not segmented, and have a complete digestive tract, including both a mouth and anus (Roberts and Janovy 2000). This is in contrast to cestodes and acanthocephala, which have no intestinal tracts and

absorb food through their cuticles; trematodes have bilateral blind ceca. A buccal capsule lined by a cuticle is present in some nematodes. The esophagus of nematodes is glandular and muscular, and lined by a cuticle. There are several types of esophagi among nematodes. Filariform esophagi have parallel muscular walls and are the most common. Strongyliform esophagi are bulbous. Rhabditiform esophagi are bulbous, with an indentation on the bulb to allow a nerve to cross over. Nematodes can be distinguished from other pseudocoelomate groups by the possession of spicules among males and a ventral excretory pore (Roberts and Janovy 2000). Taxonomy typically is based on the copulatory bursa with associated spicules and rays in males (Bush et al. 2001), the location of the vulvar opening in females, esophageal shape, the anterior adornments (e.g., buccal capsule or lips), and characteristics of cuticular bosses and ridges (Fig. 3.1). Although there is some variation in taxonomic schemes proposed for nematodes (Adamson 1987; Anderson 1992, 2000; Roberts and Janovy 2000), we use the work of Anderson (2000), which is based on the Commonwealth Institute of Helminthology (CIH) keys (Anderson et al. 1974–1983), as the foundation for a classification (App. 1: Table 1). However, biochemical, immunological, and molecular techniques increasingly are providing additional and alternative approaches for studying parasite systematics (McManus and Bowles 1996, Andrews and Chilton 1999). Nematodes have more species that cause mortality among wildlife than any other helminth group. Approximately 16,000 nematode species are described, and about 40,000 species are estimated to exist (Anderson 2000). About one-third of all nematode genera described occur as parasites of vertebrates (Anderson 1984) and many others are important as insect or plant pathogens (Bush et al. 2001). However, many nematodes are free-living and play important roles as decomposers in recycling nutrients, or in food web relationships by feeding on bacteria and other microorganisms

46   nematodes, acanthocephala, pentastomes, and leeches

figure 3.2  Ovum of Ostertagia spp., an abomasal nematode of cervids.

figure 3.3  First stage (L1) larvae of Trichinella spp. being digested out of bear muscle (Courtesy of R. M. Wood and California Department of Public Health).

(Bush et al. 2001). In one assessment, about 24,000 parasitic species of nematodes were estimated to exist (Poulin and Morand 2004). A number of pathogenic nematodes are described for both birds (Wehr 1971) and mammals (Samuel et al. 2001). Among waterfowl (Family Anatidae) alone, over 200 species of nematodes in more than 50 genera have been reported (McDonald 1974); of these, about 20 species are reported to be pathogenic (Wobeser 1997). Nematodes have several distinct life history stages, including an egg (Fig. 3.2), four larval stages (Fig. 3.3), and adults of separate sexes (Fig. 3.4). The larval and adult stages are separated by four molts (Bird and Bird 1991). After adult nematodes mate, the eggs develop first stage larvae (L 1), which successively molt into second stage larvae (L 2), third stage larvae (L 3), fourth stage larvae (L 4), and then mature

into adults. In the great majority of nematodes parasitizing vertebrates, the L 3 larvae are infective to the definitive vertebrate host (Anderson 2000). Some exceptions of stages infective to the definitive host occur with many ascarid nematodes, in which the L 2 larva is infective, Trichinella (L 1 is infective), and Eustrongylides (L 4) (Anderson 2000). There is no reproduction among any larval stages of nematodes. Some nematodes have a direct life cycle (monoxenous), without a need for any intermediate hosts. Other nematodes, especially among vertebrates, have an indirect life cycle (heteroxenous), with development from the L1 to the L3 requiring intermediate hosts (Anderson 2000). Some advantages of an intermediate host are that the pre-infective stages of nematodes are protected from the uncertainties of the external environment in a nutrient-rich environment and the survival of larvae can be extended in time and space; also, the intermediate host often is a desired prey item of the final host and thus increases the likelihood of the parasite reaching its definitive host (Anderson 2000). Although relatively uncommon, some nematodes have paratenic hosts in which the infective stage of a parasite persists without essential development and usually without growth in a host (Anderson 2000). While there are some larval changes in an intermediate host, there are no larval changes in a paratenic host. Nematodes using paratenic hosts include metastrongyloid lungworms of mustelids and members of the order Spirurida (Anderson 1992). Definitive hosts can be infected in a variety of ways. Ingestion of an infective larval stage is the most common means. Ingestion can be of infected intermediate hosts, paratenic hosts, or food or water in fecally contaminated environments. Penetration of skin is another means of transmission. Some infective nematode larvae can penetrate directly through the skin of the definitive host; hematophagous arthropods also can serve to infect a definitive host with infective larvae through the skin. Less commonly, transmammary transmission can occur to nursing young; transmission also can occur by ingestion

nematodes, acanthocephala, pentastomes, and leeches   47

figure 3.4  Adult kidney worms (Dioctophyma renale) from a coyote (Canis latrans). Note the size of the animal’s uninfected kidney on the left (Courtesy of O. Brunetti and California Department of Fish & Wildlife).

of emesis. Transplacental transmission is an occasional mechanism as well (Anderson 1992). In this chapter, we will illustrate some of the variation found among nematode life history strategies, with a focus on species of concern to managers. The approach initially will be by life cycle type, starting with direct life cycles and then focusing on nematodes with indirect life cycles. Coverage will include important intestinal nematodes, lung worms, and vascular system nematodes.

Direct Life Cycle Nematodes Intestinal nematodes amidostomum anseris (gizzard worm) c aus at i v e agen t (cl a ssific at ion, morphology)  Gizzard worms comprise several nematode species of the Superfamily Trichostrongyloidea, Family Amidostomatidae (App. 1: Table 1) (McDonald 1974); two important genera are Amidostomum and Epomidiostomum (Tuggle and Friend 1999, Fedynich and Thomas 2008). Amidostomum anseris (Amidostomatidae: Strongylida) is one of nine species reported for the genus (McDonald 1974) and is the primary focus in this section. host range and distribution  Amidostomum anseris is a nematode infecting the gizzards of wild and domestic geese (Anderson 1992); it also has been reported from ducks, swans,

ardeids, and grebes (MacNeill 1970, Wehr 1971, Gylstorff and Grimm 1987). This common parasite has been reported from North America, Eurasia, and Africa (McDonald 1974, Tuggle and Crites 1984, Gylstorff and Grimm 1987, Gicik and Arslan 2003, Borgsteede et al. 2006). life cycles and variations  Amidostomum anseris lives in the gizzard lining of its avian hosts and has a direct life cycle. The eggs are inhibited by the high body temperature of the host from developing beyond an early set of cleavage cells until shed into the external environment (Geller 1962). After developing to the first (L1) stage, the larvae undergo two more molts and become infective L3 larvae. There is some conflict in the litera ture on the extent to which larval development occurs in the egg, with references both to eggs hatching L1 larvae that immediately shed their cuticle (Cowan 1955, Tuggle and Friend 1999), and to eggs containing infective L3 stages (Enigk and Dey-Hazra 1970, Stradowski 1971). In the latter case, eggs are reported to reach the infective stage in 23 days at 20 C (Enigk and Dey-Hazra 1970). The pre-patent period (time from infection by L3 larvae to shedding of eggs by gravid females) in goslings is 14 to 31 days (Stradowski 1977). Canada goose (Branta canadensis) goslings as young as 34 days have died from infections from parasites, and adult worms have been observed in goslings even younger (Wehr and Herman 1954). Parasites arriving at the same time do not

48   nematodes, acanthocephala, pentastomes, and leeches

mature simultaneously, and infections variably have been reported to persist in birds for 120–175 days (Stradowski 1977), 18 months (Cowan 1955), to several years (Tuggle and Friend 1999). Larvae can swim and even rise to the surface of a water column, but both eggs and larvae are very susceptible to desiccation (Enigk and DeyHazra 1970). In freezing temperatures, eggs can survive but most exposed L3 larvae are killed. reservoirs and transmission Wild and domestic geese are considered the primary reservoirs for A. anseris. Most infections of susceptible hosts follow ingestion of L3 larvae. When goslings were given eggs and L3 larvae of known age, few fully developed L3 larvae still in eggs matured into adult worms, but most L3 larvae given at least one hour after hatching from eggs became established in the gizzard (Stradowski 1971). Infective larvae occasionally can penetrate intact skin; the prepatent period of worms in such cases was 15 to 18 days, similar to birds infected orally (Enigk and Dey-Hazra 1968). It appeared that after invading the skin, the larvae migrated to the lungs and appeared in the mucus of the trachea 16 to 32 hours later. It is not clear whether these worms ever were able to mature and complete the life cycle in the gizzard, and to shed fertile eggs. Among ducks, those up to 6 months of age are susceptible to infective larvae (Stradowski 1972). However, few infective larvae mature and the prepatent period is much longer; thus ducks are not as suitable as geese to host A. anseris (Stradowski 1972).

1971). Laboratory diagnosis can be through the diagnosis of eggs in the feces, but usually requires an expert to identify them. population effects  This parasite can be a serious problem among wild geese (Herman and Wehr 1954, Tuggle and Friend 1999), and often is associated with high goose densities. While A. anseris is the best-known pathogen among waterfowl (Tuggle and Friend 1999), A. acutum also is associated with severe mortality, in combination with food shortage, among adult eiders (Somateria mollissima) in Europe (Borgsteede 2001). special problems  Transmission potential is greatest in crowded, continuously used habitats. Canada geese from Pea Island National Wildlife Refuge on the coast of North Carolina have experienced considerable mortality from the effects of Amidostomum anseris infections (Herman and Wehr 1954, Herman et al. 1955). Severe impacts were associated with inclement weather forcing the birds away from the offshore breeding grounds and being crowded onto the island, with food that was lower in protein and in inadequate supply (Herman 1969). In similar circumstances, mortality occurred among Aleutian cackling geese (Branta hutchinsii leucopareia) in coastal California, at a time when this subspecies was listed as endangered (R. Botzler, personal records). control  With domestic animals, fenbadazol and carbon tetrachloride have been used (Gylstorff and Grimm 1987), as has flubendazole (Vanparijs 1984). No control efforts have been reported among wild birds.

clinical effects and identification  Adult nematodes live in the linings of the gizzard and proventriculus. Heavily infected birds experience severe necrosis of the gizzard lining and subsequent weight loss—often weighing only half as much as healthy birds; they also typically are infected with large numbers of other parasites (Herman 1969). Death follows loss of blood and digestive function in heavily infected birds (Herman and Wehr 1954, Cowan and Herman 1955). Diagnosis usually requires identifying the adult parasite in the gizzard of a carcass (Wehr

abomasal parasites c aus at i v e agen t (cl a ssific at ion, morphology)  A distinctive array of nematodes of the Family Trichostrongylidae lives in the abomasum (fourth chamber of the ruminant stomach) of many ruminants. Genera of abomasal nematodes include Haemonchus, Marshallagia, Mazamastrongylus, Obeliscoides, Ostertagia, Pseudostertagia, Spiculopteragia, Teladorsagia, and Trichostrongylus (App. 1: Table 1) At least 21 distinct species have been reported among these genera from wild bovids and cervids of North America alone (Hoberg et al. 2001),

nematodes, acanthocephala, pentastomes, and leeches   49

with many additional species in other parts of the world (Keep 1971, Wetzel and Rieck 1972, Anderson 1992, Chowdhury and Aguirre 2001). Because many of these nematodes have been associated with diseases and because they often occur in mixed infections among hosts where individual species cannot readily be assessed, they will be addressed collectively. Additional trichostrongyloid nematodes more commonly associated with the small intestine (e.g., Nematodirus spp., Cooperia spp.) also occasionally are found in the abomasa of ruminants. host range and distribution Abomasal parasites typically are found in all ruminant animals, including members of the Bovidae, Cervidae, and Giraffidae. The Camelidae (llamas, camels) have only three chambers in the ruminant stomach, but also are infected with some abomasal parasites (Hoberg et al. 2001). Abomasal parasites have a worldwide distribution (Kotrlý and Kotrla-Erhadova 1970, Keep 1971, Wetzel and Rieck 1972, Anderson 1992), although some of this extended distribution may have been influenced by translocations and introductions of hosts and parasites from Europe and other countries (Hoberg 1997). On a worldwide basis, cross-transmission of abomasal parasites between wild ruminants and domestic hosts may be common (Dunn 1969), but helminths of wild ruminants to which domestic animals are exposed appear generally to be of low pathogenicity (Hoberg et al. 2001). life cycles and variations Abomasal nematodes have direct life cycles. Eggs are passed by gravid females; the L 1 larvae hatch from the eggs (Herman and Wehr 1954) and molt to L 2 and then L 3 larvae in the pasture (Anderson 2000). The L3 larvae are infective and crawl onto herbage until ingested by their grazing definitive hosts. Some L3 larvae of abomasal parasites can survive in host feces or exposed habitat for 4 months or more (Durie 1961); however, the larvae also commonly are susceptible to high temperatures and desiccation. Once ingested, L 3 larvae exsheath in the abomasum and invade the gastric pits and glands, where they elicit host nodules and swellings in

which the larvae develop (Sommerville 1953, 1954); they molt to L 4 larvae in a few days. The L 4 larvae leave the mucosa as early as 4 days post-infection and enter the lumen to mature or, in some cases, have the fourth molt before returning to the lumen (Anderson 1992). Worms may be either free or attached to the abomasum mucosa. The prepatent period typically is 3 weeks or more (Anderson 1992). However, some trichostrongyloid larvae may undergo a development arrest for several weeks or months in their definitive hosts (Michel 1974, Gibbs 1986). reservoirs and transmission Infected ruminants are the reservoirs of abomasal nematodes, and susceptible ruminants usually acquire these parasites by ingesting infective L 3 larvae contaminating their food. Infective larvae of abomasal parasites have a tendency to climb films of moisture up onto vegetation where they become more readily available to the host (Anderson 1992). Earthworms commonly ingest L 3 larvae, which likely pass through the oligochaete intestinal canal to surrounding soil; earthworms also may greatly accelerate the breakdown of feces and contribute to a higher concentration of infective L 3 larvae available in the vicinity of the feces (Grønvold 1979). The coprophilous fungi Pilobolus spp. may contribute to the dispersal of some trichostrongyloid larvae (Bizzell and Ciordia 1965); these fungi are addressed in more detail in the discussion of the lungworm, Dictyocaulus viviparus. clinical effects and identification  Among the abomasal parasites, Haemonchus contortus can be pathogenic in low numbers and is particularly significant for young deer (Odocoileus spp.) (Davidson et al. 1980). These parasites cause blood loss and damage to the abomasal lining; anemia is a common symptom (Prestwood and Pursglove 1981). Abomasal nematodes typically can cause inflammation, edema, and necrosis of the abomasum, as well as diarrhea and emaciation (Soulsby 1968). Traditionally, abomasal parasites have been identified through species-specific features of male bursas, including size and number of spicules, presence and shape of a gubernaculums,

50   nematodes, acanthocephala, pentastomes, and leeches

figure 3.5  Male Ostertagia spp. with spicules and curled bursa from a black-tailed deer Odocoileus hemionus californicus.

and numbers and configurations of their rays (Fig. 3.5) (Little et al. 1998). Females can be identified by the structure of their longitudinal cuticular ridge system (synlophe) (Lichtenfels and Hoberg 1993). population effects  Abomasal nematodes have been important mortality factors among white-tailed deer (Odocoileus virginianus) in the southeastern U. S. (Prestwood and Pursglove 1981). Abomasal parasites, along with intestinal nematodes, long have been important mortality factors among California black-tailed deer (Odocoileus hemionus columbianus) along the northern California coast (Longhurst et al. 1952). An estimated 30,000 abomasal nematodes were found in one coastal elk of northern California (Brunetti 1960). However, there are only limited studies on the population impacts of abomasal parasites on any host of a region. special problems  Abomasal parasites have had significant impacts on white-tailed deer in the southeastern United States; in habitats where deer are above carrying capacity, they had significantly higher abomasal counts than at sites where deer populations were at or below carrying capacity (Eve and Kellogg 1977). Consequently, abomasal parasite counts (APC) of deer were proposed as an indicator of the condition of deer habitat as well as an indicator of the health and nutritional status of a given population of white-tailed deer (Eve and Kellogg 1977, Little et al. 1998).

However, conflicting data emerged with these findings. For white-tailed deer in Mississippi (Demarais et al. 1983) and Texas (Waid et al. 1985), abomasal parasite counts were not related to the physical condition of the deer. For black-tailed deer (Odocoileus hemionus columbianus) of coastal and inland Humboldt County, California, there also was no relationship between deer health measures and abomasal parasite counts (Botzler 1979); the numbers of abomasal parasite varied consistently between three habitats assessed, but the differences were independent of physical condition of deer from the three habitats. The abomasal parasite counts and physical condition of desert mule deer (Odocoileus hemionus hemionus Rafinesque) from southwestern Texas were not consistently related as they were for white-tailed deer in the southeastern U. S. (Moore and Garner 1980). This was also true for southern mule deer (Odocoileus hemionus fulginatus) in San Diego County, California, which had low abomasal parasite counts despite being in relatively poor physical condition (Ladd-Wilson et al. 2000). Host age of deer and season have been linked to variation in abomasal parasite counts. Abomasal parasites were most commonly observed among younger animals (Foreyt and Samuel 1979, Davidson et al. 1980) and in late summer and early fall (Eve and Kellogg 1977, Waid et al. 1985).

nematodes, acanthocephala, pentastomes, and leeches   51

Soil moisture is another factor that has been evaluated in relation to differences in APC. Arid conditions may influence the abomasal parasite counts. For example, the study sites for whitetailed deer in Texas (Moore and Garner 1980, Waid et al. 1985) and southern mule deer in southern California (Ladd-Wilson et al. 2000) were characterized by dry conditions; such dry conditions may reduce the survival of the freeliving phases of these direct life cycle nematodes as they wait to be ingested by the definitive hosts. Black-tailed deer in Humboldt County, from a cutover redwood temperate rainforest, were intermediate in measures of physical condition but had the highest counts of abomasal parasites; deer from a brushland habitat were in very poor physical condition but had an intermediate level of abomasal parasites (Botzler 1979). Thus, the APC may only be of consistent value for whitetailed deer in the southeastern United States (Eve 1981). Alternatively, specific standards may need to be individually tailored to particular herds, habitats, or geographic regions. However, if the animals are killed for collection, parasite loads may be a less direct method to assess herd health than directly measuring health-related physical or chemical parameters of the deer killed. control  Currently, control and treatment remain problematic (Hoberg et al. 2001). While many anthelminthics, including benzimidazoles and avermectins, reduce Haemonchus and other nematodes, few are approved for use in wild cervids and bovids; thus, effective dosages often have not been established (Hoberg et al. 2001). There also are complications associated with attempting anthelminthic therapy in free-ranging ruminants (Prestwood and Pursglove 1981). Haemonchus reportedly is reduced by rafoxanide among wild deer in Europe (Barth and Schaich 1973a, 1973b). Lungworms dictyocaulus viviparus c aus at i v e agen t (cl a ssific at ion, morphology) The lungworm Dictyocaulus viviparus is one of four species of the genus Dictyocaulus in the Family Dictyocaulidae,

Superfamily Trichostrongyloidea (Anderson 2000). Others include D. arnfieldi, a parasite of equines, D. filarial, a common parasite of sheep and goats, as well as wild antelope and deer, and D. eckerti, also in deer (Schneider et al. 1996, Anderson 2000). All of these worms are medium-sized and occur in the trachea and bronchi of their hosts. host range and distribution  Dictyocaulus viviparus is considered a cosmopolitan parasite among many domestic animals such as cattle, sheep, and goats (Soulsby 1968), as well as members of the deer family (Cervidae) (Soulsby 1968, Anderson 1992), and additional wild bovids including bison (Bison bison) (Locker 1953, Boddicker and Hugghins 1969), muskoxen (Ovibos moschatus) (Samuel and Gray 1974), and nyala (Tragelaphus angasi) (Keep 1971). life cycles and variations Adult D. viviparus live in the air passages of the lungs, rather than within the parenchyma as do most other lungworms (e.g., Protostrongylus spp.). The D. viviparus eggs hatch in the lungs, and the L 1 larvae crawl up the respiratory passages, are coughed up by the host, swallowed, and pass out with the feces (Fig. 3.6) (Anderson 1992); such L 1 larvae can be detected with a living larvae technique such as the Baermann Test (Zajac and Conboy 2006). The L 1 larvae are susceptible to desiccation and sunlight, and many die within the first 2 weeks on pasture (Anderson and Prestwood 1981). Under suitable conditions of temperature and moisture, the L 1 larvae molt to the L 2 stage in a few hours, which, in turn, molt to the infective L 3 larvae in a few more hours (Anderson 1992). The L 3 larvae are fairly resistant to cold and freezing conditions and moderate drying (Anderson 1992). reservoirs and transmission  As a direct life cycle nematode, the reservoirs are the infected cervids and bovids in the regions where D. viviparus occurs. The definitive host becomes infected by ingesting infective L 3 larvae contaminating its food. When ingested, the L 3 larvae penetrate the mesenteric lymph nodes of the ileum, cecum, and upper colon and remain for 3 to 8 days (Jarrett et al. 1957,

52   nematodes, acanthocephala, pentastomes, and leeches

ADULT PARASITES IN LUNGS OF HOST

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Larvae penetrate intestine, migrate to and develop in the mesenteric lymph glands and then migrate to lungs via bloodstream

in

INFECTIVE LARVAE INGESTED BY HOST

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LARVAE IN FECES

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Jarrett and Sharp 1963). The larvae then continue their migration to the lungs via the thoracic duct and heart (Soulsby 1968). The prepatent period in cattle is 21–30 days (Jarrett and Sharp 1963, Soulsby 1968), and the patent period is 27–72 days (Rubin and Lucker 1956). For these lungworms, transmission to susceptible hosts can have several complications. The definitive hosts are not coprophagic and many avoid feeding near fecal material; thus, once the L 1 larvae have passed out with the feces, the larvae must be able to leave the infected fecal pads or pellets to have a reasonable chance of being transmitted to the next host. The best likelihood is associated with the larvae being on vegetation used by the definitive host. While there is some evidence that they are negatively geotactic and weakly climb vegetation (Jørgensen 1980), they also are relatively inactive after being deposited with the feces (Soulsby 1968, Anderson and Prestwood 1981). It is possible that the diarrhea associated with long lush vegetation or gastrointestinal parasite infections (Soulsby 1968) can lead the

ge

figure 3.6  Life cycle of Dictyocaulus viviparus (Redrafted with permission from Food and Agriculture Organization of the United Nations, Fig. 169, Manual on meat inspection for developing countries http://www.fao. org/docrep/003/t0756e/ T0756E.htm#ch5.4.1.5).

feces to be deposited directly onto vegetation (Michel and Rose 1954). An interesting aspect of the transmission of D. viviparus is the involvement of a coprophilous fungus (Pilobolus spp.) to spread L 3 away from the feces onto surrounding herbage (Robinson 1962, Croll 1966). These fungi characteristically discharge their sporangia explosively (Buller 1934). Large numbers of D. viviparus have been observed to migrate to the upper surface of the sporangiophore of Pilobolus spp.; when the sporangia discharged, the larvae were thrown into the air up to distances of 1.2 m under laboratory conditions and up to 3.05 m from fecal pads in the field (Robinson 1962, Doncaster 1981). Cattle pastures with Pilobolus spp. had far more infective D. viviparus larvae than plots without Pilobolus spp. (Jørgensen et al. 1982). Pilobolus spp. also occurs in Yellowstone National Park, and elk (Cervus elaphus nelsoni) have a high D. viviparus prevalence there; thus the same relationship between parasite and fungus may hold here (Foos 1989). While dung beetles have been proposed as a means of mechanically

nematodes, acanthocephala, pentastomes, and leeches   53

spreading D. viviparus larvae (Poynter and Selway 1966), there also is evidence that dung beetles can reduce the likelihood of infection in hosts by trichostrongyloid nematodes (Fincher 1975, Bergstrom et al. 1976). In some cases the L 3 larvae may disappear from the herbage of infected pastures during winter but reappear the following spring or early summer (Duncan et al. 1979). This may be a result of heavy rains washing larvae from the vegetation into the soil; earthworms may contribute to the overwintering survival of D. viviparus larvae by ingesting the previous year’s L 3 larvae in the soil and returning them to the surface (Oakley 1981). clinical effects and identification  The prepatent phase in the lungs of cattle is associated with blockage of many bronchioles, eosinophilic exudate, and collapse of alveoli; this is associated with rapid shallow breathing and coughing and is sometimes followed with emphysema. Infected black-tailed deer lose weight, have increased coughing, and have an abnormal increase in the depth and rate of breathing (hyperpnea) (Presidente et al. 1973). Among white-tailed deer, gross pathologic lesions attributable to D. viviparus are confined to the respiratory tract and may range from a foamy exudate in the air passages in mild infections, to a bronchitis and peribronchitis (Goble 1941), to thick purulent exudate in the air passages, and even obstruction of the airways by infecting worms and exudate (Jarrett et al. 1957, Anderson and Prestwood 1981). Opportunistic secondary invaders are common and often may be the eventual cause of death (Anderson and Prestwood 1981); examples include Mannheimia haemolytica and Pasteurella multocida, as well as a variety of viruses and mycoplasma. population effects  Dictyocaulus viviparus infections may be relatively common in some deer populations (Prestwood et al. 1971). Among white-tailed deer in New York, D. viviparus lungworms were found in higher prevalence among winter-killed deer, compared to hunter-collected deer in the same area; however, this observation was compounded by

three other parasites, including another lungworm [Varestrongylus (5 Leptostrongylus) alpenae], also having higher prevalences among the winter-killed deer (Cheatum 1951). Thus the relative contribution of D. viviparus was difficult to ascertain. Losses to dictyocaulosis, especially among fawns, may occur annually among white-tailed deer in some areas, and dictyocaulosis has been proposed as a population control mechanism in these areas (Anderson and Prestwood 1981), but there are no known systematic studies establishing significant impacts by D. viviparus on wild populations. There have been past reports of D. viviparus having significant effects on black-tailed deer populations in California and British Columbia (Cowan 1946, Longhurst et al. 1952), but no systematic studies on their impacts. special problems  There are many reports of sporadic mortality from D. viviparus. However, there are no clearly established ongoing problems reported for these lungworms. control Infections in cattle generally occur on over-grazed pastures where the grass is short (Levine 1968). It has been observed that white-tailed deer in the southeastern U.S. with D. viviparus problems often are associated with areas of poor range condition or which are overpopulated; thus population control and range management are important. Under these circumstances it was proposed that controlled burning in the late winter or early spring may destroy larvae that overwintered on a pasture (Anderson and Prestwood 1981). Keeping range in good condition also would be a sensible strategy—both to reduce the benefit of any upward climbing by lungworm larvae and to keep deer in good health. Among penned black-tailed deer, anthelminthic therapy was not successful (Presidente et al. 1973). Direct Life Cycle Nematodes with Unusual Features baylisascaris procyonis c aus at i v e agen t (cl a ssific at ion, morphology)  Baylisascaris procyonis (Nematoda, Family Ascarididae) is a large roundworm

54   nematodes, acanthocephala, pentastomes, and leeches

of the small intestine (Kazacos and Boyce 1989, Kazacos 2001). The definitive host for adult worms is the raccoon (Procyon lotor); while young raccoons can be infected by ingesting eggs as in a direct life cycle, adult raccoons typically are infected by ingesting infected intermediate hosts or paratenic hosts. Baylisascaris procyonis is one of at least seven species in the genus (Sprent 1968, 1970). Other species in the genus include B. columnaris in skunks, B. laevis in marmots and ground squirrels, B. devosi in martens and fishers (Mustela spp.), B. transfuga in all species of bears, B. melis in European badgers (Meles meles), B. schroederi in giant pandas (Ailuropoda melanoleuca), and B. tasmaniensis in Tasmanian devils (Sarcophilus harrisii) (Sprent 1968, 1970). The genus Baylisascaris is closely related to the genus Ascaris, and to a lesser extent Toxascaris and Parascaris in the family (Zhu et al. 2000). host range and distribution  Baylisascaris procyonis adults are found primarily in adult raccoons, but also have been found in at least two other species as natural infections, including kinkajous (Potus flavus) (Overstreet 1970) and domestic dogs (Greve and O’Brien 1989); there also are several hosts in which adult parasites can form after experimental infections (Miyashita 1993, Kazacos 2001). Baylisascaris procyonis has a geographic distribution that basically corresponds with the distribution of its main host, the raccoon, and covers primarily the United States; while less common in the southeast (Kazacos 2001, Acha and Szyfres 2003), it appears to be increasing in this region (Eberhard et al. 2003). The parasite occurs among raccoons in parts of Canada and also has been reported among kinkajous of Colombia (Overstreet 1970) and raccoons in zoos or pets in Germany, Poland, Czechoslovakia, and Japan (Miyashita 1993, Kazacos 2001). Among raccoons in the northeastern United States, autumn was the period of highest prevalence of eggs in the feces (Kidder et al. 1989). There also was a significantly higher prevalence of patent infections in juveniles compared to adults (Kidder et al. 1989).

life cycles and variations Raccoons are the normal definitive host, and adult female nematodes in the small intestines of raccoons produce up to 180,000 eggs per worm per day (Fig. 3.7) (Kazacos 1982). Among these ascarid nematodes, the eggs commonly develop infective L 2 larvae in 11 to 14 days (Sakla et al. 1989), and these embryonated eggs can remain infective in the environment for years (Kazacos 2001). Infected raccoons can shed feces with over 30,000 eggs per gram, with highest egg counts occurring in the fall and the lowest in winter (Evans 2002b). In young raccoons, L2 lar vae hatching from ingested eggs penetrate and develop within the mucosa of the small intestine for several weeks before entering the intestinal lumen to mature in 50 to 76 days (Kazacos 2001). A wide variety of birds and mammals, but especially rodents, can serve as intermediate hosts (Tiner 1953, Kazacos 2001, Evans 2002a), in which ingested L 2 larvae molt to L 3 larvae. Once eggs are ingested, the L 2 larvae penetrate the intestinal wall, migrate through the liver to the lungs, and enter the pulmonary veins to gain access to the arterial circulation. The parasite produces a larval migrans, a prolonged migration and persistence of helminth larvae in the organs and tissues of infected intermediate hosts (Beaver 1969). Some larvae become encapsulated in visceral and somatic tissues in eosinophilic granulomas until ingested by raccoons (Kazacos 2001). In older raccoons, L 3 larvae from intermediate hosts develop to adults in the intestinal lumen in 32 to 38 days without evidence of any migration through other body tissues (Kazacos 2001). reservoirs and transmission In the midwestern United States, both the prevalence and intensity of B. procyonis infections are higher among raccoons from rural areas than those in urban locations. Although raccoons may be very common in urban sites, the lower levels of infection may follow from their decreased dependence on infected intermediate hosts as a source of food in urban areas and decreased exposure of young raccoons to B. procyonis eggs (Page et al. 2008).

nematodes, acanthocephala, pentastomes, and leeches   55

figure 3.7  Life cycle of Baylisascaris procyonis (Courtesy of Centers for Disease Control and Prevention’s Division of Parasitic Diseases and Malaria, www.dpd.cdc.gov/dpdx).

Transmission to raccoons is by ingesting contaminated nutrients or infected intermediate hosts. Young raccoons become infected at an early age by ingesting eggs from their mother’s contaminated teats or fur, from the contaminated den, or from raccoon latrines near their den (Kazacos 2001). Older raccoons become infected by ingesting L3 larvae in intermediate hosts (Kazacos and Boyce 1989). Transmission among raccoons can be enhanced by altering

resource distributions. Prevalences of B. procyonis in raccoons was significantly higher in a habitat with clumped food distributions compared to a similar habitat in which an equal amount of food was dispersed uniformly; the clumped food distribution was postulated to enhance intraspecific contact (Gompper and Wright 2005). In nature, most transmission to intermediate hosts occurs at raccoon latrines, preferred sites of raccoon defecation where their feces

56   nematodes, acanthocephala, pentastomes, and leeches

with B. procyonis eggs accumulate (Page et al. 1999). Large numbers of eggs can occur in these latrines, and they can become important long-term sources of infection (Kazacos 2001). Many species of mammals and birds visit these latrine sites, and a considerable number of species actively forage there (Page et al. 1999, 2001b). The development of the central nervous system disease is viewed as a benefit for B. procyonis because the debilitation or death of the host enhances transmission by predation or scavenging so as to reinfect the raccoons (Tiner 1953, Kazacos and Boyce 1989). clinical effects and identification  This nematode is a native parasite that has co-evolved to a benign relationship with raccoons of North America. However, it can be very pathogenic in many of the species that serve as intermediate hosts (Tiner 1953, Sprent 1968, Kazacos 2001, Evans 2002a). Among intermediate hosts, the L s larvae undergo a larval migrans, causing fatal central nervous system disease, pulmonary hemorrhage, and other somatic disruptions (Tiner 1953, Kazacos and Boyce 1989). Those migrating to the brain produce a progressive central nervous system disease; the onset and severity are dose related (Kazacos 2001). A single larva in the brain of a mouse or small bird usually is fatal (Tiner 1953, Sheppard and Kazacos 1997). Identification of the adults is based on morphological characters and the site of infection (small intestine) of the definitive host (Kazacos 2001). population effects This parasite has become well known as an important cause of morbidity and mortality in individuals and populations of small vertebrates sharing or frequenting the habitat of infected raccoons (Kazacos 2001). In one assessment, B. procyonis was associated with about 5% of the mortality measured among wild white-footed mice (Peromyscus leucopus) (Tiner 1954). Baylisascaris procyonis may have been an important factor in the extirpation of the Allegheny wood rat (Neotoma magister) from parts of its range in the Appalachian Mountains of the eastern United States (McGowan 1993,

LoGiudice 2003), linked to the foraging behavior of the woodrat. Woodrats carry whole raccoon feces to their nests and generally do not collect them until they are several weeks old, when the eggs had enough time to embryonate (LoGiudice 2001). LoGiudice also speculated that the nematode’s benign relationship with raccoons, combined with its highly pathogenic effects on the Allegheny woodrat, may have weakened typical density-dependent relationships in which the hosts could avoid being driven to extinction by parasites (LoGiudice 2003). The abundance of B. procyonis appears to depend more on the density of the raccoon than on that of any one species of intermediate host; thus the raccoon can function as the reservoir for the parasite by continuously reintroducing the parasite to the intermediate host and allowing it to persist in populations that otherwise might be too small to maintain it (Dobson and May 1986, LoGiudice 2003). special problems  This parasite is emerging as an important human pathogen, principally affecting young children. Raccoons increasingly are peridomestic animals living in close proximity to humans and their high densities, which, along with the high prevalence of infective eggs in their feces in raccoon latrines, has led to increased concern about this helminthic zoonotic organism (Sorvillo et al. 2002, Roussere et al. 2003). This is a particularly severe problem among children because the very young can be neurologically devastated despite treatment with anthelminthic drugs and corticosteroids (Gavin et al. 2002). Translocation to other geographic regions is a potential concern. Although this parasite is found primarily in raccoons of North America, it has been found in mammalian hosts of other countries to which raccoons have been introduced. For example, this parasite caused an outbreak of larval migrans among rabbits (Oryctolagus cuniculus) of a wildlife park in Japan (Sato et al. 2002). control Under controlled conditions, naturally and experimentally infected dogs

nematodes, acanthocephala, pentastomes, and leeches   57

harboring B. procyonis had their infections cleared or significantly reduced after administration of milbemycin oxime (Bowman et al. 2005). Decontamination of raccoon latrines in urban settings offers promise for controlling the human risk from this nematode (Page et al. 1999). The diet of raccoons may influence the risk of infection to potential intermediate hosts. For example, white-footed mice (Peromyscus leucopus) may have higher prevalences of infection when corn is present in raccoon feces (Page et al. 2001b). Likewise, differential foraging strategies can influence the risk of ingesting embryonated eggs among intermediate hosts; white-footed mice tended to select corn particles from fresh feces and had lower risk of ingesting infectious eggs than woodrats, which brought back older, whole fecal pellets to their nests containing embryonated eggs (LoGiudice 2001). Habitat disturbance also can affect the risks to intermediate hosts; prevalence, intensity of infection, and average number of larvae were significantly higher among white-footed mice in a highly fragmented, predominantly agricultural landscape, compared to a homogeneous, predominantly forested landscape (Page et al. 2001a). trichinella spp. c ausative agen t (cl assific ation, morphology)  Trichinella spp. are members of the Superfamily Trichinelloidea, which all contain a distinctive esophagus with a short muscular anterior portion and a long glandular posterior portion. The genus Trichinella is the sole member of the Family Trichinellidae. The Trichinella species generally are indistinguishable based on morphology (Dick and Pozio 2001), and species identification in much of the earlier literature is not clear. Thus much of the following discussion will be limited to the genus as a whole, with development of species and genotype differences given when adequate information is available. Trichinella spiralis is well established as a species, and for many years was considered the only accepted species in the genus (Zimmerman 1971). However, the systematics

of Trichinella has been considerably complicated by evidence that there are consistent differences in isolates from animals throughout the world where the parasite occurs (Anderson 2000). Based on isoenzymic patterns in isolates of Trichinella spp., from many hosts and sites, there are at least seven distinct electrophoretic clusters (LaRosa et al. 1989, Pozio et al. 1989). There now are at least 12 taxa, including eight species, with four additional genotypic variants not yet taxonomically defined (Gasser et al. 2004, Pozio 2013). Currently, two main clades are recognized, one comprising species that encapsulate in host muscle tissue and parasitize only mammals, and the other that does not encapsulate and whose species parasitize mammals and birds or mammals and reptiles (Pozio and Zarlenga 2005). Five species comprise those forming capsules. Trichinella spiralis has a cosmopolitan distribution in temperate and equatorial climatic zones, because it has been passively imported into most continents due to its high infectivity to swine and synanthropic rats; it infects a variety of sylvatic carnivores (Dick and Pozio 2001). It is the species causing most human infections, and it is among the most pathogenic species because of the large number of larvae produced by females (Pozio et al. 1992b); it generates a strong immune response among infected humans (Gomez-Morales et al. 2002). Trichinella nativa and the closely related genotype T6 infects sylvatic and marine carnivores living in frigid zones of Asia, North America, and Europe; the isotherm −4°C in January seems to be the southern border of this species, and larvae can survive in frozen muscles of carnivores for up to 5 years (Pozio et al. 1992a, Dick and Pozio 2001, Pozio and Zarlenga 2005). Trichinella britovi and the T8 genotype infect sylvatic carnivores of the Palearctic region in Europe, Asia, and also Africa (Pozio et al. 1992a). Trichinella murrelli infects sylvatic carnivores in temperate areas of the Nearctic region (Pozio and LaRosa 2000, Hill et al. 2008). Trichinella nelsoni infects sylvatic

58   nematodes, acanthocephala, pentastomes, and leeches

carnivores from East Africa to South Africa (Pozio et al. 1992a, Pozio and Zarlenga 2005). Three additional species do not induce capsule formation during the muscle phase of infection. Trichinella pseudospiralis is a cosmopolitan parasite of both birds and mammals, including marsupials. Trichinella papuae has been found only in Papua New Guinea; it infects both mammals and reptiles, with wild pigs appearing to be the primary reservoir. Trichinella zimbabwensis has been found only in crocodiles and monitor lizards of Africa under natural conditions and has infected several mammals in the laboratory (Pozio and Zarlenga 2005). host r ange and distribution Host ranges and distribution have been addressed in the presentation of species above. The genus Trichinella is unique among animal parasite associations because it has at least 150 species of definitive hosts (Campbell 1983). Virtually any mammal can be infected with Trichinella spp. (Dick and Pozio 2001, Pozio and Zarlenga 2005). In addition, T. pseudospiralis can infect birds, and T. papuae and T. zimbabwensis both infect reptiles. Although the various Trichinella species and genotypes have varied distributions, many of the species are cosmopolitan and the genus itself has a broad and worldwide distribution, extending from the Arctic to the tropics, including Oceania (Dick and Pozio 2001, Pozio and Zarlenga 2005). While often considered a zoonotic parasite of domestic habitats, involving pigs, synanthropic rats, and humans, there is evidence that total Trichinella spp. biomass is greater in wild than in domestic animals (Pozio 2013). There are variations among the various geographic regions. Trichinella nativa and T. spiralis both are reported in Arctic regions. Polar bears (Ursus maritimus) are a key component of transmission in the Arctic and are likely infected by cannibalism, or by scavenging carcasses of Arctic foxes and sled dogs; Trichinella larvae can survive in bears for at least several years (Born and Henriksen 1990, Dick and Pozio 2001). Walruses (Odobenus rosmarus) are another important source; larvae also have

been found in bearded seals (Erignathus barbatus), ringed seals (Phoca hispida), and Beluga whales (Delphinopterus leucas) (Rausch et al. 1956, Madsen 1961, Forbes 2000). In contrast with the Palearctic region, there are fewer reports of trichinellosis from wildlife in the Nearctic region. Black bears (Ursus americanus) are important in parts of eastern United States and New Brunswick, Canada, as well as throughout the Rocky Mountains, Arctic regions of Canada, and Alaska. Raccoons appear unimportant in central and western Canada, but are important in parts of the United States. Foxes are important in temperate regions with large urban communities and large tracts of agricultural lands as well as in the Arctic. In contrast, small furbearers such as the marten (Martes americana) appear important for transmission in areas with large tracts of relatively intact forest lands. In the Rocky Mountains, cougar (Puma concolor), bears, and perhaps marten are important hosts (Dick and Pozio 2001). Although some of these strains may be T. spiralis, most are identified as Trichinella T5 and Trichinella T6 genotypes (Dick and Pozio 2001). Details on additional geographic regions, including the temperate Palearctic region (T. britovi), the Ethiopic region (probably T. nelsoni), the Southeast Asia and Oceania region, the Central and South American region, are covered in other sources (Dick and Pozio 2001, Pozio and Zarlenga 2005). life cycles and variations  Despite the wide variety of hosts and geographic regions involved, the species and genotypes of Trichinella spp. generally have very similar life cycle strategies. As a model for the other species, Trichinella spiralis infective L 1 larvae are acquired by ingesting meat containing cysts with larvae of T. spiralis (Fig. 3.8). After exposure to gastric acid and pepsin, the L 1 larvae are released from the cysts and invade the small bowel mucosa, where they undergo four molts and develop into adult worms within 36 hours (Ali Khan 1966, Anderson 2000). The females release L 1 larvae that invade veins or lymphatics (Anderson 2000)

nematodes, acanthocephala, pentastomes, and leeches   59

figure 3.8  Life cycle of Trichinella spp. (Courtesy of Centers for Disease Control and Prevention’s Division of Parasitic Diseases and Malaria, www.dpd.cdc.gov/dpdx).

and migrate to the striated muscles, where they encyst. The muscles most commonly infected are the diaphragm, larynx, tongue, intercostal, biceps, abdomen, psoas, pectoral, gastrocnemius, and deltoid muscles (Anderson 2000). The infected muscle cell is modified into a “nurse cell,” which functions to nourish the parasite or protect it from host responses (Stewart and Giannini 1982). The concept of an intermediate host is difficult to apply in the case of Trichinella spp. Trichinelloids always infect the definitive host as L 1 larvae (Anderson 2000); in contrast, an

L 1 larva is considered the infective stage for the intermediate host in almost all other parasitic nematodes requiring an intermediate host. There is no pause in development at the L 3 larva stage, the stage commonly viewed as infective to definitive hosts. Thus, in the Trichinella spp. life cycle, the parasite’s definitive host functionally serves as its own intermediate host. reservoirs and transmission Trichinellosis is primarily a disease of sylvatic carnivores with cannibalistic and scavenger behaviors (Dick and Pozio 2001, Pozio 2013). While ingestion is established as the route for transmission,

60   nematodes, acanthocephala, pentastomes, and leeches

some aspects of transmission among marine mammals are not well established. Trichinella nativa in marine mammals has a circumpolar Arctic distribution and a narrow host range (Pozio 2013). It is most commonly found in polar bears and walruses, where it presents a significant zoonotic hazard. Cannibalism probably is the main route for maintaining infections among polar bears (Forbes 2000). Arctic carnivores as polar bears, foxes, and domestic dogs have high prevalences of Trichinella spp., and the carcasses of at least some of these animals are deposited in the ocean. While scavenging on these carcasses by walruses probably occurs, it likely does not account for the high prevalence of Trichinella spp. seen in walruses. Active predation, carrion feeding, and cannibalism have been documented for walruses, and a sylvatic cycle similar to that of bears may exist in walrus populations (Forbes 2000). Although uncommon, Trichinella is found in seals and whales; these hosts likely are infected through occasional exposure to infected carcasses, either directly by scavenging or indirectly by consuming amphipods or fish that have fed on infected carcasses. The inefficiency of such transmission may help account for the low prevalence of Trichinella larvae in seals and whales (Forbes 2000). clinical effects and identification There are few observations of clinical effects or pathology of Trichinella spp. in free-living wild animals. Diarrhea, sloughing of gut epithelium, and local hemorrhaging occur in the intestines of experimentally infected domestic and laboratory animals; domestic animals generally tolerate high levels of infection and have few clinical signs (Dick and Pozio 2001). Experimentally infected animals tend to have reduced activity and reduced reproductive success (Dick and Pozio 2001). The simplest method to recover larvae of Trichinella spp. is to compress pieces of striated muscle from the tongue, diaphragm, or intercostal muscles of mammals between glass slides and view them with a dissecting

microscope. One also can digest muscle tissue in 1% HCl/pepsin solution at 37°C for several hours (Dick and Pozio 2001). Identification of the various Trichinella spp. is complicated. Except for the existence of a capsule and some possible size differences in one of the groups, all species and genotypes of the genus are morphologically indistinguishable at all development stages; thus only biochemical or molecular methods can be used reliably to identify the genotype (Pozio and Zarlenga 2005). A number of methods have been devised, with the polymerase chain reaction (PCR) of single larvae being most common (Zarlenga et al. 1999, Gasser et al. 2004, Pozio and Zarlenga 2005). population effects  While it has been spec­ ulated that Trichinella may have some influence on carnivore population densities in Kenya and other parts of Africa (Nelson 1982), there is no clear evidence for any Trichinella spp. causing significant population effects among freeliving wildlife. special problems Emerging problems are related to non-encapsulated species of Trichinella infecting a wide spectrum of hosts (humans, other mammals including marsupials, birds, reptiles), as well as encapsulated Trichinella species infecting horses and other herbivores (Pozio 2001). It is not clear how long these newly discovered host–parasite systems have been established, nor the role humans have played in influencing their importance. The sylvatic cycle is complicated by the existence of a variety of Trichinella species and genotypes in different regions, and the existence of “new” transmission patterns. Also, game animals have become a more common source of infection for humans in developed and developing countries (Pozio 2001). control  There are no recommended means of control of trichinellosis in free-living wildlife. For domestic animals, the primary control methods are to prevent ingestion of infected meat by susceptible animals; this involves cooking garbage fed to domestic pigs for 30 minutes at 100°C, and rodent control (Fraser and Mays

nematodes, acanthocephala, pentastomes, and leeches   61

1986). Use of ACTH and adrenal corticosteroids also can serve as supportive therapy in some cases (Fraser and Mays 1986). For humans, thiabendazole can be used to treat infections. Likewise, raw pork or products lacking a governmental seal should be cooked at 58°C for about 6 minutes or be frozen at −25°C for 10 to 20 days (Fraser and Mays 1986). However, meat in the Arctic is best treated by cooking rather than freezing (Acha and Szyfres 2003).

Indirect Life Cycle Nematodes parelaphostrongylus tenuis (meningeal worm) c aus at i v e agen t (cl a ssific at ion, morphology)  The genus Parelaphostrongylus (para: near, elapho: deer, strongylus: nematode) was named based on this genus being similar to another genus, Elaphostrongylus. Parelaphostrongylus tenuis (Superfamily Metastrongyloidea, Family Protostrongylidae) is one of three members of this genus (Lankester 2001). Earlier, this parasite was called Pneumostrongylus tenuis, but was reclassified into its current genus about 1971. Two other closely related species, P. odocoilei and P. andersoni, also occur in North America; both are called “muscleworms” because the adult nematodes are found in striated muscle of the deer definitive hosts. host range and distribution  Parelaphostrongylus tenuis is a neurotropic nematode of white-tailed deer (Anderson 2000) found in most regions of the eastern United States and southern Canada (Lankester 2001). Generally, this parasite is common among deer of the deciduous forest biome and deciduous– coniferous ecotones of eastern and central North America. It is rare or absent in the coastal plains region of the southeastern United States and has not been documented in western North America (Lankester 2001). Its distribution has been extended into parts of Europe, including Holland, Scotland, Germany, the Czech Republic, Sweden, Finland, Russia (Ural Mountains), central Asia, and

south Siberia, through white-tailed deer introductions (Hörning 1975). White-tailed deer are considered the normal definitive host of this parasite (Lankester 2001). However, in North America, the parasite also has been described in mule deer and blacktailed deer (Odocoileus hemionus) (Anderson et al. 1966, Nettles et al. 1977a, Tyler et al. 1980) elk and red deer (Cervus elaphus) (Carpenter et al. 1973), moose (Alces alces) (Karns 1967), caribou and reindeer (Rangifer tarandus) (Anderson 1971), and fallow deer (Dama dama) (Nettles et al. 1977b). In Europe, this parasite also is reported from free-living fallow deer, red deer, reindeer, and moose that have come in contact with introduced infected white-tailed deer (Hörning 1975). In addition, infections have been reported in a wide variety of bovids and other ungulates in zoos or game farms situated on white-tailed deer habitats, or through experimental infections (Anderson 1992, Lankester 2001) However, the parasite rarely develops to adulthood or is able to pass infective larvae in hosts other than white-tailed deer (Lankester 2001). life cycles and variations In whitetailed deer, the usual definitive host, the adult worms most frequently are found in the veins and venous sinuses of the cranial meninges (Lankester 2001). Unembryonated eggs released by females into the venous blood are carried to the lungs, where they lodge in alveolar capillaries and hatch L 1 larvae (Fig. 3.9). These L 1 larvae move into the mucous of alveolar air space and are carried up the lungs by cilia to the pharynx, where they are swallowed and pass through the digestive tract and out with the feces (Anderson 1963). The L 1 larvae tend to concentrate in a thin layer of mucous on the surface of fecal pellets (Lankester and Anderson 1968) and can be detected by a Baermann apparatus or other living larvae technique. To develop further, L 1 larvae must penetrate or be eaten by one of many terrestrial snails or slugs that can serve as intermediate hosts (Lankester 2001). The L 1 larvae molt to L 2 and then to L 3 larvae in the

62   nematodes, acanthocephala, pentastomes, and leeches

figure 3.9  Life cycle of Parelaphostrongylus tenuis. A. abomasum. B. brain. H. heart. I. intestine. J. jugular vein. L. lung. LN. lumbar nerve. O. esophagus. PC. Peritoneal cavity. R. rumen. SC. Spinal cord. T. trachea (Anderson, 1992; Courtesy of Uta Strelive, copyright holder).

foot of the gastropod over a period of several weeks. Development is influenced by temperature and the metabolic rates of the gastropods, and the L 3 larvae probably remain viable for the life of the intermediate host (Lankester and Anderson 1968). White-tailed deer are infected by ingesting infected gastropods. The L 3 larvae penetrate the deer abomasum and other parts of the intestinal tract and migrate through the peritoneal cavity to the central nervous system (Anderson 1963, Lankester 2001). The L 3 larvae enter the dorsal horns of gray matter in the spinal cord, molt to the L 4 and then to the subadult stage as they move up the spinal subdural space.

On reaching the head, they enter the venous sinuses (Lankester 2001). The prepatent period of P. tenuis in white-tailed deer varies from 82 to 137 days (Anderson and Prestwood 1981). The worms are long-lived in white-tailed deer and may persist for the life of the host (Slomke et al. 1995, Lankester 2001). reservoirs and transmission  The whitetailed deer is the usual definitive host, although moose and elk occasionally can become patent and shed some larvae (Lankester 2001). However, it is not evident that the parasite can be established in any geographic region without the presence of the white-tailed deer as the reservoir host.

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Two key points of transmission include the L 1 larvae moving from infected deer feces to the gastropod intermediate host, and then the L 3 larvae from infected snails being ingested by a susceptible definitive host. The ability of L 1 larvae to withstand adverse natural conditions and remain infective to gastropods has not been thoroughly studied (Lankester 2001). Although they can be kept frozen in the laboratory for several months (Lankester and Anderson 1968), repeated freezing and thawing greatly reduces survival of the L 1 larvae (Shostak and Samuel 1984). The larvae also do not survive well beneath snow (Forrester and Lankester 1998). There is some inconsistency on whether transmission is more likely in specific sites within a white-tailed deer range (Lankester and Anderson 1968, Maze and Johnstone 1986, Platt 1989). Where distinct foci of infection seem to occur, distribution of infected gastropods and young deer may be contributing factors to the distribution of the parasite (Lankester 2001). clinical effects and identification In white-tailed deer, the parasite generally causes no clinical signs or pathology (Anderson 1992). However, considerable pathology can occurs in other species that serve as abnormal hosts. The parasite is highly pathogenic in moose, in which it causes a neurological disease (“moose sickness”); clinical effects include lumbar weakness, ataxia, torticollis, blindness, fearlessness, depression, paresis, paraplegia, and death (Anderson 1992). The parasite also is highly pathogenic in caribou (Trainer 1973) and reindeer (Anderson 1971). While elk and red deer are susceptible to neurologic disease, they can tolerate the infection and sometimes even pass L 1 larvae (Anderson 1992). The parasite also is pathogenic to mule deer, and partially pathogenic to mule deer and white-tailed deer hybrids (Anderson et al. 1966, Nettles et al. 1977a, Tyler et al. 1980). In abnormal hosts, the parasite causes neurological symptoms and death through meningitis (inflammation of the meninges),

encephalitis (inflammation of the brain), and meningoencephalitis. The severity of infection in abnormal hosts probably is due to the higher proportion of invading worms that reach the central nervous system, their longer developmental period in the spinal cord, their resulting larger size and coiling behavior, and frequent invasion of the membranes lining the ventricles of the brain (Anderson 1968). Recovering and identifying adult parasites from infected hosts currently is the only way to confirm infection with P. tenuis. However, the first stage larvae passing out with the feces have a distinctive dorsal cuticular spine on the tail, and this often is strong evidence for infection by P. tenuis; such larvae are detected with a Baermann apparatus or other living larvae technique. There is interest in developing blood tests using immunological and molecular techniques (Dew et al. 1992, Ogunremi et al. 1999). Reported prevalence in white-tailed deer ranges widely, from 1 to 94% (Lankester 2001). However, many of the lower estimates may be biased by a lack of care or skill in finding adult worms in venous sinuses, inclusion of traumatized heads that cannot be completely examined, or sampling young infected animals in autumn before the adult worms mature in the head or L 1 larvae are produced (Lankester 2001). Prevalences in infection based on L 1 larvae in feces also may reduce estimates of true prevalence (Lankester 2001). population effects Historically, some authors associated marked declines in moose populations and reports of sick moose with incursions by white-tailed deer into these regions; they proposed that P. tenuis brought with invading deer might have caused the moose sickness and population declines (Anderson 1972, Gilbert 1974). However, the idea that white-tailed deer were preventing moose from becoming established in the eastern United States and Canada was challenged (Nudds 1990). Later, using available data on deer and moose population dynamics in several sites, Nudds argued that, with the available information, meningeal worms could

64   nematodes, acanthocephala, pentastomes, and leeches

contribute to moose declines, but that moose could coexist with deer, at lower densities, even in the absence of habitat refuges from meningeal worm (Schmitz and Nudds 1994). In a retrospective study of moose sickness, there was not adequate data to support a relationship between white-tailed deer densities and moose declines (Whitlaw and Lankester 1994). In another study of a declining moose population in Minnesota, meningeal worms were assessed to be a mortality factor of less impact than liver flukes; while there was no relationship between annual population growth and deer abundance, the authors did view deer as potentially serving as reservoir hosts for parasites for the moose (Murray et al. 2006). The role of this parasite as a source of moose decline still is controversial. In contrast, meningeal worms clearly have been associated with the failure of several caribou or reindeer introductions in areas with white-tailed deer (Anderson 1971, Trainer 1973). Based on a review of 33 reintroduction attempts in eastern North America, no caribou reintroductions were successful where whitetailed deer had a high frequency of meningeal worm infection (Bergerud and Mercer 1989). special problems  Parelaphostrongylus tenuis is reported from free-living fallow deer, red deer, reindeer, and moose that have come in contact with introduced infected white-tailed deer in parts of Europe, including Holland, Scotland, Germany, the Czech Republic, Sweden, Finland, Russia (Ural Mountains), central Asia, and south Siberia (Hörning 1975). The parasite causes a meningoencephalitis where these free-living cervids are in contact with introduced whitetailed deer populations (Hörning 1975). control  At doses of 0.1 mg/kg, ivermectin did not prevent infection or eliminate P. tenuis in white-tailed deer, but routine use of ivermectin may temporarily reduce the number of larvae shed in the feces of infected deer (Kocan 1985). However, there is some evidence that an acquired or concomitant immunity follows low-dose infections in white-tailed deer (Slomke et al. 1995) and fallow deer (Davidson et al. 1985).

Filarial Nematodes Filarial nematodes are in the Order Spirurida, Superfamily Filarioidea, Family Onchocercidae, (Anderson 2000). Filarioid nematodes are highly specialized parasites of tissues and tissue spaces of vertebrates other than fish. Mature filarial nematode females are ovoviviparous or viviparous and produce modified L 1 larvae called microfilariae that enter the blood or lodge in the skin of the definitive host (Lee 2002), where they are available to hematophagous arthropods that serve as intermediate hosts or vectors (Anderson 2001). Elaeophora schneideri, a parasite of wild ungulates, will serve to illustrate this group. Although not addressed further in this text, a number of other species affect wildlife. Dirofilaria immitis, the canid heartworm, is transmitted by mosquitoes; although primarily described for domestic dogs, there are increasing numbers of cases being observed in wild canids and a few other species (Abraham 1988). There are a number of other poorly studied but potentially interesting filarial worms of wild mammals and birds (Anderson 2000). elaeophora schneideri (arterial worm) c aus at iv e agen t (cl a ssific at ion, morphology)  Elaeophora schneideri is a medium-sized (6 to 9 cm) filarial nematode found in the carotid and other arteries of susceptible ungulate hosts (Hibler and Adcock 1971). Although there has been some revision of the genus and the species it contains, there currently are at least five other species in the genus Elaeophora, all of which also are parasites of host circulatory systems (Santin-Durán et al. 2000). These include E. elaphi in hepatic vessels of red deer (Cervus elaphus) (Hernández-Rodríguez et al. 1986), E. poeli in the aorta and heart of bovids in Asia and Africa (Sonin 1966), E. boehmi in the arteries and limbs of equids in Europe (Supperer 1953), E. abramovi in hepatic vessels of cervids (Alces alces, Cervus elaphus canadensis, Rangifer tarandus) of Europe and Asia (Oshmarin and Belous 1951), and E. sagittus in the heart of bovids in Africa (Bain and Haesevoets 1974).

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host range and distribution  Elaeophora schneideri occurs primarily in the Odocoileus hemionus complex, comprising mule deer and black-tailed deer, and is distributed in the western United States as far east as Nebraska, and north to British Columbia, Canada (Anderson 2001, McKown et al. 2007). Less commonly, it occurs among white-tailed deer of the southeastern and southwestern United States (Prestwood and Ridgeway 1972, Forrester 1992). Infections among white-tailed deer of the southeastern U.S. are thought to have resulted from interstate movement of animals (Prestwood and Ridgeway 1972). The parasite also has been reported from less well-adapted hosts such as elk (Hibler and Adcock 1971), moose (Worley et al. 1972), bighorn sheep (Ovis canadensis) (Boyce et al. 1999), and domestic sheep and goats (Hibler and Adcock 1971) in the western and southwestern United States, as well as Barbary sheep (Ammotragus lervia) (Pence and Gray 1981) and Sika deer (Cervus nippon) (Robinson et al. 1978) in Texas. Ibex (Capra ibex) also can be parasitized (Pence 1991). life cycles and variations  In the normal definitive hosts, E. schneideri typically is found in the common carotid and internal maxillary arteries, but can occur in almost any artery large enough to hold it (Hibler and Adcock 1971). Microfilariae generally are found in capillaries of the skin on the forehead and face of infected deer (Anderson 2001). The parasite is transmitted by horseflies (Family Tabanidae) of the genera Hybomitra and Tabanus. Microfilariae invade the fat body lining the abdomen of horseflies and develop into L 1 larvae; these, in turn, move to the hemocoel and develop to L 3 larvae before moving to the fly mouthparts for transmission to the next definitive host (Hibler et al. 1971, Hibler and Metzger 1974). After transmission to mule deer from infected horseflies, the L 3 larvae may enter the venous system from the subcutaneous site of entry to be carried to the heart and lungs before moving to the arterial system. They invade the leptomeningeal arteries of the head, develop into immature adults (“L 5” larvae), and move to the carotids, where they mature. The prepatent

period is about 5.5 months (Hibler and Adcock 1971, Anderson 2001). reservoirs and transmission In the western and southwestern United States and in southern British Columbia, Canada, members of the Odocoileus hemionus complex are the reservoir (Anderson 2000) of E. schneideri. In the southeastern United States, white-tailed deer appear able to sustain the parasite population, although the host–parasite relationship appears to be tenuous (Hibler and Prestwood 1981). Transmission occurs by horseflies of the genera Tabanus and Hybomitra (Hibler and Adcock 1971). At least 16 horsefly species have been identified as intermediate hosts, including seven species of Hybomitra and nine species of Tabanus; occasionally other flies have been infected, but appear to be insignificant contributors to this life cycle (Longfellow 1984, Pence 1991). In most geographic areas, there is a single primary vector for E. schneideri, with additional species of lesser importance (Pence 1991). Horseflies generally are not host specific and often have a broad host range (Allan 2001). There is some variation in the reported elevations inhabited by horseflies. In the Gila Forest, New Mexico, horseflies occurred almost completely above 2,000 meters and were most abundant at about 2,300 meters (Hibler et al. 1971). In contrast, horseflies were collected in coastal California at elevations ranging from 610 to 760 meters (Longfellow 1984). The horseflies on South Island, South Carolina (Couvillion et al. 1984), likely also were at low coastal elevations. Depending on the site, reported prevalences in horseflies can range from 0.3% (Couvillion et al. 1984) to over 20% (Hibler et al. 1971, Clark and Hibler 1973). The variation in horsefly prevalences may reflect parasite prevalences in the reservoir hosts (Pence 1991). Horseflies in the Gila Forest, New Mexico, first emerged in early June, peaked by the second week of June, and remained stable until early July, when summer rains began (Hibler et al. 1971). Similarly, in southeastern New Mexico, horsefly numbers reached a peak

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about the third week in June and persisted for 2 to 3 weeks after that (Pence 1991). In coastal Humboldt County, California, capture dates for members of the genera Tabanus and Hybomitra ranged from 2 May to 30 September; some species peaked in June while others reached their peak in July (Longfellow 1984). Most adult tabanid flies are active during the warmer parts of the year, and activity generally occurs during the warmer hours of the day (Hibler et al. 1971, Allan 2001). There are some sites where the greatest activity has been late in the day (Longfellow 1984); also, some tabanid flies are crepuscular (Allan 2001). clinical effects and identification Mule deer and black-tailed deer in western North America are well-adapted hosts for E. schneideri, with little or no associated pathology (Hibler and Adcock 1971). Pathology in white-tailed deer typically is not severe, but may involve intimal thickening, disruption of the internal elastic lamina, and verminous thrombosis in cephalic arteries with E. schneideri (Couvillion et al. 1986). While very well adapted in its normal definitive hosts, E. schneideri can cause considerable pathology and even mortality in abnormal ungulate hosts (Anderson 2001). Most pathology in abnormal hosts is linked to physical blockage of arteries by the nematodes. In elk, blockage of the arterial system of the head can lead to ischemic damage in the brain, optic nerves and eyes, ears, muzzle, and developing antlers (Hibler and Adcock 1971). Bilateral clear-eyed blindness, circling, nystagmus, necrosis of the muzzle and nostrils, dry gangrene of the ear tips, abnormal antler growth, and emaciation are some of the characteristic clinical effects of elaeophorosis in elk (Hibler and Adcock 1971). Clinical signs in moose included blindness, circling, staggering, and ataxia (Worley et al. 1972, Madden et al. 1991). Moose also had fibromuscular intimal proliferation within the carotid and other arteries (Madden et al. 1991). Sheep, goats, and Sika deer often are hypersensitive to the microfilariae and develop

skin lesions of a raw, bloody dermatitis on the forehead or face, called “sorehead” (Robinson et al. 1978, Fraser and Mays 1986). Some thrombi may occur in cerebral and leptomeningeal arteries (Fraser and Mays 1986). Domestic sheep and goats, especially young animals, may die 3 to 5 weeks after infection following a period of incoordination, circling, and convulsions (Fraser and Mays 1986). population effects Between 1960 and 1975, there was an abundance of mule deer in the national forests of the southwestern United States. On the Gila National Forest, New Mexico, elk calf survival was reported to range from 7 to 20% during times of severe tabanid fly and E. schneideri infestation, an inadequate survival for success of the population (Hibler et al. 1969). In 1975, the mule deer population decreased markedly, and the horsefly population also decreased to 90%) are 3-host ticks, having life stages that feed on three different hosts (larvae, nymphs, and adults feed once per life stage, each typically on a different host). Larvae of 3-host ticks find a host, feed for several days until replete (meaning fully fed), and drop off the host onto the soil, leaf litter, or nesting material, where they molt into nymphs. When the season is right, resulting nymphs seek a second host, feed for several more days, and drop off to molt into adult males or females. Mating occurs either in the environment before feeding or on a host as the female feeds; interestingly, the adults of some species (including the Ixodes ricinus group of ticks that vector the spirochetes causing Lyme borreliosis) are commonly found mating in both circumstances. Finally, blood-engorged, mated females detach and drop off their hosts to oviposit their eggs in the environment and to continue the life cycles. In fact, most (again, >90%) of the life cycle of 3-host ticks is spent in the environment and separated from the hosts (Nicholson et al. 2009). The entire life cycles of tropical ixodids may include several generations per year. However, species occurring in regions with more temperate, or colder, climates may require two or more years to complete their cycles. As an example, Ixodes ricinus, an important vector of Lyme disease spirochetes, tick-borne encephalitis viruses, human anaplasmosis, and several other pathogens in Europe, may require up to four years to complete a single life cycle in the northern regions of its distribution (Hoogstraal 1985). parasitic insects, mites, and ticks 139

Figure 5.6  Comparison of representatives of the families Argasidae and Ixodidae: A and B, respectively, are drawings of an Ornithordoros sp. and of the fowl tick, Argus persicus (both argasids); C, D, and E, respectively, are drawings of a larva, nymph and adult male of the American dog tick, Dermacentor variabilis (Family Ixodidae) (from Capinera 2010, copyright © John Wiley and Sons, by permission).

Less commonly, tick life cycles are focused on only two individual hosts; such species are called 2-host ticks. For these species (including Dermacentor hunteri, several species of the genus Rhipicephalus, and some populations of Hyalomma marginatum), fed larvae remain on the first host following feeding, molt, and then re-feed on the same host as a nymph; as with 3-host ticks, the fed nymphs drop off, molt to adults in the environment, and then seek another host (Allan 2001b, Nicholson et al. 2009).

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The extreme modification of the ixodid life cycle is that of the 1-host ticks. Species, including Rhipicephalus (Boophilus) annulatus and the winter tick, Dermacentor albipictus, remain on the host during the larval and nymphal molts. Replete (mated) females drop off of the host to oviposit their large mass of eggs in the environment; these are often large-bodied ticks, and females commonly oviposit more than 4,000 eggs after a single blood meal (Allan 2001b, Nicholson et al. 2009).

Ticks of the Family Argasidae are referred to as soft ticks because they lack the dorsal scutum characteristic of the Family Ixodidae. Argasids also differ from ixodids in having ventrally oriented mouthparts (actually the entire capitulum, composed of hypostome, palps, and chelicerae) covered dorsally by a hood; the mouthparts of argasids cannot be seen when looking directly at the dorsum of the tick. In contrast, the capitulum of ixodids projects anteriorly well beyond the margin of the body (idiosoma) and is obvious when viewed from above (Nicholson et al. 2009). Most importantly for our purposes, the life histories of argasids and ixodids differ in ways that affect their potential to serve as vectors of pathogens. Argasids have life cycles with two or more nymphal stages, and females that undergo several gonotropic cycles; they feed multiple times, laying small batches of eggs after successive blood meals. Many species of ixodids and most argasids are nidicolous, being typically associated with the burrows or nests of their hosts. Nidicolous species tend to specialize on similar types of hosts; some species specialize on colonially nesting sea birds, others on burrowing rodents, and so on. Females of non-nidicolous species detach from hosts and oviposit in the environment irrespective of proximity to host burrows or nests. Nonnidicolous species tend to be host generalists, but the different stages may be found more commonly on some host groups than on others (e.g., lizards, birds, small mammals, large mammals, etc.). As examples, Ixodes pacificus and Ixodes scapularis are generalists in the western and eastern United States, respectively. Larvae and nymphs of both species feed on reptiles, birds, and small mammals, while most adults tend to feed on medium- and large-sized mammals (Nicholson et al. 2009). mesostigmatid mites are often large, semi–free-living, intermittent parasites or predators that live in nests or burrows where they seek cover between blood meals. These mites include the American bird mite Dermanyssus



americanus, rodent mites in the genus Liponyssoides, the tropical rat mite Ornithonyssus bacoti, the free-tailed bat mite Chiroptonyssus robustipes, snake mites in the genus Ophionyssus, the spiny rat mite and others in the genus Laepaps, and chiggers, including Eutrombicula spp. and Neotrombicula spp. The large Family Trombiculidae includes species that are parasitic as larvae, referred to as chiggers, on a wide range of hosts, including amphibians, reptiles, birds, and mammals (Mullen and O’Connor 2009). Many species are known only from their parasitic larvae; the prelarvae, protonymphs, and tritonymphs are inactive and do not feed, while the deutonymphs and adults are predators of other arthropods. Larval chiggers pierce areas of thin skin and feed on cells of the epidermal lining, lymph, or serum. Approximately 20 genera have larvae that are adapted to life as internal parasites of the nasal passages or respiratory airways of reptiles, birds, and mammals. Although larvae are very small, they are often encountered in large numbers, and their infestations can cause considerable annoyance and dermatitis, especially in abnormal hosts (Mullen and O’Connor 2009). prostigmatid mites  include the parasitic Families Demodicidae, Cheyletidae, and Psorergatidae. The Family Demodicidae includes follicle mites that are tiny and elongated (cigarshaped), with very short, stumpy legs (Mullen and O’Connor 2009). Follicle mites live in hair follicles and skin glands of mammals, where they pierce cell membranes to feed on cell contents. The entire life cycle of these mites takes place on the host, and host specificity is typically quite high. Transmission of mites from infested hosts to new hosts is accomplished by intimate contact such as occurs between a female and her offspring. Most individuals in a population tolerate small numbers of these mites without apparent disease. For example, approximately 20% of people 20 years of age, and nearly 100% of elderly people, have follicle mites living in the follicles of their eyelashes, eyebrows, or other facial hair. Young animals

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may develop areas of alopecia (hair loss) or abscessed follicles (depending on the species). Cheyletid mites are small, oval-shaped mites that live on organic debris or that parasitize other mites associated with feathers or fur; parasitic cheyletids of dogs and cats are often referred to as walking dandruff because they appear to be very small bits of skin moving over the pelage. Psorergatid mites include Psorergates ovis, an economically important parasite of domestic sheep, and Psorergates simplex, a parasite of house mice (Mullen and O’Connor 2009). astigmatid mites are tiny parasites that may spend their entire lives, indeed generations, on individual hosts. Examples of parasites of birds and mammals may live in the fur, in the waxy secretions of the ears, under scabs that form on the skin, in dermal burrows, or in the lungs or airways of their hosts. The Families Myobiidae, Listrophoridae, and Myocoptidae are fur mites that parasitize specific species of marsupials, insectivores, rodents, or bats. Most species attach to individual hair shafts and feed on extracellular fluids in the skin, but some myobiids pierce capillaries to feed directly on blood. Dermatitis may be associated with infestations of some species of fur mites, and individual rodents may become irritated enough to chew away areas of fur (known as barbering), but most infestations are apparently well tolerated by their hosts (Mullen and O’Connor 2009).

Examples of Diseases Caused by Insects, Mites, and Ticks Disease conditions are presented in the following section as somewhat isolated problems. However, health results from the combination of all factors influencing a host, and most parasites influence hosts in multiple ways. Bloodfeeding ectoparasites can consume enough blood to cause anemia, and infestations can be associated with inflammatory dermatitis, alopecia (hair or feather loss) with resulting

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environmental exposure and ensuing energetic costs, annoyance, unthriftiness, chronic wasting, secondary bacterial infections, transmission of pathogenic agents, etc. Thus, the following conditions might best be thought of as participants in the cascades of interactions that influence health.

Myiasis Whereas some people are repulsed by thoughts of living animals parasitized by fly maggots, the natural histories of flies with parasitic larvae can be quite fascinating. For example, larvae of many species of flies feed on dead tissue and other organic debris found at the edges of wounds, and the feeding of such species may actually help to keep wounds clean by discouraging secondary bacterial infections. In fact, maggots of several species were used historically to treat human wounds before the advent of antibiotics, and the use of medical maggots is gaining in popularity again because antibiotic resistance of bacteria leaves physicians with few options for successful treatment (Sherman et al. 2000). Myiasis is the term used to describe such parasitism, and three general categories are used to describe infestations depending on the degree to which the fly species is dependent on host tissues: accidental myiasis, facultative myiasis, and obligatory myiasis (Scholl et al. 2009). In accidental myiasis, also called pseudomyiasis, the maggots do not typically parasitize a living host; the host is just an accidental victim. Such accidental myiasis may occur when hosts consume vegetation or carrion infested with fly eggs. These eggs may hatch and the surviving larvae may pass through the alimentary tract of the host. Larvae that persist in the individual host may cause considerable disease, but they are not typically associated with population-level problems for wildlife. Signs of disease associated with accidental infestations range from no apparent signs, through nausea, vomiting, and diarrhea, to more severe

disease resulting from ulceration or perforation of the alimentary tract. In most cases, these larvae do not continue to develop or molt while in the host, so this is truly an accidental interaction (Scholl et al. 2009). In facultative myiasis, fly larvae feed either as saprophytes or as parasites, and they can be thought of as having the ability to opportunistically take advantage of many types of food sources including living flesh (Scholl et al. 2009). These species can be further characterized by their ability to initiate parasitism of living tissues. Primary facultative myiasis involves the invasion of living tissue through a small wound or through intact skin. Secondary facultative myiasis involves species that take advantage of wounds created by other maggots, but that can’t invade a host without having access to a wound. Tertiary facultative myiasis involves those species of flies that invade a host at sites with primary and secondary species prior to host death, but that lack the ability to parasitize healthy hosts. Of course, nature is not limited to such neat categories, and some species of flies with maggots utilizing tertiary facultative myiasis will only rarely feed on living tissues, while others will do so commonly. Many species, including the blow flies, lay eggs on carrion in various states of decomposition. The evolutionary step is short indeed between a carrion feeder and one that feeds on hair matted with feces, urine, or blood or on dead tissues next to wounds. The next step, involving invasion of healthy tissues, is facilitated by the availability of healthy tissues next to such wounds or in association with soiled fur or feathers. Thus, facultative myiasis can be thought of as an evolutionary transition between flies producing accidental myiasis and those whose larvae require living flesh to complete their life cycles (Scholl et al. 2009). North American species whose larvae commonly engage in some level of facultative myiasis include many species of the Families Calliphoridae [Chrysomya spp., Calliphora spp.,



Phormia regina, Cochliomyia macerllaria (the secondary screw worm), and Sarcophagidae (Sarcophaga spp.)]. Calliphorids, known collectively as blow flies or bottle flies, are moderately large flies with shiny metallic cuticles. Blow flies typically lay their eggs in carrion, but many species are attracted to purulent, necrotic wounds, chronic infections, or fur or feather soiled with urine or feces. Larvae feeding in such fetid conditions may invade healthy tissues as opportunity arises, and, if the host dies, overlapping generations of maggots may continue to feed on the dead body. Blow fly maggots drop away from the host prior to pupation in the leaf litter or loose dirt, and adults emerge approximately one week later to feed, breed, and continue their cycle (Scholl et al. 2009). Fly larvae engaged in obligatory myiasis require the tissues of a living host for development to complete their life cycles. Such species include the primary screw worms, bot flies, warble flies, and “blood sucking maggots” (Protocalliphora spp.). Cochliomyia hominivorax is known in North America as the primary screw worm (the Old World primary screw worm is Chrysomya bezziana). Females of this fly are attracted to wounds where they feed and lay eggs on the edges of such sites; screw worm flies commonly infest sites such as umbilical cords of newly born mammals, small wounds resulting from tick bites, and lacerations. Larval screw worms hatch in less than a day after oviposition, and young larvae begin feeding in the exposed wound, enlarging it for themselves and for the next cohort of maggots. The larvae develop quickly through their three instars, drop off of the host to pupate, and emerge as adults in as little as two weeks (Scholl et al. 2009). Such infestations tend to progress and can be highly invasive; death can result from screw worm maggots invading the body cavities and destroying internal organs, from secondary bacterial infections at the wound, or from subsequent septicemia. Moreover, this fly is an

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opportunistic generalist in that the species has been found on a large variety of wildlife, including Virginia opossums, cottontail rabbits, jackrabbits, American black bears, coyotes, white-tailed deer, pronghorn, and feral hogs, as well as pets, livestock, and people (Allan 2001a). The history of primary screw worms in the southern United States is an ecological story involving anthropogenic changes and an interesting method of eradication. By the early 1900s, cattle ranching and farming had altered the landscape of the southern United States, increasing the edge habitats available for deer and promoting the availability of woody legumes that are fed upon by the adult screw worm flies. The increased availability of altered habitats, the movement of cattle across long distances, and the increase in hosts available (including livestock) facilitated an increase in screw worm numbers and an extension of their geographic range (Strickland et al. 1981, Wobeser 1994). Attempts to eradicate the fly from the United States began during the mid-1950s. The strategy for eradication relied heavily on the fact that female screw worm flies mate once with a single male, and scientists reasoned that they could disrupt the fly’s life cycle by blocking the successful fertilization of the next cohort of eggs. Disruption was accomplished by rearing screw worms on artificial substrates and irradiating them to produce sterile male flies. It was important that mass rearing and irradiation did not cause loss of male sexual activity and that females remained receptive to sterile males. As many as 50 million sterile males were released from airplanes annually during the peak production of the program, achieving as high a density as 3,500 flies per km2 in Florida. Local populations declined quickly as the sterile males mated with wild females, and continued efforts eventually suppressed the species throughout the region. The initial goal was to release flies in the northern part of the geographic range and to continue releasing flies each year to push the northern border of the range to somewhere south of ranching interests in the United States. In areas where the species 144

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had been abundant, including most of Florida, Louisiana, Oklahoma, and Texas, losses to the livestock industry were estimated at over $100 million annually prior to eradication efforts (Allan 2001a). Although the species was not eradicated entirely, the suppression of screw worm populations allowed livestock ranching to thrive in the region. Not unexpectedly, such a program was very expensive, with a cost of $10 million annually in the 1950s (Strickland et al. 1981, Wobeser 1994, Allan 2001a). Although sterile male flies are still being used to suppress populations in Mexico, and to drive them further south through Central America, limited outbreaks do still occur north of the Mexican border (Wobeser 1994). Although the motivation for screw worm eradication had little to do with wildlife, many species, including white-tailed deer, benefited from the suppression of this deadly parasite. Screw worm flies were once considered the most important parasite of white-tailed deer in Texas and Florida, and, in some areas of Texas, fawn mortality was estimated to be as high as 80% during years of high fly densities, whereas only 25% died during years with low fly density (Strickland et al. 1981, Wobeser 1994, Allan 2001a). Clearly, the loss of 80% of fawns to a single mortality factor could limit populations of deer (Allan 2001a). Deer did not evolve with such significant losses to screw worms, because the increases in screw worm f ly densities resulted directly from anthropogenic changes associated with cattle ranching. Finally, suppression of screw worm populations in the southern United States was followed by a rapid expansion of deer populations (Strickland et al. 1981, Wobeser 1994), but continued increases in deer populations throughout the eastern United States suggests that many other factors (including changes in land use patterns) are responsible for the continued increase in numbers. Wohlfahrtia spp. (including W. vigil and W. opaca in the New World and W. magnifica in the Eurasia) are obligatory parasites in northern latitudes. Although rabbits and rodents are considered to be the primary reservoirs,

infestations have been reported from a wide range of hosts, including cottontail rabbits, jackrabbits, rodents, mustelids, canids, domestic pets, waterfowl, and humans (Craine and Boonstra 1986, Allan 2001a). Eggs of these species develop and hatch within the females’ bodies, and larvae are larviposited onto moist body openings, fresh wounds, or unbroken skin. The larvae then penetrate and create boillike cysts in the subdermis with a single opening in the skin through which many individual maggots breath. These species tend to parasitize young animals. Feeding by as few as four W. vigil maggots has been associated with mortality of animals as large as fox pups (Craine and Boonstra 1986, Allan 2001a), and this parasite can have significant impacts on host numbers. In one study, four of 43 nests of meadow voles, Microtus pennsylvanicus, were found to be infested with W. vigil (Craine and Boonstra 1986). Nestling voles were found moribund or dead infested with a mean of 10.8 (61.5 SE) maggots per vole, with extensive pathology involving as much as 30% of hosts’ tissues (Craine and Boonstra 1986). While the description sounds dramatic, such losses tend to be local in scope and typically do not limit vole populations. Bot flies and warble flies (Family Oestridae) represent another type of obligatory myiasis, and all species are parasites of mammals. There are many species of such flies, including the rodent and rabbit bots (Cuterebra spp.), warble flies of reindeer and caribou (Hypoderma tarandi), nasal bots of deer (Cephenemyia spp.), and the poetically named blood-sucking maggots (Protocalliphora spp.). The life cycles of the flies presented are superficially similar, but differ in the biology of the species involved, the diseases they produce, and the implications for wildlife management. Rodent and rabbit bots are maggots of Cuterebra spp. (including C. lepusculi and C. horripilum of rabbits, C. emasulator of a variety of rodents including tree squirrels, chipmunks, and deer mice in eastern North America, C. latifrons of woodrats, and C. fontinella and C. jellisoni of rodents in western North America). The adults are large, colorful, robust, bee-like flies with large

eyes and vestigial mouthparts (adults do not feed); lack of functional mouthparts results in adults that live only long enough to mate, develop eggs, and oviposit. Thus, all of the nutrition necessary to complete the life cycle must be obtained during the period of the larval instars (Colwell 2001). Breeding of Cuterebra spp. most commonly involves emergent males swarming over a prominent feature in the environment such as a hill top or large rock outcrop; females that fly through such swarms are quickly caught and mated. Females are univoltine (one generation per year) and oviparous; 1,000 to 4,000 eggs are oviposited on vegetation or substrate near the nests or warrens of their preferred hosts during summer months. After 5–7 days, the eggs hatch in response to higher temperatures and CO 2 concentrations that often indicate presence of a host. First stage larvae crawl and attach to hair of the passing host, enter the body cavities via the eyes, nares, or through a fresh skin wound, and then migrate to the nasal passages, esophagus, or trachea. Second stage larvae continue to wander until they reach the preferred site, depending on the species, in the subdermal tissues (under the skin), where they cut a small hole in the host’s skin. The larvae obtain nourishment by absorbing available host fluids while keeping their posterior ends, containing the respiratory spiracles, lodged against the breathing hole. The host reacts by surrounding the larva with a granulomatous wall or “warble” characteristic of bot fly infestations. Such lesions are most commonly observed in late summer or autumn months of August, September, and October. After growth through four successive molts, the mature larva squeezes through the breathing hole, drops to the ground, and burrows into the leaf litter or loose soil, where it pupates. Pupal diapause allows species in northern latitudes to survive until the combination of environmental characteristics and host availability is right for emergence of the adults (Cogley 1991, Colwell 2001). The impact of cuterebrid maggots on their hosts was initially assumed to be great and has been somewhat controversial parasitic insects, mites, and ticks 145

Figure 5.7  Cuterebra sp. bots in the groin and perineal region of a cotton mouse, Peromyscus gossypinus (Mullen and Catts 2002, copyright © Elsevier, by permission).

(Cogley 1991, Colwell 2001). In fact, C. emasculator, a parasite of mice and tree squirrels, was named because it was thought originally to feed on the testicles of its host. These larvae sometimes are found in subcutaneous tissues of the groin (Fig. 5.7), and their large size appears to displace the testicles, but they do not emasculate their hosts. Bot fly larvae obtain nutrients for growth and development from their hosts, and the energy depletion may impact survival or reproduction of individual hosts during times of resource scarcity. Although rare, larvae occasionally may migrate to abnormal tissues such as the nares or brain, where they may debilitate or kill individual hosts. Likewise, secondary bacterial infections may be associated with bot fly infestations, but other species of bots produce bacteriostatic secretions limiting abscess formation (Beesley 1968). Although it may seem counterintuitive, these maggots only rarely cause serious disease, and impacts on mean fitness or populations appear to be minimal. Hypoderma tarandi is a fairly large fly with a dense covering of yellow and black setal hairs. Adults of these flies, as with other oestrids, lack functional mouthparts and therefore do not feed as adults. In contrast to Cuterebra spp., and other oestrids that swarm over high points in their environment, male Hypoderma spp. swarm over low-lying areas such as dry stream beds or along roadways (Anderson et al. 1994, Colwell 2001). Following mating, females are 146

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known to travel long distances to find their hosts, especially when such hosts are migratory species such as caribou, reindeer, or wildebeest; the maximum flight range of female H. tarandi was estimated to be as far as 600–900 km (Nilssen and Anderson 1995, Colwell 2001). Females are oviparous and, unlike Cuterebra spp., the female lands on the host to oviposit. Eggs are attached firmly to the base of hair shafts via a unique structure at one end of the eggs, and females apparently select individual hair shafts on which to oviposit; the result is that eggs are securely attached and are even somewhat resistant to grooming (Colwell 2001). First stage larvae crawl to the skin surface and regurgitate enzymes that digest collagen, aiding their penetration through the skin. Larvae then migrate through the subcutaneous tissues to eventually reach a location along the dorsum of the host. As with Cuterebra spp., a hole is cut in the skin through which the maggot breathes. As they grow and develop, the larvae keep their anteriorly placed mouth deep in the granulomatous warble. Once matured, larvae exit through the breathing hole and fall to the ground to pupate during late spring or early summer. Emergent adults breed, and the cycle continues prior to the onset of autumn. Individual reindeer can have as many as 2,000 larvae (ranging in size to 2.5 cm in length) scattered under the skin along their backs. Heavy infections cause damage to the skin

and underlying tissues, allergic reactions, nutritional imbalances, secondary infections, and immunosuppression (Karter et al. 1992); Hypoderma spp. from cattle have been shown to secrete a protease enzyme that breaks down host complement (C3), decreasing the host’s potential immunologic response, but this enzyme has not been reported from H. tarandi (Colwell 2001). On warm summer days, large swarms of this fly cause reindeer to seek out patches of snow without flies, and this behavior may limit their ability to forage efficiently under such circumstances. Although it seems clear that Hypoderma spp. adversely impact milk yields in cattle (Reist et al. 2002), and one might assume energetic costs for hosts that reduce their feeding in attempts to avoid the parasites, weight gains of reindeer have been unaffected by control of warble flies (Oksanen and Nieminen 1998). Nasal bots (sometimes called head bots) of deer (Cephenemyia spp.) are robust flies approximately the size of honey bees. Females are viviparous and larviposit (squirt) first stage larvae onto the muzzle or into the eyes of deer without landing; these larvae then crawl through the mucous secretions and membranes into the eye socket, nose, or mouth to find their way to the nasal passages. These yellow to brown maggots use their mouth hooks to attach to the tissues in the nasal passages and the tonsilar pouches of mule deer, black-tailed deer, white-tailed deer, elk, and caribou. Unlike the warble flies and rodent bots that assume subcutaneous locations, nasal bots are not encapsulated by a granuloma. Maggots feed on blood and secretions in nasal passages, where they continue to grow, develop, and molt. When fully mature, the larvae migrate to the opening of the nares and drop to the ground to pupate during the following spring or early summer. Adults emerge from the pupae, mate, and continue the life cycle (Cogley and Anderson 1981). The true impact of these maggots on deer is unknown. Fully mature bots may exceed 25 mm in length, and infested deer may harbor large numbers of such maggots. Although

few clear signs of overt disease are associated with most infestations, annoyance, weight loss, and even death, associated with migration into the cranial cavity, have been reported (Colwell 2001). Surely, the maggots block the nasal passages, and their growth in confined spaces seems likely to produce annoyance and potentially pain. Although only the maggots parasitize the hosts, it may actually be the indirect effects of the adults that cause population-level phenomena by causing avoidance behavior (Anderson 1975, Karter and Folstad 1989, Moerschel and Klein 1997, Anderson 2001). Deer may stand with their noses to the ground or buried into their sides, and (reportedly) try to run away from the buzz of the females’ wing beats (Anderson 1975, 2001). Reindeer seek snow or ridge tops with cooler temperatures, stronger winds, and fewer flies. They spend more time walking and running and less time lying down and eating (Karter and Folstad 1989, Hagemoen and Reimers 2002). Reduction in the relative time spent grazing during short Arctic summers may affect body condition (and potentially overwinter survival and reproduction) of reindeer and caribou (Downes et al. 1986, Moerschel and Klein 1997). Moreover, the stress induced by oestrid flies and fly avoidance causes physiologic responses, including activation of opioid peptides that act as neurochemicals effecting a wide range of responses that influence immunologic function and fitness (Colwell 2001). The final example of myiasis involves the larvae of Protocalliphora spp., which are aptly called blood-sucking maggots. Adults of the different species of Protocalliphora lay their eggs on nestling birds or in the nests near newly hatched nestlings. The fly eggs hatch within 24 to 48 hours. The larvae of most species crawl to a place in the nest near a nestling where they can feed intermittently on nestling blood or other fluids through the skin of the feet and legs. Other species have larvae that directly enter the nostrils or ears, and P. braueri larvae burrow through the skin of nestlings to continue to feed as internal parasites parasitic insects, mites, and ticks 147

(Bennett 1957, Rogers et al. 1991, Whitworth and Bennett 1992).

Blood Loss and Hemolysis Blood is an extremely important tissue that transports oxygen, nutrients, and biochemicals (including hormones) to other tissues, carries metabolic waste products away from tissues, and contains many of the cells and proteins important to the immune system. Blood loss can result in a number of problems for hosts, and anemia (lower than normal volume or percentage of red blood cells or hemoglobin) can occur acutely when the number of red cells drops precipitously or develops more slowly with chronic disease. There are many causes of anemia, including hemolysis (the internal rupture of red blood cells, as occurs with some toxins that distort red blood cells, making them susceptible to elimination by the spleen, or with infections by protozoa causing malaria, leucocytozoonosis, hemobartonellosis, etc.). Alternatively, anemia may develop from an inability to create red blood cells (as occurs with some types of cancers and iron deficiency) or more directly from loss of blood. Blood loss can be associated with many etiologies (causes) including trauma (wounds, car collisions, etc.), the rupture of aneurysms, and internal or external parasites that feed on blood. Although ectoparasites may seem too small in relation to the host to remove enough blood to cause problems, large numbers of blood-feeding ectoparasites can result in both debility and death under the right (or wrong) circumstances. We describe three examples: feeding by bloodsucking maggots, swallow bugs, and ticks. The first example is of blood loss caused by blood-sucking maggots mentioned above under the heading “myiasis”. These are muscoid flies, related to house flies (Musca domestica), and various other species of Calliphorid blow flies (Calliphora spp., Phormia spp., Lucilia spp., etc.) that lay eggs, hatch as small larvae, undergo three larval instars as maggots, pupate, and then feed, mate, and oviposit another 148

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generation of eggs as adults. The genus is widespread in North America, and most nesting avian species, including songbirds, swallows and martins, woodpeckers, hawks, and owls appear to be susceptible to infestation by one or more species of Protocalliphora (Bennett 1957, Rogers et al. 1991, Whitworth and Bennett 1992). The effects of blood-sucking maggots on nestling survival range from unapparent effects to mortality and nest abandonment. Mature larvae of Protocalliphora spp. are large parasites in relation to nestling size, varying from about 7 to 17 mm (¼ to ¾ inch), and each larva can consume a considerable volume of blood. When nests are infested with hundreds of such larvae, the result can devastate the entire brood (Bennett 1957, Rogers et al. 1991, Whitworth and Bennett 1992). The most common effects on individual birds include those typical of blood loss: anemia, weakness, and reduced growth rates. However, maggots that invade the hosts also cause direct damage to tissues, and all species create wounds that promote secondary bacterial infections. Anemic, debilitated nestlings grow slowly, increasing the duration of time spent in the nest and the probability that nestlings will succumb to predation prior to fledging. In other instances, the presence of large numbers of blood-sucking maggots may cause the parents to desert the nest. One report noted that 5–10% of infested nestlings died of blood loss and others that survived to fledge may have been less resistant to diseases and more susceptible to predation than would be strong healthy fledglings (Rogers et al. 1991). Mature larvae undergo metamorphosis as pupae in the environment. Adults that emerge near the beginning or middle of the nesting season may repeat the cycle. Others that emerge near the end of the songbird fledging period over-winter in the nests or under bark or cracks in downed woody debris and lay eggs on nestlings during the following spring. Some species of sucking lice have been associated with blood loss. As an example, the African blue louse, Linognathus africanus,

is non-native in North America and has been associated with anemia and death of Columbian black-tailed deer (Odocoileus hemionus columbianus) and white-tailed deer (O. virginina) fawns (Brunetti and Cribbs 1971, Foreyt et al. 1986, Durden 2001) in the western United States. Swallow bugs, Oeciacus vicarious and Oeciacus hirundinis, illustrate another example of blood loss. These insects are related to human bed bugs (Order Hemiptera, Family Cimicidae), and, like bedbugs, they reside near places of rest and feed while the hosts are sleeping. Swallow bugs reside in cracks in the mud nests of cliff swallows [Petrochelidon (formally Hirundo) pyrrhonota], purple martins (Progne subis), and potentially other socially breeding species in the Family Hirundinidae. Like other hemipteran bugs, there are five nymphal stages and the adults, which develop through a life cycle that often matches the timing of their migratory hosts. Although bats and other birds sometimes use swallow nests in their absence, and probably provide occasional blood meals for the bugs, most populations of bugs in temperate zones are forced to survive the 7–9 months of host absence prior to the migratory return of their primary hosts, that is, swallows. Surviving bugs help to reestablish the bug colony during the early phases of nest occupancy, and numbers increase throughout the breeding season. Further, nymphal swallow bugs appear to be transferred between nests by birds. Eggs of Oeciacus spp. are glued to the outside of mud nests, and, in heavy infestations, large numbers of eggs plastered to the nests can be seen from a distance (Krinsky 2009a). Heavily infested nests may produce enough immature and adult swallow bugs to literally drain the nestlings of their life blood, causing reduced growth, early fledging, and sometimes death or abandonment, of entire broods (Chapman and George 1991, Loye and Zuk 1991). Of course, these are not the only blood-sucking nest parasites that take advantage of the colonial breeding swallows. There are also mites, mosquitoes, and several species of ticks, including Argas cooleyi (Acari: Argasidae), Ornithodoros concanensis

(Acari: Argasidae), and Ixodes baergi (Acari: Ixodidae), are commonly associated with swallow nests. Swallow bugs transmit at least one important virus, a togavirus (Alphaviridae) known as Buggy Creek virus, to the birds. Several authors have reported the prevalence of the virus among swallows (Hopla et al. 1993, Brown et al. 2001), but the impacts of the virus on populations of swallows is not well understood. Interestingly, both the bug population density and the prevalence of the virus in the birds are directly correlated with the swallow colony size (Brown et al. 2001). Such a correlation allows the prediction of ectoparasite populations and virus persistence at colonies of swallows, and it suggests that there are costs associated with colonial breeding that may be directly associated with colony size. When we think about the evolution of colonial breeding, the effects of parasites and disease transmission should be among the variables considered. All ticks are parasitic, and they all suck blood. However, the winter tick, Dermacentor albipictus, is notorious for the ability of large numbers of ticks to cause severely debilitating anemia and even death of large ungulates. This tick parasitizes domestic livestock, deer, elk, bighorn sheep, and moose in many areas throughout North America, from northern Mexico to approximately 60° north latitude in Alaska and Canada, but the problem becomes most extreme for moose in southern Canada (Samuel et al. 1991, Allan 2001b, Samuel 2004). Dermacentor albipictus is a 1-host tick; once larvae attach to a host they remain on that host for the remainder of their lives, only dropping off as fed females to oviposit more eggs into the environment. Especially in northern regions, this species follows a regular rhythm of seasonality with one generation per year in which adult females engorge between February and May, each producing thousands of eggs (up to 4,400!) that hatch into hungry larval “seed ticks” during autumn (Allan 2001b, Samuel 2004). Moose especially feed large numbers of parasitic insects, mites, and ticks 149

Figure 5.8  (A) A moose with alopecia and wasting associated with severe Dermacentor albipictus infestation and the condition known as ghost moose syndrome; (B) D. albipictus feeding on a moose with broken hair shafts (from Samuel 2004 and courtesy of Bill Samuel and Federation of Alberta Naturalists).

these ticks, and an “average” moose might harbor 33,000 to 50,000 ticks. Even higher numbers are found on some animals, including a bull with 150,000 ticks and a calf with 145,000 ticks (resulting in densities of 5.7 and 7.8 ticks per square centimeter or 37 and 50 ticks per square inch!) (Samuel 2004). Summarizing decades of work on winter ticks on moose, Samuel (2004) estimated blood loss in moose as follows: bulls, cows, and calves with median numbers of ticks were found to feed 5,441, 3,161, and 8,123 female ticks per year, respectively. Each female tick was estimated conservatively to remove 1 gram of blood, equating to an estimated 5.4 L, 3.2 L, and 8.1 L of blood from the median bulls, cows, and calves, respectively. Such losses result in a required minimal replacement of 16.9%, 11%, and 57.9% of the blood volume of these median animals, respectively. Moreover, this blood loss occurs during the late winter and early spring, when temperatures are coldest, food is least plentiful, and thermoregulation afforded by a dense hair coat is most important (Glines and Samuel 1989). Disease becomes apparent when heavy tick infestations cause annoyance and inflammation of the skin, resulting in grooming, scratching, and rubbing of the skin, and leading to damage of the hair coat and hair loss (alopecia). This combined nutritional and cold stress is exacerbated by the anemia caused by significant blood loss. Anemia causes animals to become weakened and lethargic, further reducing their ability to successfully forage and groom. Thus, weak animals become weaker, more ticks feed to repletion, hair loss and anemia worsen, sick animals become sicker, and some may die of malnutrition, exposure, decreased resistance to other parasites or diseases, or to reduced ability to avoid predation. In Canada, severely affected moose are called ghost moose because damage and loss of the dark (brown or black) tips of the guard hairs exposes the inner hair shafts, which tend to be gray or whitish in color (Fig. 5.8). The appearance of such moose is often that of an

undernourished animal with a ragged hair coat; the dorsal neck mane is often destroyed, there may be large patches of skin without hair or with only sparse and broken hair, and the shoulders and backs of the animals may have large gray to whitish patches (Samuel 2004). Although discussed here under blood loss, ghost moose syndrome would fit just as well with the next section.

Dermatitis The skin is an amazing first defense against potential infectious agents, but it is not impervious. Cellular immune reactions to parasites and pathogens (helminths, protozoa, fungi, bacteria, and viruses) that colonize the skin surface, or in deeper layers, involve changes in the microvasculature and migration of white blood cells into the area. These changes result in the signs we all recognize as inflammation: tissues become swollen, red, hot, and painful (or itchy, depending on the severity); refer to Chapter 2. Some types of inflammation cause blisters to form, and sometimes ulceration of the epidermis exposes the deeper layers to secondary infections. Chronic inflammation can cause skin to become thickened (hypertrophied), dry, and flaky. Hair and feathers may become brittle and break, or they may fall out, get abraded from rubbing, or be plucked out. Hypersensitivity reactions to proteins in arthropod saliva often develop as a result of repeated bites of fleas, flies, bugs, lice, mites, or ticks. Such reactions may cause intense itching to the point of disrupting normal behavior and can even result in obsessive self-mutilation. Dermatitis varies from the inconsequential reaction to a single mosquito bite to the full-body loss of hair, with thickened, cracked, bleeding, infected, and unimaginably itchy skin associated with severe sarcoptic mange. Of course, the combination of chronic itching, hair or feather loss, and altered behavior are exacerbated by secondary bacterial infections at sites with cracked, ulcerated arthropod-bite wounds, or mutilated skin, as well as to exposure to environmental parasitic insects, mites, and ticks 151

conditions, chronic fatigue, blood loss, and transmission of parasitic organisms by biting arthropods. Examples of dermatitis induced by arthropods include hypersensitivity reactions to antigens associated with blood-feeding ectoparasites (such as sucking lice, kissing bugs, fleas, mosquitoes, tabanid flies, ticks, etc.), and the skin, feather, and hair damage associated with large numbers of chewing lice. Two species of non-native chewing lice, Damalinia sp. and Bovicola tibialis, cause hair loss syndrome of deer in the western United States. Hair loss syndrome (HLS, or hair slip syndrome) caused by Damalinia sp. was first observed in populations of black-tailed deer in western Oregon and Washington in 1995 and had expanded south and east by 1998 (Bildfell et al. 2004). In 2005, another exotic louse, B. tibialis, was associated with HLS and mortality in central Washington, and both lice are now known to cause HLS as far south as the southern Sierra Nevada Mountains of California; and B. tibialis extends as far eastward as New Mexico and Texas (Westrom et al. 1976, Mertins et al. 2011). Hair loss syndrome presents as white or yellowish patches of fur of the thorax, neck, flanks, and rump of deer; discolored patches are coincident with the loss of guard hairs, revealing the under-fur. Hair loss may progress to loss of body condition, emaciation, and death. Hair loss syndrome typically causes more severe signs of disease in fawns than in adult deer, and loss of condition due to exposure has been associated with fawn mortalities that peak in late winter and early spring (Bender and Hall 2004, Bildfell et al. 2004). The causes of death of fawns with HLS often seem associated with additional health problems, including lungworm pneumonia, nutritional deficiencies, and predation, and it may be that multiple causes commonly act synergistically to result in HLS deaths (Foreyt et al. 2004). Although prevalence of lice, and mortality from HLS fawns, may be high in some deer populations, at least one study has shown that over-winter survivorship of fawns in different management units did not 152

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correlate with prevalence of HLS (Bender and Hall 2004). Obviously, more work is needed to tease apart the relative importance of lice, HLS, and other mortality factors as influences of survivorship of deer fawns in areas with HLS (Krinsky 2009a). Most lice spend their entire lives on a single host, transmission of pathogens from host to host is rare, and relatively few disease agents important to wildlife populations are transmitted by lice (Durden 2004, Durden and Lloyd 2009). However, some lice do serve as important intermediate hosts of helminth parasites, including Trichodectes canis, an intermediate host of the double-pored tapeworm, Dipylidium caninum, of wild canids (Durden 2001, Durden and Lloyd 2009), and Trinoton anserinum, an intermediate host of the swan heart worm, Sarconema eurycerca (Seegar et al. 1976, Cohen et al. 1991). Sarcoptic mange, or scabies, is caused by parasitism of mammals by the burrowing itch mite, S. scabei. The mite has been reported to cause disease in a large number of wild mammals, including marsupials, insectivores, rodents, lagomorphs, primates, carnivores, and ungulates (Bornstein et al. 2001, Pence and Ueckermann 2002, Mullen and O’Connor 2009). Mites in the genus Notoedres are superficially similar to S. scabei in morphology, disease produced, and general ecology, but they have a more narrow range of hosts; N. cati parasitizes mainly wild and domestic felids (Pence et al. 1982, 1995; Ryser-Degiorgis et al. 2002), and coatis (Valenzuela et al. 2000), and civets (Ninomiya et al. 2003), and N. centrifera causes disease in tree squirrels (Cornish et al. 2001). Development of clinical mange varies among species, by immune status of the host, and with population characteristics of the mites; it is intensely pruritic (causing intense itching) in most affected individuals, but some infested red foxes have failed to show signs of itching (Bornstein et al. 2001). Signs of disease typically include rash, dry, thickened, crusted, hyperpigmented skin, and alopecia. In addition, severe cases in wildlife may be associated with lymphadenopathy (swollen lymph

nodes), inappetence, secondary bacterial infections, behaviors allowing approach by people, and death. Mangy wildlife typically appear unthrifty, with matted, odiferous hair coats, and thickened, crusty, hyperpigmented skin. In severe cases, alopecia (hair loss) may cover most or all of the body, and death likely results from exposure, secondary bacterial infections, and the energetic costs of fighting chronic disease. While epizootics of mange seem dramatic due to loss of life and the appearance of obviously diseased animals, Pence and Ueckermann (2002) point out that mange does not typically affect long-term population dynamics of self-sustaining populations of wildlife. Of course, small or threatened populations may suffer decimating losses when facing multiple insults, when loss of individuals affects population viability, when generalist parasites such as S. scabei are maintained by a community of hosts, and when researchers facilitate transmission through a diseased population (Cornish et al. 2001, Pence and Ueckermann 2002). Psoroptes spp. are astigmatid mites that live in the ears of rabbits and ungulates, but they may spread out over the neck, shoulders, torso, and flanks when mite populations are unabated. Several species of Psoroptes were originally characterized by the host from which they were collected; Psoroptes cuniculi from rabbits, Psoroptes cervinus from elk and deer, Psoroptes ovis from sheep, Psoroptes equi from horses, and so on. However, the validity of these species has been questioned and the need for taxonomic revision has been noted (Boyce and Brown 1991, Ramey et al. 2000, Zahler et al. 2000). A recent molecular comparison of Psoroptes spp. mites from different host species did not reveal species-specific patterns, and Zahler et al. (2000) suggested that all of these mites be considered conspecifics under the binomial associated with the earliest species description, that of P. equi. Even so, there remains enough population-level adaptation that mites may be unable to survive transfer to hosts of different species.

The life cycle of the astigmatid mite P. equi (including all related strains or subspecies) is interesting in that it depends, in part, on the host response, and the disease that results varies from minor annoyance to lifethreatening illness. Many generations of these mites occur on a single host animal, and the hypersensitivity response stimulated during feeding on host fluids results in serum being exuded through the skin. The mites live on the skin surface, feeding on the serum and dead cells. The serum exacerbates local inflammation, causing ulcerated lesions that result in scabs over the lesions. Scabs provide cover for the dense mat of mites, protecting them from the elements and the grooming of the host, as well as a providing a constant food supply (Mullen and O’Connor 2009). Transmission from host to host occurs with intimate contact (Mullen and O’Connor 2009). However, mites also survive off the host (in bedding, shed fur, etc., and depending on conditions) for as long as 10 to 15 days, facilitating potential indirect transmission (Meintjes et al. 2002). The extent of the scabby lesions, and associated hair loss, depends on the density of mites present, specific characteristics of the population of mites, and the hypersensitivity response of individual hosts. Some animals may harbor only a few scab mites in their ears with little apparent pathology. Others develop scabs in their ears that have been shown to block hearing, and loss of hearing might be expected to be associated with increased risks of predation (Norrix et al. 1995). Still other hosts succumb to exposure or secondary bacterial infections resulting from the combination of scab formation and hair loss over large areas of their bodies. Domestic sheep are often carriers of P. equi (subsp. ovis), and contact between domestic sheep and bighorn sheep has been responsible for die-offs of the latter in the western United States (Mullen and O’Connor 2009). The alopecia and scabs resulting from heavy infestations of P. equi are sometimes referred to as scabies (as with “bighorn scabies”). In wild ungulates, severe disease has been reported parasitic insects, mites, and ticks 153

among bighorn sheep (Lange et al. 1980, Welsh and Bunch 1983, Boyce and Brown 1991, Mazet et al. 1992, Singer et al. 1997), but infestations of mule deer, white-tailed deer, elk, ibex, and pronghorn also occur (Wright and Glaze 1988, Garris et al. 1991, Samuel et al. 1991, Ziccardi et al. 1996, Singer et al. 1997, Yeruham et al. 1999, Mullen and O’Connor 2009). In most infestations, mite populations are limited to the ears, resulting in few signs of overt disease. In mild cases in deer, inflammation may lead to mild bleeding and oozing of serum; such mild disease may be limited to small, discrete crusts on the inner surfaces of the ear pinnae and auditory canal. Indeed, where populations of white-tailed deer in the southeastern United States commonly suffer only mild disease, crusty lesions and inapparent infestations were observed with up to 80% prevalence (Roller et al. 1978, Garris et al. 1991). When cases progress to larger crusts with hair loss in and around the ears, the inflammation and pruritus may produce behavioral signs associated with head shaking, scratching, and rubbing on objects. In more severe cases, mite populations, and the inflammatory scabs they create, promote secondary bacterial infections. Ears may become filled with purulent discharge and infections may spread to the inner ear. Inner ear infections cause obvious changes in behavior, circling, loss of balance, and incoordination, and may result in death. Otitis externa (infection of the outer ear canal by bacteria and fungi) caused by mite-induced inflammation has been associated with occlusion of the ear canal of bighorn sheep (Norrix et al. 1995), and the subsequent reduction of hearing led the authors to logically suggest a connection between loss of hearing and increased susceptibility to predation. More commonly, as populations of mites grow large, they spread out from the ears over the trunk and body. Chronic severe infestations may lead to environmental exposure, weight loss, anemia, secondary infections, and death (Mullen and O’Connor 2009), but the chronic carrier state is common in most infested populations of wild ungulates. 154

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Mites in the Family Sarcoptidae are more invasive than both P. equi and the fur mites previously discussed. These species also live for generations on individual hosts, but they are not limited to the surface of the skin. Sarcoptes scabei and Notoedres spp. are burrowers that excavate tunnels under the skin of mammals. Sarcoptes scabei infests a wide variety of wild mammals, including carnivores, ungulates, primates, and bats, and has worldwide distribution. Many researchers have commented on the potential number of species of the genus Sarcoptes associated with different host species, but others have advocated for one highly variable species (Bornstein et al. 2001). A molecular comparison of 23 isolates of S. scabei collected from four continents from dogs, foxes, lynx, raccoon, camel, chamois, a wombat, and livestock supported the existence of a single, variable species (Zahler et al. 1999). However, mite populations adapt to host species, and most transmission, which occurs via direct contact, is thought to be intraspecific rather than between species (Mullen and O’Connor 2009). Male and female S. scabei breed on the skin of the host, and fertilized females burrow into the epidermis, depositing eggs in the burrows. Females lay up to three eggs per day for a period of 2–3 weeks before dying (Mullen and O’Connor 2009). Eggs hatch into larvae after 3 to 8 days, and larvae migrate to the surface to molt twice before maturing into adults. The entire life cycle requires only 14 to 21 days (Mullen and O’Connor 2009). Although a large majority of mites live on the surface, females defecate and die in the burrows, and the egg capsules remain in the burrows after the larvae hatch. Thus, considerable antigen remains in the burrows to initiate a hypersensitivity response that creates intense itching, and this hypersensitivity response to chronically present parasite antigen can be overwhelming for individual animals. It is the host’s response, and not the mites per se, that causes the pathology associated with sarcoptic mange (Mullen and O’Connor 2009). Disease caused by

Notoedres spp. is similar to mange caused by S. scabei, but these mites are associated with felids, civets, squirrels, porcupines, and a number of Old World species of rats and mice. Mange was discussed further under the section on dermatitis. The Families Knemidocoptidae and Epidermoptidae include several genera of mites superficially similar to S. scabei and Notoedres spp., but mites in these families cause mange in birds instead of mammals. Like S. scabei, these mites live on the skin, under scales, or in feather follicles. Fertilized females burrow through the epidermis, lay eggs in the burrows, and newly hatched larvae return to continue the cycle on the surface of the skin. Hypersensitivity reactions lead to thickened, crusty skin; scaly-leg; scaly-face; and small or large areas of feather loss. Finally, the Family Oribatidae includes freeliving mites that feed on organic detritus. These mites do not directly cause disease in wildlife, but some species serve as intermediate hosts of tapeworms, including the anoplocephalid Moniezia expansa. Definitive hosts accidentally ingest these small mites along with vegetation. Later, after the tapeworms have matured and eggs have been shed with host feces, oribatid mites feeding at the soil surface may ingest tapeworm eggs as they feed on organic debris (Mullen and O’Connor 2009).

Annoyance In addition to the direct disease discussed above, ectoparasites likely cause considerable irritation and annoyance of wildlife. “Fly worry” has been a known cause of decreased weight gains and milk production in cattle (Stork 1979, Jordaan and van Ark 1990, Wieman et al. 1992, Jonsson and Mayer 1999, Campbell et al. 2001, Cilek and Hallmon 2005). Such decreases in weight gain or milk yield occur because cattle feed less and expend energy avoiding flies when large densities are present. Such arguments have been extended to reindeer that clearly avoid swarms of biting flies

during Arctic summers (Karter and Folstad 1989, Nilssen and Anderson 1995, Hagemoen and Reimers 2002). Reindeer and caribou seek snowdrifts, windy ridges, water, or other areas when fly densities are high. When unable to seek habitats with fewer flies, they may run to avoid the biting swarms. However, decreased weight gain of reindeer is not as clearly associated with annoyance as is that of cattle (Oksanen et al. 1993). Columbian black-tailed deer have been shown to behaviorally avoid flying female Cephenemyia spp. (Anderson 1975, 2001). However, the energetic impacts of avoidance of flies on the fitness of deer are not clear. While it seems logical that dense populations of ectoparasites could cause enough annoyance or irritation to reduce fitness of heavily infested individuals, such effects may be subtle, and the demonstration remains elusive. Annoyance may be expected to impact individuals during any given season, and to affect population dynamics for short periods of time under severe circumstances, but annoyance should not be thought of as a limit to populations of wildlife.

Toxicosis Toxicosis results from the envenomization of wildlife by spiders, scorpions, ants, bees, and wasps; the poisonings related to consumption of poisonous insects, and toxins in the saliva of some ectoparasites. In general, toxicosis may cause disease in individual wildlife but does not impact populations. Two of the more interesting types of toxicosis result from attachment of specific species of ticks. Tick paralysis results from neurotoxic proteins in the saliva of specific species of Ixodes, Dermacentor, Amblyomma, Rhipicephalus, Argas, and Ornithodoros ticks. In fact, at least 46 species of ticks in 10 genera have been associated with tick paralysis (Nicholson et al. 2009). In the United States, paralysis is caused by D. andersoni in the northwestern and Rocky Mountain areas, and D. variabilis in the eastern regions. In Africa, R. evertsi and parasitic insects, mites, and ticks 155

O. savignyi cause paralysis in livestock and wildlife. Paralysis most commonly affects dogs, cats, and poultry. However, other species, including livestock, wildlife, and people, also are reported to suffer occasionally from tick paralysis; such events in wildlife are likely to be grossly under-reported. Paralysis has been best described in relation to the bite of I. holocyclus in Australia, from which a salivary toxin has been isolated and described (Stone et al. 1989), and the protein causing paralysis appears to be somewhat similar among different species of ticks (Crause et al. 1994). Most cases result from large female ticks feeding on small or young animals. The signs of disease typically begin as weakness and incoordination of the limbs. As the toxin continues to affect the host, ascending flaccid paralysis progresses eventually to cause failure of the heart or diaphragm, followed rapidly by death. Removal of the tick from animals suffering tick paralysis may result in rapid improvement in people, pets, livestock, and wildlife, but cases caused by I. holocyclus may take weeks or months to resolve (Nicholson et al. 2009). Tick paralysis in wildlife has been reported in a western harvest mouse (Botzler et al. 1980), a red fox (Little et al. 1998), and at least 64 species of songbirds (Luttrell 1997). Wildlife succumbing to tick paralysis may be difficult to detect and may often die from other causes, such as predation; thus, cases of tick paralysis in wildlife are rarely reported. Tick toxicosis is a similar response to unrelated proteins in tick salivary gland secretions. The disease manifests as an acute, febrile illness ranging in scope from localized inflammatory reactions, as is often associated with the bite of I. pacificus in the western United States and of I. holocyclus in Australia, through moderately severe, potentially life-threatening reactions to bites of the soft tick O. savignyi in Africa (Mans et al. 2003). Life-threatening systemic reactions, called sweating sickness of livestock and wild ungulates, are associated with the bites of Hyalomma spp., especially 156

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H. truncatum females, in Africa. In the most severe cases, sweating sickness presents as sudden fever, followed by a wet rash over the body, necrosis of the oral mucosa, sloughing of hair and skin resulting in painful, ulcerated wounds prone to secondary bacterial infections causing emaciation, dehydration, depression, and death. Animals that survive are protected from future toxicosis by an immune response to the toxin.

Arthropods as Vectors Concepts of population persistence, metapopulations, and the importance of dispersal routes connecting semi-isolated or small subpopulations are familiar to most students of wildlife biology and management. With our increasingly patchy, fragmented landscapes, humans intensify the need for corridors to promote gene flow and to provide a rescue effect for populations of wildlife that are slowly going extinct. Parasites also face such obstacles to individual survival and population persistence, and one of the most crucial of obstacles to parasite population persistence is that involving dispersal between hosts. The vector-borne diseases of humans and domestic livestock, including the zoonotic diseases maintained in wild or domestic animal reservoirs, are explained in a large, and ever-growing, scientific literature, and related topics are reviewed in general texts on zoonotic diseases and medical or veterinary entomology (Acha and Szyfres 2001, 2003a, 2003b; Marquart et al. 2004; Mullen and Durden 2009). Likewise, arthropod vectors transmit many agents causing wildlife diseases. Disease ecologists typically use the term vector in relation to arthropods, and the pathogens transmitted by arthropod vectors are referred to as being vector-borne. However, the term vector also can be applied broadly to any animal or even a fomite that transmits an infectious agent. Thus, it is generally helpful to define such terms when used, and we will restrict our discussion of

vector-borne pathogens to those transmitted by arthropods. The nature of the relationships of a pathogen to both the vector and the host determines the details of biological cycles necessary for transmission, and knowledge of such details is required for an understanding of, and potential management of, the cycles. Vector-borne pathogens can be transmitted vertically from a female vector to her offspring or horizontally among hosts or between hosts and vectors (Reisen 2009). Transovarially transmitted pathogens are transmitted from female arthropods to their eggs, with subsequent larvae amplifying the prevalence of the pathogen through the vector population, but this requires that the pathogen not kill the developing eggs or larvae. Pathogens that are transmitted within an individual vector from stage to stage (e.g., viruses or bacteria that persist in a larval tick through the molt to the nymphal stage) are referred to as being transstadially transmitted. Phoretic transfer describes the situation where arthropods are moved between hosts by other arthropods; examples include mites and lice that hitchhike between hosts on ectoparasitic flies. The term mechanical transmission describes the movement of parasites or pathogens between hosts by arthropods; in this case, the vector functions merely as transportation for the pathogen. In contrast, biological transmission describes situations in which parasites or pathogens undergo an obligatory increases in number or development within the vector; in biological transmission, the vector serves as a required host as well as an agent for transportation (Reisen 2009). Examples of mechanical vectors include cockroaches or flies that carry fecal bacteria on their legs or mouthparts to food that is then ingested along with the agents that cause giardiasis (Giardia spp.) and avian salmonellosis (Salmonella spp.). Likewise, the deer flies and horse flies that serve as vectors of Francisella tularensis (the cause of tularemia), and the mosquitoes that transmit the viruses that causes avian pox, are mechanical vectors of the pathogens listed.

There are many examples of parasites that serve as biological vectors. The vector–parasite interactions involving biological vectors and their parasites are necessarily more complex than the interactions between parasites and mechanical vectors. Examples of parasites transmitted biologically include the arboviruses (including West Nile virus and the western and eastern equine encephalitis viruses), Borrelia burgdorferi sensu lato (the group of related spirochetes that cause Lyme borreliosis), Yersinia pestis (the causative agent of plague), various rickettsial agents (including Anaplasma spp. that cause anaplasmosis in humans and other mammals), Plasmodium and Leucocytozoon spp. (which cause avian hemosporidiosis or avian malaria), Elaeophora schneideri (an arterial worm of ungulates), and heart worms in the genera Dirofilaria, Sarconema, and Cordolobia. In biological transmission, the parasites must undergo development, multiplication, or both during the period between exposure of a biological vector to a parasite and the time when it becomes infective to a naïve host; such requirements relate directly to the life history characteristics of the parasites. When only multiplication is required, the process is termed propagative transmission. When developmental changes are required, the process is termed cyclodevelopmental, or just developmental, transmission, and when both multiplication and development are required the process is referred to as cyclopropagative transmission (Reisen 2009). Thus, the arboviruses, B. burgdorferi, Y. pestis, and Anaplasma spp. all must multiply in their vectors prior to effective propagative transmission; filarial worms must develop in their vectors prior to reaching the infective L 3 stage prior to successful developmental transmission; and the protozoa causing avian malaria must both develop and multiply in their vectors prior to transmission of infective sporozoites via cyclodevelop­ mental transmission. Biological transmission often requires that a vector remain infected and pass the infection on through either transstadial (stage to stage) parasitic insects, mites, and ticks 157

or transovarial (from a female to her offspring) transmission. There are many different potential blocks to such transmission. For instance, insects have a membrane lining their gut, the peritrophic membrane, that effectively blocks parasitic infections of many potential insect vectors (Harwood and James 1979, Gullan and Cranston 2005, Mullen and Durden 2009). In those cases of salivary transmission of pathogens (such as occurs with West Nile virus, various heart worms mentioned above, and parasites that cause avian malaria), the parasites have adapted some mechanism to cross this membrane in order to move from the gut to the salivary glands. Agents transmitted by ticks, such as Borrelia spp., have evolved characteristics favoring their retention during the complete reorganization inherent in the transstadial molts (Allan 2001b, Nicholson et al. 2009). Indeed, pathogens have evolved many creative ways to ensure their transmission among hosts. However, morphological features and life history characteristics of potential vectors also influence transmission in predictable ways (Black and Kondratieff 2004). Vectors that feed on the blood of multiple hosts during their life cycles, that feed on a wide range of host species, and that have greater dispersal abilities tend to serve as vectors for a greater diversity of pathogens than do those that feed on single hosts, that specialize on a single species (or related group of species), that feed on skin, feathers, hair, or dead tissue rather than blood, and those that have poor dispersal ability. The mouthparts of vectors directly affect the intimacy of their association with host tissues (e.g., blood, skin, mucous membranes, or contamination of host foods). Insect mouthparts have evolved from a primitive (but complicated) apparatus involving oral and pre-oral structures that facilitate manipulation of food items, ripping and tearing of organic matter, sometimes mastication, and movement of food into the pharynx (Black and Kondratieff 2004). Chewing mouthparts of the cockroaches and the grasshoppers provide well-recognized examples; anatomical structures include the labrum 158

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(a flap-like, dorsal, sensory lip), the mandibles (paired, jaw-like structures that lie beneath the labrum), maxillae (paired structures used to manipulate and chew food items before they are moved into the pharynx), and the hypopharynx (a tongue-like lobe that helps to mix salivary secretions with the chewed food bolus and then to suck the bolus into the pharynx). These basic structures have been modified in a myriad of ways, but we can group the modifications into at least eight additional categories, including: (1) a reduction of mouthparts to a vestigial state in adult bot flies (Order Diptera; Families Oestridae, Cuterebridae, Gastrophilidae, and Hypodermatidae); (2) chewing mouthparts of chewing lice; (3) piercing mouthparts of sucking lice (Order Phthiraptera; Suborder Anoplura); piercing mouthparts of true bugs (Order Hemiptera); (4) piercing mouthparts of fleas (Order Siphonaptera); (5) sponging mouthparts of some f lies (Order Diptera); (6) piercing/sucking mouthparts of some flies; (7) slashing mouthparts of horse flies and deer flies (Order Diptera; Family Tabanidae); and (8) the piercing, sucking mouthparts of butterflies and moths. In addition, tissue-feeding maggots (fly larvae) have a pair of external, anterior mouth hooks used as anchors and to scratch and tear tissue. Tissue removed by the mouth hooks is mixed with salivary secretions and pumped through the gut of the maggot (Black and Kondratieff 2004, Gullan and Cranston 2005). Each type of mouthparts allows a parasitic insect to obtain nourishment from hosts, but the different types of mouthparts, and ancillary tissues including salivary glands, result in different host–parasite interactions. Thus, the chewing mouthparts of amblycerans and ischnocerans allow the lice to feed on dead skin, feathers, and hair, but in most cases they do not allow direct access to the blood vascular system. As expected, chewing lice don’t transmit many blood-borne pathogens (Clayton et al. 2008, Durden and Lloyd 2009). The slashing mouthparts of deer flies and horse flies create painful bites and feeding episodes are often interrupted (Allan 2001a, Hall and Gerhardt

2009). As expected, these flies are very good at mechanically transmitting pathogens that don’t require long periods of feeding for transmission, such as F. tularensis, the causative agent of tularemia. Tabanid flies also serve as good biological vectors for parasites such as the arterial worm Elaeophora schneideri. The dispersal ability of different species of arthropods is determined largely by (1) the presence or absence of wings, (2) the ability to survive prolonged periods of time in the environment, and (3) the ability to use widely ranging hosts for long-distance dispersal (Harwood and James 1979). Many groups of parasitic arthropods, including lice, fleas, mites, and ticks, lack the ability to fly. These parasites typically move long distances on their hosts to colonize new populations. While these species may be good vectors, their dispersal ability is hampered by their lack of flight. Of course, many groups of insects have evolved wings. Within the Diptera, the tiny size and delicate wing structure of midges and gnats make long-distance dispersal unlikely. Other groups, including horse flies, blow flies, flesh flies, and tsetses, are very strong fliers and have correspondingly greater potential to disperse long distances. Strong-flying vectors have the ability to move long distances and move pathogens quickly among different populations. Some arthropods, such as the astigmatid mites, survive only for a relatively short time off their hosts, while others such as ticks and fleas survive for extended periods of time in the environment between meals. Non-volant species of vectors that remain on hosts during prolonged bouts of feeding, or after feeding, have the potential to move pathogens over greater distances than those that restrict their feeding to short periods in areas restricted to nests or burrows. Nest- and burrow-dwelling ectoparasites are known as nidicolous species; they typically require a narrow range of environmental conditions associated with the nests or burrows to which they are adapted, and they feed on a narrow range of hosts; nidicolous species tend to be specialists.

In contrast, ectoparasites with a more general palate are less likely to be associated with specific types of nests or burrows and are more likely to be exposed to a wide range of host species and their parasites (Durden and R. 2009, Nicholson et al. 2009). Clearly, the range of hosts to which parasites are adapted and associated varies between specialists and generalists, and affects their ability to transmit disease agents. As an example, the Ixodes ricinus group of ticks is infamous because it includes four generalist species (I. ricinus in Europe, I. persulcatus in northern Asia, I. pacificus in the western United States, and I. scapularis in the eastern and southern United States) that transmit a large number of pathogens among a wide range of species, including humans. In contrast, nidicolous specialists within the I. ricinus complex (including I. affinis, I. cookei, I. jellisoni, I. minor, I. muris, and I. spinipalpis) are known to transmit relatively few pathogens, and then among a relatively smaller number of host species (Nicholson et al. 2009). Specialists and generalists vary in their relative abilities to focus transmission within a reservoir community and to amplify infections beyond the reservoir. In the most extreme examples, such as sucking lice and some species of fleas and ticks, highly specialized ectoparasites feed mainly on the reservoir host species, and they can be very efficient vectors, causing high prevalence of disease among hosts. Examples of transmission by specialists include that of Buggy Creek virus transmitted by swallow bugs, and the heart worm Sarconema eurycerca, transmitted among swans and geese via the chewing louse intermediate host, Trinoton anserinum. In the case of Buggy Creek virus, nidicolous bugs feed mainly on swallows and martins, serving to focus transmission and as an over-winter reservoir of the virus at colony nesting sites (Hopla et al. 1993). In the case of this heart worm, the species of louse (Suborder Amblycera) is a relative specialist on geese and swans, and, unlike many chewing lice, bites the host and feeds directly on blood. The blood-feeding habit allows intimate association parasitic insects, mites, and ticks 159

between the birds’ blood and the lice in which the immature stages (microfilariae through the infective L 3) of the nematode develop, and the louse serves as an intermediate host in an enzootic transmission cycle (Seegar et al. 1976). Generalist feeders may also dilute the force of transmission by feeding on non-reservoir hosts (LoGiudice 2003); the prevalence of infection among the population of vectors may be diminished or remain low when non-reservoir hosts account for a great enough proportion of the blood meals of the vector population. Extreme cases of host dilution effects converge on a phenomenon that has been termed zooprophylaxis, in which proteins in the blood of specific species of hosts actually kill parasites within blood-feeding vectors (Matuschka et al. 1993, Lane and Quistad 1998, Kelly and Thompson 2000, Saul 2003). On the other hand, generalist vectors serve as bridge vectors, carrying pathogens to hosts beyond enzootic maintenance cycles, to amplification or deadend hosts. Bridging is exemplified by four related species of Ixodes ricinus–group ticks that carry pathogens from enzootic cycles (in which they may also serve as maintenance vectors) to a wide range of host species including humans (Brown et al. 2005, Nicholson et al. 2009). The number of times that a vector feeds on different hosts during its life cycle, and the timing of those feedings, affects vectorial capacity (or potential). Some species of flies do not feed as adults (including bot flies in the Family Oestridae), and the adults of these species are not known to transmit pathogens to hosts. In contrast, horse flies and deer flies are intermittent feeders and may fly between several successive hosts before obtaining a full blood meal (Allan 2001a, Hall and Gerhardt 2009). Such a feeding pattern coupled with the pain and blood flow associated with their bites, as discussed above, greatly facilitate their ability to serve as mechanical vectors. Likewise, the vector potential of different species of ticks is directly related to the number of hosts encountered during their life cycle. Ticks that feed on 160

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a single host [1-host ticks such as Rhipicephalus (Boophilus) spp. and D. albipictus] transmit relatively few microorganisms, whereas the 3-host ticks tend to be more efficient vectors and transmit most of the tick-borne disease agents (Allan 2001b, Nicholson et al. 2009). Thus, some groups of arthropods are notorious for transmitting disease agents, while other groups can be closely associated with infected hosts without efficiently transmitting pathogens. Vectorial capacity is a measure of an arthropod species’ ability to serve as a vector between individuals in a population or community of hosts. Vectorial capacity depends on many aspects of the host–vector–parasite interactions, including the temporal and spatial overlap of hosts and vectors, biological associations of arthropods and pathogens, and the biological adaptations of these arthropods facilitating their own propagation. Specifically, vectorial capacity is dependent on the vector:host ratio, the probability that the vector will feed on a given host species, the typical number of days between blood meals, the proportion of vectors that become infected from feeding on infective hosts, the daily survivorship of the vectors, the probability that the exposed vector will become infective to subsequent naïve hosts, and the number of days required for the vector to achieve such infectivity (Fine 1981, Black and Salman 2004). The ecology of vector-borne diseases also varies somewhat from directly transmitted diseases. Like hosts and parasites, vectors have their own niche requirements, and the incidence of vector-borne disease in wildlife populations tends to depend upon temporal and spatial overlap of hosts and vectors (Marquart et al. 2004). In temperate regions, the activity pattern of vectors tends to be seasonal, and such seasonality results in seasonal occurrence of vector-borne disease that coincides with the activity of the primary, or amplification, vectors. Thus, the incidence of West Nile fever in birds in the United States increases with mosquito activity during the spring and subsides with the cessation of mosquito activity

during autumn. Specific seasonal patterns are explained in more detail in chapters addressing bacteria, viruses, protozoa, and helminths. Clearly, a detailed understanding of the vector– host relationships and transmission ecology requires considerable information on hosts, pathogens, and vectors. Moreover, the ecological interactions that determine the variables listed are likely to vary among different communities of hosts, vectors, and pathogens, as well as through time.

Literature Cited Acha, P. N., and B. Szyfres. 2001. Zoonoses and communicable diseases common to man and animals. Vol. I: Bacterioses and mycoses. 3rd ed. Report No. 580. Pan American Health Organization, Washington, DC. Acha, P. N., and B. Szyfres. 2003a. Zoonoses and communicable diseases common to man and animals. Vol. 2: Chlamydioses, rickettsioses, and viroses. 3rd ed. Report No. 580. Pan American Health Organization, Washington, DC. Acha, P. N., and B. Szyfres. 2003b. Zoonoses and communicable diseases common to man and animals. Vol. 3: Parasitoses. 3rd ed. PReport No. 580. Pan American Health Organization, Washington, DC. Adler, P. H., and J. W. McCreadie. 2009. Black flies (Simuliidae). Pp. 189–206 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Allan, S. A. 2001a. Biting flies (Class Insecta: Order Diptera). Pp. 18–45 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Allan, S. A. 2001b. Ticks (Class Arachnida: Order Acarina). Pp. 72–106 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Anderson, J. R. 1975. The behavior of nose bot flies (Cephenemyia apicata and C. jellisoni) when attacking black-tailed deer (Odocoileus hemionus columbianus) and the resulting reactions of the deer. Canadian Journal of Zoology 53:977–992. Anderson, J. R. 2001. Larviposition by nasopharyngeal bot fly parasites of Columbian black-tailed deer: A correction. Medical and Veterinary Entomology 15:438–442.



Anderson, J. R., A. C. Nilssen, and I. Folstad. 1994. Mating behavior and thermoregulation of the reindeer warble fly, Hypoderma tarandi L. (Diptera: Oestridae). Journal of Insect Behavior 7:679–706. Beesley, W. N. 1968. Observations on the biology of the ox warble-fly (Hypoderma: Diptera, Oestridae). II: Bacteriostatic properties of larval extracts. Annals of Tropical Medicine and Parasitology 62:8–12. Bender, L. C., and P. B. Hall. 2004. Winter fawn survival in black-tailed deer populations affected by hair loss syndrome. Journal of Wildlife Diseases 40:444–451. Bennett, G. F. 1957. Studies on the genus Proto­ calliphora (Diptera: Calliphoridae). Ph.D. diss­ertation, University of Toronto, Toronto, Ontario, Canada. Bildfell, R. J., J. W. Mertins, J. A. Mortenson, and D. F. Cottam. 2004. Hair-loss syndrome in blacktailed deer of the Pacific Northwest. Journal of Wildlife Diseases 40:670–681. Black, W. C., IV, and B. C. Kondratieff. 2004. Evolution of arthropod disease vectors. Pp. 9–23 in W. C. Marquart, B. C. Kondratieff, C. G. Moore, J. E. Freier, H. H. Hagedorn, W. C. Black, A. A. James, J. Hemingway, and S. Higgs (editors), The biology of disease vectors. Elsevier Academic Press, Burlington, MA. Black, W. C., IV, and D. Salman. 2004. Molecular techniques for epidemiology and the evolution of arthropod-borne pathogens. Pp. 227–255 in W. C. Marquart, B. C. Kondratieff, C. G. Moore, J. E. Freier, H. H. Hagedorn, W. C. Black, A. A. James, J. Hemingway, and S. Higgs (editors), The biology of disease vectors. Elsevier Academic Press, Burlington, MA. Bornstein, S., T. Mörner, and W. M. Samuel. 2001. Sarcoptes scabei and sarcoptic mange. Pp. 107–119 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Botzler, R. G., J. Albrecht, and T. Schaefer. 1980. Tick paralysis in a western harvest mouse (Reithrodontomys megalotis). Journal of Wildlife Diseases 16:223–224. Boyce, W. M., and R. N. Brown. 1991. Antigenic characterization of Psoroptes spp. (Acari: Psoroptidae) mites from different hosts. Journal of Parasitology 77:675–679. Brown, C. R., N. Komar, S. B. Quick, R. A. Sethi, N. A. Panella, M. B. Brown, and M. Pfeffer. 2001. Arbovirus infection increases with group size. Proceedings of the Royal Society of London B: Biological Sciences 268:1833–1840.

parasitic insects, mites, and ticks 161

Brown, R. N., R. S. Lane, and D. T. Dennis. 2005. Geographic distributions of tick-borne diseases and their vectors. Pp. 363–391 in J. Goodman, D. T. Dennis, and D. Sonenshine (editors), Tickborne diseases of humans. ASM Press, Washington, DC. Brunetti, O., and H. Cribbs. 1971. California deer deaths due to massive infestation by the louse (Linognathus africanus). California Fish and Game 57:162–166. Campbell, J. B., S. R. Skoda, D. R. Berkebile, D. J. Boxler, G. D. Thomas, D. C. Adams, and R. Davis. 2001. Effects of stable flies (Diptera: Muscidae) on weight gains of grazing yearling cattle. Journal of Economic Entomology 94:780–783. Capinera, J. L. 2010. Insects and wildlife: Arthropods and their relationships with wild vertebrate animals. Wiley-Blackwell, Hoboken, NJ. Chapman, B. R., and J. E. George. 1991. The effects of ectoparasites on cliff swallow growth and survival. Pp. 69–92 in J. E. Loye and M. Zuk (editors), Bird-parasite interactions: Ecology, evolution, and behaviour. Oxford University Press, New York. Chilton, G., M. J. Vonhof, B. V. Peterson, and N. Wilson. 2000. Ectoparasitic insects of bats in British Columbia, Canada. Journal of Parasitology 86:191–192. Cilek, J. E., and C. F. Hallmon. 2005. The effectiveness of the mosquito magnet trap for reducing biting midge (Diptera: Ceratopogonidae) populations in coastal residential backyards. Journal of the American Mosquito Control Association 21:218–221. Clayton, D. H., R. J. Adams, and S. E. Bush. 2008. Phthiraptera, the chewing lice. Pp. 515–526 in C. T. Atkinson, N. J. Thomas, and D. B. Hunter (editors), Parasitic diseases of wild birds. Wiley-Blackwell, Ames, IA. Clayton, D. H., and K. P. Johnson. 2003. Linking coevolutionary history to ecological process: Doves and lice. Evolution 57:2335–2341. Cogley, T. P. 1991. Warble development by the rodent bot Cuterebra fontinella (Diptera: Cuterebridae) in the deer mouse. Veterinary Parasitology 38:275–288. Cogley, T. P., and J. R. Anderson. 1981. Invasion of black-tailed deer by nose bot fly larvae (Diptera: Oestridae: Oestrinae). International Journal for Parasitology 11:281–286. Cohen, S., M. T. Greenwood, and J. A. Fowler. 1991. The louse Trinoton anserinum (Amblycera: Phthiraptera), an intermediate host of Sarconema eurycerca (Filarioidea: Nematoda), a heartworm of swans. Med Vet Entomol 5:101–110.

162

parasitic insects, mites, and ticks

Colwell, D. D. 2001. Bot flies and warble flies (Order Diptera: Family Oestridae). Pp. 46–71 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Cornish, T. E., M. J. Linders, S. E. Little, and W. M. Vander Haegen. 2001. Notoedric mange in western gray squirrels from Washington. Journal of Wildlife Diseases 37:630–633. Craine, I. T., and R. Boonstra. 1986. Myiasis in Wohlfahrtia vigil in nestling Microtus pennsylvanicus. Journal of Wildlife Diseases 22:587–589. Crause, J. C., S. van Wyngaardt, R. Gothe, and A. W. Neitz. 1994. A shared epitope found in the major paralysis inducing tick species of Africa. Experimental and Applied Acarology 18:51–59. Downes, C. M., J. B. Theberge, and S. M. Smith. 1986. The influence of insects on the distribution, microhabitat choice, and behavior of the Burwash caribou herd. Canadian Journal of Zoology 64:622–629. Durden, L. 2004. Lice, the Phthiraptera. Pp. 67–76 in W. C. Marquart, B. C. Kondratieff, C. G. Moore, J. E. Freier, H. H. Hagedorn, W. C. Black, A. A. James, J. Hemingway, and S. Higgs (editors), The biology of disease vectors. Elsevier Academic Press, Burlington, MA. Durden, L. A. 2001. Lice (Phthiraptera). Pp. 3–17 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Durden, L. A., and N. C. Hinkle. 2009. Fleas (Siphonaptera). Pp. 115–135 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Durden, L. A., and J. E. Lloyd. 2009. Lice (Phthiraptera). Pp. 59–82 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Durden, L. A., and M. G. R. 2009. Introduction. Pp. 1–12 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Durden, L. A., and N. Wilson. 1991. Parasitic and phoretic arthropods of sylvatic and commensal white-footed mice (Peromyscus leucopus) in central Tennessee, with notes on Lyme disease. Journal of Parasitology 77:219–223. Fahrenholz, H. 1913. Ectoparasiten und Abstammungslehre. Zoologische Anzeiger 41:371–374. Fine, P. E. M. 1981. Epidemiological principles of vector mediated transmission. Pp. 77–91 in J. J. McKelvey, B. F. Eldridge, and K. Maramorosch (editors), Vectors of disease agents. Praeger Scientific, New York.

Foreyt, W. J., B. Hall, and L. Bender. 2004. Evaluation of ivermectin for treatment of hair loss syndrome in black-tailed deer. Journal of Wildlife Diseases 40:434–443. Foreyt, W. J., D. H. Rice, and K. C. Kim. 1986. Pediculosis of mule deer and white-tailed deer fawns in captivity. Journal of the American Veterinary Medical Association 189:1172–1173. Foster, W. A., and E. D. Walker. 2009. Mosquitoes (Culicidae). Pp. 207–259 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Fritsche, T. R. 1999. Arthropods of medical importance. Pp. 1449–1466 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. American Society for Microbiology, Washington, DC. Gage, K. L. 2004. Fleas, the Siphonaptera. Pp. 77–92 in W. C. Marquart, B. C. Kondratieff, C. G. Moore, J. E. Freier, H. H. Hagedorn, W. C. Black, A. A. James, J. Hemingway, and S. Higgs (editors), The biology of disease vectors. Elsevier Academic Press, Burlington, MA. Garris, G. I., J. B. Prullage, J. L. Prullage, F. C. Wright, and J. A. Miller. 1991. Control of Psoroptes cuniculi in captive white-tailed deer with ivermectin-treated corn. Journal of Wildlife Diseases 27:254–257. Glines, M. V., and W. M. Samuel. 1989. Effect of Dermacentor albipictus (Acari:Ixodidae) on blood composition, weight gain and hair coat of moose, Alces alces. Experimental and Applied Acarology 6:197–213. Gullan, P. J., and P. S. Cranston. 2005. The insects: An outline of entomology. 3rd ed. Blackwell Publishing, Ltd., Malden, MA. Hagemoen, R. I. M., and E. Reimers. 2002. Reindeer summer activity pattern in relation to weather and insect harassment. Journal of Animal Ecology 71:883–892. Haldane, J. B. S. 1932. Possible worlds: And other essays. Chatto and Windus, London, UK. Hall, R. D., and R. R. Gerhardt. 2009. Flies (Diptera). Pp. 137–152 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Halos, L., T. Jamal, Maillard, B. Girard, J. Guillot, B. Chomel, M. Vayssier-Taussat, and H.-J. Boulouis. 2004. Role of Hippoboscidae flies as potential vectors of Bartonella spp. infecting wild and domestic ruminants. Applied Environmental Microbiology 70:6302–6305. Harwood, R. F., and M. T. James. 1979. Entomology in human and animal health. 7th ed. Macmillan Publishing Company, New York.



Hoberg, E. P., D. R. Brooks, and D. Siegel-Causey. 1997. Host–parasite co-speciation: History, principles, and prospects. Pp. 212–235 in D. H. Clayton, and J. Moore (editors), Host– parasite evolution: General principles and avian models. Oxford University Press, Oxford, UK. Hoogstraal, H. 1985. Argasids and nuttallielid ticks as parasites and vectors. Advances in Parasitology 24:135–238. Hopla, C. E., D. B. Francy, C. H. Calisher, and J. S. Lazuick. 1993. Relationship of cliff swallows, ectoparasites, and an Alphavirus in west-central Oklahoma. Journal of Medical Entomology 30:267–272. Jongejan, F., and G. Uilenberg. 2004. The global importance of ticks. Parasitology 129(Suppl.):S3–14. Jonsson, N. N., and D. G. Mayer. 1999. Estimation of the effects of buffalo fly (Haematobia irritans exigua) on the milk production of dairy cattle based on a meta-analysis of literature data. Medical and Veterinary Entomology 13:372–376. Jordaan, L. C., and H. van Ark. 1990. A survey of annoyance of livestock by Simulium chutteri Lewis along the Orange River, South Africa (Diptera: Simuliidae). Onderstepoort Journal of Veterinary Research 57:189–195. Jovani, R., J. L. Tella, D. Sol, and D. Ventura. 2001. Are hippoboscid flies a major mode of transmission of feather mites? Journal of Parasitology 87 1187–1189 Karter, A. J., and I. Folstad. 1989. Defence behaviour of reindeer in response to flying parasitic Diptera. Rangifer 9:14–16. Karter, A. J., I. Folstad, and J. R. Anderson. 1992. Abiotic factors influencing embryonic development, egg hatching, and larval orientation in the reindeer warble fly, Hypoderma tarandi. Medical and Veterinary Entomology 6:355–362. Kelly, D. W., and C. E. Thompson. 2000. Epidemiology and optimal foraging: Modelling the ideal free distribution of insect vectors. Parasitology 120:319–327. Krinsky, W. L. 2009a. True bugs (Hemiptera). Pp. 83–99 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Krinsky, W. L. 2009b. Tsetse flies (Glossinidae). Pp. 297–308 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Lane, R. S., and G. B. Quistad. 1998. Borreliacidal factor in the blood of the western fence lizard (Sceloporus occidentalis). Journal of Parasitology 84:29–34.

parasitic insects, mites, and ticks 163

Lange, R. E., A. V. Sandoval, and W. P. Meleney. 1980. Psoroptic scabies in bignorn sheep (Ovis canadensis mexicana) in New Mexico. Journal of Wildlife Diseases 16:77–82. Little, S. E., W. R. Davidson, E. W. Howerth, P. M. Rakich, and V. F. Nettles. 1998. Diseases diagnosed in red foxes from the southeastern United States. Journal of Wildlife Diseases 34:620–624. Lloyd, J. E. 2009. Louse flies, keds, and related flies (Hippoboscoidea). Pp. 339–352 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/ Elsevier, San Diego, CA. LoGiudice, K. 2003. Trophically transmitted parasites and the conservation of small populations: Raccoon roundworm and the imperiled Allegheny woodrat. Conservation Biology 17:258–266. Loye, J. E., and M. Zuk (editors). 1991. Bird-parasite interactions. Oxford University Press, Oxford, UK. Luttrell, M. P. 1997. Avian tick paralysis. In W. R. Davidson, and V. F. Nettles (editors), Field manual of wildlife diseases in the southeastern United States. Southeastern Cooperative Wildlife Disease Study, College of Veterinary Medicine, The University of Georgia, Athens, GA. Madslien, K., B. Ytrehus, T. Vikoren, J. Malmsten, K. Isaksen, H. O. Hygen, and E. J. Solberg. 2011. Hair-loss epizootic in moose (Alces alces) associated with massive deer ked (Lipoptena cervi) infestation. Journal of Wildlife Diseases 47:893–906. Mans, B. J., A. I. Louw, and A. W. Neitz. 2003. The major tick salivary gland proteins and toxins from the soft tick, Ornithodoros savignyi, are part of the tick Lipocalin family: Implications for the origins of tick toxicoses. Molecular Biology and Evolution 20:1158–1167. Marquart, W. C., B. C. Kondratieff, C. G. Moore, J. E. Freier, H. H. Hagedorn, W. C. Black, A. A. James, J. Hemingway, and S. Higgs (editors). 2004. The biology of disease vectors. Elsevier Academic Press, Burlington, MA. Matuschka, F. R., M. Heiler, H. Eiffert, P. Fischer, H. Lotter, and A. Spielman. 1993. Diversionary role of hoofed game in the transmission of Lyme disease spirochetes. American Journal of Tropical Medicine and Hygiene 48:693–699. Mazet, J. A., W. M. Boyce, J. Mellies, I. A. Gardner, R. K. Clark, and D. A. Jessup. 1992. Exposure to Psoroptes sp. mites is common among bighorn sheep (Ovis canadensis) populations in California. Journal of Wildlife Diseases 28:542–547. Meintjes, T., L. J. Fourie, and I. G. Horak. 2002. Onhost ecology and off-host survival of the sheep scab mite Psoroptes ovis. Onderstepoort Journal of Veterinary Research 69:273–283.

164

parasitic insects, mites, and ticks

Mertins, J. W., J. A. Mortenson, J. A. Bernatowicz, and P. B. Hall. 2011. Bovicola tibialis (Phthiraptera: Trichodectidae): Occurrence of an exotic chewing louse on cervids in North America. Journal of Medical Entomology 48:1–12. Moerschel, F. M., and D. R. Klein. 1997. Effects of weather and parasitic insects on behavior and group dynamics of caribou of the Delta Herd, Alaska. Canadian Journal of Zoology 75:1659–1670. Moon, R. D. 2009. Muscid flies (Muscidae). Pp. 275–295 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Morse, S. F., K. J. Olival, M. Kosoy, S. Billeter, B. D. Patterson, C. W. Dick, and K. Dittmar. 2012. Global distribution and genetic diversity of Bartonella in bat flies (Hippoboscoidea, Streblidae, Nycteribiidae). Infection, Genetics and Evolution 12:1717–1723. Mullen, G. R. 2009. Biting midges (Ceratopogonidae). Pp. 169–188 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Mullen, G. R., and L. A. Durden (editors). 2009. Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Mullen, G. R., and B. M. O’Connor. 2009. Mites (Acari). Pp. 433–492 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Nicholson, W. L., D. E. Sonenshine, R. S. Lane, and G. Uilenberg. 2009. Ticks (Ixodida). Pp. 493–542 in G. Mullen and L. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Nilssen, A. C., and J. R. Anderson. 1995. Flight capacity of the reindeer warble fly, Hypoderma tarandi (L.), and the reindeer nose bot fly, Cephenemyia trompe (Modeer) (Diptera: Oestridae). Canadian Journal of Zoology 73:1228–1238. Ninomiya, H., M. Ogata, and T. Makino. 2003. Notoedric mange in free-ranging masked palm civets (Paguma larvata) in Japan. Veterinary Dermatology 14:339–344. Norrix, L. W., D. W. DeYoung, P. R. Krausman, R. C. Etchberger, and T. J. Glattke. 1995. Conductive hearing loss in bighorn sheep. Journal of Wildlife Diseases 31:223–227. Oksanen, A., and M. Nieminen. 1998. Moxidectin as an endectocide in reindeer. Acta Veterinaria Scandinavica 39:483–489. Oksanen, A., M. Nieminen, and T. Soveri. 1993. A comparison of topical, subcutaneous and

oral administrations of ivermectin to reindeer. Veterinary Research 133:312–314. Pence, D. B., F. D. Matthews, 3rd, and L. A. Windberg. 1982. Notoedric mange in the bobcat, Felis rufus, from south Texas. Journal of Wildlife Diseases 18:47–50. Pence, D. B., M. E. Tewes, D. B. Shindle, and D. M. Dunn. 1995. Notoedric mange in an ocelot (Felis pardalis) from southern Texas. Journal of Wildlife Diseases 31:558–561. Pence, D. B., and E. Ueckermann. 2002. Sarcoptic manage in wildlife. Review of Science and Technology 21:385–398. Proctor, H. C., and D. N. Jones. 2004. Geographical structuring of feather mite assemblages from the Australian brush-turkey (Aves: Megapodiidae). Journal of Parasitol 90:60–66. Ramey, R. R., S. T. Kelley, W. M. Boyce, and B. D. Farrell. 2000. Phylogeny and host specificity of Psoroptic mange mites (Acarina: Psoroptidae) as indicated by ITS sequence data. Journal of Medical Entomology 37:791–796. Reisen, W. K. 2009. Epidemiology of vector-borne diseases. Pp. 19–33 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Reist, M., T. D. Medjitna, U. Braun, and K. Pfister. 2002. Effect of a treatment with eprinomectin or trichlorfon on the yield and quality of milk produced by multiparous dairy cows. Veterinary Record 151:377–380. Rogers, A. R., R. J. Raleigh, J. Robertson, and B. J. Stutchbury. 1991. Patterns and effects of parasitism by Protocalliphora siala on tree swallow nestlings. Pp. 123–139 in J. E. Loye and M. Zuk (editors), Bird-parasite interactions: Ecology, evolution, and behaviour. Oxford University Press, New York. Roller, E. A. I., V. F. Nettles, W. R. Davidson, and R. R. Gerrish. 1978. Otitis media caused by Psoroptes cunniculi in white-tailed deer. Journal of the American Medican Association 173:1242–1243. Rutledge, L. C., and R. K. Gupta. 2009. Moth flies and sand flies (Psychodidae). Pp. 153–168 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Ryser-Degiorgis, M. P., A. Ryser, L. N. Bacciarini, C. Angst, B. Gottstein, M. Janovsky, and U. Breitenmoser. 2002. Notoedric and sarcoptic mange in free-ranging lynx from Switzerland. Journal of Wildlife Diseases 38:228–232. Samuel, W. M. 2004. White as a ghost: Winter ticks and moose. Vol. 1. Federation of Alberta Naturalists, Edmonton, Alberta, Canada.



Samuel, W. M., D. A. Welch, and B. L. Smith. 1991. Ectoparasites from elk (Cervus elaphus nelsoni) from Wyoming. Journal of Wildlife Diseases 27:446–451. Saul, A. 2003. Zooprophylaxis or zoopotentiation: The outcome of introducing animals on vector transmission is highly dependent on the mosquito mortality while searching. Malaria Journal 2:32. Scholl, P. J., E. P. Catts, and G. R. Mullen. 2009. Myiasis (Muscoidea, Oestridae). Pp. 309–338 in G. R. Mullen and L. A. Durden (editors), Medical and veterinary entomology. 2nd ed. Academic Press/Elsevier, San Diego, CA. Seegar, W. S., E. L. Schiller, W. J. Sladen, and M. Trpis. 1976. A mallophaga, Trinoton anserinum, as a cyclodevelopmental vector for a heartworm parasite of waterfowl. Science 194:739–741. Sherman, R. A., M. J. R. Hall, and S. Thomas. 2000. Medicinal maggots: An ancient remedy for some contemporary afflictions. Annual Review of Entomology 45:55–81. Singer, R. S., D. A. Jessup, I. A. Gardner, and W. M. Boyce. 1997. Pathogen exposure patterns among sympatric populations of bighorn sheep, mule deer and cattle. Journal of Wildlife Diseases 33:377–382. Stone, B. F., K. C. Binnington, M. Gauci, and J. H. Aylward. 1989. Tick/host interactions for Ixodes holocyclus: Role, effects, biosynthesis and nature of its toxic and allergenic oral secretions. Experimental and Applied Acarology 7:59–69. Stork, M. G. 1979. The epidemiological and economic importance of fly infestation of meat and milk producing animals in Europe. Veterinary Record 105:341–343. Strickland, R. K., R. R. Garrish, and J. S. Smith. 1981. Arthropods. Pp. 363–389 in W. R. Davidson, F. A. Hayes, V. F. Nettles, and F. E. Kellogg (editors), Diseases and parasites of white-tailed deer. Tall Timber Research Station, Tallahassee, FL. ter Hofstede, H. M., and M. B. Fenton. 2005. Relationships between roost preferences, ectoparasite density, and grooming behaviour of neotropical bats. Journal of Zoology, London 266:333–340. ter Hofstede, H. M., M. B. Fenton, and J. Whitaker, J. O. . 2004. Host and host-site specificity of batflies (Diptera: Streblidae and Nycteribiidae) on Neotropical bats (Chiroptera). Canadian Journal of Zoology 82:616–626. Valenzuela, D., G. Ceballos, and A. Garcia. 2000. Mange epizootic in white-nosed coatis in western Mexico. Journal of Wildlife Diseases 36:56–63. Welsh, G. W., and T. D. Bunch. 1983. Psoroptic scabies in desert bighorn sheep (Ovis canadensis

parasitic insects, mites, and ticks 165

nelsoni) from northwestern Arizona. Journal of Wildlife Diseases 19:342–344. Westrom, D. R., B. C. Nelson, and G. E. Connolly. 1976. Transfer of Bovicola tibialis (Piaget) Mallophaga: Trichodectidae) from the introduced fallow deer to the Columbian black-tailed deer in California. Journal of Medical Entomology 13:169–173. Whitworth, T. L., and G. F. Bennett. 1992. Pathogenicity of larval Protocalliphora (Diptera: Calliphoridae) parasitizing nestling birds. Canadian Journal of Zoology 70:2184–2191. Wieman, G. A., J. B. Campbell, J. A. Deshazer, and I. L. Berry. 1992. Effects of stable flies (Diptera: Muscidae) and heat stress on weight gain and feed efficiency of feeder cattle. Journal of Economic Entomology 85:1835–1842. Wobeser, G. A. 1994. Investigation and management of disease in wild animals. Plenum Press, New York. Wright, F. C., and R. L. Glaze. 1988. Blackbuck antelope (Antilope cervicapra), a new host for Psoroptes

166

parasitic insects, mites, and ticks

cuniculi (Acari: Psoroptidae). Journal of Wildlife Diseases 24:168–169. Yeruham, I., S. Rosen, A. Hadani, and Y. Braverman. 1999. Arthropod parasites of Nubian ibexes (Capra ibex nubiana) and gazelles (Gazella gazella) in Israel. Veterinary Parasitology 83:167–173. Zahler, M., A. Essig, R. Gothe, and H. Rinder. 1999. Molecular analyses suggest monospecificity of the genus Sarcoptes (Acari: Sarcoptidae). International Journal for Parasitology 29:759–766. Zahler, M., W. M. Hendrikx, A. Essig, H. Rinder, and R. Gothe. 2000. Species of the genus Psoroptes (Acari: Psoroptidae): A taxonomic consideration. Experimental and Applied Acarology 24:213–225. Ziccardi, M. H., W. M. Boyce, and I. A. Gardner. 1996. Seroprevalence of Psoroptes sp. mites in free-ranging elk (Cervus elaphus) as determined by kinetic ELISA. Journal of Wildlife Diseases 32:51–56.

SIX

Kingdom Protista

CONTENTS Introduction to Protozoa and Other Protista

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Life Cycle Strategies among the Protista

168

Cyst Stage Present 182 Eimeria spp.182 Indirect (Heteroxenous) Life Cycles 184 Use Vertebrates as Both Intermediate and Definitive Hosts 184 Toxoplasma gondii184 Arthropod-borne Apicomplexa 188 Plasmodium relictum (Avian Malaria) 188 Trypanosoma spp. (Nagana) 189 Helminth-borne191 Histomonas meleagridis191 Opportunistic Soil and Water Organisms 193 Acanthamoeba spp. 193

Major Groups Considered 169 Amebae169 Apicomplexa (Sporozoa) 170 Coccidia172 Malaria Parasites (Haemosporidia) 173 Piroplasms175 Ciliophora (Ciliates) 176 Excavata (Flagellates) 177 Intestinal Flagellates 177 Hemoflagellates178 Life History Types among Protozoa 180 Direct (Monoxenous) Life Cycles 180 No Cyst Stage 180 Trichomonas gallinae180

Literature Cited

1999). At least 45,000 species have been described, many of which are parasitic (Roberts and Janovy 2000). The terms “protozoa” and “protista” have undergone considerable revision in definitions and still are in a state of flux (Cox 1991, Patterson 1999, Adl et al. 2005).

Introduction to Protozoa Protozoa and other protists evolved from the first eukaryotes (Patterson 1999) and probably have been present at least twice as long as any of the major multicellular organisms (Patterson

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167

Protozoa historically were defined as a phylum of unicellular, heterotrophic eukaryotes (Kudo 1966, Patterson 2000), with their classification based primarily on their organelles of locomotion and food-acquiring structures; major groups included flagellates, amebae, ciliates, and non-motile spore-forming protozoa (Cox 1991, Sleigh 1991). More recently, based on the complement and organization of organelles within the organisms, use of 16S ribosomal-RNA sequencing data, and protein and gene sequences, it was recognized that many groups previously classified among the protozoa were polyphyletic and probably only distantly related (Patterson 2000). There has been a gradual move to replace the notion of “protozoa” with the broader concept of “protista,” recognizing that even the protista continue to contain some polyphyletic groups of eukaryotes. There still is no agreement on how various groups are related or how they should be classified. In one substantial revision, about 71 monophyletic lineages, within a set of four major lineages, were identified among the Protista, based on common patterns of cell organization (ultrastructural identity) (Patterson 1999). In a more recent revision, there has been a continued focus on emphasizing evolutionary relationships; the numbers of paraphyletic groups are further decreased from earlier studies (Adl et al. 2005). In Adl et al.’s (2005) taxonomic scheme, both unicellular and multicellular eukaryotic organisms are included, organized into six supergroups based on molecular phylogenies: Amoebozoa, Opisthokonta, Rhizaria, Archaeplastida, Chromalveolata, and Excavata (App. 1). Three of these supergroups (Amoebozoa, Chromoalveolata, Excavata) include single-celled eukaryotic organisms traditionally classified as protozoa (App. 1: Table 5). While still hierarchical, the proposed system lacks formal rank designations such as “class,” “order,” and so on; this was done to simplify the impacts of any future changes within the system (Adl et al. 2005). 168

kingdom protista

For purposes of this classification, eukaryotes are defined as organisms having a distinct nucleus bounded by a double membrane, the outer being derived from the endomembrane network, with the nuclear pore complex traversing both membranes, and with one or more linear chromosomes typically packaged by histones and usually with a centromere and telomeres (Adl et al. 2005). Protozoa are predominantly non-filamentous heterotrophic species. The term protist is used to describe eukaryotes with a unicellular level of organization and without cell differentiation into tissues (Adl et al. 2005). For this discussion, the more traditional term, “protozoa,” generally will be retained. While some protozoa are simple, others are among the most complex cells known; all of the biological and biochemical mechanisms needed for a complex lifestyle are contained within a single cell (Sleigh 1991). Reproduction may be either asexual or sexual, and often is complex. Many species alternate sexual and asexual modes in their life cycles, or may vary the pattern in relation to environmental conditions (Roberts and Janovy 2000). Asexual reproduction usually is by binary fission, but also may involve several forms of budding or multiple fission; with multiple fission there is repeated division of the nucleus and other essential organelles before cytokinesis (Roberts and Janovy 2000). Sexual reproduction involves meiosis and the process of gametogony. Many protozoa can encyst by secreting a thick resinous case around themselves and becoming dormant (Keeton and Gould 1993). For life cycles, the term cyst is restricted to a vegetative quiescent stage, while the term spore is restricted to a reproductive stage (Adl et al. 2005).

Life cycle strategies among protozoa The life cycles of some parasitic protozoa are direct (monoxenous), involving transmission from one definitive host to the next. For most monoxenous parasites, a protective cyst stage is

used to enhance parasite survival in the environment between active infections; the intestinal flagellate Trichomonas gallinae is an exception, having no cyst stage. Many parasitic protozoa, however, have an indirect (heteroxenous) life cycle, involving a definitive host and at least one intermediate host. Most heteroxenous protozoa use hematophagous arthropods. Examples include the malaria parasites, piroplasms, and the trypanosomes. In other cases, the protozoa require an intermediate vertebrate host, or may use an optional (paratenic) host. In a few cases, protozoa rely on transmission through helminth life cycles, where the protozoa are incorporated with the helminth ova and transmitted to the susceptible host with the helminth infection. In humans, the flagellate Dientamoeba fragilis is transmitted by ova of the pinworm Enterobius vermicularis (Ockert and Schmidt 1976, BonDurant and Wakenell 1994) and possibly Ascaris lumbricoides ova (Sukanahaketu 1977). In gallinaceous birds, the intestinal flagellate Histomonas meleagridis is transmitted by ova of the cecal nematode Heterakis gallinarum (Cole and Friend 1999). In addition, there are a few free-living soil and water protozoa, primarily amebae, that are opportunistic parasites; these agents generally are not transmitted directly between susceptible hosts, but involve common-source infections derived from contaminated environments (Visvesvara and Stehr-Green 1990). Examples include species of Naegleria, Acanthamoeba, Balamuthia, and Hartmannella (Ma et al. 1990, Martinez and Visvesvara 1997, John 2001). Control strategies typically are directed at perceived weak points in the parasites’ life cycles. For arthropod-borne or helminth-borne parasites, this may involve habitat manipulation or the use of chemicals. For monoxenous parasites, or those using vertebrates for intermediate hosts, breaking the chain of infection by treating infected individuals or preventing infection by habitat manipulation also is common. Examples are addressed with specific parasites.

MAJOR GROUPS CONSIDERED Amebae Amebae are protozoa generally characterized as using pseudopodia as organelles of motility (Sleigh 1991). Most amebae now are included in the Supergroup Amoebozoa; a few other species having an ameba-like stage (e.g., Dientamoeba, Naegleria) are classified in the Supergroup Excavata (Adl et al. 2005) (App. 1: Table 5). A trophozoite is any protozoan in an asexual phase of development; among amebae the trophozoite is the active feeding stage. For intestinal amebae, the trophozoite will form a cyst stage that is well suited to withstand environmental conditions outside of the host, prior to being passed out with feces. Amoebozoa are characterized by ameboid locomotion generally, with non-eruptive, morphologically variable pseudopodia (lobopodia). The cells are “naked” or testate; cells usually are uninucleate, rarely binucleate or multinucleate. Cysts are common, but can be morphologically variable among species (Adl et al. 2005). Within the amebae, there are several groups that can cause diseases among vertebrates; of these, the genus Entamoeba is the most important. Entamoeba spp. lack mitochondria, have a simple endomembrane system, and undergo mitosis that is closed with an internal spindle (Sleigh 1991, Adl et al. 2005). Trophozoites have one nucleus. Cyst stages typically have four nuclei (Fig. 6.1), although immature cysts may have only one of two nuclei (Leber and Novak 1999). Most parasitic amebae inhabit the large intestine; cyst formation is a chemicaldependent process in the colon (Eichinger 2001). Entamoeba histolytica also has been reported from nonhuman primates and occasionally canines (Kocan 2001a). This parasite can be found worldwide but is more prevalent in tropical and subtropical regions (Leber and Novak 1999). Humans are considered the primary reservoir for Entamoeba histolytica (Leber and Novak 1999), and this parasite is the third most common cause of parasite-induced deaths among humans (Martinez-Paloma 1993). kingdom protista

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figure 6.1  Entamoeba spp. multinuclear cyst.

Entamoeba invadens is an important parasite of reptiles (Frank 1976, Lucius and Loos-Frank 1997); it infects carnivorous reptiles such as snakes, where it causes disease, and herbivorous reptiles such as turtles, in which it lives as a commensal (Ratcliffe and Geiman 1938, Meerovitch 1958, Jakob and Wesemeier 1995). Entamoeba spp. also have been reported in waterfowl (Quortrup and Shillinger 1941). Opportunistic, free-living amebae that can act as opportunistic parasites include Balamuthia spp. and Acanthamoeba spp.; members of both genera have a cyst stage. Acanthamoeba spp. and Balamuthia spp. inhabit the neurological tissues of infected hosts (Martinez and Visvesvara 1997, John 2001). No animal reservoir for these potentially pathogenic free-living amebae has been identified (John 2001). Interestingly, Acanthamoeba castellanii may play a role in the maintenance of pathogenic bacteria and fungi in the environment (Essig et al. 1997, Abd et al. 2003, Steenbergen et al. 2004). The life cycle of the ameba within the Amoebozoa is simple and direct. Division among ameba trophozoites is by simple fission. In cyst formation, trophozoites typically empty their food vacuoles, round up, get a little smaller, and surround themselves with a thin, delicate wall that is relatively sensitive to 170

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desiccation but fairly resistant to many chemicals. Some nuclear multiplication typically occurs in the phase, and on infecting a new host, the cyst opens (excystment) in the small intestine before the resulting amebae migrate down to the large intestine. Some cytoplasm is associated with each nucleus in the cyst after excystment.

Apicomplexa (Sporozoa) Apicomplexa are single-celled protists named for an apical complex of organelles in the infective stages (sporozoites, merozoites) that is unique among living organisms (Atkinson and van Riper 1991, Sleigh 1991). Historically called Sporozoa (“spore animals”), these protists currently are classified in the Supergroup Chromalveolata (App. 1: Table 5) (Adl et al. 2005). Apicomplexa produce spores at some time during their life cycle. While Apicomplexa generally are characterized by the absence of obvious external motility organelles (Bush et al. 2001), locomotion can occur by gliding, body f lexion, longitudinal ridges, and/or f lagella in some stages (Adl et al. 2005). Virtually all forms are parasitic (Adl et al. 2005). The life cycles of Apicomplexa are complex and include both direct (monoxenous) and

Sporozoite

Y N

Sporocyst O

M E

Trophozoite R

G

O

O

G

R

Oocyst

O

O

N

P S Zygote

GAMETOGONY Gametes

Gametocytes

(Micro & Macro)

(Micro & Macro)

indirect (heteroxenous) life cycles; all include both sexual and asexual phases (Bush et al. 2001). We use the terms and descriptions of Baer, as modified by Olsen and Bush (Baer 1951, 1971; Olsen 1974; Bush et al. 2001) to present the basic cycle. Although complex, the same terms and processes apply to virtually all Apicomplexa life cycles. Three distinct processes of multiplication generally occur among the Apicomplexa, including one sexual and two asexual forms (Fig. 6.2). Sexual reproduction (gametogony) among Apicomplexa, where known, is by syngamy (“gamete fusion”) followed by immediate meiosis to produce haploid progeny. Some authors prefer the term “gamogony” rather than “gametogony” (Duszynski and Upton 2001). Asexual reproduction of haploid stages occurs by binary fission, merogony (a form of multiple fission also called schizogony in earlier literature), endodyogeny (binary fission within a cell), and/or endopolyogeny (multiple fission within a cell) (Adl et al. 2005). If the final products of multiplication are merozoites, the process is called merogony, and if sporozoites, it is called sporogony (Bush et al. 2001). Merogony, often called schizogony in earlier literature, is a form of multiple fission leading to the formation of merozoites. Sporogony is a form of binary fission in the spore stage, leading to development of sporozoites.

Meront/Schizont

Y Merozoite

figure 6.2  Generalized life cycle for the Apicomplexa.

Sporozoites are the infective stage for vertebrate hosts and, once in the proper vertebrate host, infect the host cells and develop into trophozoites. Through repeated division (merogony), trophozoites form meronts (earlier called schizonts), which further develop into merozoites. For some Apicomplexa, merozoites may function as trophozoites by reinvading the same kind of host cell and producing additional generations of merozoites (Bush et al. 2001). Eventually, merozoites invade new vertebrate host cells and differentiate into either micro- or macrogametes through gametogony; in many cases the merozoite may first undergo repeated division to form many microgametes. Eventually, the micro- and macrogametes fuse to form a diploid zygote, followed by repeated meiotic division of this zygote to form oocysts, a protective stage within which one or more sporocysts develop, with each sporocyst further forming one of more sporozoites within itself. There are numerous variations as to the hosts and host tissues within which these various stages occur, whether the parasite has a direct life cycle or requires intermediate hosts or vectors, but the basic life cycle stages and three multiplication processes retain considerable consistency among the Apicomplexa. Although the taxonomy of the Apicomplexa frequently has been revised, three common groups of concern for wildlife diseases include kingdom protista

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the coccidia, malaria parasites, and piroplasms (App. 1: Table 5) (Cox 1991, Patterson 2000, Adl et al. 2005).

Coccidia Coccidia, variously classified as Eimeriida (Cox 1991), Eimeriorina (Bush et al. 2001), or Coccidiasina (Adl et al. 2005) are relatively widespread among birds and mammals. Every species of vertebrate ever examined intensively over a broad geographic range has at least one coccidian species unique to it, and some may have many more species (Duszynski and Upton 2001). Thus, there are many different species of coccidia causing specific problems, but very few species of broad impact in wildlife diseases. There are at least 15 genera of coccidia from birds and mammals (Todd and Hammond 1971, Lindsay and Todd 1993, Long 1993, Cole 1999a, Friend and Franson 1999). The focus in this text primarily will be on the genera Eimeria, Toxoplasma, and Sarcocystis. However, others, including Hepatozoon spp. (Craig 2001) and Besnoitia spp. (Leighton and Gajadhar 2001), can be of importance as well. Oocyst morphology is an important means by which various members of the coccidia can be identified, particularly with the numbers and combinations of sporocysts and sporozoites within each oocyst (Bush et al. 2001). For example, oocysts of members of Eimeria have four sporocysts, each of which has two sporozoites; in contrast, members of the genus Isospora have two sporocysts, each of which has four sporozoites. Among coccidia (e.g., Eimeria, Cryptosporidium, Cyclospora) with a monoxenous life cycle (Fig. 6.3), the infective sporozoite typically is ingested from contaminated soil or water and inhabits the intestine or liver, leaving again as undeveloped oocysts. Once out of the host, the oocysts sporulate to develop sporocysts and sporozoites in the ambient environment, and the sporozoites become infectious for a new host. Monoxenous coccidia most commonly are reported among herbivorous hosts. 172

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Sporozoite

In ambient environment

In herbivore host

Oocyst

Zygote

Merozoite

figure 6.3  Generalized direct (monoxenous) life cycle for coccidia.

Among coccidia (e.g., Sarcocystis, Toxoplasma) with a heteroxenous life cycle (Fig. 6.4), the sporozoite is ingested by the intermediate vertebrate host and develops to a trophozoite stage. When the intermediate host is eaten by the predatory definitive host, the trophozoite stage develops further to the oocyst stage, which is shed into the ambient environment. Following sporulation, the life cycle is completed when oocysts containing infective sporozoites are ingested by a susceptible host. Heteroxenous coccidia typically use carnivores as definitive hosts and herbivorous prey species as intermediate hosts. A few coccidian parasites (e.g., Isospora) can use optional (paratenic) vertebrate hosts (Dubey 1993). In paratenic hosts, the ingested sporozoite may become encysted (unizoite cyst) before being ingested by the predatory definitive host (Dubey 1993). There is evidence for an evolutionary transition among coccidia genera from monoxenous to heteroxenous life cycles (Belova and Krylov 2003, Šlapeta et al. 2003). Genera such as Sarcocystis, Frenkelia, and Besnoitia parasitize carnivores as definitive hosts and have required intermediate hosts (Dubey 1993). The closely related genus Toxoplasma functions as a transitional form; while it can complete a one-host life cycle in the cat, it primarily uses intermediate hosts to facilitate completion of its life cycle (Sanger 1971, Jindrichova et al. 1975, Dubey and Odening 2001). Some members of the genus Isospora

Sporozoite In herbivore intermediate host

In ambient environment

Trophozoite

In carnivore definitive host Oocyst

Zygote

also have this pattern (Dubey 1993). Details of these evolutionary relations are incomplete (Šlapeta et al. 2003).

Malaria Parasites (Haemosporidia) Malaria parasites are blood parasites found in a wide variety of mammals, birds, and reptiles; all are transmitted by hematophagous arthropods. The term “malaria” stems from earlier beliefs that bad air or gases (“mal-air”) from swamps caused the disease in humans. Variously identified as Haemosporina (Atkinson and van Riper 1991), Haemospororina (Bush et al. 2001), and Haemospororida (Cox 1991, Adl et al. 2005), all malarial parasites are vectorborne and involve a hematophagous invertebrate. The prefix “haemo” is the Greek root for blood, and “sporo” is the root for spore or seed. Genera of importance to wildlife include Haemoproteus, Leucocytozoon, and Plasmodium among birds (Atkinson and van Riper 1991, van Riper et al. 1994) and, to a lesser extent, Plasmodium in reptiles, rodents, and primates (Collins and Aikawa 1993, Cox 1993). Two genera causing avian malaria each have a large number of species, with each species often parasitizing a relatively limited host range; Leucocytozoon has 60 and Haemoproteus has 133 species (Bennett et al. 1994). In contrast,

Merozoite

figure 6.4  Generalized indirect (heteroxenous) life cycle for coccidia.

Plasmodium spp. generally have broader host specificity (Bennett et al. 1982). There are about 38 species of Plasmodium described from mammals (Collins and Aikawa 1993, Cox 1993, López-Antuñano and Schmunis 1993), 34 species from birds (Bennett et al. 1993), and 87 species from reptiles (Telford 1994). Malarial parasites have been recorded in about 68% of the avian species examined (Atkinson and van Riper 1991). Haemoproteus is the most common genus and has been reported in about 67% of malaria-infected birds, followed by Plasmodium (42%) and Leucocytozoon (39%) (Atkinson and van Riper 1991). Some host families (e.g., Fringillidae, Columbidae, Phasianidae) have relatively high prevalences of infection, whereas others (e.g., Laridae, Scolopacidae, Charadriidae) are only rarely infected with blood parasites (Atkinson and van Riper 1991). The reasons for these differences may involve behavior, habitat, suitable vectors, climate, environmental conditions, and host specificities (Atkinson and van Riper 1991). Malarial parasite life cycles involve asexual reproduction in both the vertebrate host and arthropod vector, and sexual reproduction in the arthropod (Fig. 6.5) (van Riper et al. 1994). The arthropod hosts for Haemoproteus are members of the Family Hippoboscidae for upland birds and members of Culicoides kingdom protista

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Sporozoite

Trophozoite

Oocyst

Zygote (Ookinete)

Meront/ Schizont

Invertebrate

Vertebrate

Gametes

Gametocytes

for waterfowl (Bush et al. 2001). Members of Leucocytozoon are transmitted by black flies (Family Simuliidae) (Cook 1971b). Among Plasmodium spp., mosquitoes of the genus Anopheles transmit all species infecting mammals as well as a few bird malarias; however, species of Culex transmit most avian malarias of Plasmodium spp. (van Riper et al. 1994). The invertebrate is the definitive host because it is the site of sexual reproduction and thus is where the adult stage of the parasite occurs. In contrast, the vertebrate is the intermediate host since only asexual reproduction occurs within it. The infective sporozoite stage enters the susceptible vertebrate host with salivary gland secretions during the bite of the hematophagous vector (Atkinson and van Riper 1991). After invading the host and undergoing intracellular growth, development, and asexual reproduction (merogony) in the host tissues, merozoites are formed. Merozoites invade circulating blood cells and may undergo additional merogony (e.g., Plasmodium) or undergo gametogony to form male (micro-) or female (macro-) gametocytes that are infective for another susceptible hematophagous vector (Atkinson and van Riper 1991). 174

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Merozoite

figure 6.5  Generalized malarial life cycle.

Once ingested by the invertebrate host, gametocytes form gametes that fuse in the arthropod midgut to form motile zygotes (ookinetes), which, in turn, penetrate the midgut epithelial cells (Atkinson and van Riper 1991). There, ookinetes round up into oocysts and undergo merogony to produce numerous sporozoites. Sporozoites rupture from the oocysts into the hemocoel of the vector and move to the salivary glands for injection into the next susceptible host (Atkinson and van Riper 1991). While similar, the life cycles of Plasmodium, Leucocytozoon, and Haemoproteus differ in the vertebrate host tissues they use for schizogony. Leucocytozoon and Haemoproteus undergo schizogony only in fixed, non-circulating cells in the host such as hepatocytes (liver cells) and vascular endothelial cells (cells lining blood vessels); further, only the gametocyte stage is observed in the circulating blood for these genera (Atkinson and van Riper 1991). In contrast, Plasmodium undergoes merogony in both fixed non-circulating cells (exoerythrocytic merogony) and circulating blood cells (erythrocytic merogony); thus, both gametocytes and meronts may occur in erythrocytes (Atkinson and van Riper 1991). The occurrence

figure 6.6  Description of a malarial infection (from Parasitic protozoa, Volume 7, C. van Riper III et al., Plasmodia of birds, pp. 73-140, J. Kreier (ed.), Copyright Academic Press 1994).

of erythrocytic meronts is a key difference of Plasmodium from Leucocytozoon and Haemoproteus (Atkinson and van Riper 1991). A distinct vocabulary is associated with malarial infections. The occurrence of parasites in the blood is termed “parasitemia” (Fig. 6.6). Following the bite of an infected hematophagous vector, the period of time between initial infection and when parasites are detected in the blood is termed the prepatent period; this is the time during which parasites are developing in various host tissues before invading the blood (Atkinson and van Riper 1991). The patent period is defined as the time when parasites can be observed in the blood, generally with a thick blood smear. The acute phase is when parasites first appear in the blood and rapidly increase in number. The crisis phase occurs when the parasitemia in circulating blood and the resulting physiological stresses reach a peak. Following the patent period, a latent or subpatent phase occurs when immune responses have reduced the parasitemia to low levels, where the parasites no longer are detectable in blood smears, and the surviving hosts have few or no overt signs of infection (Atkinson and van Riper 1991, van Riper et al. 1994). For species

typical of temperate areas, a relapse of parasites in the vertebrate host’s blood typically occurs the following spring, and serves to infect the new generation of hematophagous vectors. While some species of Haemoproteus sometimes are associated with host pathology (Julian and Galt 1980, Atkinson et al. 1988, Cannell et al. 2013), many species have little or no pathogenic effect (Cook 1971a, Bennett et al. 1988). Plasmodium and Leucocytozoon more commonly are associated with diseases among infected hosts (Atkinson 1999).

Piroplasms Piroplasms are sporozoan parasites of mammals and birds passed by ticks of the Family Ixodidae (Allan 2001, Kocan and Waldrup 2001). All cause a hemolytic anemia in the vertebrate host. Two families are recognized: Babesidae and Theileridae. At least 73 species of Babesia have been reported from mammals (Kakoma and Mehlhorn 1994), as well as 13 species from birds (Peirce 2000), and several from reptiles (Bush et al. 2001). Among mammals, species of kingdom protista

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Babesia infect primarily cervids and bovids, as well as some carnivores and rodents around the world. The 13 recognized species of Theileria all occur in mammals, primarily wild and domestic ungulates (Mehlhorn et al. 1994). The taxonomic status of the genus Cytauxzoon currently is unclear, as some scholars combine it with Theileria (Mehlhorn et al. 1994) and others keep it separate (Kocan and Waldrup 2001). Species of Theileria and Cytauxzoon occur in a variety of mammals throughout the world (Barnett and Brocklesby 1969), Theileria cervi, T. annulata, and Cytauxzoon felis are known from wild mammals of North America (Kocan and Waldrup 2001). The vectors for Babesia spp. have been identified for only 17 of the 73 recognized species; all are ticks (Kakoma and Mehlhorn 1994). Known ixodid tick vectors for Babesia include species of Boophilus, Dermacentor, Haemaphysalis, Rhipicephalus, and Hyalomma (Kakoma and Mehlhorn 1994). For Theileria, known ixodid tick vectors include species of Rhipicephalus, Hyalomma, and Amblyomma (Mehlhorn et al. 1994). The life cycles of piroplasms involve processes similar to those of malaria parasites (Fig. 6.5). Merogony occurs in the vertebrate host, whereas gamete maturation, fusion, and formation of a motile zygote (kinete) occur in the tick intestine (Bush et al. 2001). Piroplasm sporozoites are injected into susceptible hosts with the saliva of infected ixodid ticks (Kakoma and Mehlhorn 1994, Mehlhorn et al. 1994). Babesia spp. sporozoites invade the cytoplasm of erythrocytes of the vertebrate hosts and undergo several cycles of merogony to form merozoites (Kakoma and Mehlhorn 1994). Theileria spp. sporozoites invade the cytoplasm of erythrocytes, as well as newly dividing lymphocytes, which continue to divide together with the parasite (Mehlhorn et al. 1994, Bush et al. 2001). Sexual reproduction occurs in the tick vector (Mackenstedt et al. 1990). For both genera, the newly formed merozoites invade other erythrocytes and are then ingested by the appropriate tick vectors, in which they undergo 176

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gamogony (gametogony) and form gametes and kinetes (motile zygotes) in the tick gut. Babesia spp. kinetes can penetrate a number of tick cell types and divide by multiple fission to produce additional kinetes (Kakoma and Mehlhorn 1994). Kinetes eventually migrate to the tick salivary glands, where sporogony results in the production of many sporozoites, which, in turn, are transmitted to the next generation of vertebrate hosts (Bush et al. 2001). For Theileria, kinetes develop only in the salivary glands of the tick hosts (Mehlhorn et al. 1994). Members of the Theileridae and Babesidae differ in a number of ways. Theileridae undergo merogony in lymphocytic cells in addition to their merogony in erythrocytes; in contrast, members of the Babesidae generally undergo asexual multiplication (merogony) only in erythrocytes of the vertebrate host. However at least two species of Babesia also may invade lymphocytes as sporozoites (Moltmann et al. 1983, Mehlhorn et al. 1986, Kakoma and Mehlhorn 1994). Members of the Babesidae undergo both transstadial (across stages of the tick vector life cycle) and transovarial transmission in the tick vector; in contrast, only transstadial transmission is known to occur in the Theileridae (Kocan and Waldrup 2001). Thus, transmission of Theileria requires multi-host ticks whereas Babesia, which can be transmitted transovarially, can be transmitted by 1-host ticks (Mehlhorn et al. 1994). For Babesia, ticks are biological vectors; mechanical transmission by inoculation of infective blood by flying insects does not occur (Kakoma and Mehlhorn 1994). In contrast to Babesia, Theileria also can be transmitted mechanically by the bite of bloodsucking insects (Mehlhorn et al. 1994).

Ciliophora (Ciliates) The Ciliophora (ciliates) are a large and diverse group of protista related to the Apicomplexa, which are also in the Alveolata (App. 1: Table 5) (Adl et al. 2005). All members have cilia as organelles of motility; species are distinguished on the basis of their pattern of

cilia and associated cortical structure, and on the basis of their nuclear structure and function (Bush et al. 2001). Ciliates commonly divide by binary fission, although conjugation can occur (Bush et al. 2001). In contrast to the Apicomplexa, there are very few ciliates that cause disease among vertebrates (Bush et al. 2001). All ciliates have a direct (monoxenous) life cycle. Among wildlife, only Balantidium coli has been reported to cause disease with any regularity (Lucius and Loos-Frank 1997, Kocan 2001a). Some ciliates, including B. coli, produce resistant cyst stages (Bush et al. 2001).

Excavata (Flagellates) A diverse variety of eukaryotic single-celled organisms have a life history stage using one or more flagellae as organelles of motility. Parasitic f lagellates infect most animal phyla and occupy a variety of host habitats, including the intestines, reproductive tract, deep body tissues, as well as intracellularly and extracellularly in the blood vascular system (Bush et al. 2001). Until recently, flagellates were believed to be closely related to amebae (Bush et al. 2001). However, most flagellates currently are classified within the Supergroup Excavata (Adl et al. 2005). Further, within the Excavata, flagellates are broadly distributed among three of the six First Rank subgroups, including Fornicata, Parabasilia, and Euglenozoa. From a clinical perspective, flagellates often have been distinguished as intestinal flagellates (included in Fornicata and Parabasilia) or hemoflagellates (in Euglenozoa) (Chandler and Read 1961, Kocan 2001a). Intestinal Flagellates Most intestinal flagellates of concern to wildlife fall within the Trichomonadida subgroup of the Parabasilia (Adl et al. 2005). Important genera include Histomonas and Trichomonas. Their flagellae are often associated with a lamellar undulating membrane. Another member of

this group, Dientamoeba fragilis, no longer has a flagellum (BonDurant and Wakenell 1994). Most trichomonad species are not established pathogens for mammals or birds (BonDurant and Honigberg 1994). However, a few are of importance, including Histomonas meleagridis and Trichomonas gallinae in birds (BonDurant and Honigberg 1994, BonDurant and Wakenell 1994). Another group among the Excavata, the Fornicata, lack typical mitochondria and generally have either a single nucleus or pair of nuclei (Adl et al. 2005). Giardia is one important genus in this group, infecting amphibians, reptiles, birds, and mammals, including humans (Olson and Buret 2001). Giardia spp. have two nuclei in their trophozoites and four nuclei in their cysts (Garcia and Bruckner 1997). They lack a functional feeding apparatus; each organism has one posteriorly directed flagellum running through the length of the cell axially and within the cytoplasm (Adl et al. 2005). While classification of Giardia species still is quite controversial (Kulda and Nohýnková 1994), members of the genus cause serious diarrheal disease in humans as well as diarrhea and allergies in domestic animals; Giardia also infects a variety of wild animals, but usually with fewer clinical signs (Olson and Buret 2001). Life cycles among the intestinal flagellates are varied, but most generally are simple. The intestinal flagellates divide by binary fission, and most, including Giardia and Trichomonas, have a direct life cycle; Giardia forms cysts, whereas Trichomonas and Histomonas lack cyst stages (BonDurant and Honigberg 1994, BonDurant and Wakenell 1994, Kulda and Nohýnková 1994). Transmission is direct, including ingestion of trophozoites (e.g., Trichomonas gallinae) or encysted stages (e.g., Giardia), or sexual intercourse (e.g., Trichomonas vaginalis). Rarely, transmission also may be indirect, carried through helminth infections; both Histomonas meleagridis (Graybill and Smith 1920) and Dientamoeba fragilis (Burrows and Swerdlow 1956, Ockert and Schmidt 1976, Sukanahaketu 1977) are kingdom protista

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figure 6.7  Trypanosoma evansi in mouse blood (Courtesy of W. Frank, Universität Hohenheim, Germany).

helminth-transmitted parasites. For H. meleagridis, the nematode Heterakis gallinae is critical to the natural transmission of the parasite; earthworms further serve as paratenic hosts in this complex (BonDurant and Wakenell 1994). For D. fragilis, the human pinworm Enterobius vermicularis plays an important role in the life cycle. Hemoflagellates Among the Excavata, the Euglenozoa include parasitic hemoflagellates of importance to wildlife (Adl et al. 2005). Two important parasite genera in this group, Trypanosoma and Leishmania, both have a single flagellum; they multiply by binary fission and only rarely undergo sexual reproduction (Bush et al. 2001, Adl et al. 2005). These hemoflagellates live in the blood, lymph, and other tissues of vertebrates (Fig. 6.7). Morphological features such as length, shape, and location within the parasite of organelles such as the basal body, kinetoplast, nucleus, and flagellum are used to distinguish various species (Logan-Henfrey et al. 1992). Most hemoflagellates are found in tropical and subtropical regions of the world. Species of Leishmania are found in Central and South America, Asia, India, and China, with occasional reports in North America (Lainson 1982). These parasites live in the macrophages of the lymph nodes, spleen, and liver of their vertebrate hosts and are parasites of mammals; carnivores and rodents commonly serve as important vertebrate reservoirs. The 178

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invertebrate vectors are sand flies (Phlebotomus spp.) (Lainson 1982). Species of Trypanosoma infect a wide variety of vertebrates and arthropods in Africa, Asia, and North and South America (Wells and Lumsden 1969). Although most wildlife-related studies with Trypanosoma have involved mammals (Molyneux 1982), all vertebrate classes are parasitized by species of Trypanosoma (Herman 1969, Bardsley and Harmsen 1973, Baker 1976, Lom 1979, Telford 1995). Avian hosts include raptors, waterfowl, columbiforms, galliforms, and passerine birds (Gylstorff and Grimm 1987). The taxonomy of Trypanosoma spp. is controversial. Members of the genus historically have been divided into a) species that develop in the midgut or hindgut of their insect vectors and are passed by scratching infective stages from contaminated feces into a wound produced by the bite of an insect vector (Stercoraria), and b) species that develop in the midgut or mouth parts of their insect vectors and are passed directly by arthropod bite (Salivaria) (Hoare 1966). In more recent years, parasites in the genus Trypanosoma have been viewed as a complex taxonomic unit that likely includes many different genera; species designations often include reference to a new proposed genus, as in Trypanosoma (Schizotrypanum) cruzi, T. (Trypanozoon) brucei, T. (Nannomonas) congolense, and T. (Duttonella) vivax (Bush et al. 2001). However, we continue using the standard Trypanosoma genus in this text. Trypanosoma cruzi, a Stercoraria, is a significant human problem in Central and South America; it develops in the hind-gut of its hosts and infects its vertebrate host through the vector’s excrement, which is rubbed into the bite wound by the host (Acha and Szyfres 2003). About 150 mammal species can serve as vertebrate hosts, ranging from marsupials (Didelphis marsupialis) to primates; prevalence can be high in cats, dogs, rodents, and both domestic and wild lagomorphs (Acha and Szyfres 2003). About 100 species of triatomid bugs (Hemiptera: Reduviidae) can transmit these parasites,

figure 6.8  Key developmental stages of hemoflagellates. a. trypomastigote; b. opisthomastigote; c. epimastigote; d. promastigote; e. choanomastigote, f. amastigote (Lucius and Loos-Frank, 1997; courtesy R. Lucius, Humboldt University Berlin).

but only about three are considered as significant vectors for human disease (Sousa and Johnson 1973, Cedillos 1975, Acha and Szyfres 2003). Trypanosomiasis in Africa is one of the most important diseases of humans and livestock (Wells and Lumsden 1969, LoganHenfrey et al. 1992). Wildlife serve as reservoir hosts (Kocan 2001b). Trypanosoma brucei is one important species; two subspecies, T. brucei gambiense and T. brucei rhodesiense, are important causes of sleeping sickness among humans (Wells and Lumsden 1969). Another subspecies, T. brucei brucei, along with T. vivax and T. congolense, are important parasites, causing considerable morbidity and mortality with a disease called nagana among cattle of Africa (Wells and Lumsden 1969). All of these use species of tsetse flies (Glossina spp.) as vectors (Wells and Lumsden 1969). Trypanosoma and Leishmania have relatively complex life cycles involving both vertebrates and hematophagous insects (Lucius and LoosFrank 1997, Bush et al. 2001). Several distinct life history stages occur among the vertebrate and invertebrate hosts (Hoare and Wallace 1966) (Fig. 6.8). The names of the life cycle stages each are derived from a combination of the Greek root “mastigote,” meaning whip, and an appropriate prefix (Hoare and Wallace 1966). The main morphological features distinguishing the developmental stages in trypanosomes

are the position of the kinetoplast (a specialized region of the mitochondrion containing a dense network of DNA), the way in which the flagellum is attached to the body of the cell, and the extent of the flagellum’s extension anteriorly (Logan-Henfrey et al. 1992). When present, the trypomastigote and amastigote forms typically develop in vertebrates, and the epimastigote and promastigote develop in the intestinal tract of the infected insects (Lucius and Loos-Frank 1997). The trypomastigote stage is the common form in the peripheral blood of the vertebrate host during early stages of infection for Trypanosoma (Bush et al. 2001, Kocan 2001b). “Metacyclic trypomastigote” is a term for the stage in the invertebrate host that is infective for the susceptible vertebrate host. Amastigotes are stages of some trypanosomes infecting vertebrate tissues (Bush et al. 2001). Promastigotes are stages typically found in the midgut of infected insects for Leishmania and some Trypanosoma. In Trypanosoma, the epimastigote stages occur in the insect midgut; epimastigotes divide by binary fission (Bush et al. 2001). The sphaeromastigote (not pictured) is a stage identified in the T. cruzi life cycle as occurring between amastigote and epimastigotes stages in mammalian hosts (Bush et al. 2001). Collectively, metacyclic stages are defined as the final stage of development of parasitic kingdom protista

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Table 6.1 Life History Types Found among Protista, with Representative Genera for Each

Life History Type

Representative Genera

Direct (Homoxenous) Life Cycles No cyst stage

Trichomonas

Cyst stage

Entamoeba, Giardia, Eimeria, Balantidium

May use paratenic host

Isospora

Indirect (Heteroxenous) Life Cycles Vertebrate Intermediate host

Toxoplasma, Sarcocystis

Arthropod-borne

Plasmodium, Leucocytozoon, Babesia, Hepatocystis, Trypanosoma, Leishmania

Helminth-borne

Histomonas, Dientamoeba

Free-living Soil and Water Protista (Opportunistic parasites)

Naegleria, Acanthamoeba, Balamuthia

kinetoplastid flagellates during the invertebrate (or vector) phase of their life cycle and are the infective stages for their vertebrate host (Bush et al. 2001). In T. cruzi, it is the infective stage that develops in the reduviid rectum from epimastigotes. In Salivaria, the infective (trypomastigote) stage develops in the mouthparts of the insect (Bush et al. 2001); these stages do not divide. The trypomastigote and epimastigote stages are found only in the genus Trypanosoma; in contrast, the amastigote and promastigote stages are represented in some species among both Trypanosoma and Leishmania (Bush et al. 2001). Trypomastigote stages occur among all members of Trypanosoma; amastigotes are tissue stages found among the infected vertebrates with T. cruzi, Leishmania spp., and others. A procyclic stage refers to any developmental stage in the life cycle of a hemoflagellate in the invertebrate host. These most often are found in the midgut of the host (Bush et al. 2001). The role of wildlife is complex for these parasites. Most infections by hemoflagellates in wildlife are benign (Baker 1969, Kocan 2001b), but some infecting humans and domestic animals can cause severe diseases (Molyneux 1982, Bruckner and Labarca 1999). In southern

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Mexico, and in Central and South America, Trypanosoma cruzi is a significant cause of morbidity and mortality among humans (Chagas’ disease) (Acha and Szyfres 2003). There are not many records of trypanosomes being pathogenic to African ungulates (Baker 1969); this could be due to tolerance of these vertebrates to the parasites or to predation of any unfit animal (Bertram 1973, Carmichael and Hobday 1975, Molyneux 1982). Based on laboratory experiments with captive African ungulates, some animals have lesions but generally appear to be unaffected (Molyneux 1982).

LIFE HISTORY TYPES AMONG PROTOZOA There is a wide variety of life history strategies used by parasitic protozoa to be successful. The major ones of importance to wildlife are presented in this section (Table 6.1).

Direct (Homoxenous) Life Cycles No Cyst Stage trichomonas gallinae c ausative agent (cl assific ation, morphology) The genus Trichomonas and other trichomonads are classified among

the Parabasilia of the Supergroup Excavata (App. 1: Table 5) (Adl et al. 2005). The genus Trichomonas includes a number of species infecting birds and mammals, of which one, Trichomonas gallinae, is a wildlife pathogen that infects the upper respiratory tract of birds (Cole 1999b). Another, T. foetus, is a parasite of the urogenital system of cattle (BonDurant and Honigberg 1994), and T. vaginalis infects the urogenital system of humans (Leber and Novak 1999). host range and distribution  Trichomonas gallinae has a worldwide distribution, with the domestic pigeon (Columba livia) serving as primary host (BonDurant and Honigberg 1994). In North America, it is speculated that T. gallinae generally occurs wherever domestic pigeons and mourning doves (Zenaida macroura) are found (Stabler 1954, Cole 1999b). This parasite also infects other columbiforms, as well as falcons and hawks, owls, passerine birds, upland birds, waterfowl, and gulls (BonDurant and Honigberg 1994, Cole 1999b). life cycle and variations Trichomonads have the simplest kind of protozoan life cycle, in which the organism occurs only as a trophozoite, with no cyst. These parasites multiply by binary fission (Leber and Novak 1999). reservoirs and tr ansmission The common pigeon (Columba livia) is believed to be the ultimate source of T. gallinae infection throughout the world (BonDurant and Honigberg 1994). The parasite was introduced to the United States with the pigeons and doves brought by European settlers (Stabler 1954). Most nestlings become infected and, once acquired, the infections can last up to 2 years (BonDurant and Honigberg 1994). Even though individual birds can lose an infection, the pattern of communal living helps ensure that most birds remain infected (BonDurant and Honigberg 1994). Because there is no resistant cyst stage, transmission from host to host must be relatively direct. In pigeons, the parasite is spread during courtship behavior and infection of young through the process of regurgitive feeding of

partially digested foodstuffs (“pigeon’s milk”) produced in the crops of the adults (BonDurant and Honigberg 1994). Because T. gallinae is not found in the intestinal tract past the muscular stomach, it cannot be transmitted in fecal droppings; rather, it is transmitted only from the mouth, and secondarily the nares and eyes (BonDurant and Honigberg 1994). Transmission to species other than pigeons is through ingestion of parasites from water contaminated by infected pigeons, or through ingestion of infected hosts (Stabler 1954, Pokras et al. 1993, Cole 1999b). clinic a l effec t s and di agnosis  Although trichomoniasis is a general term for an infection, the disease also is termed “canker” in columbiform birds and “frounce” in raptors (Pokras et al. 1993). For columbiforms, most T. gallinae strains are nonpathogenic or only moderately pathogenic for their avian hosts; however, there also are a number of virulent strains (BonDurant and Honigberg 1994). Virulent strains kill young birds, and thus often are selected out of a population. Benign strains may provide immunity against the more virulent strains. The normal sites of infections in columbiforms are the mouth, pharynx, esophagus, and crop. Virulent strains may cause lesions in the upper digestive tract and spread to other parts of the body through these ulcerations (BonDurant and Honigberg 1994). Infected birds often appear listless, with ruffled feathers, and with yellowish caseous lesions around the beak or eyes. Wet, sticky discharges and nodules within the mouth are characteristic of acute disease, whereas hard caseous lesions are associated with chronic infections (Cole 1999b). Lesions in the pharynx or crop may spread to the esophagus and eventually block it, resulting in starvation or even suffocation (Cole 1999b). Trichomonas gallinae can be diagnosed in wet smears from the mouth and oropharyngeal cavity of pigeons or by cultivation of swabs taken from those areas (BonDurant and Honigberg 1994). Stabler argued that examining samples directly in kingdom protista

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saline solution was better than trying to culture organisms. He described the following technique: Sit, place the pigeon on its back on the examiner’s left leg; place the floor of the thumb against the tip of the lower mandible and pull back gently, thus opening the bird’s mouth; with a pair of curved forceps, stroke the roof of the mouth back of the palatal flaps in an effort to remove saliva for examination (the curve of the forceps should stroke and lift the rear of the flaps); dab saliva in a drop of saline and examine on ’scope. This procedure, when mastered, is quick, easy, and much more efficient than all the swabbing and culturing! (Stabler 1975)

We found this technique to work very well. More recently, the InPouch TF culture system (BioMed Diagnostics, San Jose, California) has been very effective compared with other techniques, including wet mount and direct microscopic examinations, for detecting T. gallinae (Cover et al. 1994, Bunbury et al. 2005). population effects  Trichomonas gallinae is a very common parasite of pigeons and sometimes can cause mortality involving thousands to tens of thousands of birds (Cole 1999b, Höfle et al. 2004). Trichomoniasis is considered by many avian disease specialists to be the most important disease of mourning doves (Zenaida macroura) in North America (Conti 1993). Band-tailed pigeons (Columba fasciata) also have experienced considerable losses (Cole 1999b). The role of T. gallinae as an agent causing declines or suppression of wild populations is not clear, but in one study on mourning doves T. gallinae was not a significant contributor to a population decline (Ostrand et al. 1995). Trichomoniasis in raptors (frounce) has caused mortality in falcons, and less commonly in hawks and owls, for hundreds of years (Bert 1619, Stabler 1954, Cooper 2002), and may have serious impacts on some raptors populations (Cooper and Petty 1988, Pepler and Oettlé 1992). Trichomoniasis among wild raptors is linked to their feeding on urban pigeons. With Cooper’s hawks (Accipiter cooperii), prevalence of T. gallinae was significantly higher among urban nestlings (85%) than nestlings in undeveloped natural areas (exurban sites) (9%) (Boal et al. 1998); 182

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nestling mortality also was higher in urban than in exurban nests (Boal and Mannan 1999). In contrast, very few adults were found infected (Boal et al. 1998). In a study among northern goshawks (Accipiter gentilis), overall nestling prevalence among urban birds was 65% and tended to increase with age of nestlings and brood size (Krone et al. 2005). However, prevalences of T. gallinae in raptors can vary considerably by site (Rosenfield et al. 2002), and the overall impact of this parasite on raptors is not well understood. special problems Although the parasite has a very broad distribution, introduction of T. gallinae to new sites has been of some concern. Domestic pigeons were introduced to the Galápagos Islands in 1972 or 1973 (Harmon et al. 1987). A high prevalence later was found among these introduced birds, along with T. gallinae infections among endemic Galápagos doves (Zenaida galapagoensis); doves examined on pigeon-free islands were not infected (Harmon et al. 1987). control For virulent T. gallinae strains, removing infected individuals and any other source of infection is an effective strategy to protect susceptible birds (BonDurant and Honigberg 1994). Preventing concentrations of doves at bird feeders and artificial watering sites can help reduce transmission among wild populations (Cole 1999b). Although environmental persistence of T. gallinae is of short duration, bird feeders and other sites of columbiform concentrations should be kept clean and fresh (Cole 1999b). Most common antibiotics are ineffective against T. gallinae infections and are not practical for wild populations (BonDurant and Honigberg 1994, Cole 1999b). Several nitroimidazole compounds are effective when administered in drinking water, but this group of drugs cannot be used for any food-producing animals in the United States (McDougald 1991, BonDurant and Honigberg 1994). Cyst Stage Present eimeria spp. c aus at i v e agen t (cl a ssific at ion, morphology)  Members of the genus Eimeria fall within the Supergroup Chromoalveolata

(App. 1: Table 5) (Adl et al. 2005). As Apicomplexa, all are parasitic (Lindsay and Todd 1993). Most Eimeria spp. are parasites of the gastrointestinal tract, although some also invade other tissues (Lindsay and Todd 1993). host range and distribution Members of the genus Eimeria have a worldwide distribution, with many species affecting a wide variety of hosts among all classes of vertebrates (Gylstorff and Grimm 1987, Cole 1999a, Friend and Franson 1999, Bush et al. 2001, Duszynski and Upton 2001). Well over 800 species of Eimeria are reported from mammals alone, and this may be only a fraction of the total in mammals (Duszynski and Upton 2001). While at least 60 avian species of Eimeria have been reported (Todd and Hammond 1971), many of these species are not well understood. Much of the information on avian coccidiosis is through work on domestic birds. There currently are seven recognized species of Eimeria in chickens, at least four species in domestic turkeys, 10 species in pheasants, two species in guinea fowl, at least six from partridge and quail, and about 15 species in geese (Long 1993). At least six species have been described from raptors (Cawthorn 1993). life cycle and variations  All members of the genus Eimeria have a direct life cycle (Fig. 6.3) that commonly requires about a 1-week period. Eimeria spp. sporozoites infect cells of the intestinal mucosa and may cause serious disease in some hosts. Thick-walled and environmentally resistant oocysts are produced and excreted into the environment. Each mature oocysts contains four sporocysts; each sporocyst has two sporozoites (Bush et al. 2001). Sporogony requires moisture and oxygen, and is most effective between 10 and 35°C (Lindsay and Todd 1993). reservoirs and transmission It is not clear whether most species of Eimeria have only a single vertebrate host (Lindsay and Todd 1993, Bush et al. 2001) or if they regularly can infect more than one species (Duszynski and Upton 2001). Among wild mammals, members of the Orders Rodentia and Artiodactyla have

the largest numbers of known Eimeria species, followed by Lagomorpha, Carnivora, Diprodontia, Insectivora, and Chiroptera (Duszynski and Upton 2001). Eimeria spp. infections are reported among wild birds of the Orders Anseriformes, Galliformes, and Passeriformes (Todd and Hammond 1971). Transmission typically is by ingestion of a mature oocyst containing infective sporozoites (Duszynski and Upton 2001). clinical effects and diagnosis Enteritis, associated with diarrhea, is a common clinical syndrome. Some Eimeria spp. also may invade the liver, leading to hepatitis. However, pathogenicity of coccidian parasites in wild mammals may not be of great significance, and appears to be a disease due primarily to human intervention (Duszynski and Upton 2001). Among geese, there is a renal and an intestinal form of coccidiosis (Long 1993). Renal coccidiosis, caused by E. truncata, develops only in the kidneys and the cloaca near its junction with the ureters; the route to infecting the kidneys is unknown (Long 1993). Most other avian species of Eimeria are not considered severely pathogenic (Long 1993). When a host is not killed, diagnosis of a coccidian infection typically depends on finding oocysts in the feces and observing them completely sporulate (Duszynski and Upton 2001). The polymerase chain reaction and random amplified polymorphic DNA tests also are used to detect and identify coccidia (Comes et al. 1996, Barta et al. 1997, Duszynski and Upton 2001). population effects Among mammals, Eimeria stiedae, for example, may be a limiting factor on young wild rabbits (Oryctolagus cuniculus) of Australia in normal and wet years, but not in dry years (Dunsmore 1971). Most wild birds pass small numbers of oocysts in their feces without apparent ill effects, but coccidiosis becomes important as a disease when animals live under crowded or dense populations that allow buildup of infective oocysts in the environment and ingestion by susceptible hosts (Long 1993). Among domestic birds, it is speculated that the poultry industry is dependent on continuous medication in the feed (Long 1993). kingdom protista

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Renal coccidiosis, caused by E. truncata, has caused losses as high as 87% among infected, presumably domestic, flocks of birds in Iowa (Long 1993). This parasite also has been associated with mortality among Canada geese in North America (Critcher 1950, Farr 1954, Hanson et al. 1957) special problems No ongoing special problems are noted for Eimeria spp. control Among domestic birds, there are a wide variety of anti-coccidial chemotherapies used to combat Eimeria infections (Long 1993). There also have been some efforts to immunize domestic birds by use of attenuated strains of Eimeria, as well as vaccinating with specific antigens (Long 1993). Immunity does not play a significant role; among domestic birds immunity lasts only 4 to 6 weeks in birds recovering from infection (Long 1993).

Indirect (Heteroxenous) Life Cycles Use Vertebrates as Both Intermediate and Definitive Hosts toxoplasma gondii c aus at i v e agen t (cl a ssific at ion, morphology)  Toxoplasma gondii is classified as a coccidian parasite, with Eimeria and Sarcocystis, among the Apicomplexa in the Chromoalveolata (Dubey and Odening 2001). The name Toxoplasma (toxon: arc, plasma: form) is derived from the crescent shape of the tachyzoite stage (Hill et al. 2005). Toxoplasma gondii is the sole member of the genus (Hill et al. 2005); currently, three dominant genotypes (I, II, III) are recognized in the species (Grigg et al. 2001, Su et al. 2003, Volkman and Hartl 2003). Recently, another type (X) has been observed in marine mammals (Miller et al. 2004). host range and distribution Although it can only complete its life cycle in felids (Dubey and Odening 2001), T. gondii can infect nearly all mammal and bird species and has a worldwide distribution (Lehmann et al. 2006). In recent years, it has been described 184

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and become widely reported among marine mammals (Dubey 2004). Toxoplasma gondii is postulated to originally have been a parasite of South American felids that underwent a relatively recent worldwide expansion from the effects of both migratory birds and the expansion of maritime travel; in particular, the transatlantic slave trade may have played a considerable role in this expansion (Lehmann et al. 2006). life cycle and variations Wild and domestic felids are the only known definitive hosts; oocysts that successfully sporulate in the environment and infect susceptible hosts are formed only in cats (Fig. 6.9: 1 ) (Dubey et al. 1970). In contrast to other coccidia, several life history stages of T. gondii can infect both intermediate hosts and definitive hosts. The three infectious stages of T. gondii are sporozoites, tachyzoites (trophozoites), and bradyzoites, developing in meront-like tissue cysts called pseudocysts) (Dubey 2004). Most natural infections probably are acquired by ingesting sporulated oocysts with sporozoites in food or water contaminated with cat feces, or by ingesting pseudocysts in infected meat (Hill et al. 2005). Oocysts transform into tachyzoites shortly after ingestion by intermediate hosts (Fig. 6.9: 2 ). Tachyzoites rapidly proliferate in pseudocysts by binary fission (Bush et al. 2001) and destroy infected cells during acute infection. Tachyzoites localize in neural and muscle tissue and develop into bradyzoites within pseudocysts (Fig. 6.9: 3 ). Bradyzoites multiply slowly in tissue cysts (Bush et al. 2001). Cats become infected by consuming intermediate hosts with these tissue cysts (Fig. 6.9: 4 ). Toxoplasma gondii also may spread by infected macrophages to mesenteric lymph nodes and then to distant organs by invasion of the lymphatic system and blood (Hill et al. 2005). The host cells die and release tachyzoites, which invade adjacent cells and continue the process (Wilson and McAuley 1999, Hill et al. 2005). Tachyzoites are pressured by host immune responses to transform into bradyzoites and to form meront-like pseudocysts (Wilson and McAuley 1999). Although

figure 6.9  Life cycle of Toxoplasma gondii. Three sources of infection (a, b, c) occur for both intermediate and definitive hosts. Bradyzoites in pseudocysts will become tachyzoites if ingested by a new host, and then form bradyzoites in that intermediate host; or they will go on to form gametes and oocysts if they are in a felid definitive host (Courtesy of Centers for Disease Control and Prevention’s Division of Parasitic Diseases and Malaria, www.dpd.cdc.gov/dpdx).

pseudocysts containing bradyzoites may develop in visceral organs, including lungs, liver, and kidneys, they are more prevalent in muscular and neural tissues, including the brain, eye, skeletal, and cardiac muscle (Hill et al. 2005). Infected intermediate hosts may contain infective cysts for their lifetime and shed tachyzoites, but they cannot complete the life cycle (Hill et al. 2005). Generally, cats are more readily infected by ingesting bradyzoites than oocysts (Dubey 2004); ingesting as few as one bradyzoite can lead to a cat shedding millions of oocysts (Dubey 2001). Virtually all cats ingesting pseudocysts containing bradyzoites will shed oocysts, whereas less than half of the cats ingesting tachyzoites or oocysts will complete the life cycle (Dubey and Frenkel 1972). When felids ingest pseudocysts from infected prey, the released bradyzoites penetrate

epithelial cells of the small intestine (lamina propria) (Dubey 2004). In the epithelial cells, T. gondii multiplies (enteroepithelial cycle) and forms meronts (also called schizonts), which, in turn, continue developing to merozoites, gametes, zygotes, and oocysts (Dubey and Frenkel 1972, Hill et al. 2005). Formed oocysts are discharged into the intestinal tract by rupture of the epithelial cells and sporulate about 1–5 days after passing into the environment (Hill et al. 2005). As this enteroepithelial cycle progresses, bradyzoites also penetrate the feline intestine and multiply as tachyzoites, eventually disseminating to other tissues in the cat. Both intestinal and extraintestinal stages of T. gondii may occur for several months, or even remain for the life time of the cat (Hill et al. 2005). Often, wild or domestic animals can become infected (Fig. 6.9: 5 ). Humans become infected kingdom protista

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by eating these animals, as well as by eating food or water contaminated by cat feces, including from cat litter boxes (Fig. 6.9: 6 ). In a more serious problem, developing fetuses may be infected transplacentally, sometimes with fetal mortality or severe neurological consequences after birth. reservoirs and transmission One collective population estimate of domestic and feral cats in the United States ranges from about 93 to 133 million cats [Introduced Species Summary Project: Domestic Cat (Felis catus), www .columbia.edu/itc/cerc/danoff-burg/invasion_bio /inv_spp_summ/Felis_catus.html]. Up to 2% of all cats are estimated to be shedding T. gondii oocysts at any given time, with occasional higher prevalences reported (Dubey 2004). Three known routes of transmission for T. gondii include transmission through oocysts shed in the feces of wild and domestic felids, ingestion of tissue cysts from meat, and congenital transmission (Fayer et al. 2004). The oocysts are the only stage to enter the general environment, and they can remain infectious for months to years under moist temperate conditions while being widely dispersed in large numbers (Yilmaz and Hopkins 1972, Frenkel et al. 1975); thus, oocysts are viewed as the most likely means of transmission to non-carnivores (Fayer et al. 2004). Land-based surface runoff is a likely source of T. gondii oocysts for marine mammals (Miller et al. 2002a). Immunity does not eradicate infection, and T. gondii tissue cysts persist several years after acute infections (Hill et al. 2005). It is not clear how T. gondii is destroyed in immune cells (Renold et al. 1992). Extracellular forms of the parasite are directly affected by antibody, but intracellular forms are not; it is believed that T-lymphocytes and lymphokines are more important than humoral factors in immune destruction of T. gondii (Renold et al. 1992). clinical effects and diagnosis Pathogenicity of T. gondii is determined by many factors, including the susceptibility of the host species, virulence of the parasitic strain, and the life cycle stage ingested; oocyst-induced infections are the most severe clinically in intermediate hosts, 186

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independent of dose (Dubey 2004). Although cats of any age can die of toxoplasmosis, kittens and those with depressed immunity are the most susceptible (Dubey and Carpenter 1993a, 1993b). Toxoplasmosis can be a severe disease in domestic animals, and may cause embryonic and fetal death, abortion, stillbirth, and neonatal death (Dubey 2004). Clinical signs of toxoplasmosis are nonspecific and mimic those of several other diseases (Dubey and Odening 2001). Trophozoites can invade many tissues, leading to fever, splenomegaly, myocarditis, pneumonitis, and encephalitis. The pathology described among marine mammals includes meningoencephalitis, with lesser observations of hepatitis, myocarditis, and lymphadenitis, as well as disseminated and congenital infections (Dubey et al. 2003, Kreuder et al. 2003, Dubey et al. 2004, Miller et al. 2004). Diagnosis of toxoplasmosis is made by biologic, serologic, or histologic methods, or by their combination (Hill et al. 2005). Oocysts in infected cats are diagnosed by concentration methods such as flotation in high-density sucrose solutions (Ruiz and Frenkel 1980). Isolation of T. gondii from animals by inoculation of laboratory animals and tissue cultures can lead to definitive diagnoses (Dubey and Odening 2001). However, for epizootiological surveys, serological prevalence is a better measure of T. gondii prevalence (Dubey 2004). Serological tests include the dye test, indirect fluorescent antibody test (Miller et al. 2002b), modified agglutination test (MAT), ELISA, and immunosorbent agglutination assay test (Dubey and Beattie 1988). Some work with polymerase chain reaction is promising (Hill et al. 2005). population effects  Generally, the parasite causes few problems among wildlife populations. One exception may be with sea otters (Enhydra lutris), and this is addressed in the next section. special problems  Toxoplasma gondii has been detected in a wide variety of geographic areas from seals (Lambourn et al. 2001, Dubey et al. 2003, Measures et al. 2004, Honnold

et al. 2005), dolphins (Dubey et al. 2003, Cabezón et al. 2004, Dubey et al. 2005), sea otters (Cole et al. 2000, Lindsay et al. 2001b, Miller et al. 2002b, Dubey et al. 2003), and other marine mammals (Cruikshank et al. 1990, Dubey et al. 2003, Philippa et al. 2004). Pathology and mortality have been described from dolphins (Inskeep et al. 1990, Jardine and Dubey 2002, Resendes et al. 2002), a seal (Dubey et al. 2004), and sea otters (Kreuder et al. 2003, Miller et al. 2004). In an experimental infection, Toxoplasma established patent infections in gray seals (Halichoerus grypus), but did not cause pathology (Gajadhar et al. 2004). The southern sea otter (Enhydra lutris nereis) is listed as threatened under the Endangered Species Act and is considered a keystone species strongly influencing the abundance and diversity of other species within the kelp forest ecosystem (Jessup et al. 2004). There has been a recent decline as a consequence of high mortality rather than low recruitment, with considerable concern among biologists and managers (Estes et al. 2003, Gerber et al. 2004). Current evidence is that toxoplasmosis is making a significant contribution to this mortality (Kreuder et al. 2003, Hill et al. 2005). Of the T. gondii evaluated from sea otters in one study, 40% were Type II and 60% were Type X; no Types I or III were observed (Miller et al. 2004). Subsequent work has an estimate of 72% Type X in sea otters, with the observation that the same Type X has been isolated from other marine mammals as well (Conrad et al. 2005). Of the potential sources of infection, ingestion of sporulated oocysts seems most likely; sea otters do not prey on warm-blooded animals, which are the usual intermediate hosts of T. gondii (Conrad et al. 2005). There is evidence that Toxoplasma gondii has invaded the marine ecosystem through runoff from terrestrial sources (Miller et al. 2002a, Fayer et al. 2004). Sea otters in areas of high freshwater runoff are more likely to be exposed to T. gondii than otters from low or medium freshwater runoff (Miller et al.

2002a). Increased risk of sea otter mortality has been linked to increased exposure to T. gondii (Miller et al. 2002a, Kreuder et al. 2003, Conrad et al. 2005). Collectively, there is good evidence that Toxoplasma oocysts are spread from terrestrial runoff into the marine ecosystems, where they are concentrated in the resident shellfish; shellfish are a major food source of the California sea otters and serve as a means of transmission (Mitchell and Sinai 2006), or as paratenic hosts (Arkush et al. 2003). Eastern oysters (Crassostrea virginica) are able to remove T. gondii oocysts from seawater readily and retain their infectivity for at least 85 days (Lindsay et al. 2001a, 2004). Similarly, another bivalve mussel, Mytilus galloprovincialis, also is able to take up and retain infective T. gondii oocysts (Arkush et al. 2003). It also has been observed that anchovies (Family Engraulidae), a small common saltwater fish, can be infected with T. gondii after exposure (Massie and Black 2008); these fish are common prey items of marine mammals. Thus far, natural infections have not been reported in these species, however, and their actual role in the epizootiology of the disease is unclear (Conrad et al. 2005). Interestingly, the oocysts also sporulate readily and remain viable in seawater for at least 6 months (Lindsay et al. 2003). Reasons for the unusual morbidity and mortality from toxoplasmosis in sea otters, compared to the subclinical or mild infections seen in most immunocompetent terrestrial animals, are unclear (Conrad et al. 2005). control  Prevention of Toxoplasma among humans and domestic animals generally is by maintaining cleanliness and reducing ingestion of contaminated food or water (Hill et al. 2005). There are no specific recommendations to prevent transmission of T. gondii by drinking water because the level of contamination is unknown and because it can be transmitted by several modes (Dubey 2004). At present there is no vaccine to control toxoplasmosis in any species (Dubey and Odening 2001). kingdom protista

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Arthropod-borne Apicomplexa plasmodium relictum (avian malaria) causative agent As Apicomplexa (Adl et al. 2005), all members of the genus Plasmodium are parasitic. There are about 34 recognized species of Plasmodium described from birds (Bennett et al. 1993). Over 25% of all avian host species are parasitized by members of the genus Plasmodium (Bennett et al. 1982, 1994). host range and distribution  Plasmodium relictum has a worldwide distribution (Bennett et al. 1993, Peirce 2005, Kimura et al. 2006); it is the most common avian Plasmodium species and has been reported from over 300 species and at least 70 families of birds (Bennett et al. 1993, Valkiūnas 2005). life cycle and variations Using the general malarial life cycle model (Fig. 6.5), P. relictum sporozoites are introduced to susceptible birds by the bites of infected culicine mosquitoes. Sporozoites invade a variety of cells, including the Malpighian body, and later the lungs, brain, and spleen (van Riper et al. 1994). After three or more generations of merogony in these tissues, the merozoites invade erythrocytes, where they also undergo merogony; in older literature this was referred to as exoerythrocytic (out of erythrocyte) schizogony and erythrocytic schizogony. The resulting merozoites then develop into gametocytes (van Riper et al. 1994). Gametocytes ingested by Culex spp. mosquitoes form gametes that fuse into zygotes that, in turn, become motile (ookinetes); ookinetes penetrate midgut epithelial cells and form an oocyst on the hemocoel side of the gut. Following sporogony, oocysts rupture and sporozoites are released into the hemocoel of the mosquito; from here they migrate to the salivary glands to be injected into the new avian host during a blood meal by the mosquito (van Riper et al. 1994). reservoirs and transmission  The avian malaria cycle requires the avian vertebrate host and culicine mosquitoes as the invertebrate hosts (van Riper et al. 1994). Although P. relictum completes its development in at least 188

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26 species of mosquitoes (Huff 1965, Garnham 1966), culicine mosquitoes are judged to be the most likely transmitters under natural conditions (van Riper et al. 1994). clinical effects and diagnosis Clinically ill birds may appear listless, have ruffled feathers, lose their appetite, and have difficulty breathing; pathologic changes often involve anemia as well as enlargement of the spleen and liver (van Riper et al. 1994). Some infected birds may show few if any symptoms. population effects  The parasite P. relictum capistranoae has been associated with severe population declines of native birds in Hawaii, including extinction of many native species of Hawaiian honeycreepers (Drepanidinae). These events followed introduction of the vector, the house mosquito, Culex quinquefasciatus, in approximately 1826 and introduction of the parasite through migratory birds (Warner 1968, van Riper et al. 1986). special problems Endemic Hawaiian forest birds are highly susceptible to infection with Plasmodium relictum capistranoae, with some having mortality rates of 65–90% after being bitten by a single infective mosquito (Atkinson et al. 1995, 2000, 2001b). High susceptibility of native Hawaiian honeycreepers (Family Fringillidae, Subfamily Drepanidinae) to avian malaria is believed to be one of the primary factors responsible for the disappearance of these birds from lowland habitats on the main Hawaiian islands (Atkinson et al. 2001a). There is compelling evidence that the near absence of native forest birds below elevations of 900 m in Hawaii was due to the Culex spp. vector and parasite transmission (van Riper et al. 1986). With further work, it was established that malaria transmission was greatest in mid-elevation forests where the highly susceptible native forest birds overlapped in distribution with mosquitoes and where these native birds themselves served as the primary reservoirs of infection (van Riper et al. 1986, Atkinson et al. 2001a, Woodworth et al. 2005). The native birds were almost entirely absent from low-elevation study areas

because of high mosquito densities; malaria prevalence in the primarily exotic, diseaseresistant lowland avifauna was low (van Riper et al. 1986, Woodworth et al. 2005). There is evidence that at least one native Hawaiian forest bird, the amakihi (Hemignathus virens), has endured the parasite, has mounted an immune response (Atkinson et al. 2001a), and is even recolonizing low-elevation habitats that contain higher densities of the culicine vector (Woodworth et al. 2005). control  No specific measures are taken to control avian malaria in wild birds. trypanosoma spp. (nagana) c ausative agent (cl assific ation, morphology) The nagana disease complex involves three species of trypanosomes (T. vivax, T. congolense, T. brucei brucei) (Logan-Henfrey et al. 1992); they are classified in the Kinetoplastea among the Euglenozoa in the Excavata Supergroup (Adl et al. 2005). These are long, spindleshaped cells (about 2 × 20 mµ) with a single flagellum (Logan-Henfrey et al. 1992). All three are salivarian parasites typically passed through the bites of tsetse flies. host range and distribution Distribution of this group of parasites is between 14°N and 29°S on the African continent (Wells and Lumsden 1969). Distribution is closely linked to the distribution of tsetse flies (Glossina spp.), which infest about 11 million km2 of Africa, or about one-half of the available arable land. A wide variety of African ungulates are infected with this complex of organisms (Ashcroft 1959, Baker et al. 1967, Geigy et al. 1967, Baker 1969). Most of these wild ungulates are resistant to the trypanosomes, but occasional pathology and disease has been reported among them (Molyneux 1982). life cycle and variations  The trypomastigote and the epimastigote (Fig. 6.8) stages are found among hosts of the African salivarian trypanosomes (Logan-Henfrey et al. 1992). Infected tsetse flies inoculate metacyclic trypomastigotes into the skin of animals, where the trypanosomes grow for several days and cause

local swellings. They enter the lymph nodes and then the bloodstream, where they divide rapidly by binary fission (Fraser and Mays 1986). After the tsetse fly feeds on a host infected with trypanosomes, the bloodstream trypomastigotes develop into procyclic trypomastigotes in the midgut, then into epimastigotes, and then into metacyclic trypomastigotes (LoganHenfrey et al. 1992). There also are a number of variations in the timing of the life cycle, the various stages developed in the hosts, and the host sites at which the parasites concentrate among the trypanosome species (Fig. 6.10) (Logan-Henfrey et al. 1992). reservoirs and transmission These African trypanosomes typically are passed by the bite of an infected tsetse fly. There are potentially 23 tsetse fly species involved, classified into three taxonomic groups according to their preferred ecological habitat. The Glossina morsitans group typically is found in savanna, the G. palpalis group prefers areas around rivers and lakes, and the G. fusca group lives in high forest areas (Logan-Henfrey et al. 1992). The limits of tsetse infestation are determined primarily by climate and secondarily by vegetation (Ford 1971, Logan-Henfrey et al. 1992). In addition to the typical transmission by hematophagous arthropods, there also is evidence that transmission of trypanosomes to predators such as lions (Panthera leo) and hyenas (Crocuta crocuta) can occur by ingestion of infected ungulates (Baker et al. 1967, Baker 1969, Geigy and Kauffman 1973). This insight is supported by laboratory studies (Heisch 1963, Moloo et al. 1973). clinical effects and diagnosis The severity of disease varies with the species of animal and the trypanosome species involved (Molyneux 1982). There are individual case reports of wild ungulate species in which lesions were attributed to trypanosome infections, but generally African ungulates have an innate tolerance of trypanosome infections (Molyneux 1982). For cattle and other susceptible species, the primary clinical sign is anemia, with an associated stunting and wasting kingdom protista

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figure 6.10  Life Cycle of a Salivaria (Trypanosoma brucei gambiense) (Courtesy of Centers for Disease Control and Prevention’s Division of Parasitic Diseases and Malaria, www.dpd.cdc.gov/dpdx).

(Molyneux 1982). Most infected cattle have a chronic disease course with high mortality (Fraser and Mays 1986); mortality usually is associated with circulatory disorder, damage to the heart, and the anemia (Molyneux 1982). There are few consistent clinical signs or internal lesions specific to the disease, and other diseases in Africa (e.g., anaplasmosis, babesiosis, theileriosis) can be confused with trypanosomiasis (Logan-Henfrey et al. 1992). Trypanosome infections may be detected directly by microscopic observation of the parasites, by immunoassays for trypanosome antigens, or by biochemical techniques that detect trypanosomal DNA. Identifying trypanosome infections also is based on serological detection of anti-trypanosomal antibodies (LoganHenfrey et al. 1992). population effects  There is little overall impact on indigenous wild ungulate species. However, much of Africa between 14°N and 190

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29°S was excluded for cattle due to the severe impacts of trypanosomiasis. There is virtually no overlap between the cattle-raising regions of Africa and the range of tsetse flies and the trypanosomes (Donelson and Turner 1985). special problems  Cattle have been denied vast areas of Africa by the occurrence of trypanosomiasis. Trypanosomiasis is regarded economically as the most important livestock disease in Africa (Jawara 1990), with an estimated annual loss to African farmers in meat production alone estimated at US$5 billion in the 1990s (Gyening 1990, Logan-Henfrey et al. 1992). control  Historically, risk of trypanosomiasis was minimized by avoiding tsetse-infected areas. More recently, control of trypanosomiasis in livestock relies on the treatment of the infected host and on vector control (LoganHenfrey et al. 1992). In the early 1900s, it was observed that the mortality among wild ruminants from

rinderpest was so high that the tsetse fly, especially Glossina morsitans, was severely reduced in some areas such as Kruger National Park, from lack of suitable hosts (Stevenson-Hamilton 1911, Duke 1919, Carmichael 1933). At that time, “game elimination” became a technique used to limit the sources of blood meals available to the tsetse flies, and to eliminate trypanosome reservoirs (Logan-Henfrey et al. 1992); this in turn was intended to reduce tsetse fly populations, specifically Glossina morsitans, and to enhanced success of cattle in the region; there was some evidence for success with this approach (Clarke 1964, Ford 1971). Between the 1920s and the 1960s, it was estimated that 1.3 million game animals were killed in tsetse control programs (Matthiessen and Douthwaite 1985). However, the public outcry against wildlife hunting as a means of tsetse control (Cockbill 1967) and the developing evidence for its overall inefficiency (Glover 1965), along with the advent of insecticides, largely ended the practice by the early 1970s (Matthiessen and Douthwaite 1985). At that time there also was an increasing appreciation of the game animals as valuable in themselves and as important national resources (Molyneux 1982). Vector control historically involved bush clearing in an effort to destroy tsetse fly habitat, to limit the sources of blood meals available to the tsetse flies, and to eliminate trypanosome reservoirs (Logan-Henfrey et al. 1992). Pastoralists and arable farmers tend to clear brush as part of their land use, making the land unsuitable for the flies (Matthiessen and Douthwaite 1985). Some savanna woodland clearing was done to destroy G. morsitans; while there is evidence that this has been effective in some regions, there is no overall assessment of its effects, but there is speculation that continued deforestation and other habitat changes changed and reduced the occurrence of tsetse flies, and trypanosomiasis, in parts of Africa (Molyneux 1982). Game destruction and bush clearing were replaced by insecticides for tsetse control (Jordan 1974, Allsopp 1984). A variety of insecticides have been used, but generally have had

limited effectiveness and often have had significant side effects on nontarget species (Du Toit 1954, Graham 1964, Wilson 1972, Molyneux 1982, Matthiessen 1985, Matthiessen and Douthwaite 1985). In recent years, use of traps and screens impregnated with pyrethrum has been effective in reducing tsetse flies of the G. palpalis and G. fusca groups; also, a number of attractants are effective for capture of the G. morsitans group of flies (Logan-Henfrey et al. 1992). There has been considerable interest in biological control, including use of sterilized males, but no effective techniques have emerged (Logan-Henfrey et al. 1992). There also is exploration of developing more trypanotolerant animals, but these animals represent only a small portion of the cattle in Africa (LoganHenfrey et al. 1992). Treatment of individually affected animals can be accomplished with a number of drugs, including dimazene aceturate, quinapuramine sulfate, and homidium bromide, among others (Logan-Henfrey et al. 1992). There is little likelihood of an effective vaccine because of the unique ability of trypanosomes to evade the immune system while in the blood stream by switching on new genes to encode new surface antigens (Donelson and Turner 1985). Trypanosomes may possess up to 1,000 variable surface glycoprotein (VSG) genes and, from these, they select certain ones to express when they are at the metacyclic stage of development, others as trypomastigotes in the early stages of infection in the blood, and still others during the later stages of chronic (Boothroyd 1985, Borst 1986, Van der Ploeg 1987, Pays and Steinert 1988). Helminth-borne histomonas meleagridis causative agent (cl assification, morphology)  Histomonas meleagridis is included among the Trichomonadida of the Parabasalia, in the Supergroup Excavata (Adl et al. 2005). Histomonas meleagridis is one of two species in this genus and the only one of pathogenic significance. Two forms of H. meleagridis kingdom protista

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have been described, a lumen form with one or two flagellae, and a tissue form without flagellae; both range from 6 to 20 mµ in size (McDougald and Reid 1978). There is also some evidence for a third form parasitizing gonads of larvae of the cecal nematode Heterakis gallinae; a 3 µm form has been observed in cecal worm eggs (BonDurant and Wakenell 1994). cost r ange and distribution Most, if not all, gallinaceous birds are susceptible hosts (Cole and Friend 1999), including chukar (Alectoris chukar), pheasant (Phasianus colchicus), quail (Order Galliformes), grouse (Family Phasianidae), wild turkeys (Meleagris gallopavo), jungle fowl (Gallus gallus), guinea fowl (Family Numididae), and peafowl (Pavo spp.) (BonDurant and Wakenell 1994). Histomonas meleagridis essentially has a worldwide distribution (Cole and Friend 1999). life cycle and variations Histomonads multiply by binary fission. The H. meleagridis life cycle is unique in that it is one of the rare examples of a helminth-borne life history pattern; in addition, earthworms typically are used as paratenic hosts (Cole and Friend 1999). Histomonas meleagridis uses the cecal nematode Heterakis gallinae as a vector for entry into the avian hosts (BonDurant and Wakenell 1994). Although histomonads can be passed directly under optimal circumstances, H. meleagridis quickly loses viability upon cooling outside the host or inside a dead host; thus, development of some resting stage inside the nematode eggs appears essential for survival of H. gallinae, and this nematode is considered critical to the natural transmission of H. meleagridis (BonDurant and Wakenell 1994). While the histomonads are carried inside the nematode egg (Tyzzer 1934), it still is not clear how the parasite becomes incorporated into the egg and the nature of the form in which it may exist for several years (BonDurant and Wakenell 1994). Both the male and female H. gallinae become infected with Histomonas; the histomonads then become incorporated into the eggs of the female (Lee 1970, McDougald 1991). There is some evidence that 192

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the male Heterakis may become infected first and transfer the infection to the female during copulation (Springer et al. 1969, McDougald 1991). Unembryonated Heterakis eggs are not infective; also, the processes of hatching or physical destruction of the egg seem important in release of infective histomonads (BonDurant and Wakenell 1994). reservoirs and transmission Heterakis gallinae, the nematode involved, is an ascarid with a direct life cycle, but one that also commonly uses earthworms as paratenic hosts. After the histomonad eggs are shed into the environment, the embryos develop to second stage larvae (L 2). On being ingested, the nematode larvae develop to adulthood in the ceca of their avian hosts. The histomonads accompanying the egg also infect the birds and may cause pathology before infecting the new generation of helminth ova produced by infecting H. gallinae. In addition, earthworms often ingest fecally contaminated soil containing H. gallinae eggs infected with histomonads (Kemp and Franson 1975, BonDurant and Wakenell 1994). Thus earthworms concentrate the nematode eggs and enhance transmission; earthworms also take the eggs to new sites as they migrate in the soil and help preserve the eggs from fungi and predation by other soil invertebrates (Lund et al. 1966). After earthworms ingest Heterakis gallinarum eggs, L 2 larvae hatch out and invade the earthworm tissues; thus the earthworm concentrates the H. gallinarum larvae as well as the histomonads present (Lund et al. 1966). Susceptible birds consuming these earthworms become infected by both the Heterakis gallinarum nematode and Histomonas meleagridis. Thus, while earthworms are not required for the nematode or histomonad life cycle, they are considered to play a critical role in enhancing natural transmission of each (BonDurant and Wakenell 1994). clinical effects and diagnosis  The disease histomoniasis also is called blackhead because infections often cause cyanosis, a blue or black appearance on the skin of the head on turkeys and some other birds due to an excess

of reduced hemoglobin in the blood (Cole and Friend 1999). There are no clinical signs specific to histomoniasis. Infected wild turkeys often are listless, amd have ruffled feathers and drooped wings. Feces are often sulfur-yellow in color and, combined with other field signs, are highly suggestive of histomoniasis (Cole and Friend 1999). The term enterohepatitis also is applied, reflecting their invasion of the cecum and, in more serious cases, causing multifocal necrosis in the liver (BonDurant and Wakenell 1994) Turkeys, grouse, and partridge develop severe disease and have mortality rates that can exceed 75% of the birds infected (Cole and Friend 1999). In contrast, histomoniasis is less severe in gray partridge (Perdix perdix) and northern bobwhites (Colinus virginianus) (Cole and Friend 1999). Pathologic effects of Histomonas appear to involve an interaction with fecal bacteria. Pathologic effects are fully present only with the full complement of fecal bacteria and are reduced in the presence of single species or limited combinations of fecal bacteria (Franker and Doll 1964, Springer et al. 1970). Histomonas does not colonize germ-free animals (Franker and Doll 1964, Kemp 1974). A presumptive diagnosis can be based on the presence of characteristic gross pathology and history. Laboratory confirmation generally involves microscopic examination for living flagellates with one or two flagellae, an axostylelike projection, and a 6–20 mµ diameter size; these organisms also can be cultured in Dwyer’s medium (BonDurant and Wakenell 1994). population effects  Turkeys are highly susceptible to Histomonas meleagridis infections (histomoniasis, blackhead). The parasite can cause catastrophic losses in both wild and domestic turkey populations (Cole and Friend 1999). special problems One interesting case study involving Histomonas in wildlife occurred in the southeastern United States; it illustrates the complexity of parasitic interactions with their hosts. In the 1960s, there was evidence that wild turkey populations in the southeastern United States were declining; it was proposed that the

turkeys were suffering from histomoniasis transmitted from bobwhites (Colinus virginianus) infected with Histomonas-infected Heterakis gallinarum eggs (Kellogg and Reid 1970). But the bobwhites carried Heterakis bonasae rather than H. gallinarum, and only H. gallinarum is known to carry Histomonas (Kellogg and Reid 1970). Thus bobwhites seemed relatively unimportant in contaminating soil with Histomonas-bearing Heterakis eggs (Lund and Chute 1971). However, Indian red jungle fowl (Gallus gallus murghi) previously released in the range of eastern wild turkey (Meleagris gallopavo silvestris) carried Histomonas spp. and were considered a more likely reservoir host for histomoniasis (Kellogg et al. 1972). control A variety of anthelminthics are used in commercial operations (BonDurant and Wakenell 1994). Because of the high susceptibility of turkeys compared to chickens, wild and domestic turkeys should not have exposure to habitat frequented by chickens; the development in the turkey industry of confinement rearing has reduced exposure to histomonads (BonDurant and Wakenell 1994).

Opportunistic Soil and Water Organisms acanthamoeba spp. c aus at i v e agen t (cl a ssific at ion, morphology)  Acanthamoeba is a genus of amebae that generally are free living, but include several species of that can be pathogenic to wildlife, domestic animals, and humans (Visvesvara 1999, John 2001). The genus derives its name from the distinctive feature of its acanthopodia (Gr. acanth: spine or thorn), tapering spike-like pseudopodia, and is classified in the Amoebozoa (Adl et al. 2005). Trophozoites average about 24 to 56 µm in length (John 1993). Species often have been named on the basis of variation in cyst morphology among Acanthamoeba (John 1993), but are not easily distinguished (Costas and Griffiths 1986, John 1993). host range and distribution  Acanthamoeba are distributed worldwide in soil and water (John 1993). Although generally kingdom protista

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free-living amebae, they are able to cause disease and even death in numerous animals, including humans (John 2001). life cycle and variations  The life cycle is simple and includes a motile feeding trophozoite (ameba stage) undergoing binary fission and a resting cyst stage (John 2001). reservoirs and tr ansmission Under most conditions, Acanthamoeba spp. are freeliving phagotrophs in freshwater ponds, rivers, lakes, soil, sewage, and sludge, feeding mainly on bacteria (Ma et al. 1990, Visvesvara and Stehr-Green 1990, John 2001). They occasionally also have been isolated from brackish water and seawater (Sawyer et al. 1976). There is no evidence for an animal reservoir or carrier host for Acanthamoeba (John 1993). Infections result from environmental exposure; animal-to-animal transmission is unknown (John 2001). Some infections are associated with aquatic exposure (John 2001). In other cases, infections may occur through the lower respiratory tract or through ulcers of the skin or mucosa, or direct contact with the eye (John 1993). clinical effects and diagnosis  Acanthamoeba infections can involve the central nervous system, eye, and other organs; invasion of the central nervous system appears to be by the circulation, with the initial focus of amebic infection occurring elsewhere in the body (John 2001). Syndromes can include an amebic meningoencephalitis, granulomatous amebic encephalitis, and keratitis (John 1993). Both the trophozoite and cyst stages can occur in tissues of infected animals (John 2001). Primary amebic meningoencephalitis usually is fatal in humans (John 1993), and generally is diagnosed by microscopic detection of living or stained amebae in cerebrospinal fluid or by culture (John 2001). Granulomatous amebic encephalitis is diagnosed by microscopic detection of trophozoites in cerebrospinal fluid or detection of trophozoites or cysts in brain tissue. Corneal infections can be diagnosed by microscopic detection of amebae from corneal scrapings or histologic examination of corneal tissue (John 2001). 194

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Acanthamoeba can be difficult to culture from infected animals (John 2001). Species of Acanthamoeba are diagnosed by analysis of isoenzyme patterns with electrophoresis and isoelectric focusing (Daggett et al. 1985), restriction enzyme analysis of mitochondrial DNA (Byers et al. 1983, Costas et al. 1983), as well as restriction endonuclease digestion of whole-cell DNA (McLaughlin et al. 1988). In general, there is not a good correlation between pathogenicity and species of Acanthamoeba (John 1993). However, there are genetic markers to help distinguish pathogenic from nonpathogenic strains of Acanthamoeba (Howe et al. 1997). population effects Although numerous wild and domestic animals are susceptible to infection, there are no reported population effects of any significance (John 2001). special problems  Acanthamoeba spp. can serve as host for a number of pathogenic agents, and enhance their survival in soil or water. The capacity for some bacteria not normally considered soil and water organisms to survive in the environment for extended periods is influenced by their ability to successfully infect free-living Acanthamoeba. After being ingested by amebae, the bacteria grow, multiply, and eventually lyse the amebae and are released back into the environment. In addition to serving as a host for the soil saprophyte Listeria monocytogenes (Ly and Müller 1990), this unique relationship has been linked with extended environmental survival for Francisella tularensis (Abd et al. 2003), Salmonella enterica (Tezcan-Merdol et al. 2004), Chlamydia pneumoniae (Essig et al. 1997), Pasteurella multocida (Hundt and Ruffolo 2005), Mycobacterium avium (Cirillo et al. 1997, Steinert et al. 1998), and Mycobacterium leprae (Jadin 1975), among others. Thus, their extended capacity to survive in the environment by parasitizing amebae complicates the development of strategies to control these bacteria. There also is evidence that Acanthamoeba can serve as suitable hosts for echoviruses and polioviruses (Danes and Červa 1981). control All work with prevention and control has been directed toward human cases.

There is no known satisfactory treatment for primary or granulomatous amebic meningoencephalitis (John 2001). There are several treatments for corneal infections, including topical treatments, epithelial debridement, and systemic ketoconazole (John 2001), as well as protective treatment of contact lenses (Hughes et al. 2003). No work has been reported with wildlife or domestic animals. Literature Cited Abd, H., T. Johansson, I. Golovliov, G. Sandström, and M. Forsman. 2003. Survival and growth of Francisella tularensis in Acanthamoeba castellanii. Applied and Environmental Microbiology 69:600–606. Acha, P. N., and B. Szyfres. 2003. Zoonoses and communicable diseases common to man and animals. Vol. III: Parasitoses. 3rd ed. Pan American Health Organization. Report No. 580. Adl, S. M., A. G. B. Simpson, M. A. Farmer, R. A. Andersen, O. R. Andersen, J. R. Barta, S. S. Bowser, G. Brugerolle, R. A. Fensome, S. Fredericq, T. Y. James, S. Karpov, P. Kugrens, J. Krug, C. E. Lane, L. A. Lewis, J. Lodge, D. H. Lynn, D. G. Mann, R. M. McCourt, L. Mendoza, Ø. Moestrup, S. E. MozleyStandridge, T. A. Nerad, C. A. Shearer, A. V. Smirnov, F. W. Spiegel, and M. F. J. R. Taylor. 2005. The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. Journal of Eukaryotic Microbiology 52:399–451. Allan, S. A. 2001. Biting flies (Class Insecta: Order Diptera). Pp. 18–45 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Allsopp, R. 1984. Control of tsetse flies (Diptera: Glossinidae) using insecticides: A review and future prospects. Bulletin of Entomological Research 74:1–23. Arkush, K. D., M. A. Miller, C. M. Leutenegger, I. A. Gardner, A. E. Packham, A. R. Heckeroth, A. M. Tenter, B. C. Barr, and P. A. Conrad. 2003. Molecular and bioassay-based detection of Toxoplasma gondii oocyst uptake by mussels (Mytilus galloprovincialis). International Journal for Parasitology 33:1087–1097. Ashcroft, M. T. 1959. The importance of African wild animals as reservoirs of trypanosomes. East African Medical Journal 36:289–297. Atkinson, C. T. 1999. Hemosporidiosis. Pp. 193–199 in M. Friend and J. C. Franson (editors), Field



manual of wildlife diseases: General field procedures and diseases of birds. U.S. Geological Survey, Biological Resources Division, Washington, DC. Atkinson, C. T., R. J. Dusek, and J. K. Lease. 2001a. Serological responses and immunity to superinfection with avian malaria in experimentallyinfected Hawaii amakihi. Journal of Wildlife Diseases 37:20–27. Atkinson, C. T., R. J. Dusek, K. L. Woods, and W. M. Iko. 2000. Pathogenicity of avian malaria in experimentally-infected Hawaii amakihi. Journal of Wildlife Diseases 36:197–204. Atkinson, C. T., D. J. Forrester, and E. C. Greiner. 1988. Pathogenicity of Haemoproteus meleagridis (Protozoa: Haemoproteidae) in experimentally infected domestic turkeys. Journal of Parasitology 74:228–239. Atkinson, C. T., J. K. Lease, B. M. Drake, and N. P. Shema. 2001b. Pathogenicity, serological responses, and diagnosis of experimental and natural malarial infections in native Hawaiian thrushes. The Condor 103:209–218. Atkinson, C. T., and C. van Riper III. 1991. Pathogenicity and epizootiology of avian haematozoa: Plasmodium, Leucocytozoon, and Haemoproteus. Pp. 19–48 in J. E. Loye and M. Zuk (editors), Bird–parasite interactions. Oxford University Press, Oxford, UK. Atkinson, C. T., K. L. Woods, R. J. Dusek, L. S. Sileo, and W. M. Iko. 1995. Wildlife disease and conservation in Hawaii: Pathogenicity of avian malaria (Plasmodium relictum) in experimentally infected iiwi (Vestiaria coccinea). Parasitology 111(Suppl.):S59–S69. Baer, J. G. 1951. Ecology of animal parasites. University of Illinois Press, Urbana, IL. Baer, J. G. 1971. Animal parasites. World University Library, McGraw-Hill Book Company, New York. Baker, J. R. 1969. Trypanosomes of wild mammals in the neighbourhood of the Serengeti National Park. Pp. 147–155 in A. McDiarmid (editor), Diseases in free-living wild animals. Symposia of the Zoological Society of London No. 24. Academic Press, London, UK. Baker, J. R. 1976. Biology of trypanosomes of birds. Pp. 131–174 in W. H. R. Lumsden and D. A. Evans (editors), Biology of the Kinetoplastida 1. Academic Press, London, UK. Baker, J. R., R. Sachs, and I. Laufer. 1967. Trypanosomes of wild mammals in an area northwest of Serengeti National Park, Tanzania. Zeitschrift für Tropenmedizin und Parasitologie 18:280–284. Bardsley, J. E., and R. Harmsen. 1973. The trypanosomes of Anura. Advances in Parasitology 11:1–63.

kingdom protista

195

Barnett, S. F., and D. W. Brocklesby. 1969. Some piroplasms of wild mammals. Pp. 159–176 in A. McDiarmid (editor), Diseases in free-living wild mammals. Symposia of the Zoological Society of London No. 24. Academic Press, London, UK. Barta, J. R., D. S. Martin, P. A. Liberator, M. Dashkevicz, J. W. Anderson, S. D. Feighner, A. Elbrecht, A. Perkins-Barrow, M. C. Jenkins, H. H. Danforth, M. D. Ruff, and H. Profous-Juchelka. 1997. Phylogenetic relationships among eight Eimeria species infecting domestic fowl inferred using complete small subunit ribosomal DNA sequences. Journal of Parasitology 83:262–271. Belova, L. M., and M. V. Krylov. 2003. The problems of evolution and phylogeny of coccidia with the eimerioran-type and isosporan-type of oocysts. Parazitologiya 37:324–332. Bennett, G. F., M. A. Bishop, and M. A. Peirce. 1993. Checklist of the avian species of Plasmodium Marchiafava & Celli, 1885 (Apicomplexa) and their distribution by avian family and Wallacean life zones. Systematic Parasitology 26:171–179. Bennett, G. F., J. R. Caines, and M. A. Bishop. 1988. Influence of blood parasites on the body mass of passeriform birds. Journal of Wildlife Diseases 24:339–343. Bennett, G. F., M. A. Peirce, and R. A. Earlé. 1994. An annotated checklist of the valid avian species of Haemoproteus, Leucocytozoon (Apicomplexa: Haemosporida) and Hepatozoon (Apicomplexa: Haemogregarinidae). Systematic Parasitology 29:61–73. Bennett, G. F., M. Whiteway, and C. WoodworthLynas. 1982. A host-parasite catalogue of the avian haematozoa. Memorial University of Newfoundland Occasional Papers in Biology, No. 5, St. John’s, Newfoundland, Canada. Bert, E. 1619. An approved treatise of hawkes and hawking. Richard Moore Quaritch, London, England. Bertram, B. C. R. 1973. Sleeping sickness survey in the Serengeti area (Tanzania) 1971. III: Discussion of the relevance of the trypanosome survey to the biology of large mammals in the Serengeti. Acta Tropica 30:36–48. Boal, C. W., and R. W. Mannan. 1999. Comparative breeding ecology of Cooper’s hawks in urban and exurban areas of southeastern Arizona. Journal of Wildlife Management 63:77–84. Boal, C. W., R. W. Mannan, and K. S. Hudelson. 1998. Trichomoniasis in Cooper’s hawks from Arizona. Journal of Wildlife Diseases 34:590–593. BonDurant, R. H., and B. M. Honigberg. 1994. Trichomonads of veterinary importance.

196

kingdom protista

Pp. 111–188 in J. P. Kreier (editor). Parasitic protozoa, Vol. 9. Academic Press, San Diego, CA. BonDurant, R. H., and P. S. Wakenell. 1994. Histomonas meleagridis and relatives. Pp. 189–206 in J. P. Kreier (editor), Parasitic protozoa, Vol. 9. Academic Press, San Diego, CA. Boothroyd, J. C. 1985. Antigenic variation in African trypanosomes. Annual Review of Microbiology 39:475–502. Borst, P. 1986. Discontinuous transcription and antigenic variation in trypanosomes. Annual Review of Biochemistry 55:701–732. Bruckner, D. A., and J. A. Labarca. 1999. Leishmania and Trypanosoma. Pp. 1365–1373 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Bunbury, N., D. Bell, C. Jones, A. Greenwood, and P. Hunter. 2005. Comparison of the InPouch TF culture system and wet-mount microscopy for diagnosis of Trichomonas gallinae infections in the pink pigeon Columba mayeri. Journal of Clinical Microbiology 43:1005–1006. Burrows, R. G., and M. A. Swerdlow. 1956. Enterobius vermicularis as a probable vector of Dientamoeba fragilis. American Journal of Tropical Medicine and Hygiene 5:258–265. Bush, A. O., J. C. Fernández, G. W. Esch, and J. R. Seed. 2001. Parasitism: The diversity and ecology of animal parasites. Cambridge University Press, Cambridge, UK. Byers, T. J., S. A. Bogler, and L. L. Burianek. 1983. Analysis of mitochondrial DNA variation as an approach to systematic relationships in the genus Acanthamoeba. Journal of Protozoology 30:198–203. Cabezón, O., A. R. Resendes, M. Domingo, J. A. Raga, C. Agustí, F. Alegre, J. L. Mons, J. P. Dubey, and S. Almería. 2004. Seroprevalence of Toxoplasma gondii antibodies in wild dolphins from the Spanish Mediterranean coast. Journal of Parasitology 90:643–644. Cannell, B. L., K. V. Krasnec, K. Campbell, H. I. Jones, R. D. Miller, and N. Stephens. 2013. The pathology and pathogenicity of a novel Haemoproteus spp. infection in wild little penguins (Eudyptula minor). Veterinary Parasitology. doi: 10.1016/j. vetpar.2013.1004.1025. Carmichael, I. H., and E. Hobday. 1975. Blood parasites of some wild Bovidae in Botswana. Onderstepoort Journal of Veterinary Research 42:55–62. Carmichael, J. 1933. The virus of rinderpest and its relation to Glossina morsitans (Weston). Bulletin of Entomological Research 24:337–342.

Cawthorn, R. J. 1993. Cyst-forming coccidia of raptors: Significant pathogens or not? Pp. 14–20 in P. T. Redig, J. E. Cooper, J. D. Remple, D. B. Hunter, and T. Hahn (editors), Raptor biomedicine. University of Minnesota Press, Minneapolis, MN. Cedillos, R. A. 1975. Chagas’ disease in El Salvador. Bulletin of the Pan American Health Organization 9:135–141. Chandler, A. S., and C. P. Read. 1961. Introduction to parasitology. 10th ed. John Wiley and Sons, New York. Cirillo, J. D., S. Falkow, L. S. Tompkins, and L. E. Bermudez. 1997. Interaction of Mycobacterium avium with environmental amoebae enhances virulence. Infection and Immunity 65:3759–3767. Clarke, J. E. 1964. Game elimination as a means of tsetse control with special reference to host preferences. The Puku 2:67–75. Cockbill, G. F. 1967. The history and significance of trypanosomiasis problems in Rhodesia. Proceedings and Transactions of the Rhodesia Scientific Association 52:7–15. Cole, R. A. 1999a. Renal coccidiosis. Pp. 215–218 in M. Friend and J. C. Franson (editors), Field manual of wildlife diseases: General field procedures and diseases of birds. Biological Resources Division Information and Technology Report 1999-001. U.S. Geological Survey, Washington, DC. Cole, R. A. 1999b. Trichomoniasis. Pp. 201–206 in M. Friend and J. C. Franson (editors), Biological Resources Division Information and Technology Report 1999-001. U.S. Geological Survey, Washington, DC. Cole, R. A., and M. Friend. 1999. Miscellaneous parasitic diseases. Pp. 249–258 in M. Friend and J. C. Franson (editors), Field manual of wildlife diseases: General field procedures and diseases of birds. Biological Resources Division Information and Technology Report 1999-001. U.S. Geological Survey Washington, DC. Cole, R. A., D. S. Lindsay, D. K. Howe, C. L. Roderick, J. P. Dubey, N. J. Thomas, and L. A. Baeten. 2000. Biological and molecular characterizations of Toxoplasma gondii strains obtained from southern sea otters (Enhydra lutris nereis). Journal of Parasitology 86:526–530. Collins, E., and M. Aikawa. 1993. Plasmodia of nonhuman primates. Pp. 105–134 in J. P. Kreier (editor), Parasitic protozoa, Vol. 5. Academic Press, San Diego, CA. Comes, A. M., J. F. Humbert, J. Cabaret, and L. Élard. 1996. Using molecular tools for diagnosis in veterinary parasitology. Veterinary Research 27:333–342.



Conrad, P. A., M. A. Miller, C. Kreuder, E. R. James, J. Mazet, H. Dabritz, D. A. Jessup, F. Gulland, and M. E. Grigg. 2005. Transmission of Toxoplasma: Clues from the study of sea otters as sentinels of Toxoplasma gondii flow into the marine environment. International Journal for Parasitology 35:1155–1168. Conti, J. A. 1993. Diseases, parasites and contaminants. Pp. 205–224 in T. S. Baskett, M. W. Sayre, R. E. Tomlinson, and R. E. Mirarchi (editors), Ecology and management of the mourning dove. Stackpole Books, Harrisburg, PA. Cook, R. S. 1971a. Haemoproteus. Pp. 300–308 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Cook, R. S. 1971b. Leucocytozoon Danilewsky 1890. Pp. 291–299 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Cooper, J. E. 2002. Birds of prey: Health and disease. 3rd ed. Blackwell Science, Oxford, UK. Cooper, J. E., and S. J. Petty. 1988. Trichomoniasis in free-living goshawks (Accipiter gentilis gentilis) from Great Britain. Journal of Wildlife Diseases 24:80–87. Costas, M., and A. J. Griffiths. 1986. Physiological characterizations of Acanthamoeba strains. Journal of Protozoology 32:604–607. Costas, M., S. W. Edwards, D. Lloyd, A. J. Griffiths, and G. Turner. 1983. Restriction enzyme analysis of mitochondrial DNA of members of the genus Acanthamoeba as an aid in taxonomy. FEMS (Federation of European Microbiology Societies) Microbiology Letters 17:231–234. Cover, A. J., W. M. Harmon, and M. W. Thomas. 1994. A new method for the diagnosis of Trichomonas gallinae infection by culture. Journal of Wildlife Diseases 30:457–457. Cox, F. E. G. 1991. Systematics of parasitic protozoa. Pp. 55–80 in J. P. Kreier and J. R. Baker (editors), Parasitic protozoa. Academic Press, San Diego, CA. Cox, F. E. G. 1993. Plasmodia of rodents. Pp. 49–105 in J. P. Kreier (editor), Parasitic protozoa, Vol. 5. Academic Press, San Diego, CA. Craig, T. M. 2001. Hepatozoon spp. and hepatozoonosis. Pp. 462–468 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Critcher, S. 1950. Renal coccidiosis in Pea Island Canada geese. Wildlife of North Carolina 14:14–15.

kingdom protista

197

Cruikshank, J. J., D. M. Haines, N. C. Palmer, and D. J. S. Aubin. 1990. Cysts of a Toxoplasma-like organism in an Atlantic bottlenose dolphin. Canadian Veterinary Journal 31:213–215. Daggett, P.-M., D. Lipscomb, T. K. Sawyer, and T. A. Nerad. 1985. A molecular approach to the phylogeny of Acanthamoeba. BioSystems 18:399–405. Danes, L., and L. Červa. 1981. Survival of polioviruses and echoviruses in Acanthamoeba castellanii cultivated in vitro. Journal of Hygiene, Epidemiology, Microbiology and Immunology 25:169–174. Donelson, J. E., and M. J. Turner. 1985. How the trypanosome changes its coat. Scientific American 252:44–51. Dubey, J. P. 1993. Toxoplasma, Neospora, Sarcocystis, and other tissue cyst-forming coccidia of humans and animals. Pp. 1–158 in J. P. Kreier (editor), Parasitic protozoa, Vol. 6. Academic Press, San Diego, CA. Dubey, J. P. 2001. Oocyst shedding by cats fed isolated bradyzoites and comparison of infectivity of bradyzoites of the VEG strain Toxoplasma gondii to cats and mice. Journal of Parasitology 87:215–219. Dubey, J. P. 2004. Toxoplasmosis: A waterborne zoonosis. Veterinary Parasitology 126:57–72. Dubey, J. P., and C. P. Beattie. 1988. Toxoplasmosis of animals and man. CRC Press, Boca Raton, FL. Dubey, J. P., and J. L. Carpenter. 1993a. Histologically confirmed clinical toxoplasmosis in cats: 100 cases (1952–1990). Journal of the American Veterinary Medical Association 203:1556–1566. Dubey, J. P., and J. L. Carpenter. 1993b. Neonatal toxoplasmosis in littermate cats. Journal of the American Veterinary Medical Association 203:1546–1549. Dubey, J. P., P. A. Fair, G. D. Bossart, D. Hill, R. Fayer, C. Sreekumar, O. C. H. Kwok, and P. Thulliez. 2005. A comparison of several serologic tests to detect antibodies of Toxoplasma gondii in naturally exposed bottlenose dolphins (Tursiops truncatus). Journal of Parasitology 91:1074–1081. Dubey, J. P., and J. K. Frenkel. 1972. Cyst-induced toxoplasmosis in cats. Journal of Protozoology 19:155–177. Dubey, J. P., R. P. Lipscomb, and M. Mense. 2004. Toxoplasmosis in an elephant seal (Mirounga angustirostris). Journal of Parasitology 90:410–411. Dubey, J. P., N. L. Miller, and D. K. Frenkel. 1970. Toxoplasma gondii life cycle in cats. Journal of the American Veterinary Medical Association 157:1767–1770.

198

kingdom protista

Dubey, J. P., and K. Odening. 2001. Toxoplasmosis and related infections. Pp. 478–519 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Dubey, J. P., R. Zarnke, N. J. Thomas, S. K. Wong, W. V. Bonn, M. Briggs, J. W. Davis, R. Ewing, M. Mense, O. C. H. Kwok, S. Romand, and P. Thulliez. 2003. Toxoplasma gondii, Neospora caninum, Sarcocystis neurona, and Sarcocystis canis-like infections in marine mammals. Veterinary Parasitology 116:275–296. Duke, H. L. 1919. An enquiry into the relations of Glossina morsitans and ungulate game, with special reference to rinderpest. Bulletin of Entomological Research 10:7–20. Dunsmore, J. D. 1971. A study of the biology of the wild rabbit in climatically different regions in eastern Australia. IV: The rabbit in the south coastal region of New South Wales, an area in which parasites appear to exert a populationregulating effect. Australian Journal of Zoology 19:355–370. Duszynski, D. W., and S. J. Upton. 2001. Cyclospora, Eimeria, Isospora, and Cryptosporidium spp. Pp. 416–459 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Du Toit, R. 1954. Trypanosomiasis in Zululand and the control of tsetse flies by chemical means. Onderstepoort Journal of Veterinary Research 26:317–387. Eichinger, D. 2001. A role for a galactose lectin and its ligands during encystment of Entamoeba. Journal of Eukaryotic Microbiology 48:17–21. Essig, A., M. Heinemann, U. Simnacher, and R. Marre. 1997. Infection of Acanthamoeba castellanii by Chlamydia pneumoniae. Applied and Environmental Microbiology 63:1396–1399. Estes, J. A., B. B. Hatfield, K. Ralls, and J. Ames. 2003. Causes of mortality in California sea otters during periods of population growth and decline. Marine Mammal Science 19:198–216. Farr, M. M. 1954. Renal coccidiosis of Canada geese. Journal of Parasitology 40(Suppl.):46. Fayer, R., J. P. Dubey, and D. S. Lindsay. 2004. Zoonotic protozoa: From land to sea. Trends in Parasitology 20:531–536. Ford, J. 1971. The role of trypanosomiasis in African Ecology. Clarendon Press, Oxford, UK. Frank, W. 1976. Parasitologie. Verlag Eugen Ulmer, Stuttgart, Germany. Franker, C. K., and J. P. Doll. 1964. Experimental histomoniasis in gnotobiotic turkeys. II: Effects

of some cecal bacteria on pathogenesis. Journal of Parasitology 50:636–640. Fraser, C. M., and A. Mays. 1986. The Merck veterinary manual. 6th ed. Merck & Co., Inc., Rahway, NJ. Frenkel, J. K., A. Ruiz, and M. Chinchilla. 1975. Soil survival of Toxoplasma oocysts in Kansas and Costa Rica. American Journal of Tropical Medicine and Hygiene 24:439–443. Friend, M., and J. C. Franson. 1999. Intestinal coccidiosis. Pp. 207–213 in M. Friend and J. C. Franson (editors), Field manual of wildlife diseases: General field procedures and diseases of birds. Biological Resources Division, U.S. Geological Survey, Washington, DC. Gajadhar, A. A., L. Measures, L. B. Forbes, C. Kapel, and J. P. Dubey. 2004. Experimental Toxoplasma gondii infection in grey seals (Halichoerus grypus). Journal of Parasitology 90:255–259. Garcia, L. S., and D. A. Bruckner. 1997. Diagnostic medical parasitology. 3rd ed. American Society for Microbiology, Washington, DC. Garnham, P. C. C. 1966. Malaria parasites and other Haemosporidia. Blackwell Scientific, Oxford, UK. Geigy, R., and M. Kauffman. 1973. Sleeping sickness survey in the Serengeti area (Tanzania) 1971. I: Examination of large mammals for trypanosomiasis. Acta Tropica 30:12–23. Geigy, R., M. Kauffman, and R. Beglinger. 1967. A survey of wild animals as potential reservoirs of trypanosomiasis in the Ulanga District (Tanzania). Acta Tropica 24:97–108. Gerber, L. R., M. T. Tinker, D. F. Doak, and J. A. Estes. 2004. Mortality sensitivity in life-stage simulation analysis: A case study of southern sea otters. Ecological Applications 14:1554–1565. Glover, P. E. 1965. A review of recent knowledge on the relationship between the tsetse fly and its vertebrates hosts. Fauna Preservation Society, London, UK. Graham, P. 1964. Destruction of birds and other wildlife by dieldrex spraying against tsetse fly in Bechuanaland. Arnoldia (Rhodesia) 1:1–4. Graybill, H. W., and T. Smith. 1920. Production of fatal blackhead in turkeys by feeding embryonated eggs of Heterakis papillosa. Journal of Experimental Medicine 31:647–655. Grigg, M. E., S. Bonnefoy, A. B. Hehl, Y. Suzuki, and J. C. Boothroyd. 2001. Success and virulence in Toxoplasma as the result of sexual recombination between two distinct ancestries. Science 294:161–165. Gyening, K. O. 1990. Organizational perceptions and current approaches to trypanosomiasis



problems undertaken by FAO: Consultation on trypanosomiasis research and programmes in Africa, 30 April–4 May 1990. International Laboratory for Research on Animals Diseases, Nairobi, Kenya. Gylstorff, I., and F. Grimm. 1987. Vogelkrankheiten. Verlag Eugen Ulmer, Stuttgart, Germany. Hanson, H. C., N. D. Levine, and V. Ivens. 1957. Coccidia (Protozoa: Eimeriidae) of North American wild geese and swans. Canadian Journal of Zoology 35:715–733. Harmon, W. M., W. A. Clark, A. C. Hawbecker, and M. Stafford. 1987. Trichomonas gallinae in columbiform birds from the Galapagos Islands. Journal of Wildlife Diseases 23:492–494. Heisch, R. G. 1963. Presence of trypanosomes in bush babies after eating infected rats. Nature (London) 169:118. Herman, C. M. 1969. Blood protozoa of free-living birds. In A. McDiarmid (editor), Diseases in freeliving wild animals. Symposia of the Zoological Society of London No. 24. Academic Press, London, UK. Hill, D. E., S. Chirukandoth, and J. P. Dubey. 2005. Biology and epidemiology of Toxoplasma gondii in man and animals. Animal Health Research Reviews 6:41–61. Hoare, C. A. 1966. The classification of mammalian trypanosomes. Ergebnisse der Mikrobiologie, Immunitätsforschung und experimentellen Therapie 39:43–57. Hoare, C. A., and F. G. Wallace. 1966. Developmental stages of trypanosomatid flagellates: A new terminology. Nature 212:1385–1386. Höfle, U., C. Gortazar, J. A. Ortíz, B. Knispel, and E. F. Kaleta. 2004. Outbreak of trichomoniasis in a woodpigeon (Columba palumbus) wintering roost. European Journal of Wildlife Research 50:73–77. Honnold, S. P., R. Braun, D. P. Scott, C. Sreekumar, and J. P. Dubey. 2005. Toxoplasmosis in a Hawaiian monk seal (Monachus schauinslandi). Journal of Parasitology 91:695–697. Howe, D. K., M. H. Vodkin, R. J. Novak, G. S. Visvesvara, and G. L. McLaughlin. 1997. Identification of two genetic markers that distinguish pathogenic and nonpathogenic strains of Acanthamoeba spp. Parasitology Research 83:345–348. Huff, C. G. 1965. Susceptibility of mosquitoes to avian malaria. Experimental Parasitology 16:107–132. Hughes, R., P. W. Andrew, and S. Kilvington. 2003. Enhanced killing of Acanthamoeba cysts with a plant peroxidase-hydrogen peroxide-halide

kingdom protista

199

antimicrobial system. Applied and Environmental Microbiology 69:2563–2567. Hundt, M. J., and C. G. Ruffolo. 2005. Interaction of Pasteurella multocida with free-living amoeba. Applied and Environmental Microbiology 71:5458–5464. Inskeep, W., 2nd, C. H. Gardiner, R. K. Harris, J. P. Dubey, and R. T. Goldston. 1990. Toxoplasmosis in Atlantic bottle-nosed dolphins (Tursiops truncatus). Journal of Wildlife Diseases 26:377–382. Jadin, J. B. 1975. Amibes “Limax” vecteurs possibles des Mycobactéries et de Mycobacterium leprae. Acta Leprologica 59–60:57–67. Jakob, W., and H. H. Wesemeier. 1995. Intestinal inflammation associated with flagellates in snakes. Journal of Comparative Pathology 112:417–421. Jardine, J. E., and J. P. Dubey. 2002. Congenital toxoplasmosis in a Indo-Pacific bottlenose dolphin (Tursiops aduncus). Journal of Parasitology 88:197–199. Jawara, A. S. D. K. 1990. Animal disease as a factor limiting economic development in Africa. Cornell Veterinarian 80:17–25. Jessup, D. A., M. Miller, J. Ames, M. Harris, C. Kreuder, P. A. Conrad, and J. A. K. Mazet. 2004. Southern sea otter as a sentinel of marine ecosystem health. EcoHealth 1:239–245. Jindrichova, J., K. Kramarova, B. Rosicky, J. Jira, and A. Simko. 1975. The cat as a possible source of Toxoplasma infection for man. Folia Parasitologica 22:309–315. John, D. T. 1993. Opportunistically pathogenic freeliving amebae, Vol. 3. Pp. 143–246 in J. P. Kreier and J. R. Baker (editors), Parasitic protozoa. Academic Press, San Diego, CA. John, D. T. 2001. Opportunistic amoebae. Pp. 460–462 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Jordan, A. M. 1974. Recent developments in the ecology and methods of control of tsetse flies (Glossina spp.) (Dipt. Glossinidae): A review. Bulletin of Entomological Research 63:361–399. Julian, R. J., and D. E. Galt. 1980. Mortality in muscovy ducks (Cairina moschata) caused by Haemoproteus infection. Journal of Wildlife Diseases 16:39–44. Kakoma, İ., and H. Mehlhorn. 1994. Babesia of domestic animals. Pp. 141–216 in J. P. Kreier (editor), Parasitic protozoa, Vol. 7. Academic Press, San Diego, CA. Keeton, W. T., and J. L. Gould. 1993. Biological science, Vol. 2. 5th ed. W. W. Norton & Company, New York.

200

kingdom protista

Kellogg, F. E., T. H. Eleazer, and T. R. Colvin. 1972. Transmission of blackhead from jungle fowl to turkey. Proceedings of the Annual Conference of the Southeastern Association of Fish and Wildlife Agencies 32:378–379. Kellogg, F. E., and W. M. Reid. 1970. Bobwhites as possible reservoir hosts for blackhead in wild turkeys. Journal of Wildlife Management 34:155–159. Kemp, R. L. 1974. The failure of Histomonas meleagridis to establish in germ-free ceca in normal poults. Avian Diseases 18:452–455. Kemp, R. L., and J. C. Franson. 1975. Transmission of Histomonas meleagridis to domestic fowl by means of earthworms recovered from pheasant yard soil. Avian Diseases 19:741–744. Kimura, M., A. A. Dhondt, and I. J. Lovette. 2006. Phylogeographic structuring of Plasmodium lineages across the North American range of the house finch (Carpodacus mexicanus). Journal of Parasitology 92:1043–1049. Kocan, A. A. 2001a. Amebiasis, balantidiasis, and enteric trichomoniasis. Pp. 397–399 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Kocan, A. A. 2001b. Blood-inhabiting protozoan parasites. Pp. 520–524 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Kocan, A. A., and K. A. Waldrup. 2001. Piroplasms (Theileria spp., Cytauxzoon spp., and Babesia spp.). Pp. 524–536 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Kreuder, C., M. A. Miller, D. A. Jessup, L. J. Lowenstine, M. D. Harris, J. A. Ames, T. E. Carpenter, P. A. Conrad, and J. A. K. Mazet. 2003. Patterns of mortality in southern sea otters (Enhydra lutris nereis) from 1998–2001. Journal of Wildlife Diseases 39:495–509. Krone, O., R. Altenkamp, and N. Kenntner. 2005. Prevalence of Trichomonas gallinae in northern goshawks from the Berlin area of northeastern Germany. Journal of Wildlife Diseases 41:304–309. Kudo, R. R. 1966. Protozoology. Charles C. Thomas, New York. Kulda, J., and E. Nohýnková. 1994. Giardia in humans and animals. Pp. 225–422 in J. P. Kreier (editor), Parasitic protozoa, Vol. 10. Academic Press, San Diego, CA. Lainson, R. 1982. Leishmania parasites of mammals in relation to human health. Pp. 137–179 in

M. A. Edwards and U. McDonnell (editors), Animal disease in relation to animal conservation. Symposia of the Zoological Society of London No. 50. Academic Press, London, UK. Lambourn, D. M., S. J. Jeffries, and J. P. Dubey. 2001. Seroprevalence of Toxoplasma gondii in harbor seals (Phoca vitulina) in southern Puget Sound, Washington. Journal of Parasitology 87:1196–1197. Leber, A. L., and S. M. Novak. 1999. Intestinal and urogenital amebae, flagellates, and ciliates. Pp. 1391–1405 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Lee, D. H. 1970. The structure and development of Histomonas meleagridis (Mastigamoebidae: Protozoa) in the female reproductive tract and its intermediate host, Heterakis gallinarum (Nematoda). Parasitology 59:877–884. Lehmann, T., P. L. Marcet, D. H. Graham, E. R. Dahl, and J. P. Dubey. 2006. Globalization and the population structure of Toxoplasma gondii. Proceedings of the National Academy of Sciences of the United States of America 103:11423–11428. Leighton, F. A., and A. A. Gajadhar. 2001. Besnoitia spp. and besnoitiosis. Pp. 468–478 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Lindsay, D. S., M. V. Collins, S. M. Mitchell, R. A. Cole, G. J. Flick, C. N. Wetch, A. Lindquist, and J. P. Dubey. 2003. Sporulation and survival of Toxoplasma gondii oocysts in seawater. Journal of Eukaryotic Microbiology 50:687–688. Lindsay, D. S., M. V. Collins, S. M. Mitchell, C. N. Wetch, C. N. Wetch, A. C. Rosypal, G. J. Flick, A. M. Zajac, A. Lindquist, and J. P. Dubey. 2004. Survival of Toxoplasma gondii oocysts in eastern oysters (Crassostrea virginica). Journal of Parasitology 90:1054–1057. Lindsay, D. S., K. K. Phelps, S. A. Smith, G. Flick, S. S. Sumner, and J. P. Dubey. 2001a. Removal of Toxoplasma gondii oocysts from sea water by eastern oysters (Crassostrea virginica). Journal of Eukaryotic Microbiology 48:197s–198s. Lindsay, D. S., N. J. Thomas, A. C. Rosypal, and J. P. Dubey. 2001b. Dual Sarcocystis neurona and Toxoplasma gondii infection in a northern sea otter from Washington State, USA. Veterinary Parasitology 97:319–327. Lindsay, D. S., and K. S. Todd. 1993. Coccidia of mammals. Pp. 89–131 in J. P. Kreier (editor), Parasitic protozoa, Vol. 4. Academic Press, San Diego, CA.



Logan-Henfrey, L. L., P. R. Gardiner, and M. M. Mahmoud. 1992. Animal trypanosomiasis in sub-Sahara Africa. Pp. 157–276 in J. P. Kreier and J. R. Baker (editors), Parasitic protozoa, Vol. 2. Academic Press, San Diego, CA. Lom, J. 1979. Biology of the trypanosomes and trypanoplasms of fish. Pp. 269–337 in W. H. R. Lumsden and D. A. Evans (editors), Biology of the Kinetoplastida 2. Academic Press, London, UK. Long, P. F. 1993. Avian coccidiosis. Pp. 1–88 in J. P. Kreier (editor), Parasitic protozoa, Vol. 4. Academic Press, San Diego, CA. López-Antuñano, F. J., and G. A. Schmunis. 1993. Plasmodia of humans. Pp. 135–266 in J. P. Kreier (editor), Parasitic protozoa, Vol. 5. Academic Press, San Diego, CA. Lucius, R., and B. Loos-Frank. 1997. Parasitologie. Spektrum Akademischer Verlag GmbH, Heidelberg, Germany. Lund, E. E., and A. M. Chute. 1971. Bobwhite, Colinus virginianus, as host for Heterakis and Histomonas. Journal of Wildlife Diseases 7:70–75. Lund, E. E., E. E. Wehr, and D. J. Ellis. 1966. Earthworm transmission of Heterakis and Histomonas to turkeys and chickens. Journal of Parasitology 52:899–902. Ly, T. M., and H. E. Müller. 1990. Ingested Listeria monocytogenes survive and multiply in protozoa. Journal of Medical Microbiology 33:51–54. Ma, P., G. S. Visvesvara, A. J. Martinez, F. H. Theodore, P. M. Daggett, and T. K. Sawyer. 1990. Naegleria and Acanthamoeba infections: review. Review of Infectious Diseases 12:490–513. Mackenstedt, U., M. Gauer, H. Mehlhorn, E. Schein, and S. Hauschild. 1990. Sexual cycle of Babesia divergens confirmed by DNA measurements. Parasitology Research 76:199–206. Martinez, A. J., and G. S. Visvesvara. 1997. Freeliving, amphizoic and opportunistic amebas. Brain Pathology 7:583–598. Martinez-Paloma, A. 1993. Parasitic amoebas in the intestinal tract. Pp. 65–141 in J. P. Kreier and J. R. Baker (editors), Parasitic protozoa. Academic Press, New York. Massie, G., and M. Black. 2008. Unraveling the mystery of the kitty litter parasite in marine mammals. Science Daily, American Society for Microbiology (June 5). . Matthiessen, P. 1985. Contamination of wildlife with DDT insecticide residues in relation to tsetse fly control operations in Zimbabwe. Environmental Pollution Series B: Chemical and Physical 10:189–211.

kingdom protista

201

Matthiessen, P., and B. Douthwaite. 1985. The impact of tsetse fly control campaigns on African wildlife. Oryx 19:202–209. McDougald, L. R. 1991. Other protozoal diseases of the intestinal tract. Pp. 804–809 in B. W. Calnek, H. J. Barnes, C. W. Beard, W. M. Reid, and H. W. Yoder (editors), Diseases of poultry. Iowa State University Press, Ames, IA. McDougald, L. R., and W. M. Reid. 1978. Histomonas meleagridis and its relatives. Pp. 139–161 in J. P. Kreier (editor), Parasitic protozoa, Vol. II. Academic Press, New York. McLaughlin, G. L., F. H. Brandt, and G. S. Visvesvara. 1988. Restriction fragment length polymorphisms of the DNA of selected Naegleria and Acanthamoeba amebae. Journal of Clinical Microbiology 26:1655–1658. Measures, L. N., J. P. Dubey, P. Labelle, and D. Martineau. 2004. Seroprevalence of Toxoplasma gondii in Canadian pinnipeds. Journal of Wildlife Diseases 40:294–300. Meerovitch, E. 1958. Some biological requirements and host parasite relations of Entamoeba invadens. Canadian Journal of Zoology 36:513–523. Mehlhorn, H., W. Raether, E. Schein, M. Weber, and M. Uphoff. 1986. Licht und elektronenmikrskopische Untersuchungen zum Entwicklungszyklus und Einfluss von Pentamichin auf die Morphologie der intraerythrozytären Stadien von Babesia microti. Deutsche Tierärtzliche Wochenschrift 93:377–464. Mehlhorn, H., E. Schein, and J. S. Ahmed. 1994. Theileria. Pp. 217–304 in J. P. Kreier (editor), Parasitic protozoa, Vol. 7. Academic Press, San Diego, CA. Miller, M. A., I. A. Gardner, C. Kreuder, D. M. Paradies, K. R. Worcester, D. A. Jessup, E. Dodd, M. D. Harris, J. A. Ames, A. E. Packham, and P. A. Conrad. 2002a. Coastal freshwater runoff is a risk factor for Toxoplasma gondii infection of southern sea otters (Enhydra lutris nereis). International Journal for Parasitology 32:997–1006. Miller, M. A., I. A. Gardner, A. Packham, J. K. Mazet, K. D. Hanni, D. Jessup, J. Estes, R. Jameson, E. Dodd, B. C. Barr, L. J. Lowenstine, F. M. Gulland, and P. A. Conrad. 2002b. Evaluation of an indirect fluorescent antibody test (IFAT) for demonstration of antibodies to Toxoplasma gondii in the sea otter (Enhydra lutris). Journal of Parasitology 88:594–599. Miller, M. A., M. E. Grigg, C. Kreuder, E. R. James, A. C. Melli, P. R. Crosbie, D. A. Jessup, J. C. Boothroyd, D. Brownstein, and P. A. Conrad. 2004. An unusual genotype of Toxoplasma gondii

202

kingdom protista

is common in California sea otters (Enhydra lutris nereis) and is a cause of mortality. International Journal for Parasitology 34:275–284. Mitchell, S., and A. Sinai. 2006. Toxoplasma gondii: Presentations at the IX International Workshop on Opportunistic Protists and the International Society of Protistologists 57th Annual Meetings. Journal of Eukaryotic Microbiology 53:S159. Moloo, S. K., G. J. Losos, and S. S. Katuza. 1973. Transmission of Trypanosoma brucei to cats and dogs by feeding on infected goats. Annals of Tropical Medicine and Parasitology 67:331–334. Moltmann, U. G., H. Mehlhorn, E. Schein, G. Rehbein, W. P. Voigt, and E. Zweygarth. 1983. Fine structure of Babesia equi (Laveran 1901) within lymphocytes and erythrocytes of horses: An in vivo and in vitro study. Journal of Parasitology 69:111–120. Molyneux, D. H. 1982. Trypanosomes, trypanosomiasis and tsetse control: Impact on wildlife and its conservation. Pp. 29–55 in M. A. Edwards and U. McDonnell (editors), Animal disease in relation to animal conservation. Symposia of the Zoological Society of London No. 50. Academic Press, London, UK. Ockert, G., and T. Schmidt. 1976. On the epidemiology of Dientamoeba fragilis Jepps and Dobell 1918. 4th communication: Evidence of Dientamoeba fragilis in Enterobius eggs using isoelectric point determination [in German]. Journal of Hygiene Epidemiology Microbiology and Immunology (Prague) 20:76–81. Olsen, O. W. 1974. Animal parasites: Their life cycles and ecology. 3rd ed. University Park Press, Baltimore, MD. Olson, M. E., and A. G. Buret. 2001. Giardia and giardiasis. Pp. 399–416 in W. M. Samuel, M. J. Pybus, and A. A. Kocan (editors), Parasitic diseases of wild mammals. Iowa State University Press, Ames, IA. Ostrand, W. D., J. A. Bissonette, and M. R. Conover. 1995. Trichomoniasis as a factor in mourning dove population decline in Fillmore, Utah. Journal of Wildlife Diseases 31:87–89. Patterson, D. J. 1999. The diversity of eukaryotes. American Naturalist 65:S96–S124. Patterson, D. J. 2000. Changing views on protistan systematics: The taxonomy of protozoa—an overview. Pp. 2–9 in J. J. Lee, G. F. Leedale, and P. Bradbury (editors), An illustrated guide to the protozoa: Organisms traditionally referred to as protozoa, or newly discovered groups. Society of Protozoologists, Lawrence, KS. Pays, E., and M. Steinert. 1988. Control of antigen gene expression in African trypanosomes. Annual Review of Genetics 22:107–126.

Peirce, M. A. 2000. A taxonomic review of avian piroplasms of the genus Babesia Starcovici, 1893 (Apicomplexa: Piroplasmorida: Babesidae). Journal of Natural History 34:317–332. Peirce, M. A. 2005. Pathogenic subspecies of Plasmodium relictum found in African birds. Veterinary Record 156:328. Pepler, D., and E. E. Oettlé. 1992. Trichomonas gallinae in wild raptors on the Cape Peninsula. South African Journal of Wildlife Research 22:87–88. Philippa, J. D. W., F. A. Leighton, P. Y. Daoust, O. Nielsen, H. Schwantje, T. Shury, R. v. Herwijnen, B. E. E. Martina, T. Kuiken, M. W. G. V. d. Bildt, and A. D. M. E. Osterhaus. 2004. Antibodies to selected pathogens in free-ranging terrestrial carnivores and marine mammals in Canada. Veterinary Record 155:135–140. Pokras, M. A., E. B. Wheeldon, and C. J. Sedgwick. 1993. Trichomoniasis in owls: Report on a number of clinical cases and a survey of the literature. Pp. 88–91 in P. T. Redig, J. E. Cooper, J. D. Remple, D. B. Hunter, and T. Hahn (editors), Raptor biomedicine. University of Minnesota Press, Minneapolis, MN. Quortrup, E. R., and J. E. Shillinger. 1941. 3,000 wild bird autopsies on western lake areas. Journal of the American Veterinary Medical Association 99:382–387. Ratcliffe, H. L., and Q. M. Geiman. 1938. Spontaneous and experimental amebic infections in reptiles. Archives of Pathology 25:160–184. Renold, C., A. Sugar, J. P. Chave, L. Perrin, J. Delavelle, G. Pizzolato, P. Burkhardt, V. Gabriel, and B. Hirschel. 1992. Toxoplasma encephalitis in patients with the acquired immunodeficiency syndrome. Medicine 71. Resendes, A. R., S. Almería, J. P. Dubey, E. Obón, C. Juan-Sallés, E. Degollada, F. Alegre, O. Cabezón, S. Pont, and M. Domingo. 2002. Disseminated toxoplasmosis in a Mediterranean pregnant Risso’s dolphin (Grampus griseus) with transplacental fetal infection. Journal of Parasitology 88:1029–1032. Roberts, L. S., and J. Janovy. 2000. Gerald D. Schmidt and Larry S. Roberts’ Foundations of parasitology. 6th ed. McGraw-Hill, Boston, MA. Rosenfield, R. N., J. Bielefeldt, L. J. Rosenfield, S. J. Taft, R. K. Murphy, and A. C. Stewart. 2002. Prevalence of Trichomonas gallinae in nestling Cooper’s hawks among three North American populations. Wilson Bulletin 114:145–147. Ruiz, A., and J. K. Frenkel. 1980. Toxoplasma gondii in Costa Rican cats. American Journal of Tropical Medicine and Hygiene 29:1150–1160.



Sanger, V. L. 1971. Toxoplasmosis. Pp. 313–316 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Sawyer, T. K., G. S. Visvesvara, and B. A. Harke. 1976. Pathogenic amebas from brackish and ocean sediments with a description of Acanthamoeba hatchetti, n. sp. Science 196:1324–1325. Šlapeta, J. R., D. Modrý, J. Votýpka, M. Jirků, J. Lukeš, and B. Koudela. 2003. Evolutionary relationships among cyst-forming coccidia Sarcocystis spp. (Alveolata: Apicomplexa: Coccidea) in endemic African tree vipers and perspective for evolution of heteroxenous life cycle. Molecular Phylogenetics and Evolution 27:464–475. Sleigh, M. A. 1991. The nature of protozoa. Pp. 1–53 in J. P. Kreier and J. R. Baker (editors), Parasitic protozoa. Academic Press, San Diego, CA. Sousa, O. E., and C. M. Johnson. 1973. Prevalence of Trypanosoma cruzi and Trypanosoma rangeli in triatomines (Hemiptera: Reduviidae) collected in the Republic of Panama. American Journal of Tropical Medicine and Hygiene 22:18–23. Springer, W. T., J. Johnson, and W. M. Reid. 1969. Transmission of histomoniasis with male Heterakis gallinarum (Nematoda). Parasitology 59:401–405. Springer, W. T., J. Johnson, and W. M. Reid. 1970. Histomoniasis in gnotobiotic chickens and turkeys: Biological aspects of the role of bacteria in the etiology. Experimental Parasitology 28:383–392. Stabler, R. M. 1954. Trichomonas gallinae: A review. Experimental Parasitology 3:368–402. Stabler, R. M. 1975. Personal letter, 21 May 1975. In R. G. Botzler (editor). Humboldt State University, Arcata, CA. Steenbergen, J. N., J. D. Nosanchuk, S. D. Malliaris, and A. Casadevall. 2004. Interaction of Blastomyces dermatitidis, Sporothrix schenckii, and Histoplasma capsulatum with Acanthamoeba castellanii. Infection and Immunity 72:3478–3488. Steinert, M., K. Birkness, E. White, B. Fields, and F. Quinn. 1998. Mycobacterium avium bacilli grow saprozoically in coculture with Acanthamoeba polyphaga and survive within cyst walls. Applied and Environmental Microbiology 64:2256–2261. Stevenson-Hamilton, J. 1911. The relation between game and tsetse flies. Bulletin of Entomological Research 2:113–118. Su, C., D. Evans, R. H. Cole, J. C. Kissinger, J. W. Ajioka, and L. D. Sibley. 2003. Recent

kingdom protista

203

expansion of Toxoplasma through enhanced oral transmission. Science 299:414–416. Sukanahaketu, S. 1977. The presence of Dientamoeba fragilis in the Ascaris lumbricoides ova: The first report from Thailand. Journal of the Medical Association of Thailand 60:256–258. Telford, S. R., Jr. 1994. Plasmodia of reptiles. Pp. 1–71 in J. P. Kreier (editor), Parasitic protozoa, Vol. 7. Academic Press, San Diego, CA. Telford, S. R., Jr. 1995. The kinetoplastid hemoflagellates of reptiles. Pp. 161–223 in J. P. Kreier (editor), Parasitic protozoa, Vol 10. Academic Press, San Diego, CA. Tezcan-Merdol, D., M. Ljungström, J. WinieckaKrusnell, E. Linder, L. Engstrand, and M. Rhen. 2004. Uptake and replication of Salmonella enterica in Acanthamoeba rhysodes. Applied and Environmental Microbiology 70:3706–3714. Todd, K. S., and D. M. Hammond. 1971. Coccidia of Anseriformes, Galliformes, and Passeriformes. Pp. 234–281 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Tyzzer, E. E. 1934. Studies on histomoniasis or “blackhead” infection, in the chicken and turkey. Proceedings of the American Academy of Arts and Science 69:189–264. Valkiūnas, G. 2005. Avian malaria parasites and other haemosporidia. CRC Press, Boca Raton, FL. Van der Ploeg, L. H. T. 1987. Control of variant surface antigens switching in trypanosomes. Cell 51:159–161. van Riper, C., III, C. T. Atkinson, and T. M. Seed. 1994. Plasmodia of birds. Pp. 73–140 in J. P. Kreier (editor), Parasitic protozoa, Vol. 7. Academic Press, San Diego, CA. van Riper, C. H., III, S. G. van Riper, M. L. Goff, and M. Laird. 1986. The epizootiology and ecological significance of malaria in Hawaiian land birds. Ecological Monographs 56:327–344.

204

kingdom protista

Visvesvara, G. S. 1999. Pathogenic and opportunistic free-living amebae. Pp. 1383–1390 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. 7th ed. ASM Press, Washington, DC. Visvesvara, G. S., and J. K. Stehr-Green. 1990. Epidemiology of free-living ameba infections. Journal of Protozoology 37:25S–33S. Volkman, S. K., and D. L. Hartl. 2003. A game of cat and mouth. Science 299:353–354. Warner, R. E. 1968. The role of introduced diseases in the extinction of the endemic Hawaiian avifauna. The Condor 70:101–120. Wells, E. A., and W. H. R. Lumsden. 1969. Trypanosome infections of wild mammals in relation to trypanosome diseases of man and his domestic stock. Pp. 135–145 in A. McDiarmid (editor), Diseases in free-living wild animals. Symposia of the Zoological Society of London No. 24. Academic Press, London, UK. Wilson, M. R., and J. B. McAuley. 1999. Toxoplasma. Pp. 1374–1382 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Wilson, V. J. 1972. Observations on the effect of dieldrin on wildlife during tsetse fly Glossina morsitans control operations in eastern Zambia. Arnoldia (Rhodesia) 5:1–12. Woodworth, B. L., C. T. Atkinson, D. A. LaPointe, P. J. Hart, C. S. Spiegel, E. J. Tweed, C. Henneman, J. LeBrun, T. Denette, R. DeMots, K. L. Kozar, D. Triglia, D. Lease, A. Gregor, T. Smith, and D. Duffy. 2005. Host population persistence in the face of introduced vector-borne diseases: Hawaii amakihi and avian malaria. Proceedings of the National Academy of Sciences of the United States of America 102:1531–1536. Yilmaz, S. M., and S. H. Hopkins. 1972. Effects of different conditions on duration and infectivity of Toxoplasma gondii oocysts. Journal of Parasitology 58:939–939.

SEVEN

Kingdom Fungi

CONTENTS Introduction to Pathogenic Fungi Fungal Systematics Fungal Identification Fungal Diseases

205 206 207 208

Histoplasma capsulatum 214 Subcutaneous Diseases  215 Sporothrix schenckii 215 Lacazia (Loboa) loboi216

Saprophytic Fungi 208 Systemic Diseases 208 Aspergillus spp. (Aspergillosis) 208 Coccidioides immitis and C. posadasii (Coccidioidomycosis)210 Other Saprophytic Systemic Fungi Occasionally Affecting Wildlife 214 Emmonsia spp. (Chrysosporium parvum)214

Infective Fungi 216 Batrachochytrium dendrobatidis (Bd) 217 Pseudogymnoascus (Geomyces) destructans221 Miscellaneous Dermatophytes 224 Mycotoxins224 Aflatoxins (Aspergillus f   lavus)225 Literature Cited

1999). Fungi have cell walls with chitin, cellulose, or both, but lack chlorophyll, roots, leaves, stems, xylem, or phloem. Almost all are non-motile and reproduce by means of spores (Alexopoulos et al. 1996). Many fungi have multinucleated cytoplasms. Most fungi form branched, tube-like filaments

INTRODUCTION TO PATHOGENIC FUNGI Fungi are a subgroup of the Supergroup Opisthokonta (Adl et al. 2005) and comprise a Kingdom of eukaryotic organisms distinguished by their unique cellular structures (Alexopoulos et al. 1996, Dixon et al.



225



205

(hyphae) that can develop into multicellular complexes, each of which is called a mycelium. Although most parts of a fungus are potentially capable of growth, the hyphae constituting the body (thallus) of the fungus typically elongate by apical growth (Alexopoulos et al. 1996). Each partition in a fungal hypha is called a septum. While most fungi produce mycelia, the network of branched septated hyphae, some exist as single-celled yeasts that reproduce quickly by budding or fission (Alexopoulos et al. 1996). Species that have the capacity to both produce mycelia and live as single cells are dimorphic; dimorphism is common among species that cause diseases in animals (Alexopoulos et al. 1996). All fungi can reproduce by asexual (somatic) reproduction, and most are known to reproduce sexually. The most common method of asexual reproduction is the production of mitotic spores, with each spore germinating and forming hyphae that then form mycelia. Some fungal spores are produced and borne in sporangia, sac-like structures whose entire contents are converted through cleavage into one or many spores; such spores are called sporangiospores. In contrast, spores produced at the tips or sides of hyphae are called conidia. Most spores are non-motile; however, those of the Phylum Chytridiomycota have a flagellum and are called zoospores (Alexopoulos et al. 1996). Fungal spores can vary considerably in morphology. Other methods of asexual reproduction involve fragmentation of the vegetative form, with each fragment growing into a new individual, fission of vegetative cells into daughter cells, and budding of somatic cells or spores with each bud producing a new individual (Alexopoulos et al. 1996). Sexual reproduction involves the union of two compatible nuclei and usually results in the formation of specialized spores, including oospores, zygospores, ascospores, and basidiospores (Alexopoulos et al. 1996). Sexual stages have not been described for some species of fungi or have been lost, especially among ascomycetes and basidiomycetes. The term teleomorph is used to identify a meiotic sexual form that produces 206  kingdom fungi

ascospores or basidiospores; the term anamorph refers to a mitotic asexual form that does not produce spores (Alexopoulos et al. 1996). Fungi undergoing sexual reproduction fall into one of three morphological categories (Alexopoulos et al. 1996). Most fungi are sexually undifferentiated, and sexually functional structures are produced that are morphologically indistinguishable as male or female. In contrast, monoecious (hermaphroditic) fungi have thalli that each bear both male and female organs that may (or may not) be compatible. Rarely, there are dioecious fungi in which some thalli bear only male and some bear only female organs. Asexual reproduction is important for rapidly colonizing a new habitat because it results in the production of a large number of individuals in a short time. Sexual reproduction results in a high incidence of recombination and formation of new genotypes that enable the fungi to adapt more readily to new conditions (Alexopoulos et al. 1996). Overall, fungi are very successful organisms, as evidenced by their ubiquity in nature; they function primarily as decomposers and saprophytes, and less commonly as parasites of living hosts.

Fungal Systematics Although once classified with plants, the emergence of molecular phylogenetics as a discipline has made it clear that fungi are more closely related to animals (Adl et al. 2005). Because the usual definition of species as members of sexually interbreeding groups cannot always be applied, the taxonomy of fungi, including yeasts, historically has been based on morphology, along with some physiological characterizations, and some specialized tests adapted from clinical bacteriology (Dixon et al. 1999, Merz and Roberts 1999). Fungal systematics have changed considerably over the years, being particularly influenced by acceptance of phylogenetic-based systems of classification, development of molecular techniques, and discovery of new taxa, including fossils. There is a proposal for a major revision of the phylogenetic classification of the Kingdom Fungi

that calls for one subkingdom, Dikarya, comprising the Ascomycetes and Basidiomycetes, as well as seven phyla (Hibbett et al. 2007). However, until it is clear where the majority of pathogenic fungi are placed in this revision, we currently follow the more traditional format of recognizing four phyla in the Kingdom Fungi: Chytridiomycota, Zygomycota, Ascomycota, and Basidiomycota (App. 1: Table 6). The Form-group Deuteromycota (App. 1: Table 6), also called Mitosporic Fungi (Howard 2003), was established by taxonomists to include organisms known only in anamorph form. In recent years, molecular phylogeny has been applied extensively to members of the Kingdom Fungi (Lutzoni et al. 2004, James et al. 2006, Spatafora et al. 2006), and many asexual fungi have been linked with their sexual relatives. For example, members of the pathogenic genera Blastomyces, Emmonsia, Histoplasma, and Paracoccidioides are closely related, with about 5% base differences, and with even less of a difference between Blastomyces and Emmonsia (Gueho et al. 1997). All now are known to be closely allied with other members of the Family Onygenaceae in the Class Ascomycetes (Howard 2003). In this text, members of the Deuteromycota are included with their close sexual relatives wherever possible. The Phylum Chytridiomycota comprises those members of the Kingdom Fungi that produce motile cells in their life cycles (Alexopoulos et al. 1996). These flagellated spores (zoospores) are produced in four of the five orders within the single Class Chytridiomycetes (Alexopoulos et al. 1996) and are homologous to the non-fungal opisthokonts (App. 1) (Barr 1992). Current evidence is that the Chytridiomycota are polyphyletic, consisting of some early diverging lineages that retained the zoospore (James et al. 2006). However, the euchytrids, a large clade of orders within the Chytridiomycota, appear to be monophyletic (James et al. 2006). The Phylum Zygomycota contains those fungi that produce zygospores, formed from morphologically identical gametes, as a result

of sexual reproduction (Howard 2003). Current evidence is that the Zygomycota also are polyphyletic (James et al. 2006). All representatives of the Class Trichomycetes are obligate parasites of arthropods and will not be addressed further. The Class Zygomycetes has two orders, Mucorales and Entomophthorales, both of which include important animal pathogens. The Mucorales generally produce nonseptate hyphae, while the Entomophthorales usually have septated hyphae (Howard 2003). The Phylum Ascomycota comprises fungi that reproduce sexually by means of ascospores that are contained in a specialized sac called an ascus (Howard 2003). Molecular phylogeny studies have allowed associations of many former members of the Deuteromycota to be classified, even when known teleomorphs were not identified for the specific anamorph forms (Lutzoni et al. 2004); these are included when applicable. Two classes commonly are recognized. The Endomycetes undergo asexual reproduction by budding or fission of somatic cells, whereas Ascomycetes form blastic or thallic conidia (Howard 2003). Current genetic phylogeny of the Kingdom Fungi supports the monophyly of the Ascomycota (James et al. 2006). Members of the Phylum Basidiomycota all produce spores (basidiospores) on the outside of a club-shaped to elongate structure (basidium). Pathogenic yeasts are of most concern in animal diseases (Howard 2003). The Class Hymenomycetes includes mushrooms (e.g., Order Agaricales), some of which can be toxic (Howard 2003). Current genetic phylogeny of the Kingdom Fungi supports the monophyly of the Basidiomycota (James et al. 2006).

Fungal Identification Laboratory identification of mycoses calls for the initial collection and storage of specimens and the proper identification, which generally is based on direct microscopic examination to determine size and morphology, antigenic detection, finding fungus-specific metabolites, detection of cell wall components, detection of kingdom fungi   207

fungus-specific nucleic acids, and direct culturing on fungal media (Merz and Roberts 1999). Direct examination of fungi within clinical specimens can be done with potassium hydroxide, India ink, or calcofluor white, as well as Giemsa and a few other staining techniques (Kwon-Chung and Bennett 1992). A variety of media are used for culturing fungi from clinical specimens, and there are a number of laboratory techniques for their identification (Kwon-Chung and Bennett 1992). Many of these are similar to techniques used for the cultivation and identification of pathogenic bacteria (Gough 1997).

Fungi cause several types of diseases. The most common type of fungal disease is a mycosis, the direct growth of fungi on or in host cells. Pathogenic fungi have varied growth patterns on or in their hosts, and often are labeled as superficial, cutaneous, subcutaneous, or systemic (Friend 1999b). In contrast, a mycotoxicosis is a disease resulting from fungal metabolites acquired through ingestion, inhalation, or abrasion (Bennett and Klich 2003). Allergic diseases may result from fungal spores or colonies developing in the lungs.

Fungal Diseases

Systemic Diseases

While the number of fungi pathogenic to vertebrates and invertebrates is not certain, it probably is not high. For example, of the estimated 250,000 species (Dixon et al. 1999) to 1.5 million species (Hawksworth 1991) of fungi known, only about 120 species pathogenic to humans are known (McGinnis et al. 1999). Rarely, fungi are contagious; examples include epidermal parasites such as Pseudogymnoascus (Geomyces) destructans (cause of white-nose syndrome in bats) and Candida albicans, a systemic parasite. But most pathogenic fungi are primarily free-living saprophytes that do not require an animal host for survival or propagation (Hogan et al. 1996); they are opportunists of low virulence that only rarely cause disease in healthy individuals. They are more likely to affect animals with compromised immunity and typically reach a dead end in their hosts (Burek 2001). Thus, most cases of fungal infections are acquired from some point source in the environment rather than from another infected animal (Dixon et al. 1999). It is not clear why there is a selective pressure for continued animal pathogenicity among fungi well adapted to abiotic environments. However, soil amebae may ingest the yeast forms of some soil fungi and help select and maintain traits that confer virulence in these pathogenic fungi for mammals (Steenbergen et al. 2004).

aspergillus spp. (aspergillosis)

208  kingdom fungi

SAPROPHYTIC FUNGI

causative agent (cl assification, morphology) The genus Aspergillus is a member of the Family Trichocomaceae, Order Eurotiales, Class Ascomycetes (App. 1: Table 6) (Howard 2003). This genus includes at least 185 species, of which 20 are known to be pathogenic to plants or animals (Converse 2007) The most common animal pathogens are A. fumigatus and A. niger, which cause aspergillosis, and A. flavus, which produces an aflatoxin (Chute et al. 1965). Aspergillus fumigatus is the most commonly reported species and is most pathogenic for both wild birds (Converse 2007) and mammals (Burek 2001). Pathogenic members of the genus Aspergillus include anamorphs among at least seven teleomorph genera: Emericella, Eurotium, Fennellia, Hemicarpenteles, Neopetromyces, Neosartorya, and Petromyces (Howard 2003). Because species of Aspergillus often are not clearly identified, the genus will be addressed as a whole. host r ange and geogr aphic distribution  Aspergillus spp. have a worldwide distribution (Burek 2001) and commonly are found among free-living birds (McDiarmid 1955, Beer 1963, Rosen 1964, O’Meara and Witter 1971) and mammals (Burek 2001). In North America, infection by Aspergillus spp. are considered common in waterfowl, gulls, and corvids; occasional

in songbirds, upland game birds, and blackbirds; and infrequent or rare in wild raptors, herons, and shorebirds (Friend 1999a). On a worldwide basis, waterfowl, raptors, and gulls represent the majority of free-living hosts reported with aspergillosis (Converse 2007). Although less frequently reported in mammals, Aspergillus spp. have occurred in farmed deer (Odocoileus spp.) (Jensen et al. 1989), alpacas (Vicugna pacos) (Pickett et al. 1985, Severo et al. 1989), camels (Camelus dromedarius) (El-Khouly et al. 1992), dolphins (Tursiops truncatus) (Reidarson et al. 1998b), and wild felids (Peden et al. 1985). It also is reported in domestic birds and mammals (Fraser and Mays 1986, Acha and Szyfres 2001). life cycles and variations Aspergillus spp. reproduce asexually by forming aerial fruiting bodies that bear conidiospores, asexual spores formed on the tip of specialized hyphae called conidiophores; these spores are easily inhaled into lungs and air sacs (Converse 2007). Some Aspergillus spp. produce mycotoxins (aflatoxins) (Converse 2007); this topic is addressed later in this chapter. reservoirs and transmission  Aspergillus spp. generally are ubiquitous saprophytes found in soil, decaying vegetation, and agricultural wastes such as spilled grain and corn silage (O’Meara and Witter 1971, Converse 2007). Aspergillus spp. generally grow over a wide temperature range (18 to 30°C); most will sporulate at 23 to 26°C (Raper and Fennell 1965). Growth and sporulation of Aspergillus spp. are encouraged by moist conditions and decaying vegetation (Burek 2001). Spores commonly are released when fungal fruiting bodies are broken by animal movements (Bellrose 1945). Primary transmission is through inhalation, which typically requires one to several million air-borne spores (Austwick 1968, Atasever and Gumussoy 2004). Disturbance of infected soil and vegetation results in aerosols, allowing for aspiration and deposition of spores in bronchioles and pulmonary alveoli (Burek 2001). Transmission also can occur by puncture wounds contaminated with hyphae or spores, especially if the lungs or air sacs are penetrated (O’Meara

and Witter 1971). Aspergillus spp. spores can penetrate intact egg shells, resulting in mortality of the embryos or birds hatched (O’Meara and Chute 1959). Ingestion is an unlikely route of transmission (Chute et al. 1965). clinical effects and identification  Aspergillosis is a noncontagious disease usually caused by inhalation of Aspergillus sp. spores (Converse 2007). Most reports are among birds. Two hypotheses for the development of avian aspergillosis are that the initial dose of spores is very high, exceeding the natural resistance of a host, or that birds may carry the fungi but become ill after stresses lower the immune response (Friend and Trainer 1969). Both acute and chronic forms of aspergillosis occur (Converse 2007). In acute cases, birds die in a few days from severe respiratory compromise following a generalized lung infection; birds generally are in good body condition with no other evident problems (O’Meara and Witter 1971, Redig et al. 1980). With chronic aspergillosis, birds develop a slowly progressing disease and may become severely debilitated before dying. Signs often include emaciation, reduced activity, inability to fly, dyspnea, and diarrhea (Converse 2007). Characteristic lesions include firm nodules and sometimes colored conidiospores in the respiratory tract (O’Meara and Witter 1971, Redig et al. 1980). There typically is a strong inflammatory response with many nodules disseminated through the respiratory tract; in time any internal organ can be infected via the vascular system. Birds with chronic aspergillosis also may have evidence of trauma, parasitism, and other diseases, as well as a history of capture, captivity, or rehabilitation (Converse 2007). Avian aspergillosis losses often are associated with changing environmental conditions, such as increases in temperature and moisture, which can enhance development of fungi on waste grain in agricultural fields (Converse 2007). Among mammals, clinical signs and pathology vary considerably by the organ system involved (Burek 2001). With a primary respiratory disease, there are miliary nodules kingdom fungi   209

in the lungs containing fungal hyphae; in disseminated disease, nodules may be scattered throughout the body. The fungi can cause considerable tissue damage associated with vascular invasion and thrombosis; the hyphae proliferate in necrotic tissue (Burek 2001). Antemortem (before death) diagnosis is based on clinical signs, serological tests, culture of tracheal washes, endoscopy, and radiographs (Converse 2007). Diagnostic tests include an enzyme-linked immunosorbent assay (ELISA) (Converse 2007) and polymerase chain reaction (PCR) (Katz et al. 1996). Adult free-ranging female birds can be tested indirectly by testing for IgG antibodies in their eggs against Aspergillus spp. (Graczyk and Cranfield 1995); titers in eggs are correlated with specific antibody levels in the mothers (Graczyk and Cranfield 1996). Because Aspergillus spp. are common contaminants, positive culture should correlate with appropriate histopathologic evidence of tissue invasion (Chandler and Watts 1987). Postmortem diagnosis of aspergillosis is based on the presence of compatible lesions at necropsy and identification of Aspergillus spp. in tissues by isolation or identification of characteristic fungal hyphae (Converse 2007). The fungi can be grown on several media (Kunkle and Richard 1998). However, the fungal hyphae need to be distinguished from other invasive fungi such as Fusarium spp. (Chandler and Watts 1987). Monoclonal and polyclonal antibodies can be used with an immunohistochemical label to identify Aspergillus spp. (Carrasco et al. 1993). population effects Epizootics of acute aspergillosis have been reported among freeliving waterfowl (Bowes 1990), including mallards (Anas platyrhynchos) (Neff 1955, Adrian et al. 1978), as well as common crows (Corvus brachyrhynchos) (Zinkl et al. 1977). Mortality may disproportionately affect younger animals (Brand et al. 1988), and may be influenced by severity and duration of exposure, and stress from concurrent infections of other parasites (O’Meara and Witter 1971). Chronic aspergillosis is associated with individual bird deaths rather than group mortality (Converse 2007). 210  kingdom fungi

Aspergillosis was the most common (23%) finding among dead or moribund common loons (Gavia immer) in New York from 1972 to 1999, with ingestion of lead fishing weights being the second most common diagnosis (21%) (Stone and Okoniewski 2001). This prevalence of lead may be significant as aspergillosis has been associated with lead poisoning in other areas (Locke et al. 1969, Bair et al. 1988, Souza and Degernes 2005). Aspergillosis also has been associated with tracheal trematodes (Pennycott 1999) and thiamine deficiency (Friend and Trainer 1969). Aspergillosis has caused mortality in a number of marine mammals (Sweeney et al. 1976). Pneumonia caused by Aspergillus spp. may account for most pulmonary mycoses in marine mammals (Reidarson et al. 1998b). special problems  No specific persistent problems are reported. control  Immunity to Aspergillus spp. can develop among birds. Adult birds are more resistant than chicks to aspergillosis; there is evidence for innate resistance, but little definitive data are available (Converse 2007). Birds are able to produce both IgM and IgG antibodies to Aspergillus spp. (Martinez-Quesada et al. 1993), and mammals produce IgE (Kwon-Chung and Bennett 1992), but the role of antibodies in protecting infected animals or enhancing recovery is not clear (Converse 2007). There is evidence among birds that maternal antibodies are protective for hatchlings (Graczyk and Cranfield 1995). Among mammals, itraconazole can be used for treatment of systemic aspergillosis (Denning and Stevens 1990), but cure is uncommon (Burek 2001). Decreasing the numbers of spores contaminating an environment and controlling other predisposing conditions also are considered important (Pier and Richard 1992). coccidioides immitis and c. posadasii (coccidioidomycosis) causative agent (cl assification, morphology)  Coccidioides spp. are ascomycetes found in the Family Onygenaceae, Order

Map 7.1.  Distribution of endemic Coccidioides immitis and C. posadasii (Bultman et al. 2004; courtesy of M. W. Bultman and the U.S. Geological Service).

Onygenales (App. 1: Table 6) (Howard 2003, Clemons et al. 2007). Until recently only one species, C. immitis, was recognized. However, there now is phylogenetic evidence for a second species, C. posadasii (Fisher et al. 2002, Tintelnot et al. 2007). The teleomorph form is not certain (Saubolle et al. 2007), but Uncinocarpus spp. has been proposed (Sigler 2003). host range and geographic distribution  Coccidioides spp. can infect many mammal species (Burek 2001, Laniado-Laborin 2007) and rarely reptiles (Timm et al. 1988). No reports have yet been made of avian infections (Laniado-Laborin 2007). Most reports from wild species are from captive animals in zoos or exhibits in enzootic regions; death from coccidioidomycosis has been reported from a variety of nondomestic canids and felids, as well as bats, wallabies, kangaroos, tapirs, and several primates (Shubitz 2007). Among free-living wildlife, Coccidioides spp. has been reported

from desert bighorns (Ovis canadensis nelsoni) (Jessup et al. 1989), mountain lions (Puma concolor) (Clyde et al. 1990, Adaska 1999), and coyotes (Canis latrans) (Straub et al. 1961). Coccidioidomycosis also has been reported from free-living marine mammals, including California sea lions (Zalophus californianus) (Fauquier et al. 1996), a bottle-nosed dolphin (Tursiops truncatus) (Reidarson et al. 1998a), and a sea otter (Enhydra lutris) (Cornell et al. 1979). Among domestic mammals, lesions containing Coccidioides spp. have been found in cattle, sheep, and swine, but these species rarely develop overt disease (Maddy 1959). Coccidioides immitis and C. posadasii are endemic in the Western Hemisphere, almost exclusively between 40°N and 40°S latitudes (Burek 2001) in the Lower Sonoran Life Zone (Map 7.1); these areas seldom have freezing temperatures during the year, and the climate is arid, with yearly rainfall ranging from 11 to 44 cm kingdom fungi   211

(Galgiani 1993). Coccidioides spp. survives well in saline soil (Fauquier et al. 1996), and elevated salinity may inhibit some microbial competitors (Egeberg et al. 1964). Presence of fine sand and silt in the soil is the single characteristic common in all areas in which the organism is found (Saubolle et al. 2007). In enzootic areas, the mean annual soil temperatures of enzootic regions range from 15°C to over 22°C; the soil is alkaline, with adequate pore space in the upper 20 cm for moisture, oxygen, and room for growth. Endemic foci have been identified in semiarid areas of the United States and Mexico, as well as some areas of Central and South America (Laniado-Laborin 2007). There is evidence that Coccidioides spp. may have had a larger range at one time in North America, or that migratory animals from other regions were infected as they passed through enzootic areas (Morrow 2006). With the division of species, C. immitis now is geographically restricted to the Central Valley of California and southern California; in contrast, C. posadasii occurs in all other regions of North, Central, and South America where infections have been reported (Fisher et al. 2002). Both species appear to be have considerable genetic diversity with frequent genetic recombination; both species also have biogeographically distinct populations (Barker et al. 2007). The genetic diversity of Coccidioides spp. in North America has clear geographic partitions, whereas the populations of Coccidioides spp. in South America are genetically similar and appear to be founded on a North American population centered in Texas; one hypothesis is that South America was colonized by Amerindians carrying this fungus (Fisher et al. 2001). life cycles and variations  The life cycle of Coccidioides spp. is unique among pathogenic fungi (Kirkland and Fierer 1996). These dimorphic fungi are considered true pathogens and readily change from a saprophytic mycelial form in soil into a round, thick-walled spherule/endospore form in an animal host (Shubitz 2007). In soil, the fungus grows on decaying organic matter and is characterized by mycelia 212  kingdom fungi

with branching segmented hyphae. Here the fungus reproduces asexually by disarticulating the hyphae into small, environmentally resistant structures called arthrospores, also called arthroconidia; the arthroconidia are released from mycelia and dispersed in the air (Hector and Laniado-Laborin 2005). The saprophytic mycelial cycle does not depend on a reversion to the parasitic form as it contains large amounts of arthroconidia that are dispersed by the wind to colonize new sites. However, upon inhalation by a host, the conidia are converted by isotropic growth into spherules (Cole and Hung 2001). Each spherule divides to produce hundreds of endospores, which then disperse into surrounding tissue and produce a second generation of spherules (San-Blas and NiñoVega 2004). The endospores again can revert to the saprophytic mycelial stage if they reach the soil through death of the infected animal or by body secretions (Bultman et al. 2004). reservoirs and transmission  The reservoirs are the soils and infected animals of the Lower Sonoran Life Zone. Coccidioides spp. spores present in dry soil first may require moisture to germinate and grow, and then require a dry period for the fungal hyphae to desiccate and form arthroconidia (Kolivras and Comrie 2003). Cases of coccidioidomycosis are linked to both the amount of dust in the air and the cumulative rainfall over the preceding several months (Park et al. 2005). In one study, precipitation during the normally arid periods in late spring and early summer 1.5 to 2 years before the season of exposure was the dominant predictor of coccidioidomycosis cases for all seasons (Comrie 2005). Work is continuing to clarify the environmental factors leading to the success of Coccidioides spp. throughout its range (Baptista-Rosas et al. 2007, Fisher et al. 2007). Inhalation of arthrospores is the principal means of transmission (Davis 1981); in the mouse, only a few arthrospores administered intranasally are required for infection (Galgiani 1993). The arthroconidia developing in the Coccidioides spp. hyphae commonly are

separated by empty, thin-walled brittle cells (disjunctors) formed by autolysis. The spores are easily released into the air by soil disruption and wind (Saubolle et al. 2007), and spread of the fungus by wind is an important local process (Bultman et al. 2004). However, there are distinct genetic clades of Coccidioides spp.; thus arthroconidia probably do not travel over long distances to new sites (Fisher et al. 2001). clinical effects and identification  Coccidioidomycosis, the disease resulting from Coccidioides spp. infection, is not to be confused with the disease “coccidiosis,” caused by members of the sporozoan protozoa (App. 1: Table 5) (Galgiani 1999). Virulence of Coccidioides spp. is a function of a spherule outer wall glycoprotein that reduces cell-mediated immunity, reduces glycoprotein occurrence on endospores to prevent host recognition, and produces of chemicals contributing to tissue damage at the sites of infection (Hung et al. 2007). Clinical signs vary greatly, ranging from a benign, selflimited upper respiratory infection, to chronic pulmonary disease, to disseminated fatal disease (Burek 2001). In humans, symptoms of coccidioidomycosis usually begin 7 to 21 days after inhalation of arthroconidia and may include fever, cough, chest discomfort, malaise, and fatigue (Saubolle et al. 2007). Symptoms of pneumonia generally last under 3 weeks, but can be longer. Skin rashes can be seen in 10 to 50% of infected humans. Disseminated coccidioidomycosis occurs in less than 5% of symptomatic patients; those of black or Filipino ethnic backgrounds, pregnant women, and immunocompromised patients have higher risks for disseminated disease (Saubolle et al. 2007). Detection and diagnosis in animals, including humans, often is based on direct microscopy, culture, and serologic findings (Saubolle 2007, Parish and Blair 2008). Fungal identification can be based on phenotypic characters such as the presence of spherules in hosts, the presence of arthroconidia in culture, or genotypic characteristics (Saubolle 2007). Enzyme immunoassays and immunodiffusion methods commonly

are used for detection of both IgM and IgG antibodies (Saubolle 2007). Polymerase chain reaction tests (PCR) and restriction fragment length polymorphism (RFLP) have been used to identify and differentiate Coccidioides spp. (Umeyama et al. 2006, Tintelnot et al. 2007). Detection of C. immitis in the soil is difficult. Traditionally, mice have been inoculated with isolates from suspect soil and their organs later evaluated for evidence of the unique spherules characteristic of the parasitic form of the fungus (Bultman et al. 2004). Although DNA analysis is used, there are no standardized procedures as yet (Bultman et al. 2004). population effects  There are no known ongoing significant population impacts among wildlife. special problems  Coccidioidomycosis is a common infectious disease among humans in the southwestern United States, Mexico, Central America, and South America (DiCaudo 2006), with upwards of 100,000 primary coccidial infections of humans each year in the United States alone. While recent increases in human cases in Arizona were thought to be linked to development of a hypervirulent strain, this does not appear to be the case (Jewell et al. 2008). About 3,000 inmates were required to be moved out of the Central Valley of California to other prisons in 2013 to reduce the risk of coccidioidomycosis among high-risk prisoners (http://articles.latimes.com/2013/apr/30/local/ la-me-prisons-valley-fever-20130430). The occurrence of coccidioidomycosis in marine mammals is intriguing but most likely is the result of marine mammals inhaling arthroconidia blown from land over the water (Shubitz 2007). Coccidioides spp. can survive for an extended period in seawater (Fauquier et al. 1996). control  Macrophages producing cytokines against spherules are one important host defense (Viriyakosol et al. 2005). Humoral antibodies generally are not considered protective against Coccidioides spp. (Saubolle et al. 2007); however, B-lymphocytes may play a role in vaccine-induced immunity (Magee et al. 2005). kingdom fungi   213

Orally administered azole and other antifungal agents are the primary therapy used for captive animals (Shubitz 2007). Among humans, coccidioidomycosis usually is treated with azole or amphotericin B (Parish and Blair 2008). Recovery from coccidioidal infection confers lifelong protection against reinfection (Smith and Pappagianis 1961, Pappagianis 2001), and there is considerable interest in developing effective vaccines, especially for humans (Johnson et al. 2007). A vaccine based on whole killed spherules is effective for some nonhumans (Levine et al. 1965). Other Saprophytic Systemic Fungi Occasionally Affecting Wildlife emmonsia spp. (chrysosporium parvum) Emmonsia crescens and E. parva are ascomycetes (Family Onygenaceae, Order Onygenales) (App. 1: Table 6) that infect small mammals. These parasites also have been classified as subspecies of Chrysosporium parvum (Burek 2001), but currently are identified as two distinct species of Emmonsia (Sigler 2003). Both species are dimorphic fungi that live in the soil and produce self-limiting pulmonary mycotic infections in a wide variety of small mammals that burrow or produce ground nests; they also infect mustelids (Dvorek et al. 1973, Jellison 1981). They appear to have a worldwide distribution (Jellison and Lord 1964, Dvorek et al. 1973, Jellison 1981, Hubálek et al. 2005), with the exception of Australia and Antarctica (Emmons and Jellison 1960). Transmission is primarily through inhalation of spores, called adiaspores, from the freeliving soil forms (Burek 2001). The predator– prey relation may help in the spread of the fungi since adiaspores passed in the feces are viable and germinate (Křivanec and Otčenášek 1977). The fungi are associated with mammalian species using forest–grassland ecotones (Leighton and Wobeser 1978), especially near windbreaks (Hubálek et al. 1991, Hubálek et al. 1998), but can occur in other uncultivated soils (Fischer 2001). Peak fungal proliferation 214  kingdom fungi

occurs when mean monthly soil temperatures 5 cm below the surface range between 3 and 12°C (Hubálek et al. 1993). In mammalian tissues, the adiaspores develop into thick-walled, gray-white fungal spherules which are surrounded by granulomatous infiltrates; the disease is called adiaspiromycosis. There appears to be no multiplication of the fungus in host species, even though extensive infections have been observed in small mammals (Burek 2001). Lesions usually are restricted to lung inflammations. In heavy infections there may be some pulmonary compromise (Chandler and Watts 1987). No control methods are proposed. his topl asma c apsul atum Histoplasma capsulatum, the cause of histoplasmosis, is an ascomycete (Family Onygenaceae, Order Onygenales) (App. 1: Table 6); this dimorphic fungus develops hyphae in soil and a yeast stage in host tissues (Burek 2001). This parasite has a worldwide distribution and is found in both temperate and tropical zones (Sanger 1981). Only mammals are known to be naturally infected; of these, rodents and bats are most important (Otčenášek et al. 1967). Currently three subspecies are recognized (McGinnis et al. 1999), of which H. capsulatum var. capsulatum is of greatest significance and is the focus of this summary. The soil is the primary natural habitat and source of infection of H. capsulatum (Otčenášek et al. 1967). Interestingly, the fungi often are associated with soils enriched by bird feces (Ajello 1964) or with bat guano (DiSalvo et al. 1970). Despite past justification of histoplasmosis as a basis for instituting “blackbird control” programs (Anonymous 1975), birds are not able to sustain infections of H. capsulatum, likely because of their high (42°C) body temperatures (Ajello 1967); overall, there is no evidence that birds regularly carry the fungus or can seed it into the soil (Sanger 1981). Bats also are susceptible to histoplasmosis (Ajello 1967), and the frequent involvement of the intestinal tract in bats is a mechanism

for seeding the environment with the fungus (Shacklette et al. 1967, Greer and McMurray 1981). In Mexico, H. capsulatum has been found primarily in caves, abandoned mines, and houses with bat guano (Velasco-Castrejon and Gonzalez-Ochoa 1977). Further, the fungi can grow in bat guano; thus bats can excrete H. capsulatum in a medium in which the fungus can propagate itself (Isbister et al. 1976). It is likely that bats are the animals of greatest significance in the epizootiology of the disease, but other factors also are important (Ajello 1967), including temperatures under 40°C and soil pH of 5 to 10 (Goodman 1965). Histoplasma capsulatum also may parasitize and grow in soil amebae; parasitism of soil amebae may select for features that increase pathogenicity of H. capsulatum for mammals (Steenbergen et al. 2004). Transmission to susceptible hosts occurs by inhalation of spores from soil (Otčenášek et al. 1967), and rarely by ingestion or skin contact with infective spores of H. capsulatum (Burek 2001). Bat caves can have high concentrations of H. capsulatum in their atmosphere (Shacklette et al. 1967). Foci of infection occur in the lungs and regional lymph nodes. In disseminated histoplasmosis, many tissues are affected, including lymph nodes, spleen, bone marrow, liver, and adrenals (Burek 2001). Many infected animals on enzootic sites have clinically inapparent infections (Burek 2001); various stresses have been implicated as contributing factors for immunosuppression and emergence of overt disease in wildlife (Quandt and Nesbit 1992). Diagnosis is based on finding organisms in monocytes and macrophages of blood smears, bone marrow aspirates, or biopsy samples of lymph nodes, liver, intestine, or rectum (Burek 2001). Thoracic radiographs, a Histoplasma skin test, and serologic methods can be used (Quandt and Nesbit 1992), as well as ELISA and radioimmunoassay (Durkin et al. 1997). Restriction fragment length polymorphisms and polymerase chain reaction (PCR)–based DNA fingerprinting also have

been successful tools to identify and characterize strains (Kersulyte et al. 1992, Bialek et al. 2002). There are no control methods for histoplasmosis in wildlife. However, ketoconazole has been used with domestic pets (Legendre 1995), and several other antibiotics appear to have promise (Li et al. 2000).

Subcutaneous Diseases A number of fungi cause subcutaneous infections among mammals (Burek 2001), including humans (Schell et al. 1999), and birds (Friend 1999b). Until recently, most of these fungi were classified among the Fungi Imperfecti, but the relationship for many of them to other wellestablished groups is becoming clearer (App. 1: Table 6). Two species causing some pathology among wildlife are Sporothrix schenckii and Lacazia (Loboa) loboi. sporothrix schenckii  Sporotrichosis is a subcutaneous mycosis caused by Sporothrix schenckii (Order Ophiostomatales, Class Ascomycetes), a thermally dimorphic fungus that forms mycelia in decaying vegetation and fungal media at 25 to 30°C, but yeast-like stages in tissues and fungal media at 37°C (Schell et al. 1999). The parasite has a worldwide distribution, but is more common in temperate and tropical climates (Lopes-Bezerra et al. 2006). Based on mitochondrial DNA analysis using restriction fragment length polymorphism patterns, there are 24 different genotypes divided into two main types (Mesa-Arango et al. 2002). Type A is found predominantly in North and South America and South Africa (Ishizaki et al. 1998, 2000); Type B is found primarily is Spain, Asia, and Australia (Ishizaki et al. 2000, Mesa-Arango et al. 2002). Sporothrix schenckii occurs in both marine (Migaki et al. 1978) and terrestrial mammals (Jones and Hunt 1983, Costa et al. 1994, Werner and Werner 1994). Though it has also been isolated from birds (Ziólkowska and Tokarzewski 2007), S. schenckii is not known kingdom fungi   215

to cause pathology or mortality among birds. This fungus typically is inoculated through a scratch or into a previous wound by plant material, particularly of a grassy or straw-like nature (Mackinnon et al. 1969, Windingstad 1990). In its most common form, lymphocutaneous lesions are characterized by small, firm nodules on the head, extremities, and tail that ulcerate and discharge hemorrhagic or serous exudate (Burek 2001). Diagnosis is based on clinical signs, histopathology, fungal cultures, and serologic tests (Goad and Pecquet-Goad 1986, Hu et al. 2003, Bernardes-Engemann et al. 2005). In captive hosts, the infection can be successfully treated with potassium iodide (Scott et al. 1995) or several antibiotics such as itraconazole and ketoconazole (Legendre 1995). lacazia (loboa) loboi  Lacazia (Loboa) loboi is an ascomycete (Family Onygenaceae, Order Onygenales) (Herr et al. 2001) that is closely related to the fungal parasite Paracoccidioides brasiliensis (Vilela et al. 2005). Lacazia loboi causes subcutaneous mycoses in bottle-nosed dolphins (Tursiops truncatus) (Migaki et al. 1971, Caldwell et al. 1975, Simõse-Lopes et al. 1993) and the Guiana dolphin (Sotalia fluviatilis guianensis) (DeVries and Laarmann 1973), as well as humans (Burns et al. 2000, Opromolla et al. 2000). It is considered enzootic to the South and Central American tropics (Pfaller and Diekema 2005). Natural infections have been reported only in dolphins and humans, but experimental infections occur in hamsters (Opromolla and Nogueira 2000) and armadillos (Euphractus sexcinctus) (Sampaio and Braga-Dias 1977). There may be both an aquatic and a terrestrial reservoir. The occurrence of lobomycosis in dolphins from marine and freshwater habitats of Florida (Reif et al. 2006), Texas (Cowan 1993), the Spanish–French coast (Symmers 1983), the south Brazilian coast (Simõse-Lopes et al. 1993), and the Surinam River Estuary of South America (DeVries and Laarmann 1973) is evidence that an aquatic reservoir is important. Lacazia loboi also may be a saprophyte of 216  kingdom fungi

soil or vegetation (Pfaller and Diekema 2005). Based on consistent differences in fungal cell size between dolphins and humans, there may be strain differences between the human (terrestrial) and dolphin (aquatic) strains (Haubold et al. 2000). Cutaneous trauma is the likely mode of infection (Pfaller and Diekema 2005). There is one possible case of dolphin-to-human transmission (Borelli 1961). In dolphins and humans, the disease is characterized by slowly developing cutaneous nodules and hard plaques arising on injured areas of skin (Caldwell et al. 1975, SimõseLopes et al. 1993, Pfaller and Diekema 2005). Diagnosis is based on finding the characteristic yeast cells in lesion exudates or tissue sections (Burns et al. 2000); a vinyl adhesive tape method has proved simple and useful for diagnosis (Miranda and Silva 2005). Western blotting analysis is used to detect dolphin and human antibodies against L. loboi, and may be useful for further epizootiological studies on the parasite (Mendoza et al. 2008). Although lobomycosis generally has been reported as isolated observations among dolphins, there is evidence for a developing epizootic among bottle-nosed dolphins in a Florida lagoon. Environmental stress caused by freshwater intrusions are speculated to have contributed to the high (30%) prevalence of infections observed (Reif et al. 2006). Several antifungal agents have been used in humans and other animals (Brun 1999). Generally, treatment of free-ranging animals is not likely or warranted (Burek 2001).

INFECTIVE FUNGI Many authors distinguish between superficial fungi, to which there is little cellular response from the host and which have only a general cosmetic impact, from cutaneous fungi (dermatophytes) found on the skin and appendages and which can invade keratinous tissues (skin, hair, nails, etc.) of living animals (Friend 1999b, Kane and Summerbell 1999). It often is not clear for dermatophytes if they are transmitted

directly between susceptible hosts (infectious) or arise from point source infections with soil and water as the reservoir. For Batrachochytrium dendrobatidis (Bd), the chytrid fungus infecting amphibians, there is ample evidence of transmission between amphibians through the water, even though the vast majority of species within the Class Chytridiomycetes (chytrid fungi) are saprophytes widespread in soil and water (Sparrow 1960, Olsen et al. 2004); Bd is the only member of its clade known to attack vertebrates (Joneson et al. 2011). Pseudogymnoascus destructans, the cause of white-nose syndrome in bats, also appears to be transmitted between individuals, but is isolated from soil of bat hibernacula as well (Lorch et al. 2013). Future studies may clarify some of these issues further. batrachochytrium dendrobatidis (bd) causative agent (morphology and classification)  The genus Batrachochytrium is one of at least 100 genera in the Class Chytridiomycetes (chytrids). Most chytrid species are saprophytes, decomposing cellulose, chitin, keratin, other fungi, algae, and various types of plant material, although a few species are parasitic upon selected invertebrates such as insects (Sparrow 1960, Powell 1993, Longcore et al. 1999). Batrachochytrium dendrobatidis is the only species in this genus (Carey et al. 2006). ho s t r a nge and geo gr a phic distribution  Batrachochytrium dendrobatidis (Bd) infects at least 516 (42%) of 1,240 amphibian species among 35 of 40 families of Anura and six of eight families of Caudata (Olson et al. 2013), as well as four families of caecilians (Gymnophiona) (Gower et al. 2013). While it has been found on most land masses with amphibians, the parasite appears to be an emerging infectious disease and only recently has been found in most parts of the world (Weldon et al. 2004, Garner et al. 2005). The first known record for the fungus is from an African clawed frog (Xenopus laevis) collected in 1938 from South Africa; Bd may have originated in Africa and become disseminated through

international trade in X. laevis that began in the mid-1930s (Weldon et al. 2004). It now is identified in at least 52 of 82 countries sampled, on all major land masses (Olson et al. 2013). After Africa, continents with infected amphibians first were observed in 1961 in Québec, Canada (Ouellet et al. 2005), and shortly thereafter in Colorado (Ouellet et al. 2005), in 1978 in Europe (Garner et al. 2005) and Australia, in 1986 in Ecuador, in 1999 in New Zealand (Johnson and Speare 2003), in 2007 in Japan (http://www.promedmail .org/; http://www.asahi.com/english/Heraldasahi/TKY200706120087.html), and in 2013 in Southeast Asia (Gilbert et al. 2013). Despite a broad distribution around the world, this parasite has little evidence for genetic diversity among isolates collected in various locations (Morehouse et al. 2003). There is considerable variation of Bd prevalences and intensities among families of amphibians (Olson et al. 2013). life cycles and variations  This aquatic organism has at least two distinct life stages. A sessile monocentric thallus develops into a reproductive, host-dependent zoosporangium; then motile, uniflagellate, water-borne zoospores are released from the zoosporangium and serve as the infective stage to vertebrates (Berger et al. 2005a). After a period of up to 24 hours of motility, zoospores encyst on a susceptible host, resorb their flagella, and form stages called germlings; in turn, these form rhizoids and subsequently develop thalli that again grow and form mature zoosporangia over a few days. The contents of the enlarged thallus become multinucleate by mitotic divisions, and the entire contents divide again into the next generation of zoospores. Discharge tubes form in the sporangia. When the zoospores are mature, they are released as the plugs in the discharge tube liquefy (Berger et al. 2005a). The development of Bd is affected by ambient temperature and moisture. Under experimental conditions, optimal growth occurs between 17 and 25°C. Death of the pathogen occurs above 29°C, below 0°C, or after kingdom fungi   217

prolonged desiccation (Piotrowski et al. 2004); Bd can survive in tap water for 3 weeks, in deionized water for 4 weeks, and in lake water for up to 7 weeks after inoculation (Johnson and Speare 2003). In field studies, prevalence varies by season, elevation, or region (Ron 2005); increased prevalence is associated with cooler temperatures and moister conditions (Woodhams et al. 2003; Retallick et al. 2004; Kriger and Hero 2006, 2007). reservoirs and tr ansmission The fungi are transmitted through water (Kriger and Hero 2006). Among boreal toads (Bufo boreas), transmission can occur by contact with water containing zoospores; no physical contact with infected animals is required (Carey et al. 2006). Among yellow-legged frogs (Rana muscosa), tadpoles transmit the fungus among themselves and to postmetamorphic animals (Rachowicz and Vredenburg 2004). Transmission among yellow-legged frogs does not vary with degree of crowding or temperature (Rachowicz and Briggs 2007). Batrachochytrium dendrobatidis survives up to 3 months in sterile, moist river sand and can attach and grow on sterile feathers; it is proposed that this fungus may be translocated by movement of moist river sand, and birds may carry it between amphibian habitats (Johnson and Speare 2005). Waterfowl such as geese (Branta canadensis, Anser anser domesticus) also are able to carry the chytrid on their toes and probably transmit the fungus between sites (Garmyn et al. 2012). The parasite also has been identified among crayfish (Procambarus spp. and Orconectes virilis) of Louisiana and Colorado (McMahon et al. 2012). Some amphibians such as bullfrogs (Rana catesbeiana) (Hanselmann et al. 2004) and tiger salamanders (Ambystoma tigrinum) (Davidson et al. 2003) can carry this fungus on their epidermis without developing lethal infections and may serve as reservoirs for the transmission of this parasite to susceptible species (Daszak et al. 2004). The bullfrog is the most commonly farmed amphibian and regularly escapes and establishes feral populations; 218  kingdom fungi

it is speculated to be an important source for the spread and establishment of this fungus throughout the world (Garner et al. 2006). The Pacific chorus frog (Pseudacris regilla) also appears unaffected by Bd and survives at sites where other species have been extirpated (Reeder et al. 2012). Batrachochytrium dendrobatidis also can persist in an endemic state among healthy frog populations once an epidemic wave has passed through those populations (Retallick et al. 2004, McDonald et al. 2005). At some sites where Bd has been endemic for at least 10 years, it occurred in two species of stream frogs in very low prevalences, but was not present in any environmental samples of these sites (Rowley et al. 2007). In Québec, Canada, there was an overall 18% prevalence among amphibians in 1999 through 2001 among many apparently healthy populations, with no significant differences when compared to the prevalences of infection of 1960 through 1969 (Ouellet et al. 2005). This pathogen is widely distributed and apparently enzootic in seemingly healthy amphibians in parts of eastern North America (Ouellet et al. 2005). Infected frogs are more frequently associated with permanent than with temporary water bodies (Kriger and Hero 2007). Of these, stream breeders had a higher prevalence of infection than pond breeders; however, the intensity of frog infections does not differ significantly between these groups (Kriger and Hero 2007). Based on ecological niche modeling for Bd, suitable habitats in the New World are extensive, including pine–oak forests, dry forests, moist forests, temperate forests, and tropical rainforests throughout the United States, Central America, South America, and the Caribbean (Ron 2005). Also, for Old World localities, Bd was found in 56 of 59 habitats with high predicted suitability (Ron 2005). clinical effects and identification  This fungus infects only keratinized tissues of amphibians (Daszak et al. 1999, Longcore et al. 1999, Pessier et al. 1999), and interferes with

normal skin function of susceptible amphibian species, leading to disruption of osmoregulation, subsequent electrolyte imbalance, and eventual death (Berger et al. 1998, Voyles et al. 2009). The disease, chytridiomycosis, often involves sloughing of dermatitis (Longcore et al. 1999, Pessier et al. 1999). Although amphibian larvae lack keratin in their epidermis, the fungus occurs in the keratinized mouthparts of tadpoles and toes of premetamorphic tadpoles in some species (Berger et al. 1998, Fellers et al. 2001, Marantelli et al. 2004, Rachowicz and Vredenburg 2004). There is considerable interspecific variation in susceptibility to this parasite among frog tadpoles (Blaustein et al. 2005). Unique Bd-specific genes containing pathogenic factors have been identified (Joneson et al. 2011). Despite low genetic diversity (Morehouse et al. 2003), there are some strain differences among Bd regarding impacts on amphibians (Retallick and Miera 2007). Among postmetamorphic boreal toads (Bufo boreas), the time between exposure and death is influenced by the dose of infectious zoospores, duration of exposure, and body size of the toad (Carey et al. 2006); about 107 to 108 sporangia appears to be a minimum threshold needed to cause death in boreal toads. Once infected, variation in air temperature between 12 and 23°C had no significant effect on survival time among infected animals (Carey et al. 2006). Among adult Australian green tree frogs (Litoria caerulea), large numbers of sporangia occurred in all areas of ventral skin and toes; few or no sporangia occurred on dorsal skin. This difference may be due to the dryness of the dorsal skin or the greater numbers of serous glands there, which produce antifungal peptides (Berger et al. 2005b). Thus sampling of skin cells typically is made on ventral surfaces and on the toes of tested animals. There are several studies on methods to identify Bd in amphibians (Brem et al. 2007, Hyatt et al. 2007). Polymerase chain-reaction (PCR)–based assays are used to detect both zoospores and infections in skin samples (Annis et al. 2004, Boyle et al. 2004). Although less

sensitive than some other methods, a PCR test also can be used for non-lethal detection of Bd from swabbed tadpole mouth parts (Retallick et al. 2006). Congo red is effective for finding zoosporangia, zoospores, and germling stages of Bd in epidermal skin swabs of frogs (Briggs and Burgin 2004). Using an indirect immunoperoxidase test, polyclonal antibodies can be used for diagnosing Bd in amphibians by staining walls, cytoplasm, rhizoids, and zoospores (Berger et al. 2002). This method has been combined with Hollande’s Trichrome keratin stain to simultaneously detect Bd and keratin (Olsen et al. 2004). popul ation effects Batrachochytrium dendrobatidis has been associated with extirpations and worldwide population declines in many wild amphibian populations (Daszak et al. 2003, Stuart et al. 2004, Johnson 2006). In several studies, healthy amphibian populations occurred at sites in the absence of Bd, while acute mortalities and population declines occurred immediately after detection of the pathogen (Rachowicz et al. 2005, Lips et al. 2006). Thus the impacts are caused by long-lived or saprophytic free-living stages of this pathogen, and there is evidence that Bd has caused direct extinctions of many amphibians (Schloegel et al. 2006, Mitchell et al. 2008). Epizootics in Central and South America are explained by multiple introductions of Bd because the epizootics move across the affected regions in the wave-like pattern characteristic of many disease systems (Lips et al. 2008). However, not all are convinced that Bd is the primary cause of mortality, even in regions where it is commonly reported (Burgin et al. 2005). The role of weather also has been assessed. Weather-driven simulations of pathogen growth potential with Pearson’s green tree frog (Litoria pearsoniana) were positively related to both prevalence and intensity of Bd infections (Murray et al. 2013). Also, high rainfall, numbers of rain days, and temperatures between 3 and 30°C were positively correlated with infection levels of the quacking frog (Crinia kingdom fungi   219

georgiana) in Australia (Riley et al. 2013). More generally, global warming and contaminants also may exacerbate chytrid outbreaks (Pounds et al. 2006). special problems  Batrachochytrium dendrobatidis is likely a significant contributor to the amphibian declines in many parts of the world, including North America (Bradley et al. 2002, Green et al. 2002), Europe (Bosch et al. 2001), South America (Ron and Merino 2000, Ron et al. 2003, Lampo et al. 2007), Central America (Berger et al. 1998, Lips et al. 2006), the Caribbean (Lips et al. 2003, Alemu et al. 2008), Australia (Berger et al. 1998), and New Zealand (Bishop 2000). This is further addressed in Case Study on Amphibian Declines, Chapter 11. The parasite appears to follow a densityindependent pattern of spread, which increases extinction risk in affected populations (Collins 2010), especially among critically endangered species such as the harlequin frog (Atelopus mucubajiensis) (Lampo et al. 2007), Bloody Bay frog (Mannophryne olmonae) (Alemu et al. 2008), and the Sardinian newt (Euproctus platycephalus, Order Urodela) (Bovero et al. 2008). Interestingly, this fungus has been considered as a means of controlling an invasive frog (Eleutherodactylus coqui) on Hawaii. Such an introduction may be of less risk to other native fauna because there are no native amphibians on Hawaii (Beard and O’Neill 2005). control  Amphibians have a variety of antimicrobial defenses including skin peptides that inhibit the growth of mature B. dendrobatidis cells (Woodhams et al. 2007). While antimicrobial peptides often are effective at both 10°C and 22°C, many are more effective at 10°C (RollinsSmith et al. 2002). Some infected frogs can clear their own B. dendrobatidis infections entirely (Kriger and Hero 2006). Salamanders also have antimicrobial peptides that inhibit B. dendrobatidis and other microorganisms (Sheafor et al. 2008). At least eight genera of cutaneous bacteria found on amphibians inhibit the growth of B. dendrobatidis (Harris et al. 2006). For 220  kingdom fungi

example, Pedobacter cryoconitis, a bacterium found on the skin of red-backed salamanders (Plethodon spp.), lessens the effects of Bd when transferred to the skin of yellow-legged frogs, (Lam et al. 2008). Bioaugmentation of individual amphibians and of amphibian habitats with carefully selected, locally occurring, anti-chytrid microbes may be of potential value in areas under threat from Bd. Several sampling strategies and filtering protocols to identify promising probiotics have been proposed (Bletz et al. 2013). Manipulating environmental conditions can influence success of this fungus. Low temperatures, toxic chemicals, and stress inhibit the immune system and may impair natural defenses against B. dendrobatidis (RollinsSmith et al. 2011). Infected red-eyed tree frogs (Litoria chloris) artificially held at 37°C can be cleared of B. dendrobatidis in less than 16 hours; thus elevated body temperatures to clear frogs of chytrid infection might be used to eliminate the fungal pathogen from captive populations and reduce the likelihood of accidental spread when animals are translocated or released from captivity (Woodhams et al. 2003). In another study where frogs were cleared of infection by the use of heat, mortality ceased and the frogs gained weight; in contrast, frogs in which infection was not cleared gained less weight or continued to die (Retallick and Miera 2007). Among laboratory-infected yellow-legged frogs, fewer frogs died when housed at 22°C than at 17°C; since both temperatures were within the optimal range for growth of B. dendrobatidis, the authors proposed that the difference was from the effect of temperature on the host’s resistance to chytridiomycosis rather than any effect on the fungus alone (Andre et al. 2008). In the laboratory, a number of chemicals, including Path-X TM, the quaternary ammonium compound 128, and itraconazole (Jones et al. 2012), are effective to disinfect in vitro cultures of Bd. In contrast, UV light was ineffective at the wavelengths tested. Under experimental conditions, cultures were sensitive to heating for 4 hours at 37°C, 30 min at 47°C,

and 5 min at 60°C (Johnson et al. 2003). Such treatments are applicable to prevent spread by cleaning items contacting amphibians or water (e.g., nets, boots), in captive husbandry, and in the laboratory. More broadly, the worldwide emergence of this fungal disease has been linked to the spread of infected animals, introducing nonnative infected animals into naïve populations, and amplifying infections of amphibians by co-housing of mixed populations, as well as discharge of wastewater containing untreated discharge of infectious zoospores into natural water supplies. In general, the trade in amphibians probably has been an important contributor to the wide distribution of this parasite, and special effort must be made to prevent introduction of this pathogen into remaining uninfected areas (Fisher and Garner 2007). Effective management of chytridiomycosis will depend on different countries and regions recognizing the disease as a “threatening process”, defined as a disease that threatens, or may threaten the survival, abundance, or evolutionary development of a native species or ecological community, and that requires strategies being implemented for its control (Hyatt et al. 2007). Managerial strategies ultimately will involve detection of infected populations of both laboratory-housed and free-ranging animals, identification of infected geographical areas, and control of human-mediated movement of animals from one location to another (Hyatt et al. 2007). Needed steps include identification of the parasite in adults and tadpoles from captive and free-ranging animals, estimation of prevalence of infection of these populations, identifying specific infected animals or groups for the purposes of control, identifying diseasefree zones, and demonstrating the eradication of infections from individuals undergoing treatment (Hyatt et al. 2007). Raising and maintaining chytrid-free laboratory populations of vulnerable species until there are mechanisms for safely introducing them back into natural habitats also is being attempted (Stone 2013). Hundreds of frogs were

collected from El Valle, an inactive volcano in Panama (Goodman 2006). The Houston Zoo, with the assistance of other zoological groups, is constructing an ex situ facility in Panama’s El Valle de Anton region to house Atelopus zeteki and several other native species, serving as a repository and conservation breeding center, a treatment facility, and a nature education center for Panamanians and foreign tourists (www.houstonzoo.org/Golden_Frogs.aqf). pseudogymnoascus (geomyces) destructans [white-nose syndrome (geomycosis)] causative agent  Pseudog ymnoascus (Geomyces) destructans (Order Onygenales, Family Onygenaceae) is a recently described psychrophilic (cold-loving) ascomycete (App. 1: Table 6) (Blehert et al. 2009). It causes, white-nose syndrome (WNS), a fatal disease in many hibernating bats (Order Chiroptera) (Gargas 2009, Lorch et al. 2011). host r ange and geogr a phic distribution  In North America, the fungus has been identified in at least 19 states throughout the northeastern and mid-Atlantic regions of the United States as well as the provinces of Ontario and Québec in Canada (Frick et al. 2010, Cryan 2011). At least six bat species of North America currently are known to be adversely affected by WNS, including four species of Myotis (M. leibii, M. lucifugus, M. septentrionalis, and M. sodalis), Perimyotis subfalvus, and Eptesicus fuscus (Cryan 2011). It is possible that many of the 25 species of hibernating bats in North America could be vulnerable to P. destructans (Cryan 2011, Foley et al. 2011). The fungus is also widespread in Europe and has been observed in at least six species of Myotis in eight countries (Martínková et al. 2010, Puechmaille et al. 2010, Wibbelt et al. 2010a, Šimonovičov et al. 2011). However, despite extensive monitoring, no major mortality events have been documented among European bats (Puechmaille et al. 2011). This intercontinental difference is associated with differences in susceptibility of resident bats kingdom fungi   221

in these regions rather than differences in the pathogen (Cryan et al. 2013). reservoirs and tr ansmission As of 2011, only bats were reported as hosts for P. destructans. There is evidence that the parasite in North America originated in Europe and was transported by human trade or travel (Warnecke et al. 2012). Transmission occurs through direct bat-to-bat contact (Lorch et al. 2011) but also may occur by point source infections from exposure to soil in which fungi are present (Lindner et al. 2010). The fungus has been called a host-generalist pathogen with an abiotic reservoir in caves (Eskew and Todd 2013). Spread of the fungus to new geographic regions and to other vulnerable bat species may result from social and spatial mixing of bats during migration or by social contact, including mixed groups, during hibernation or some more long-distance movements (Frick et al. 2010, Lorch et al. 2011). The role of soil as a reservoir for P. destructans is not clear. The fungus has been isolated from soil of hibernacula in areas where whitenose syndrome occurs; there is a very abundant and diverse group of the closely related Geomyces spp. cultured among soil organisms sampled in bat hibernacula, many of which are undescribed taxa (Lorch et al. 2013). clinical effects and identification  While some species of Geomyces can colonize skin (Gianni et al. 2003, Foley et al. 2011), P. destructans can invade, digest, and erode the skin of hibernating bats (Meteyer et al. 2009). Known characteristics of WNS include a white filamentous or powdery growth on the nose, ears, and wing membranes, and emaciation, (Blehert et al. 2009); early emergence from hibernacula in mid-winter (Wibbelt et al. 2010b); and ulcerated, necrotic, and scarred wing membranes among bats recently emerged from hibernation (Reichard and Kunz 2009). The disease also may be associated with elevated metabolism of bats, reduced flora in the their digestive tracts, and some disruptions in their immune systems (Turner and Reeder 2009). Mortality is linked to rapid depletion 222  kingdom fungi

of fat reserves during hibernation (Boyles and Willis 2009), disruption of wing-dependent physiological functions after infections (Cryan et al. 2010), and by evaporative water loss (Willis et al. 2011). Mortality from WNS does not become evident until about 120 days after bats enter hibernation and assume a cold physiological state conducive to proliferation of P. destructans; mortality peaks about 180 days after bats first enter hibernacula (about March) (Lorch et al. 2011). But with supportive laboratory care, infected bats can recover from infection (Meteyer et al. 2011). Many hibernating animals have a cycle of suppression of cellular immune response during hibernation and reactivation of the immune response after hibernation (Bouma et al. 2010). The lack of a visible cellular immune response to P. destructans during hibernation, with subsequent neutrophil recruitment and sequestration of P. destructans in homeothermic bats, suggests that bats also have this type of immune regulation (Meteyer et al. 2011). Such a model makes them vulnerable to infections at low temperatures and, while their immune response is enhanced on becoming homeothermic, it calls upon considerable expenditure of fat reserves to establish the body temperatures needed for effective response. Some mortality is associated with an immune reconstitution inflammatory syndrome, in which immune suppression during hibernation is followed by an intense neutrophil inflammatory response to the P. destructans, resulting in severe pathology and mortality (Meteyer et al. 2012). Diagnosis of the disease is based on histopathology (Meteyer et al. 2009), polymerase chain reaction tests, and laboratory culture (Lorch et al. 2010, Chaturvedi et al. 2011, Foley et al. 2011). population effects First observed in North America near Albany, New York, in February 2006 (Blehert et al. 2009), this explosive disease has caused unprecedented reductions in the abundance of hibernating bat species in the eastern United States, with up to

95% mortality in some hibernacula and an estimated mortality of 6 million or more bats dying from WNS (Froschauer and Coleman 2012). Bat population declines in the northeastern United States since the emergence of WNS may exceed 80 percent (Turner et al. 2011). Some winter colonies that were stable or increasing in numbers for decades have virtually disappeared (Reichard and Kunz 2009); the little brown bat (Myotis lucifugus) has been particularly severely affected (Frick et al. 2010). Population decreases at infected hibernacula range from 30 to 99% annually, with a regional mean of 73%, and all surveyed sites have become infected within 2 years of the disease arriving in their region (Frick et al. 2010). Since its original identification as a disease, WNS has expanded at least 2,000 km westward in North America (Cryan 2011). Because most affected bat species are long-lived and have only one offspring per year, bat populations affected by WNS are not expected to recover quickly (Cryan 2011). About 25 of the 45 insectivorous bat species in the United States and Canada rely on hibernation as a primary strategy to over-winter, including four species and subspecies listed as endangered and an additional 13 listed as federal species of concern (Cryan 2011). The increasing impact of this pathogenic fungus in hibernating bats potentially could undermine the basic survival strategy of over half of the bat species in the United States and 18 of the 22 bat species living above 40°N in North America (Cryan 2011). special problems  The natural cycle of bat hibernation, with reduced metabolism and immune function, has allowed P. destructans to become very successful in bats (Cryan et al. 2010). Pseudogymnoascus (Geomyces) destructans thrives in darkness, low temperatures (5–10°C), and high humidity (>90%), and it cannot grow above 20°C (Cryan 2011); thus it appears to be well suited to persist in the hibernacula of bat caves and mines. It establishes itself in the bats and invades wing membranes and other skin tissue when the bat body temperatures are lowered

to 2–10°C during hibernation (Cryan 2011). Fungal infiltration of the wing membranes of bats may be problematic because the wing surfaces cover about 85% of the bat’s total surface area, and healthy wing membranes are vital to regulation of body temperature, blood pressure, water balance, and gas exchange (Cryan 2011). Behavioral strategies of bats also may contribute to their vulnerability to infections. For example, selection of humid areas of hibernacula or dense clustering to conserve energy and decrease moisture loss could further enhance fungal colonization, growth, and conidial amplification by elevating humidity, as well as by increasing infection prevalence and dispersal of P. destructans through increased contact with infected individuals (Cryan et al. 2010). Finally, the natural reduction of immune function in hibernating species may allow the fungus to invade body tissues without facing a significant immune response (Bouma et al. 2010). Bats play a very important role in North American ecosystems through predation of nocturnal insects, including many crop and forest pests. Economic impacts on agriculture from loss of bats is estimated at about $22.9 billion per year, with a range of extremes $3.7 to $53 billion per year (Boyles et al. 2011). These estimates incorporate the reduced costs of pesticide applications not needed to suppress insects consumed by bats (Cleveland et al. 2006). However, such estimates do not include suppression of forest insects, which also is believed to be considerable (Kalka et al. 2008). control  There are no known strategies to reduce the spread or to control WNS in bats. There is evidence that bat hibernacula closer to the site of fungal origin, as well as those of larger size, tend to have higher risk for infection (Wilder et al. 2011). One proposal for reducing WNS impacts includes culling of bats (Szymanski et al. 2009); however, disease models do not support this approach (Hallam and McCracken 2011). Other suggestions for control include providing artificial localized warm areas inside cold caves (Boyles and Willis 2009), treatment of individual bats, enhancing kingdom fungi   223

resistance of key populations through vaccines or immunomodulators, nutritional support to reduce starvation or dehydration from the effects of the disease, modifying hibernacula environments to reduce P. destructans, and informing the public to reduce anthropogenic spread (Foley et al. 2011). Most efforts have focused on implementing universal precautions, including restricting access of humans to sensitive bat hibernation sites and decontaminating equipment and clothing when sites are accessed for disease surveillance, research, or recreational purposes (Blehert 2012). miscell aneous dermatophy tes  Three other genera of dermatophytes generally are recognized: Trichophyton, Microsporum, and Epidermophyton; all are ascomycetes in the Family Arthodermataceae, Order Onygenales (App. 1: Table 6) (Kane and Summerbell 1999). Some species in each genus infect humans (anthropophilic), with little or no transmission to other species; some species of Trichophyton and Microsporum are pathogens primarily of nonhuman mammals or birds (zoophilic), whereas other species of these two genera are primarily soil-associated organisms (geophilic) that only occasionally infect animal hosts (Kane and Summerbell 1999). Trichophyton gallinae is the cause of ringworm; it also is called fowl flavus in wild birds and poultry (Friend 1999b). Trichophyton mentagrophytes, another ringworm fungus, has been reported among wild foxes (Vulpes vulpes) (Knudtson et al. 1980), opossums (Didelphis marsupialis) (Menges and Georg 1955, McKeever et al. 1958), and as a serious problem among young muskrats (Erethizon zibethicus) (Errington 1963). Among muskrats, there was an 8–12% prevalence among the litters studied, and mortality of about 50–60% among infected young; infected muskrats usually had hairless patches and a dandruff-like scurf (Errington 1963). Although infected muskrats were observed only in Iowa and Maryland, this fungus was assumed to be widespread over North America (Errington 1963). There also 224  kingdom fungi

are both zoophilic and anthropophilic strains of this species (Knudtson et al. 1980). Infections among red foxes (Vulpes vulpes) were successfully treated with griseofulvin in the feed (Knudtson et al. 1980). Transmission of infectious dermatophytes generally occurs by direct contact with carrier animals or contaminated fomites (Kane and Summerbell 1999, Burek 2001). In addition, chewing and sucking lice can be mechanical vectors in some cases (Durden and Musser 1994). Animal host infections typically require some slight trauma or continued moisture or maceration of the skin for the parasites to become established (Burek 2001). Skin lesions typically begin as focal, round areas of hair or feather loss and may progress to redness, skin depigmentation, and excess keratin production (Burek 2001). Diagnosis often is based on clinical lesions and finding arthroconidia or hyphae in skin scrapings digested in 10% potassium hydroxide (Kane and Summerbell 1999). For captive wildlife, systemic therapies include griseofulvin and other drugs (Knudtson et al. 1980, Burek 2001). Most infections are self-limiting, with lesions requiring a few weeks to several months to regress, depending on the species of fungus and host, and individual host responses (Burek 2001).

MYCOTOXINS Mycotoxin is a general term for a variety of diverse but potent toxins produced by fungi that affect birds, mammals, and other vertebrates (Quist et al. 2007). There are a number of mycotoxins of importance to vertebrates (Bennett and Klich 2003). For example, aflatoxins are produced by several species of Aspergillus, notably A. flavus. Citrinin is produced by several species of both Penicillium and Aspergillus. Ergot alkaloids are produced in species of Claviceps, which are common pathogens of various grass species. Fumonisins are produced by a number of Fusarium species, including F. verticillioides, as well as Alternaria alternaria; Fusarium verticillioides is a common corn

endophyte. Ochratoxin is produced by several species of Aspergillus, including A. niger. Patulin is produced by a number of different molds, including Penicillium. Trichotecenes is a family of metabolites produced by a number of genera, including Fusarium, Myrothecium, Phomopsis, Stachybotrys, Trichoderma, and Trichothecium. Although classified as a mycotoxin by some, zearalenone is a secondary metabolite from Fusarium graminearum better classified as a nonsteroidal estrogen or mycoestrogen (Bennett and Klich 2003). Most genera of fungi producing mycotoxins are found in the Phylum Ascomycetes. Types of mycotoxicosis in freeliving birds include aflatoxins as well as trichothecenes (Creekmore 1999). In contrast to mycoses, which are infections, mycotoxicoses are pathologies analogous to poisonings by pesticides or heavy metal residues (Bennett and Klich 2003). Like mycoses, mycotoxins generally are not communicable from host to host (Bennett and Klich 2003). Aflatoxins are used as an example of a mycotoxin. aflatoxins (aspergillus flavus) c aus at i v e agen t (cl a ssific at ion, morphology)  Aflatoxins are one of at least nine types of closely related mycotoxins produced by members of the genus Aspergillus spp. (Pier and Richard 1992). The term “af latoxin” is an acronym for A spergillus f lavus toxin (O’Hara 1996). The best-known examples of aflatoxins are produced by Aspergillus f lavus and A. parasiticus (Creekmore 1999). A number of aflatoxin types, including B1, B2, G1, G2, M1, and M2, are important for wildlife (Quist et al. 2007). Structurally, aflatoxins are ringed compounds consisting of a coumarin nucleus fused to a bifuran, and either a pentanone (for B1 or B2) or lactone (for G1 or G2) ring (Palmgren and Hayes 1987). Aflatoxin B1 is the most prevalent and most toxic (Moss 2002), and the most common cause of mycotoxicoses (Quist et al. 2007). Although host susceptibility varies considerably among species (Arafa et al. 1981), no vertebrate species appears to be completely resistant to aflatoxins (Quist et al. 2007). Among free-living

birds, aflatoxins most commonly are reported among waterfowl (Robinson et al. 1982, Quist et al. 2007). Galliform birds also are susceptible (Ruff et al. 1990, 1992; Quist et al. 2007), as are some fish (Hussain et al. 1993). Most reports of aflatoxin have been from wild birds, and from Texas (Robinson et al. 1982), with some observations among wild passerine birds of Great Britain (Lawson et al. 2006) and in eggs of green sea turtles (Chelonia mydas) of Oman (Elshafie et al. 2007). There is little information about aflatoxicosis among wild mammals. Most mycotoxicoses result from eating contaminated foods, although skin contact with mold-infested substrates and inhalation of spore-borne toxins can be important sources of exposure (Bennett and Klich 2003). Aflatoxins can reduce immune response and increase the severity of avian diseases such as coccidiosis (Edds et al. 1973, Rao et al. 1990). Aflatoxin ingestion also increases susceptibility to the organophosphate malathion (Ehrlich et al. 1985), as well as to Salmonella spp., Candida spp., and Treponema spp. Immunity to Pasteurella multocida also is impaired (Pier 1992), and aflatoxin ingestion may exacerbate avian mortality events such as avian cholera among wild birds (Smith et al. 1990). Literature Cited Acha, P. N., and B. Szyfres. 2001. Zoonoses and communicable diseases common to man and animals. Vol. I: Bacterioses and mycoses. 3rd ed. Report No. 580. Pan American Health Organization, Washington, DC. Adaska, J. M. 1999. Peritoneal coccidioidomycosis in a mountain lion in California. Journal of Wildlife Diseases 35:75–77. Adl, S. M., A. G. B. Simpson, M. A. Farmer, R. A. Andersen, O. R. Andersen, J. R. Barta, S. S. Bowser, G. Brugerolle, R. A. Fensome, S. Fredericq, T. Y. James, S. Karpov, P. Kugrens, J. Krug, C. E. Lane, L. A. Lewis, J. Lodge, D. H. Lynn, D. G. Mann, R. M. McCourt, L. Mendoza, Ø. Moestrup, S. E. Mozley-Standridge, T. A. Nerad, C. A. Shearer, A. V. Smirnov, F. W. Spiegel, and M. F. J. R. Taylor. 2005. The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. Journal of Eukaryotic Microbiology 52:399–451.

kingdom fungi   225

Adrian, W. J., T. R. Spraker, and R. B. Davies. 1978. Epornitics of aspergillosis in mallards (Anas platyrhynchos) in north central Colorado. Journal of Wildlife Diseases 14:212–217. Ajello, L. 1964. Relationship of Histoplasma capsulatum to avian habitats. Public Health Reports 79:266–270. Ajello, L. 1967. Comparative ecology of respiratory mycotic agents. Bacteriological Reviews 31:6–24. Alemu, J. B., M. N. E. Cazabon, L. Dempewolf, A. Hailey, R. M. Lehtinen, R. P. Mannette, K. T. Naranjit, and A. C. J. Roach. 2008. Presence of the chytrid fungus Batrachochytrium dendrobatidis in populations of the critically endangered frog Mannophryne olomonae in Tobago, West Indies. EcoHealth 5:34–39. Alexopoulos, C. J., C. W. Mims, and M. Blackwell. 1996. Introductory mycology. 4th ed. John Wiley & Sons, New York. Andre, S. E., J. Parker, and C. J. Briggs. 2008. Effect of temperature on host response to Batrachochytrium dendrobatidis infection in the mountain yellow-legged frog (Rana muscosa). Journal of Wildlife Diseases 44:716–720. Annis, S. L., F. P. Dastoor, H. Ziel, P. Daszak, and J. E. Longcore. 2004. A DNA-based assay identifies Batrachochytrium dendrobatidis in amphibians. Journal of Wildlife Diseases 40:420–428. Anonymous. 1975. Blackbird control on two Army Installations: Fort Campbell, Kentucky and Milan, Tennessee AAP, Environmental Impact Statement. U.S. Army, Washington, DC. Arafa, A. S., R. J. Bloomer, H. R. Wilson, C. F. Simpson, and R. H. Harms. 1981. Susceptibility of various poultry species to dietary aflatoxins. British Poultry Science 22:431–436. Atasever, A., and K. S. Gumussoy. 2004. Pathological, clinical and mycological findings in experimentally aspergillosis infections in starlings. Journal of Veterinary Medicine Series A 51:19–22. Austwick, P. K. C. 1968. Mycotic infections. Symposia of the Zoological Society of London 24:249–271. Bair, W. C., S. G. Simpson, and R. M. Windingstad. 1988. Acute aspergillosis in mallards at Oahe Seep near Pierre, South Dakota. Prairie Naturalist 20:153–156. Baptista-Rosas, R., A. Hinojosa, and M. Riquelme. 2007. Ecological niche modeling of Coccidioides spp. in western North American deserts. Annals of the New York Academy of Sciences 111:35–46. Barker, B. M., K. A. Jewell, S. Kroken, and M. J. Orbach. 2007. The population biology of Coccidioides: Epidemiologic implications for

226  kingdom fungi

disease outbreaks. Annals of the New York Academy of Sciences 1111:147–163. Barr, D. J. S. 1992. Evolution and kingdoms of organisms from the perspective of a mycologist. Mycologia 84:1–11. Beard, K. H., and E. M. O’Neill. 2005. Infection of an invasive frog Eleutherodactylus coqui by the chytrid fungus Batrachochytrium dendrobatidis in Hawaii. Biological Conservation 126:591–595. Beer, J. V. 1963. The incidence of Aspergillus fumigatus in the throats of wild geese and gulls. Sabouraudia 2:238–247. Bellrose, F. C. 1945. Aspergillosis in wood ducks. Journal of Wildlife Management 9:325–326. Bennett, J. W., and M. Klich. 2003. Mycotoxins. Clinical Microbiology Reviews 16:497–516. Berger, L., A. D. Hyatt, V. Olsen, S. G. Hengstberger, D. Boyle, G. Marantelli, K. Humphreys, and J. E. Longcore. 2002. Production of polyclonal antibodies to Batrachochytrium dendrobatidis and their use in an immunoperoxidase test for chytridiomycosis in amphibians. Diseases of Aquatic Organisms 48:213–220. Berger, L., A. D. Hyatt, R. Speare, and J. E. Longcore. 2005a. Life cycle stages of the amphibian chytrid Batrachochytrium dendrobatidis. Diseases of Aquatic Organisms 68:51–63. Berger, L., R. Speare, P. Daszak, D. E. Green, A. A. Cunningham, C. L. Goggin, R. Slocombe, M. A. Ragan, A. D. Hyatt, K. R. McDonald, H. B. Hines, K. R. Lips, G. Marantelli, and H. Parkes. 1998. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proceedings of the National Academy of Sciences of the United States of America 95:9031–9036. Berger, L., R. Speare, and L. F. Skerratt. 2005b. Distribution of Batrachochytrium dendrobatidis and pathology in the skin of green tree frogs Litoria caerulea with severe chytridiomycosis. Diseases of Aquatic Organisms 68:65–70. Bernardes-Engemann, A. R., R. C. O. Costa, B. P. Miguens, C. V. L. Penha, E. Neves, B. A. S. Pereira, C. M. P. Dias, M. Mattos, M. C. Gutierrez, A. Schubach, M. P. O. Neto, M. Lazéra, and L. M. Lopes-Bezerra. 2005. Development of an enzyme-linked immunosorbent assay for the serodiagnosis of several clinical forms of sporotrichosis. Medical Mycology 43:487–493. Bialek, A. Feucht, C. Aepinus, F. Just-Nübling, V. J. Robertson, J. Knobloch, and R. Hohle. 2002. Evaluation of two nested PCR assays for detection of Histoplasma capsulatum DNA in human tissue. Journal of Clinical Microbiology 40:1644–1647.

Bishop, P. 2000. Chytrid fungi identified from dying frogs in New Zealand. Froglog 39:1–2. Blaustein, A. R., J. M. Romansic, E. A. Scheessele, B. A. Han, A. P. Pessier, and J. E. Longcore. 2005. Interspecific variation in susceptibility of frog tadpoles to the pathogenic fungus Batrachochytrium dendrobatidis. Conservation Biology 19:1460–1468. Blehert, D. S. 2012. Fungal disease and the developing story of bat white-nose syndrome. Public Library of Science Pathogens 8:e1002779. Blehert, D. S., A. C. Hicks, M. Behr, C. U. Meteyer, B. M. Berlowski-Zier, E. L. Buckles, J. T. H. Coleman, S. R. Darling, A. Gargas, R. Niver, J. C. Okoniewski, R. J. Rudd, and W. B. Stone. 2009. Bat white-nose syndrome: An emerging fungal pathogen? Science 323:227. Bletz, M. C., A. H. Loudon, M. H. Becker, S. C. Bell, D. C. Woodhams, K. P. C. Minbiole, and R. N. Harris. 2013. Mitigating amphibian chytridiomycosis with bioaugmentation: Characteristics of effective probiotics and strategies for their selection and use. Ecology Letters. doi: 10.1111/ele.12099. Borelli, D. 1961. Lobomicosis experimental. Dermatologia Venezolana 3:72–82. Bosch, J., I. Martínez-Solano, and M. García-Paris. 2001. Evidence of a chytrid fungus infection involved in the decline of the common midwife toad (Alytes obstetricans) in protected areas of central Spain. Biological Conservation 97:331–337. Bouma, H. R., H. V. Carey, and F. G. M. Kroese. 2010. Hibernation: The immune system at rest? Journal of Leukocyte Biology 88:1–6. Bovero, S., G. Sotgiu, C. Angelini, S. Doglio, E. Gazzaniga, A. A. Cunningham, and T. W. J. Garner. 2008. Detection of chytridiomycosis caused by Batrachochytrium dendrobatidis in the endangered Sardinian newt (Euproctus platycephalus) in southern Sardinia, Italy. Journal of Wildlife Diseases 44:712–715. Bowes, V. A. 1990. An outbreak of aspergillosis in wild waterfowl. Canadian Veterinary Journal 31:303–304. Boyle, D. G., D. B. Boyle, V. Olsen, J. A. T. Morgan, and A. D. Hyatt. 2004. Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatidis) in amphibian samples using real-time Taqman PCR assay. Diseases of Aquatic Organisms 60:141–148. Boyles, J. G., P. M. Cryan, G. F. McCracken, and T. H. Kunz. 2011. Economic importance of bats in agriculture. Science 332:41–42. Boyles, J. G., and C. K. R. Willis. 2009. Could localized warm areas inside cold caves reduce



mortality of hibernating bats affected by white-nose syndrome? Frontiers in Ecology and the Environment 7:92–98. Bradley, G. A., P. C. Rosen, M. J. Sredl, T. R. Jones, and J. E. Longcore. 2002. Chytridiomycosis in native Arizona frogs. Journal of Wildlife Diseases 38:206–212. Brand, C. J., R. M. Windingstad, L. M. Siegfried, R. M. Duncan, and R. M. Cook. 1988. Avian mortality from botulism, aspergillosis, and salmonellosis at Jamaica Bay Wildlife Refuge, New York, U.S.A. Colonia Waterbirds 11:284–292. Brem, F., I. Joseph T. Mendelson, and K. R. Lips. 2007. Field-sampling protocol for Batrachochytrium dendrobatidis from living amphibians, using alcohol preserved swabs, Version 1.0. Conservation International, Arlington, VA. . Briggs, C., and S. Burgin. 2004. Congo red, an effective stain for revealing the chytrid fungus, Batrachochytrium dendrobatidis, in epidermal skin scrapings from frogs. Mycologist 18:98–103. Brun, A. M. 1999. Lobomycosis in three Venezuelan patients. International Journal of Dermatology 38:298–305. Bultman, M. W., F. S. Fisher, and M. E. Gettings. 2004. Coccidioidomycosis: Mitigating the risk. GeoHealth News 3:2–6. Burek, K. 2001. Mycotic diseases. Pp. 514–531 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Burgin, S., C. B. Schell, and C. Briggs. 2005. Is Batrachochytrium dendrobatidis really the proximate cause of frog decline? Acta Zoologica Sinica 51:344–348. Burns, R. A., J. S. Roy, C. Woods, A. A. Padhye, and D. W. Warnock. 2000. Report of the first human case of lobomycosis in the United States. Journal of Clinical Microbiology 38:1283–1285. Caldwell, D. K., M. C. Caldwell, J. C. Woodard, L. Ajello, W. Kaplan, and H. M. McClure. 1975. Lobomycosis as a disease of the Atlantic bottle-nosed dolphin (Tursiops truncatus Montagu, 1821). American Journal of Tropical Medicine and Hygiene 24:105–114. Carey, C., J. E. Bruzgul, L. J. Liva, M. L. Walling, K. A. Kuehl, B. F. Dixon, A. P. Pessier, R. A. Alford, and K. B. Rogers. 2006. Experimental exposures of boreal toads (Bufo boreas) to a pathogenic chytrid fungus (Batrachochytrium dendrobatidis). EcoHealth 3:5–21. Carrasco, L., M. J. Bautista, J. M. deLasMulas, and H. E. Jensen. 1993. Application of enzyme-immunohistochemistry for the

kingdom fungi   227

diagnosis of aspergillosis, candidiasis and zygomycosis in three lovebirds. Avian Diseases 37:923–927. Chandler, F. W., and J. C. Watts. 1987. Pathologic diagnosis of fungal infections. American Society of Clinical Pathologists, Chicago, IL. Chaturvedi, S., R. J. Rudd, A. Davis, T. R. Victor, X. Li, K. A. Appler, S. S. Rajkumar, and V. Chaturvedi. 2011. Rapid real-time PCR assay for culture and tissue identification of Geomyces destructans: The etiologic agent of bat geomycosis (white nose syndrome). Mycopathologia 172:246–256. Chute, H. L., S. L. Hollander, E. S. Barden, and D. C. O’Meara. 1965. The pathology of mycotoxicosis of certain fungi in chickens. Avian Diseases 9:57–66. Clemons, K. V., R. Laniado-Laborin, and D. A. Stevens. 2007. Coccidioidomycosis: Sixth International Symposium. Annals of the New York Academy of Sciences 1111:1–462. Cleveland, C. J., M. Betke, P. Federico, J. D. Grank, T. G. Hallam, J. Horn, J. López, Juan D., G. F. McCracken, R. A. Medillín, A. Moreno-Valdez, C. G. Sansone, J. K. Westbrook, and T. H. Kunz. 2006. Economic value of pest control service provided by Brazilian free-tailed bats in south-central Texas. Frontiers in Ecology and the Environment 4:238–243. Clyde, V. L., G. V. Kollias, M. E. Roelke, and M. R. Wells. 1990. Disseminated coccidioidomycosis in a western cougar (Felis concolor). Journal of Zoo and Wildlife Medicine 21:200–205. Cole, G. T., and C. Y. Hung. 2001. The parasitic cell wall of Coccidioides immitis. Medical Mycology 39:31–40. Collins, J. P. 2010. Amphibian decline and extinction: What we know and what we need to learn. Diseases of Aquatic Organisms 92:93–99. Comrie, A. C. 2005. Climate factors influencing coccidioidomycosis seasonality and outbreaks. Environmental Health Perspectives 113:688–692. Converse, K. A. 2007. Aspergillosis. Pp. 360–374 in N. J. Thomas, D. B. Hunter, and C. T. Atkinson (editors), Infectious diseases of wild birds. Blackwell Publishing, Ames, IA. Cornell, L. H., K. G. Osborn, J. E. Antrim, and J. G. Simpson. 1979. Coccidioidomycosis in a California sea otter. Journal of Wildlife Diseases 15:373–378. Costa, E. O., L. S. M. Diniz, C. F. Netto, C. Arruda, and M. L. A. Dagli. 1994. Epidemiological study of sporotrichosis and histoplasmosis in captive

228  kingdom fungi

Latin American wild mammals, Sao Paulo, Brazil. Mycopathologia 125:19–22. Cowan, D. F. 1993. Lobo’s disease in a bottlenose dolphin (Tursiops truncatus) from Matagorda Bay, Texas. Journal of Wildlife Diseases 29:488–489. Creekmore, L. H. 1999. Mycotoxins. Pp. 267–270 in M. Friend and J. C. Franson (editors), Field manual of wildlife diseases: General field procedures and diseases of birds. U.S. Geological Survey, Washington, DC. Cryan, P. 2011. White-nose syndrome threatens the survival of hibernating bats in North America. In U.S. Geological Survey, Fort Collins Science Center. . Cryan, P. M., C. U. Meteyer, J. G. Boyles, and D. S. Blehert. 2010. Wing pathology of whitenose syndrome in bats suggests life-threatening disruption of physiology. BMC Biology 8:135. Cryan, P. M., C. U. Meteyer, J. G. Boyles, and D. S. Blehert. 2013. White-nose syndrome in bats: Illuminating the darkness. BMC Biology. . Daszak, P., L. Berger, A. A. Cunningham, A. D. Hyatt, D. E. Green, and R. Speare. 1999. Emerging infectious diseases in amphibian declines. Emerging Infectious Diseases 5:735–748. Daszak, P., A. A. Cunningham, and A. D. Hyatt. 2003. Infectious disease and amphibian population declines. Diversity and Distributions 9:141–150. Daszak, P., A. Strieby, A. A. Cunningham, J. E. Longcore, C. C. Brown, and D. Porter. 2004. Experimental evidence that the bullfrog (Rana catesbeiana) is a potential carrier of chytridiomycosis, an emerging fungal disease of amphibians. Herpetological Journal 14:201–207. Davidson, E. W., M. Parris, J. P. Collins, J. E. Longcore, A. P. Pessier, and J. Brunner. 2003. Pathogenicity and transmission of chytridiomycosis in tiger salamanders (Ambystoma tigrinum). Copeia 2003:601–607. Davis, J. W. 1981. Coccidioidomycosis. Pp. 361–365 in J. W. Davis, L. H. Karstad, and D. O. Trainer (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Denning, D. W., and D. A. Stevens. 1990. Antifungal and surgical treatment of invasive aspergillosis: Review of 2121 published cases. Review of Infectious Diseases 12:1147–1201. DeVries, G. A., and J. J. Laarmann. 1973. A case of Lobo’s disease in the dolphin Sotalia guianensis. Aquatic Mammals 1:26–33. DiCaudo, D. J. 2006. Coccidioidomycosis: A review and update. Journal of the American Academy of Dermatology 55:929–942.

DiSalvo, A. F., W. J. Bigler, L. Ajello, J. E. Johnson, and J. Palmer. 1970. Bat and soil studies for sources of histoplasmosis in Florida. Public Health Reports 85:1063–1069. Dixon, D. M., J. C. Rhodes, and R. A. Fromtling. 1999. Taxonomy, classification, and morphology of the fungi. Pp. 1161–1166 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. American Society for Microbiology, Washington, DC. Durden, L. A., and G. G. Musser. 1994. The mammalian hosts of the sucking lice (Anoplura) of the world: A host–parasite list. Bulletin of the Society for Vector Ecology 19:130–168. Durkin, M. M., P. A. Connolly, and L. J. Wheat. 1997. Comparison of radioimmunoassay and enzymelinked immunoassay methods for detection of Histoplasma capsulatum var. capsulatum antigen. Journal of Clinical Microbiology 35:2252–2255. Dvorek, J., M. Otčenášek, and B. Rosicky. 1973. Adiaspiromycosis caused by Emmonsia crescens, Emmons and Jellison 1960. Acta Universitatis Palackianae Olomucensis Facultatits Medicae 70:1–120. Edds, G. T., K. P. C. Nair, and C. F. Simpson. 1973. Effect of aflatoxin B1 on resistance in poultry against cecal coccidiosis and Marek’s disease. American Journal of Veterinary Research 34:819–826. Egeberg, R. O., A. E. Elconin, and M. C. Egeberg. 1964. Effect of salinity and temperature on Coccidioides immitis and three antagonistic soil saprophytes. Journal of Bacteriology 88:473–476. Ehrlich, M., C. Driscoll, and W. B. Gross. 1985. Effect of dietary exposure to aflatoxin B1 on resistance of young chickens to organophosphate pesticide challenge. Avian Diseases 29:715–720. El-Khouly, A. B., F. A. Gadir, D. D. Cluer, and G. W. Manefield. 1992. Aspergillosis in camels affected with a specific respiratory and enteric syndrome. Australian Veterinary Journal 69:182–186. Elshafie, A., S. N. Al-Bahry, A. Y. AlKindi, T. Ba-Omar, and I. Mahmoud. 2007. Mycoflora and aflatoxins in soil, eggshells, and failed eggs of Chelonia mydas at Ras Al-Jinz, Oman. Chelonian Conservation and Biology 6:267–270. Emmons, C. W., and W. L. Jellison. 1960. Emmonsia crescens sp. n. and adiaspiromycosis (haplomycosis) in mammals. Annals of the New York Academy of Sciences 89:91–101. Errington, P. L. 1963. Muskrat populations. Iowa State University Press, Ames, IA.



Eskew, E. A., and B. D. Todd. 2013. Parallels in amphibian and bat declines from pathogenic fungi. Emerging Infectious Diseases 19:379–385. Fauquier, D. A., F. M. D. Gulland, J. G. Trupkiewicz, T. R. Spraker, and L. J. Lowenstine. 1996. Coccidioidomycosis in free-living California sea lions (Zalophus californianus) in central California. Journal of Wildlife Diseases 32:707–710. Fellers, G. M., D. E. Green, and J. E. Longcore. 2001. Oral chytridiomycosis in the mountain yellow-legged frog (Rana muscosa). Copeia 2001:945–953. Fischer, O. A. 2001. Adiaspores of Emmonsia parva var. crescens in lungs of small rodents in a rural area. Acta Veterinaria Brno 70:345–352. Fisher, F. S., M. W. Bultman, S. M. Johnson, D. Pappagianis, and E. Zaborsky. 2007. Coccidioides niches and habitat parameters in the southwestern United States: A matter of scale. Annals of the New York Academy of Sciences 1111:47–72. Fisher, M. C., and T. W. J. Garner. 2007. The relationship between the emergence of Batrachochytrium dendrobatidis, the international trade in amphibians and introduced amphibian species. Fungal Biology Reviews 21:2–9. Fisher, M. C., G. L. Koenig, T. J. White, G. San-Blas, R. Negroni, I. G. Alvarez, B. Wanke, and J. W. Taylor. 2001. Biogeographic range expansion into South America by Coccidioides immitis mirrors New World patterns of human migration. Proceedings of the National Academy of Sciences of the United States of America 98:4558–4562. Fisher, M. C., G. L. Koenig, T. J. White, and J. W. Taylor. 2002. Molecular and phenotypic description of Coccidioides posadasii sp. nov., previously recognized as the non-California population of Coccidioides immitis. Mycologia 94:73–84. Foley, J., D. Clifford, K. Castle, P. Cryan, and R. S. Ostfeld. 2011. Investigating and managing the rapid emergence of white-nose syndrome, a novel fatal, infectious disease of hibernating bats. Conservation Biology 25:223–231. Fraser, C. M., and A. Mays. 1986. The Merck veterinary manual. 6th ed. Merck & Co., Inc., Rahway, NJ. Frick, W. F., J. F. Pollock, A. C. Hicks, K. E. Langwig, D. S. Reynolds, G. G. Turner, C. M. Butchkoski, and T. H. Kunz. 2010. An emerging disease causes regional population collapse of a common North American bat species. Science 329:679–682. Friend, M. 1999a. Aspergillosis. Pp. 129–133 in M. Friend and J. C. Franson (editors), Field

kingdom fungi   229

manual of wildlife diseases: General field procedures and diseases of birds. U.S. Geological Survey, Washington, DC. Friend, M. 1999b. Miscellaneous fungal diseases. Pp. 137–138 in M. Friend and J. C. Franson (editors), Field manual of wildlife diseases: General field procedures and diseases of birds. U.S. Geological Survey, Washington, DC. Friend, M., and D. O. Trainer. 1969. Aspergillosis in captive herring gulls. Bulletin of the Wildlife Disease Association 5:271–275. Froschauer, A., and J. Coleman. 2012. North American bat death toll exceeds 5.5 million from white-nose syndrome. U. S. Fish and Wildlife Service, Office of Communications. . Galgiani, J. N. 1993. Conferences and reviews: Coccidioidomycosis. Western Journal of Medicine 159:153–171. Galgiani, J. N. 1999. Coccidioidomycosis: A regional disease of national importance; rethinking our approaches to its control. Annals of Internal Medicine 130:293–300. Gargas, A., M. T. Trest, M. Christensen, T. J. Volk, and D. S. Blehert. 2009. Geomyces destructans sp. nov. associated with bat white-nose syndrome. Mycotaxon 108:147–154. Garmyn, A., P. V. Rooij, F. Pasmans, T. Hellebuyck, W. V. D. Broeck, F. Haesebrouck, and A. Martel. 2012. Waterfowl: Potential environmental reservoirs of the chytrid fungus Batrachochytrium dendrobatidis. PLoS One 7:e35038. Garner, T. W. J., M. W. Perkins, P. Govindarajulu, D. Seglie, S. Walker, A. A. Cunningham, and M. C. Fisher. 2006. The emerging amphibian pathogen Batrachochytrium dendrobatidis globally infects introduced populations of the North American bullfrog, Rana catesbeiana. Biology Letters 2:455–459. Garner, T. W. J., S. Walker, J. Bosch, A. D. Hyatt, A. A. Cunningham, and M. C. Fisher. 2005. Chytrid fungus in Europe. Emerging Infectious Diseases 11:1639–1641. Gianni, C., G. Caretta, and C. Romano. 2003. Skin infection due to Geomyces pannorum var. pannorum. Mycoses 46:430–432. Gilbert, M., D. Bickford, L. Clark, A. Johnson, P. H. Joyner, L. O. Keats, K. Khammavong, L. N. Van, A. Newton, T. P. W. Seow, S. Roberton, S. Silithammavong, S. Singhalath, A. Yang, and T. A. Seimon. 2013. Amphibian pathogens in Southeast Asian frog trade. EcoHealth 9:386–398. Goad, D. L., and M. E. Pecquet-Goad. 1986. Osteoarticular sporotrichosis in a dog. Journal

230  kingdom fungi

of the American Veterinary Medical Association 189:1326–1328. Goodman, B. 2006. To stem widespread extinction, scientists airlift frogs in carry-on bags. The New York Times. . Goodman, L. L. 1965. Environmental studies on Histoplasma capsulatum, Ph.D. dissertation. University of Oklahoma, Norman, OK (cited in Ajello 1967). Gough, R. E. 1997. Collection and identification of avian viruses, bacteria, and fungi. Pp. 379–395 in D. H. Clayton and J. Moore (editors), Host– parasite evolution: General principles and avian models. Oxford University Press, Oxford, UK. Gower, D. G., T. Doherty-Bone, S. P. Loader, M. Wilkinson, M. T. Kouete, B. Tapley, F. Orton, O. Z. Daniel, F. Wynne, E. Flach, H. Müller, M. Menegon, I. Stephen, R. K. Browne, M. C. Fisher, A. A. Cunningham, and T. W. J. Garner. 2013. Batrachochytrium dendrobatidis infection and lethal chytridiomycosis in caecilian amphibians (Gymnophiona). EcoHealth. doi: 10.1007/s10393-10013-10831-10399. Graczyk, T. K., and M. R. Cranfield. 1995. Maternal transfer of anti-Aspergillus spp. immunoglobulins in African black-footed penguins (Spheniscus demersus). Journal of Wildlife Diseases 31:545–549. Graczyk, T. K., and M. R. Cranfield. 1996. A model for the prediction of relative titres of avian malaria and Aspergillus spp. IgG in jackass penguin (Spheniscus demersus) females based on IgG in egg-yolk. International Journal for Parasitology 26:749–754. Green, D. E., K. A. Converse, and A. K. Schrader. 2002. Epizootiology of sixty-four amphibian morbidity and mortality events in the USA, 1996–2001. Annals of the New York Academy of Sciences 969:323–339. Greer, D. L., and D. N. McMurray. 1981. Pathogenesis of experimental histoplasmosis in the bat, Artibeus lituratus. American Journal of Tropical Medicine and Hygiene 30:653–659. Gueho, E., M. C. Leclerc, G. S. d. Hoog, and B. Dupont. 1997. Molecular taxonomy and epidemiology of Blastomyces and Histoplasma species. Mycoses 40:69–81. Hallam, T. G., and G. F. McCracken. 2011. Management of the panzootic white-nose syndrome through culling of bats. Conservation Biology 25:189–194. Hanselmann, R., A. Rodriguez, M. Lampo, L. Fajardo-Ramos, A. A. Aguirre, A. M. Kilpatrick, J. P. Rodríguez, and P. Daszak. 2004. Presence of an emerging pathogen of amphibians in

introduced bullfrogs Rana catesbeiana in Venezuela. Biological Conservation 120:115–119. Harris, R. N., T. Y. James, A. Lauer, M. A. Simon, and A. Patel. 2006. Amphibian pathogen Batrachochytrium dendrobatidis is inhibited by cutaneous bacteria of amphibian species. EcoHealth 3:53–56. Haubold, E. M., C. R. Cooper, J. W. Wen, M. R. McGinnis, and D. F. Cowan. 2000. Comparative morphology of Lacazia loboi (syn. Loboa loboi) in dolphins and humans. Medical Mycology 38:9–14. Hawksworth, D. L. 1991. The fungal dimension of biodiversity: Magnitude, significance and conservation. Mycological Research 95:641–655 Hector, R. F., and R. Laniado-Laborin. 2005. Coccidioidomycosis: A fungal disease of the Americas. Public Library of Science Medicine 2:15–18. Herr, R. A., E. J. Tarcha, P. R. Taborda, J. W. Taylor, L. Ajello, and L. Mendoza. 2001. Phylogenetic analysis of Lacazia loboi places this previously uncharacterized pathogen within the dimorphic Onygenales. Journal of Clinical Microbiology 39:309–314. Hibbett, D. S., M. Binder, J. F. Bischoff, M. Blackwell, P. F. Cannon, O. E. Eriksson, S. Huhndorf, T. James, P. M. Kirk, R. Lücking, H. T. Lumbsch, F. Lutzoni, P. B. Matheny, K. J. McLaughlin, M. J. Powell, S. Redhead, C. L. Schoch, J. W. Spatafora, J. A. Stalpers, R. Vilgalys, M. C. Aime, A. Aptroot, R. Bauer, D. Begerow, G. L. Benny, L. A. Castlebury, P. W. Crous, Y.-C. Dai, W. Gams, D. M. Geiser, G. W. Griffith, C. Gueidan, D. L. Hawksworth, G. Hestmark, K. Hasaka, R. A. Humber, K. D. Hyde, J. E. Ironside, U. Kõljalg, C. P. Kurtzman, K.-H. Larsson, R. Lichtwardt, J. Longcore, J. Miądlikowska, A. Miller, J.-M. Moncalvo, S. Mozley-Standridge, F. Oberwinkler, E. Parmasto, V. Reeb, J. D. Rogers, C. Roux, L. Ryvarden, J. P. Sampaio, A. Schüßler, J. Sugiyama, R. G. Thorn, L. Tibell, W. A. Untereiner, C. Walker, Z. Wang, A. Weir, M. Weiss, M. M. White, K. Winka, Y.-J. Yao, and N. Zhang. 2007. A higher-level phylogenetic classification of the Fungi. Mycological Research 111:509–547. Hogan, L. H., S. M. Levitz, and B. S. Klein. 1996. Virulence factors of medically important fungi. Clinical Microbioly Reviews 9:469–488. Howard, D. H. 2003. An introduction to the taxonomy of zoopathogenic fungi. Pp. 1–16 in D. H. Howard (editor), Pathogenic fungi in humans and animals. Marcel Dekker, New York.



Hu, S., W. H. Chung, S. I. Hung, H. C. Ho, Z. W. Wang, C. H. Chen, S. C. Lu, T. T. Kuo, and H. S. Hong. 2003. Detection of Sporothrix schenckii in clinical samples by a nested PCR assay. Journal of Clinical Microbiology 41:1414–1418. Hubálek, Z., H. Burda, A. Scharff, G. Heth, E. Nevo, R. Šumbera, J. Peško, and J. Zima. 2005. Emmonsiosis of subterranean rodents (Bathyergidae, Spalacidae) in Africa and Israel. Medical Mycology 43:691–697. Hubálek, Z., J. Nesvadbová, and J. Halouska. 1998. Emmonsiosis of rodents in an agroecosystem. Medical Mycology 36:387–390. Hubálek, Z., J. Zejda, J. Nesvadbova, and B. Rynchovsky. 1991. Adiaspiromycosis: A widespread disease of rodents in southern Moravia. Folia Zoologica (Brno) 40:107–116. Hubálek, Z., J. Zejda, S. Svobodova, and J. Kucera. 1993. Seasonality of rodent adiaspiromycosis in a lowland forest. Journal of Medical and Veterinary Mycology 31:359–366. Hung, C.-Y., J. Xue, and G. T. Cole. 2007. Virulence mechanisms of Coccidioides. Annals of the New York Academy of Sciences 1111:225–235. Hussain, M., M. A. Gabal, T. Wilson, and R. C. Summerfelt. 1993. Effect of aflatoxincontaminated feed on morbidity and residues in walleye fish. Veterinary and Human Toxicology 35:396–398. Hyatt, A. D., D. G. Boyle, V. Olsen, D. B. Boyle, L. Berger, D. Obendorf, A. Dalton, K. Kriger, M. Hero, H. Hines, R. Phillott, R. Campbell, G. Marantelli, F. Gleason, and A. Colling. 2007. Diagnostic assays and sampling protocols for the detection of Batrachochytrium dendrobatidis. Diseases of Aquatic Organisms 73:175–192. Isbister, J., M. Elliott, and S. Nogrady. 1976. Histoplasmosis: An outbreak occurring among young men who visited one cave. Medical Journal of Australia 2:243–248. Ishizaki, H., M. Kawasaki, M. Aoki, T. Matsumoto, A. A. Padhye, M. Mendoza, and R. Negroni. 1998. Mitochondrial DNA analysis of Sporothrix schenckii in North and South America. Mycopathologia 142:115–118. Ishizaki, H., M. Kawasaki, M. Aoki, H. Vismer, and D. Muir. 2000. Mitochondrial DNA analysis of Sporothrix schenckii in south Africa and Australia. Medical Mycology 38:433–436. James, T. Y., F. Kauff, C. L. Schoch, P. B. Matheny, V. Hofstetter, C. J. Cox, G. Celio, C. Gueidan, E. Fraker, J. Miadlikowska, H. T. Lumbsch, A. Rauhut, V. Reeb, A. E. Arnold, A. Amtoft, J. E. Stajich, K. Hosaka, G. H. Sung, D. Johnson,

kingdom fungi   231

B. O’Rourke, M. Crockett, M. Binder, J. M. Curtis, J. C. Slot, Z. Wang, A. W. Wilson, A. Schüssler, J. E. Longcore, K. O’Donnell, S. Mozley-Standridge, D. Porter, P. M. Letcher, M. J. Powell, J. W. Taylor, M. M. White, G. W. Griffith, D. R. Davies, R. A. Humber, J. B. Morton, J. Sugiyama, A. Y. Rossman, J. D. Rogers, D. H. Pfister, D. Hewitt, K. Hansen, S. Hambleton, R. A. Shoemaker, J. Kohlmeyer, B. Volkmann-Kohlmeyer, R. A. Spotts, M. Serdani, P. W. Crous, K. W. Hughes, K. Matsuura, E. Langer, G. Langer, W. A. Untereiner, R. Lücking, B. Büdel, D. M. Geiser, A. Aptroot, P. Diederich, I. Schmitt, M. Schultz, R. Yahr, D. S. Hibbett, F. Lutzoni, D. J. McLaughlin, J. W. Spatafora, and R. Vilgalys. 2006. Reconstructing the early evolution of fungi using a six-gene phylogeny. Nature 443:818–822. Jellison, W. L. 1981. Adiaspiromycosis. Pp. 366–368 in J. W. Davis, L. H. Karstad, and D. O. Trainer (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Jellison, W. L., and R. D. Lord. 1964. Adiaspiromycosis in Argentine mammals. Mycologia 56:374–383. Jensen, H. E., J. B. Jorgensen, and H. Schonheyder. 1989. Pulmonary mycosis in farmed deer: Allergic zygomycosis and invasive aspergillosis. Journal of Medical and Veterinary Mycology 27:329–334. Jessup, D. A., N. Kock, and M. Berbach. 1989. Coccidioidomycosis in a desert bighorn sheep (Ovis canadensis nelsoni) from California. Journal of Zoo and Wildlife Medicine 20:471–473. Jewell, K., R. Cheshier, and G. D. Cage. 2008. Genetic diversity among clinical Coccidioides spp. isolates in Arizona. Medical Mycology 46:449–455. Johnson, M. L., L. Berger, L. Philips, and R. Speare. 2003. Fungicidal effects of chemical disinfectants, UV light, desiccation and heat on the amphibian chytrid Batrachochytrium dendrobatidis. Diseases of Aquatic Organisms 57:255–260. Johnson, M. L., and R. Speare. 2003. Survival of Batrachochytrium dendrobatidis in water: Quarantine and disease control implications. Emerging Infectious Diseases 9:922–925. Johnson, M. L., and R. Speare. 2005. Possible modes of dissemination of the amphibian chytrid Batrachochytrium dendrobatidis in the environment. Diseases of Aquatic Organisms 65:181–186. Johnson, P. T. J. 2006. Amphibian diversity: Decimation by disease. Proceedings of the National Academy of Sciences of the United States of America 103:3011–3012.

232  kingdom fungi

Johnson, S. M., K. M. Kerekes, J. M. Lunetta, and D. Pappagianis. 2007. Characteristics of the protective subcellular coccidioidal T27K vaccine. Annals of the New York Academy of Sciences 1111:275–289. Jones, M. E. B., D. Paddock, L. Bender, J. L. Allen, M. S. Schrenzel, and A. P. Pessiert. 2012. Treatment of chytridiomycosis with reduced-dose itraconazole. Diseases of Aquatic Organisms 99:243–249. Jones, T. C., and R. D. Hunt. 1983. Diseases caused by higher bacteria and fungi. Pp. 638–718 in Veterinary pathology. Lea and Febiger, Philadelphia, PA. Joneson, S., J. E. Stajich, S.-H. Shiu, and E. B. Rosenblum. 2011. Genomic transition to pathogenicity in chytrid fungi. PLoS Pathogens 7:e1002338. Kalka, M. B., A. R. Smith, and E. K. V. Kaldo. 2008. Bats limit arthropods and herbivory in a tropical forest. Science 320:71. Kane, J., and R. C. Summerbell. 1999. Trichophyton, Microsporum, Epidermophyton, and agents of superficial mycoses. Pp. 1275–1294 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Katz, M. E., S. C. J. Love, H. S. Gill, and B. F. Cheetham. 1996. Development of a method for identification using the polymerase chain reaction, of Aspergillus fumigatus isolated from ostriches. Australian Veterinary Journal 74:50–54. Kersulyte, D., J. P. Woods, E. J. Keath, W. E. Goldman, and D. E. Berg. 1992. Diversity among clinical isolates of Histoplasma capsulatum detected by polymerase chain reaction with arbitrary primers. Journal of Bacteriology 174:7075–7079. Kirkland, T. N., and J. Fierer. 1996. Coccidioidomycosis: A reemerging infectious disease. Emerging Infectious Diseases 2:192–199. Knudtson, W. U., C. E. Gates, and G. R. Ruth. 1980. Trichophyton mentagrophytes dermatophytosis in wild fox. Journal of Wildlife Diseases 16:465–468. Kolivras, K. N., and A. C. Comrie. 2003. Modeling valley fever incidence based on climate conditions in Pima County, Arizona. International Journal of Biometeorology 47:87–101. Kriger, K. M., and J.-M. Hero. 2006. Survivorship in wild frogs infected with chytridiomycosis. EcoHealth 3:171–177. Kriger, K. M., and J.-M. Hero. 2007. The chytrid fungus Batrachochytrium dendrobatidis is

non-randomly distributed across amphibian breeding habitats. Diversity and Distributions 13:781–788. Křivanec, K., and M. Otčenášek. 1977. Importance of free living mustelid carnivores in circulation of adiaspiromycosis. Mycopathologia 60:139–144. Kunkle, R. A., and J. L. Richard. 1998. Mycoses and mycotoxicoses. P. 311 in D. E. Swayne, J. R. Glisson, M. W. Jackwood, J. E. Pearson, and W. M. Reed (editors), A laboratory manual for the isolation and identification of avian pathogens. American Association of Avian Pathologists, University of Pennsylvania, Kennett Square, PA. Kwon-Chung, K. J., and J. E. Bennett. 1992. Medical mycology. Lea & Febiger, Philadelphia, PA. Lam, B., D. Woodhams, R. Harris, and V. Vredenburg. 2008. Probiotic bacteria protect endangered frogs from lethal skin disease. Science Daily, American Society for Microbiology (June 5). . Lampo, M., C. Barrio-Amorós, and B. Han. 2007. Batrachochytrium dendrobatidis infection in the recently rediscovered Atelopus mucubajiensis (Anura, Bufonidae), a critically endangered from the Venezuelan Andes. EcoHealth 3:299–307. Laniado-Laborin, R. 2007. Expanding understanding of epidemiology of coccidioidomycosis in the Western Hemisphere. Annals of the New York Academy of Sciences 1111:19–34. Lawson, B., S. MacDonald, T. Howard, S. Macgregor, and A. Cunningham. 2006. Exposure of garden birds to aflatoxins in Britain. Science of the Total Environment 361:124–131. Legendre, A. M. 1995. Antimycotic drug therapy. Pp. 327–331 in J. D. Bonagura and R. W. Kirk (editors), Kirk’s current veterinary therapy. XII: Small animal practice. W. B. Saunders, Philadelphia, PA. Leighton, F. A., and G. Wobeser. 1978. The prevalence of adiaspiromycosis in three sympatric species of ground squirrels. Journal of Wildlife Diseases 14:362–365. Levine, H. B., Y.-C. M. Kong, and C. E. Smith. 1965. Immunization of mice to Coccidioides immitis: Dose, regimen and spherulation stage of the killed spherule vaccine. Journal of Immunology 94:132–142. Li, R.-K., M. A. Ciblak, N. Nordoff, L. Pasarell, D. W. Warnock, and M. R. McGinnis. 2000. In vitro activities of voriconazole, itraconazole, and amphotericin B against Blastomyces dermatitidis, Coccidioides immitis, and Histoplasma capsulatum. Antimicrobial Agents and Chemotherapy 44:1734–1736. Lindner, D. L., A. Gargas, J. M. Lorch, M. T. Banik, J. Glaeser, T. H. Kunz, and D. S. Blehert. 2010.



DNA-based detection of the fungal pathogen Geomyces destructans in soils from bat hibernacula. Mycologia 103:241–246. . Lips, K. R., F. Brem, R. Brenes, J. D. Reeve, R. A. Alford, J. Voyles, C. Carey, L. Livo, A. P. Pessier, and J. P. Collins. 2006. Emerging infectious disease and the loss of biodiversity in a Neotropical amphibian community. Proceedings of the National Academy of Sciences of the United States of America 103:3165–3170. Lips, K. R., J. Diffendorfer, J. R. Mendelson, and M. W. Sears. 2008. Riding the wave: Reconciling the roles of disease and climate change in amphibian declines. Public Library of Science Biology 6:441–454 (e472). Lips, K. R., D. E. Green, and R. Papendick. 2003. Chytridiomycosis in wild frogs from southern Costa Rica. Journal of Herpetology 37:215–218. Locke, L. N., G. E. Bagley, D. N. Frickie, and L. T. Young. 1969. Lead poisoning and aspergillosis in an Andean condor. Journal of the American Veterinary Association 155:1052–1056. Longcore, J. E., A. P. Pessier, and D. K. Nichols. 1999. Batrachochytrium dendrobatidis gen. et sp. nov., a chytrid pathogenic to amphibians. Mycologia 91:219–227. Lopes-Bezerra, L. M., A. Schubach, and R. O. Costa. 2006. Sporothrix schenckii and sporotrichosis. Anais da Academia Brasileira de Ciências 78:293–308. Lorch, J. M., A. Gargas, C. U. Meteyer, B. M. Berlowski-Zier, D. E. Green, V. Shearn-Bochsler, N. J. Thomas, and D. S. Blehert. 2010. Rapid polymerase chain reaction diagnosis of white-nose syndrome in bats. Journal of Veterinary Diagnostic Investigation 22:224–230. Lorch, J. M., D. L. Lindner, A. Gargas, L. K. Muller, A. M. Minnis, and D. S. Blehert. 2013. A culturebased survey of fungi in soil from bat hibernacula in the eastern United States and its implications for detection of Geomyces destructans, the causal agent of bat white-nose syndrome. Mycologia 105:237–252. Lorch, J. M., C. U. Meteyer, M. J. Behr, J. G. Boyles, P. M. Cryan, A. C. Hicks, A. E. Ballmann, J. T. H. Coleman, D. N. Redell, D. M. Reeder, and D. S. Blehert. 2011. Experimental infection of bats with Geomyces destructans causes white-nose syndrome. Nature. doi: 10.1038/nature10590. Lutzoni, F., F. Kauff, C. J. Cox, D. McLaughlin, G. Celio, B. Dentinger, M. Padamsee, D. Hibbett, T. Y. James, E. Baloch, M. Grube, V. Reeb, V. Hofstetter, C. Schoch, A. E. Arnold, J. Miadlikowska, J. Spatafora, D. Johnson,

kingdom fungi   233

S. Hambleton, M. Crockett, R. Shoemaker, G.-H. Sung, R. Lücking, T. Lumbsch, K. O’Donnell, M. Binder, P. Diederich, D. Ertz, C. Gueidan, K. Hansen, R. C. Harris, K. Hosaka, Y.-W. Lim, B. Matheny, H. Nishida, D. Pfister, J. Rogers, A. Rossman, I. Schmitt, H. Sipman, J. Stone, J. Sugiyama, R. Yahr, and R. Vilgalys. 2004. Assembling the fungal tree of life: Progress, classification, and evolution of subcellular traits. American Journal of Botany 91:1446–1480. Mackinnon, J. E., I. A. Conti-Diaz, E. Gezuele, E. Civila, and S. daLuz. 1969. Isolation of Sporothrix schenckii from nature and considerations on its pathogenicity and ecology. Sabouraudia 7:38–45. Maddy, K. T. 1959. Coccidioidomycosis in animals. Veterinary Medicine 54:233–242. Magee, D. M., R. L. Friedberg, M. D. Woitaske, S. A. Johnston, and R. A. Cox. 2005. Role of B cells in vaccine-induced immunity against coccidioidomycosis. Infection and Immunity 73:7011–7013. Marantelli, G., L. Berger, R. Speare, and L. Keegan. 2004. Distribution of the amphibian chytrid Batrachochytrium dendrobatidis and keratin during tadpole development. Pacific Conservation Biology 10:173–179. Martinez-Quesada, J., A. Nieto-Cadenazzi, and J. M. Torres-Rodriguez. 1993. Humoral immunoresponse of pigeons to Aspergillus fumigatus antigens. Mycopathologia 124:131–137. Martínková, N., P. Bačkor, T. Bartonička, P. Blažková, J. Červený, L. Falteisek, J. Gaisler, V. Hanzal, D. Horáček, Z. Hubálek, H. Jahelková, M. Kolařik, L. u. Korytár, A. Kubátová, B. Lehotská, R. Lehotský, R. K. Lučan, O. Májek, J. Matĕjů, Z. Řehák, J. Šafář, P. Tájek, E. Tkadlec, M. Uhrin, J. Wagner, D. Weinfurtová, J. Zima, J. Zukal, and I. Horáček. 2010. Increasing incidence of Geomyces destructans fungus in bats from the Czech Republic and Slovakia. PLoS One 5:e13853. doi: 13810.13137/journal.pane.0013853. McDiarmid, A. 1955. Aspergillosis in free-living birds. Journal of Comparative Pathology 65:246–249. McDonald, K. R., D. Méndez, R. Müller, A. B. Freeman, and R. Speare. 2005. Decline in the prevalence of chytridiomycosis in frog populations in North Queensland, Australia. Pacific Conservation Biology 11:114–120. McGinnis, M. R., L. Sigler, and M. G. Rinaldi. 1999. Some medically important fungi and their common synonyms and names of uncertain application. Clinical Infectious Diseases 29:728–730.

234  kingdom fungi

McKeever, S., W. Kaplan, and L. Ajello. 1958. Ringworm fungi of large wild mammals in southwestern Georgia and northwestern Florida. American Journal of Veterinary Research 19:973–975. McMahon, T. A., L. A. Brannelly, M. W. H. Chatfield, P. T. J. Johnson, M. B. Joseph, V. J. M. C. L. RichardsZawacki, M. D. Venesky, and J. R. Rohr. 2012. Chytrid fungus Batrachochytrium dendrobatidis has nonamphibian hosts and releases chemicals that cause pathology in the absence of infection. Proceedings of the National Academcy of Sciences. . Mendoza, L., A. F. F. Belone, R. Vilela, M. Rehtanz, G. D. Bossart, J. S. Reif, P. A. Fair, W. N. Durden, J. S. Leger, L. R. Travassos, and P. S. Rosa. 2008. Use of sera from humans and dolphins with lacaziosis and sera from experimentally infected mice for Western blot analyses of Lacazia loboi antigens. Clinical and Vaccine Immunology 15:164–167. Menges, R. W., and L. K. Georg. 1955. Animal ringworm study. Veterinary Medicine 50:293–297. Merz, W. G., and G. D. Roberts. 1999. Algorithms for detection and identification of fungi. Pp. 1167–1183 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Mesa-Arango, A. C., M. delRocíoReyes-Montes, A. Pérez-Mejía, H. Navarro-Barranco, V. Souza, G. Zúñiga, and C. Toriello. 2002. Phenotyping and genotyping of Sporothrix schenckii isolates according to geographic origin and clinical form of sporotrichosis. Journal of Clinical Microbiology 40:3004–3011. Meteyer, C. U., D. Barber, and J. N. Mandl. 2012. Pathology in euthermic bats with white nose syndrome suggests a natural manifestation of immune reconstitution inflammatory syndrome. Virulence 3:1–6. Meteyer, C. U., E. L. Buckles, D. S. Blehert, A. C. Hicks, D. E. Green, V. Shearn-Bochsler, N. J. Thomas, A. Gargas, and M. J. Behr. 2009. Histopathologic criteria to confirm white-nose syndrome in bats. Journal of Veterinary Diagnostic Investigation 21:411–414. Meteyer, C. U., M. Valent, J. Kasmer, E. L. Buckles, J. M. Lorch, D. S. Blehert, A. Lollar, D. Berndt, E. Wheeler, C. L. White, and A. E. Ballmann. 2011. Recovery of little brown bats (Myotis lucifugus) from natural infection with Geomyces destructans, white-nose syndrome. Journal of Wildlife Diseases 47:618–626.

Migaki, G., R. L. Font, W. Kaplan, and E. D. Asper. 1978. Sporotrichosis in a Pacific white-sided dolphin (Lagenorhynchus obliquidens). American Journal of Veterinary Research 39:1916–1919. Migaki, G., M. G. Valerio, B. Irvine, and F. M. Garner. 1971. Lobo’s disease in an Atlantic bottle-nosed dolphin. Journal of the American Veterinary Medical Association 159:578–582. Miranda, M. F., and A. J. Silva. 2005. Vinyl adhesive tape also effective for direct microscopy diagnosis of chromomycosis, lobomycosis, and paracoccidioidomycosis. Diagnostic Microbiology and Infectious Diseases 52:39–43. Mitchell, K. M., T. S. Churcher, T. W. J. Garner, and M. C. Fisher. 2008. Persistence of the emerging pathogen Batrachochytrium dendrobatidis outside the amphibian host greatly increases the probability of host extinction. Proceedings of the Royal Society B: Biological Sciences 275:329–334. Morehouse, E. A., T. Y. James, A. R. D. Ganley, R. Vilgalys, L. Berger, P. Murphy, and J. E. Longcore. 2003. Multilocus sequence typing suggests the chytrid pathogen of amphibians is a recently emerged clone. Molecular Ecology 12:395–403. Morrow, W. 2006. Holocene coccidioidomycosis: Valley fever in early holocene bison (Bison antiquus). Mycologia 98:669–677. Moss, M. O. 2002. Mycotoxin review. 1: Aspergillus and Penicillium. Mycologist 16:116–119. Murray, D. A., L. F. Skerratt, S. Garland, D. Kriticos, and H. McCallum. 2013. Whether the weather drives patterns of endemic amphibian chytridiomycosis: A pathogen proliferation approach. PLoS One 8:e61061. Neff, J. A. 1955. Outbreak of aspergillosis in mallards. Journal of Wildlife Management 19:415–416. O’Hara, T. M. 1996. Mycotoxins. Pp. 24–30 in A. Fairbrother, L. N. Locke, and G. L. Hoff (editors), Noninfectious diseases of wildlife. Iowa State University Press, Ames, IA. Olsen, V., A. D. Hyatt, D. G. Boyle, and D. Mendez. 2004. Co-localisation of Batrachochytrium dendrobatidis and keratin for enhanced diagnosis of chytridiomycosis in frogs. Diseases of Aquatic Organisms 61:85–88. Olson, D. H., D. M. Aanensen, K. L. Ronnenberg, C. I. Powell, S. F. Walker, J. Bielby, T. W. J. Garner, G. Weaver, T. B. M. Group, and M. C. Fisher. 2013. Mapping the global emergence of Batrachochytrium dendrobatidis, the amphibian chytrid fungus. PLoS One 8:e56802. O’Meara, D. C., and H. L. Chute. 1959. Aspergillosis experimentally produced in hatching chicks. Avian Diseases 3:404–406.



O’Meara, D. C., and J. F. Witter. 1971. Aspergillosis. Pp. 153–162 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Opromolla, D. V. A., A. F. F. Belone, P. R. O. Taborda, and V. B. A. Taborda. 2000. Clinic-pathological correlation in 40 cases of lobomycosis. Clinical, Laboratory and Therapeutic Investigation 75:425–434. Opromolla, D. V. A., and M. E. S. Nogueira. 2000. Inoculation of Lacazia loboi into the subcutaneous tissue of the hamster cheek pouch. Revista do Instituto de Medicina Tropical de São Paulo 42:119–123. Otčenášek, M., J. Dvořák, and B. Rosický. 1967. Some epidemiological aspects of histoplasmosis in the light of Pavlovsky’s theory. Folia Parasitologica (Praha) 14:193–206. Ouellet, M., I. Mikaelian, B. D. Pauli, J. Rodrigue, and D. M. Green. 2005. Historical evidence of widespread chytrid infection in North American amphibian populations. Conservation Biology 19:1431–1440. Palmgren, M. s., and A. W. Hayes. 1987. Aflatoxins in food. Pp. 65–95 in P. Krogh (editor), Mycotoxins in food. Academic Press, New York. Pappagianis, D. 2001. Seeking a vaccine against Coccidioides immitis and serologic studies: Expectations and realities. Fungal Genetics and Biology 32:1–9. Parish, J. M., and J. E. Blair. 2008. Coccidioidomycosis. Mayo Clinic Proceedings 83:343–348. Park, B. J., K. Sigel, V. Vaz, K. Komatsu, C. McRill, M. Phelan, T. Colman, A. C. Comrie, D. W. Warnock, J. N. Galgiani, and R. A. Hajjeh. 2005. An epidemic of coccidioidomycosis in Arizona associated with climatic changes, 1998–2001. Journal of Infectious Diseases 191:1981–1987. Peden, W. M., J. L. Richard, D. W. Trampel, and R. E. Brannian. 1985. Mycotic pneumonia and meningoencephalitis due to Aspergillus terreus in a neonatal snow leopard (Panthera uncia). Journal of Wildlife Diseases 21:301–305. Pennycott, T. W. 1999. Causes of mortality in mute swans (Cygnus olor) in Scotland 1995–1996. Wildfowl 50:11–20. Pessier, A. P., D. K. Nichols, J. E. Longcore, and M. S. Fuller. 1999. Cutaneous chytridiomycosis in poison dart frogs (Dendrobates spp.) and White’s tree frogs (Litoria caerulea). Journal of Veterinary Diagnostic Investigation 11:194–199. Pfaller, M. A., and D. J. Diekema. 2005. Unusual fungal and pseudofungal infections of humans. Journal of Clinical Microbiology 43:1495–1504.

kingdom fungi   235

Pickett, J. P., C. P. Moore, B. A. Beehler, A. GendronFitzpatrick, and R. R. Dubielzig. 1985. Bilateral chorioretinitis secondary to disseminated aspergillosis in an alpaca. Journal of the American Veterinary Medical Association 187:1241–1243. Pier, A. C. 1992. Major biological consequences of aflatoxicosis in animal production. Journal of Animal Science 70:3964–3967. Pier, A. C., and J. L. Richard. 1992. Mycoses and mycotoxicoses of animals caused by aspergilli. Biotechnology 23:233–248. Piotrowski, J. S., S. L. Annis, and J. E. Longcore. 2004. Physiology of Batrachochytrium dendrobatidis, a chytrid pathogen of amphibians. Mycologia 96:9–15. Pounds, J. A., M. R. Bustamante, L. A. Coloma, J. A. Consuegra, M. P. L. Fogden, P. N. Foster, E. LaMarca, K. L. Masters, A. Merino-Viteri, R. Puschendorf, S. R. Ron, G. A. Sánchez-Azofeifa, C. J. Still, and B. E. Young. 2006. Widespread amphibian extinctions from epidemic disease driven by global warming. Nature 439:161–167. Powell, M. J. 1993. Looking at mycology with a Janus face: A glimpse at Chytridiomycetes in the environment. Mycologia 85:1–20. Puechmaille, S., J. Wibbelt, Gudrun , V. Korn, H. Fuller, F. Forget, K. Mühldorfer, A. Kurth, W. Bogdanowicz, C. Borel, T. Bosch, T. Cherezy, M. Drebet, T. Görföl, A.-J. Haarsma, F. Herhaus, G. Hallart, M. Hammer, C. Jungmann, Y. Le Bris, L. Lutsar, M. Masing, B. Mulkens, K. Passior, M. Starrach, A. Wojtaszewski, U. Zöphel, and E. C. Teeling. 2011. Pan-European distribution of white-nose syndrome fungus (Geomyces destructans) not associated with mass mortality. PLoS One 6:e19167. doi: 19110.11371/ journal.pone.0019167. Puechmaille, S. J., P. Verdeyroux, H. Fuller, M. A. Gouilh, M. Bekaert, and E. C. Teeling. 2010. White-nose syndrome fungus (Geomyces destructans) in bat, France. Emerging Infectious Diseases 16:290–293 (serial on the Internet). Quandt, S. K. F., and J. W. Nesbit. 1992. Histoplasmosis in a 2-toed sloth (Choloepus didactylus). Journal of Zoo and Wildlife Medicine 23:369–373. Quist, C. F., T. Cornish, and R. D. Wyatt. 2007. Mycotoxicosis. Pp. 417–430 in N. J. Thomas, D. B. Hunter, and C. T. Atkinson (editors), Infectious diseases of wild birds. Blackwell Publishing Professional, Ames, IA. Rachowicz, L. J., and C. J. Briggs. 2007. Quantifying the disease transmission function: Effects of density on Batrachochytrium dendrobatidis transmission in the mountain yellow-legged

236  kingdom fungi

frog Rana mucosa. Journal of Animal Ecology 76:711–721. Rachowicz, L. J., J.-M. Hero, R. A. Alford, J. W. Taylor, J. A. T. Morgan, V. T. Vredenburg, J. P. Collins, and C. J. Briggs. 2005. The novel and endemic pathogen hypotheses: Competing explanations for the origin of emerging infectious diseases of wildlife. Conservation Biology 19:1441–1448. Rachowicz, L. J., and V. T. Vredenburg. 2004. Transmission of Batrachochytrium dendrobatidis within and between amphibian life stages. Diseases of Aquatic Organisms 61:75–83. Rao, J. R., N. N. Sharma, P. K. R. Iyer, and A. K. Sharma. 1990. Interaction between Eimeria uzura infection and aflatoxicosis in Japanese quail (Coturnix coturnix japonica). Veterinary Parasitology 35:259–267. Raper, K. B., and D. I. Fennell. 1965. The genus Aspergillus. Williams and Wilkins, Baltimore, MD. Redig, P. T., R. R. Fuller, and D. L. Evans. 1980. Prevalence of Aspergillus fumigatus in free-living goshawks (Accipiter gentilis atricapillus). Journal of Wildlife Diseases 16:169–174. Reeder, N. M. M., A. P. Pessier, and V. T. Vredenburg. 2012. A reservoir species for the emerging amphibian pathogen Batrachochytrium dendrobatidis thrives in a landscape decimated by disease. PLoS One 7:e33567. Reichard, J. D., and T. H. Kunz. 2009. White-nose syndrome inflicts lasting injuries to the wings of little brown myotis (Myotis lucifugus). Acta Chiropterologica 11:457–464. Reidarson, T. H., L. A. Griner, D. Pappagianis, and J. McBain. 1998a. Coccidioidomycosis in a bottlenose dolphin. Journal of Wildlife Diseases 34:629–631. Reidarson, T. H., J. H. Harrell, M. G. Rinaldi, and J. McBain. 1998b. Bronchoscopic and serologic diagnosis of Aspergillus fumicatus pulmonary infection in a bottlenose dolphin (Tursiops truncatus). Journal of Zoo and Wildlife Medicine 29:451–455. Reif, J. S., M. S. Mazzoil, S. D. McCulloch, R. A. Varela, J. D. Goldstein, P. A. Fair, and G. D. Bossart. 2006. Lobomycosis in Atlanta bottlenose dolphins from the Indian River Lagoon, Florida. Journal of the American Veterinary Medical Association 228:104–108. Retallick, R. W., and V. Miera. 2007. Strain differences in the amphibian chytrid Batrachochytrium dendrobatidis and non-permanent, sub-lethal effects of infection. Diseases of Aquatic Organisms 75:201–207. Retallick, R. W. R., H. McCallum, and R. Speare. 2004. Endemic infection of the amphibian

chytrid fungus in a frog community post-decline. PLoS Biology 2:e351. Retallick, R. W. R., V. Miera, K. L. Richard, K. J. Field, and J. P. Collins. 2006. A non-lethal technique for detecting the chytrid fungus Batrachochytrium dendrobatidis on tadpoles. Diseases of Aquatic Organisms 72:77–85. Riley, K., O. F. Berry, and J. D. Roberts. 2013. Do global models predicting environmental suitability for the amphibian fungus, Batrachochytrium dendrobatidis, have local value to conseravation managers? Journal of Applied Ecology. doi: 10.1111/1365-2664.12091. Robinson, R. M., A. C. Ray, J. C. Reagor, and L. A. Holland. 1982. Waterfowl mortality caused by aflatoxicosis in Texas. Journal of Wildlife Diseases 18:311–313. Rollins-Smith, L. A., C. Carey, J. Longcore, J. K. Doersam, A. Boutte, J. E. Bruzgal, and J. M. Conlon. 2002. Activity of antimicrobial skin peptides from ranid frogs against Batrachochytrium dendrobatidis, the chytrid fungus associated with global amphibian declines. Developmental and Comparative Immunology 26:471–479. Rollins-Smith, L. A., J. P. Ramsey, J. D. Pask, L. K. Reinert, and D. C. Woodhams. 2011. Amphibian immune defenses against chytridiomycosis: Impacts of changing environments. Integrative and Comparative Biology 51:552–562. Ron, S. 2005. Predicting the distribution of the amphibian pathogen Batrachochytrium dendrobatidis in the New World. Biotropica 37:209–221. Ron, S. R., W. E. Duellman, L. A. Coloma, and M. R. Bustamante. 2003. Population decline of the Jambato toad Atelopus ignescens (Anura: Bufonidae) in the Andes of Ecuador. Journal of Herpetology 37:116–126. Ron, S. R., and A. Merino. 2000. Amphibian declines in Ecuador: Overview and first report of chytridiomycosis from South America. Froglog 42:2–3. Rosen, M. N. 1964. Aspergillosis in wild and domestic fowl. Avian Diseases 8:1–6. Rowley, J. J., L. F. Skerratt, R. A. Alford, and R. Campbell. 2007. Retreat sites of rain forest stream frogs are not a reservoir for Batrachochytrium dendrobatidis in northern Queensland, Australia. Diseases of Aquatic Organisms 74:7–12. Ruff, M. D., W. E. Huff, and G. C. Wilkins. 1990. Characterization of the toxicity of the mycotoxins aflatoxin, ochratoxin, and T-2 toxin in game birds. I: Chukar partridge. Avian Diseases 34:717–720.



Ruff, M. D., W. E. Huff, and G. C. Wilkins. 1992. Characterization of the toxicity of the mycotoxins aflatoxin, ochratoxin, and T-2 toxin in game birds. III: Bobwhite and Japanese quail. Avian Diseases 36:34–39. Sampaio, M. M., and L. Braga-Dias. 1977. The armadillo Euphractus sexcinctus as a suitable animal for experimental studies of Jorge Lobo’s disease. Revista do Instituto de Medicina Tropical de São Paulo 19:215–220. San-Blas, G., and G. Niño-Vega. 2004. Morphogenesis in other agents of systemic mycoses. Pp. 167–220 in G. San-Blas and R. A. Calderone (editors), Pathogenic fungi: Structural biology and taxonomy. Caister Academic Press, Norfolk, UK. Sanger, V. L. 1981. Histoplasmosis. Pp. 356–360 in J. W. Davis, L. H. Karstad, and D. O. Trainer (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Saubolle, M. A. 2007. Laboratory aspects in the diagnosis of coccidioidomycosis. Annals of the New York Academy of Sciences 1111:301–314. Saubolle, M. A., P. P. McKellar, and D. Sussland. 2007. Epidemiologic, clinical, and diagnostic aspects of coccidioidomycosis. Journal of Clinical Microbiology 45:26–30. Schell, W. A., I. F. Salkin, L. Pasarell, and M. R. McGinnis. 1999. Bipolaris, Exophiala, Scedosporium, Sporothrix, and other dematiaceous fungi. Pp. 1295–1317 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Schloegel, L. M., J.-M. Hero, L. Berger, R. Speare, K. McDonald, and P. Daszak. 2006. The decline of the sharp-snouted day frog (Taudactylus acutirostris): The first documented case of extinction by infection in a free-ranging wildlife species? EcoHealth 3:35–40. Scott, D. W., W. H. Miller, and C. E. Griffin. 1995. Muller and Kirk’s Small animal dermatology. W. B. Saunders, Philadelphia, PA. Severo, L. C., J. C. Bohrer, G. R. Geyer, and L. Ferreiro. 1989. Invasive aspergillosis in an alpaca (Lama pacos). Journal of Medical and Veterinary Mycology 27:193–195. Shacklette, M. H., H. F. Hasenclever, and E. A. Miranda. 1967. The natural occurrence of Histoplasma capsulatum in a cave. 2: Ecologic aspects. American Journal of Epidemiology 86:246–252. Sheafor, B., E. W. Davidson, L. Parr, and L. Rollins-Smith. 2008. Antimicrobial peptide defenses in the salamander, Ambystoma tigrinum,

kingdom fungi   237

against emerging amphibian pathogens. Journal of Wildlife Diseases 44:226–236. Shubitz, L. F. 2007. Comparative aspects of coccidioidomycosis in animals and humans. Annals of the New York Academy of Sciences 1111:395–403. Sigler, L. 2003. Ascomycetes: The Onygenaceae and other fungi from the Order Onygenales. Pp. 195–236 in D. H. Howard (editor), Pathogenic fungi in humans and animals. Marcel Dekker, New York. Šimonovičov, A., D. Pangallo, K. Chovanová, and B. Lehotská. 2011. Geomyces destructans associated with bat disease WNS detected in Slovakia. Biologia 66:562–564. Simõse-Lopes, P. C., G. S. Paula, M. C. Both, F. M. Xavier, and A. C. Scaramello. 1993. First case of lobomycosis in a bottlenose dolphin from southern Brazil. Marine Mammal Science 9:329–331. Smith, B. J., K. F. Higgins, and W. L. Tucker. 1990. Precipitation, waterfowl densities and mycotoxins: Their potential effect on avian cholera epizootics in the Nebraska Rainwater Basin area. Transactions of the North American Wildlife and Natural Resources Conference 55:269–282. Smith, C. E., and D. Pappagianis. 1961. Human coccidioidomycosis. Bacteriological Reviews 25:310–320. Souza, M. J., and L. A. Degernes. 2005. Mortality due to aspergillosis in wild swans in northeast Washington State, 2000–2002. Journal of Avian Medicine and Surgery 19:98–106. Sparrow, F. K. 1960. The aquatic phycomycetes. University of Michigan Press, Ann Arbor, MI. Spatafora, J. W., G.-H. Sung, D. Johnson, C. Hesse, B. O’Rourke, M. Serdani, R. Spotts, F. Lutzoni, V. Hofstetter, J. Miadlikowska, V. Reeb, C. Gueidan, E. Fraker, T. Lumbsch, R. Lücking, I. Schmitt, K. Hosaka, A. Aptroot, C. Roux, A. N. Miller, D. M. Geiser, J. Hafellner, G. Hestmark, A. E. Arnold, B. Büdel, A. Rauhut, D. Hewitt, W. A. Untereiner, M. S. Cole, C. Scheidegger, M. Schultz, H. Sipman, and C. L. Schoch. 2006. A five-gene phylogeny of Pezizomycotina. Mycologia 98:1018–1028. Steenbergen, J. N., J. D. Nosanchuk, S. D. Malliaris, and A. Casadevall. 2004. Interaction of Blastomyces dermatitidis, Sporothrix schenckii, and Histoplasma capsulatum with Acanthamoeba castellanii. Infection and Immunity 72:3478–3488. Stone, R. 2013. A rescue mission for amphibians at the brink of extinction. Science 339:1371.

238  kingdom fungi

Stone, W. B., and J. C. Okoniewski. 2001. Necropsy findings and environmental contaminants in common loons from New York. Journal of Wildlife Diseases 37:178–184. Straub, M., R. J. Trautman, and J. W. Greene. 1961. Coccidioidomycosis in 3 coyotes. American Journal of Veterinary Research 22:811–813. Stuart, S. N., J. S. Chanson, N. A. Cox, B. E. Young, A. S. L. Rodrigues, D. L. Fischman, and R. W. Waller. 2004. Status and trends of amphibian declines and extinctions worldwide. Science 306:1783–1786. Sweeney, J. C., G. Migaki, P. M. Vainik, and R. H. Conklin. 1976. Systemic mycoses in marine mammals. Journal of the American Veterinary Medical Association 169:946–948. Symmers, W. S. 1983. A possible case of Lobo’s disease acquired in Europe from a bottle-nosed dolphin (Tursiops truncatus). Bulletin de la Société de Pathologie Exotique et de ses Filiales 76:777–784. Szymanski, J. A., M. C. Runge, M. J. Parkin, and M. Armstrong. 2009. White-nose syndrome management: Report on structured decision making initiative. WNS SDM Report, October 2009. U. S. Fish and Wildlife Service and State Natural Resource Agencies. Timm, K. I., R. J. Sonn, and B. D. Hultgren. 1988. Coccidioidomycosis in a Sonoran gopher snake, Pituophis melanoleucus affinis. Journal of Medical and Veterinary Mycology 26:101–104. Tintelnot, K., G. S. DeHoog, E. Antweiler, H. Losert, M. Seibold, M. A. Brandt, A. H. G. GerritsVanDenEnde, and M. C. Fisher. 2007. Taxonomic and diagnostic markers for identification of Coccidioides immitis and Coccidioides posadasii. Medical Mycology 45:385–393. Turner, G. G., and D. M. Reeder. 2009. Update of white-nose syndrome in bats. Bat Research News 50:47–53. Turner, G. G., D. M. Reeder, and J. T. H. Coleman. 2011. A five-year assessment of mortality and geographic spread of white-nose syndrome in North American bats and a look to the future. Bat Research News 52:13–27. Umeyama, T., A. Sano, K. Kamei, M. Niimi, K. Nishimura, and Y. Uehara. 2006. Novel approach to designing primers for identification and distinction of the human pathogenic fungi Coccidioides immitis and Coccidioides posadasii by PCR amplification. Journal of Clinical Microbiology 44:1859–1862. Velasco-Castrejon, O., and A. Gonzalez-Ochoa. 1977. Primary pulmonary epidemic of

histoplasmosis in an abandoned mine. Mykosen 20:393–399. Vilela, R., L. Mendoza, P. S. Rosa, A. F. F. Belone, S. Madeira, D. V. A. Opromolla, and M. A. deResende. 2005. Molecular model for studying the uncultivated fungal pathogen Lacazia loboi. Journal of Clinical Microbiology 43:3657–3661. Viriyakosol, S., J. Fierer, G. D. Brown, and T. N. Kirkland. 2005. Innate immunity to the pathogenic fungus Coccidioides posadasii is dependent on toll-like receptor 2 and dectin-1. Infection and Immunity 73:1553–1560. Voyles, J., S. Young, L. Berger, C. Campbell, V. W. F., A. Dinudom, D. Cook, R. Webb, R. A. Alford, L. F. Skerratt, and R. Speare. 2009. Pathogenesis of chytridiomycosis, a cause of catastrophic amphibian declines. Science 236:585–592. Warnecke, L., J. M. Turner, T. K. Bollinger, J. M. Lorch, V. Misra, P. M. Cryan, G. Wibbelt, D. S. Blehert, and C. K. R. Willis. 2012. Inoculation of bats with European Geomyces destructans supports the novel pathogen hypothesis for the origin of white-nose syndrome. Proceedings of the National Academy of Sciences of the United States of America 109:6999–7003. Weldon, C., L. H. duPreez, A. D. Hyatt, R. Muller, and R. Speare. 2004. Origin of the amphibian chytrid fungus. Emerging Infectious Diseases 10:2100–2105. Werner, A. H., and B. E. Werner. 1994. Sporotrichosis in man and animals. International Journal of Dermatology 33:692–700. Wibbelt, G., A. Kurth, D. Hellmann, M. Weishaar, A. Borlow, M. Veith, J. Prüger, T. Görföl, L. Grosche, G. Bontadina, U. Zöphel, H.-P. Seidl, P. M. Cryan, and D. S. Blehert. 2010a.



White-nose syndrome fungus (Geomyces destructans) in bats, Europe. Emerging Infectious Diseases 16:1237–1242. Wibbelt, G., M. S. Moore, T. Schountz, and C. C. Voigt. 2010b. Meeting report: Emerging diseases in Chiroptera: Why bats? Biology Letters 6:438–440. Wilder, A., P. , W. F. Frick, K. E. Langwig, and T. H. Kunz. 2011. Risk factors associated with mortality from white-nose syndrome among hibernating bat colonies. Biology Letters 7:950–953. Willis, C. K. R., A. K. Menzies, J. G. Boyles, and M. S. Wojciechowski. 2011. Evaporative water loss is a plausible explanation for mortality of bats from white-nose syndrome. Integrative and Comparative Biology 51:364–373. Windingstad, R. M. 1990. Contaminated hay used as goose nest structures cause fungal infection in refuge staff. U.S. Fish and Wildlife Service Research Information Bulletin 90-27. Woodhams, D. C., R. A. Alford, and G. Marantelli. 2003. Emerging disease of amphibians cured by elevated body temperature. Diseases of Aquatic Organisms 55:65–67. Woodhams, D. C., K. Ardipradja, R. A. Alford, G. Marantelli, L. K. Reinert, and L. A. RollinsSmith. 2007. Resistance to chytridiomycosis varies among amphibian species and is correlated with skin peptide defenses. Animal Conservation 10:409–417. Zinkl, J. G., J. M. Hyland, and J. J. Hurt. 1977. Aspergillosis in common crows in Nebraska. Journal of Wildlife Diseases 13:191–193. Ziólkowska, G., and S. Tokarzewski. 2007. Occurrence of moulds in reproductive goose flocks in southern-eastern Poland. Bulletin of the Veterinary Institute in Pulawy 51:553–561.

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EIGHT

Introduction to Non-eukaryotic Agents

CONTENTS Bacteria241 Bacterial Identification 242

Reservoir Types Used by Bacteria and Viruses Vertebrate-dependent Reservoirs Latent Infections (Apparently Healthy Carriers) Clinically Active Infections Invertebrate–Vertebrate Reservoirs Arthropod–Vertebrate Reservoir Helminth–Vertebrate Reservoir Soil and Water as Reservoirs

Viruses243 Viral Identification 243 Bacterial and Viral Transmission Patterns 243 Physical Contact 244 Ingestion244 Aerosol Transmission 245 Arthropod Transmission 245 Helminth-mediated Transmission 245

Literature Cited

247 248 249 249 250 251 252

transmission and the basic types of reservoirs used by pathogenic bacteria and viruses. We illustrate these patterns through discussion of specific bacteria and viruses in Chapters 9 and 10, respectively.

Bacteria and viruses, while very different kinds of infectious agents, share many similarities in their basic life history strategies, including their means of transmission to susceptible hosts and the basic types of reservoirs they use for their long-term survival and success. To reduce redundancy, and to accentuate these important patterns, we give a brief introduction to both bacteria and viruses in this chapter, followed by an overview of the most common patterns of



246 247

Bacteria The Kingdom Monera (Whittaker 1969) includes both the Archaebacteria and



241

Eubacteria (“true bacteria”) (Keeton and Gould 1993b). Of these, only the Eubacteria are considered of importance in the study of wildlife diseases, and the term “bacteria” generally will be used for this group. Bacteria are single-celled prokaryotic organisms or associations of simple cells with cellular rather than organismal properties (Holt et al. 1994); occasional filamentous or mycelial forms also may occur. Bacteria generally have single-stranded DNA; cell division normally is by binary fission rather than mitosis or meiosis, and does not involve any cyclical changes in the nucleoplasm or cytoplasm (Holt et al. 1994). Bacteria have ribosomes, and the 70S type ribosome is dispersed in the cytoplasm. Most have a rigid cell wall, although this is lacking in some forms such as rickettsiae, Mycoplasma spp., and Chlamydia spp. Bacteria lack nuclear membranes, endoplasmic reticula, Golgi apparatuses, mitochondria, lysosomes, and most other cellular organelles. Bacteria are ancient organisms (Keeton and Gould 1993a, 1993b), with evidence for their presence in the Precambrian Period. Collectively, they occur in a broader variety of habitats and lifestyles than do eukaryotic organisms (Keeton and Gould 1993a). Historically, there has been little consensus on eubacterial evolution. Bacteria were among the first life forms, and one form recently was proposed as the last universal common ancestor (Ciccarelli et al. 2006). Classification now is largely phylogenetically based, and molecular methods are used to establish these relationships (Grimont 1999). Currently, about 35 major groups are recognized among the bacteria, of which about 17 have species of specific interest to the study of wildlife diseases (App. 1: Table 7). However, these phylogenetic relationships can become complicated due to the common mobility of bacterial genes among unrelated groups (Narra and Ochman 2006, Bohannon 2008, Sheppard et al. 2008). Bergey’s Manual of Systematic Bacteriology (Holt 1984), Bergey’s Manual of Determinative Bacteriology (Holt et al. 1994), and the American Society for Microbiology’s Manual of Clinical Microbiology (Murray et al. 1999) contain contemporary 242   introduction to non-eukaryotic agents

bacterial classification schemes. A new multivolume series of Bergey’s Manual of Systematic Bacteriology recently has become available.

Bacterial Identification Identification of bacteria is based on their metabolic activities, as determined by the enzymes they carry, as well as their cell wall characteristics and internal biochemistry. Typically, bacterial samples from a diseased host or other source are collected first; the suspect bacteria then are cultivated and isolated on bacteriological media (Miller and Holmes 1999, Reisner et al. 1999). The gram stain, based on differential presence of lipopolysaccharides in the cell wall, is probably the most important of many stains used in bacterial identification (Chapin and Murray 1999). In addition, bacterial identification typically involves establishing the presence of key enzymes such as those used to catabolize sugars such as glucose, lactose, or sucrose; there are a wide variety of manual and automated systems for establishing profiles for key enzymes of bacteria (Miller and O’Hara 1999). Also, the presence of a bacterial agent may be determined by amplifying small remnants of its DNA through PCR until the DNA can be detected and identified (App. 2). The species and serological group (serotype) to which a bacterium belongs often can be determined from specific serological tests, including immunof luorescence, immunoperoxidase, enzyme-linked immunosorbent assays (ELISA), histochemistry, and immunodiffusion tests (Gough 1997, Mahony and Chernesky 1999); some of the more commonly used ones are as described in Appendix 2. An increasing number of molecular techniques have emerged in recent years (Gough 1997), including the polymerase chain reaction (PCR), DNA sequencing, molecular hybridization, and restriction fragment length polymorphism; a few also are summarized in Appendix 2. In addition to isolating and identifying bacteria associated with a disease process, past bacterial infections also may be ascertained by

sampling hosts for the presence of antibodies to specific bacterial agents by the use of a number of serological tests, as outlined in Appendix 2.

Viruses While viruses possess some properties of living systems, including having a genome (Villarreal 2004), they probably are better described as nonliving infectious entities than as living microorganisms (van Regenmortel and Mahy 2004). Viruses generally lack ribosomes and any independent metabolic activities; they cannot generate ATP, and they cannot reproduce in the absence of a host. All viruses are obligate intracellular parasites and generally have no evidence of living qualities outside of an infected host. However, the finding that some viruses can be infected by other viruses and become “sick” in the process complicates the separation of viruses from other living organisms (Ogata and Claverie 2008). Free viral particles generally consist of a nucleic acid core surrounded by a protein coat (capsid); the complete structure is termed a nucleocapsid, and the individual protein units of the capsid are termed capsomeres (Voyles 2002). Nucleocapsids typically vary from 20 to 300 nanometers in size. Many animal viruses also have an envelope surrounding the capsid. When present, this envelope is derived from the cellular membrane of the host in which the virus replicated; the envelope also has additional viral proteins or glycoproteins used to bind to host cell receptors (Voyles 2002).

Viral Identification Individual viruses contain either DNA or RNA, but not both; single- and double-stranded forms of both RNA and DNA viruses occur. Classification of viruses is determined by their capsid size and shape (helical, icosahedral, complex), kind of viral nucleic acid (RNA or DNA), nucleic acid strandedness and direction of the nucleic acid twist, number of base pairs and base sequences of their RNA or DNA, and the

presence or absence of a lipoprotein envelope. Other taxonomically important features include the mode of replication, as well as the general chemical composition (antigenic properties) of the capsid (Melnick 1999, Murphy et al. 1999). Viral isolation often first requires isolating and cultivating the viruses in living, susceptible cells such as laboratory animals, embryonated hens’ eggs, or tissue culture cells. Viral identification is based on identifying the viral size and morphology through electron microscopy techniques; characterizing the viruses based on gel electrophoresis; DNA profiling by restriction endonuclease analysis, DNA hybridization, or polymerase chain reactions (PCR) (Gough 1997); and using specific antigen-detecting tests (App. 2). Many of the methods used to diagnose viruses are similar in principle to those applied to bacterial infections, particularly regarding the use of serology and recently developed molecular techniques; the primary differences between diagnosing bacterial and viral infections lies in the ability to isolate and cultivate bacteria in prepared sterile growth media, whereas viruses require susceptible living cells in which to replicate (Gough 1997). Viruses currently are separated into about 73 families and groups (Melnick 1999). Current taxonomic schemes for viruses have been maintained by the International Committee on Taxonomy of Viruses (ICTV) (BüchenOsmond 2003). Although the species concept for viruses is controversial and complicated (van Regenmortel and Mahy 2004), the ICTV currently recognizes about 3,000 viral species; however, at least 30,000 different viral strains and isolates have been identified by virologists (Büchen-Osmond 2003). A considerable number of viruses have been identified as important to the study of wildlife diseases (App. 1: Table 8).

Bacterial and Viral Transmission Patterns There is a rich variation in the strategies used by both bacteria and viruses for transmission to susceptible hosts (Smith 1982, Moore 1995, introduction to non-eukaryotic agents   243

Graczyk 2002, Moore 2002, Swinton et al. 2002, Wobeser 2006). One important distinction is between horizontal and vertical forms of transmission. Horizontal transmission refers to transmission within the same age cohort (same generation) of hosts; in contrast, vertical transmission is transmission between generations of host (e.g., mother to young). Among mammals, vertical transmission may occur through an intrauterine route or by ingestion of contaminated milk; among egg-laying animals, transmission can occur through infected eggs. Overall, vertical transmission is relatively uncommon among most wildlife diseases. Some reported examples include transmission of the bacterium Bartonella spp. in rodents (Kosoy et al. 1998) and the malignant catarrhal fever virus among wildebeest (Connochaetes spp.) (Plowright 1965) by an intrauterine route. The bacteria Mycobacterium bovis (Smith 1982) and Brucella abortus (Davis et al. 1990, Meyer and Meagher 1995, Thorne 2001) can be transmitted through infected milk among nursing ungulates. Also, duck plague virus (Burgess and Yuill 1981) and the bacterium Salmonella typhimurium (Friend 1999b) can be transmitted via infected eggs. Horizontal forms of transmission are far more common and involve a wide variety of strategies. In an effort to simplify this complexity, we organize the various forms of horizontal transmission into five broad patterns: physical contact, ingestion, aerosol transmission, arthropod transmission, and helminth-mediated transmission.

Physical Contact A variety of bacteria and viruses can be transmitted by direct physical contact including sexual activity, biting, and fomites. Examples of transmission by physical contact include papillomavirus infections in mammals (Sundberg et al. 2001), duck plague in waterfowl (Leibovitz 1971), rinderpest virus in ungulates (Rossiter 2001), pseudorabies virus in feral swine (Stallknecht and Howerth 2001), 244   introduction to non-eukaryotic agents

and malignant catarrhal fever virus (alcelaphine herpesvirus 1) in ungulates (Plowright 1965). Bites typically play an important role in transmission of rabiesvirus from infected to susceptible animals; also, the bacterium Pasteurella multocida can be transmitted from felids to wild birds during failed predation attempts (Smit et al. 1980, Korbel 1990). Chlamydia spp. can be transmitted by the venereal route (Whittington 2001). Direct urinary contamination of skin also can occur and may be particularly important in transmission of bacteria such as Leptospira spp. (Smith 1982). Some transmission also may occur through direct contact with fomites. Fomites are inanimate objects that can become contaminated by infectious material and serve as a source of these infective agents to susceptible hosts coming in contact with the contaminated surface. A bird feeder having perches contaminated with the bacterium Mycoplasma spp. from infected songbirds may serve as a source of these bacteria to other songbirds (Friend 1999a). The bacterium Yersinia pseudotuberculosis also is transmitted via fomites (Wetzler 1971)

Ingestion Ingestion of contaminated food or water is an important means of transmission for many bacteria and viruses (Smith 1982), especially intestinal bacteria and viruses transmitted by an fecal–oral route with contaminated food or water (Bopp et al. 1999, Farmer 1999). Predation or scavenging of infected prey or carcasses also are means by which bacteria and viruses are ingested (Rosen and Morse 1959, Smith 1982), as is the casual licking of infected carcasses by curious ungulates. Although a rare occurrence, bacteria and viruses can be transmitted by cannibalism; epizootics of bubonic plague, caused by the bacterium Yersinia pestis, can be sustained in colonies of laboratory rats through cannibalism alone (Rust et al. 1972). Francisella tularensis, the cause of tularemia, also can be spread among wild rodents by cannibalism (Olsuf’ev and Shlygina 1979).

Canine parvoviruses commonly are transmitted by ingestion of fecally contaminated food or water, or by contact with contaminated fomites near latrine sites, marking sites, or other areas contaminated by virus shedders (Barker and Parrish 2001).

Aerosol Transmission Some infective agents can be transmitted by means of contaminated aerosols through inhalation, nasal infection, or aerosol contact with the eyes. Among viruses, influenza virus (Slemon and Brugh 1994) and canine distemper virus (Williams 2001) commonly are transmitted in this fashion. Among bacteria, Pasteurella multocida can be transmitted to waterfowl through aerosol inhalation in the laboratory (Titche 1979); Mycoplasma conjunctivae also can be transmitted among susceptible ungulates through conjunctival contamination (Giacometti et al. 2002).

Arthropod Transmission Many viruses and bacteria can be transmitted by arthropods such as fleas, mosquitoes, lice, biting flies, ticks, or mites (Smith 1982). Viruses transmitted by arthropods include a variety of arboviruses (arthropod-borne viruses), including flaviviruses such as the West Nile virus, orthomyxoviruses such as eastern and western equine encephalitis viruses, as well as poxviruses such as myxoma and avian poxviruses. Among the bacteria using arthropod transmission are Borrelia burgdorferi, the cause of Lyme disease, Yersinia pestis, the cause of bubonic plague, Francisella tularensis, the cause of tularemia (Smith 1982, Williams and Barker 2001), and most rickettsiae. Most of these transmissions occur through the bite of hematophagous (“blood-feeding”) arthropods, but some also can result from surface feeding or excretions by flies or other infected arthropods (Smith 1982). For example, besides aerosol transmission, Mycoplasma conjunctivae also can be transmitted among

susceptible ungulates by flies feeding on eye secretions (Giacometti et al. 2002). Thus, in this reservoir, it becomes important to distinguish mere mechanical transmission by arthropods (mechanical vectors) from a relationship in which the bacteria and viruses invade and multiply in the arthropod tissues, and use the arthropod as an essential part of its development (biological vectors). As outlined in Chapter 5 (on Arthropoda), parasites using arthropods may use one of several types of transmission: mechanical, multiplicative (propagative), developmental, and cyclopropagative. Because there are no changes in life history stages among viruses and bacteria, their transmission is limited to the mechanical and propagative types.

Helminth-mediated Transmission Only a few bacteria and viruses are transmitted through the life cycle of a helminth parasite. However, members of the bacterial genus Neorickettsia are consistently transmitted through the life cycles of several intestinal trematodes (Rikihisa et al. 2004). Another bacterium, Brucella spp., may be transmitted by infected lungworms (Parafilaroides spp.) to harbor seals (Phoca vitulina richardsi) (Garner et al. 1997, Dunn et al. 2001). Viruses that have used helminths for transmission include swine influenza virus and hog cholera virus, both of which are associated with transmission through the swine lungworm, Metastrongylus spp. (Shope 1941, 1943a, 1943b, 1958; Sen et al. 1961; Shope 1965). The lymphocytic choriomeningitis virus may be transmitted through the life cycle of Trichinella spiralis (Syverton et al. 1947). Transmission of microorganisms to susceptible vertebrate hosts in this transmission pattern involves the same mechanisms as found for acquiring the respective affected helminths, such as ingestion of, or penetration by, infective helminth stages. Not surprisingly, some pathogens may use two or even several means of transmission. Francisella tularensis, the agent of tularemia, introduction to non-eukaryotic agents   245

can be transmitted by infected ticks, ingestion of infected prey or contaminated water, bacterial physical contact with lacerated or even intact skin, through ocular mucous membranes, or by inhalation of infected aerosols (Jellison 1974, Mörner and Addison 2001). Yersinia pestis most typically is transmitted by flea bite, but also can be transmitted by inhalation, as well as ingestion routes through predation, scavenging, and cannibalism; lacerations of mucous membranes by chewing on infected bones can enhance this transmission (Gasper and Watson 2001). Hosts have developed a number of important strategies to reduce the success of various parasite transmission strategies. Immune systems are an important component in fending off parasite attack. Physical barriers to parasite entry are also important. These were developed more fully in Chapter 2 (on immunity). Social barriers are an additional defense against successful transmission of infective agents (Loehle 1995). In contrast, habitat fragmentation and other anthropogenic disturbances have been influential in changing patterns of transmission (Goldberg et al. 2008).

Reservoirs Types used by Bacteria and viruses As with transmission patterns, there also can be considerable variation in the life history strategies used by bacteria and viruses for their long-term survival; transmission patterns and reservoir models often are closely linked. Applying the notion of reservoir (defined in Chapter 1 and the Glossary) to bacteria and viruses, we refer to an ecologic system that supports the survival needs of a population of bacteria or viruses over the long term; reservoirs are characterized by the combination of hosts and environmental factors the bacteria and viruses exploit for their survival and success. Reservoirs also must allow for a level of multiplication adequate to overcome the continual attrition bacteria and viruses experience. We simplify this complexity by emphasizing major types of reservoirs observed and 246   introduction to non-eukaryotic agents

providing salient examples of pathogens using these models. We have built on some of the initial insights of Shope (1965) regarding epidemiological patterns among viruses and rickettsiae, and have modified them in light of more recent knowledge of wildlife pathogens. There always is a risk in simplifying such complex matters. For example, reducing such complexity into simple broad patterns does not always allow a full appreciation of the rich and subtle variations that may occur among the bacteria and viruses classified within a reservoir type as they work out their own specific strategies for success. Also, this simplification does not always fully acknowledge that some bacteria and viruses may opportunistically use more than one reservoir type for their survival and success. Despite the risks of simplified systems, most pathogenic bacteria and viruses of importance to wildlife diseases can be reasonably described and understood in the context of one (or more) of these reservoirs, and there are advantages to thinking in these broader patterns. These patterns help clarify broader principles for understanding the ecology of the parasites and the strategies they use to survive. Recognizing that only a few basic patterns represent the strategies for survival and success of a wide diversity of viruses and bacteria emphasizes the importance of those reservoir types to bacteria and viruses as life history strategies. Understanding the characteristics, strengths, and weaknesses of those strategies is the basis for developing possible approaches for control, as well as insights on habitat changes or management practices that may exacerbate or reduce disease problems. And we hope that in the process of addressing some of the key species classified into these reservoir types, we can also give examples of some of the important variations found among the species within each reservoir type. We define five basic reservoir types. We first consider two reservoir models in which the microparasite depends primarily or exclusively on infected vertebrate hosts; one is a reservoir

dependent on apparently healthy carriers (latent infections), and the other is a reservoir dependent on the continuing presence of clinically active infections among the vertebrate hosts. We then consider two reservoir models in which the bacteria and viruses require both a vertebrate and an invertebrate host. One is a reservoir requiring both a vertebrate and an arthropod; in the other reservoir type, the microparasite uses a combination of a vertebrate and a helminth capable of infecting that vertebrate as a reservoir of the microparasite. Finally, we address bacteria and viruses that use soil or water as an integral part of their reservoir. Within these basic life history models, there is considerable variation in the specific strategies these bacteria and viruses employ for their long-term survival and success. In contrast to most eukaryotic parasites, for example, many bacteria and viruses are able to capitalize on two or even more life history models, as their circumstances warrant. Also, overt disease may not play a significant role in the strategies some bacteria and viruses use for survival and success. We first give a brief overview of these patterns in this introduction, and then develop each of them more fully in Chapters 9 (on bacteria) and 10 (on viruses). The particular bacteria and viruses addressed are often, but not always, significant mortality factors among wildlife. In other cases, the bacteria and viruses may be particularly good examples of microparasites using a specific reservoir, are important emerging diseases, or may illustrate special problems managers encounter with wildlife diseases.

Vertebrate-dependent Reservoirs Latent Infections (Apparently Healthy Carriers) In the latent infection reservoir, bacteria and viruses persist in the tissues and organs of a vertebrate host for extended periods, sometimes for the life of the infected host, and are shed by the infected carriers as the source of infection to other susceptible hosts. This type of reservoir is used by many pathogens

infecting the intestinal or respiratory tracts of vertebrates. Most commonly, newly infected animals become clinically ill on initial infection by the microparasite, recover, and then become recovered carriers, serving as a source of infection for other susceptible animals. There are a limited number of cases in which parasites live in apparently healthy carrier hosts that typically do not experience clinical illness (i.e., silent carriers), but these carriers then serve as a source of infection for other, susceptible hosts (i.e., indicator hosts) (Shope 1965). Latent infections may be the single most common reservoir type among bacteria. Examples of bacteria frequently using this life history pattern are found in a wide variety of taxonomic groups, and include Salmonella spp. (Matyas 1988), Brucella abortus (Thorne 2001), Chlamydophila (Chlamydia) psittaci (Meyer and Eddie 1932–33), Pasteurella multocida and Mannheimia (Pasteurella) haemolytica (Miller 2001), Mycoplasma spp. (Friend 1999a, Whithear 2001), and Fusobacterium necrophorum (Leighton 2001). This reservoir type also is common among viruses and involves many herpesviruses (Burnet and Williams 1939), including duck plague virus (Burgess et al. 1979), as well as hantaviruses (Childs et al. 1994), some retroviruses (Worley 2001), foot-and-mouth disease viruses (Thomson et al. 2001), and influenza viruses (Acha and Szyfres 2003). Most known examples of silent carrier infections involve viruses, and include herpesviruses such as malignant catarrhal fever virus (alcelaphine herpesvirus) (de Kock and Neitz 1950, Heuschele and Reid 2001), pigeon herpesvirus (Vindevogel and Pastoret 1981, Aini et al. 1993), and the Herpes B virus (Sabin and Wright 1934); another example is the African swine fever virus (Montgomery 1921). Viruses associated with silent carriers typically are contracted at an early age, cause mild or inapparent infections, and may persist indefinitely in the silent carrier hosts. In some cases, the bacteria and viruses may be difficult to detect in silent carriers (Shope 1965). introduction to non-eukaryotic agents   247

Depending on the site of infection in the vertebrate, the bacteria and viruses may be shed in feces, urine, mucous membranes, saliva, or conjunctival fluids. Typically, susceptible animals are infected by direct or close contact with an infected carrier, an infective aerosol, or ingestion of food or water contaminated by a shedding animal. The coexistence of infection and immunity often is a delicate balance that can easily shift from the immune system successfully suppressing the infective agent to the agent causing a relapse. Clinical relapses often occur among recovered carriers and can be exacerbated by stress (Shope 1965). Although this reservoir type is widespread among bacteria and viruses, there still is only limited information on effective prevention or control methods. One general strategy is to separate susceptible animals from sources of infection, including infected carrier animals and habitats contaminated with infective agents by carriers. Approaches used sometimes involve finding and eliminating carriers through test and slaughter or vaccination programs; this often is used among the hosts for bacteria and viruses believed to be a risk for humans or domestic animals. Alternatively, one can treat known carriers with antibiotics if they are part of a sensitive population or a high-profile species. Clinically Active Infections In this reservoir type, the bacteria and viruses are dependent on an unbroken series of clinically active infections among susceptible hosts to sustain the parasite population among vertebrate hosts of a region. Currently infected hosts or recovered immune hosts are not considered part of the pool of susceptible hosts that must be available (Swinton et al. 2002). Very few bacteria depend primarily on clinical infections as their primary reservoir for long-term sustainability. Mycobacterium spp., such as M. bovis in free-ranging deer (Schmitt et al. 1997), are presented as one example. Neisseria gonorrhoeae, a venereal disease among 248   introduction to non-eukaryotic agents

humans, as well as some virulent strains of Streptococcus spp., also can fit into this reservoir type (Knapp and Koumans 1999, Ruoff et al. 1999). This reservoir type is found more commonly among viruses, including rabies and other lyssaviruses (Baer 1975), some avian poxviruses (Karstad 1971b), morbilliviruses such as rinderpest virus (Rossiter 2001), and many papillomaviruses (Sundberg et al. 2001). Bacteria and viruses in this life history group typically are transmitted through direct contact with infected animals or contaminated fomites, ingestion of fecally contaminated food or water, droplet inhalation, or salivary contamination, including bites. Parasites in this reservoir type tend to be density-dependent, and their risk of transmission is increased as the density of susceptible animals is increased; conversely, the risk of transmission tends to decline as the density of susceptible hosts declines. Overall, the effects of immunity are the same as death; they remove susceptible hosts and effectively reduce the density of the population susceptible to infection. For infections typically leading to rapid disease outcomes (immunity or death), as caused by some viruses, success of the microparasite is increased with relatively shorter incubation times between initial infection and the start of shedding, relatively longer time periods during which hosts are able to shed the bacteria and viruses, higher numbers (intensity) of bacteria and viruses shed by the clinically ill host over time, and an efficient means of transmission to new susceptible hosts. With such parasites, epizootics may occur at regular intervals, partly depending on the rate at which the host population increases during recovery from a previous epizootic. New epizootics begin both when the population of susceptible animals becomes dense enough to support a chain of transmission and when the infective agent is reintroduced from another, infected host population. Among many fast-acting viral diseases, surviving hosts have a strong, even lifelong immunity and do not contribute further to maintaining the parasite (Swinton et al. 2002).

In such cases the susceptible host populations often are composed of relatively young and previously unexposed hosts. For a few bacteria and viruses, such as mycobacteriae and poxviruses, clinical diseases tend to develop slowly and persistently, eventually resulting in either a domination by the parasite and death of the host, or domination of the parasite by the host and resolution of the disease through host immunity; yet in many cases the host may die from other causes before the disease balance is resolved. In such slowdeveloping diseases as tuberculosis, host mortality typically is well below the replacement rate from typical host reproductive success and there is an adequate addition of young, previously unexposed hosts to sustain the bacterium. Distinguishing whether bacteria and viruses are using a recovered carrier or a clinical infection type of reservoir occasionally can be difficult, as when there appears to be a very low prevalence of carriers detected in an infected population. This can occur because the detectable numbers of microparasites in the recovered carriers drop below detection levels based on current laboratory techniques, as with Salmonella spp.; it may only be after a host is stressed that the microparasite numbers rise again to become detectable (Williams and Newell 1970). But in general, bacteria and viruses in the clinically active infection reservoir do not use previously infected hosts as continuing sources of infections following their recovery, nor do they generally causes clinical relapses in these former hosts, in contrast to bacteria and viruses using a carrier reservoir. In some cases, bacteria and viruses may have a carrier state in one species, but can cause epizootics in another. Examples include some avian influenza viruses (Webster et al. 1992) and Mycoplasma gallisepticum (Dhondt et al. 1998) in birds. Generally, however, hosts experiencing these epizootics are not able to sustain the microparasite in the absence of the other infected carrier species. In such cases, the bacteria and viruses still are classified as part of the latent infection reservoir.

Control typically is directed at breaking the chain of infection by reducing the density of susceptible hosts to a level below the parasite’s requirement to maintain a chain of infection. Two strategies commonly have been employed. Population reduction has been used to reduce the density of infected and susceptible animals (e.g., rabies, rinderpest). Also, vaccination programs typically have been used with captive wildlife, domestic animals, and humans, when vaccines are available. Use of vaccination in free-living wildlife is an emerging technology that holds considerable promise.

Invertebrate–Vertebrate Reservoirs Two additional models are addressed under this general category: arthropod–vertebrate and helminth–vertebrate reservoirs. Both reservoirs are complicated from an ecological standpoint because they involve interactions with two hosts (invertebrate, vertebrate), each with very different physiological and ecological characteristics. With the helminth–vertebrate system, the physiological and ecological constraints superimposed by any helminth intermediate hosts add an even greater complexity to the relationships considered. Arthropod–Vertebrate Reservoir This reservoir type generally entails the combination of a vertebrate host and a hematophagous arthropod. Common examples of arthropods involved include fleas, mosquitoes, lice, biting flies, ticks, and mites. Occasionally, nonhematophagous arthropods such as flies feeding on conjunctival fluids or other infected materials can regularly transmit infectious agents. Bacteria and viruses using this reservoir typically multiply in their invertebrate host, and thus have a propagative form of transmission. Examples of bacteria classified in this reservoir type include tick-borne parasites such as Borrelia burgdorferi, the cause of Lyme disease (Brown and Burgess 2001), Francisella tularensis, the cause of tularemia (Mörner and Addison 2001), Anaplasma spp, the cause of granulocytic introduction to non-eukaryotic agents   249

ehrlichiosis (Davidson and Goff 2001), and the flea-borne bacterium Yersinia pestis, causative agent of sylvatic and bubonic plague (Gasper and Watson 2001). Examples of viruses using this reservoir type include numerous arboviruses (Karstad 1971a, Yuill and Seymour 2001), such as the mosquito-borne flaviviruses of West Nile disease and eastern and western equine encephalitis, the Culicoides-borne hemorrhagic disease orbiviruses of cervids (Howerth et al. 2001), and the flea-borne myxoma and other, mosquito-borne, poxviruses (Karstad 1971b, Robinson and Kerr 2001). There are important distinctions in the disease ecology between bacteria and viruses transmitted by free-flying arthropods or ticks, compared to those transmitted by more hostbound ectoparasites such as lice and fleas (Smith 1982). Arthropods that live much of their life independent of the host often are more affected by the ecological constraints of their specific habitat requirements, including seasonality. This is particularly true regarding the seasonal availability of arthropods in more temperate regions where over-wintering strategies may become an important problem for many of these bacteria and viruses. Consequently, the success of parasites using this reservoir type is affected by the specific ecological and habitat constraints of both vertebrate and invertebrate. Control strategies typically are aimed at reducing arthropod populations or preventing infected arthropods from reaching susceptible hosts. Less commonly, treating or immunizing susceptible vertebrate hosts is used. Helminth–Vertebrate Reservoir Bacteria and viruses in this reservoir model are in a complex relationship involving both vertebrate hosts and one or more helminth parasites infecting that vertebrate. As with the arthropod–helminth model, the ecological relationships in this model can be complicated by the special physiological and ecological factors resulting from involvement of both helminths and vertebrates, in addition to any intermediate hosts for the carrier helminth. One important 250   introduction to non-eukaryotic agents

feature proposed for this model is that the microparasite may survive for extended periods in the helminth between periods in which it causes clinical disease among vertebrates (Syverton et al. 1947, Shope 1965). One well-established example involving an obligatory relationship with helminths and vertebrates involves members of the bacterial genus Neorickettsia (Rikihisa et al. 2004). Claims of possible helminth–vertebrate reservoirs have been made for some viruses (Shope 1965); both the classic swine fever (hog cholera) virus (Shope 1958) and swine influenza virus (Shope 1941, 1943a, 1943b; Sen et al. 1961) cycle between swine and a swine lungworm, Metastrongylus spp. A similar claim for a helminth– vertebrate reservoir involving swine influenza virus among rats and mice, with a rat nematode (Strongyloides ratti), also has been made (Shotts et al. 1968). However, the evidence that helminths are an essential part of the reservoir for these viruses has been seriously challenged (Wallace 1977). Another bacterium, Brucella spp., recovered from the harbor seal (Phoca vitulina richardsi), may use infected lungworms (Parafilaroides spp.) as part of its reservoir (Garner et al. 1997, Dunn et al. 2001). One recent example of interest is the San Miguel Sea Lion Virus 5, a virus that may be maintained by a combination of marine mammals and lungworm Parafilaroides decorus, as well as a liver fluke (Zalophatrema spp.) (Smith et al. 1978, 1980; Smith and Boyt 1990; Smith et al. 1998; Kennedy-Stoskopf 2001). However, there is a need for further clarification on the role of the helminth as part of the viral reservoir. Other than for Neorickettsia spp., the overall importance of helminths in the reservoirs of these bacteria and viruses has been unclear. While a number of viruses and bacteria may capitalize on helminths as one of several means for transmission, it is more likely that only a few bacteria and viruses require the helminth as an essential part of their reservoir. For many of the viruses, the helminth–vertebrate relationship may be only one part of a more complex reservoir system that also involves latent infections

in one or more vertebrate species. For example, waterfowl and shorebird carriers are an important source for many influenza viruses affecting other species (Stallknecht and Shane 1988, Webster et al. 1992, Campen and Early 2001). There also is evidence that the San Miguel sea lion virus may be a parasite of the opaleye perch (Girella nigricans), with the perch playing an important role in the ongoing occurrence of this virus among marine mammals (Smith and Boyt 1990, Kennedy-Stoskopf 2001). Thus, this reservoir type currently involves few clearly established examples, and for most bacteria and viruses proposed for this reservoir type, the role of the helminth in the survival and success of these microparasites must be more clearly established. Interestingly, this reservoir type has been used successfully by eukaryotic as well as prokaryotic organisms of concern to wildlife. The flagellated protozoon Histomonas meleagridis is closely tied to the life cycle of the nematode Heterakis gallinae for its survival and success (Graybill and Smith 1920, Wehr 1971); this parasite was addressed in Chapter 6 (on Protista). However, with the apparent success of a helminth–vertebrate relationship among at least a few bacteria and viruses, it seems likely that, with searching, other examples will emerge. For obligatory helminth–vertebrate relationships, control likely would be the same as that directed toward control of helminth infections; an attempt could be made to break the helminth life cycle. For relationships involving enhanced transmission, but non-obligatory relations between the microparasite and helminth, helminth control would have only limited success. In all cases, one could immunize or treat individual hosts if they were of particular importance to human or domestic animals health, or were members of listed species.

Soil and Water as Reservoirs In this reservoir model, we include bacteria and viruses with a range of ecological capabilities. All established examples occur among bacteria.

Such organisms share a capacity to survive for extended periods in soil and water, and can use this capacity as an important part of their longterm strategy for survival and success. A few bacteria in this reservoir live independently as true saprophytes in the environment, including Listeria monocytogenes and Nocardia spp. For these cases, infections in vertebrate hosts are an insignificant aspect of the bacterial reservoir because infected animals are “dead-end” hosts; these bacteria rarely are transmitted directly from infected animals to other susceptible hosts. In addition, Clostridium botulinum, a bacterium causing an intoxication from a neurotoxin rather than a true bacterial infection, is considered in this section because of its long survival in wetlands; it also is saprophytic, depending on decaying animal matter (Bell et al. 1955, Dodds 1992). We also include in this reservoir bacteria and viruses that may depend upon another reservoir, but also have the ability to survive and endure in soil or water for such an extended period that the environment becomes an important part of their strategy for success and, for management purposes, it becomes sensible to consider soil or water as a significant part of their overall reservoir. These examples are virtually all bacteria. Some of these bacteria are linked to carrier animals and ultimately depend on them as a source of organisms to contaminate the environment, but also have extended survival in the environment. In these cases, the infected animals and soil and water can serve as important complements in the reservoir. Examples include Bacillus anthracis (Logan and Turnbull 1999), Mycobacterium spp. (Metchock et al. 1999), Aeromonas hydrophila (Altwegg 1999), and Salmonella spp. (Winfield and Groisman 2003). Further, we note the extended capacity of a few viruses to survive in the environment as an important element of their endurance between active infections and transmission, even though they are inert and lack any evidence for metabolism or replication outside of a living host cell. Examples include parvoviruses, avian introduction to non-eukaryotic agents   251

influenza viruses, enteroviruses, and hepatitis A virus (Hurst et al. 1980, Yates et al. 1985, Gordon and Angrick 1986, Stallknecht et al. 1990, Gantzer et al. 1998, Brown et al. 2007). Bacteria and viruses using soil or water as part of their reservoir typically are transmitted to susceptible hosts by ingestion of food or water contaminated from environmental sources. Occasionally these bacteria and viruses can be transmitted through wounds that came into contact with contaminated soil or water. Diseases associated with soil and water reservoirs are classically density-independent diseases. Varying the density of the hosts has little or no impact on the risk of infection to an individual in a population. The capacity for some bacteria not normally considered soil and water organisms to survive in the environment for extended periods can be influenced by their ability to successfully infect free-living amebae; after being ingested by the amebae, the bacteria grow, multiply, and eventually lyse the amebae and are released back into the environment. In addition to occurring with the soil saprophyte Listeria monocytogenes (Ly and Müller 1990), this unique relationship has been linked with extended environmental survival for Francisella tularensis (Abd et al. 2003), Salmonella enterica (Tezcan-Merdol et al. 2004), Chlamydia pneumoniae (Essig et al. 1997), Pasteurella multocida (Hundt and Ruffolo 2005), and Mycobacterium avium (Cirillo et al. 1997, Steinert et al. 1998), among others. Thus, even where these bacteria are not true soil and water organisms, their extended capacity to survive in the environment by parasitizing protozoa complicates the development of strategies to control them. Control strategies most commonly involve keeping susceptible hosts out of environmentally contaminated areas. In some cases, one can treat hosts contracting the illness. Both strategies have been used in the case of avian botulism. As noted in the previous discussion, there is evidence that some pathogens regularly use more than one type of reservoir. As an example, 252   introduction to non-eukaryotic agents

Francisella tularensis may use recovered carrier animals such as voles (Microtus spp.) (Olsuf’ev and Shlygina 1979, Shlygina and Olsuf’ev 1982), can be part of an arthropod–vertebrate reservoir (Jellison 1974), and can live in natural waters in the absence of known infections among animals (Parker et al. 1951, Ul’yanova et al. 1982, Mörner and Addison 2001). Literature Cited Abd, H., T. Johansson, I. Golovliov, G. Sandström, and M. Forsman. 2003. Survival and growth of Francisella tularensis in Acanthamoeba castellanii. Applied and Environmental Microbiology 69:600–606. Acha, P. N., and B. Szyfres. 2003. Influenza. Pp. 155–172 in Zoonoses and communicable disease common to man and animals. Scientific and Technical Publication No. 580. Pan American Health Organization, Washington, DC. Aini, I., L. M. Shih, A. E. Castro, and Y. C. Zee. 1993. Comparison of herpesvirus isolates from falcons, pigeons, and psittacines by restriction endonuclease analysis. Journal of Wildlife Diseases 29:196–202. Altwegg, M. 1999. Aeromonas and Plesiomonas. Pp. 507–516 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Baer, G. M. (editor). 1975. The natural history of rabies. Academic Press, New York. Barker, I. K., and C. R. Parrish. 2001. Parvovirus infections. Pp. 131–146 in I. K. Barker and C. R. Parrish (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Bell, J. F., G. W. Sciple, and A. A. Hubert. 1955. A microenvironment concept of the epizoology of avian botulism. Journal of Wildlife Management 19:352–357. Bohannon, J. 2008. Confusing kinships. Science 320:1031–1033. Bopp, C. A., F. W. Brenner, J. G. Wells, and N. A. Strockbine. 1999. Escherichia, Shigella, and Salmonella. . Pp. 459–474 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken, editors. Manual of clinical microbiology. ASM Press, Washington, DC. Brown, J. D., D. E. Swayne, R. J. Cooper, R. E. Burns, and D. E. Stallknecht. 2007. Persistence of H5 and H7 avian influenza viruses in water. Avian Diseases 51:285–289.

Brown, R. N., and E. C. Burgess. 2001. Lyme borreliosis. Pp. 435–454 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Büchen-Osmond, C. 2003. Taxonomy and classification of viruses. Pp. 1217–1226 in Manual of clinical microbiology. ASM Press, Washington, DC. Burgess, E. C., J. Ossa, and T. M. Yuill. 1979. Duck plague: A carrier state in waterfowl. Avian Diseases 23:940–949. Burgess, E. C., and T. M. Yuill. 1981. Vertical transmission of duck plague virus (DPV) by apparently healthy DPV carrier waterfowl. Avian Diseases 25:795–800. Burnet, F. M., and S. W. Williams. 1939. Herpes simplex: A new point of view. Medical Journal of Australia 1:637–642. Campen, V., and G. Early. 2001. Orthomyxovirus and paramyxovirus infections. Pp. 271–279 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Chapin, K. C., and P. R. Murray. 1999. Stains. Pp. 1674–1686 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Childs, J. E., T. G. Ksiazek, C. F. Spiropoulou, J. W. Krebs, S. Morzunov, G. O. Maupin, K. L. Gage, P. W. Rollin, J. Sarisky, R. E. Enscore, et al. 1994. Serologic and genetic identification of Peromyscus maniculatus as the primary rodent reservoir for a new hantavirus in the southwestern United States. Journal of Infectious Diseases 169:1271–1280. Ciccarelli, F. D., T. Doerks, C. vonMering, C. J. Creevey, B. Snel, and P. Bork. 2006. Toward automatic reconstruction of a highly resolved tree of life. Science 311:1283–1287. Cirillo, J. D., S. Falkow, L. S. Tompkins, and L. E. Bermudez. 1997. Interaction of Mycobacterium avium with environmental amoebae enhances virulence. Infection and Immunity 65:3759–3767. Davidson, W. R., and W. L. Goff. 2001. Anaplasmosis. Pp. 455–466 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Davis, D. S., J. W. Templeton, T. A. Ficht, J. D. Williams, J. D. Kopec, and L. G. Adams. 1990. Brucella abortus in captive bison. I: Serology, bacteriology, pathogenesis, and transmission to cattle. Journal of Wildlife Diseases 26:360–371. de Kock, G., and W. O. Neitz. 1950. Sheep as a reservoir of snotsiekte (or bovine malignant catarrhal

fever) of cattle in South Africa. South African Journal of Science 46:176–180. Dhondt, A. A., D. L. Tessaglia, and R. L. Slothower. 1998. Epidemic mycoplasmal conjunctivitis in house finches from eastern North America. Journal of Wildlife Diseases 34:265–280. Dodds, K. L. 1992. Clostridium botulinum in the environment. Pp. 21–52 in A. H. W. Hauschild and K. L. Dodds (editors), Clostridium botulinum: Ecology and control in foods. Marcell Dekker, New York. Dunn, J. L., J. D. Buck, and T. R. Robeck. 2001. Bacterial diseases of cetaceans and pinnipeds. Pp. 309–335 in L. A. Dierauf and F. M. D. Gulland (editors), CRC handbook of marine mammal medicine. CRC Press, Boca Raton, FL. Essig, A., M. Heinemann, U. Simnacher, and R. Marre. 1997. Infection of Acanthamoeba castellanii by Chlamydia pneumoniae. Applied and Environmental Microbiology 63:1396–1399. Farmer, J. J., III. 1999. Enterobacteriaceae: Introduction and identification. Pp. 442–458 in P. R. Murray, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Friend, M. 1999a. Mycoplasmosis. Pp. 115–119 in M. Friend and J. C. Franson (editors), Field manual of wildlife diseases: General field procedures and diseases of birds. Information and Technology Report 1999-001. U.S. Geological Survey, Washington, DC. Friend, M. 1999b. Salmonellosis. Pp. 99–109 in M. Friend and J. C. Franson (editors), Field manual of wildlife diseases: General field procedures and disease of birds. Information and Technology Report 1999-001. U.S. Geological Survey, Washington, DC. Gantzer, C., E. Dubois, J.-M. Crance, S. Billaudel, H. Kopecka, L. Schwartzbrod, M. Pommepuy, and F. L. Guader. 1998. Influence of environmental factors on the survival of enteric viruses in seawater. Oceanologica Acta 21:983–992. Garner, M. M., D. M. Lambourn, S. J. Jeffries, P. B. Hall, J. C. Rhyan, D. R. Ewalt, L. M. Polzin, and N. F. Cheville. 1997. Evidence of Brucella infection in Parafilaroides lungworms in a Pacific harbor seal (Phoca vitulina richardsi). Journal of Veterinary Diagnostic Investigation 9:298–303. Gasper, P. W., and R. P. Watson. 2001. Plague and yersiniosis. Pp. 313–329 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Giacometti, M., M. Janovsky, L. Belloy, and J. Frey. 2002. Infectious keratoconjunctivitis of ibex, chamois and other Caprinae. Revue Scientifique

introduction to non-eukaryotic agents   253

et Technique Office International des Epizooties 21:335–345. Goldberg, T. L., T. R. Gillespie, I. B. Rwego, E. L. Estoff, and C. A. Chapman. 2008. Forest fragmentation as cause of bacterial transmission among nonhuman primates, humans, and livestock, Uganda. Emerging Infectious Diseases 14:1375–1382. Gordon, J. C., and E. J. Angrick. 1986. Canine parvovirus: Environmental effects on infectivity. American Journal of Veterinary Research 47:1464–1467. Gough, R. E. 1997. Collection and identification of avian viruses, bacteria, and fungi. Pp. 379–395 in D. H. Clayton and J. Moore (editors), Host– parasite evolution: General principles and avian models. Oxford University Press, Oxford, UK. Graczyk, T. K. 2002. Zoonotic infections and conservation. Pp. 220–228 in R. S. O. A. A. Aguirre, G. M. Tabor, C. House, and M. C. Pearl (editors), Conservation medicine: Ecological health in practice. Oxford University Press, New York. Graybill, H. W., and T. Smith. 1920. Production of fatal blackhead in turkeys by feeding embryonated eggs of Heterakis papillosa. Journal of Experimental Medicine 31:647–655. Grimont, P. A. D. 1999. Taxonomy and classification of bacteria. Pp. 249–261 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Heuschele, W. P., and H. W. Reid. 2001. Malignant catarrhal fever. Pp. 157–164 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Holt, J. G. (editor-in-chief). 1984. Bergey’s Manual of systematic bacteriology. Williams & Wilkins, Baltimore, MD. Holt, J. G., N. R. Krieg, P. H. A. Sneath, J. T. Staley, and S. T. Williams. 1994. Bergey’s Manual of determinative bacteriology. 9th ed. Lippincott Williams & Wilkins, Philadelphia, PA. Howerth, E. W., D. E. Stallknecht, and P. D. Kirkland. 2001. Bluetongue, epizootic hemorrhagic disease, and other orbivirus-related diseases. Pp. 77–97 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Hundt, M. J., and C. G. Ruffolo. 2005. Interaction of Pasteurella multocida with free-living amoeba. Applied and Environmental Microbiology 71:5458–5464. Hurst, C. J., C. P. Gerba, and I. Cech. 1980. Effects of environmental variables and soil characteristics on virus survival in soil. Applied and Environmental Microbiology 40:1067–1079.

254   introduction to non-eukaryotic agents

Jellison, W. L. 1974. Tularemia in North America, 1930–1974. University of Montana, Missoula, MT. Karstad, L. 1971a. Arboviruses. Pp. 17–21 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Karstad, L. 1971b. Pox. Pp. 34–41 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Keeton, W. T., and J. L. Gould. 1993a. Biological science. 5th ed. Vol. 1. W. W. Norton & Company, New York. Keeton, W. T., and J. L. Gould. 1993b. Biological science. 5th ed. Vol. 2. W. W. Norton & Company, New York. Kennedy-Stoskopf, S. 2001. Viral diseases. Pp. 285–307 in L. A. Dierauf and F. M. D. Gulland (editors), CRC handbook of marine mammal medicine. CRC Press, Boca Raton, FL. Knapp, J. S., and E. H. Koumans. 1999. Neisseria and Branhamella. Pp. 586–603 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Korbel, R. 1990. Epizootiologie, Klinik und Therapie der Pasteurella-multocida-Infektion beim Vogelpatienten nach Katzenbiss. Tierärtzliches Praxis 18:365–376. Kosoy, M. Y., R. L. Regnery, O. I. Kosaya, D. C. Jones, E. L. Marston, and J. E. Childs. 1998. Isolation of Bartonella spp. from embryos and neonates of naturally infected rodents. Journal of Wildlife Diseases 34:305–308. Leibovitz, L. 1971. Duck plague. Pp. 22–33 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Leighton, F. A. 2001. Fusobacterium necrophorum infection. Pp. 493–496 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Loehle, C. 1995. Social barriers to pathogen transmission in wild animal populations. Ecology 76:326–335. Logan, N. A., and P. C. B. Turnbull. 1999. Bacillus and recently derived genera. Pp. 357–369 in P. R. Murray, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Ly, T. M., and H. E. Müller. 1990. Ingested Listeria monocytogenes survive and multiply in protozoa. Journal of Medical Microbiology 33:51–54.

Mahony, J. B., and M. A. Chernesky. 1999. Immunoassays for the diagnosis of infectious diseases. Pp. 202–214 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Matyas, Z. (chair, Expert Committee). 1988. Salmonellosis control: The role of animal and product hygiene. Technical Report Series 774. 92 4 120774 4. World Health Organization, Geneva, Switzerland. Melnick, J. L. 1999. Taxonomy and classification of viruses. Pp. 835–842 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Metchock, B. G., F. S. Nolte, and J. R. J. Wallace. 1999. Mycobacterium. Pp. 399–437 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Meyer, K. F., and B. Eddie. 1932–33. Latent psittacosis infection in shell parakeets. Proceedings of the Society for Experimental Biology and Medicine 30:484–488. Meyer, M. E., and M. Meagher. 1995. Brucellosis in free-ranging bison (Bison bison) in Yellowstone, Grand Teton, and Wood Bison National Parks: A review. Journal of Wildlife Diseases 31:579–598. Miller, J. M., and H. T. Holmes. 1999. Specimen collection, transport, and storage. Pp. 33–63 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Miller, J. M., and C. M. O’Hara. 1999. Manual and automated systems for microbial identification. Pp. 193–201 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Miller, M. W. 2001. Pasteurellosis. Pp. 330–339 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Montgomery, R. E. 1921. On a form of swine fever occurring in British East Africa (Kenya Colony). Journal of Comparative Pathology and Therapeutics 34:159–191, 243–262. Moore, J. 1995. The behavior of parasitized animals. BioScience 45:89–96. Moore, J. 2002. Parasites and the behavior of animals. Oxford University Press, Oxford, UK. Mörner, T., and E. Addison. 2001. Tularemia. Pp. 303–312 in E. S. Williams and I. K. Barker

(editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Murphy, F. A., E. P. J. Gibbs, M. C. Horzinek, and M. J. Studdert. 1999. Veterinary virology. 3rd ed. Academic Press, San Diego, CA. Murray, P. R., E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken. 1999. Manual of clinical microbiology. 7th ed. American Society for Microbiology, Washington, DC. Narra, H. P., and H. Ochman. 2006. Of what use is sex to bacteria? Current Biology 16:R705–R710. Ogata, H., and J.-M. Claverie. 2008. How to infect a mimivirus. Science Express 321:1305–1306. Olsuf’ev, N. G., and K. N. Shlygina. 1979. Role of cannibalism in the development of tularemia with nonlethal outcome in common voles highly sensitive to tularemia [in Russian]. Zoologicheskii Zhurnal 58:933–936. Parker, R. R., E. A. Steinhaus, G. M. Kohls, and W. L. Jellison. 1951. Contamination of natural waters and mud with Pasteurella tularensis and tularemia in beavers and muskrats in the northwestern United States. National Institutes of Health Bulletin 193:1–61. Plowright, W. 1965. Malignant catarrhal fever in East Africa. II: Observations on wildebeest calves at the laboratory and contract transmission of the infection to cattle. Research in Veterinary Science 6:69–83. Reisner, B. S., G. L. Woods, J. R. B. Thomson, D. H. Larone, L. S. Garcia, and R. Y. Shimizu. 1999. Specimen processing. Pp. 64–104 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Rikihisa, Y., C. Zhang, M. Kanter, Z. Cheng, N. Ohashi, and T. Fukuda. 2004. Analysis of p51, groESL, and the major antigen P51 in various species of Neorickettsia, and obligatory intracellular bacterium that infects trematodes and mammals. Journal of Clinical Microbiology 42:3823–3826. Robinson, A. J., and P. J. Kerr. 2001. Poxvirus infections. Pp. 179–201 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Rosen, M. N., and E. E. Morse. 1959. An interspecies chain in a fowl cholera epizootic. California Fish and Game 45:51–56. Rossiter, P. 2001. Rinderpest. Pp. 37–45 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Ruoff, K. L., R. A. Whiley, and D. Beighton. 1999. Streptococcus. Pp. 283–296 in P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and

introduction to non-eukaryotic agents   255

R. H. Yolken (editors), Manual of clinical microbiology. ASM Press, Washington, DC. Rust, J. H., Jr., D. N. Harrison, and J. J. D. Marshall. 1972. Susceptibility of rodents to oral plague infection: A mechanism for the persistence of plague in inter-epidemic periods. Journal of Wildlife Diseases 8:127–133. Sabin, A. B., and A. M. Wright. 1934. Acute ascending myelitis following a monkey bite, with the isolation of a virus capable of reproducing the disease. Journal of Experimental Medicine 59:115–136. Schmitt, S. M., S. D. Fitzgerald, T. M. Cooley, C. S. Bruning-Fann, L. Sullivan, D. Berry, T. Carlson, R. B. Minnis, J. B. Payeur, and J. Sikarskie. 1997. Bovine tuberculosis in freeranging white-tailed deer from Michigan. Journal of Wildlife Diseases 33:749–758. Sen, J. G., G. W. Kelley, N. R. Underdahl, and G. A. Young. 1961. Transmission of swine influenza virus by lungworm migration. Journal of Experimental Medicine 113:517–520. Sheppard, S. K., N. D. McCarthy, D. Falush, and M. C. J. Maiden. 2008. Convergence of Campylobacter species: Implications for bacterial evolution. Science 320:237–239. Shlygina, K. N., and N. G. Olsuf’ev. 1982. Latent tularemia in common voles (Microtus subarvalis) in an experiment [in Russian]. Zhurnal Microbiologii, Epidemiologii i Immunologii no. 4:101–104 (cited in Mörner and Addison 2001). Shope, R. E. 1941. The swine lungworm as a reservoir and intermediate host for swine influenza virus. II: The transmission of swine influenza virus by the swine lungworm. Journal of Experimental Medicine 74:49–68. Shope, R. E. 1943a. The swine lungworm as a reservoir and intermediate host for swine influenza virus. IV: The demonstration of transmission of the virus and the provocation of influenza. Journal of Experimental Medicine 77:111–126. Shope, R. E. 1943b. The swine lungworm as a reservoir and intermediate host for swine influenza virus. IV: The demonstration of masked swine influenza virus in lungworm larvae and swine under natural conditions. Journal of Experimental Medicine 77:127–138. Shope, R. E. 1958. The swine lungworm as a reservoir and intermediate host for hog cholera virus. I: The provocation of masked hog cholera virus in lungworm-infested swine by Ascaris larvae. Journal of Experimental Medicine 107:609–622. Shope, R. E. 1965. Transmission of viruses and epidemiology of viral infections. Pp. 385–404 in F. L. Horsfall and I. Tamm (editors), Viral and

256   introduction to non-eukaryotic agents

rickettsial infections of man. J. B. Lippincott Company, Philadelphia, PA. Shotts, E. B., J. W. Foster, M. Brugh, H. E. Jordan, and J. L. McQueen. 1968. An intestinal threadworm as a reservoir and intermediate host for swine influenza virus: A confirmation and amplification of Shope’s syndrome. Journal of Experimental Medicine 127:359–369. Slemon, R. D., and M. Brugh. 1994. Influenza. Pp. 385–395 in G. W. Beran and J. H. Steele (editors), Handbook of zoonoses, Section B: Viral. CRC Press, Boca Raton, FL. Smit, T., L. G. Moraal, and T. Bakhuizen. 1980. Pasteurella multocida infecties bij vogels na een kattebeet. Tijdschrift voor Diergeneeskunde 106:327–329. Smith, A. W., and P. M. Boyt. 1990. Caliciviruses of ocean origin: A review. Journal of Zoo and Wildlife Medicine 23:3–23. Smith, A. W., D. E. Skilling, and R. J. Brown. 1980. Preliminary investigation of a possible lung worm (Parafilaroides decorus), fish (Girella nigricans), and marine mammal (Callorhinus ursinus) cycle for San Miguel sea lion virus type 5. American Journal of Veterinary Research 41:1846–1850. Smith, A. W., D. E. Skilling, N. Cherry, J. H. Mead, and D. O. Matson. 1998. Calicivirus emergence from ocean reservoirs: Zoonotic and interspecies movements. Emerging Infectious Diseases 4:13–20. Smith, A. W., D. E. Skilling, and A. E. Ritchie. 1978. Immunoelectron microscopic comparisons of caliciviruses. American Journal of Veterinary Research 39:1531–1533. Smith, C. E. G. 1982. Major factors in the spread of infections. Pp. 207–235 in M. A. Edwards and U. McDonnell (editors), Animal disease in relation to animal conservation. Symposia of the Zoological Society of London Number No. 50. Academic Press, London, UK. Stallknecht, D. E., and E. W. Howerth. 2001. Pseudorabies (Aujeszky’s disease). Pp. 164–170 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Stallknecht, D. E., M. T. Kearney, S. M. Shane, and P. J. Zwank. 1990. Effects of pH, temperature, and salinity on persistence of avian influenza viruses in water. Avian Diseases 34:412–418. Stallknecht, D. E., and S. M. Shane. 1988. Host range of avian influenza virus in free-living birds. Veterinary Research Communications 12:125–141. Steinert, M., K. Birkness, E. White, B. Fields, and F. Quinn. 1998. Mycobacterium avium bacilli grow saprozoically in coculture with

Acanthamoeba polyphaga and survive within cyst walls. Applied and Environmental Microbiology 64:2256–2261. Sundberg, J. P., M. vanRanst, and A. B. Jenson. 2001. Papillomavirus infections. Pp. 223–231 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Swinton, J., M. E. J. Woolhouse, M. E. Begon, A. P. Dobson, E. Ferroglio, B. T. Grenfell, V. Guberti, R. S. Hails, J. A. P. Heesterbeek, A. Lavazza, M. G. Roberts, P. J. White, and K. Wilson. 2002. Microparasite transmission and persistence. Pp. 83–101 in P. J. Hudson, A. Rizzoli, B. T. Grenfell, H. Heesterbeek, and A. P. Dobson (editors), The ecology of wildlife diseases. Oxford University Press, Oxford, UK. Syverton, J. T., O. R. McCoy, and J. Koomen. 1947. The transmission of the virus of lymphocytic choriomeningitis by Trichinella spiralis. Journal of Experimental Medicine 85:759–769. Tezcan-Merdol, D., M. Ljungström, J. WinieckaKrusnell, E. Linder, L. Engstrand, and M. Rhen. 2004. Uptake and replication of Salmonella enterica in Acanthamoeba rhysodes. Applied and Environmental Microbiology 70:3706–3714. Thomson, G. R., R. G. Bengis, and C. C. Brown. 2001. Picornavirus infections. Pp. 119–130 in E. S. Williams and I. K. Barker, editors. Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Thorne, E. T. 2001. Brucellosis. Pp. 372–395 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Titche, A. R. 1979. Avian cholera in California. California Department of Fish and Game, Wildlife Management Branch. Ul’yanova, N. I., M. A. Bessonova, L. N. Panasik, V. N. Svimonishvili, and L. S. Grishina. 1982. Results of the prolonged study of backwater foci of tularemia and its prevention (based on the materials obtained in the Leningrad oblast, Russian SFSR, USSR) [in Russian]. Zhurnal Microbiologii, Epidemiologii i Immunologii 2:104–107. van Regenmortel, M. H. V., and B. W. J. Mahy. 2004. Emerging issues in virus taxonomy. Emerging Infectious Diseases 10:8–13. Villarreal, L. P. 2004. Are viruses alive? Scientific American 291(6):101–105. Vindevogel, H., and P. P. Pastoret. 1981. Pathogenesis of pigeon herpesvirus infection. Journal of Comparative Pathology 92:415–426. Voyles, B. A. 2002. The biology of viruses. 2nd ed. McGraw-Hill, Boston, MA

Wallace, G. W. 1977. Swine influenza and lungworms. Journal of Infectious Diseases 135:490–492. Webster, R. G., W. J. Bean, O. T. Gorman, T. M. Chambers, and Y. Kawaoka. 1992. Evolution and ecology of influenza A viruses. Microbiological Reviews 56:152–179. Wehr, E. E. 1971. Nematodes. Pp. 185–233 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Wetzler, T. F. 1971. Pseudotuberculosis. Pp. 75–88 in J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (editors), Infectious and parasitic diseases of wild birds. Iowa State University Press, Ames, IA. Whithear, K. 2001. Diseases due to mycoplasmas. Pp. 413–422 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Whittaker, R. H. 1969. New concepts of kingdoms of organisms. Science 163:150–160. Whittington, R. 2001. Chlamydiosis of koalas. Pp. 423–434 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Williams, E. S. 2001. Canine distemper. Pp. 50–59 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Williams, E. S., and I. K. Barker (editors). 2001. Infectious diseases of wild mammals. 3rd ed. Iowa State University Press, Ames, IA. Williams, L. P., and K. W. Newell. 1970. Salmonella excretion in joy-riding pigs. American Journal of Public Health 60:926–929. Winfield, M. D., and E. A. Groisman. 2003. Role of nonhost environments in the lifestyles of Salmonella and Escherichia coli. Applied and Environmental Microbiology 69:3687–3694. Wobeser, G. A. 2006. Essentials of disease in wild animals. Blackwell Publishing Professional, Ames, IA. Worley, M. 2001. Retrovirus infections. Pp. 213–222 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA. Yates, M. V., C. P. Gerba, and L. M. Kelley. 1985. Virus persistence in groundwater. Applied and Environmental Microbiology 49:778–781. Yuill, T. M., and C. Seymour. 2001. Arbovirus infections. Pp. 98–118 in E. S. Williams and I. K. Barker (editors), Infectious diseases of wild mammals. Iowa State University Press, Ames, IA.

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NINE

Eubacteria

CONTENTS Francisella tularensis275 Yersinia pestis278 Borrelia burgdorferi284

Latent Infections (Apparently Healthy Carriers) 259 Salmonella spp.260 Brucella spp.262 Pasteurella spp. and Mannheimia spp. 265 Mycoplasma spp.268

Helminth–Vertebrate Reservoir 288 Neorickettsia helminthoeca289

Bacteria Maintained by Hosts with Clinically Active Infections 270 Mycobacterium spp.270

Soil and Water as Reservoirs 290 Listeria monocytogenes290 Bacillus anthracis291 Clostridium botulinum293 Other organisms 296

Vector-borne Bacteria

Literature Cited

275

life of the infected host. The bacteria shed from this infected host become the source of infection for other susceptible hosts. Typically, the carrier has recovered from a clinical disease (i.e., recovered carrier) but continues to shed the bacteria to other members of its own species or other susceptible species. There also are occasional cases in which parasites live in healthy carrier hosts that typically do not

Latent Infections (Apparently Healthy Carriers) Latent infections (apparently healthy carriers) may be the most common type of reservoir among pathogenic bacteria, and examples are found among a wide variety of bacterial groups. In this reservoir, the bacteria persist within an infected host for extended periods, up to the



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259

experience clinical illness for years at a time, or for their lifetimes (i.e., silent carriers). salmonella spp. causative agent Salmonella spp. are gram-negative bacteria (Family Enterobacteriaceae; App. 1: Table 7) shaped as short rods; they are well-adapted intestinal bacteria of many birds, mammals, reptiles, and amphibians. At least 2,435 serotypes (serovars) of Salmonella spp. have been identified, based on their surface and somatic antigens (Bopp et al. 1999, Mörner 2001b); most of these serotypes show little specificity for their host species (Daoust and Prescott 2007). Until the 1970s, species within the genus Salmonella were defined by their epidemiology, host range, biochemical reactions, and antigenic structure. These former species were reduced to serotypes of two species: S. enterica and S. bongori; S. enterica contains five subspecies and the vast majority of serovars (Murray 1991, Clark and Gyles 1993, Bopp et al. 1999, Farmer 1999, Brenner et al. 2000). Most of the older literature used the serotype name as a species name (e.g., Salmonella typhimurium), and for convenience some scholars still continue to do so (Gast 2003b). host range and distribution Salmonellae have a worldwide distribution (Mörner 2001b), and the occurrence of salmonellae in the intestinal tract of wild animals often is correlated with the proximity of their habitats to those of humans or livestock (Daoust and Prescott 2007). Serovars Enteritidis and Typhimurium are the two most widespread salmonellae (Acha and Szyfres 2001). Among mammals, salmonellae are found among a wide variety of taxonomic groups throughout the world, many of which are able to serve as recovered carriers; examples include wild pigs (Sus scrofa), many deer species, hares, rodents, marsupials, insectivores, primates, and carnivores (Mörner 2001b). The potential avian host range of the genus Salmonella appears unlimited (Daoust and Prescott 2007). Among North American birds, salmonellae are reported regularly among 260 eubacteria

gulls and terns, songbirds, waterfowl, herons and egrets, and pigeons and doves (Friend 1999c); salmonellosis has been a significant contributor to mortality, particularly among the Passeriformes and Piciformes, but also among the Ciconiiformes, Pelecaniformes, and Gaviiformes (Hall and Saito 2008). Salmonellae also commonly are reported from reptiles, fish, and insects (Janssen and Meyers 1968, Murray 1991). While salmonellae infect humans (Bopp et al. 1999), humans rarely acquire salmonellae from wildlife (Murray 1991, Acha and Szyfres 2001). reservoirs and transmission  Although salmonellae can survive for extended periods in the environment, the animal host is believed to be the primary habitat of Salmonella spp. (Winfield and Groisman 2003); salmonellae carry a number of genes that aid in the invasion of and survival in host cells (Scherer and Miller 2001). Many specific serotypes of Salmonella spp. lack special host adaptations and are capable of colonizing a wide variety of hosts (Foltz 1969). Other serotypes are highly host adapted and can cause high mortality in their respective hosts, but also establish long-term carrier states in some of these hosts to ensure their continued transmission (Daoust and Prescott 2007). Asymptomatic carriers commonly serve as the reservoir and can occur in a variety of species. For example, about 3.4% of free-living wild mammals in Panama had detectable levels of Salmonella spp. in a variety of hosts (Kourany et al. 1976), principally by serovar Enteritidis; the highest prevalence (12%) occurred among marsupials, opossums of the genera Philander spp., and Didelphis spp. Among wild birds, 10% of apparently healthy coots (Fulica americana) in the Imperial Valley of California had Salmonella spp. in their feces (Rosen et al. 1957). However, most surveys of Salmonella in wild birds has shown