Infectious Diseases and Pathology of Reptiles. Color Atlas and Text 2006051177

Table of contents :
Contents
Preface
ACKNO WLEDGMENTS
About the Editor
Contributors
Chapter 1. Overview of Reptile Biology, anatomy, and Histology
Chapter 2. Reptile Immunology
Chapter 3. Circulating Inflammatory Cells
Chapter 4. Reptile Necropsy Techniques
Chapter 5. Host Response to Infectious Agents and Identification of Pathogens in Tissue Section
Chapter 6. Identifying Reptile Pathogens Using Electron Microscopy
Chapter 7. Molecular Diagnostics
Chapter 8. Serodiagnostics
Chapter 9. Viruses and Viral Diseases of Reptiles
Chapter 10. Bacterial Diseases of Reptiles
Chapter 11. Mycotic Diseases of Reptiles
Chapter 12. Parasites and Parasitic Diseases of Reptiles
Chapter 13. Isolation of Pathogens
Index
Back cover

Citation preview

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Jogfdujpvt!Ejtfbtft!boe Qbuipmphz!pg!Sfqujmft Dpmps!Bumbt!boe!Ufyu

FMMJPUU!S/!KBDPCTPO University of Florida College of Veterinary Medicine Gainesville, Florida

CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487‑2742 © 2007 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid‑free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number‑10: 0‑8493‑2321‑5 (Hardcover) International Standard Book Number‑13: 978‑0‑8493‑2321‑8 (Hardcover) This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978‑750‑8400. CCC is a not‑for‑profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation with‑ out intent to infringe. Library of Congress Cataloging‑in‑Publication Data Infectious diseases and pathology of reptiles : color atlas and text / edited by Elliott Jacobson. p. cm. Includes bibliographical references and index. ISBN 0‑8493‑2321‑5 (alk. paper) 1. Reptiles‑‑Infections. 2. Reptiles‑‑Infections‑‑Atlases. I. Jacobson, Elliott. SF997.5.R4I54 2007 639.3’9‑‑dc22 Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

2006051177

Contents

Preface............................................................................................................................................ vii Acknowledgments.......................................................................................................................... ix About the Editor............................................................................................................................. xi Contributors.................................................................................................................................. xiii 1

Overview of Reptile Biology, Anatomy, and Histology..................................................................1 Elliott R. Jacobson

2

Reptile Immunology.....................................................................................................................131 Francesco C. Origgi

3

Circulating Inflammatory Cells....................................................................................................167 Nicole I. Strik, A. Rick Alleman, and Kendal E. Harr

4

Reptile Necropsy Techniques......................................................................................................219 Scott P. Terrell and Brian A. Stacy

5

Host Response to Infectious Agents and Identification of Pathogens in Tissue Section..........257 Brian A. Stacy and Allan P. Pessier

6

Identifying Reptile Pathogens Using Electron Microscopy....................................................... 299 Elliott R. Jacobson and Don A. Samuelson

7

Molecular Diagnostics..................................................................................................................351 April J. Johnson, Francesco C. Origgi, and James F.X. Wellehan, Jr.

Infectious Diseases and Pathology of Reptiles  

vi  Contents

8

Serodiagnostics .......................................................................................................................... 381 Elliott R. Jacobson and Francesco C. Origgi

9

Viruses and Viral Diseases of Reptiles....................................................................................... 395 Elliott R. Jacobson

10

Bacterial Diseases of Reptiles......................................................................................................461 Elliott R. Jacobson

11

Mycotic Diseases of Reptiles........................................................................................................527 Jean A. Paré and Elliott R. Jacobson

12

Parasites and Parasitic Diseases of Reptiles................................................................................571 Elliott R. Jacobson

13

Isolation of Pathogens..................................................................................................................667 Francesco C. Origgi and Jean A. Paré

Index .......................................................................................................................................................681

Preface

Why this book and why now? As I approached my 30th year following graduation from veterinary college I decided the time was ripe for a color atlas including some of the best of the tens of thousands of images I had taken documenting infectious diseases and pathology of reptiles. Although descriptive reports of reptile pathology date back to the mid-1800s, and whereas many of the recent texts on reptile medicine and disease cover various aspects of this topic, it was my feeling that a more definitive text with more inclusive images covering infectious diseases and pathology of reptiles was needed. Given that much of my clinical and research career has centered on this topic, I decided to take sabbatical leave in July 2004 to gather as much of the most relevant material that I had collected and published over the past 30 years into a book. As with many books, the time it took to conclude this project was far greater than originally anticipated. Two major hurricane seasons later it is done. Here I have selected a number of topics that are relevant to infectious diseases and pathology of reptiles. Because understanding the biology of reptiles, particularly anatomy and histology, is critical in understanding and interpreting pathology, this book starts with a general review of the biology of the Reptilia in Chapter 1. All major systems are reviewed, and in-depth anatomy and histology are provided. This represents the most complete single source of color images of normal reptile histology. Scientific names are first given as the currently accepted name followed by the former name originally published in the older literature. In each chapter, the scientific name follows the common name the first time the common name is used. Thereafter, only the common name is used. The following served as sources of information for the currently accepted common, scientific, and family names used in this book: EMBL Reptile Database (http://www.embl-heidelberg.de/~uetz/Reptiles.html); Norman Frank and Erica Ramus, 1995, A Complete Guide to Scientific and Common Names of Reptiles and Amphibians, NG Publishing, Pottsville, Pennsylvania; Tim Halliday and Kraig Adler, 2002, Firefly Encyclopedia of Reptiles and Amphibians, Firefly Books, Buffalo, New York; and George R. Zug, Laurie J. Vitt, and Janalee P. Caldwell, 2001, Herpetology: An Introductory Biology of Amphibians and Reptiles, Academic Press, San Francisco, California. Immunology is a special component of reptile biology, and because of its role in the response of reptiles to pathogens, this is reviewed as a separate topic in Chapter 2. Reptiles have a number of circulating inflammatory cells that are critical in their defense against invading pathogens, and while only a few reptiles have been studied in any vii

viii  Preface

detail, there is enough information to be synthesized from the literature to merit having this as a separate topic in Chapter 3. Postmortem evaluation of reptiles is critical in determining causes of mortality, and those working with reptiles have made some modifications to go along with the different body plans of this group. Chapter 4 provides an approach and will be useful to both trainees and seasoned pathologists having limited contact with reptiles. The host response to pathogens is often key in making a diagnosis, and while reptiles as a group show many similarities, differences between groups do exist. Chapter 5 presents the most up-to-date information on this topic. Because many pathogens are not easy to isolate and difficult or impossible to specifically identify using light microscopy, electron microscopy is often used in determining the presence and nature of certain infectious agents. Chapter 6 provides an overview of techniques and methods used in electron microscopy and many electron photomicrographs of reptile pathogens are included. Although isolation of a particular pathogen is still important when trying to identify the cause of a disease, many pathogens are extremely fastidious or impossible to culture, and necessitate the use of molecular approaches. Chapter 7 brings together and reviews this topic. Serodiagnostics have come a long way over the last 15 years with the development of immunological reagents specifically produced against reptile immunoglobulins and their use in such tests as the indirect enzyme-linked immunosorbent assays (ELISA). Chapter 8 reviews those serological assays used in determining the presence of pathogen-specific antibodies in reptiles. Chapters 9, 10, 11, and 12 review viral, bacterial, fungal, and parasitic diseases, respectively, and present what is known about these major groups of pathogens in reptiles. Finally, methods for isolating viruses, bacteria, and fungi are reviewed in Chapter 13. Many books become outdated very quickly, while a few provide worthwhile information for many years to come. Given the expertise of the various contributors, we expect this book to serve as a valuable source of information for future generations.

ACKNOWLEDGMENTS

This book was a team effort and special thanks go to those colleagues who authored and coauthored chapters. Many images in this book have been graciously provided by many friends and I want them to know how much I appreciate their willingness to share this material with me. I am grateful to the University of Florida, which approved my sabbatical leave from July to January 2004. This provided me with the much-needed time for organizing and starting this project. My laboratory technician, April Childress, obtained many of the electronic and hard copies of papers referenced in various chapters. I want to thank Pat Lewis, an outstanding histotechnician. Pat prepared numerous microscopic slides that were used for obtaining photomicrographs in this book. The University of Florida Electron Microscopy Core Laboratory processed tissues for electron microscopy and provided most of the electron photomicrographs presented in this book. Many of the images in this book are of cases submitted to the Zoological Medicine Service, University of Florida Medical Center, for diagnostics and treatment. These cases were supported, in part, by the Batchelor Foundation, University of Florida. Several colleagues graciously reviewed chapters and they are acknowledged at the end of the chapters they reviewed. Thanks go to John Sulzycki, Pat Roberson, and Gail Renard of CRC Press for their editorial and production help. And most important, a very special thanks goes to my wonderful wife Stephanie Pierce for her understanding, encouragement, and patience, Our home was taken over by the many hundreds of scientific papers and books that were used as references in this project. She also had incredible tolerance for the hum of the slide scanner that ran for several hours on countless evenings over the 2½ years I worked on this book. Luckily she is a deep sleeper. She is, and always will be, the most important person in my life.

ix

About the Editor

Elliott Jacobson is a native of Brooklyn, New York, and despite growing up in an inner city, he became fascinated with reptiles at a very early age. Endless days were spent looking at reptiles in collections at the Staten Island Zoo, Bronx Zoo, and American Museum of Natural History. He was extremely fortunate to have parents who encouraged this interest and allowed him to keep various reptiles and amphibians as pets. He majored in biology at Brooklyn College of the City University of New York and earned his B.S. degree in 1967. He eventually went on to earn an M.S. degree at New Mexico State University in 1969 where he worked on physiological ecology of snakes. The two years spent there shaped his professional and personal life in many different ways. He then went to graduate school at the University of Missouri where, for his doctoral research, he worked on glucose regulation in the mudpuppy, Necturus maculosus. Illness and disease in his research animals opened his eyes to a career in veterinary medicine, a career that he hoped would give him a better understanding of disease processes in these animals. He dually enrolled in graduate school and veterinary school and earned his D.V.M. and Ph.D. in zoology in 1975. From 1975 to 1977 he was a faculty member in the Veterinary Science Department at the University of Maryland in College Park, and wildlife veterinarian for the state of Maryland. Elliott Jacobson arrived at the University of Florida in Gainesville in 1977 and is currently a professor of zoological medicine in the Department of Small Animal Clinical Sciences in the College of Veterinary Medicine. He is also a member of the Zoological Medicine Service at the Veterinary Medical Center at the University of Florida where he serves as a clinician and teaches veterinary students and graduate veterinarians in a zoological medicine residency training program. Since 1979, Dr. Jacobson has advised 30 residents and has advised or served on the committees of 18 graduate students. Almost all of his former residents are employed in major zoological institutions and aquariums scattered across the United States. In 1985 he became a diplomate of the American College of Zoological Medicine. He is the recipient of numerous awards including the Fredric L. Frye Lifetime Achievement Award given by the Association of Reptilian and Amphibian Veterinarians (May 2004). Over the last 31 years Dr. Jacobson has been studying health problems of reptiles, both in the wild and in captivity. His laboratory focuses on infectious diseases of reptiles including the development of serologic and molecular assays used to determine exposure to and infection with certain pathogens. xi

xii  About the Editor

He has also studied and published on pharmacokinetics of antimicrobials and parasiticides in reptiles. Former and current graduate students are responsible for developing many of the assays currently employed in his laboratory. He has authored or coauthored 228 refereed scientific papers, 37 chapters in texts, edited and coedited three books, and has been either the principal or coprincipal investigator on 83 funded projects since 1978. Many of his papers represent the first description of certain infectious agents in a reptile. Several of these descriptive reports have evolved into long-term research projects. Reptiles continue to be a challenging group of animals to study with respect to infectious diseases and pathology. There is so much to be done that it is hard to picture a time when everything will be known about disease processes in these animals. Elliott Jacobson is married and has two sons. His household is full of pets including fish, amphibians, dogs, and numerous reptiles. This would not be possible if his wife, Stephanie Pierce, did not equally enjoy this way of life. She is a graduate of the Santa Fe Community College Zoo Program and is a registered nurse. All the author can say is that life has been good. A hobby turned into a career and both will remain closely connected as long as he is alive.

Contributors A. Rick Alleman, D.V.M., Ph.D. Diplomate, American College of Veterinary Pathologists Diplomate, American Board of Veterinary Practitioners Department of Physiological Sciences College of Veterinary Medicine University of Florida Gainesville, Florida [email protected] Kendal E. Harr, D.V.M., M.S. Diplomate, American College of Veterinary Pathologists FVP Consultants, Inc. Gainesville, Florida [email protected]

Francesco C. Origgi, D.V.M., Ph.D. Department of Infectious Diseases and Pathology College of Veterinary Medicine University of Florida Gainesville, Florida [email protected] Jean A. Paré, D.M.V., D.V.Sc. Diplomate, American College of Zoological Medicine Staff Veterinarian Animal Health Centre Toronto Zoo Scarborough, Ontario, Canada [email protected]

Elliott R. Jacobson, D.V.M., M.S., Ph.D. Diplomate, American College of Zoological Medicine Department of Small Animal Clinical Sciences College of Veterinary Medicine University of Florida Gainesville, Florida [email protected]

Allan P. Pessier, D.V.M. Diplomate, American College of Veterinary Pathologists Wildlife Disease Laboratories Conservation and Research for Endangered Species Zoological Society of San Diego San Diego, California [email protected]

April J. Johnson, D.V.M., M.P.H., Ph.D. Department of Small Animal Clinical Sciences College of Veterinary Medicine University of Florida Gainesville, Florida [email protected]

Don A. Samuelson, Ph.D. Department of Small Animal Clinical Sciences College of Veterinary Medicine University of Florida Gainesville, Florida [email protected]

Contributors  xiii

xiv  Contributors

Brian A. Stacy, D.V.M. Diplomate, American College of Veterinary Pathologists Department of Small Animal Clinical Sciences College of Veterinary Medicine University of Florida Gainesville, Florida [email protected] Nicole I. Strik, Dr.med.vet Clinical Pathology Service College of Veterinary Medicine University of Florida Gainesville, Florida [email protected]

Scott P. Terrell, D.V.M. Diplomate, American College of Veterinary Pathologists Veterinary Services, Disney’s Animal Kingdom Bay Lake, Florida [email protected] James F. X. Wellehan, Jr., D.V.M., M.S. Diplomate, American College of Zoological Medicine Diplomate, American College of Veterinary Microbiologists Department of Small Animal Clinical Sciences College of Veterinary Medicine University of Florida Gainesville, Florida [email protected]

1 Overview of Reptile Biology, anatomy, and Histology Elliott R. Jacobson

1.1 General Concepts

Contents 1.1 General Concepts...................................................... 1 1.2 Extant Taxonomic Orders of the Reptilia................. 2 1.2.1 Chelonia......................................................... 2 1.2.2 Crocodylia...................................................... 2 1.2.3 Rhynchocephalia........................................... 3 1.2.4 Squamata....................................................... 3 1.3 Thermal Biology........................................................ 4 1.4 Review by System and Organs................................. 4 1.4.1 Integumentary System................................... 4 1.4.2 Musculoskeletal System................................. 6 1.4.3 Digestive System............................................ 8 1.4.4 Respiratory System...................................... 12 1.4.5 Urinary System............................................ 13 1.4.6 Reproductive System................................... 14 1.4.7 Cardiovascular System................................. 18 1.4.8 Hemopoietic System.................................... 19 1.4.9 Endocrine Organs....................................... 19 1.4.10 Nervous System........................................... 20 1.4.11 Eye................................................................ 21 1.4.12 Ear................................................................ 23 1.4.13 Vomeronasal Organ..................................... 24 1.4.14 Salt Glands................................................... 24 1.4.15 Infrared Detection Organs.......................... 24 Acknowledgments.............................................................. 24 References........................................................................... 25

The class Reptilia consists of the following four extant orders: Chelonia (tortoises and turtles), Crocodylia (alligators, caimans, crocodiles, gharial), Rhynchocephalia (tuataras), and Squamata (lizards, worm lizards, and snakes). During the Mesozoic, often called the Age of Reptiles, there were 17 orders. Thus, current day reptiles are far less diverse and represent a smaller number of species than those present during that time. The exact phylogenetic relationship among chelonians, crocodilians, and squamata remains unclear. Particularly, the relationship between chelonians and the reptile clade Diapsida (crocodilians, lizards, and snakes) remains unsettled. For example, molecular studies suggest a relationship between chelonians and crocodilians (Zardoya and Meyer, 1998; Hedges and Poling, 1999), whereas analyses of osteological data indicated that turtles are nested within the diapsids as a sister group of the Sauopterygia, a group of Mesozoic reptiles (Rieppel and Reisz, 1999). The development of the shelled (cleidoic or amniotic) eggs places reptiles at the crossroads of vertebrate evolution. Reptiles evolved from amphibians, and based on certain shared morphologic features, Anthracosaurs are probably the ancestors of the early reptiles. The first reptiles appeared during the Paleozoic era, approximately 300 million years ago. The fundamental difference between primitive reptiles and their amphibian ancestor(s) is believed to be in their reproductive strategies: amphibians produce anamniotic eggs, whereas reptiles produce amniotic eggs. The amphibian anamniotic egg is gelatinous, lacks extraembryonic membranes, and is thus very susceptible to desiccation. For the most part, amphibians have to lay their eggs in a wet or very moist environment. The following extraembryonic membranes characterize the amniotic egg: chorion, allantois, amnion, and yolk sac. This

Infectious Diseases and Pathology of Reptiles  

  Overview of Reptile Biology, Anatomy, and Histology

egg probably developed in a series of steps over time (Packard and Seymour, 1997). The reptile egg is far more resistant to desiccation compared to the amphibian egg, and the large amount of internal yolk provides an energy source and supplies maternal immunoglobulins that may last up to a year following hatching (Schumacher et al., 1999). It is the amniotic egg, in addition to certain morphologic and physiologic modifications such as the development of scales, evolution of the metanephric kidney, and musculoskeletal modifications for terrestrial locomotion, that unite reptiles and separate them from the Amphibia. Ecothermy, a body covered by scales, and a lack of feathers and hair separate reptiles from birds and mammals, respectively. Birds and mammals evolved from certain reptiles as separate lines of evolution. These two classes are more closely related to reptiles than they are to each other.

1.2 Extant Taxonomic Orders of the Reptilia There are approximately 7500 species of present-day reptiles (Zug et al., 2001), which are categorized into four orders. Each order is briefly reviewed below. Scientific names are first given in this chapter and throughout the book as the currently accepted name, followed by the former name originally published in the older literature. The following served as sources of information for currently accepted common, scientific, and family names used in this book: EMBL Reptile Database (http://www.reptileweb.org), Frank and Ramus (1995), Halliday and Adler (2002), and Zug et al. (2001).

1.2.1 Chelonia The Chelonia (Testudines) includes the turtles and tortoises (13 families, more than 285 species). Phylogenetically, this is the oldest group of reptiles. While all chelonians are united by the presence of a bony shell, they do show diversity. This very conservative group is subdivided into two suborders. The suborder Pleurodira includes side-neck and snake-neck turtles, which have anatomical structures that allow them to fold their head and neck across the front of the shell and under the overhanging carapace (Figure 1.1). Two of the 13 families of chelonians are within this suborder. In contrast, the Cryptodira are more advanced and have evolved structures that allow them to withdraw their cervical vertebrae rostrocaudally within the margin of the shell (Figure 1.2). Eleven of 13 families of chelonians have this capability. Chelonians are the only tetrapods with the pectoral girdles internal to the ribs (Figure 1.3); in all other tetrapods it is external to their ribs. Except for cervical and caudal vertebrae, all others (along with ribs) are fused with the dermal bone and fibrous connective tissue of the carapace. A tympanic membrane is located caudal and ventral to the eye (Figure 1.4). All chelonians lay eggs.

The majority of chelonians are aquatic or semiaquatic. Two families and five genera of marine-adapted turtles are found primarily in tropical and subtropical regions of the world. The marine turtle family Dermochelyidae includes one species, the leatherback (Dermochelys coriacea). The other family of sea turtles, Cheloniidae, includes the green (Chelonia mydas), loggerhead (Caretta caretta), hawksbill (Eretmochelys imbricata), Atlantic or Kemp’s ridley (Lepidochelys kempii), and olive ridley (Lepidochelys olivacea) sea turtle. Tortoises (approximately 50 species) are all within the family Testudinidae and as a group have adapted anatomic structures and physiologic mechanisms for surviving in arid and semiarid environments. For instance, in proportion to their size, tortoises have the largest urinary bladder of all chelonians. In tortoises the urinary bladder serves as a storage site for water and a site where certain ions such as potassium are concentrated during periods of drought. In contrast, the urinary bladder of sea turtles is much smaller and the wall is thicker. Most of the common freshwater turtles are in the family Emydidae.

1.2.2 Crocodylia The Crocodylia (23 species) consist of the following 3 families: Alligatoridae (American alligator [Alligator mississippiensis], Chinese alligator [Alligator sinensis], and caimans (Caiman spp. Melanosuchus niger, Paleonsuchus spp.)], Crocodylidae (crocodiles [Crocodylus spp. Osteolaemus tetraspis, Tomistoma schlegelii]), and Gavialidae (gharial [Gavialis gangeticus]). This is also considered a conservative group because their overall morphology has remained the same for millions of years. Crocodilians are part of an evolutionary lineage that gave rise to the dinosaurs, and that line also gave rise to the class Aves. Crocodilians have several anatomic firsts for the reptiles. Crocodilians have armored integument (primarily dorsally) that contain osteoderms. They are the first vertebrates to have a four-chambered heart and the first to have a complete hard palate. The glottis is located at the angle of the jaw, directly posterior to overlapping dorsal and ventral folds (Figure 1.5). Together, these folds are called the gular valve and anatomically separate the oral and pharyngeal cavities (Putterill and Soley, 2006). For further details, see Section 1.4.3. A complete hard palate and a posteriorly located glottis are adaptations to feeding in water. A well-developed nictitans (Figures 1.6–1.7) covers and protects the entire globe when submerged and feeding in water. A tympanic membrane is located in a depression behind the eyes and is protected by specialized integumentary structures (Figure 1.8). A pseudodiaphragm separates the coelomic cavity into a cranial space where the heart, liver, and lungs are located, and a caudal space where all the remaining viscera are located (Figure 1.9). A unique fat body is located directly caudal to the pseudodiaphragm, in the right quadrant of the caudal coelomic cavity (Figures 1.10, 4.9). When healthy, this structure is relatively large and distinct. In times of illness or

Overview of Reptile Biology, Anatomy, and Histology  

when anorexic, this fat body will atrophy. Of all the crocodilians, only the gharial has external characteristics that can be used for distinguishing sexes. In male gharials, a large bulbous structure develops at the rostral end of the upper jaw when they are approximately 13 years old. All crocodilians lay eggs.

1.2.3 Rhynchocephalia The remaining two orders (Rhynchocephalia and Squamata) of reptiles are somewhat closely related and are placed within a single subclass, Lepidosauria. The Rhynchocephalia consists of the following two species of tuataras, which are monotypic at the generic level: tuatara (Sphenodon punctatus) and Brother Island tuatara (S. guntheri). While historically they were quite diverse, being found around the world, modern tuataras are confined to a few dozen islands off the coast of New Zealand. Tuataras are extremely long lived, with records around 100 years of age. They are somewhat unusual in that they are adapted to a cool environment with an ideal ambient temperature range of 12°C to 16°C. Tuataras have a very well developed parietal eye and pineal gland (Ung and Molteno, 2004), and although they have internal fertilization, males lack a copulatory organ. While superficially resembling lizards, sharing some morphologic characteristics such as fracture planes in the tail vertebrae, they lack the middle ear cavity and tympanic membrane (Figure 1.11). Tuataras also have gastralia (abdominal ribs).

1.2.4 Squamata The fourth, and most diversified order of reptiles, is the Squamata (approximately 7200 species) including Sauria (Lacertilia; lizards: 4300 species), Amphisbaenia (worm lizards: 140 species), and Ophidia (Serpentes; snakes: 2900 species). While some lizards range into temperate to cool areas of the world, the greatest diversity is in tropical and warm desert regions. With 16 families, including those without limbs, lizards have the widest range in morphology of all reptiles. While many species have keratinized skin thrown into folded distinctive scales that cover their body, some species have less distinctive scalation. Lizards have a well-developed tympanic membrane that covers the middle ear canal (Figure 1.12). Some lizards, such as certain geckoes, have evolved modified eyelids in the form of a spectacle that covers and protects the cornea (see Section 1.4.11.1 in this chapter). Although the spectacle is heavily vascularized, this is not appreciated in normal healthy lizards. While the majority of lizards are tetrapods, some groups show ranges in limb reduction (some skinks) and others have completely lost their limbs (glass lizards, legless lizards). One family of lizards, Helodermatidae, has members (beaded lizard [Heloderma horridum] and gila monster [Heloderma suspectum]) that evolved venom glands associated with their lower jaws. In a recent report (Fry et al., 2006), venom toxins were reported to occur in Varanidae

(monitors) and the agamid, Pogona barbata. The majority of lizards are oviparous. However, some have developed placentation and are viviparous. The suborder Amphisbaenia consists of the worm lizards. These exclusively subterranean reptiles are found in subtropical to tropical regions of the Americas, Africa, and western Asia. Three of the four families of worm lizards totally lack limbs. One family, Bipedidae from Baja and southwest coastal Mexico, has enlarged forelimbs that are used in burrowing (Figure 1.13). Other adaptations for digging include a blunt head with compact, hard cranial bones, large scales on the anterior portion of the head, fused eyelid skin with eyes beneath, and no external ear openings. The scales covering the body are arranged in rings that give these animals a segmented or annulated appearance. The Florida worm lizard (Rhineura floridana) is easily confused with a worm when dug up during gardening (Figure 1.14). The suborder Ophidia (Serpentes) is the most recent of the present day reptiles. Snakes are distributed around the world with the greatest diversity in deserts and tropical regions. Depending upon the classification scheme used, the number of families can vary from 14 to 17. For instance, while some classifications have boas and pythons in the family Boidae, others have assigned members to two distinct families: Boidae and Pythonidae. For purposes of this book, the latter categories are used for these snakes. Snakes probably evolved from a lizard group that became subterranean and lost their eyes. The eye of snakes is embryologically distinct from that of all other reptiles (Underwood, 1970; Walls, 1942). For details see Section 1.4.11.4 of this chapter. Snakes totally lack all limbs, lack tympanic membranes (Figure 1.15), and lack middle ear cavities. The tongue is elongated, forked, and serves as a mechanical structure to collect particles in the air that are delivered to the vomeronasal organ (see Section 1.4.13 in this chapter) in the roof of the mouth. The vomeronasal organ, also called Jacobson’s organ, is a chemoreceptor. The total body plan of snakes is elongate, with vertebrae numbering several hundred in most species. The body is covered by scales, with several rows of smaller scales laterally and dorsally and a single row of large scales ventrally. Compared to other reptiles, all major organs in snakes are more linearly arranged. All snakes lack a urinary bladder, with urine formed in the kidney ultimately being transported from the ureter to the cloaca, and then retrograde into the colon. Water in the colon is reabsorbed and the urates condense into a semisolid mass that is eliminated when snakes defecate. While most snakes are oviparous, many have developed primitive placentation and are viviparous. Venom glands and a venom injection apparatus involving front fangs have evolved in two families (Viperidae [vipers and pit vipers] and Elapidae [cobras, kraits, mambas, coral snakes, sea snakes) of snakes. The family Colubridae has some members having caudally located fangs and venom proteins.

  Overview of Reptile Biology, Anatomy, and Histology

1.3 Thermal Biology In addition to anatomic and physiologic differences, it is the source of thermal energy that can be used to distinguish reptiles (ectotherms) from mammals and birds (endotherms). The studies of Cowles and Bogert (1944) demonstrated that behavioral thermoregulation is an important aspect of the thermal biology of many reptile species. Whereas birds and mammals (within limits) can control their body temperature within a fairly narrow zone by shifts in metabolic rates, reptiles are dependent upon external sources for regulation of body temperature. All reptiles have a preferred optimum temperature zone (POTZ) that is fairly characteristic of the species, being regulated by behavioral and physiologic mechanisms. The limits of this zone, particularly with temperate species, may fluctuate with seasons of the year. Many physiologic functions appear to have evolved in unison with the thermal biology of reptiles. The temperature zone below the POTZ has been termed the critical thermal minimum (CTMin) and is defined as the temperature that causes a cold narcosis and effectually prevents locomotion. The temperature zone above the POTZ is the critical thermal maximum (CTMax) and may be visualized as a value that is the arithmetic mean of the collective thermal points at which locomotive activity becomes disorganized and the animal loses its ability to escape from conditions that will promptly lead to its death (Lowe and Vance, 1955).

1.4 Review by System and Organs 1.4.1 Integumentary System The integument plays an important role in the conservation of body fluids by forming a protective barrier between deeper tissues and the dehydrating environment. It also functions to protect the animal from invading pathogens. The integument consists of an outer epidermis and underlying dermis. The epidermis, including the stratum corneum, is much thicker in reptiles compared to amphibians. Scales represent a folding of the epidermis, and for the most part cover most of the reptile integument. The greatest folding is seen in snakes where adjacent scales overlap and are joined by a flexible hinge region (Figure 1.16). The integument of reptiles is covered by either α- or β-keratin. In chelonians and crocodilians, α- and β-keratins in epidermal scales alternate horizontally, while in squamates the keratins in the outer portion of scales alternate vertically, with β-keratin overlying α-keratin (Maderson, 1985). In most chelonians the shell is covered by β-keratin. However in soft-shelled turtles (Apalone spp.) and leatherback sea turtles (Dermochelys coriacea), α-keratin covers the carapace and plastron. The shell, consisting of a dorsal carapace and ventral plastron, is a unique structure that distinguishes chelonians

from other reptile groups. Specialized epidermal hard parts called scutes cover the surface of the shell of most chelonians (Figure 1.17). The major epidermal component of each scute is a multilayered β-keratin that covers a continuous layer of pseudostratified columnar epithelial cells (Figures 1.18– 1.19). These cells have a basal surface with thin processes that extend into and interdigitate with the underlying connective tissue (Figure 1.20). When chelonians hatch from the egg, they are born with scutes called embryonic shields (Figure 1.21). As chelonians grow, new keratin in the shell is formed in seams, areas of the shell where two scutes come together. As rings of new keratin are formed around embryonic shields, formerly adjacent embryonic shields become separated (Figure 1.22). At the seams the epidermis invaginates into the dermis (Figure 1.23). Differentiation of basal cells into keratin-forming cells occurs in the deepest portion of the invagination. Scutes overlay a dermis that is unique in that much of it is ossified (Figure 1.24). The outer dermis of the chelonian shell consists of collagen fibers, melanophores, vessels, and nerves. Underneath the dermal connective tissue is dermal bone (Figure 1.25). Outer and inner layers of the dermal bone plates are compact and “sandwich” a middle layer of trabecular or spongy bone (Wronski et al., 1992). In young growing desert tortoises, osteoid surfaces were devoid of adjacent osteoblasts, the cells that normally deposit the unmineralized bone matrix. This is in contrast to mammals in which nearly all osteoid surfaces are lined by osteoblasts. Ribs and vertebrae (except cervical and caudal) are embedded in the dermal bone. In reptiles, epidermal growth and replacement of the outer old with a new inner epidermis, is either continuous (turtles and crocodilians) or discontinuous (lepidosaurs: tuataras, lizards, and snakes). In lepidosaurs there is a distinct cycle of ecdysis, with periodic formation of a new inner epidermal generation and loss (shedding) of the old outer epidermal generation. While snakes and some lizards may shed an entire old outer generation at once, some lizards (varanids, helodermatids) will lose portions of the outer generation over a one- to two-week period (Figure 1.26). This is a cyclic process that is synchronized to occur over a discrete period of time across the entire body surface. A cycle consists of a resting stage (consisting of 3 subdivisions) followed by 5 stages of renewal (Landmann, 1986; Maderson, 1965; Maderson et al., 1970a, 1970b, 1998). During the resting stage the skin is bright in coloration (Figure 1.27). As a snake begins epidermal renewal, the skin and spectacle covering the cornea become dull and develop a bluish tinge (Figure 1.28). This reaches its deepest blue color in 3 to 7 days (Figure 1.29). Approximately 3 to 7 days later, the snake’s color rapidly brightens and the spectacle of the eye completely clears (Figure 1.30). In another 3 to 7 days, shedding occurs (Figures 1.31–1.32). This is followed by the next resting stage. The stages of ecdysis can be distinguished at a light microscopic level and have been diagrammatically summarized by Landmann (1979; 1986) (Figure 1.33). An entire cycle con-

Overview of Reptile Biology, Anatomy, and Histology  

sists of a resting phase (stage 1) followed by five stages of epidermal renewal. Stage 1 begins following shedding with the immediate post-shedding period, during which most of the α-layer is formed. This is followed by a perfect resting condition, at which time there is little cellular differentiation and proliferation. The final part of stage 1 represents a completion of the outer generation, with the formation of the lacunar tissue and clear cells. At the end of stage 1, the outer epidermal generation is complete and the following layers are present in the outer portion of a scale in order of the most outer to inner epidermal layers: oberhautchen, β-, mesos-, and α-layers (Figure 1.34). The lepidosaurian integument is unique because cells in the outer portion of a scale contain an outer layer of β-keratins and an inner layer of α-keratins. The inner portion of a scale and the hinge region differ from the outer portion in that only α-keratin is present (Figure 1.35). Beta-keratin determines shape and provides stiffness to snake skin while α-keratin allows flexibility and distensibility. When a snake feeds on a large prey item, the distension of the integument and separation of adjacent scales are achieved because of the nature of the properties of α-keratin within this region. At this time the hinge region is exposed. The mesos layer, which is an important barrier in the prevention of water loss (Landmann, 1979), is thicker in the inner portion of a scale compared to the outer (Figure 1.35). In California king snakes (Lampropeltis getula), this layer was found to become thicker and to have increased deposition of lamellar lipids following the first neonatal shed (often 24 to 36 h following birth or hatching from an egg). This accounted for the twofold increase in skin resistance to transepidermal water loss (TEWL), which was seen following the first shed (Tu et al., 2002). Skin resistance to TEWL continued through the second shedding cycle. The oberhautchen is the most outer portion of the β-layer and is characterized by the presence of serrations, surface ornaments, and pits. This is better seen using scanning electron microscopy then light microscopy (Irish et al., 1988) (Figures 1.36–1.37). Although the lacunar tissue and clear layer may or may not be distinguishable at a light microscopic level at the end of stage 1, they are distinguishable using electron microscopy (Landmann, 1979). During the subsequent renewal phase, a new inner generation emerges, with the stratum germinativum differentiating into inner oberhautchen, β-, mesos, and part of the α-layers (Figures 1.38–1.40). In snakes, starting in late stage 3, heterophils can be seen migrating through the epidermis (Paul Maderson, personal communication) (Figure 1.39). Shedding takes place when the oberhautchen of the inner generation separates from the clear layer of the outer generation. While a final synchronization of cellular events occurs at the time of shedding, during the renewal phase prior to shedding, epidermal cytology at any one time varies among different regions of a scale. Thus, regions of each scale may be at different stages of a renewal cycle. A “zip fastener” model (Maderson, 1966; Maderson et al., 1998) explains how the outer generation is held in place while the inner generation is forming. This model involves

interdigitation between the cell membranes of the clear layer of the outer generation and the oberhautchen of the inner generation. This complex is another unique feature of the lepidosaurian integument (Maderson et al., 1998). There is a heightened loss of intracellular fluids at the time of shedding that softens the outer generation and allows the shedding complex to “unzip” (Maderson et al., 1998). Once shedding takes place, the inner generation becomes the new outer generation and the resting stage commences once again. From research that has been performed on determining mechanisms for controlling squamate shedding, it appears this is under the influence of the pituitary–thyroid axis (Maderson, 1985). In snakes, hypophysectomy or thyroidectomy will result in repeated sequential renewal phases (Chiu and Lynn, 1972; Chiu et al., 1983). While thyroid hormones have an inhibitory effect on shedding in snakes, they have a stimulatory effect in lizards (Maderson, 1985). In a healthy snake, the resting phase can last from a few weeks to months. Age, frequency and amount of food consumption, and temperature influence shedding frequency. The author has observed certain skin diseases associated with increased frequency of shedding. A Florida king snake (Lampropeltis getula) with a fungal skin disease entered a new cycle of renewal that overlapped with shedding. Tuataras also produce and lose epidermal generations. However, Alibardi and Maderson (2003) found the following six differences, through histochemistry and transmission electron microscopy, which distinguished skin shedding in tuataras from squamates: (1) absence of a well-defined shedding complex; (2) persistence of plasma membranes throughout the β-layer; (3) presence of lipogenic lamellar bodies and PASpositive (periodic acid-Schiff stain) mucus granules in α-keratinizing cells; (4) presence of contents of these organelles in intercellular spaces of tissues homologous to the squamate mesos, α-, and lacunar cells; (5) few lamellated lipid deposits in the domains of these tissues; (6) presence of keratohyalin-like granules in the presumptive lacunar, clear, and oberhautchen cells. Compared to amphibians, reptiles have relatively few glands associated with their integument. Overall, the reptile integument is dry. Musk glands are associated with the cloaca of many species, mental or chin glands are present in different chelonians (with greater development in males compared to females) (Figures 1.41–1.42), musk glands are medial to the dentary bone of crocodilians, and specialized glands are associated with the angle of the jaw of chameleons. Femoral pores are present and better developed in males of certain lizards such as iguanids (Figures 1.43–1.44) (Jacobson, 2003). They are the openings to a type of holocrine gland, with the waxy secretion seen protruding from the pore (Figure 1.45) and originating from entire cells that are derived from those that line the base of the gland (Figure 1.46). This material may aid males in grasping females during copulation. Some lizards, such as geckos, have precloacal glands arranged in a

  Overview of Reptile Biology, Anatomy, and Histology

chevron of pores (Figures 1.47–1.48). However, many lizards lack femoral pores or precloacal pores. A basement membrane separates the epidermis from the underlying dermis. Within the dermis is a complex pigment system consisting of melanophores containing melanin, erythrophores and xanthophores containing pteridines and carotenoids, and iridophores containing reflecting platelets of guanine, adenine, hypoxanthine, and uric acid (Alexander and Fahrenback, 2005; Bagnara and Hadley, 1973). The arrangement of these cells, both vertical and horizontally, will determine the animal’s coloration and skin pattern (Figures 1.49–1.52). Iridophores, with their reflecting platelets of purines, can be identified using polarizing light microscopy (Figure 1.53).

1.4.2 Musculoskeletal System The skeletal system of reptiles shows many advances over amphibians for feeding on more diverse food items and ambulating in a dry environment. Those aquatic reptiles have further modifications for swimming and feeding in water. The great variety of reptiles is reflected by a very diverse array of musculoskeletal patterns. Major divergences are found among the four orders and differences also are seen within orders. For details on the skeletal system of reptiles see Romer (1956). As in other vertebrates, the skull of reptiles is composed of the chondrocranium, splanchnocranium, and the dermatocranium. Overall, it is more ossified, longer, narrower, and higher than that of amphibians. The dermatocranium is the major component of the reptile skull and includes the nasals, prefrontals, frontals, and parietals. The premaxillae, maxillae, and mandibles also derived from the dermatocranium. The mandible consists of the following bones: dentary, splenial, angular, surangular, coronoid, prearticular, and articular. The articular is the only cartilage replacement bone in the mandible. All teeth on the lower jaw are confined to the dentary. The quadrate (part of the cranium and contains the articular surface for the lower jaw) and stapes (middle ear bone) is derived from the splanchnocranium. Within each clade, these basic components have differentially evolved and exhibit great diversity around several major patterns. A good example is the temporal region of the skull. The skulls of chelonians lack openings in the temporal region, a condition that is classified as anapsid (Figures 1.54–1.55). Embryonically, crocodilians, tuataras, lizards, worm lizards, and snakes have two openings that are separated by the postorbital and squamosal bones. In adult reptiles, this condition can be best seen in tuataras and crocodilians. Tuatara skulls have arches forming the lower boundaries of the two openings and the quadrate is fixed in position (Figures 1.56–1.57). In other reptiles there is a wide range in modification of the skull, especially in this region. In crocodilians (Figures 1.58– 1.59) both temporal openings are generally present, with the upper opening small or closed. In contrast, worm lizards have evolved a skull adapted to burrowing, with many of

the bones fused (Figure 1.60). The shape of the skull of lizards varies from those that are high and domed (for example the green iguana [Iguana iguana]), to those that are laterally flattened, to those that are dorsoventrally flattened and elongate (Figures 1.61–1.67). The persistence of the temporal arches varies in squamates, with the upper arch present in most lizards and the ventral border of the lower opening absent. In some lizards and snakes, both arches are absent (Figures 1.66–1.74). When present, the temporal openings are sites of muscle attachment, and along with various hinges between different regions of the skull, are involved in cranial kinesis. A loose attachment between the quadrate and skull of snakes allows rotation of the quadrate in different directions (streptostyle). This accounts for snakes having the ability to swallow large prey items. Chelonians, while lacking teeth, have specialized keratinized hard parts (rhamphotheca) that cover the maxillae and mandibles (Figure 1.75). Most of the other reptile taxa have teeth. Different types of dentition have evolved between and within the other major groups of reptiles. Crocodilians are the only reptile group having thecodont dentition, an attachment where teeth are set in sockets (Figure 1.76). The dentition of tuataras is acrodont, having teeth directly attached to the biting surfaces of the jaws (Figure 1.77). In lizards, teeth are either acrodont (Figures 1.63, 1.65, 1.78) or pleurodont (Figures 1.61, 1.66, 1.79–1.81). In pleurodont dentition, the teeth sit on a ledge on the lingual side of the jaws. Traditionally snake teeth have been characterized as pleurodont, but a more current and accurate description is that each snake tooth is actually ankylosed to the rim of a low socket — a type of modified thecodont dentition (Lee, 1997; 1998) (Figures 1.68–1.74). Crocodilians and lizards have two rows of teeth associated with the dentary bone of the mandibles and maxillae. Snakes have two additional upper rows of teeth that are attached to the palatine and pterygoid bones (Figures 1.69–1.71). Compared to nonvenomous snakes, in frontfanged snakes the maxillary bone is reduced in size and only supports the fangs (Figures 1.72, 1.74). While elapid snakes (cobras, kraits, mambas) do not have the ability to erect their fangs to the extent that viperids (vipers and pit vipers) can, some elapids are capable of maxillary and palatine erection while feeding (Duefel and Cundall, 2003). A secondary palate is partially developed in chelonians (Figure 1.82) and is complete in crocodilians (Figure 1.83). The secondary palate evolved as an adaptation for feeding in water. All reptiles have a single occipital condyle that articulates with the first cervical vertebra (atlas), while amphibians have two. Cranial muscle groups are primarily involved with jaw movements and tongue movements. A well-developed hyoid apparatus, which shows anatomic variation among the various reptile groups, supports the tongue and glottis. In those lizards with dewlaps, such as many iguanid lizards (Figure 1.84), ceratobranchial II of the hyoid apparatus is paired and extends into the anterior margins of the dewlap (Figure 1.85).

Overview of Reptile Biology, Anatomy, and Histology  

In reptiles, the axial skeleton (consisting of vertebrae) shows greater regional differentiation than in amphibians. Divisions of the axial skeleton can be made based on presence or absence of ribs, and when present, the type of associated rib. Major divisions are cervical, trunk, sacral, and caudal vertebrae. Each vertebra consists of a body or centrum (formed around the embryologic notochord), and a separately ossified neural arch (that surrounds the spinal cord) along with its dorsal neural spine. A ventral wedge-shaped bone, the intercentrum, which is present between the vertebrae of early reptiles, is present in between the first two cervical vertebrae and caudal vertebrae of all reptiles; it is between the trunk vertebrae of only geckos and the tuatara. Various processes project from the vertebrae and are termed apophyses. Those resulting in articulation of adjacent neural arches are the zygapophyses. Adjacent centra typically articulate with a ball-and-socket joint or fibrous joints between amphiplatyan vertebrae. Where the anterior portion of a centrum forms the socket and the posterior portion forms the convexity (ball), the condition is termed procoelous. Most extant reptiles have procoelous vertebrae. If the anterior portion of a centrum forms the convexity and the posterior portion forms the socket, the condition is termed opisthocoelous (many chelonians). Those with sockets at both ends, such as the tuatara and gekkonid lizards, are amphicoelous. One proximal caudal vertebra of crocodilians is biconvex; the subsequent vertebrae are procoelous or amphiplatyan. Intervertebral discs (derived from the embryonic notochord), which are present between successive surfaces of vertebral centra of mammals, are absent in reptiles. Instead, the intervertebral bodies forming the convexities on either the anterior or posterior surface of the centra serve to prevent excessive forces being applied to the spinal cord during flexion of the vertebrae. The first two vertebrae, the atlas and axis, are modified compared to other cervical vertebrae. Chelonians have 8 distinct cervical vertebrae, crocodilians have 9, and the tuatara and the majority of lizards have 8 (Romer, 1956). Varanid lizards have 9 and chameleons have 3 to 5. Trunk vertebrae are not separated into distinct thoracic and lumbar vertebrae; all trunk vertebrae have ribs. Almost all chelonians have 10 trunk vertebrae that, along with their ribs, are fixed in place. The neural processes and ribs are fused with the overlying dermal plates of the shell (Figure 1.86A, B). In the leatherback sea turtle, the ribs and vertebrae are separate from the carapace. Because the ribs of most chelonians are fixed in place, they cannot be used in moving air into and out of the lungs. Instead, movements of the limbs are involved in the mechanics of respiration. In some reptiles such as crocodilians, “freefloating” abdominal ribs also are present. In snakes, the ribs have a semicircular shape and are attached to the connective tissue that is continuous with the enlarged ventral scales. Reptiles have two to four sacral vertebrae. Fracture planes have evolved in the centrum and part of the neural arch of the caudal vertebrae of many lizards. This adaptation to pre-

dation allows the lizard to lose a portion or most of the tail. This process is called autotomy. In glass lizards (Ophisaurus spp.), almost half the length of the animal is tail. Tail loss is metabolically costly to lizards since it represents a loss of tissue that will be replaced when the tail is regenerated. The regenerated tail has a different morphology to its scalation and bone is replaced with cartilage. Limbs of reptiles vary widely and show modifications to meet the demands of their ecology and ability to ambulate within that ecologic setting. Limbs in some reptiles, such as tortoises, have been modified for digging burrows and for protection from predators. In sea turtles the forelimbs have been modified into flippers for swimming, and in snakes and certain lizards, limbs have been completely lost; other mechanisms for locomotion have evolved. Some worm lizards such as Bipes (Figure 1.13) have only forelimbs. Those reptiles with limbs generally have five digits on the fore- and hindlimbs. A patella is present in some lizards. The pectoral girdle is well developed in all reptiles except snakes where it is totally absent. Chelonians are unique, having lost the sternum and have a pectoral girdle that is internal to the ribs (Figures 1.3, 1.86A). The chelonian pectoral girdle consists of the scapula, acromion (an elongated process of the scapula), and coracoid. In aquatic turtles such as sea turtles, the procoracoid is more elongate than in other chelonians (Figure 1.87). This translates into a greater surface area, allowing muscle attachment needed for sustained swimming. The pelvic girdle consists of three bones: the ischium, ilium, and pubis (Figure 1.86A, B). It is well developed in all reptiles except snakes, where a vestigial girdle is seen in members of the families Boidae and Pythonidae. Pelvic spurs are femoral remnants (Figure 1.88) and in some boids they are noticeably larger in males than females. Bone is a living, changing tissue consisting of a matrix of hydroxyapatites and collagen. There is no single pattern of bone structure for all groups of reptiles, with much variation existing among and within groups. Based on structure, Enlow (1969) categorized reptile bone as the following: 1. Haversian bone. The Haversian system or osteone, the structural unit of compact bone of mammals, is uncommon in reptiles. It is absent in lizards and snakes and in chelonians and crocodilians is limited to certain regions of the cortex. Haversian systems are more common in cortical bone of dinosaurs and some therapsids. 2. Primary vascular bone and nonvascular bone. While some reptiles lack vascular channels in their cortical bone, most reptiles have vascular channels. Presence and absence of channels can vary from one region of cortical bone to another. 4. Lamellar and nonlamellar bone. The calcified matrix of cortical bone typically has a lamellar appearance. This is generally seen in slow growing bone. Nonlamellar (woven) bone also occurs and is particu-

  Overview of Reptile Biology, Anatomy, and Histology









larly prominent in fast-growing skeletal structures. In lamellar bone, osteocytes are arranged in a regular fashion while in nonlamellar bone they are randomly arranged. 6. Lamina bone. This refers to the addition of a compacta composed of a circumferential series of regularly arranged strata (lamina) superimposed over the layering that results from remodeling. 7. Plexiform bone. This type of bone is common in certain prehistoric reptiles, but is uncommon in presentday reptiles. It has been seen in young crocodilians. It is characterized by a regularly arranged threedimensional plexus of primary vascular channels. 8. Compacted coarse-cancellous bone. This type of bone is formed on the endosteal surface of long bones as cancellous bone, and is incorporated as compact bone into the inward-growing moving cortex. This bone has an irregular convoluted appearance. 9. Periosteal and endosteal bone. Bone will be preferentially deposited and reabsorbed from periosteal and endosteal surfaces depending upon the direction of growth in a given region of bone. The entire thickness of a bone could be made entirely of one or the other, or a combination of the two.

In long bones of turtles and crocodilians, there is a large cartilaginous epiphysis that overlies a growth plate (Figures 1.89–1.90). Early in life in long bone of these two groups there is a temporary large cartilaginous cone below the growth plate at both ends of the shaft. In squamates, within the epiphysis of long bones, secondary centers of ossification develop, which are followed by bone formation (Figure 1.91). A growth plate is present where the epiphysis joins the shaft (Figure 1.92) and consists of several zones of endochondral bone formation: zone of reserve cartilage, zone of multiplication, zone of hypertrophy, zone of calcification, and zone of ossification (Figure 1.93). The diaphysis consists of periosteal and endosteal surfaces, with cortical compact bone in between (Figure 1.94). Histologically, arrest lines (lines of arrested growth) may be seen in cortical bone (Figure 1.94). Arrest lines form during certain periods such as hibernation. The medullary canal consists of spongy bone and vascular spaces. Bone marrow is not consistently present in the long bones of all reptiles. Vertebrae have a similar microscopic appearance as long bone and consist of compact bone, trabecular bone, and growth plates. Vertebrae often contain marrow. In healthy snakes, remodeling of vertebral bone is seen as irregularly arranged cement or reversal lines. This results in a “mosaic” appearance of normal bone (Figure 5.45). For the most part, the musculature of reptiles is far more developed than that of amphibians. This includes muscles used in supporting the body and used in locomotion, those used in jaw and tongue movements, and those used in pulmonary respiration. The musculature associated with ribs is

also better developed because the ribs of amphibians are either reduced in size or absent. While chelonians have well developed musculature associated with their girdles and associated limbs, because of the shell, trunk musculature is quite reduced. In contrast, and due to their locomotion, the trunk musculature of snakes is very well developed. Skeletal muscle of reptiles (Figure 1.95) is histologically similar to that of birds and mammals. Detailed descriptions of the musculature of reptiles can be found elsewhere (Gasc, 1981; Guthe, 1981). Homologous muscle groups are quite similar among the different vertebrates. However while in reptiles (as in birds and amphibians) many muscle fibers are tonic, tonic fibers in mammals are uncommon (Guthe, 1981). Tonic fibers show a graded response to a stimulus rather than an all-or-none response that is seen in a twitch fiber. Compared to twitch fibers, tonic fibers tend to be smaller, have less distinct fibrils, less sarcoplasmic reticulum, and are innervated by fine axons with grapelike endings rather than those with single end plates. While special staining with gold chloride can be used to demonstrate nerve endings, they also can be seen using transmission electron microscopy. Similar to mammals, muscle fibers in reptiles also are categorized as either broad white (paler fibers of wider diameter), narrow red (small dark granular fibers), or intermediate in size and color. Histochemical stains for certain enzymes such as succinic dehydrogenase and stains used for demonstrating myoglobin can be used to distinguish these fibers. Skeletal muscle fibers consist of both specialized intrafusal fibers (forming neuromuscular spindles) and typical extrafusal fibers. Differences that are seen among homologous muscles of different reptiles may reflect specific adaptations to the animal’s ecology.

1.4.3 Digestive System The digestive system contains all structures from the oral cavity to the cloaca. The mouthparts of reptiles are generally more complex than those of amphibians. Histologically, the oral cavity varies between different groups of reptiles. For most, the oral cavity is lined by a mucous membrane consisting of squamous non-keratinized epithelial cells, ciliated epithelial cells, goblet cells and columnar epithelial cells, with lingual, sublingual, and labial glands diffusely scattered about the submucosa (Figure 1.96). In the Nile crocodile (Crocodylus niloticus), the palate is lined by a keratinized stratified squamous epithelium that is continuous with that of the maxillary gingiva (Putterill and Soley, 2003). Pacinian-like corpuscles were found in both the gingiva and palate of the Nile crocodile. The tongue of reptiles varies in complexity. In some reptiles such as monitors and crocodilians it is keratinized. In the Nile crocodile, the dorsal aspect of the tongue in the midline was found to have a few shallow folds that increased in complexity and numbers toward the lateral borders (Putterill and Soley, 2004). Intraepithelial structures resembling taste buds

Overview of Reptile Biology, Anatomy, and Histology  

were also found. In the posterior two-thirds of the tongue of the Nile crocodile there were salivary glands and associated lymphoid tissue. In other reptiles such as the green iguana, the dorsal surface has papillary projections of a non-keratinized stratified squamous epithelium and a smooth ventral surface covered by a keratinized squamous epithelium (Figure 1.97). In tortoises and certain iguanid, anguid, and gekkonid lizards the tongue is richly invested with diffusely arranged salivary glands (Figure 1.98). The tongue of snakes is long, forked, muscular, aglandular, and is located in a sheath that is below the glottis (Figure 1.99). There are several species of crocodilians, such as the salt-water crocodile (Crocodylus porosus), having salt glands in the tongue. Sublingual salt secreting glands are present in the oral cavity of sea snakes. As previously mentioned (Section 1.2.2), crocodilians have a dorsal flap and ventral flap that together form a gular valve that separates the oral and pharyngeal cavities (Putterill and Soley, 2006). The glottis is located on the ventral floor of the pharyngeal cavity directly posterior to this valve (Figure 1.83). The dorsal fold represents an area of transition from the lightly keratinized epithelium of the palate to a pseudostratified respiratory epithelium of the pharyngeal cavity that consists of ciliated epithelial cells, goblet cells, and mucus-secreting glands. In the Nile crocodile, dorsally there were diffuse and nodular accumulations of lymphoid tissue that were scattered among the mucus-secreting glands. A similar transition was seen with the ventral fold. However, the ventral fold contained very little lymphoid tissue. Specialized oral glands include the labial venom glands of the Gila monster and beaded lizard, which are associated with the lower jaws and venom glands of venomous snakes, which are associated with the upper jaws. Venomous snakes in the family Colubridae have fangs in the caudal portion of maxillary bone and are considered rear fanged (opisthoglyphs). The venom gland of these snakes is a modified parotid gland and has been named Duvernoy’s gland. All members of the families Viperidae (vipers and pit vipers) and Elapidae (cobras, kraits, mambas, sea snakes) are front fanged. The fangs of viperids (Figure 1.72) are capable of rotating and are classified as solenoglyphs. Those of elapids are more fixed in position and are classified as proteroglyphs (Figure 1.74). However, as previously mentioned, some elapids are capable of maxillary and palatine erection while feeding (Duefel and Cundall, 2003). In front-fanged snakes a venom gland is located behind the eye (Figure 1.100). The structure and histology of venom glands differs among members of the Elapidae, Viperidae, and Atractaspididae (mole vipers) (Kochva, 1978). In elapids, the external adductor superficialis muscle attaches to the gland (Figure 1.100). The gland consists of simple to compound tubules that drain into a small central lumen in the center of the gland. A single duct drains the gland and runs though an accessory gland that attaches to the anterior surface of the venom gland. The duct fuses with the fang sheath. In mole vipers the gland is elongate, extending either slightly or significantly beyond

the base of the head. Tubules are distinctive in that they are arranged radially around an elongate central lumen. Mole vipers lack a separate accessory gland. In viperids the gland is attached to several bones by three ligaments. The major muscle that attaches to the gland is the compressor glandulae. The gland also receives insertions from the adductor externus profundus and pterygoideus muscles. The venom gland of viperids has the following four parts: main venom gland, the primary duct, the accessory gland (having two histologically distinct parts), and the secondary duct. The main venom gland consists of branching tubules (Figures 1.101–1.102) that empty into a central lumen that eventually becomes the primary duct. The primary duct at its terminus fuses with the fang sheath. Reptile venoms are complex mixtures containing inorganic ions, phospholipases, alkaline phosphatase, phosphodiesterase, toxins, neurotoxins, nucleosides, biogenic amines, peptides, polypeptides, proteases, endonucleases (enzymes affecting RNA and DNA), enzymes affecting cell membranes (lecithinases), enzymes affecting cell to cell attachment (hyaluronidases), enzymes affecting synaptic transmission, (acetylcholinases), and enzymes affecting clotting mechanisms (Elliott, 1978). Venom composition varies both among families and within families and even within genera and among species from different populations. The alimentary tract has essentially the same components as in higher vertebrates. The esophagus is a transport system carrying food to the stomach and shows modification from group to group. In marine turtles the esophagus is lined with a series of heavily keratinized papillae (Figure 1.103) that protect the animal from potential damage from a diet that may include highly abrasive foods such as spiculated sponges, jellyfish, and silicaceous plants. These structures also serve as filtering devices, somewhat similar to the baleen of whales. In sea turtles and some freshwater turtles, the esophagus makes an S-shaped bend prior to entering the stomach. Whereas the esophagus of sea turtles is lined with a multilayered keratinized squamous epithelium arranged into papillae, the esophagus of many reptiles is thrown into folds and is lined with a mucosal epithelium consisting of ciliated and goblet epithelial cells (Figures 1.04–1.07). In snakes, the posterior region of the esophagus is lined primarily with goblet cells. Glands may be found in the submucosa along with gut-associated lymphoid tissue (GALT). In boid snakes the GALT is organized into esophageal tonsils (Jacobson and Collins, 1980) (Figures 1.08–1.110). The esophagus in snakes is rather thin, especially in the cranial portion, and highly distensible, adapted to accommodating large prey items. In the specialized egg-eating snake, the anterior esophagus is penetrated by ventral vertebral hypopophyses, which function in perforating and crushing the shell of ingested eggs. The amount of muscle in the wall of the esophagus varies among major reptile groups. While muscularis mucosa is lacking in most chelonians, it is present in some tortoises (Figure 1.104) and sea turtles and is found in the posterior portion of the

10  Overview of Reptile Biology, Anatomy, and Histology

esophagus in the other orders of reptiles. In snakes the cranial esophagus is thin to allow distension for ingesting large prey items, and caudal to the heart the esophagus becomes more muscular. The esophagus leads into the stomach where digestion commences with mechanical and chemical enzymatic breakdown of food. The stomachs of turtles have both greater and lesser curvatures, while those of crocodilians are somewhat saccular. Lizards have ovoid stomachs, while those of snakes are linear and elongate, often located in the left coelomic cavity near the caudal pole of the liver. A sphincter demarcates the esophageal–gastric transition. The mucosa of the stomach is thrown up into longitudinal folds and the coloration is a light brown compared to a paler coloration of the esophagus. The extent of folding varies among groups. In crocodilians and many lizards there is relatively little folding with the stomach appearing smooth. Two components of the stomach are generally distinguishable: the corpus (fundus) and pars pylorica (Luppa, 1977). The multilayered mucosal epithelium of the esophagus abruptly transform into a glandular epithelium indicative of the stomach. The mucosa of the fundic stomach consists of rows of tubular, often branching glands that can be divided into a pit and a glandular body (Figure 1.111). Neck cells are present in most snakes and absent in most lizards and the tuatara. Depending on the group of reptiles, the glandular region contains either one or two cell types: dark serous (oxyntico-peptic) cells or clear (mucous) cells. Snakes and some lizards have neck cells and only dark serous cells in the glandular region (Figure 1.112). Using PAS stain, the neck cells and clear cells stain positive while the dark cells stain negative (Figure 1.113). The ratios of the two cell types vary among different regions of a given species and among species. The dark cells function as combined parietal and chief cells. Additional cells that are scattered about the glandular epithelium of the stomach and the intestinal tract are enteroendocrine cells. These cells contain membrane- delimited vesicles of secretory proteins, peptide hormones, and biogenic amines. Using novel monoclonal antibodies, the gastrointestinal tract of the Italian wall lizard (Podacris sicula) contained many cells that had chromagranin A (CgA) and chromogranin B (CgB) immunoreactivity (D’Este et al., 2004). Almost all cells that were immunoreactive for chromogranins were also argyrophilic. Immunoreactivity for CgA or CgB (or both) was found in almost all cells that were also immunoreactive for serotonin, histamine, substance P, and gastric peptide tyrosine-tyrosine. However, cells that were immunoreactive for neurotensin, gastrin/cholecystokinin, pancreatic polypeptide, and intestinal tyrosine cells did not show any co-reactivity for chromogranins. Below the epithelial cells covering the surface and lining glands, the following structures are seen from superficial to deeper layers: lamina propria, muscularis mucosae (internal circular and outer longitudinal layers), submucosa, tunica muscularis (inner thicker circular and outer thinner longitudinal layers) and serosa (Figures 1.111–1.115).

The tubular glands of the pars pylorica are shorter, less branched than in the fundus, and are lined with mucous cells (Figure 1.116). In chelonians the pylorus is distinct and muscular. In snakes it is less distinct. Dark cells are absent, and argentaffin and argyrophil cells are numerous in certain species. From the stomach, digesta moves into the small intestine where digestion is completed. In the duodenum the mixture of stomach, bile, pancreatic enzymes, and intestinal juices is usually alkaline or slightly acidic. The small intestine may be highly convoluted in turtles and lizards, while relatively straight with minor convolutions in snakes. The surface varies among groups and may be thrown into longitudinal and transverse folds, have a zigzag pattern or netlike or honeycomb appearance, or consist of fine striations (Parsons and Cameron, 1977). The epithelium is organized into villi, with poorly developed crypt-like glandular epithelium present in crocodilians, turtles, and lizards (Luppa, 1977) (Figures 1.117–1.118). Crypts are absent in snakes (Figure 1.119). The epithelium consists primarily of absorptive epithelial cells and goblet cells. The absorptive cells are PAS-negative and the goblet cells are PAS-positive (Figure 1.120). Paneth cells and, as previously mentioned, enteroendocrine cells have been identified in the intestinal epithelium of reptiles. Using the Grimelius silver nitrate technique, numerous enteroendocrine cells were identified in the intestine of the Iberian wall lizard (P. hispanica) (Burrell et al., 1991). Using immunocytochemistry, 11 types of immunoreactive endocrine cells were identified. Immunoreactive enteroendocrine cells were also identified in the intestine of the red-eared slider (Ku et al., 2001). In mammals, proliferative activity of intestinal cells is confined to the crypts of Lieberkühn where undifferentiated cells give rise to new cells that continually replace those that are lost at the tips of each villus, In reptiles it is unknown whether replacement occurs within crypt-like structures at the base of villi or if cellular division occurs along a greater length of each villus. Below the epithelium is a lamina propria consisting of vessels and connective tissue. The muscularis mucosae is very thin and consists of a single layer of muscle cells. The submucosa is below the muscularis mucosae, followed by the tunica muscularis and the serosa (Figure 1.121). Beyond the cranial duodenum, regions of the small intestine are not readily distinguishable. In herbivorous reptiles the length of the small intestine is generally longer than that of carnivorous reptiles. From the small intestine digesta passes into the colon. In some species, the transition between the two is not easy to define. As with the small intestine, the colon of herbivorous reptiles also has a much greater volume and length compared to that of carnivorous species. The surface of the colon may be thrown into folds or when filled with food, appears to be smooth. In herbivorous turtles and tortoises it is heavily convoluted and crosses from the right side to the left, and from dorsal to ventral. A distinct caecum is found just past the junction of the small intestine and colon in pythons. Herbivo-

Overview of Reptile Biology, Anatomy, and Histology  11

rous lizards and herbivorous chelonians may have partitions in the cranial colon that subdivides it into compartments. In herbivorous reptiles, oxyurid nematodes are common in this location. While some crypts are in the cranial colon of some species, in most they are minimal or completely absent. In most species the epithelium lining the lumen consists of a single layer of absorptive cells and mucin-containing columnar epithelial cells (Figure 1.122). In certain tortoises there are distinct glandular-like structures in the submucosa of the colon (Figure 1.123). Feces produced in the colon enter the cloaca, which is a terminal chamber common to both the digestive and urogenital systems. The cloaca is lined with columnar epithelial cells consisting of marginal and goblet cells. A variety of glands have been reported in the cloaca of reptiles. The liver of reptiles is bilobed, dark brown to black in coloration in healthy adult animals, and surrounded by a connective tissue capsule, Glisson’s capsule (Schaffner, 1998). In neonates and young reptiles the liver tends to be paler. In chelonians the left lobe is considerably smaller than the right and is closely associated with the lesser curvature of the stomach (Figures 1.124–1.125); the gastrohepatic ligament attaches the stomach to the left lobe of the liver. The right lobe of the liver of chelonians is attached to the duodenum by the hepatoduodenal ligament and the gallbladder is within the right lobe. In crocodilians, the liver resides in the cranial coelomic cavity with the heart located between both lobes and the lungs located dorsally. A pseudodiaphragm (posthepatic septum) (Figures 1.9, 1.126) separates the cranial (lungs, heart, and liver) from the caudal coelomic cavity of crocodilians. In most lizards the right lobe is also larger than the left, with the heart directly cranial to the liver (Figure 1.127). The liver may be located cranially in the coelomic cavity in some lizards such as iguanas and more caudally in others such as monitors. The gallbladder is also in the right lobe of the liver of lizards (Figure 1.27). Bile ducts transport bile from the gallbladder to the duodenum (Figure 1.128). In snakes, the liver is elongate and somewhat flattened (Figure 1.129). The lung is attached to the dorsal surface of the liver and gradually transforms into an air sac. Two vessels, the portal vein and the hepatic vein, are located dorsally and ventrally, respectively, and run the length of the liver, dividing it into its two lobes (Figure 1.130). There is a narrowing of the hepatic parenchyma between the two vessels. Upon leaving the liver the hepatic vein joins the post cava. The gallbladder of snakes is located caudal to the liver and is closely associated with the pancreas and spleen (Figures 1.131–1.133; 4.30). The mucosa of the gallbladder is thrown into folds and consists of a simple or pseudostratified layer of columnar epithelial cells (Figures 1.134–1.36). A lamina propria consisting of fibrous connective tissue and blood vessels is below the epithelium and muscle fibers are found outside the lamina propria.

Histologically, the liver is not as distinctly organized into lobules as in mammals (Schaffner, 1998). In some reptiles, hepatic cords may be arranged radially around a central vein (Figures 1.137–1.138). In snakes, a radial arrangement of hepatic plates around a central vein is generally not apparent (Figure 1.139). A distinct layer of connective tissue does not separate adjacent lobules. The periphery of a lobule can be defined by a bile ductule, branches of the hepatic vein, and small amounts of connective tissue. A branch of the hepatic artery may or may not be seen (Figures 1.140–1.143). As in mammals, hepatic cords are separated by sinusoids that are lined with Kupffer cells and endothelial cells (Figure 1.144). Stellate or Ito cells have been identified in hepatic sinusoids of reptiles. The Kupffer cell serves as a sinusoidal macrophage and the Ito cell is the main storage site of vitamin A. Melanomacrophages are commonly seen in the liver of reptiles and are extremely numerous in some species (Figures 1.145–1.146, 5.8). Clusters of melanomacrophages are sometimes referred to as melanomacrophage centers. While most snakes have few melanomacrophages in the liver (Figure 1.147), in some they may be numerous (Figure 1.148). These cells function as macrophages and free radical scavengers. The brown pigment seen in melanomacrophages stained with hematoxylin and eosin (H&E) (Figure 1.149) consists of both iron (Figure 1.150) and melanin granules (Figure 1.151). In many diseases, especially chronic illnesses, they will increase in size and number. The pancreas of reptiles is located along the mesenteric border of the duodenum (Miller and Lagios, 1970). In most species it is closely associated with the spleen. In the few pleurodiran (sideneck and snakeneck) turtles studied, the spleen and pancreas are separated (Figure 1.152). In cryptodiran chelonians, crocodilians, and lizards it is elongated (Figures 1.128, 1.153) and in snakes it is triangular in shape (Figures 1.131–1.133). The exocrine pancreas consists of branching tubules containing epithelial cells with zymogen granules (Figures 1.154–1.155). In contrast to mammals, there is no intercalated or transitional segment and the islets of Langerhans lack a distinct demarcation from the exocrine pancreas (Figures 1.156–1.158). In some reptiles, the cytoplasm of both the islets of Langerhans and exocrine pancreas may stain eosinophilic with H&E staining (Figure 1.159). Lizards have an elongate pancreas (Moscona, 1990), which is considered a more primitive trait than the compact pancreas of snakes. In snakes there is a concentration of the islets of Langerhans adjacent to and surrounded by the spleen (Figure 1.160). In some snakes, islet tissue is within the spleen. In snakes in the families Typhlopidae, Boidae, and Pythonidae, the dorsal lobe of the pancreas is connected to the ventral lobe by an isthmus and is closely associated with the spleen. This anterior portion has been named the juxtasplenic body. It consists primarily of islet cells. Based on structure of the lobes and ducts, their spatial relationship with spleen and gallbladder, and the location of islet cells, the pancreas of 18 species of snakes was categorized into five types

12  Overview of Reptile Biology, Anatomy, and Histology

(Moscona, 1990). Lizards in the genus Varanus (monitors) appear to have a pancreas that is transitional between that of lizards and snakes. While islet cells are scattered within the dorsal lobe of the pancreas (Figure 1.161), there is a concentration in a juxtasplenic body adjacent to the spleen (Figures 1.162–1.163). In squamates, α and β cells, the main endocrine cells of the islets of Langerhans, are not separated into distinct groups and lie together along vascular channels (Miller and Lagios, 1970). In chelonians and crocodilians, α and β cells are segregated, with β cells centrally located and surrounded by α cells. The spectacled caiman (Caiman fuscus), Nile crocodile, American alligator, American or green anole (Anolis carolinensis), and garter snake (Thamnophis sirtalis) also have somatostatin-containing and polypeptidecontaining cells in the pancreas (Jackintell and Lance, 1994; Rhoten, 1984; Rhoten, 1987a,b; Rhoten and Hall, 1981, 1982). The physiology of the gastrointestinal tract is affected by environmental and ultimately, the body temperature of reptiles. Decomposition rates of ingested food, passage rates of food through the gastrointestinal tract, motility of the tract, and rates of absorption are affected by body temperature. For the most part, no digestion takes place below 7°C, and digestion takes place extremely slowly in the temperature range from 10 to 15°C. Hibernating species typically enter hibernation with the proximal part of the alimentary tract empty. Digestion may also slow down near the highest temperatures tolerated. Within the preferred optimum temperature range, higher temperatures raise the animal’s general metabolism and increase the rate of secretion of digestive juices resulting in increased amounts of enzymes in the digestive tract. Burmese pythons that consume large meals after long periods of time, showed an increase in oxygen consumption and intestinal nutrient uptake rates and capacities, within 1 to 3 days of ingestion (Secor and Diamond, 1995). This is an example of up-regulation. After they feed, Burmese pythons can also activate gastric functions well beyond that of mammals (Secor, 2003; Secor et al., 2001). Much of the energy expended was measured before the prey energy was absorbed (Secor and Diamond, 1995). The mass of the small intestine and other organs increased during this period (Secor et al., 1994; Starck and Beese, 2001). All these values returned to fasting levels 8 to 14 days later. Smaller snakes such as sidewinders (Crotalus cerastes) also increased their metabolic rates following feeding, but not to the same extent as larger pythons (Secor et al., 1994). In red-sided garter snakes (T. s. parietalis), the patterns of up- and down-regulation of the small intestine were similar to that for pythons but not to the same extent (Starck and Beese, 2002). In large pythons, within 24 h following feeding, circulating concentrations of the following biochemicals increased: cholecystokinin (CCK), glucose dependent insulinotropic peptide (GIP), glucagon, and neurotensin (Secor et al., 2001). During digestion, the peptides neurotensin, somatostatin, motilin, and vasoactive intestinal peptide were found primarily in the stomach, GIP and glucagon in the pancreas, and CCK and substance P in

the small intestine. Following feeding, there was a decline in tissue CCK, GIP, and neurotensin (Secor et al., 2001). The white-throated monitor (Varanus albigularis), another reptile that may feed intermittently, increased oxygen consumption following feeding to rates 7 to 10 times the prefeeding values (Secor and Phillips, 1997). Reptiles that feed upon large prey items after long periods of not feeding may serve as important models of feeding-induced extreme physiological regulation (Secor and Diamond, 1998).

1.4.4 Respiratory System The respiratory tract can be divided into upper and lower components. The upper component is the nose, consisting of the external nares, vestibulum nasi, cavum nasi proprium, conchae, ductus nasopharyngeus, and internal nares (Parsons, 1970). Associated with the nose is the vomeronasal organ (see Section 1.4.13). Reptiles normally breathe with their mouth closed, with air entering the upper respiratory tract through the external nares. In chelonians, the vestibulum nasi is keratinized and opens into a large cavum nasi proprium, which occupies the area of the head cranial to the eyes (Figures 1.164, 10.46). This cavity is lined with a mucous epithelium ventrally (Figure 10.47) and a multilayered olfactory epithelium dorsally (Figure 10.48). (Jacobson et al., 1991). A mucous-lined ventral recess comes off the ventral nasal passageway with the latter opening into the roof of the pharynx at the choanae. Right and left nasal cavities are separated by a cartilaginous septum. Numerous mucous glands are associated with the nasal cavity. Conchae are absent in chelonians and are present in crocodilians, tuataras, lizards, and snakes (Figures 1.165–1.166). The nasal cavities of crocodilians are more complex than any other reptile. There are three conchal formations on the lateral wall of the cavum nasi proprium. The lower respiratory tract consists of the glottis, larynx, trachea, paired bronchi, and paired lungs. Reptiles breathe primarily by lungs, but some aquatic turtles have supplemental cloacal, pharyngeal, or cutaneous respiration. The lungs of living reptiles vary widely among groups in the degree of complexity, being generally more complex and efficient than those of amphibians and less complex than those of birds and mammals. The lungs can be categorized based on structure, the type of parenchyma, and the pattern of intrapulmonary distribution of the gas-exchange component (Perry, 1998). The extent of partitioning of the lung has also been used to categorize different types of the reptile lung. The structural types include unicameral (single chamber), multicameral (multiple chambers), and transitional lungs (while septa are present, the chambers remain confluent). In chelonians the lungs are equal (or nearly equal) in size and occupy the dorsal coelomic cavity, with the dorsal pleura adherent to the peritoneum. In turtles the lungs extend to the cranial poles of the kidneys (Figures 1.67, 4.43). All chelonians have bronchi, with each bronchus remaining unbranched

Overview of Reptile Biology, Anatomy, and Histology  13

and running the entire length of the lung. Depending upon the family of chelonians, the lung is subdivided into from 3 to 11 chambers (Perry, 1998). The chambers open into the central intrapulmonary bronchus (Fleetwood and Munnell, 1996). In crocodilians the lungs are equal in size, relatively short and located in the cranial coelomic cavity with the liver and heart, and separated from the caudal coelomic cavity by a pseudodiaphragm (Figures 1.9, 1.126). All crocodilians have multichambered lungs. The tuatara lacks bronchi and has a single chambered lung. In most lizards the right and left lungs are comparable in size and occupy the cranial half of the coelomic cavity (Figure 1.168). Lizards vary in the structure of their lungs. Members (the few studied) of the families Iguanidae, Agamidae, and Chamaelontidae have transitional lungs (Figure 1.169). Chameleons have tentacular diverticula projecting from the lungs (Figure 1.170). Of lizards, varanids and helodermatids are unique in that they have multichambered lungs. In snakes the lung is elongate, gradually merging into an air sac that terminates in the intestinal mesentery in the vicinity of the gallbladder in terrestrial species and to the cloaca in some aquatic species such as sea snakes. In boid and colubrid snakes, the respiratory portion of the lung lies between the heart and cranial pole of the liver (Figure 1.171), while in most viperids and elapids it is situated cranial to the heart. In all snakes the right lung is larger than the left, with the left lung being fairly well developed in boids and only vestigial in colubrids. Snakes in the families Anomalepididae, Typhlopidae, and Acrochordidae have multichambered lungs. In other snakes they are single chambered. The lung gradually transforms into an air sac along the surface of the liver (Figures 1.172, 4.29). Trachea and bronchi are lined with ciliated, nonciliated secretory, and basal epithelial cells. These cells continue into the lung of those reptiles having intrapulmonary bronchi. The respiratory region of the lung is divided and subdivided by interconnecting septae or trabeculae into terminal gas exchange units that are named either faveolae or ediculae for air exchange chambers that are either deeper than they are wide or wider than they are deep, respectively. This gives the lung a honeycomb appearance when opened, flattened, and viewed from above (Figure 1.173). Trabecular parenchyma has also been described in which the trabeculae are fused with the inner wall of the lung and do not support free septae. In snakes the lung runs across the surface of the liver, where it slowly transforms into a nonrespiratory air sac. The air sac extends, depending on the species, from the level of the gallbladder to the cloaca. Faveolae and ediculae are absent from the air sac. When viewed by light microscopy in cross-section, the faveolar parenchyma of snake lungs opens into a large central chamber (Figure 1.174). Adjacent faveolae and ediculae are separated by a connective tissue septum containing blood vessels and occasionally lymphoid aggregates (Figures 1.175–1.76). At the luminal end of each septum, and surrounding each faveola and edicula, is a cord of smooth

muscle cells (Figures 1.174, 1.176–1.77). In cross-section this appears as a discrete muscle bundle. The luminal septal–surface overlying the smooth muscle bundle projecting into the central chamber is covered by cuboidal to columnar ciliated epithelial cells, nonciliated secretory epithelial cells, and basal cells. Faveolae and ediculae are lined primarily by squamous (type I cells) epithelial cells, and to a lesser degree by cuboidal (type II) epithelial cells (Figure 1.178). Overlying capillary beds where gas exchange occurs are type I epithelial cells. Type II cells contain punctate staining granules, which by electron microscopy represent lamellar material (Figure 1.179). In some reptiles, between ciliated epithelial cells and type I and II epithelial cells are hedge cells. These cells contain microvilli and may be involved in reabsorption of fluid on the surface of the lung. In other reptiles, hedge cells are absent and serous epithelial cells are found in this location (Luchtel and Kardong, 1981). Neuroendocrine cells are scattered about the lining epithelium, either singly or as domelike bodies (Fleetwood and Munnell, 1996; Perry, 1988; Perry et al., 1989). As previously mentioned, on the surface of the liver the respiratory portion of the lung slowly transforms into a thin-walled nonrespiratory air sac, which is lined with a columnar to squamous epithelium (Figures 1.180–1.181). Whereas the lung volume of reptiles is quite large in comparison with that of mammals, the surface area is 10 to 20% as large as a comparably sized mammal (Perry, 1998). This is consistent with the basal metabolic rate of reptiles ranging from one-tenth to one-third that of mammals of comparable weight. In snakes, a major portion of this volume is due to the presence of an air sac. This air sac may act as a reservoir for oxygen during periods of apnea. In aquatic forms it may also act as a buoyancy organ.

1.4.5 Urinary System The evolution of reptiles resulted in significant differences between their urinary system and that of amphibians. In amphibians the reproductive system is more intimately associated with the urinary system than in reptiles. Differences in the urinary system of reptiles compared to amphibians reflect adaptation for conserving body water in a generally dehydrating environment, the evolutionary trend toward anatomic separation of excretory and reproductive tracts, and development of more precise control over the animal’s internal environment. Components of the urinary system include paired kidneys, ureters, and urinary bladders in chelonians and many lizards. Crocodilians and snakes lack a urinary bladder. While most lizards have a urinary bladder, in some it is rudimentary and in others it is absent (Beuchat, 1986). The nitrogenous excretions of reptiles may be in the form of ammonia, urea, or uric acid. The proportions of these products vary with the lifestyle of the particular species. Marine and highly aquatic freshwater turtles and crocodilians excrete up to 25% ammonia as a percent of total urinary nitrogen. Amphibious pond and swamp turtles excrete

14  Overview of Reptile Biology, Anatomy, and Histology

approximately two to four times as much urea as ammonia or uric acid. Tortoises, the tuatara, lizards, and snakes (especially those found in deserts) excrete primarily uric acid. Reptiles as a group cannot concentrate urine above blood osmolarity, thus the ability to produce and excrete uric acid, which is insoluble in water, serves as a mechanism for conserving water. Also many desert lizards have evolved salt glands that allow excess salt to be eliminated without losing needed water. Kidneys of reptiles of different groups show both similarities and differences. All reptiles have lobulated, paired kidneys that are approximately equal in size in most species. The color ranges from light to dark brown. In snakes, the kidneys are located in the caudal coelomic cavity, with the right kidney situated more cranial than the left kidney. While in most species the kidneys are of equal size, in some the right kidney may be larger than the left. Kidneys of snakes are elongated, lobulated, and some are longer and thinner and others shorter and wider relative to the size of the snake (Figures 1.182–1.183). In female snakes the kidney is dark reddish-brown in coloration, with streaks of urates throughout. Kidneys in adult male snakes, especially during periods of breeding, have a creamy pale coloration (Figures 1.184, 4.32) due to hypertrophy of a segment of the nephron called the sexual segment (see below). This may be confused with a “gouty” kidney. In chelonians, crocodilians, and lizards the kidneys are shorter, broader, and located near the pelvic canal (Figures 1.185–1.187, 4.44). In some lizards such as the green iguana the caudal pole of the kidney extends into the base of the tail, just caudal to the caudal margins of where the hindlimbs join the body wall (Figures 1.187, 4.19). In lizards, the ureter and oviduct open separately into the cloaca while in males the ureter and vas deferens open conjointly into the cloaca. In chelonians, crocodilians, and snakes the ureter and both oviduct and vas deferens open separately into the cloaca. As in other vertebrate groups, the structural and functional unit of the reptile kidney is the nephron. While the human kidney has approximately 2 million nephrons in each kidney (Fawcett, 1986), reptile kidneys typically have only a few thousand nephrons (Fox, 1977). Renal corpuscles, consisting of a Bowman’s capsule and glomerulus, are present in the majority of reptiles; some lizards and snakes have aglomerular tubules. Bowman’s capsule consists of an outer capsular (parietal) epithelium and an inner glomerular (visceral) epithelium. While in most reptiles the capsular epithelium is squamous (Figure 1.188), in some such as the green iguana it is cuboidal (Figure 1.189). Compared to those of amphibians, there is a definite reduction in the size of glomeruli in reptiles; lizards typically have the smallest. This is an adaptation that conserves water by reducing the flow of urine into tubules. Glomeruli of reptiles are not as vascular as those of birds and mammals. In crocodilians many renal corpuscles uniquely line up within the parenchyma equidistant from the capsule (Figure 1.190). Several

segments of the reptile nephron can be histologically distinguished (Bishop, 1959). Beginning with Bowman’s capsule, the nonsecretory neck segment composed of cuboidal cells (many having cilia) continues as the next segment of the reptile nephron (Figures 1.191–1.192). The nuclei of neck segment cells occupy most of the cytoplasm of the cell. The neck segment is followed by the proximal segment consisting of proximal tubules (PT). The PT is lined by cuboidal cells that, with hematoxylin and eosin staining, have an eosinophilic staining cytoplasm (Figures 1.188–1.89, 1.191). These cells lack cilia but have well-developed microvilli on their luminal surface. With PAS staining, the brush border and small granules within the cytoplasm stain with PAS (Figure 1.192). The next segment, the intermediate segment, has an initial ciliated region that is followed by an area of mucus cells. The cells of this segment, while similar in appearance to those of the neck segment, stain basophilic with H&E, and have a smallerdiameter tubule than the PT (Figures 1.193–1.194). This leads to the distal segment (Figures 1.188, 1.189, 1.193, 1.195), followed by the collecting tubules (Figure 1.196). The kidney of lizards and snakes is sexually dimorphic, with males having an enlarged portion called the sexual segment (Bishop, 1959), which is located between the distal segment and collecting tubules (Figures 1.197–1.199). This segment produces a secretory product that is incorporated into the seminal fluid. Varying degrees of seasonal changes occur in this segment, with lizards appearing to have greater seasonal changes compared to snakes. Following sexual maturity, while the northern water snake (Nerodia sipedon sipedon) maintains a level of sexual segment hypertrophy throughout the year, changes in granule appearance correlate with changes in concentration of plasma androgens (Krohmer, 2004). In the black swamp snake (Seminatrix pygaea), the sexual segment of the kidney does not go through an extended period of inactivity, but does show a cycle of synthesis and secretion that was found to be related to the spermatogenic cycle and mating activity of this snake (Sever et al., 2002). The urinary bladder is absent in crocodilians, some lizards, and snakes. It is present in all chelonians and most lizards. In aquatic turtles the wall is relatively smaller and thicker (Figure 1.200) than in tortoises (Figure 1.201). It is highly distensible, contains thin bands of muscle, and is lined with a transitional epithelium (Figures 1.202–1.204). In tortoises the bladder is a major site of fluid and ion (potassium) storage during periods of drought. The urinary bladder opens into the cloaca near the opening of the ureters and reproductive tract openings.

1.4.6 Reproductive System Reptilian gonads (testes and ovaries) are paired organs and are derived from the germinal ridge, with the testes derived from the medulla and the ovaries from the cortex. They are located in the abdominal cavity, and in most species are in close proximity to the cranial poles of the kidneys (cheloni-

Overview of Reptile Biology, Anatomy, and Histology  15

ans, crocodilians, and many lizards). In snakes the gonads are caudal to the gallbladder.

cranial to the cloacal opening, and the hemipenes of lizards and snakes are inverted within the base of the tail.

1.4.6.1 Male Reproductive System — Anatomy and Histology  The external color of the testes varies within

1.4.6.2 Female Reproductive System — Anatomy and Histology  The female reproductive tract consists of paired

and among the orders of reptiles and includes those that are white, yellow, brown, and black (Figures 1.186–1.187, 1.205–1.212). In many snakes and lizards the right testes is cranial to the left, whereas the gonads are symmetrically arranged in crocodilians and chelonians. The testes of most reptiles are smooth and ovoid to elongate, although those of blind snakes in the genus Leptotyphlops are multilobed. Testes size generally varies seasonally, with increased size reflecting spermatogenesis. In those reptiles that hibernate, the testes may be of maximum size at the time of emergence in the spring, while in others it will be in the early summer. Testes are surrounded by a tunica albuginea. The interior consists of convoluted seminiferous tubules (Figures 1.213– 1.216) with the interstitium containing fibroblasts, blood vessels, lymphatics, and interstitial cells (Figures 1.217–1.218). Interstitial cells vary in size, number, and appearance depending on the stage of the reproductive cycle. In some stages they are easy to miss. They may occur either singly or in groups. Some lizards have a collar of interstitial cells below the tunica albuginea (Fox, 1977). Seminiferous tubules are lined with seminiferous epithelium consisting of Sertoli cells and developing germ cells. The germ cells can be categorized into the following three layers of different germ cell types: (1) spermatogonia near the basement membrane, (2) spermatocytes underneath the spermatids, and (3) centrally located spermatids and spermatozoa (Figures 1.214–1.215). In birds and mammals, germ cells are synchronized in their spatial development with different layers containing germ cells at a stage of development. This ultimately results in waves of spermatogenesis during the breeding season. In contrast, the slider turtle (Trachemys scripta) and the European wall lizard (Podarcis muralis) have a strategy of germ cell development in which a temporal progression of the germ cell population results in a discrete period of spermatozoa production and release (Gribbins and Gist, 2003; Gribbins et al., 2003). Because so few reptiles have been studied, it is unknown if this pattern of spermatogenesis is typical for most reptiles. Seminiferous tubules ultimately empty into ductuli efferentia (Figure 1.219) that are lined by a single layer of flattened cells. From here sperm enters the ductuli epididymides, followed by the ductus epididymis, and then the ductus deferens (Figure 1.220). Both ciliated and nonciliated cells line these structures and muscle is located in the wall of the vas deferens. In some reptiles the vas deferens may join the ureters at a common opening in the cloaca. Sperm are conveyed by the vas deferens to the penis of chelonians (Figure 1.221) and crocodilians (Figure 1.222), and the paired hemipenes of lizards and snakes (Figures 1.223–1.224). The tuatara has no specialized copulatory organ. The penis of chelonians and crocodilians is

ovaries and, in most species, a pair of reproductive ducts. Ovaries vary in their location and may be found in the midto caudal-coelomic cavity in chelonians, crocodilians, and lizards (Figures 1.225–1.229, 4.18), and near the gallbladder in snakes (Figure 1.132). The ovaries of snakes are elongated (Figures 1.230–1.231). The ovary contains a hierarchy of follicles at different stages of development and atresia. Vitellogenic follicles can fill the coelomic cavity of lizards and are commonly removed if they fail to enter the oviducts and develop a shell (Figure 1.232). The complement of follicles and the relative proportions of different sizes and stages depend on the species and stage of the reproductive cycle. As with spermatogenesis, follicular development is influenced by season. Follicular development in reptiles is divided into previtellogenic and vitellogenic phases. Vitellogenesis refers to the accumulation of yolk within the developing oocyte following synthesis of yolk precursors in the liver. Grossly, previtellogenic and vitellogenic follicles can be distinguished by size and color. Previtellogenic follicles are small and white and become yellow and enlarge as they are recruited into vitellogenesis (Figures 1.233–2.235). In many species, the ovaries are quiescent for most of the year and most follicles are small and previtellogenic. Oogenesis and the histology of the ovary and oviduct have been reported for several species of reptiles (Callebaut et al., 1997; Guillette et al., 1989; Guraya, 1968; Hubert, 1985; Motz and Callard, 1991; Palmer and Guillette, 1988; Palmer and Guillette, 1992; Uribe and Guillette, 2000). Primordial follicles consist of a nucleus, ooplasm, and a layer of simple squamous follicular epithelial cells (Figure 1.236). Previtellogenic follicles are devoid of yolk (Figures 1.236–1.238), whereas yolk platelets are present in vitellogenic follicles (Figures 1.239–1.241). The acellular zona pellucida separates the ooplasm from the granulosa (Figures 1.239–1.240). Developing follicles undergo dramatic changes during development and there are notable differences among groups of reptiles. Oocytes are surrounded by a single layer of flattened or cuboidal granulosa cells during early previtellogenesis (Figures 1.236, 1.238, 1.242). As development progresses, the granulosa of squamates becomes more complex and assumes a polymorphic appearance (Figures 1.243–1.244). At this stage, granulosa cells are classified as pyriform cells, intermediate cells, and small cells. As vitellogenesis begins, however, the granulosa transitions back into a monomorphic single layer of flattened or cuboidal cells (Figures 1.245– 1.246). In contrast, the granulosa of chelonians and crocodilians (Figures 1.239–1.240) remains a single monomorphic layer throughout follicular development. Changes also occur in other structures. The theca interna and externa become distinct as the follicle matures and the zona pellucida 

16  Overview of Reptile Biology, Anatomy, and Histology

differentiates into an inner striated layer and an outer homogeneous layer. As in other vertebrates, follicles either ovulate or undergo atresia. Following ovulation, corpora lutea are formed (Figures 1.233, 1.247, 1.248) and persist throughout gravidity. In squamates, early in corpus luteum formation there is a central luteal cavity that is lined by proliferating granulosa cells (Figure 1.249) that eventually fill this cavity (Guraya, 1989) (Figure 1.250). Chelonians have a similar appearing corpus luteum (Figure 1.251). The corpora lutea of crocodilians and tuataras, however, have a proportionately larger thecal component and the luteal mass never refills the follicle (Guillette and Cree, 1997; Guillette et al., 1995). In addition, the ovulation aperture never closes as it does in squamates. Following oviposition or parturition, the corpora lutea regress and become corpora albicans, which may persist for months or years. Corpora albicans are scarlike structures that are typically comprised of abundant stroma and may contain entrapped pigment-laden cells (Figure 1.252). Alternatively, follicles may undergo atresia at any stage of development, including late vitellogenesis. When vitellogenic follicles undergo atresia, the granulosa undergoes dramatic hypertrophy and hyperplasia and becomes highly vacuolated as granulosa cells phagocytize the yolk (Figures 1.253–1.257). Histiocytes and other leukocytes also may infiltrate the follicle and participate in yolk absorption. The frequency of follicular atresia varies by species and season and may be common or very rare (Guraya, 1989). The oviduct (uterus) (Figures 1.227, 1.234) courses from the region adjacent to the ovary to the cloaca, where in most species they open independently. The wall of the oviducts consists of two layers of smooth muscle surrounded by an outer serosa. The following five regions of the generalized reptile oviduct are recognized (Girling, 2002): (1) infundibulum (anterior and posterior portions in some reptiles) (Figure 1.258), (2) the uterine tube, (3) the isthmus (aglandular portion), (4) the uterus (American alligator has two regions similar to those seen in birds [Palmer and Guillette 1992]), and (5) the vagina (thick muscular region). Not all five regions are present in all reptiles and some reptiles have additional regions. In snakes in the genus Typhlops and Leptotyphlops, the left oviduct is missing (Fox, 1977). Oviducts are lined with ciliated and nonciliated mucous cells, with various glands found below the lining epithelium in the uterine tube and uterus of certain reptiles. In chelonians and crocodilians the mucosa is packed with glands that are thought to produce albumen (Figures 1.259–1.261). Tuataras, lizards, and snakes do not produce an albumen layer in their eggs, and thus lack glands in this segment of the oviduct. Still, albumens have been detected in the eggs of some squamates. The origin of this albumen is unknown. The mucosal glands that produce the calcareous and fibrous components of the eggshell and the shell membrane are in the mucosa of the uterus of oviparous reptiles (Figures 1.262–1.263). Few glands are present in the uterus of viviparous species (Figure 1.264). In those

segments of the oviduct having glands, their extent of development will vary with the reproductive status of the female (Figure 1.265). The uterus grades into the vagina, which is lined by ciliated cuboidal cells; no glands are present in the submucosa (Figure 1.266). The vagina opens into the cloaca either through a common urogenital opening or through separate cloacal openings.

1.4.6.3 Fertilization  Fertilization is internal in all reptiles and takes place near the cranial end of the oviduct. This occurs prior to deposition of the egg envelopes by glands lining the oviduct. Based upon the type of postfertilization developmental patterns, reptiles can be grouped as those that are egg layers (oviparous) and those that are live-bearers (viviparous) (Packard et al., 1977). Oviparity is believed to represent the ancestral mode of reproduction in the class Reptilia. Oviparous reproduction characterizes all living chelonians, crocodilians, the tuatara, and most species of lizards and snakes. Many species of lizards and snakes are viviparous and viviparity is believed to have evolved in squamate reptiles over 100 times (Shine, 1985). A primitive placenta is present in viviparous reptiles. Several species of reptiles have been shown to possess seminal receptacles in the female reproductive tract (Conner and Crews, 1980; Fox, 1956), and it is known that at least some reptiles can store sperm up to several years, producing viable eggs or offspring even though not in direct contact with a male. Seminal receptacles are located in either the posterior uterine tube, isthmus, or anterior vagina (Palmer et al., 1993; Perkins and Palmer, 1996; Sakar et al., 2003). 1.4.6.4 Reproductive Cycles  Much has been published on reproductive cycles of reptiles. Many species show cyclical changes in the development of the reproductive organs, which is timed in each species (or population) to gain maximum benefits from favorable climatic conditions and food sources (Crews and Garrick, 1980). Thus, reptiles from different regions and habitats employ a variety of reproductive strategies. While a complete review of reproductive strategies is beyond the scope of this chapter, the information below will provide some examples of different cycles encountered in various species. In temperate regions many species breed soon after hibernation ends, with young delivered or hatching later in the season when food resources are plentiful. Many lizards, such as members of the genus Sceloporus and the sideblotched lizard (Uta stansburiana), emerge from hibernation with their testes at maximum size, with testicular atrophy progressing through the summer (Fox, 1977). In other lizards the testes may be small after emergence from hibernation, with maximal size from spring to early summer. In lizards of the genus Emoia from New Hebrides, the reproductive cycles are almost continuous, with minimal and maximal periods of May to June and November to December, respectively. In garter snakes (Thamnophis spp) the testes are smallest between

Overview of Reptile Biology, Anatomy, and Histology  17

December and May and largest between July and October. Desert tortoises of the southwestern United States often breed in the fall when the testes are well developed and sperm is in the epididymis. In this species sperm remains in the epididymis through the winter and is still present in the spring when the tortoise emerges from hibernation. Copulatory behavior does not necessarily coincide with height of testicular development. At least in some garter snakes, mating takes place in the spring when the testes are small and inactive. Sperm used for reproduction is stored in the epididymis and derived from the previous year’s testicular activity during the summer (Cieslak, 1945). There are many species differences in the ovarian cycle of reptiles. Furthermore, geographic variation may occur within the same species. Follicular enlargement does not necessarily coincide with active spermatogenesis, as stored sperm from previous months is used for breeding by some species. In these species, ovulation coincides with discharge of spermatozoa from the epididymis. Ovarian cycles have been studied in a variety of reptiles and ovulation schemes have been classified as polyautochronic, monoautochronic, and monoallochronic (Etches and Petitte, 1990; Guraya, 1989). Polyautochronic refers to simultaneous ovulation of many ova from both ovaries. Monoautochronic reptiles, which include most gecko species, ovulate a single ovum simultaneously from each ovary, whereas monoallochronic reptiles, such as Anolis spp., ovulate one ovum from either the right or left ovary, and alternate between ovaries for each single-egg clutch. In snakes and turtles, many ova are ovulated from a single ovary (Aldridge, 1982; Licht, 1982).

1.4.6.5 Reproduction — Endocrine Control and Environmental Influences  Reproductive behavior and gonadal activity are influenced by a milieu of environmental, nutritional, and endocrine factors. Clutch size and number are likely controlled by similar mechanisms that operate within anatomical and physiological limitations of any given species. Different aspects of hormonal regulation of reproduction have been studied in many reptiles. It is thought that follicular development and ovulation, vitellogenesis and steroid hormone secretion are regulated by pituitary gonadotropins, as in other vertebrates. Gonadotropins similar to follicle stimulating hormone (FSH) and luteinizing hormone (LH) appear to be secreted in crocodilians and chelonians (Guaraya, 1989). Furthermore, an ovulatory surge of LH and progesterone, comparable to that of mammals and birds, has been documented in multiple chelonians (Licht, 1982). In contrast, only one gonadotropin, an FSH-like compound, is secreted by the pituitary gland of squamates (Guraya, 1989). Steroid hormones, which include estrogen, testosterone, and progesterone, have many important roles in reptile reproduction. An example is the stimulation of vitellogenesis by estrogen. Also, progesterone has many effects that support egg and embryo development, including expression of egg-white proteins and suppression of myometrial activity (Custodia-Lora, 2002).

The reptile for which cyclical reproduction patterns and endocrine control mechanism has been best described is the green anole. In this species, as with most animals in general, an array of internal and external factors works in concert to control the patterns of reproductive biology. From late September to late January, both males and females are reproductively quiescent. In late January (this varies with the geographic range of this species, which is throughout the southeastern U.S.), the males emerge from hibernation (winter dormancy) and establish breeding territories. About 1 month later, the females emerge, and by May they are laying a single-shelled egg every 10 to 14 days. The annual reproductive cycle of the female has been divided into three distinct periods (Crews, 1975). Winter ovaries contain both previtellogenic (unyolked) follicles and atretic follicles, with the former being arranged in a stepwise size hierarchy (Jones et al., 1973). Beginning in March, yolk deposition commences and results in these follicles becoming vitellogenic follicles. Ovulation occurs at a diameter of about 8 mm. Production and secretion of intrafollicular estrogen stimulates follicular hyperemia and subsequent follicular growth. This increased vascularity may lead to the greatest yolk deposition in the most hyperemic follicle (Jones et al., 1975). Initiation of vitellogenesis is correlated with rising spring temperatures, which initiate FSH release by the pituitary. During the ensuing breeding season in green anoles, a single follicle matures and is alternatively ovulated between the two ovaries every 10 to 14 days. In late August, in the last period of the ovarian cycle, vitellogenesis ceases and the yolking follicles commence a rapid degeneration resulting in the formation of corpora atretica.

1.4.6.6 Parthenogenesis  Several lizards and snakes have developed parthenogenesis as a successful reproductive strategy (Cole, 1975; Darevksy et al., 1985). These animals provide a unique source of individuals that show little genetic variation between one another and are the only known vertebrates to reproduce normally by this method. The best studied of these reptiles involve the teiid whiptail lizards, including Cnemidophorus uniparens, C. velox, and C. tesselatus, all from the western United States. This may have evolved along geographic lines of hybridization between closely related species. Using genetic fingerprinting, it was determined that captive Komodo dragon (Varanus komodoensis) neonates were parthenogenetically derived (Watts et al., 2006). Further, these lizards can alternate between sexual and asexual reproduction depending upon the availability of a male. A Burmese python that was isolated from a male in a zoo produced fertile eggs in five consecutive years (Groot et al., 2003). Using molecular genetic methods, parental analysis revealed that this female reproduced parthogenetically. Other snakes that can shift between sexual and asexual reproduction are the Western terrestrial garter snake (Thamnophis elegans), checkered garter snake (T. marcianus), timber rattlesnake (Crotalus horridus), Aruba Island

18  Overview of Reptile Biology, Anatomy, and Histology

rattlesnake (C. unicolor) (Schuett et al., 1997), and Arafura file snake (Acrochordus arafurae) (Dubach et al., 1997).

1.4.6.7 Incubation Temperature and Sex Ratios  It has been shown for several species of chelonians, crocodilians, and lizards that sex ratios of neonates hatching from a clutch of eggs are temperature dependent. Although heteromorphic sex chromosomes have evolved independently in reptiles many times, there are numerous species of squamates and a few chelonians that lack these chromosomes. Thus, depending upon the clutch temperature, either all females or all males may be produced from a single clutch of eggs in those species lacking heteromorphic sex chromosomes (Vogt and Bull, 1982).

1.4.7 Cardiovascular System Many of the differences between the reptilian and amphibian circulatory systems are associated with loss of functional gills and the need for an efficient pulmonary circulation to and from the lungs. Whereas many amphibians meet oxygen needs by cutaneous and pharyngeal routes in addition to pulmonary exchange, in reptiles the pulmonary system is the main site of oxygen uptake. Thus, it is not surprising that the reptile lung is larger and more complicated than the amphibian lung. With this increased pulmonary complexity, the circulatory system underwent elaborations upon the amphibian design to accommodate this increased oxygen uptake and transport to tissues. The location of the heart in the coelomic cavity varies both within and between groups. The reptile heart is closely associated with the liver in chelonians, crocodilians, and lizards (Figures 1.9, 1.125–1.127, 1.129, 1.267). In some lizards such as the green iguana, the heart is located at the level of the forelimbs (Figure 1.127). In monitors it is more caudally located in the coelomic cavity. In all snakes the heart is located cranial to the liver. In most nonvenomous snakes the respiratory portion of the lung is situated between the heart and cranial pole of the liver (Figure 1.268) while in others the apex of the heart is near, or is in contact with, the cranial pole of the liver (Figure 1.269). In snakes, the location of the heart reflects its lifestyle (Lillywhite, 1987; Seymour 1987). In terrestrial and arboreal species the heart is approximately 15 to 25% of the total body length from the head, while in totally aquatic species it is approximately 25 to 45%. The reptile heart consists of the sinus venosus, right and left atrium, and single ventricle in all groups except crocodilians where there is a complete ventricular septum (White, 1959; White, 1976). The three caval veins enter the right side of the heart through the sinus venosus. A sinoatrial valve marks the opening of the sinus venosus into the right atria. The right and left auricles are completely separate in all reptiles (Figure 1.270), and as in higher vertebrates, oxygenated blood returning to the heart from the lungs flows through the pulmonary veins into the left auricle; deoxygenated blood

returning from systemic sites flows into the right auricle. The right and left atria are of equal size in some species while in others the right is larger than the left. A much larger right atria is commonly seen in snakes (Figure 1.269). All reptiles possess two aortae: a right and left (Figures 1.271–1.272). The right aorta originates from the left side of the heart and the left aorta originates from the right side of the heart. There are nearly all degrees of partitioning of the ventricle in living reptiles, varying from the situation (in some lizards) of having practically no interventricular septum to the situation (in crocodilians) of possessing a complete interventricular septum. The diversity of the reptile heart is a reflection of the diversity of reptiles as a group. Based upon the anatomy of the reptile heart (except crocodilians) several patterns of circulation of blood returning to the heart from the systemic and pulmonary circulation have been proposed: (1) complete mixing, (2) partial mixing, and (3) a high degree of separation between unoxygenated and oxygenated blood being distributed differentially to the two aortae. Additionally, factors such as temperature, respiratory status, and circuit resistances may alter the pattern within an individual animal. Investigative studies using radiographic techniques have yielded information from complete mixing in some species to a small right-to-left shunt directed to the left aortic arch in others. Studies measuring oxygen concentration of the atria and great vessels of snakes and lizards suggest that there may be a high degree of separation of blood entering the great vessels (White, 1959). A unique feature of reptile circulation is the ability to bypass the lungs while perfusing their systemic circulation (Farrell et al., 1998). The major vessels comprising the circulatory system of reptiles also show further modification on the amphibian plan. Reptiles are the first group of vertebrates to have evolved a well-developed coronary artery system. The left aorta is smaller than the right, and both join beyond the heart to form a common dorsal aorta. The esophagus of snakes passes through a ring (Figures 1.271–1.272) produced by the joining of the aorta, and specialized cardiovascular patterns in snakes must have evolved along with the ability to feed upon large-sized prey species. The reptile heart is composed of a lining endocardium, muscular, and connective tissue containing myocardium, and outer epicardium (Figure 1.273). The pericardial sac covers the heart. In some groups the apex of the heart is anchored to the pericardium by a ligamentous gubernaculum cordis while in others it is free floating. The endocardial lining is continuous with the endothelial lining of the great vessels at the base of the heart. The myocardium of the atria is thinner than that of the ventricle and the musculature of the left atria is thinner than the right. The ventricle is less compact than in mammals and birds, with muscular bundles organized in different directions and separated by spaces (Figure 1.273), and trabeculae and ridges extending into the lumen. This subdivides the ventricular cavity into three major chambers: cavum pulmonale, cavum venosum, and cavum arteriosum

Overview of Reptile Biology, Anatomy, and Histology  19

(Farrell et al., 1998). There is relatively little connective tissue in the ventricle. The outer myocardium is compact, and the inner myocardium is spongy.

1.4.8 Hemopoietic System The blood of reptiles consists of cellular and acellular components. The whole blood volume shows species variability with 7.3 ml/100 gm body weight for the American alligator and 9.1 m1/100 gm for the red-eared slider (Trachemys [formerly Pseudemys] scripta elegans) (Kaplan, 1974). The cellular components, comprising a packed cell volume of 20 to 40%, consist of erythrocytes, granulocytes, lymphocytes, monocytes, plasma cells, and thrombocytes. The acellular fraction of the blood, the plasma, comprising 60 to 80% of blood volume, is a colorless or straw-colored fluid in many species and a yellow, deeply pigmented fluid containing carotenoid pigments in others. Suspended within the plasma are a variety of inorganic electrolytes and a variety of organic compounds. Historically there is confusion in terminology with respect to cell types of reptilian blood and hemopoietic tissues. The inherent problems in naming cells according to staining abilities (basophil, eosinophil, neutrophil) and function (macrophage) have led to discrepancies in naming of cells among different investigators. Although no studies have completely documented the ontogenetic lineage of mature circulating blood cells, Pienaar (1962) in his monograph Hematology of Some South African Reptiles presents the most detailed picture. For information, also see Chapter 3. There is some variation with regard to hemopoietic centers both specifically (with age) and interspecifically. The centers of blood cell formation include the bone marrow, liver, and spleen. Although erythroid and granulocytic series are predominantly produced in bone marrow, the spleen also maintains a variable granulocytopoietic and erythropoietic activity. The myeloid stem cells are multipotential cells, giving rise to all the cell types found in the bone marrow. Mature erythrocytes are nucleated and oval. Immature erythrocytes are more rounded with a basophilic cytoplasm and a less chromophilic nucleus than that of the mature cell when stained with Romanowsky stains. Reticulocytes can be demonstrated by supravital staining procedures. Senile red cells are larger than mature cells, with pale staining cytoplasm and pyknotic nuclei. Dimensions for erythrocytes vary both interspecifically and intraspecifically. While the teiid lizard, Ameiva ameiva, has a mean least erythrocytic diameter of 7.6 µm, the tuatara, Sphenodon punctatus, has the largest red blood cells for a reptile, with a mean greatest diameter of 23.3 µm (Saint Girons, 1970a). Similarly, erythrocyte counts show a tremendous amount of variation both intraspecifically and interspecifically. Counts have been shown to vary with age, sex, season, altitude, nutritional status, and disease. Erythrocyte counting techniques have been described in detail elsewhere (Campbell, 2004; Chapter 3 this volume).

The white blood cell group consists of a variety of cells with unknown homology to higher vertebrate cell lines. The granulocytes are a complex group of cells, particularly the eosinophilic and azurophilic granulocytes. White blood cell morphology is reviewed and discussed in detail in Chapter 3 of this book.

1.4.9 Endocrine Organs 1.4.9.1 Pituitary Gland  The pituitary gland (hypophysis) consists of the neurohypophysis (median eminence, infundibular stalk, and infundibular process [or pars nervosa], which originates from the ventral portion of the diencephalons) and the adenohypophysis (pars intermedia, pars tuberalis, and pars distalis), which originates from the roof of the embryonic pharynx (Saint Girons, 1970b) (Figures 1.274–1.276). It is located in the sella turcica of the sphenoid bone of the ventral braincase, immediately caudal to the optic chiasma. At least nine hormones are produced, with some having local effects on the pituitary itself and also systemic effects on both endocrine and nonendocrine tissues. The pars distalis is composed of glandular cells arranged in cords or clumps. Glandular cells are classified as either chromophobic or chromophilic depending upon their affinity or lack of affinity for certain histologic stains. Historically, chromophilic cells were classified as either acidophilic or basophilic depending on their tinctorial properties of cytoplasmic granules with hematoxylin and eosin staining. More recently, immunohistochemical staining has been used to distinguish cells that produce specific hormones. Electron microscopy has also been used to distinguish between cell types based on size and morphology of cytoplasmic granules. The richly innervated pars intermedia is only a few cells wide and produces melanocyte-stimulating hormone as its main hormone. While the highly vascular pars tuberalis consists of epithelial cells, specific hormones produced by this structure have not been reported. The neurohypophysis consists primarily of axons of neurons, with cell bodies higher in the hypothalamus. The neurohypophysis serves as the major neuroendocrine regulatory center of the brain. Neurohypophyseal hormones and releasing hormones are produced and released by the hypothalamic neurons.

1.4.9.2 Thyroid Gland  In chelonians and snakes the thyroid gland is generally located ventral to the trachea and near the base of the heart (Figures 1.268, 1.271, 1.277–1.279, 4.24, 4.40). The tuatara has a single transversely elongate thyroid gland. In crocodilians it is bilobed, with an isthmus connecting the two lobes. In lizards it may be single, bilobed, or completely paired among different members of the same family (Lynn, 1970) (Figures 4.12–4.13). The functional unit of the thyroid gland is the thyroid follicle. Follicles produce and release thyroid hormones that are the same as in other

20  Overview of Reptile Biology, Anatomy, and Histology

vertebrates. Each follicle is separated by a single layer of epithelium cells, which show seasonal changes in the dimension of individual cells, extent of enclosed colloid material (containing thyroglobulin), presence of desquamated epithelial cells and blood cells, varying numbers of colloid droplets, and varying staining properties of the colloid. Within a single thyroid gland, follicles may either be of rather uniform dimensions or show a range in diameters (Figure 1.280). A capsule surrounds the entire gland and varying amounts of connective tissue separate adjacent follicles.

1.4.9.3 Parathyroid Gland  Reptiles have one or two pairs of parathyroid glands that may be either cranial or caudal to the thyroid (Clark, 1970). These glands are quite small and may be difficult to locate, even in large animals. Fat adjacent to the parathyroid gland may obscure their location. In chelonians, the cranial pair lies within the thymus while the caudal pair is near the aortic arch (Figure 1.281). In crocodilians one or two pairs may be present and are immediately anterior to the heart near the common carotid. Lizards may have one or two pairs of parathyroid glands; the cranial pair lies near the bifurcation of the common carotid, while the caudal pair is immediately posterior to the cranial pair (Figures 4.13–4.14). Snakes typically have two pairs of parathyroid glands; the caudal pair is between and often medial to the anterior and posterior lobes of the thymus. The cranial pair is at the bifurcation of the carotid artery and is often difficult to locate. The histology of the reptile parathyroid gland is similar to that of mammals. The glands are surrounded by connective tissue with fine strands dissecting through the gland. Epithelial cells are arranged in cords, clusters of cells, or follicles (Figures 1.282–1.283). Cysts are commonly seen in parathyroid glands of reptiles (Figures 1.284–2.285).

1.4.9.4 Ultimobranchial Body  The ultimobranchial body of reptiles is located anterior to the heart and near the thyroid and parathyroid glands (Clark, 1971). They are generally not grossly visible and are larger or only present on the left side of the neck. In many lizards, crocodilians, and some snakes, only the left gland is present. In the sand boa (Eryx johnii) the glands are paired and of equal size and are located midway between the rostral and caudal pairs of the parathyroid glands (Singh and Kar, 1983). In chelonians, both glands are present. Cells are arranged as cords, follicles, or clusters of cells, with the follicular epithelium containing both goblet and ciliated epithelial cells. Ultimobranchial gland follicles share morphologic features with thyroid follicles. Using an antiserum raised against salmon calcitonin, immunoreactivity was identified in clumps of cells in the ultimobranchial gland of the red-eared slider (Boudbid et al., 1987). Cells lining follicles did not stain. In a study with the Japanese grass lizard (Takydromus takydromoides) and the striped snake (Elaphe quadrivirgata), there was positive staining when antibody against pig calcitonin was used but not with antibody against synthesized human calcitonin (Yamada et al.,

2001). The mammalian equivalents of the ultimobranchial gland are the C cells, which are dispersed in the thyroid, and at least two additional types of calcitonin have been identified in mammals. The ultimobranchial gland of the green iguana also consists of clumps of cells and epithelial-lined follicles (Figure 1.286). Using antisalmon calcitonin antibody, immunoreactivity also has been seen in the clumped cells (Figure 1.287).

1.4.9.5 Adrenal Gland  All reptiles have a pair of adrenal glands that range in color from light yellow to pink to red (Gabe, 1970). Those of turtles are flattened dorsoventrally, and found within, against, or near the cranial poles of the kidneys (Figures 1.186, 1.206, 1.288). In crocodilians the adrenal gland is retroperitoneal, lying dorsal and lateral to the gonads and genital ducts (Figures 1.289). Squamates have their adrenals incorporated into the mesorchium of the male (Figures 1.208–1.209) and mesovarium of the female (Figures 1.235, 1.290); in snakes the adrenals are elongated (Figures 1.291–1.292). The adrenal gland of reptiles is composed of both chromaffin and interrenal cells; chromaffin tissue may be scattered throughout the interrenal tissue or may be at the periphery of the gland (Figures 1.293– 1.296). Chromaffin tissue of the reptile adrenal represents the adrenal medulla of mammals and contains cells producing catecholamines. Using hematoxylin and eosin staining, these cells have numerous basophilic-staining granules and appear to be arranged in clusters (Figures 1.295–1.296). In contrast, the interrenal component of the adrenal corresponds to the adrenal cortex of mammals and contains foamy corticoid containing pale staining cells that are arranged in cords (Figures 1.295–1.296).

1.4.9.6 Pancreatic Islets of Langerhans  See information on the pancreas in Section 1.4.3 (Digestive System) of this chapter.

1.4.10 Nervous System Different terminology has been used to separate the brain into various divisions. From rostral to caudal, the main divisions of the reptilian brain are the prosencephalon or forebrain (olfactory bulbs and tracts), cerebrum (telencephalon), diencephalon (rostral epithalmus, dorsal thalamus, ventral thalamus, and hypothalamus; caudal pretectum and posterior tuberculum), mesencephalon or midbrain (dorsal sensory optic tectum and torus semicircularis and ventral motor tegmentum), and rhombencephalon (hindbrain). These subdivisions can be distinguished grossly (Figures 1.297–1.301) and microscopically (Figures 1.302–1.322). Traditionally, 12 cranial nerves have been described, with each designated by Roman numerals I through XII. The olfactory nerve(s) (cranial nerve I) travel from olfactory receptors in the nasal cavity to synapse with mitral cells in the olfactory bulbs (Figures 1.303–1.304). Olfactory tracts project from the

Overview of Reptile Biology, Anatomy, and Histology 21

olfactory bulb to olfactory cortex of the telencephalon (Figures 1.302, 1.306, 1.307). In squamates, a parallel vomeronasal tract travels from the vomeronasal organ (see below) to the accessory olfactory bulb of the brain. The nasal septum is innervated by the terminal nerve (nervus terminalis), a small nerve having fibers that contain gonadotropin- releasing hormone. While in most reptiles the neocortex is lacking, in chelonians there is some evidence that the dorsal cortex is homologous to the neocortex of mammals (Belekhova, 1979). The cytoarchitecture of the telencephalon has been studied in a variety of reptiles (Ulinski, 1990) (Figures 1.309–1.312). The rhombencephalon is that part of the brain above the notochord and includes the metencepahlon (pons and cerebellum) (Figures 1.315, 1.316, 1.319, 1.320) and myelencephalon (medulla oblongata) (Figures 1.321–1.322). The cerebellum is an outgrowth of the rhombencephalon and shows considerable variation in anatomy and histology among various groups of reptiles. The visual system consists of the eyes and projections of the optic nerves (cranial nerve II) onto the mesencephalic tectum (Figures 1.313–1.314, 1.317–1.318). Ramón (1896) described 14 layers of the optic tectum for the lizard Lacerta. Other naming schemes followed (Huber and Crosby, 1933; Northcutt, 1984). Additional sensory systems associated with the brainstem are the acoustic system and vestibular (cochlear) system. The ventral root of nerve VIII is considered acoustic and the dorsal root is considered vestibular. In most reptiles the projections are bilateral. The medulla oblongata fuses ventrally with the tegmentum, and rostrally and caudally it grades respectively into the mesencephalon and the spinal cord. The brainstem consists of the mesencephalon and rhombencephalon. The spinal cord is surrounded by the vertebral column and in chelonians it is within the dermal bone of the carapace. As in other vertebrates, it consists of an exterior zone of acellular white matter and an interior zone of cellular gray matter. The gray matter consists of dorsal and ventral horns (Figures 1.323–1.325). Meninges consisting of the endomeninx and ectomenix surround both the brain and spinal cord. Melanophores are common in the meninges of reptiles. Phylogenetically, reptiles are the first vertebrate group to possess 12 pairs of cranial nerves. However, snakes lack a spinal accessory nerve (XI). The relative position and size of the sense organs (olfactory, optic, and otic and vestibular organs) exert the primary effects on brain shape. In-depth reviews and original descriptions can be found elsewhere (Balaban and Ulinski, 1981; Butler and Hodos, 2005; Butler and Northcutt, 1973; Cruce, 1974; Cruce and Cruce, 1975; Ebbesson and Voneida, 1969; Gans et al., 1979a, 1979b; Gans and Ulinski, 1992; Halpern, 1980; Schecter and Ulinski, 1979; Sereno, 1985; Sereno and Ulinski, 1985; Ulinski 1974; Ulinski, 1990). Often included with discussions of the brain of reptiles is the parietal-pineal complex. These morphologically and functionally interrelated structures are located on the roof of the diencephalon. Chelonians, some lizards (41%

of examined lizards lack a parietal eye), and snakes have the pineal alone, and many lizards and the tuatara have both (Quay, 1979). The complex is absent in crocodilians. In chelonians the pineal gland is composed of thick-walled epithelial vesicles that communicate with each other and the third ventricle of the brain, or is a thick-walled epithelial sac that is closed off from the third ventricle and has a solid stalk containing nerve fibers. In lizards having a parietal eye–pineal complex, the epidermis within the interparietal scale, which is over the parietal eye, is modified into a cornea (Figure 1.326). The parietal foramen serves as an opening in the skull to the parietal eye or its tracts and may be located in the parietal bone, frontal bone, or near the frontoparietal suture. In the green iguana it is within the caudal frontal bone and adjacent to the suture with the parietal bone (Figure 1.62). The parietal eye is usually dorsal to the telencephalon and cranial to the pineal gland (Figures 1.327–1.328). In lizards it may be within the overlying bone or extracranial. The parietal eye is organized into a single epithelial vesicle with a dorsal lens and ventral retina consisting of sensory photoreceptor cells, supportive, and ganglionic cells (Figure 1.329). The pineal organ (epiphysis) is generally located posterior to the parietal eye and above the diencephalon (Figures 1.327, 1.330). Anterior to the pineal and extending from the roof of the diencephalon are the dorsal sac and paraphysis (Figure 1.330). In the tuatara the parietal eye is located in the lower portion of the parietal foramen of the skull (Figure 1.57). As with lizards, it consists of an outer epithelial lens and an inner retina (Ung and Molteno, 2004) (Figures 1.331–1.332). The pineal organ of snakes is somewhat more compact than that of lizards (Figure 1.333). A stalk is present that connects the gland with the brain. Chemicals identified in the parietal–pineal complex include catecholamines, 5hydroxytryptzmine (serotonin), and melatonin. The parietal eye–pineal complex may affect behavior, gonadal activity, thermoregulation, and color change.

1.4.11 Eye In chelonians, crocodilians, lizards, and tuataras, there is general anatomic similarity of the eyes and adnexa, with minor variations between the groups (Underwood 1970; Walls, 1942). The eye of snakes is distinct, having many embryologic and structural differences with the eyes of other reptiles. This is thought to reflect their evolutionary development when they existed as fossorial, burrowing animals with eyes reduced to small vestigial and functionally unimportant sense organs. This can still be seen in the fossorial blind snakes (Typhlops). Following their reinvasion of terrestrial habitats, the first snakes redeveloped their ocular anatomy so that what is seen today has changed considerably in structure and function from their immediate ancestors. Each group will be briefly reviewed with lizards serving as the basic plan for all groups except snakes.

22  Overview of Reptile Biology, Anatomy, and Histology

1.4.11.1 Sauria  The eyelids are well developed in most families of lizards, with the lower lid more moveable than the upper. Most species have a third eyelid. In some lizards of the families Lacertidae, Teiidae, and Scincidae, certain scales of the lower eyelid have become transparent, enabling limited vision even when the lids are closed. This modification reaches its extreme form in Pygopodidae, Dibamidae, certain members of the family Gekkonidae, and some members of the Scincidae in which the two lids are fused to form a clear spectacle (Figure 1.334). The spectacle is separated from the cornea by the subspectacular space, which is lined in part with corneal and conjunctival epithelium. Members of the family Eublepharidae, the eyelid geckoes, lack spectacles (Figure 1.335). A Harderian gland is located ventromedially and a lacrimal gland dorsotemporally to the globe, although the latter gland is lacking in geckos and chameleons. The nasolacrimal duct drains from the medial canthus and enters the roof of the mouth, either just behind or at the base of the duct of the vomeronasal organ. The orbits are separated in the sagittal plane by a thin cartilage septum, which would provide little resistance to infection or neoplasia on either side. The sclera is thin and supported by hyaline cartilage extending from the posterior pole to the equator. Cranial to this is a ring of 14 scleral ossicles, which provide shape for the anterior segment of the globe and leverage for the ciliary muscle at the edge of the cornea. The lens is soft and has a thickened pad of epithelial cells at the equator, which abuts on the ciliary body. During accommodation for near vision, contraction of the ciliary muscle causes forward and inward movement of the ciliary body, which compresses the lens to reduce its focal length, a method shared with birds. The striated transversalis muscle, which arises in the closed fetal fissure ventrally and inserts via a ligament onto the undersurface of the lens, rotates the latter nasally to affect convergence in most species during accommodation. The iris has a welldeveloped sphincter of striated muscle, giving considerable pupillary movement, which can be both rapid in response to light and modified by voluntary control. The striated muscle of the iris, as in birds, makes the reptile eye unresponsive to the conventional mydriatic drugs used in mammals. The pupil shape varies from a vertical ellipse in nocturnal forms (Figures 1.334–1.335) to round in most diurnal species (Figure 1.336). The iridocorneal angle has some similarities to that in mammals, although it is less well developed in reptiles. Ciliary processes are absent in lizards. The retina is avascular, being nourished from the choroidal vessels and the conus papillaris, a vascular projection from the optic nerve head projecting into the vitreous, even as far as the posterior lens capsule in some species. This is the reptilian equivalent of the avian pecten. The retina contains rods and cones, reflecting in proportions the nocturnal or diurnal habits of the various species. In some of the burrowing, legless lizards, the eyes have become vestigial structures.

1.4.11.2 Chelonia  The adnexal structures include welldeveloped eyelids and large Harderian and lacrimal glands. In marine turtles, the lacrimal glands are especially large; they function as extrarenal sites of salt secretion (Figures 1.337–1.338). Chelonians do not have a nasolacrimal duct, the tears being lost by evaporation, absorption across the conjunctiva, or overflowing from the conjunctival sac. Scleral ossicles vary in numbers between species. The pupil is round (Figures 1.339), the retina is avascular, and cones are the predominant photoreceptor type.

1.4.11.3 Crocodylia  Crocodilian eyes are adapted to nocturnal vision in a semiaquatic environment. The eyelids are well developed, with the upper eyelid containing a bony tarsus that is capable of powerful closure. The third eyelid (membrana nictitans) (Figure 1.7), which is fairly transparent in young animals, can cover the globe while the lids remain open. This makes eye examination of crocodilians extremely difficult. Accessory glands in the conjunctiva augment secretions of the Harderian and lacrimal glands. Scleral ossicles are absent, although this deficit is made up by the more anterior extension of the scleral cartilage to the ora serrata. The pupil is elliptical (Figure 1.340) and responsive to light, contracting down to a small vertical slit. The retina is avascular, although small capillary loops can be seen on the optic nerve head by ophthalmoscopy. The primary visual cells are rods. A retinal tapetum is formed by the accumulation of guanine crystals in the retinal pigment epithelial cells.

1.4.11.4 Ophidia  The eyes of all snakes are covered by a transparent spectacle (Figure 1.341), formed embryologically by the fusion of the eyelids. In all reptiles with a spectacle, the spectacle has been demonstrated by microsilicone injections to be highly vascular (Mead, 1976). This is significant when differentiating normal vascular patterns in the spectacle from neovascularization of the spectacle or cornea. The spectacle is shed along with the rest of the skin at each cycle of ecdysis (Figures 1.342–1.343). Snakes have a well-developed Harderian gland, but no lacrimal gland. The nasolacrimal duct leaves the subspectacular space to enter the mouth as in lizards. The duct opens adjacent to the vomeronasal organ, in the cranial roof of the oral cavity. Thus, the subspectacular space communicates directly with the mouth. There is no cartilage in the sclera of snakes, unlike all other reptiles. The iris sphincter is formed by striated muscle that is rapidly responsive to light. Snakes have either round (Figure 1.344) or elliptical (Figure 1.345) pupils. Consensual light reflex cannot be elicited. Focal accommodation in snakes is accomplished by forward movement of the lens due to pressure exerted on the vitreous by the ciliary muscle. The retina is avascular, being supplied from the choroid by a branching array of vessels, the membrana vasculosa retinae, which run in the posterior vitreous before leaving the eye at the optic nerve. Blood flow in these vessels is from the periphery of

Overview of Reptile Biology, Anatomy, and Histology 23

the retina toward the optic nerve. Rods and cones vary in types and proportions, depending on the species.

1.4.12 Ear The reptile ear is located caudal and ventral to the eye. As in other animals, it is involved in both reception of sound and equilibrium. The extent of development varies between the different groups of reptiles. The three major subdivisions are the external ear (short recess from lateral surface of head to tympanic membrane), middle ear (tympanic membrane and structures for transmission of sound), and inner ear (otic capsule). A brief review of the major components of the ear is given below. Of reptiles, the lizard ear has been studied the best and a “generic” lizard ear will be described and used to make comparisons with other orders of reptiles. However, not all lizards fit this pattern. For detailed descriptions, see Baird (1970) and Wever (1978).

1.4.12.1 Sauria  In most lizards the external ear consists of a slight depression, with scales lining the recess up to the tympanic membrane, which is devoid of scales. In some burrowing forms the external ear canal, tympanic membrane, and tympanic cavity are either vestigial or are missing. The tympanic cavity, the major component of the middle ear, is lined with squamous epithelium on its exterior and a mucous membrane on its interior surface consisting of squamous, cuboidal, and smaller numbers of mucous epithelial cells. Cells lining the middle ear are continuous with those lining the auditory (Eustachian) tube. The middle ear is bounded dorsally by the quadrate bone and paraoccipital process and ventrally by the retroarticular process. The columella consists of the extrastapes (cartilage) and stapes (bone). The stapes traverses the middle ear and eventually expands as an opening in the otic capsule of the inner ear. At its attachment to the oval window, the stapes expands to form a discoidal footplate that terminates on the oval window of the otic capsule. The otic capsule consists of interconnected bony cavities and canals containing fluid-filled channels and sacs (membranous labyrinth) organized into otic and periotic labyrinths (Figure 1.346). The major component of the otic cavity is the vestibule. Anteriorly and posteriorly the vestibule communicates with the anterior and posterior osseus ampulla. The anterior semicircular canals originate from the anterior ampulla, arches dorsally, and ultimately joins the posterior semicircular canals, which communicate with the posterior ampulla. Lateral semicircular canals originate from the lateral ampulla. The major chambers within the otic labyrinth are the utricle, saccule, and cochlear duct. The utricle is medially located and is an elongate arched tube. The macula is the major sensory epithelium of this structure and functions as an equilibrium receptor. Cells within this receptor are distinguishable from the squamous epithelial cells that line other

areas of the utricle. The macula consists of tall columnar cells and sensory cilia; it is innervated by a branch of the vestibulocochlear nerve. A smaller structure, also consisting of sensory epithelium, is the macula neglecta. Continuous with the utricle are three semicircular ducts and their associated ampulla. The ducts pass through their corresponding semicircular canals. Each otic ampulla has sensory epithelium and receives branches from the vestibulocochlear nerve. These receptors are also involved with equilibrium. A large saccule is medially located, lateral to the utricle, and is surrounded by the semicircular ducts. The saccule and utricle communicate by an utriculosaccular duct. As with the utricle, the saccule contains a sensory epithelium. In reptiles, the lagena (a chamber projecting from the saccule) elongates to form the cochlear duct, which is an extension of the posterior aspect of the saccule. It contains specialized receptors (papilla basilaris and tectorial membrane) consisting of columnar hair cells and supporting cells; a branch of the vestibulocochlear nerve supplies them. The papilla basilaris is the receptor organ for hearing.

1.4.12.2 Ophidia  Snakes lack an external ear, tympanic membrane, tympanic cavity, and auditory canals. The columella apparatus articulates with the quadrate. The quadrate has a lose attachment between the lower jaw and dorsolateral skull. The quadrate bone acts as a receiving surface for sound waves transmitted to the columella. The stapes has a footplate that is relatively larger than that of terrestrial lizards. The otic labyrinth (Figure 1.347) has all previously mentioned components seen in lizards. The utricle and semicircular ducts are the largest part of the otic labyrinth in snakes. 1.4.12.3 Tuatara  Tuataras lack an external ear, the tympanic membrane is degenerate, and there is no tympanic cavity. The inner ear shows the least specialization of all reptiles. 1.4.12.4 Chelonia  In chelonians, the external ear is lacking and the tympanic membrane is thickened in terrestrial species such as tortoises (Figure 1.4) and thinner in aquatic species. The quadrate is ventrally elongate and divides the middle ear into a lateral tympanic cavity and a medial recessus cavi tympani. The medial part of the tympanic cavity contains a unique specialized fluid-filled structure, the paracapsular sinus.

1.4.12.5 Crocodylia  Crocodilians have a well-developed, shallow external ear. Two folds of skin overlay the opening, obscuring the tympanic membrane from visualization (Figure 1.8). Contraction of muscles in the dorsal fold firmly brings it into apposition to the ventral fold. Compared to other reptiles, the tympanic cavity is more firmly surrounded by bone, resulting in a greater compartmentalization. The two Eustachian tubes open into the pharynx in close proximity to the tonsil.

24  Overview of Reptile Biology, Anatomy, and Histology

1.4.13 Vomeronasal Organ The vomeronasal organ (VNO), which is also called Jacobson’s organ, is a chemoreceptive structure located in either the roof of the mouth or associated with the choanae of certain reptiles (Figure 1.348). The nasolacrimal ducts open either within or adjacent to the VNO duct. The VNO is absent in adult crocodilians and present in tuataras, lizards, and snakes. In some lizards such as chameleons, the VNO is vestigial. In chelonians, there is not a clear consensus about the presence or absence of the VNO. Some authors believe that a homologue of the VNO is present in the chelonian nose (Parsons, 1970). In lizards and snakes, the VNO is separated from the nose by the extension of the palate. It is best developed in varanid lizards and snakes that use their tongue to mechanically pick up particles in the air and deposit them on the duct of the VNO. The dorsal and lateral walls of the VNO are lined with a sensory epithelium and a branch of the olfactory nerve innervates it. A structure called the mushroom body invaginates from the wall of the VNO into its center (Figure 1.349).

1.4.14 Salt Glands Because reptiles cannot concentrate urine hypersomotic to blood, numerous species have evolved extrarenal sites of salt secretion as a homeostatic mechanism. This is particularly true of marine species (sea turtles, sea snakes, marine crocodiles) and certain desert species (chuckawalla [Sauromallus spp.], desert iguana [Dipsosaurus dorsalis.]). These glands serve as the major route of electrolyte excretion in those species possessing them. They have evolved independently among the reptiles at least five times and represent nonhomologous structures between the different groups (Dunson, 1976). In the diamondback terrapin (Malaclemys terrapin) and sea turtles, the lacrimal gland in the posterior orbit has been modified into a salt gland (Figure 1.337). While numerous ducts drain the gland in diamondback terrapins and open individually along the lower lid, all the lobular ducts of sea turtles join into a single duct (Figure 1.338). In those lizards (Amblyrhynchus, Iguana, Dipsosaurus, Sauromalus, Uromastyx, and Ctenosaura) with salt glands, it is lateral-nasal in location. The individual tubules empty into a common duct that discharges its contents into the nasal cavity. In sea snakes the location is sublingual, and in the saltwater crocodile it is in the tongue. While light microscopy cannot determine which of the various cephalic glands are salt-secreting glands, they can be identified using electron microscopy. The salt-secret-

ing cells (principal cells) of different reptile salt glands have similar ultrastructure. They have numerous mitochondria and the cytoplasmic membrane is thrown into numerous lateral projections (microvilli) that loosely interdigitate between adjacent cells. Shorter microvilli are on the apical surface, projecting into the tubular lumen.

1.4.15 Infrared Detection Organs Infrared receptors (pit organs) have evolved in the snake families Boidae (boas) and Pythonidae (pythons), and in the viperid subfamily Crotalinae (pit vipers). The shape, number, and location of infrared detectors in boas and pythons differ from those of crotalids. In crotalids, the pit organ is a cavity in the skin adjacent to the external nares (Figure 1.350). The pit receptor consists of free nerve endings within a concave sensory membrane that is suspended within the cavity (Figure 1.351). In boas and pythons, the receptors are found in multiple supra- and infralabial scales. In some boas, such as the boa constrictor (Boa constrictor), there are no specializations, with receptors within individual scales. In other boas, such as the emerald tree boa (Corallus caninus), the receptors are located in infralabial and supralabial pits (Figure 1.352). In contrast to crotalids, the receptors are not contained in a suspended membrane, but instead are located at the bottom of the labial pits. The pit organs are supplied by branches of the trigeminal nerve. For more detailed information, see Barrett (1970) and Molenaar (1992).

Acknowledgments The author thanks Mark Hoffenberg for photographing many of the skeletal preparations seen in this chapter. Thanks also to the Florida Museum of Natural History for allowing me to photograph many of the skulls used in this chapter. Jeanette Wyneken, Michael M. Garner, and Brian Stacy graciously reviewed this chapter. Paul Maderson made helpful comments regarding the identification of histological components of the skin of snakes presented in this chapter. Randall Morrison helped identify pigment cells. Others who graciously provided me with images are recognized in the figure legends. Bill Brant, Eugene Bessette, Stephen Hernandez-Divers, and Kenney Krysko provided reptiles used for anatomical and histological preparations. Many images were taken of client-owned animals evaluated by the Zoological Medicine Service, College of Veterinary Medicine, University of Florida at Gainesville.

Overview of Reptile Biology, Anatomy, and Histology 25

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30  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.1   Australian snakeneck turtle, Chelodina longicollis. Chelidae. The head of this chelid turtle cannot be withdrawn into the margins of the shell. Instead it is positioned laterally and below the anterior scutes. Courtesy of Darryl Heard.

Figure 1.2   Columbian slider, Trachemys scripta callirostris. Emydidae. This emydine turtle can retract its head within the margins of the shell. Courtesy of John Behler.

Figure 1.3   Red-eared slider, Trachemys scripta elegans. Emydidae. Ventrodorsal view of a skeletal preparation. The ribs are within the carapace and the pectoral girdle is internal to the ribs. The triradiate pectoral girdle consists of the acromion process (AC) of the scapula, scapula (S), and procoracoid (PC). Also seen are the cervical vertebrae (CV), humerus (H), radius (RA), and ulna (UL).

Figure 1.4   Desert tortoise, Gopherus agassizii. Testudinidae. Lateral view of the head. The tympanic membrane (TM) is covered by a modified epidermis.

Overview of Reptile Biology, Anatomy, and Histology 31

Figure 1.5   American alligator, Alligator mississippiensis. Alligatoridae. A complete hard palate can be seen forming the roof of the oral cavity. A dorsal flap and ventral flap together form the gular valve that separates the oral and pharyngeal cavities. Courtesy of Darryl Heard.

Figure 1.6   American alligator, Alligator mississippiensis. Alligatoridae. The edge of the nictitans (arrow) can be seen covering the anterior margins of the cornea. Courtesy of Darryl Heard.

Figure 1.7   Saltwater crocodile, Crocodylus porosus. Crocodylidae. The crocodile is underwater and the nictitans is covering the globe. Blood vessels are seen in the nictitans. Courtesy of Darryl Heard.

32  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.8   American crocodile. Crocodylus acutus. Crocodylidae. The ear is located behind the eye and specialized flaps of skin (arrows) surround the opening to a shallow ear canal.

Figure 1.9   American alligator, Alligator mississippiensis. Alligatoridae. A pseudodiaphragm (margins indicated by arrows) separates the liver (left liver lobe [LL], right liver lobe [RL]) and heart (HT) from a specialized coelomic fat body (FB), stomach (ST), and intestinal tract (IN).

Figure 1.10   American alligator, Alligator mississippiensis. Alligatoridae. A unique fat body is exteriorized from the coelomic cavity. The fat body is located on the right side of the coelomic cavity, directly behind the pseudodiaphragm, which separates the fat body from the right lobe of the liver.

Overview of Reptile Biology, Anatomy, and Histology 33

Figure 1.11   Tuatara, Sphenodon punctatus. Sphenodontidae. No tympanic membrane is present behind the eye. Courtesy of Ronald Goellner.

Figure 1.12   Green iguana, Iguana iguana. Iguanidae. The tympanic membrane (arrows) is located caudal to the angle of the jaws.

Figure 1.13   Five-toed worm lizard, Bipes biporus. Bipedidae. This worm lizard is from Baja, Mexico. It has reduced eyes and the forelimbs are designed for digging. Courtesy of Theodore Papenfuss.

Figure 1.14   Florida worm lizard, Rhineura floridana. Rhineuridae. This worm lizard is from central Florida, in the U.S. Scales encircle the body, and there are no eyes or limbs. Courtesy of Kenny Krysko.

Figure 1.15   Corn snake, Elaphe guttata guttata. Colubridae. Head of a corn snake. Snakes lack tympanic membranes and ear cavities.

34  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.16  Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of the hinge-region (arrow) between adjacent scutes. H&E stain.

Figure 1.17  Radiated tortoise, Geochelone radiata. Testudinidae. The surface of the shell is covered by multiple scutes having β-keratin. Embryonic shields are the yellow structures within the central portion of each scute.

Figure 1.18  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of a scute overlying dermis. The keratin (K) contains melanosomes and overlies a pseudostratified to stratified epidermis (E). Below the epidermis is the dermis (D) consisting of collagen and blood vessels. H&E stain.

Figure 1.19  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of a scute overlying dermis. The keratin (K) stains yellow and epidermal cells (E) stains red. Within the dermis (D), there is green-staining collagen. The black particles in the keratin are melanosomes. Masson’s trichrome stain.

Overview of Reptile Biology, Anatomy, and Histology 35

Figure 1.20  Desert tortoise, Gopherus agassizii. Testudinidae. Higher magnification photomicrograph of Figure 1.19. The basal layer of the epidermis has foot-like processes (arrows) that interdigitate with the collagen in the dermis. Masson’s trichrome stain.

Figure 1.21  Leopard tortoise, Geochelone pardalis. Testudinidae. The scutes of neonate tortoises consist of embryonic shields (arrows). As a tortoise grows, new keratin develops at the seams between adjacent scutes. New growth eventually forms rings around each embryonic shield.

Figure 1.22  Leopard tortoise, Geochelone pardalis. Testudinidae. In the plastron of this adult growing tortoise, recently formed keratin (arrows) is seen at seams between adjacent scutes. New keratin is lighter in coloration than older keratin. The embryonic shield (ES) is the oldest portion of each scute. Keratin is not symmetrically deposited around each embryonic shield.

Figure 1.23  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the seam between adjacent scutes. The epidermis is seen invaginating into the dermis. Masson’s trichrome stain.

36  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.24  Indian star tortoise, Geochelone elegans. Testudinidae. The shell consists of epidermal scutes overlying dermal bone. Courtesy of Darryl Heard.

Figure 1.25  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the shell below the epidermis. Dermal bone (DB) is seen between outer (OCT) and inner (ICT) connective tissue. Below the inner connective tissue is skeletal muscle (M). Masson’s trichrome stain.

Figure 1.26.  Gila monster, Heloderma suspectum. Helodermatidae. Certain lizards, such as gila monsters and beaded lizards, shed in patches and the entire old skin is sloughed over a period of weeks.

Figure 1.27  Boa constrictor, Boa constrictor. Boidae. Resting stage of a cycle of ecdysis. The skin is bright in coloration and the spectacle is clear.

Overview of Reptile Biology, Anatomy, and Histology 37

Figure 1.28  Boa constrictor, Boa constrictor. Boidae. As a snake enters the earliest stage of renewal the skin and eyes become dull and develop a light bluish tinge.

Figure 1.29  Boa constrictor, Boa constrictor. Boidae. During the mid-stages of a cycle of renewal, the skin and spectacle develop a deep bluish tinge.

Figure 1.30  Boa constrictor, Boa constrictor. Boidae. Several days prior to shedding, the color of the skin and spectacle lose the bluish tinge and become more normal in coloration.

Figure 1.31  Boa constrictor, Boa constrictor. Boidae. As shedding occurs, a new cycle of ecdysis begins. The old spectacle (arrows) is shed along with the skin. Over the next few days the new epidermal generation of skin will continue to mature.

Figure 1.32  Boa constrictor, Boa constrictor. Boidae. As a snake crawls, the old shed skin inverts.

38  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.33  European grass snake, Natrix natrix. Colubridae. Schematic representation of the histological changes in a single shedding cycle. In A the 6 stages of the cycle are listed from left to right. In B the histological changes are listed for the 6 stages. The time intervals for the 6 stages do not reflect actual times for each stage. For instance, the resting stage makes up approximately three-fourths of a complete cycle. The following layers and stages are seen: α-layer of inner generation (AI); α-layer of outer generation (AO); β-layer of inner generation (BI); β-layer of outer generation (BO); clear layer (CL); completion of outer generation (COG); inner generation (IG); immediate post-shedding period (IPS); lacunar tissue (LT); mesos layer of inner generation (MI); mesos layer of outer generation (MO); inner oberhautchen (OBI); outer oberhautchen (OBO); outer generation (OG); perfect resting condition (PRC); stratum basale (SB); stratum germinativum (SG). Courtesy of Lukas Landmann. (From Landmann L. 1979. J Morphol 162:93-126. With permission.)

Figure 1.34  Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of the skin during the resting stage. The following layers are seen: α-layer (AO); β-layer (BO); dermis (D); mesos layer (MO); oberhautchen (OBO); stratum germinativum (SG). H&E stain.

Figure 1.35  Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of the skin during the resting stage consisting of outer and inner regions of a scale. The inner region of a scale is continuous with the hinge region (see Figure 1.16). The β-layer (BO) is absent from the inner region of the scale and the mesoslayer (MO) is easier to visualize in the inner region compared to the outer region. The mesos layer is a barrier to water loss. The oberhautchen (OBO), while not distinguishable on a light microscopic level, covers the mesos-layer in the inner region and the BO in the outer region of a scale. The oberhautchen is the only β-keratogenic tissue present in the inner scale surface. The following layers are also seen: α-layer of outer generation (AO); immature cells (IM) that may be incorporated into the α-layer; stratum germinativum (SG). H&E stain.

Overview of Reptile Biology, Anatomy, and Histology 39

Figure 1.36  Dumeril’s ground boa, Acrantophis dumerili. Boidae. Scanning electron photomicrograph of the oberhautchen surface of a biopsied scale. Spinulae (arrows) and pits can be seen.

Figure 1.37  Dumeril’s ground boa, Acrantophis dumerili. Boidae. Higher magnification scanning electron photomicrograph of Figure 1.36. The oberhautchen surface has rows of spinulae (arrows) that surround “bare” areas covered by shallow pits.

Figure 1.38  Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a biopsied scale at early stage 4 of epidermal renewal. This is approximately halfway through the renewal phase. The following layers are seen: α-layer of outer generation (AO); β-layer of inner generation (BI); β-layer of outer generation (BO); clear layer of outer generation (CLO); lacunar tissue of the outer generation (LTO); mesos layer of outer generation (MO); inner oberhautchen (OBI); outer oberhautchen (OBO); stratum germinativum (SG). The immature (IM) cells will either become the deepest component of the β-layer or the most superficial components of the mesos-layer. H&E stain.

40  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.39  Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a biopsied scale at late stage 4 of epidermal renewal. The following layers are seen: α-layer of outer generation (AO); β-layer of inner generation (BI); β-layer of outer generation (BO); clear layer of outer generation (CLO); dermis (D); lacunar tissue of the outer generation (LTO); mesos layer of outer generation (MO); inner oberhautchen (OBI); outer oberhautchen (OBO); stratum germinativum (SG). The immature (IM) cells will either become the deepest component of the β-layer or the most superficial components of the mesos-layer. Heterophils (arrows) are seen within the lacunar tissue layer. H&E stain.

Figure 1.40  Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a biopsied scale at stage 5 of epidermal renewal, approximately 2 days prior to shedding. The outer generation is artifactually separated from the inner generation due to biopsy and processing.The following layers are seen: α-layer of outer generation (AO); α-layer of inner generation (AI); β-layer of inner generation (BI); β-layer of outer generation (BO); clear layer of outer generation (CLO); dermis (D); lacunar tissue of the outer generation (LTO); mesos layer of outer generation (MO); inner oberhautchen (OBI); outer oberhautchen (OBO); presumptive α-cells of the inner generation (PAI); stratum germinativum (SG). H&E stain.

Figure 1.41  Desert tortoise, Gopherus agassizii. Testudinidae. Mental (chin) glands (arrows) are seen medial to the mandibles. Courtesy of John Roberts.

Overview of Reptile Biology, Anatomy, and Histology 41

Figure 1.42  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the secretory portion of a mental (chin) gland. This is a holocrine gland since the secretion results from disintegration of its own cells. During periods of activity, new cells form at the base. The cells are cuboidal to polyhedral and the cytoplasm is lightly basophilic. Toward the surface, the cells become vacuolated. H&E stain.

Figure 1.43   Green iguana, Iguana iguana. Iguanidae. Femoral pores of a male produce a thick secretion that may help in grasping females during copulation. (From Jacobson ER. 2003. Biology, Husbandry, and Medicine of the Green Iguana. Krieger Publishing Company, Malabar, FL. With permission.)

Figure 1.44   Green iguana, Iguana iguana. Iguanidae. Femoral pores of a female are considerably smaller than males. (From Jacobson ER, 2003. Biology, Husbandry, and Medicine of the Green Iguana. Krieger Publishing Company, Malabar, FL. With permission.)

42  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.45   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a femoral pore. Eosinophilic staining material is secreted through the femoral pore. This is a holocrine gland because the secretion results from destruction of entire cells. H&E stain.

Figure 1.46   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a femoral pore. At the base of the gland the cells are cuboidal and have intracytoplasmic granules of eosinophilic material. With degeneration of the cells, the material coalesces and is secreted through the pore. H&E stain.

Overview of Reptile Biology, Anatomy, and Histology 43

Figure 1.47   Leopard gecko, Eublepharis macularius. Eublepharidae. A male lizard has well-developed precloacal pores (arrows) arranged in a chevron.

Figure 1.48   Leopard gecko, Eublepharis macularius. Eublepharidae. Female leopard geckoes do not have precloacal pores.

Figure 1.49   Brown anole, Anolis sagrei. Iguanidae. Photomicrograph of the skin. Two types of pigment cells are seen in the dermis. Melanophores (ME) are the dark pigment cells containing melanin, and the pigment cells in the superficial dermis having a clear cytoplasm are xanthophores (XA). In the epidermis the α-layer (AO), β-layer (BO), mesos layer (MO), and stratum germinativum (SG) are seen. H&E stain.

Figure 1.50   Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Photomicrograph of the skin. Several types of pigment cells are seen in the dermis. H&E stain.

44  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.51   Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Higher magnification photomicrograph of the skin in Figure 1.50. The epidermis consists of an α-layer (AO), βlayer (BO), mesos layer (MO), and stratum germinativum (SG). Starting in the superficial dermis and progressing to deeper levels, the following pigment cells can be seen: xanthophores (XA); iridophores containing small platelets (IR1); iridophores with larger platelets (IR2); melanophores (ME). H&E stain.

Figure 1.52   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a biopsied scale. The epidermis consists of an α-layer (AO), β-layer (BO), mesos layer (MO), oberhautchen (OB), and stratum germinativum (SG).Two types of pigment cells are seen in the dermis: melanophores (M) and iridophores (IR). There is a sharp boundary between these two populations of pigment cells. H&E stain.

Figure 1.53   Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Photomicrograph of skin using polarizing microscopy. Iridophores are readily identified by their birefringent platelets. H&E stain.

Overview of Reptile Biology, Anatomy, and Histology 45

Figure 1.54   Hawksbill sea turtle, Eretmochelys imbricata. Cheloniidae. Lateral view of the skull and mandible with major bones identified. There are no openings in the temporal region of the skull. The upper and lower jaws are covered by keratinized material, the rhamphothecae. Abbreviations for major structures seen here and for Figures 1.55–1.73: angular (A); articular (AR); basioccipital (BO); basisphenoid (BS); coronoid (CO); composite (COM) fused bone consisting of the articular, prearticular, angular, surangular; dentary (D); ectopterygoid (EC); external mandibular foramen (EMF); epipterygoid (EPI); external nares (EXN); frontal (F); jugal (J); lacrimal (L); lateral temporal fenestra (LTF); maxilla (M); nasal (N); orbit (OB); parietal (PA); parietal foramen (PAF); palatine (PAL); postfrontal (PF); prearticular (PAR); premaxilla (PM); postorbital (PO); parasphenoid (PS); prootic (PR); prefrontal (PRF); pterygoid (PT); quadrate (Q); quadratojugal (QJ); retroarticular process (RAP); rhamphotheca over dentary (RD); rhamphotheca over maxilla (RM); rhamphotheca over premaxilla (RPM); septomaxillae (SM); splenial (SP); squamosal (SQ); stapes (S); supraoccipital (SOC); supraorbital (SO); supratemporal (ST); superior temporal fenestra (STF); surangular (SA); vomer (VO). Skull courtesy of Michael Sapper.

Figure 1.55   Common snapping turtle, Chelydra serpentina. Chelydridae. Lateral view of the skull and mandible with major bones identified. There are no openings in the temporal region of the skull. The upper and lower jaws are covered by keratinized material, the rhamphothecae. Abbreviations as in Figure 1.54. Skull courtesy of Michael Sapper.

Figure 1.56   Tuatara, Sphenodon punctatus. Sphenodontidae. Lateral view of the skull with major bones identified. The premaxillae (PM) are well developed. The lacrimal and supratemporal are absent. There are two complete openings in the temporal region of the skull. The jugal (J) and quadratojugal (QJ) form the ventral borders of the lateral temporal fenestra, demarcated by double-headed arrow. The postorbital and squamosal form the lateral border of the superior temporal fenestra. The dentition is acrodont. Abbreviations as in Figure 1.54. Courtesy of David Kizirian and the American Museum of Natural History, New York, NY.

46  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.57   Tuatara, Sphenodon punctatus. Sphenodontidae. Dorsal view of the skull with major bones identified. There is a well-developed premaxilla (PM) and parietal foramen (PAF). The superior temporal fenestra (STF; demarcated by arrows) is complete. Abbreviations as in Figure 1.54. Courtesy of David Kizirian and the American Museum of Natural History, New York, NY.

Figure 1.58   American alligator, Alligator mississippiensis. Alligatoridae. Lateral view of the skull and mandible with major bones identified. The temporal region of the skull has two openings (lateral temporal fenestra [LTF] and superior temporal fenestra [STF]). Crocodilians have an elongated upper jaw (including the bony palate). A large external mandibular foramen (EMF) is in the caudal aspect of the mandible. Skull courtesy of the Florida Museum of Natural History, Gainesville. Abbreviations as in Figure 1.54.

Figure 1.59   American alligator, Alligator mississippiensis. Alligatoridae. Dorsal view of the skull and mandible with major bones identified. All bones of the upper jaw are firmly attached to the skull. Both lateral (LTF) and superior temporal fenestrae (STF) are present. Skull courtesy of the Florida Museum of Natural History, Gainesville. Abbreviations as in Figure 1.54.

Figure 1.60   Five-toed worm lizard, Bipes biporus. Bipedidae. Lateral view of the skull and mandible with major bones identified. This fossorial squamate shows marked fusion of skull bones. The postfrontal and squamosal are absent. The quadrate (Q) is fixed. Adjacent bones have grown over the temporal openings. The dentition is pleurodont. Abbreviations as in Figure 1.54. Courtesy of Theodore J. Papenfuss.

Overview of Reptile Biology, Anatomy, and Histology 47

Figure 1.61   Green iguana, Iguana iguana. Iguanidae. Lateral view of the skull and mandible with major bones identified. A lacrimal (L) bone is present. The ventral boundary of the lateral temporal fenestra (between the jugal [J] and quadrate [Q] bones) is absent resulting in an open area on the side of the skull. The quadratojugal is absent and the quadrate is movable. The epipterygoids (EPI) are well developed. The dentition is pleurodont. Abbreviations as in Figure 1.54. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.62   Green iguana, Iguana iguana. Iguanidae. Dorsal view of the skull with major bones identified. The premaxilla (PM) is well developed. A complete superior temporal fenestra (STF) and parietal fossa (PAF), which accommodates the parietal eye, can be seen. Abbreviations as in Figure 1.54. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.63   Bearded dragon, Pogona vitticeps. Agamidae. Lateral view of the skull and mandible with major bones identified. The lower temporal bar (quadratojugal) of the lateral temporal fenestra (double-headed arrow) is absent and results in an open area on the side of the skull. The quadrate (Q) is movable. Dentition is acrodont. Abbreviations as in Figure 1.54. Skull courtesy of Jeanette Wyneken.

Figure 1.64   Bearded dragon, Pogona vitticeps. Agamidae. Dorsal view of the skull with major bones identified. The superior temporal fenestra (STF) is complete. Abbreviations as in Figure 1.54. Skull courtesy of Jeanette Wyneken.

48  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.65   Jackson’s chameleon, Chamaeleo jacksoni. Chamaeleonidae. Lateral view of skull and mandible. The skull is flattened laterally. The upper horns are supported by the prefrontal bones and the lower horn by the premaxillary bone. The ventral boundary of the lateral temporal fenestra (double-headed arrow) is absent resulting in an open area on the side of the skull. Dentition is acrodont. Abbreviations as in Figure 1.54. Skull courtesy of the Florida Museum of Natural History, Gainesville. Figure 1.66   Nile monitor, Varanus niloticus. Varanidae. Lateral view of the skull, which is dorsoventrally flattened and elongate. The jugal (J) is thin and extends caudally upward. The lateral temporal fenestra is poorly defined. Dentition is pleurodont. Abbreviations as in Figure 1.54. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.67   Nile monitor, Varanus niloticus. Varanidae. Dorsal view of the skull and the mandible. The nasals (N) are thin and the bony openings (double-headed arrow) for the external nares are long and large resulting in exposure of the septomaxillae (SM). The superior temporal fenestrae are poorly defined. Dentition is pleurodont. Abbreviations as in Figure 1.54. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.68   Boa constrictor, Boa constrictor. Boidae. Dorsal view of the skull and lateral profiles of the mandibles. There is no supraorbital and no contact between the premaxillae (PM) and maxillae (M). As in all snakes, there are no lacrimals, jugals, quadratojugals, or squamosals. The dentary (D) bones are not fused at a symphysis, allowing widening of the lower jaws for prey ingestion. The maxillae can move independent of the brain case. The quadrate (Q) is strap-like and, in life, articulates with the pterygoids (PT), supratemporal (ST), and the articular process (AR) of the mandible. Temporal fenestrae have merged. Although snake teeth have been traditionally characterized as pleurodont, a more current and accurate description is that each snake tooth is actually ankylosed to the rim of a low socket, a type of modified thecodont dentition. Abbreviations as in Figure 1.54. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Overview of Reptile Biology, Anatomy, and Histology 49

Figure 1.69   Boa constrictor, Boa constrictor. Boidae. Palatal view of the skull and medial profiles of the mandibles are seen in the lower left image. There are no teeth on the premaxillae. Teeth are present on the maxillae (M), palatines (PAL), pterygoids (PT), and dentary (D) bones. Although snake teeth have been traditionally characterized as pleurodont, a more current and accurate description is that each snake tooth is actually ankylosed to the rim of a low socket, a type of modified thecodont dentition. The quadrate (Q) is short and strap-like and, in life, articulates with the pterygoids (PT), supratemporal (ST), and the articular process (AR) of the mandible. The stapes (S), the only middle ear ossicle in reptiles, is closely associated with the supratemporal (ST) bone. The relationship between S and ST is better visualized in the upper right macro image of this region of the skull. Abbreviations as in Figure 1.54. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.70   Reticulated python, Python reticulatus. Pythonidae. Dorsal view of the skull and medial profile of the mandibles. The premaxillae (PM) are independent of the maxillae (M). A supraorbital (SO) bone is present. The quadrate (Q) is short and stout and articulates with the supratemporal (ST) dorsally and the articular process (AR) of the mandible ventrally. Abbreviations as in Figure 1.54. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.71   Reticulated python, Python reticulatus. Pythonidae. Palatal view of the skull and medial profiles of the mandibles. Teeth are present on the premaxillae (PM), maxillae (M), palatines (PAL), pterygoids (pt), and dentary (D) bones. While snakes are considered to have pleurodont dentition, teeth are set in shallow sockets and may represent a type of thecodont dentition. Abbreviations as in Figure 1.54. Skull courtesy ofthe Florida Museum of Natural History, Gainesville.

Figure 1.72   Eastern diamondback rattlesnake, Crotalus adamanetus. Viperidae. Lateral view of the skull and mandibles. No lateral temporal fenestra is present. The maxillary (M) bone is short, only supports the fangs, and rotates off the prefrontal (PF) in fang erection. Regarding their fangs, rattlesnakes and other vipers are solenoglyphs. The rostral end of the ectopterygoid (EC) functions as a lever in fang erection. The quadrate (Q) is long and extends caudally and ventrally from the supratemporal (ST) to the articular process (AR) of the mandible. The stapes (S) is closely associated with the quadrate. The pterygoid (PT) also is long and articulates with the palatine (PAL) anteriorly and the quadrate–articular process posteriorly. Most of the bones in the mandible are fused. Abbreviations as in Figure 1.54. Skull courtesy of Michael Sapper.

50  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.73   Eastern diamondback rattlesnake, Crotalus adamanetus. Viperidae. Dorsal view of the skull. There is no superior temporal fenestra. The frontal (F) and parietal (PA) are fused. The nasal (N) is independent of the frontal (F), and the premaxillae (PM) are independent of the maxillae (M). The anterior end of the ectopterygoid (EC) abuts on the maxilla (M). Abbreviations as in Figure 1.54. Skull courtesy of Michael Sapper.

Figure 1.74   Black mamba. Dendroaspis polylepis. Elapidae. Lateral view of the skull and mandibles. The maxillae (M) supports the fangs and is relatively long compared to maxillae of vipers and pit vipers. Regarding their fangs, mambas are proteroglyphs. The palatine (PAL) and maxillary bones are capable of erection. Skull courtesy of Michael Sapper. Abbreviations as in Figure 1.54.

Figure 1.75   Hawksbill sea turtle, Eretmochelys imbricata. Cheloniidae. Mandibles are covered by a heavily keratinized rhamphotheca.

Figure 1.76   American alligator, Alligator mississippiensis. Alligatoridae. Teeth are set in sockets and are categorized as thecodont. Jaw courtesy of the Florida Museum of Natural History, Gainesville.

Overview of Reptile Biology, Anatomy, and Histology 51

Figure 1.77   Tuatara, Sphenodon punctatus. Sphenodontidae. Teeth are on the biting surfaces of the maxillary (M) and palatine (PAL) bones and are categorized as acrodont. Also seen are the vomer (VO) and pterygoid (PT) bones. Courtesy of David Kizirian and the American Museum of Natural History, NY.

Figure 1.78   Jackson’s chameleon, Chamaeleo jacksoni. Chamaeleonidae. Teeth are on the biting surfaces of the maxillary and palatine bones and are categorized as acrodont. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.79   Green iguana, Iguana iguana. Iguanidae. Teeth are on a ledge on the medial aspect of the jaw and are categorized as pleurodont dentition. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.80   Marine iguana, Amblyrhynchus cristatus. Iguanidae. Teeth are on a ledge on the medial aspect of the jaw and are categorized as pleurodont dentition. Skull courtesy of the Florida Museum of Natural History, Gainesville.

Figure 1.81   Nile monitor, Varanus niloticus. Varanidae. Teeth are on a ledge on medial aspect of the jaw and are categorized as pleurodont dentition. Skull courtesy of the Florida Museum of Natural History, Gainesville.

52  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.82   Green turtle, Chelonia mydas. Cheloniidae. Palatal view. The hard palate is partially developed.

Figure 1.83   American alligator, Alligator mississippiensis. Alligatoridae. Oral cavity. The hard palate is complete. A dorsal flap (DF) of soft tissue is located at the posterior aspect of the hard palate that overlaps with the ventral flap (VF) located at the base of the tongue. Directly posterior to the dorsal flap are the internal nares (IN) and the tonsil (T). The glottis (G) is posterior to the ventral flap.

Figure 1.84   Green iguana, Iguana iguana. Iguanidae. Extended orange dewlap of a male green iguana.

Overview of Reptile Biology, Anatomy, and Histology 53

Figure 1.85   Green iguana, Iguana iguana. Iguanidae. Lateral radiograph of the head and dewlap of a green iguana. Paired ceratobranchial II extends into the dewlap. One of the ceratobranchials is fractured (arrow).

Figure 1.86A   Red-eared slider, Trachemys scripta elegans. Emydidae. Ventral view of a skeletal preparation. Axial skeleton and girdles can be seen. The neural processes of vertebrae and ribs are fused with the dermal bone of the carapace. Abbreviations for major structures: acromion process (AC); coracoid (CO); epipubic cartilage (EPC); femur (FE); fibula (FI); humerus (H); ischium (IS); lateral pubic process (LPP); neural process (NP) of vertebra; pubis (PU); radius (RA); rib (RI); scapula (SC); thyroid (puboischiadic) fenestra (TF); tibia (T); ulna (U); vertebral body (VB). The ilium cannot be seen in this ventral view.

Figure 1.86B  Red-eared slider, Trachemys scripta elegans. Emydidae. Ventral oblique view of the pelvic girdle and vertebrae (V). The relationship between the ilium (IL) and other pelvic girdle bones and hindlimb bones can be seen. See Figure 1.86A for abbreviations.

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Figure 1.87   Green turtle, Chelonia mydas. Cheloniidae. Triradiate pectoral girdle consisting of scapula (S), acromion process (AC) of the scapula, and procoracoid (PC). The glenoid cavity (GC) is the shoulder joint and articulates with the head of the humerus.

Figure 1.88   Ball python, Python regius. Pythonidae. Paired spurs (arrows), representing remnants of the femur, are located adjacent to the cloaca.

Figure 1.89   Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph of the epiphysis (EP) of a long bone. In chelonians, the EP of long bones is cartilaginous. Spongy bone (SB) forms at the diaphyseal end of the epiphyseal growth plate (EPGP). H&E stain.

Overview of Reptile Biology, Anatomy, and Histology 55

Figure 1.90   Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph of the epiphyseal growth plate (EPGP) of a long bone. The EPGP consists of chondrocytes arranged in longitudinal columns that are separated from adjacent columns by a hyaline matrix. A zone of reserve cartilage (ZR) is adjacent to the epiphysis (EP). Moving toward the diaphysis there is a zone of proliferation (ZP), zone of hypertrophy (ZH), and zone of calcification resulting in the formation of spongy bone (SB). H&E stain.

Figure 1.91   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the epiphysis of a long bone. The epiphysis contains spongy bone formed by a secondary center of ossification. The surface is covered by a thick layer of cartilage containing chondrocytes. H&E stain.

I

Figure 1.92   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a long bone. An epiphyseal growth plate (EPGP) is between the epiphysis (EP) and diaphysis (DI). H&E stain.

Figure 1.93   Green iguana, Iguana iguana. Iguanidae. Higher magnification photomicrograph of the epiphyseal growth plate (EPGP) seen in Figure 1.92. The EPGP is between the epiphysis (EP) and diaphysis (DI) and consists of a zone of reserve cartilage (ZR), zone of proliferation (ZP), zone of hypertrophy (ZH), and zone of calcification (ZC). H&E stain.

56  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.94   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a long bone. The diaphysis has avascular compact cortical bone, with the periosteum (PE) on the outer surface and the endosteum (EN) on the inner surface. Skeletal muscle (SM) is attached to the periosteum. Osteocytes are scattered throughout and arrest lines (arrows) can be seen. H&E stain.

Figure 1.95   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a skeletal muscle. Two skeletal muscle fascicles (SMFA) are separated by perimysium (arrows). Fascicles consist of skeletal muscle fibers (SMFI) with peripherally located nuclei. Vessels (V) are seen within the perimysium. H&E stain.

Figure 1.96   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a cross-section of the lateral aspects of the head. There is an outer epidermis (EP) consisting of keratin and epithelial cells and an inner oral mucosa (OM) consisting of mucous epithelial cells. Labial glands (LG) surrounded by connective tissue, teeth (T), jawbones (B), and braincase bone (BC) are seen. H&E stain.

Overview of Reptile Biology, Anatomy, and Histology 57

Figure 1.97   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the tongue. The upper surface consists of papillary projections of a nonkeratinized stratified squamous epithelium supported by fibrovascular stroma (FV). The lower surface is covered by a keratinized epithelium (KEP). Fascicles of skeletal muscle (SM) are oriented in various directions. H&E stain.

Figure 1.98   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the tongue. The surface consists of papillary projections of a thick stratified squamous epithelium supported by fibrovascular connective tissue. Numerous lingual mucous glands are scattered throughout this area of the tongue. H&E stain.

Figure 1.99   Western diamondback rattlesnake, Crotalus atrox. Viperidae. Photomicrograph of a cross-section of the lower portion of the anterior oral cavity. The tongue is located in the midline within the tongue sheath (TS) and consists of fascicles of skeletal muscle (SM) that are oriented in various directions. The glottis (G) is above the tongue and the mucous epithelium (MU) lining the oral cavity is lateral to the tongue. H&E stain.

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Figure 1.100   Death adder, Acanthophis antarcticus. Elapidae. The venom gland (VG) is located caudal and ventral to the eye and is surrounded by a connective tissue capsule. Venom from the gland drains into the venom gland duct (VD), which courses below the eye. At its terminus, the duct fuses with the sheath surrounding the fang. The external adductor superficialis muscle (AS) attaches to the gland. The adductor profundus muscle (AP) is dorsal and caudal to the AS.

Figure 1.101   Palestine viper, Vipera (Daboia) palaestinae. Viperidae. Photomicrograph of a venom gland. The venom gland is a branched tubular gland consisting of dilated acini containing venom (V). A connective tissue capsule (CT) surrounds the venom gland. Adjacent to the venom gland is skeletal muscle (SM) that is surrounded by the epimysium (EP). H&E stain.

Figure 1.102  Neotropical rattlesnake, Crotalus durissus. Viperidae. Photomicrograph of the venom gland. Approximately 80% of the cells within acini are cuboidal secretory cells. An eosinophilic globular material (venom) is being released from the surface of secretory cells and is accumulating in the lumens of individual glands. H&E stain.

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Figure 1.103   Leatherback sea turtle, Dermochelys coriacea. Dermochelyidae. Keratinized papillae are seen lining the esophagus. These are unique structures of the esophagus of sea turtles.

Figure 1.104   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the esophagus. The mucosa (MU) consists of stratified cuboidal to columnar ciliated epithelial cells, and goblet cells. Vessels (V) are scattered about the lamina propria (LP) and the muscularis mucosae (MM) is thin. The submucosa (SM) is below the MM. H&E stain.

Figure 1.105   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the esophagus. The surface is thrown into folds and the mucosa (MU) consists of stratified ciliated epithelial cells and goblet cells. The following structures are also seen: lamina propria (LP); submucosa (SM); muscularis externa, inner circular (IC); muscularis externa, outer longitudinal (OL); serosa (S). H&E stain.

Figure 1.106   Horned viper, Cerastes cerastes. Viperidae. Photomicrograph of the esophagus. The surface is thrown into folds and the mucosa (MU) consists of stratified cuboidal epithelial cells. The following structures are also seen: lamina propria (LP); muscularis mucosae (MM); submucosa (SM). H&E stain.

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Figure 1.107   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of the esophagus. The surface is thrown into folds and the mucosa (MU) consists of a stratified columnar to cuboidal mucous epithelium consisting of numerous goblet cells and ciliated epithelial cells. The following structures are also seen: lamina propria (LP); submucosa (SM); external muscularis, inner circular (IC); external muscularis, outer longitudinal (OL); serosa (SE). H&E stain.

Figure 1.108   Reticulated python, Python reticulatus. Pythonidae. Esophageal tonsils (arrows) are raised ovoid lymphoid structures having a central cleft. They are particularly prominent in boas and pythons.

Figure 1.109   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of an esophageal tonsil. A mucous epithelium consisting of ciliated epithelial cells and goblet cells covers an aggregate of lymphoid tissue, histiocytes, plasma cells, heterophils, and blood vessels. H&E stain.

Figure 1.110   Emerald tree boa, Corallus caninus. Boidae. Photomicrograph of an esophageal tonsil. A mucous epithelium covers an aggregate of lymphoid tissue. H&E stain.

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Figure 1.111   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of fundic gland region of the stomach. Gastric glands (GG) open into gastric pits (GP) that are lined by columnar epithelial cells. Adjacent pits are connected by ridges of epithelial cells (ER) that are of the same type as those lining the pits. Gastric glands consist of dark (serous or oxyntopeptic) cells. Strands of connective tissue (arrows) within the lamina propria (LP) separate adjacent glands. Below the lamina propria the muscularis mucosae (MM) consists of inner circular and outer longitudinal layers of smooth muscle fibers. The submucosa (SM) is below the muscularis mucosa and contains blood vessels (V). H&E stain.

Figure 1.112   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a mucosal fold in the fundic gland region of the stomach. Specialized neck cells (NC) are seen between gastric glands (GG) and gastric pits (GP). Only dark cells occur in the gastric epithelium of most snakes. Ridges of epithelial cells (ER) separate adjacent gastric pits. The lamina propria (LP) at the base of the glands extends between adjacent gastric glands as thin strands of connective tissue. The muscularis mucosae (MM) consist of inner circular and outer longitudinal layers of smooth muscle fibers. The submucosa (SM) contains connective tissue and blood vessels (V). H&E stain.

Figure 1.113   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of the fundic region of the stomach. This section is through a fold of mucosa. Epithelial cells covering ridges (ER) and lining gastric pits (GP) are deeply PAS-positive. Neck cells (NC) open into the gastric pits and are moderately PAS-positive. The epithelial cells of the gastric glands (GG) are all dark cells and are PAS-negative. Only dark cells occur in the gastric epithelium of most snakes. Additional structures seen are: lamina propria (LP); muscularis mucosae (MM); submucosa (SM); vessels (V). PAS stain.

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Figure 1.114   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of the fundic region of the stomach. This section is through a fold of mucosa. Nuclei stain deep blue to black, dark cells of gastric glands (GG), and muscle cells of muscularis mucosae (MM) stain red. Apical portions of epithelial cells covering ridges (ER) and connective tissue in lamina propria (LP) and submucosa (SM) stain green. The neck cells (NC) stain light green. Masson’s trichrome stain.

Figure 1.115   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a crosssection through the fundic region of the stomach. Nuclei stain deep blue to black and dark cells of gastric glands (GG), muscle cells of the muscularis mucosae (MM), and the inner circular (IC) and outer longitudinal (OL) layers of the muscularis externa stain red. The outer longitudinal layer is thin. Apical portions of epithelial cells covering ridges (ER) and connective tissue in lamina propria (LP), submucosa (SM), and serosa stain green. The neck cells (NC) stain light green. Masson’s trichrome stain.

Figure 1.116   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of the pars pylorica region of the stomach. The gastric glands (GG) are short and consist only of positive-staining mucous cells. The apical portion of mucosal epithelial cells (EP) is also PAS-positive. PAS stain.

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Figure 1.117   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of small intestine villi. The epithelium (EP) consists of absorptive cells and goblet cells. Lacteals (L) are present within the lamina propria (LP) of villi. Crypt-like structures (CR) are seen at the base of adjacent villi. The muscularis mucosae consists of isolated bundles (arrows) of smooth muscle cells. The submucosa (SM) is thin and is surrounded by the inner circular layer (IC) of the muscularis externa. H&E stain.

Figure 1.118   Green iguana, Iguana iguana. Iguanidae. Small intestine. Photomicrograph of villi within the small intestine. The intestinal epithelium (EP) consists of absorptive cells and goblet cells. The lamina propria (LP) of villi contains fibrous connective tissue, vessels, and lacteals. Primitive crypt-like structures (CR) are at the base of some villi. Lymphoid tissue (LT) is present within the lamina propria surrounding the base of a crypt. H&E stain.

Figure 1.119   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of the small intestine. The intestinal epithelium (EP) consists of absorptive cells and goblet cells. Crypts are absent. There is a subtle boundary (arrows) between the connective tissue of the lamina propria (LP) and that of the submucosa (SM). The muscularis mucosae are absent. The muscularis externa consist of an inner circular layer (IC) and an outer longitudinal layer (OL) of smooth muscle fibers. H&E stain.

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Figure 1.120   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of small intestine. Numerous PAS-positive goblet cells are seen within the intestinal mucosal epithelium. Absorptive cells are PAS-negative. PAS stain.

Figure 1.121   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of small intestine. Mucinous material within goblet cells and connective tissue in lamina propria (LP), submucosa (SM), between inner (IL) and outer (OL) layers of muscularis externa, and serosa (S) stain green. Mucosal absorptive epithelial cells and smooth muscle cells stain red. The muscularis mucosae (MM) is thin and consists of a single layer of muscle cells. Masson’s trichrome stain.

Figure 1.122   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of the colon. The colonic epithelium (CE) consists of PAS positivestaining goblet cells (GC) and absorptive columnar epithelial cells (AC). The lamina propria is thin and vessels are within the submucosa (SM). The muscularis externa consists of inner circular (IC) and outer longitudinal (OL) layers. The serosa (S) is on the outer surface of the colon. PAS stain. Figure 1.123   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the colon. The colonic epithelium (CE) is stratified and consists primarily of goblet cells. Glands (G) are present within the lamina propria. H&E stain.

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Figure 1.124.  Red-eared slider, Trachemys scripta elegans. Emydidae. Ventral view of the coelomic cavity. The gallbladder (GB) is in the right lobe (RL) of the liver. The pancreas (arrow) is attached to the serosal surface of the pylorus (PY) and duodenum (DU). The following structures are also seen: colon (CO); left lobe of the liver (LL); ovarian follicles (OF); small intestine (SI); urinary bladder (UB).

Figure 1.125   Green turtle, Chelonia mydas. Cheloniidae. Ventral view of the coelomic cavity. The gallbladder (GB) is in the right liver lobe (RL). The following structures are also seen: colon (CO); duodenum (DU); heart (HT); left liver lobe (LL); pylorus (PY); stomach (ST).

Figure 1.126   American alligator, Alligator mississippiensis. Alligatoridae. Ventral view of the coelomic cavity. A pseudodiaphragm (arrows) separates the lung (LU), liver (right lobe [RL]; left lobe [LL]), and heart (H) from the viscera (gallbladder [GB], stomach [ST]) of the posterior coelomic cavity.

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Figure 1.127   Green iguana, Iguana iguana. Iguanidae. Ventral view of the anterior coelomic cavity. The heart (HT) is located between the forelegs, with the apex between the cranial margins of the two liver lobes (left lobe [LL]; right lobe [RL]). The gallbladder (GB) is in the right lobe of the liver. The following structures are also seen: colon (CO); stomach (ST).

Figure 1.128   Green iguana, Iguana iguana. Iguanidae. Dorsal surface of the right liver lobe (RL). Numerous bile ducts (arrows) are seen coursing from the area of the gallbladder to the duodenum. The pancreas (PA) is seen on the serosal surface of the duodenum, with a caudal portion covering the spleen (SP). A small caudal portion of the pancreas is on the opposite surface of the duodenum, caudal to the spleen. The spleen is located between the colon and duodenum (DU).

Figure 1.129   Ball python, Python regius. Pythonidae. Ventral surface of the liver. The caudal vena cava (arrows) is seen on the ventral surface and the hepatic vein is on the dorsal surface (not seen in this image). These vessels divide the liver into two lobes. The respiratory portion of the lung (LU) is between the heart (HT) and cranial pole (CP) of the liver (LI). The lung continues on the dorsal surface of the liver and gradually transforms into an air sac (AS).

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Figure 1.130   Fer-de-lance, Bothrops atrox. Viperidae. Photomicrograph of a crosssection of the liver. The hepatic portal vein (PV) is on the dorsal surface and the hepatic vein (HV) is on the ventral surface. Numerous bile ducts (BD) are adjacent to the PV.

Figure 1.131   Western diamondback rattlesnake, Crotalus atrox. Viperidae. Ventral view of the mid-coelomic cavity. The gallbladder (GB), spleen (SP), and pancreas (PA) form a triad and are caudal to the liver (LI) and adjacent to the pyloroduodenal junction (PDJ). The following structures are also seen: small intestine (SI); stomach (ST).

Figure 1.132   Boa constrictor, Boa constrictor. Boidae. Ventral view of the mid-coelomic cavity. The gallbladder (GB), spleen (SP), and pancreas (PA) are closely associated and are adjacent to the small intestine (SI). The ovary (OV) is also near this triad. While closely associated, the spleen is separated from the body of the exocrine pancreas in boid snakes.

Figure 1.133   Corn snake, Elaphe guttata guttata. Colubridae. Ventral view of the mid-coelomic cavity. The gallbladder (GB), spleen (SP), and pancreas (PA) are closely associated, and are adjacent to the small intestine (SI) and caudal to the pylorus (PY) of the stomach. In colubrid snakes, the spleen is associated with the anterior margin of the pancreas.

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Figure 1.134   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph showing the spatial relationship of the pancreas (PA), spleen (SP) and gallbladder (GB). H&E stain.

Figure 1.135   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the gallbladder. The gallbladder is within the right lobe of the liver. The mucosa consists of a columnar epithelium (EP) and a vascular lamina propria (LP), which is often thrown into folds. A perimuscular layer (MU) is between the lamina propria and the liver (LI). H&E stain.

Figure 1.136   Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of the gallbladder. The cytoplasm of epithelial cells (EP) lining the gallbladder stains deep red, nuclei stain deep blue to black, connective tissue in the lamina propria (LP) and elsewhere stains blue, and the muscularis (MU) stains light red. Cords of hepatocytes can be seen within the liver (LI). Masson’s trichrome stain.

Figure 1.137   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of a transverse section through a liver lobule. Plates (two cells thick) of hepatocytes are seen radiating from a central vein (CV) that is filled with red blood cells. H&E stain.

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Figure 1.138   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a transverse section through a liver lobule. Plates of hepatocytes (HP) are seen radiating from a central vein. Plates are separated by sinusoids (S) that drain into the central vein (CV). H&E stain.

Figure 1.139   Burmese python, Python molurus bivitattus. Pythonidae. Photomicrograph of a transverse section through the liver. There is no apparent organization of the liver into lobules. Hepatic plates are not radially arranged around the central vein (CV). H&E stain.

Figure 1.140   Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of a transverse section through a portal tract in the liver. In this section the portal tract includes a branch of the portal vein (PV), bile ductules (BD), and melanomacrophages (MM). H&E stain.

Figure 1.141   Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of a transverse section through a portal tract in the liver. In this section the portal tract includes a branch of the portal vein (PV), bile ductules (BD), and melanomacrophages (MM). Red-staining smooth muscle cells are in the wall of the bile ductule. Masson’s trichrome stain.

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Figure 1.142   Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of a transverse section through a portal tract in the liver. In this section the portal tract includes a branch of the portal vein (PV), bile ductules (BD), and a branch of the hepatic artery (AR). Redstaining smooth muscle cells are in the wall of the bile ductule and artery. Masson’s trichrome stain.

Figure 1.143   Neotropical rattlesnake, Crotalus durissus. Viperidae. Photomicrograph of a transverse section through the liver. In this section the portal tract includes a branch of the portal vein (PV), bile ductules (BD), and a branch of the hepatic artery (AR). H&E stain.

Figure 1.144   Monocled cobra, Naja kaouthia. Elapidae. Photomicrograph of a transverse section through the liver. Adjacent plates of hepatocytes are separated by sinusoids (SI). Sinusoids are dilated and are lined with endothelial and Kupffer cells. Heterophils (HE) and red blood cells (RBC) are seen within sinusoids. H&E stain.

Figure 1.145   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the liver. Melanomacrophages are peripheral to the central vein (CV). H&E stain.

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Figure 1.146   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the liver showing numerous melanomacrophages. H&E stain.

Figure 1.147   Cobra, Naja sp. Elapidae. Photomicrograph of a transverse section through a portal tract in the liver. In this section the portal tract includes a branch of the portal vein (PV) and bile ductules (BD). No melanomacrophages are seen. H&E stain.

Figure 1.148   Madagascan tree boa, Sanzinia madagascariensis. Boidae. Photomicrograph of the liver showing numerous melanomacrophages. H&E stain.

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Figure 1.149   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the liver. Large melanomacrophages are seen containing a brown-staining material. H&E stain.

Figure 1.150   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the liver. Iron is seen as blue-staining granular material within melanomacrophages and hepatocytes. Prussian blue stain.

Figure 1.151   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the liver. Melanin is seen as black-staining particulate material within melanomacrophages. Fontana’s method.

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Figure 1.152   New Guinea snakeneck turtle, Chelodina novaeguineae. Chelidae. The spleen (SP) and pancreas (PA) are separated. The following structures are also seen: duodenum (DU); pylorus (PY); fundus (FU).

Figure 1.153   Green turtle, Chelonia mydas. Cheloniidae. The pancreas (arrows) is adjacent to the duodenum (DU), with the posterior portion associated with the spleen (SP). Also seen is the pylorus (PY). Courtesy of Brian Stacy.

Figure 1.154   Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of exocrine acinar cells in the pancreas. The apical portions of cells are eosinophilic and the basal portions are basophilic. H&E stain.

Figure 1.155   Red-eared slider, Trachemys scripta elegans. Emydidae. Higher magnification photomicrograph of exocrine acinar cells in the pancreas of Figure 1.154. Cross-sections of exocrine acinar cells. The apical eosinophilic portions of cells consist of eosinophilic zymogen granules and the basophilic basal portions are nuclei. H&E stain.

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Figure 1.156   Rhinoceros viper, Bitis nasicornis. Viperidae. Photomicrograph of the pancreas. The islets of Langerhans (IL) are not clearly demarcated from the more basophilic exocrine pancreas. H&E stain.

Figure 1.157   Monocled cobra, Naja kaouthia. Elapidae. Photomicrograph of the pancreas. The islets of Langerhans (IL) stain more eosinophilic, but otherwise are not clearly demarcated from the more basophilic exocrine pancreas.

Figure 1.158   Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Photomicrograph of the pancreas. A transition can be seen (arrows) between the islets of Langerhans (IL) and the exocrine pancreas (EX). H&E stain.

Figure 1.159   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of the pancreas. Whereas the islets of Langerhans (IL) and exocrine pancreas acinar cells stain eosinophilic, the IL stains more deeply eosinophilic and has smaller granules. H&E stain.

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Figure 1.160   Gaboon viper, Bitis gabonica. Viperidae. Photomicrograph showing the spatial relationship of the spleen (SP) and pancreas (islets of Langerhans [IL] and exocrine pancreas [EX]). A band of connective tissue (CT) separates the SP from the pancreas. H&E stain.

Figure 1.161   White-throated monitor, Varanus albigularis. Photomicrograph of the pancreas. Islets of Langerhans (IL) are scattered within the dorsal lobe of the pancreas. H&E stain. Courtesy of John Roberts.

Figure 1.162   Savannah monitor, Varanus exanthematicus. Varanidae. The juxtasplenic body (JX), spleen (SP), and exocrine pancreas (EXP) are closely associated and are adjacent to the junction of the stomach and small intestine. The intestinal tract is relatively short. Courtesy of Barbara Sheppard.

Figure 1.163   White-throated monitor, Varanus albigularis. Varanidae. Photomicrograph of the pancreas and spleen. The juxtasplenic body (JX) consists of islet tissue and is surrounded by splenic lobules (SP). H&E stain. Courtesy of John Roberts.

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Figure 1.164   Desert tortoise, Gopherus agassizii. Testudinidae. Nasal cavity. Diagrammatic representation of cross-sections through the head cranial to the eyes. A large nasal cavity (arrows) is seen, with the dimensions changing from cranial (A) to caudal (D). (From Jacobson ER et al. 1991. J Wildl Dis 27: 296–316. With permission.)

Figure 1.165   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a cross-section through the head cranial to the eyes. The concha (CO) divides the cavity into a lateral extraconchal space (ECS) and a medial space (MS). The following structures are also seen: epidermis (EP), labial glands (LG), maxillary bone (MA), oral mucosa (OM), pulp (P), palatine (PAL) bone, and teeth (T).

Figure 1.166   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Higher magnification photomicrograph of a cross-section of the nasal cavity seen in Figure 1.165. The concha is covered by an olfactory epithelium (OE) dorsally and a mucous epithelium (ME) ventrally. Cartilage (CA) supports the concha. Serous glands (SG) and nerves (NE) are below the olfactory epithelium.

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Figure 1.167   Green turtle, Chelonia mydas. Cheloniidae. Ventral view of the coelomic cavity with viscera removed. The lungs extend to the anterior poles of the kidneys (KI), near the adrenal gland (AD). Ovaries (OV) of this juvenile turtle also are seen on the ventral surface of the kidneys.

Figure 1.168   Black and white tegu, Tupinambis merianae. Teiidae. Ventral view of the coelomic cavity. The lungs are located in the anterior half of the coelomic cavity.

Figure 1.169   Green iguana, Iguana iguana. Iguanidae. View of the medial surface of the lung. The lung is a transitional type. Paired bronchi (BR) enter the lungs at the hilus (HI). An intercameral septum (SP) divides the lung into two confluent chambers (CH). Cranial edicular and more caudal faveolar parenchyma are present. The lung is also divided into a smaller prehilar (PRH) and larger posthilar (POH) lung.

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Figure 1.170   Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Left lateral view of the body. The skin and ribs have been removed and the lungs have been inflated. Chameleons have intrapulmonary septae and sac-like diverticulae. Courtesy of Robert Coke.

Figure 1.171   Corn snake, Elaphe guttata guttata. Colubridae. View of the ventral coelomic cavity. The lung (LU) is between the heart (HT) and cranial pole of the liver (LI). The lung transforms into a nonrespiratory air sac (AS) on the craniodorsal surface of the liver.

Figure 1.172   Diamond python, Morelia spilota spilota. Pythonidae. Ventral view of the cranial coelomic cavity. The respiratory portion of the lung (RL) is posterior to the heart (not seen in this image), adjacent to the esophagus (ES), and gradually transforms into an air sac (AS) on the surface of the liver (LI). Whereas boid snakes have paired lungs, the left lung is shorter than the right.

Figure 1.173   Burmese python, Python molurus bivittatus, Pythonidae. Respiratory portion of the lung. The lung is unicameral and faveolar parenchyma imparts a honeycomb appearance to the lung. Image courtesy of John Roberts.

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Figure 1.174   King snake, Lampropeltis getula. Colubridae. Photomicrograph of a cross-section of the lung. Faveoli (FV) open into a central lumen (CL), which is continuous with the bronchus (BR). The bronchial wall is supported by cartilage (CA) and the lumen is lined with ciliated and mucous epithelial cells. Adjacent faveoli are separated by a connective tissue septum and the luminal end of each septum has a bundle of smooth muscle cells (SM). The esophagus (ES) is adjacent to the lung. H&E stain.

Figure 1.175   Western diamondback rattlesnake, Crotalus atrox. Viperidae. Photomicrograph of the lung. Connective tissue septae (SP) separate adjacent faveoli (FV). Red blood cells are seen within capillaries lining the faveolar spaces. H&E stain.

Figure 1.176   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the lung. A bundle of smooth muscle (SM) is located at the central luminal (CL) aspect of each septa (SP), which separate adjacent faveolae (FV). H&E stain.

Figure 1.177   Bush viper, Atheris squamiger. Viperidae. Photomicrograph of the lung. A bundle of smooth muscle (SM) is located at the central luminal aspect of each septa (SP), which separates adjacent faveolae (FV). H&E stain.

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Figure 1.178   Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Photomicrograph of cells lining a faveolus in a semithin section of the lung. Squamous alveolar type I (T1) and cuboidal type II (T2) cells containing punctate bodies within vacuoles are seen. Red blood cells (RBC) within endothelial lined capillaries (END) are also seen. Toluidine blue stain.

Figure 1.179   Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Transmission electron photomicrograph of cells lining a faveolus. Alveolar type II (T2) cells contain lamellar material within vacuoles. The cytoplasm of squamous alveolar type I (T1) cells cover much of the septal surface. Uranyl acetate and lead citrate.

Figure 1.180   Neotropical rattlesnake, Crotalus durissus. Viperidae. Photomicrograph of the liver (LI) and air sac (AS). The air sac extends from the midline, over the hepatic portal vein (PV), to the right dorsolateral margins of the liver. Masson’s trichrome stain.

Figure 1.181   Neotropical rattlesnake, Crotalus durissus. Viperidae. Higher magnification photomicrograph of the air sac in Figure 1.180. This portion of the air sac is lined with a ciliated columnar epithelium. Masson’s trichrome stain.

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Figure 1.182   Water snake, Nerodia sipedon. Colubridae. Ventral view of the mid-coelomic cavity. The left kidney (LK) is exposed and consists of multiple lobules. The right kidney (RK) is more cranial than the left kidney. The colon (CO) is between the kidneys. The belly scales of this snake are blue indicating the snake is in a skin renewal phase of a cycle of ecdysis.

Figure 1.183   Corn snake, Elaphe guttata guttata. Colubridae. Kidneys of a male snake with inactive testes. The right kidney (RK) is cranial to the left kidney (LK) and the ductus deferens (DD) is adjacent to the lateral margins of the kidneys. The colon (CO) is between the kidneys.

Figure 1.184   Tentacled snake, Erpeton tentaculum. Colubridae. Kidney during a reproductive period has a pale creamy coloration due to hypertrophy of the sexual segment. The lobules are not as apparent as in females, immature males, and males during nonreproductive periods. Courtesy of Brian Stacy.

Figure 1.185   Green turtle, Chelonia mydas. Cheloniidae. Ventral view of the coelomic cavity. The pelvic girdle is reflected. The kidneys (arrow) and the urinary bladder (UB) are adjacent to the hind limbs and within the pelvic canal.

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Figure 1.186   Gopher tortoise, Gopherus polyphemus. Testudinidae. Ventral view of the coelomic cavity near the pelvic canal. The pelvic girdle has been removed. The kidneys (KI) are on either side of the colon (CO) and the testes (T) are in contact with the cranial surface of kidneys. The adrenal glands (AD) are associated with the medial aspect of the kidneys. The urinary bladder (UB) is bilobed.

Figure 1.187   Green iguana, Iguana iguana. Iguanidae. Ventral view of the coelomic cavity with the pelvic girdle, pectoral girdle, and gastrointestinal tract removed. The kidneys (KI) are within the pelvic canal with the posterior poles extending into the base of the tail. The following structures are also seen: ductus deferens (DD); gallbladder (GB); hemipenes (HE); heart (HT); left liver lobe (LL); left lung (LLU); right liver lobe (RL); right lung (RLU); testes (TE). Courtesy of Jeanette Wyneken.

Figure 1.188   Western diamondback rattlesnake, Crotalus atrox. Viperidae. Photomicrograph of the kidney. Two renal corpuscles are seen, and each consists of a tuft of capillaries (glomerulus) covered by the visceral layer (VL) of Bowman’s capsule. The parietal layer (PL) forms the outer margins of Bowman’s capsule and the capsular or uniferous space (CS) is between the two layers. Proximal tubules (PT) have epithelial cells with an eosinophilic cytoplasm and distal tubules (DT) have epithelial cells with a basophilic cytoplasm. H&E stain.

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Figure 1.189   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the kidney. The parietal layer (PL) of Bowman’s capsule surrounds a glomerulus (GL) and consists of cuboidal epithelial cells. An afferent or efferent arteriole (AR) is seen between two distal tubules (DT). Proximal tubules (PT) have epithelial cells with an eosinophilic cytoplasm and DT have epithelial cells with a basophilic cytoplasm. H&E stain.

Figure 1.190   American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of the kidney. Glomeruli (GL) appear to be aligned in a row. H&E stain.

Figure 1.191   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of the kidney. The parietal layer (PL) of Bowman’s capsule surrounding the glomerulus (GL) is continuous with the short neck segment (NS) of the nephron. The neck segment is ciliated and is continuous with the nonciliated proximal tubule (PT). Cross-sections of another neck segment and several proximal tubules are seen. H&E stain.

Figure 1.192   Spitting cobra, Naja sp. Elapidae. Photomicrograph of the kidney. The parietal layer (PL) of Bowman’s capsule of a renal corpuscle is continuous with the short neck segment (NS) of the nephron. Connective tissue (CT) in the center of the glomeruli stains positive. Proximal tubules (PT) have brush borders and cytoplasmic granules that also stain with PAS. PAS stain.

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Figure 1.193   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of the kidney. Several renal corpuscles (RC) are seen. Proximal tubules (PT) are scattered throughout the section and are more eosinophilic compared to other components of the nephron. Intermediate segments (IS) have a basophilic staining cytoplasm and are about half the diameter of distal tubules (DT), which also have a basophilic staining cytoplasm. H&E stain. Figure 1.194   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of the kidney. Proximal tubules (PT) are large and have positive-staining granules throughout the cytoplasm. The height of cells in the intermediate segment (IS) are shorter than those in the proximal and distal tubules, and the nucleus to cytoplasmic ratio is greater than that of proximal and distal tubular epithelial cells. Cilia project into the lumen of cells in the IS. PAS stain.

Figure 1.195   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of the kidney. Proximal tubules (PT) are large and have a PAS-positive brush border (BB). The height of cells in the intermediate segment (IS) are shorter than those in the proximal and distal tubule, and the nucleus-tocytoplasmic ratio is greater. Cilia are seen in the lumen of the intermediate segment. The next segment, which is the distal tubule (DT), has distinct PAS-positive material around the luminal border and within the cytoplasm. This material is lacking in the IS. PAS stain.

Figure 1.196   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the kidney. Numerous collecting ducts (CD) are seen within the base of the kidney. H&E stain.

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Figure 1.197   Bush viper, Atheris squamiger. Viperidae. Photomicrograph of the kidney of a male snake. A distal tubule (DT) is continuous with a sexual segment (SS), which has numerous eosinophilic granules within the cytoplasm. The cytoplasm of proximal tubular (PT) epithelial cells is eosinophilic. H&E stain.

Figure 1.198   Spitting cobra, Naja sp. Elapidae. Photomicrograph of the kidney of a male snake. A cross- section of a tubule within the sexual segment (SS) in an immature snake is adjacent to a tubule of the distal segment (DT). The nuclei in cells of the sexual segment are in a basal position and positive-staining granules are within the cytoplasm around the lumen. The distal tubule also has positive-staining material within the cytoplasm around the lumen. Portions of proximal tubules (PT) also are seen. PAS stain.

Figure 1.199   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of the kidney of a male snake. Deep red-staining granules are seen within the cytoplasm of the sexual segment (SS). Masson’s trichrome stain.

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Figure 1.200   Loggerhead sea turtle, Caretta caretta. Cheloniidae. Ventral view of the posterior coelomic cavity near the pelvic canal. The urinary bladder (UB) is relatively small and has a thick wall. Also seen are the testes (TE) and colon (CO). The kidney (arrows) is covered by peritoneum.

Figure 1.201   Desert tortoise, Gopherus agassizii. Testudinidae. The urinary bladder (UB) is large, with a relatively thin wall and occupies much of the ventral coelomic cavity.

Figure 1.202   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the urinary bladder. The mucosa consists of a transitional epithelium (TE). The muscularis (MU) consists of multiple bundles of irregularly arranged smooth muscle fibers. H&E stain.

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Figure 1.203   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the urinary bladder. The mucosa consists of a transitional epithelium (TE) that lines the lumen, a lamina propria (LP) comprised of connective tissue and vessels, and a muscularis mucosae (MM). The muscularis (MU) consists of multiple bundles of smooth muscle cells. The outer surface is covered by the serosa (SE). Masson’s trichrome stain.

Figure 1.204   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the urinary bladder. The transitional epithelium (TE) of the mucosa, lamina propria (LP), muscularis mucosae (MM), and muscularis (MU) are seen. H&E stain.

Figure 1.205  Gopher tortoise, Gopherus agassizii. Testudinidae. View of the testis through a rigid endoscope inserted into the coelomic cavity through the skin surrounding a hind limb. The testis is light brown in color.

Figure 1.206   Hermann’s tortoise, Testudo hermanni. Testudinidae. View of a testis (TE) through a rigid endoscope inserted into the coelomic cavity through skin surrounding a hind limb. The TE of this tortoise is yellow and is closely associated with the kidney (KI). The adrenal gland (AD) is associated with the medial aspect of cranial pole of the kidney.

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Figure 1.207   Red-eared slider, Trachemys scripta elegans. Emydidae. Ventral view of the caudal coelomic cavity. The testes (TE) are yellow and closely associated with the kidneys (KI). The adrenal glands (AD) are paired and closely associated with the medial aspects of the cranial poles of the kidneys. Courtesy of April Johnson.

Figure 1.208   Green iguana, Iguana iguana. Iguanidae. The testes (TE) are pale and located cranial to the kidneys. The adrenal glands are closely associated with the testes. The left adrenal gland (LAD) is located in the suspensory ligament (mesorchium) between the testes and renal vein, and the right adrenal, while not visible in this image, is located on the dorsal surface of the right renal vein. The epididymis (EP) and ductus deferens (DD) are also seen.

Figure 1.209   Green iguana, Iguana iguana. Iguanidae. The right renal vein (RRV) is located between the right testes (RTE) and right adrenal gland (RAD).

Figure 1.210   Senegal chameleon, Chamaeleo senegalensis. Chamaeleonidae. The testes (TE) are located at the cranial poles of the kidneys (KI) and are covered by a black serosa/tunica. The colon (CO) is also seen. Courtesy of Rob Coke.

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Figure 1.211   Corn snake, Elaphe guttata guttata. Colubridae. Ventral view of the coelomic cavity. The testes are adjacent to the small intestine (SI). The right testis (RT) is cranial to the left testis (LT).

Figure 1.212   Corn snake, Elaphe guttata guttata. Colubridae. The testis (TE) is elongate, pale tan, and adjacent to the small intestine. The adrenal gland (AD) is thin and is located in the mesorchium.

Figure 1.213   Green iguana, iguana iguana. Iguanidae. Photomicrograph of the testis. Numerous seminiferous tubules (ST) are separated by an interstitium consisting of connective tissue, blood vessels, and interstitial cells (IC). H&E stain.

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Figure 1.214   Green iguana, iguana iguana. Iguanidae. Higher magnification photomicrograph of the testis in Figure 1.213. Spermatozoa (SP) are seen within the lumen of a seminiferous tubule. Spermatogonium (SG) are along the basement membrane of the epithelium and give rise to primary spermatocytes (PSP). Immediately following formation, the primary spermatocyte enters the prophase of the first maturation division of meiosis. Primary spermatocytes give rise to secondary spermatocytes, which are rarely seen on a light microscopic level. Secondary spermatocytes undergo the second meiotic division and give rise to haploid spermatids (SMT). Spermatids mature into spermatozoa. H&E stain.

Figure 1.215  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of a seminiferous tubule near the end of a period of spermiogenesis. A small number of spermatozoa are seen in the central lumen. Spermatogonia (SG) along the basement membrane give rise to primary spermatocytes (PSP). Secondary spermatocytes are rarely observed at a light microscopic level and give rise to haploid spermatids. A few sertoli cells (ST) are seen. H&E stain.

Figure 1.216   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the testis. Immature cells are seen within the lumina of inactive seminiferous tubules. H&E stain.

Figure  1.217   Green iguana, iguana iguana. Iguanidae. Photomicrograph of the testis. Interstitial cells (IC) have eosinophilic cytoplasm and are seen within the interstitium. H&E stain.

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Figure 1.218   Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the testis. Interstitial cells (IC) having a light-staining cytoplasm containing small eosinophilic granules are seen within the interstitium between adjacent seminiferous tubules. H&E stain.

Figure 1.219   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the efferent ductules of the testis. Spermatozoa are seen within ductular lumens. H&E stain.

Figure 1.220   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the ductus deferens. Spermatozoa (SP) are seen within the lumen. A pseudostratified epithelium (EP) lines the duct. Below the epithelium is the lamina propria (LP), submucosa (SM), and muscularis (MU). H&E stain.

Figure 1.221   Indian star tortoise, Geochelone elegans. Testudinidae. Male tortoise with the penis everted from the cloaca.

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Figure 1.222   America alligator, Alligator mississippiensis. Alligatoridae. American alligator with the penis everted from the cloaca. Papillomatous growths are present on the distal end.

Figure 1.223   Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Hemipenes are everted from the base of the tail.

Figure 1.224   Corn snake, Elaphe guttata guttata. Colubridae. Hemipenes are everted from the base of the tail.

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Figure 1.225   Gopher tortoise, Gopherus polyphemus. Testudinidae. View of an ovary (OV) of an immature tortoise through a rigid endoscope inserted into the coelomic cavity through skin surrounding a hind limb. The ovary is adjacent to the cranial pole of the kidney (KI).

Figure 1.226   Red-eared slider, Trachemys scripta elegans. Emydidae. Previtellogenic follicles in the ovary (OV) of a nonreproductive turtle. The adrenal glands (AD) are medial to the ovaries and the kidneys (KI). Courtesy of April Johnson.

Figure 1.227   Red-eared slider, Trachemys scripta elegans. Emydidae. Vitellogenic follicles in a reproductive turtle. On the right side, additional ovarian tissue (OV) can be seen at the base of the vitellogenic follicles. The following structures are also seen: left lobe of the liver (LL); right lobe of the liver (RL); uterus (UT); stomach (ST). Courtesy of John Roberts.

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Figure 1.228   American alligator, Alligator mississippiensis. Alligatoridae. Ovaries (OV) of an immature alligator are located near the cranial pole of the kidneys (KI). The KI are retroperitoneal and only the right kidney can be seen. Oviducts (OD) are adjacent to the ovaries.

Figure 1.229   Green iguana, Iguana iguana. Iguanidae. Numerous previtellogenic follicles (PVF) are seen in the ovaries of a mature green iguana. The right adrenal gland (RAD) and oviducts (OD) are also seen.

Figure 1.230   Reticulated python, Python reticulatus. Pythonidae. Multiple previtellogenic (PVF) and vitellogenic (VF) follicles are seen in the ovary.

Figure 1.231   Corn snake, Elaphe guttata guatta. Colubridae. Vitellogenic follicles (VF) and a caseated follicle (CF) are seen in the ovary of a corn snake that had yolk coelomitis. Courtesy of Stephen Barten.

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Figure 1.232   Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. The ovaries, which include numerous vitellogenic follicles, have been removed from a mature veiled chameleon with dystocia. Courtesy of Stephen Barten.

Figure 1.233   Loggerhead sea turtle, Caretta caretta. Cheloniidae. Ovary from a female that had oviductal eggs. Multiple vitellogenic follicles (VF) and corpora lutea (CL) are forming. The next progression of large vitellogenic follicles is present as is a myriad of small previtellogenic follicles (PVF). A single atretic follicle (AF) is recognized by its darker coloration and prominent vasculature. H&E stain. Courtesy of Brian Stacy.

Figure 1.234   American alligator, Alligator mississippiensis. Alligatoridae. Reproductive tract of an adult female including ovary with previtellogenic (PVF) and vitellogenic (VF) follicles, uterine tube (UT), and uterus (UTR). The two uteri empty independently into the cloaca (CL). The colon (CO) has been severed where it joins the CL.

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Figure 1.235   Green iguana, Iguana iguana. Iguanidae. Numerous vitellogenic (VI) and previtellogenic (PV) follicles are seen in the ovary of a mature green iguana. The adrenal (AD) is also seen in the suspensory ligament. Courtesy of Stephen Barten.

Figure 1.236   Siamese crocodile, Crocodylus siamensis. Crocodylidae. Photomicrograph of the ovary of a juvenile animal. All follicles are primordial follicles (PMF) and are comprised of a nucleus (NU) and ooplasm (OP) surrounded by a layer of flattened follicular cells (FC). H&E stain. Courtesy of Brian Stacy.

Figure 1.237   American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of the ovary. Early primordial oocytes (EPO), previtellogenic follicles (PVF), and lacunae (LA) are within the stroma. H&E stain. Courtesy of Mari Carmen Uribe A.

Figure 1.238   Mexican spiny-tailed iguana, Ctenosaura pectinata. Iguanidae. Photomicrograph of ovary with early primary oocytes (EPO), previtellogenic follicles (PVF), and associated stromal connective tissue (ST). H&E stain. Courtesy of Mari Carmen Uribe A.

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Figure 1.239   American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of the vegetal pole of a vitellogenic follicle. The ooplasm, which consists of vacuoles and dense yolk platelets, is surrounded by the zona pellucida (ZP), follicular epithelium (FE), and theca (TH). Masson’s trichrome stain. Courtesy of Mari Carmen Uribe A.

Figure 1.240   American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of the animal pole of a vitellogenic follicle. The ooplasm, which consists of vacuoles and more diffuse yolk platelets (arrows) than the vegetal pole, is surrounded by the zona pellucida (ZP), follicular epithelium (FE), and theca (TH). H&E stain. Courtesy of Mari Carmen Uribe A.

Figure 1.241   Mexican spiny-tailed iguana, Ctenosaura pectinata. Iguanidae. Photomicrograph of a vitellogenic follicle. Platelets of yolk are seen within the follicle. The peripheral ooplasm (PO) lacks yolk platelets. The follicular epithelium is between the zona pellucida (ZP) and the theca (TH). Gallego’s trichrome stain. Courtesy of Mari Carmen Uribe A.

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Figure 1.242   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the ovary with multiple follicles developing from the germinal bed (GB). Two early previtellogenic follicles (PVF) are present in which the granulosa (GR) is developing into a polymorphic layer. Compare these follicles with the single layer of granulosa cells in the smaller primordial follicle (PMF). H&E stain. Courtesy of Brian Stacy.

Figure 1.243   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a previtellogenic follicle. The following three recognized cell types comprise the polymorphic granulosa: small cell (SC), intermediate cell (IC), and a pyriform cell (PY). This morphology is found only in squamates. Note the zona pellucida (ZP), which separates the ooplasm and granulosa. The fibrous theca (TH) surrounds the granulosa. H&E stain. Courtesy of Brian Stacy.

Figure 1.244   Mexican spinytailed iguana, Ctenosaura pectinata. Iguanidae. Photomicrograph of previtellogenic follicle. The following three recognized cell types comprise the polymorphic granulosa: small cell (SC), intermediate cell (IC), and a pyriform cell (PY). This morphology is found only in squamates. Note the zona pellucida (ZP), which separates the ooplasm and granulosa. The fibrous theca (TH) surrounds the granulosa. H&E stain. Courtesy of Mari Carmen Uribe A. Figure 1.245   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a vitellogenic follicle. In this phase of early vitellogenesis, the granulosa (GR) is transitioning back into a single, monomorphic layer. Yolk globules are within the center of the follicle, the perivitelline zone is outside the yolk globules, and the zona pellucida (ZP) is inside the granulosa. The theca consists of an interna (TIN) and externa (TEX). H&E stain. Courtesy of Brian Stacy.

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Figure 1.246   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a vitellogenic follicle. During late vitellogenesis, the granulosa (GR) resumes a single-layered monomorphic morphology. The zona pellucida (ZP) is inside the granulosa and the theca (TH) surrounds the granulosa. H&E stain. Courtesy of Brian Stacy.

Figure 1.247   Green iguana, Iguana iguana. Iguanidae. Shelled eggs are within the oviduct and corpora lutea (CL) are seen within the ovary. Courtesy of Stephen Barten.

Figure 1.248   Green iguana, Iguana iguana. Iguanidae. Reproductive tract and ovaries surgically removed from a captive green iguana. Shelled eggs are in the oviduct and corpora lutea are seen in both ovaries (OV).

Figure 1.249   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of an early corpus luteum. The luteal cavity (LC) is lined by proliferating granulosa cells (GR) that are surrounded by a thick theca consisting of fibrous connective tissue (TH). Inset: Higher magnification image of proliferating GR and fibrous TH. H&E stain.

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Figure 1.250   Karasberg tree skink, Trachylepis sparsa. Scincidae. Corpus luteum consisting of a theca externa (TE) and granulosa cells. H&E stain. Courtesy of Stephen Goldberg.

Figure 1.251  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of the corpus luteum from a female turtle that had oviductal eggs. The corpus luteum consists of inner granulosa cells (GR) and an outer theca externa (TE). The theca interna (arrow) is a narrow area of more dense cells that separates the granulosa cells from the theca externa. Inset: Higher magnification image of granulosa cells, theca interna (arrow) and theca externa. H&E stain. Courtesy of Brian Stacy

Figure 1.252   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the corpus albicans. The corpus albicans is comprised of abundant fibrous stroma surrounding entrapped pigment-laden cells (arrows). H&E stain. Courtesy of Brian Stacy.

Figure 1.253   Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of an atretic vitellogenic follicle. This follicle is collapsed and is lined with a thick layer of vacuolated granulosa cells (arrows). The center is filled with abundant yolk. H&E stain. Courtesy of Brian Stacy.

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Figure 1.254   Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of an atretic vitellogenic follicle. The inner vacuolated cells (arrows) of the granulosa (GR) are actively phagocytizing the yolk (YK). H&E stain. Courtesy of Brian Stacy.

Figure 1.255   American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of an atretic follicle. The follicular epithelium (FE), no longer forming a distinct layer, is interspersed among the yolk (YK) platelets. A connective tissue containing theca (TH) is also seen. Gallego’s trichrome stain. Courtesy of Mari Carmen Uribe A.

Figure 1.256   Mexican spinytailed iguana, Ctenosaura pectinata. Iguanidae. Photomicrograph of an atretic follicle. H&E stain. Courtesy of Mari Carmen Uribe A.

Figure 1.257   Mexican spiny-tailed iguana, Ctenosaura pectinata. Iguanidae. Photomicrograph of an atretic follicle. Phagocytic cells (PC) are within the lumen. Masson’s trichrome stain. Courtesy of Mari Carmen Uribe A.

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Figure 1.258   Mexican spinytailed iguana, Ctenosaura pectinata. Iguanidae. Photomicrograph of the infundibulum. The lumen (LU) is lined by ciliated cuboidal epithelium and the serosal side is covered by squamous mesothelium (SM). H&E stain. Courtesy of Mari Carmen Uribe A. (From Palmer BD, et al. 1997. The Biology, Husbandry and Health Care of Reptiles, Vols I, II, and III. Ackerman L (Ed.), TFH Publications, Inc., Neptune City, NJ. With permission.)

Figure 1.259   Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of the uterine tube of a turtle during vitellogenesis. The endometrium contains numerous tubular glands (GL) containing cells with eosinophilic granules. Simple columnar epithelial cells (EP) line the lumen (LU). The muscularis (MU) is below the glandular layer. H&E stain. Courtesy of John Roberts.

Figure 1.260   Red-eared slider, Trachemys scripta elegans. Emydidae. Higher magnification photomicrograph of the uterine tube of a vitellogenic turtle seen in Figure 1.259. The endometrium contains numerous tubular glands (GL) containing cells with eosinophilic granules. Simple columnar epithelial cells (EP) line the lumen (LU). The muscularis (MU) is below the glandular layer. H&E stain. Courtesy of John Roberts.

Figure 1.261   Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of the uterine tube of a turtle that had shelled eggs in its oviduct. The lumen (LU) is lined by simple columnar epithelium (EP) and submucosal glands (GL) containing eosinophilic granules are prominent. H&E stain. Courtesy of Brian Stacy.

Overview of Reptile Biology, Anatomy, and Histology  103

Figure 1.262   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the uterus of a vitellogenic iguana. Numerous uterine glands (GL) are seen. The lumen (LU) is lined with a simple cuboidal epithelium. The myometrium (MY) varies in number of layers and thickness across the section of this image. The serosa (SE) covers the outer surface. H&E stain.

Figure 1.263  Tolucan lined ground snake, Toluca lineata. Colubridae. Photomicrograph of the uterus. The lumen (LU) is lined by a cuboidal to columnar epithelial cells (EP). Uterine glands (GL) are seen in the endometrium. Alcian blue stain. Courtesy of Mari Carmen Uribe A.

Figure 1.264   Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Photomicrograph of the uterus of a previtellogenic adult snake. Glands (GL) are scattered about the endometrium. The lumen (LU) is lined by a simple cuboidal epithelium. The myometrium (MY) surrounds the endometrium. H&E stain.

Figure 1.265   Burmese python, Python molurus bivittatus. Pythonidae. Photomicrograph of the uterus of a previtellogenic adult snake. Glands (GL) are scattered about the endometrium. The lumen (LU) is lined by a simple cuboidal epithelium. The myometrium (MY) surrounds the endometrium. H&E stain.

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Figure 1.266   Mexican spinytailed iguana, Ctenosaura pectinata. Iguanidae. Crosssection of the vagina. Sperm (SP) is within the lumen. The lumen is lined by ciliated cubodial cells (CC) and no glands are present in the submucosa. H&E stain. Courtesy of Mari Carmen Uribe A.

Figure 1.267   Gopher tortoise, Gopherus polyphemus. Testudinidae. Ventral view of the anterior coelomic cavity. The heart (HT) is located between the left lobe (LL) and right lobe (RL) of the liver. Also seen are the stomach (ST) and thymus (TH).

Figure 1.268   Boa constrictor, Boa constrictor. Boidae. The heart (HT) is cranial to the liver (LI), and the respiratory portion of the lung (LU) is between the two.

Figure 1.269   Rhinoceros viper, Bitis nasicornis. Viperidae. The apex of the heart (HT) is adjacent to the anterior pole of the liver (LI). Numerous granulomas (GR) are scattered throughout the liver. Courtesy of John Roberts.

Overview of Reptile Biology, Anatomy, and Histology  105

Figure 1.270   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of heart base. The right atrium (RA), left atrium (LA), left aorta (LAO), right aorta (RAO), pulmonary artery (PA), and ventricle (VN) are seen. A portion of the thymus (TH) and the single thyroid gland (TG) are cranial to the base of the heart.

Figure 1.271   Reticulated python, Python reticulates. Pythonidae. The left (LA) and right (RA) aortae join caudal to the heart and form a single dorsal aorta (DA). The esophagus (ES) passes through the vascular ring formed by the joining of the two aortae.

Figure 1.272   Corn snake, Elaphe guttata guttata. Colubridae. The left (LA) and right (RA) aortae join caudal to the heart and form a single dorsal aorta (DA). The esophagus (ES) passes trough the vascular ring formed by the joining of the aortae. The thyroid gland (TG) is anterior to the heart.

Figure 1.273   Green iguana, Iguana iguana. Iguanidae. The surface of the heart is covered by the epicardium (EP). The myocardium (MY) consists of bundles of myocardial cells that are separated by spaces (MS), which form the ventricular myocardial chamber.

106  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.274   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the pituitary gland. The infundibular cavity (IC), infundibular stalk (IS), pars distalis (PD), pars intermedia (PI), and pars nervosa (PN) are seen. H&E stain.

Figure 1.275   Island night lizard, Klauberina riversiana. Xantusiidae. Photomicrograph of the pituitary gland. The infundibular cavity (IC), infundibular stalk (IS), pars distalis (PD), pars intermedia (PI), and pars nervosa (PN) are seen. Alcoholic Alcian Blue-PAS-Orange-G stain. Courtesy of Hank Adams.

Figure 1.276   Neotropical rattlesnake, Crotalus durissus. Viperidae. Photomicrograph of the pituitary. The pars distalis (PD), pars intermedia (PI), and pars nervosa (PN) are seen. H&E stain.

Figure 1.277   Green turtle, Chelonia mydas. Cheloniidae. The thyroid gland (arrows) is located cranial to the heart (HT).

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Figure 1.278   Boa constrictor, Boa constrictor. Boidae. The thyroid gland (TG) is located cranial to the heart (HT) and caudal to the thymus (TH).

Figure 1.279   Corn snake, Elaphe guttata guttata. Colubridae. The thyroid gland (TG) is located cranial to the heart (HT). Also seen are the parathyroid gland (PG), posterior vena cava (PVC), and right lung (LU).

Figure 1.280   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the thyroid gland. Follicles are filled with colloid and are variable in size. H&E stain.

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Figure 1.281   Desert tortoise, Gopherus agassizii. Testudinidae. The parathyroid gland (arrow) is located on the carotid artery (CA), which is cranial to the heart (HT) and adjacent to the trachea (TR) and the multilobed thymus (TH).

Figure 1.282   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of the parathyroid gland (PG) and thymus (TH). The parathyroid gland consists of cords of cells. H&E stain.

Figure 1.283   Burmese python, Python molurus bivittatus. Pythonidae. Photomicrograph of the parathyroid gland (PG) consisting of a cluster of cells that is adjacent to and separated from the thymus (TH) by connective tissue. H&E stain.

Figure 1.284   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of the parathyroid gland (PG), which is surrounded by lobes of thymus (TH). Two cysts (CY) are seen within the parathyroid gland. H&E stain.

Overview of Reptile Biology, Anatomy, and Histology  109

Figure 1.285   Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Photomicrograph of the parathyroid gland. Several cysts (CY) are seen. H&E stain. Courtesy of John Roberts.

Figure 1.286   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the ultimobranchial body consisting of follicles (FO) and clusters of cells (CL). H&E stain. Courtesy of Tanja S. Zabka.

Figure 1.287   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the ultimobranchial body immunohistochemically stained for calcitonin with a rabbit antihuman calcitonin antibody. Immunoreactive cells have red-staining antigen in the cytoplasm. For the most part, cells lining follicles (FO) are negative. An immunoreactive cell (arrow) is at the periphery of a follicle. Immunoperoxidase stain. Courtesy of Tanja S. Zabka and Diane Naydan.

110  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.288   Loggerhead sea turtle, Caretta caretta. Cheloniidae. The adrenal gland (arrow) is located adjacent to the ovary (OV) and at the medial aspect of the cranial pole of the kidney (KI). The kidney is covered by peritoneum and cannot be seen in this image.

Figure 1.289   American alligator, Alligator mississippiensis. Alligatoridae. View of the ventral posterior coelomic cavity. The adrenal gland (AD) is elongate and is adjacent to the ovary (OV) and at the medial aspect of the cranial pole of the kidney (KI). The oviduct (OD) also can be seen.

Figure 1.290   Green iguana, Iguana iguana. Iguanidae. The ovary consists of previtellogenic (PV) and vitellogenic (VI) follicles. The adrenal gland (AD) is located in the mesovarium.

Figure 1.291   Burmese python, Python molurus bivittatus. Pythonidae. The adrenal gland is elongate and is located adjacent to the ovary and oviduct.

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Figure 1.292   Corn Snake, Elaphe guttata guttata. Colubridae. The adrenal gland (AD) is in the suspensory ligament (mesorchium) and is adjacent to the small intestine (SI) and the caudal pole of the testis (TE).

Figure 1.293   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the adrenal gland. Cords of pale-staining cortical cells (CO) and clusters of basophilicstaining chromaffin cells (CR) are seen. H&E stain.

Figure 1.294   Monocled cobra, Naja sp. Viperidae. Photomicrograph of the adrenal gland. Cords of pale-staining cortical cells (CO) and clusters of basophilic-staining chromaffin cells (CR) are seen. H&E stain.

Figure 1.295   Gaboon Viper, Bitis gabonica. Viperidae. Photomicrograph of adrenal cortical cells (CO) and chromaffin cells (CR). Cortical cells have a foamy pale-staining cytoplasm and chromaffin cells have basophilic granules within the cytoplasm. H&E staining. Figure 1.296   Timber rattlesnake, Crotalus horridus. Viperidae. Photomicrograph of adrenal cortical cells (CO) and a few clumps of chromaffin cells (CR). Cortical cells have a foamy pale-staining cytoplasm and chromaffin cells have basophilic granules within the cytoplasm. H&E staining.

112  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.297   Gopher tortoise, Gopherus polyphemus. Testudinidae. Dorsal view of the brain and anterior spinal cord (SP). Major components of the brain include the olfactory bulb (OB), telencephalon (TE), optic tectum (OT), cerebellum (CB), and medulla (ME). Calcareous material (CA) is within the otic sac, which is adjacent to the cerebellum. Nerve fibers from the olfactory sensory cells in the olfactory mucosa of the nasal cavity (NC) travel to the olfactory bulb where they synapse with mitral cells.

Figure 1.298   American alligator, Alligator mississippiensis. Alligatoridae. Dorsal view of the brain and anterior spinal cord (SP). Major components of the brain include the olfactory bulb (OB), olfactory tract (OTR), telencephalon (TE), optic tectum (OT), cerebellum (CB), and medulla (ME).

Figure 1.299   Green iguana, Iguana iguana. Iguanidae. Dorsal view of the brain and anterior spinal cord (SP). Major components of the brain include the olfactory bulb (OB), olfactory tract (OTR), telencephalon (TE), pineal gland (PI), optic tectum (OT), cerebellum (CB), and medulla (ME). The olfactory nerve travels from the nasal cavity (NC) to the olfactory bulb.

Overview of Reptile Biology, Anatomy, and Histology  113

Figure 1.300   Death adder, Acanthophis antarcticus. Elapidae. Dorsal view of the brain and anterior spinal cord (SP). Major components of the brain include the olfactory bulb (OB), olfactory tract (OTR), telencephalon (TE), optic tectum (OT), cerebellum (CB), and medulla (ME). The olfactory nerve travels from the nasal cavity (NC) to the olfactory bulb.

Figure 1.301   Burmese python, Python molurus, Pythonidae. Dorsal view of the brain and anterior spinal cord (SP). Major components of the brain include the olfactory bulb (OB), olfactory tract (OTR), telencephalon (TE), optic tectum (OT), cerebellum (CB), and medulla (ME). The olfactory nerve travels from the nasal cavity (NC) to the olfactory bulb.

Figure 1.302   Green iguana, Iguana iguana. Iguanidae. Subgross sagittal section of the head. The following structures are seen: cerebellum (CB), diencephalon (DI), medulla (ME), nasal cavity (NC), olfactory bulb (OB), olfactory nerve (NI), olfactory tract (OTR), optic nerve (NII), optic tectum (OT), pineal gland (PI), pituitary gland (PIT), telencephalon (TE), and spinal cord (SP). H&E stain.

114  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.303   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a sagittal section through the nasal cavity mucosa (NCM) and olfactory nerves (NI). H&E stain.

Figure 1.304   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a sagittal section through the olfactory nerves (NI) and olfactory bulb (OLB). H&E stain.

Figure 1.305   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of a transverse section of the head through the nasal cavity (NC). Olfactory nerves (NI) are below the olfactory mucosa (OLM). H&E stain.

Figure 1.306   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of a transverse section of the mid-olfactory tracts. The lateral ventricles (arrows) are seen within each tract. H&E stain.

Overview of Reptile Biology, Anatomy, and Histology  115

Figure 1.307   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of a transverse section of the olfactory tract near the telencephalon. The olfactory ventricles (arrows) are also seen. H&E stain.

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Figure 1.308   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a transverse hemisection of the olfactory bulb. An olfactory ventricle is also seen. H&E stain.

Figure 1.309   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a transverse hemisection of the anterior telencephalon. The following structures are seen: dorsal cortex (DC), dorsomedial cortex (DMC), dorsal ventricular ridge (DVR), lateral cortex (LC), medial cortex (MC), septum (SP), and lateral ventricle (VE). H&E stain.

116  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.310   Green iguana, Iguana iguana. Iguanidae. Testudinidae. Photomicrograph of a transverse section of the anterior telencephalon. The following structures are seen: dorsal cortex (DC), dorsomedial cortex (DMC), dorsal ventricular ridge (DVR), lateral cortex (LC), medial cortex (MC), and septum (SP). H&E stain.

Figure 1.311   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a transverse hemisection of the caudal telencephalon (CTE), rostral optic tectum (OT), diencephalon (DI), and third venricle (V3). H&E stain.

Figure 1.312   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of a transverse section of the brain through the posterior dorsal cortex (DC), diencephalon (DI), nucleus sphericus (NS), and pineal gland (PI). The cortex consists of the three continuous layers 1, 2, and 3. The third ventricle (V3) is in the midline. H&E stain.

Figure 1.313   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a transverse section of the mesencephalon. The dorsal optic tectum (OT), ventral tegmentum (TEG), and tectal ventricle are seen. H&E stain.

Overview of Reptile Biology, Anatomy, and Histology  117

Figure 1.314   Gopher tortoise, Gopherus polyphemus. Testudinidae. Higher magnification photomicrograph of the optic tectum in Figure 1.312. Multiple alternating layers of nerve fibers and cells are seen. H&E stain.

Figure 1.315   Green iguana, Iguana iguana. Iguanidae. Sagittal section of the hindbrain. The base of the cerebellum (CB), fourth ventricle (V4), medulla (ME), and tegmentum (TE) are seen. H&E stain.

Figure 1.316   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a sagittal section through the optic tectum (OT) and cerebellum (CB). Compared to other reptiles, the cerebellum in lizards is reverse curved, with the tip pointing anteriorly. H&E stain.

118  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.317   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a sagittal section through the optic tectum. Multiple alternating layers of nerve fibers and cells are seen. H&E stain.

Figure 1.318   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a transverse section of the optic tectum. Multiple alternating layers of nerve fibers and cells are seen. H&E stain.

Figure 1.319   Green iguana, Iguana iguana. Iguanidae. Higher magnification photomicrograph of a sagittal section of the cerebellar cortex seen in Figure 1.315. The three major layers are the molecular layer (MO), Purkinje cell layer (PU), and granular layer (GR). Ependymal cells (EP) line the ventricular side of the cerebellum.

Overview of Reptile Biology, Anatomy, and Histology  119

Figure 1.320   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a transverse section of the cerebellum. The three major layers are the molecular layer (MO), Purkinje cell layer (PU), and granular layer (GR). Ependymal cells (EP) line the ventricular side of the cerebellum. H&E stain.

Figure 1.321   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a transverse section of the hindbrain through the anterior medulla (ME) and fourth ventricle (V4). H&E stain.

Figure 1.322   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of a transverse section of the posterior medulla. H&E stain.

120  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.323   Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a transverse section of the spinal cord. The following structures are seen: central canal (CC), dorsal columns (DC), lateral columns (LC), ventral columns (VC), dorsal horn (DH), and ventral horn (VH). The horns are grey matter and the axon columns are white matter. H&E stain.

Figure 1.324   Burmese python, Python molurus. Pythonidae. Photomicrograph of a transverse section of the spinal cord. Major structures are the central canal (arrow), dorsal columns (DC), lateral columns (LC), ventral columns (VC), dorsal horn (DH), and ventral horn (VH). The horns are grey matter and the axon columns are white matter. H&E stain.

Figure 1.325   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of a transverse section of the spinal cord. Major structures are the central canal (arrow), dorsal columns (DC), lateral columns (LC), ventral columns (VC), dorsal horn (DH), and ventral horn (VH). The horns are grey matter and the axon columns are white matter. H&E stain.

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Figure 1.326   Green iguana, Iguana iguana. Iguanidae. The epidermis of the interparietal scale is modified into a cornealike structure (arrow). (From Jacobson ER. 2003. Biology, Husbandry, and Medicine of the Green Iguana. Kreiger Publishing, Malabar, FL. With permission.)

Figure 1.327   Green iguana, Iguana iguana. Iguanidae. Subgross sagittal section of the brain. The following structures are seen: cerebellum (CB), diencephalon (DI), medulla (ME), olfactory nerve (NI), optic nerve (NII), optic tectum (OT), parietal eye (PE), pineal gland (PI), and telencephalon (TE). Masson’s trichrome stain.

Figure 1.328   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a sagittal section of the parietal eye (PE) embedded in cartilage in the skull. The PE is dorsal to the anterior margins of the telencephalon (TE). Masson’s trichrome stain.

Figure 1.329   Green iguana, Iguana iguana. Iguanidae. Higher magnification photomicrograph of a sagittal section of the parietal eye seen in Figure 1.327. The vitreal cavity (VC) and retina (R) are seen. Masson’s trichrome stain.

122  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.330   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the sagittal section of the roof of the diencephalon. The paraphysis (PA), dorsal sac (DS), and pineal gland (PI) are seen. These structures are caudal to the parietal eye. Masson’s trichrome stain.

Figure 1.331   Tuatara, Sphenodon punctatus. Sphenodontidae. Photomicrograph of a transverse section of the head of a neonate through the parietal eye. Overlying the parietal eye (PE) is the parietal plug (PP). Below the parietal eye is the telencephalon (TE), lateral ventricles (LV) containing the choroid (CH), superior habenular ganglia (HA), and thalamus (TH). Epidermis (EP) covers the outer upper surface. H&E stain. (From Ung C Y-J and Molteno ACB. 2004. Clin Exp Ophthamol 32:614–618. With permission.)

Overview of Reptile Biology, Anatomy, and Histology  123

Figure 1.332   Tuatara, Sphenodon punctatus. Sphenodontidae. Photomicrograph of a transverse section of parietal eye of a neonate. The following structures are seen: lens (LE), parietal plug (PP), retina (RE), and vitreal cavity (VC). (From Ung C Y-J and Molteno ACB. 2004. Clin Exp Ophthamol 32:614–618. With permission.)

Figure 1.333   Dumeril’s ground boa, Acrantophis dumerili. Boidae. Photomicrograph of the pineal gland, which is located on the roof of the brain above the diencephalon.

124  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.334   New Caledonian bumpy (gargoyle) gecko, Rhacodactylus auriculatus. Gekkonidae. Example of a gecko with no eyelids, elliptical pupils, and a spectacle. Courtesy of Nicholas Millichamp.

Figure 1.335   Leopard gecko, Eublepharis macularius, Eublepharidae. Example of a gecko with eyelids, elliptical pupils, and no spectacle. Courtesy of Nicholas Millichamp.

Figure 1.336   Bearded dragon, Pogona vitticeps. Agamidae. The pupil is round and no spectacle is present. Courtesy of Nicholas Millichamp.

Overview of Reptile Biology, Anatomy, and Histology  125

Figure 1.337.   Loggerhead sea turtle, Caretta caretta. Cheloniidae. Large salt glands (arrows) are behind each eye (not visible) and lateral to the dura covering the brain (BR).

Figure 1.338   Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph of the salt gland. Serous epithelial cells discharge their salt secretions into a central lumen, which empties into a single duct. H&E stain.

Figure 1.339   Painted wood turtle, Rhinoclemmys pulcherrima. Emydidae. The pupil is round. Courtesy of Nicholas Millichamp.

126  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.340   American alligator, Alligator mississippiensis. Alligatoridae. The pupil is elliptical. Courtesy of Nicholas Millichamp.

Figure 1.341   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of the eye. A cornea (CO), lens, periocular scale (POS), spectacle (SP), and subspectacular space (SSP) are seen. H&E stain.

Figure 1.342   Green anaconda, Eunectes murinus. Boidae. The shed spectacle (SP) can be seen in the shed skin above and behind the eye. Courtesy of Stephen Barten.

Figure 1.343   Emerald tree boa, Corallus caninus. Boidae. The shed spectacle (SP) can be seen in the shed skin above and behind the eye.

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Figure 1.344   Trans-Pecos rat snake, Bogertophis subocularis. Colubridae. This crepuscular snake characteristically has oval pupils and protruding globes.

Figure 1.345   Green tree python, Chondropython viridis. Pythonidae. The pupil is elliptical. Courtesy of Nicholas Millichamp.

128  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.346   Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a transverse section through the osseous and otic labyrinth and the middle ear cavity (ME). The specialized sensory epithelium (SE) is located within the labyrinth chamber. H&E stain.

Figure 1.347   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of a transverse section through the osseous and otic labyrinth. The vestibulocochlear nerve (NVIII) provides fibers (NF) to the specialized sensory epithelium (SE) within the labyrinth chamber. H&E stain.

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Figure 1.348   Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of a coronal section through the anterior head. The vomeronasal organ (VNO) has paired chambers and is located below the nasal cavity (NC). H&E stain.

Figure 1.349   Corn snake, Elaphe guttata guttata. Colubridae. Higher magnification photomicrograph of the vomeronasal organ (VNO) in Figure 1.348. The VNO consists of a peripheral sensory epithelium (SE) and a mushroom body (MB). The MB consists of supporting cartilage (C) and a nonsensory surface epithelium (NSE) projecting into a lumen (LU). The lumen empties into a vomeronasal duct. H&E stain.

130  Overview of Reptile Biology, Anatomy, and Histology

Figure 1.350   Copperhead, Agkistrodon contortrix. Viperidae. The pit organ (arrow) is located between the eye and naris (N). Courtesy of Darryl Heard.

Figure 1.351   Western diamondback rattlesnake, Crotalus atrox. Viperidae. Photomicrograph of a pit organ. Nerves (NE) supplied by the maxillary and ophthalmic divisions of the trigeminal nerve extend between the two layers of keratinized surface epithelium of the pit membrane (PM). The PM divides the pit into anterior (AC) and posterior (PC) chambers. H&E stain. Courtesy of John Roberts.

Figure 1.352   Emerald tree boa, Corallus caninus. Boidae. Multiple labial pits (LP) can be seen.

2 Reptile Immunology Francesco C. Origgi

Contents

2.1 General Concepts

2.1 General Concepts ....................................................... 131

The immune system is comprised of organs, structures, cells, and factors that are directly involved in the defense activity of the host against pathogens (viruses, bacteria, fungi, and parasites) and transformed cells (tumors). Evolution has finely shaped and molded this system producing very different levels of organization, spanning from the very simple and primitive systems of the most elementary multicellular invertebrate organisms, to the complex and highly specialized systems of higher vertebrates. The immune system in vertebrates is traditionally divided into innate and adaptive immunity. Adaptive immunity and parts of innate immunity are based on the recognition of nonself (not of the host) molecules or structures that have come in contact with specific receptors on immune cells. Innate immunity is generally the first to come into action because it does not require any additional activation beyond that provided by the antigen itself. This first line of defense is composed of phagocytic cells that can process (and eventually dispose of) the antigen, and of nonspecific effector molecules, such as the complement, lysozyme, opsonin, defensin, and antimicrobial peptides (Brown, 2002), which directly interact with foreign organisms and neutralize them. The elements of the innate immune system do not change or shape themselves in response to different pathogens. Instead, they recognize conserved structural motifs or biochemical pathways of different microorganisms, which have been maintained during their evolution. These molecular determinants represent the signals that are both necessary and sufficient for the activation of the innate immunity effectors. While many microorganisms are neutralized and eliminated by innate immunity alone, more virulent pathogens can efficiently escape its control. When this happens, the adaptive immunity is activated and a series of complex cellto-cell interactions takes place to set up defenses tailored

2.2 Innate Defense Mechanisms........................................132 2.2.1 Surface Barriers.................................................132 2.2.2 Nonspecific Humoral Factors .........................133 2.2.3 Nonspecific Cellular Factors.............................134 2.2.4 Monocytes, Azurophilic Monocytes,   Eosinophils, Basophils ....................................136 2.2.5 Chemical Mediators of Inflammation .............136 2.3 Specific Defense Mechanisms.....................................136 2.3.1 Lymphocytes ....................................................136 2.3.2 Lymphoid Organs ............................................137 2.3.3 Immunoglobulins (Antibodies) ...................... 140 2.3.4 Antibody Response to Antigens....................... 145 2.3.5 Cell-Mediated Immune Responses................... 146 2.3.6 Memory ............................................................ 147 2.3.7 Factors Affecting the Immune Response .......148 2.3.8 The Immune Response during Bacterial   Diseases ........................................................... 151 2.3.9 The Immune Response during Viral   Diseases............................................................. 151 2.3.10 The Immune Response during Parasitic   Diseases............................................................. 151 2.3.11 Vaccination ...................................................... 151 2.3.12 Future ............................................................... 151 References............................................................................. 151

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to the specific pathogen. The adaptive immune system is comprised of cell-mediated and humoral responses. The cell-mediated response is primarily based on the cytotoxic activity of two specialized groups of cells, cytotoxic T-lymphocytes (CD8+T-cells in humans), and natural killer cells (NK). These effector cells can target and kill infected cells via specific signals that pathogen-infected cells expose on their membranes. Effector T-cells recognize the presence of a pathogen in a host cell using their T-cell receptor (TCR). TCRs engage specific major histocompatibility (MHC) molecules (MHC class I for CD8 + T-cells), which are expressed on the cell membrane of the host cells. MHCs present specific small portions of the invading pathogen in a specialized groove that is accessible to the circulating effectors cells’ TCRs (see also Section 2.3.1). The most important function of the humoral immune response is the production of soluble molecules (antibodies or immunoglobulins), which are synthesized and secreted by the B-lymphocytes. Antibodies bind to specific structural antigens of the invading pathogen. Following stimulation by the pathogen, antibodies are synthesized and released into the bloodstream where they circulate until they encounter and bind to the specific pathogen. The humoral and cell-mediated responses interact during the immune response. The innate immune response also interacts with the adaptive immune response. After the primary immune response against the invading pathogen, a selected group of lymphocytes will form a resting memory-cell pool that will circulate in the host’s bloodstream, ready to sense the presence of the pathogen that originally evoked the primary adaptive immune response. This group of memory cells will become reactivated every time the pathogen with which they were primed is detected. The immune response is a complex process that involves specialized cells and effector molecules. Many of these interactions and effector functions are likely to exist in the reptile immune system, but very few have been investigated and even fewer have been identified. Unfortunately, after a very promising and productive research period that lasted from the early 1970s to the early 1980s (see Sections 2.3.3.2 and 2.3.4 of this chapter), investigative interest in the reptilian immune system has waned. Since then, fewer research articles have been published on this topic every year. The scientific community now officially recognizes its limited knowledge of the reptile immune system (Warr et al., 2003). Reptile veterinary medicine will not be able to call itself modern until this gap in knowledge is filled. The evolutionary position of reptiles also makes the Reptilia a very important group in comparative and evolutionary immunology. In this chapter, the most important information on reptile immunity is summarized with the hope of stimulating new interest in this fascinating and underinvestigated topic.

2.2 Innate Defense Mechanisms Innate defense mechanisms may have a passive role, such as that played by natural surface barriers such as skin and mucosal surfaces, or an active role such as that played by nonspecific humoral and cellular factors.

2.2.1 Surface Barriers 2.2.1.1 Skin and Mucosal Surfaces   Skin is the first physical barrier between the reptile and the external world. The outer keratin layer of skin found in reptiles has no equal in other vertebrates. Reptiles have an exceptionally thick keratin layer, which offers resistance against external, mechanical, and microbiological insults, and a physiological renewal of their integument called ecdysis, which is a periodic shedding of the external keratinized portion of the skin (for details see Chapter 1, Section 1.4.1 of this book). In snakes, this process is characterized by a sloughing of the superficial skin layer in a single piece over a short time period. Lizards may eliminate the old outer skin layer as a complete piece or may lose it in patches over a one- to two-week period. While some chelonians periodically shed their outer keratin, others do not. The ability to replace the skin on a regular basis is intuitively very functional as a defense strategy against diseases preferentially targeting this tissue or using the skin as a port of entry into the body. Indirect evidence of this has been given by Harkewicz (2001), who reported on a reduction of the interecdysis (rest) interval length with certain parasitic skin diseases. Reptilian mucosa generally lacks the outer keratin layer and this makes these surfaces more delicate and vulnerable to mechanical events and microbiological agents. An exception is the keratinized papillae lining the esophagus of some turtles, such as sea turtles. Still, in the healthy animal the mucosal barriers do not appear to be very vulnerable to microorganisms. Probably, a complex network of active compounds (some of those discussed below) and physiological mechanisms efficiently protect the host. An interesting example of one of these possible mechanisms is that described by Pasmans and Haesebrouck (2004). They found that Salmonella enterica serovar Muenchen was unable to invade and pass through the intestinal wall when orally administered to red-eared sliders (Trachemys scripta elegans) that were kept at 26°C. When kept at 37°C, Salmonella was able to invade the intestinal wall and to colonize the liver and the spleen of two out of six orally infected turtles. Salmonella was able to adhere mainly to the mucus of chelonian intestinal explants. Salmonella successfully adhered to the explants both at 30°C (in a higher number) and 37°C (in a lower number), but slightly more bacteria were able to invade the explants at 37°C. Thus, it appears that when red-eared sliders are maintained at an optimum temperature, Salmonella cannot invade the intestinal wall efficiently. This tem-

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perature-dependent mechanism could possibly be shared by other reptiles, and could help to explain why Salmonella is generally not a primary pathogen of the alimentary tract of many reptiles.

2.2.2 Nonspecific Humoral Factors Secretions, body fluids, and mucosal surfaces contain several nonadaptive immune compounds effective against invasive microorganisms. Some of these factors are constitutively present and ready to neutralize invading pathogens. Others are activated or freed in the body only upon detection of foreign organisms (Brown, 2002). An overview of nonspecific humoral factors known to be present in reptiles is given below.

2.2.2.1 Interferons   Interferons are a family of cytokines best known for their antiviral activity. In mammals, they are divided into type I and II interferons. Type I interferons (α, β) are comprised of several proteins having antiviral activity, while type II interferons (γ) are involved in the immune response to intracellular pathogens, macrophage activation, and CD4+ T (see Section 2.3.1) cell maturation. Interferons appear to be key modulators of the innate and adaptive immune responses during infection and inflammation. They have also been recognized as potent regulators of cell growth (Schultz et al., 2004). The antiviral effect that is induced by type I interferon is produced by blocking both viral protein synthesis (translation) and viral replication through activated degradation of viral RNA. Additionally, interferons (type I) increase the expression of class I major histocompatibility complex (MHC-I) (see below) molecules on the surface of all cells and activate natural killer cells (NK). The increased expression of MHC-I upregulates the cellular immune response against viruses (and other intracellular pathogens) through the enhanced presentation of viral peptides to cytotoxic T-cells. The existence of an acid-stable, heat-resistant, 33-kDa soluble factor with interferon (IFN)-like antiviral activity in reptiles was first reported by Galabov (1981), Galabov and Savov (1973), and Galabov and Velichkova (1975), who detected an IFN-like activity in primary tortoise kidney cells (Testudo graeca) in response to viral infection (West Nile virus, Semliki Forest virus, Newcastle disease virus, and Sendai virus). Normal tortoise cells were found to be resistant to viral challenge in the presence of this factor. In a second report, Mathews and Vorndam (1982) described the production of an IFN-like factor by Terrapene heart (TH-1) cells infected with Saint Louis encephalitis virus. The physical and chemical characteristics of this factor were similar to those known for mammalian and avian IFNs.

2.2.2.2 Transferrin   Transferrins are found in the plasma of all vertebrates including reptiles. Transferrins, together with albumins, account for the 95% of the mass of small-

molecular-weight proteins in reptilian plasma. Their molecular weights span from 70 and 90 kDa and they have been reported to be similar in all major groups of reptiles (Dessauer, 1970). Transferrins have a very high binding capacity for iron, which is an essential growth element for all organisms. By sequestering iron and depriving microorganisms of this essential element for growth, transferrins have bacteriostatic and fungistatic activity. Investigations into some of the features of reptilian transferrins have been conducted by Gorman and Dessauer (1965) in the Martinique anole (Anolis roquet) and in the Grenada tree anole (A. richardi), and by George and Dessauer (1970) in colubrid snakes. Indirect evidence of the role of transferrin in the reptile innate response against pathogens has been provided by Hacker et al. (1981) and Grieger and Kluger (1978) during their investigations of the effects of Aeromonas hydrophila injections in the desert iguana (Dipsosaurus dorsalis). The iron level in the plasma decreased in bacteria-injected lizards at febrile temperatures, suggesting that the combination of low iron levels in the plasma and higher body temperatures had detrimental effects on bacterial growth (Grieger and Kluger, 1978). The plasma level of other metals such as zinc has also been investigated, but the meaning of its variation during bacterial infection is still unclear (Hacker et al., 1981).

2.2.2.3 Lysozyme   Lysozyme is an antimicrobial factor that is primarily produced by the monocyte/macrophage cell line. It degrades peptidoglycan in the bacterial cell wall and is primarily bactericidal against Gram-positive bacteria. After inoculation with Leishmania agamae, serum-lysozyme levels increase two- to fivefold in the European green lizard (Lacerta viridis) (Ingram and Molineux, 1983a), and threefold in the spiny-tailed agama (Agama caudospinosum) (Ingram and Molineux, 1983 b). In contrast, with Gram-negative bacteria, Schwab and Reeves (1966) concluded that while lysozyme may be responsible for the lysis of previously killed Escherichia coli, it was not essential for killing bacteria, and Charon et al. (1975) had similar results with Leptospira spp.

2.2.2.4 Complement   The term complement (C) is used to define a complex of 30 soluble factors and membrane-bound molecules that are part of what is known as the complement inflammatory cascade (Gasque, 2004). This cascade is phylogenetically ancient, and can also be found in invertebrates (Iwanaga and Lee, 2005). In mammals, where it has been best characterized, it is has been shown to have activities such as (I) direct killing of pathogens through the formation of the membrane–attack complex (C5b9 complex), (II) generation of chemotactic factors, (III) facilitation of the elimination of immune complexes and other toxic cell debris produced during inflammation, and (IV) enhancing adaptive immunity by lowering the threshold for activation of B-cells (mainly through the binding of C3dg to the B-cell complement receptor CR2) (Gasque, 2004). Complement cascade activation can occur through three distinct pathways called the classical,

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lectin, and alternative pathways. In general the classical pathway is antibody primed, while the lectin pathway is primed by carbohydrate-binding proteins called lectins, which bind to pathogens, which may be produced as part of innate immunity. Differently, the alternative pathway is constantly activated, but it is normally blocked by specific cell surface molecules of the host that are absent on the pathogens. The reptilian complement system is comprised of multiple isoforms of C3. This is speculated to expand their immune recognition capabilities (Brown, 2002; Sunyer and Lambris 1998). The existence of the complement-related bactericidal activity factor has been known in reptiles since the 1960s and 1970s with the publications, among others, of Schwab and Reeves (1966) and Charon et al. (1975). Similar results were obtained more recently with the work of Merchant et al. (2003/2004) where the inhibitory effect of American alligator (Alligator mississippiensis) serum against E. coli and Naegleria gruberi, was found to be likely complement mediated. Recently, possible antiviral activity of complement in American alligator serum has been investigated (Merchant et al., 2005a). Kuo et al. (2000) showed how the alternative complement pathway was responsible for the complement-mediated killing of the Lyme disease spirochete (Borrelia burgdorferi) in the western fence lizard (Sceloporus occidentalis) and the southern alligator lizard (Elgaria multicarinata). Vogel and Muller-Eberhard (1985) investigated the membrane–attack complex of the cobra complement (Naja sp.). Identification of the alternative complement activation pathway was recently reported also in the American alligator (Merchant et al., 2005b).

2.2.2.5 Other Miscellaneous Factors   Other innate defenses such as antimicrobial peptide and toll-like pathogenrecognition receptors have yet to be investigated in reptiles.

2.2.3 Nonspecific Cellular Factors 2.2.3.1 Phagocytes   Phagocytosis is the capture and ingestion of particulate material, such as bacteria and cell debris, by specialized cells. The cells that can perform phagocytosis include macrophages, neutrophils (heterophils), and dendritic cells, collectively called phagocytes (see Chapter 3, Section 3.3). The cell membrane of the phagocyte engulfs the foreign material, forming phagocytic vesicles. These fuse with cellular lysosomes, giving rise to the phagolysosome. The ingested material is degraded in the phagolysosome by enzymes and an acidic environmental pH. Killing of the phagocytosed microorganism occurs due to the generation of toxic compounds such as oxygen, hydrogen peroxide, and free radicals. Activation of the production of toxic compounds is called the respiratory burst. 2.2.3.1.1 Macrophages and Dendritic Cells   Macrophages are tissue phagocytes derived from circulating monocytes. They contribute to both innate and adaptive immunity, forming a junction between the two. Macrophages actively

phagocytize bacteria, parasites, cellular debris, and other particulate material. Following phagocytosis, the microorganisms are digested and their structural determinants are presented on MHCs to circulating T-cells to activate the adaptive response.  The phagocytic activity of reptile macrophages has been investigated by several authors. Roy and Rai (2004) showed that low levels of catecholamines could enhance the phagocytic activity of splenic macrophages of the Indian leaf-toed gecko (Hemidactylus flaviviridis). At high levels, however, they were found to act as immunosuppressors, showing how stress can influence reptile immunity. Mondal and Rai (2002b) showed that the time of exposure and the dose of glucocorticoids affected phagocytic activity and nitrite release of lipopolysaccharide (LPS)-activated splenic macrophages of Indian leaf-toed geckos. Very low concentrations (10 −13 M, roughly equivalent to 0.00005 mg/kg in an animal) of hydrocortisone sodium succinate were shown to significantly impair phagocytosis and nitric oxide production, further showing how stress can influence reptile immunity. The inhibitory activity of sex hormones on the phagocytic activity of splenic macrophages of the Indian leaf-toed geckos was also reported by Mondal and Rai (2002a), who showed that both male and female hormones significantly inhibited nitrite release, with serious impairment of the cytotoxic activity of macrophages, via a receptor-mediated system (Mondal and Rai, 1999). Interestingly, males showed a lower phagocytic index than females (Mondal and Rai, 1999), suggesting a differential sexually dependent hormonal influence. Temperature was seen to affect phagocytic activity of splenic macrophages of Indian leaf-toed geckos, with phagocytosis, phagocytic index, and cytotoxic activity of the macrophage being higher at 25°C than at higher or lower temperatures (Mondal and Rai, 2001). Pasmans et al. (2002) investigated the intracellular events that followed the phagocytosis of Salmonella enterica (ser. Muenchen) in the yellow-bellied slider (Trachemys scripta scripta). Salmonella enterica is known to survive the intracellular killing factors of macrophages in mammals and birds. The authors showed how turtle macrophages actively phagocytize Salmonella enterica both at 30°C and 37°C. Despite the initial killing of a number of bacteria, some survived the active oxygen compounds and nitrogen intermediates. The infected macrophages were eventually killed by Salmonella both at 30°C and 37°C, revealing no substantial differences between the Salmonella–host interaction of birds and mammals (endotherms) and turtles (ectotherms). Phagocytic activity has also been studied in melanomacrophages, a group of macrophages characterized by the intracellular presence of melanosomes. Johnson et al. (1999) compared the phagocytic activity of melanomacrophages obtained from turtles to that of mammalian macrophages at different temperatures using E. coli as target microorganisms. At low temperatures, turtle melanomacrophages showed more phagocytic activity than mammalian macrophages, as might be expected in a heterothermic animal.

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Dendritic cells are specialized antigen-presenting cells (APCs) that phagocytize, process, and present (via-MHCs) antigens to T-cells. Immature dendritic cells have active phagocytic activity, while mature dendritic cells lose this capacity, limiting them to presentation of antigens previously phagocytized during their immature stage. In mammals, the passage from the immature to the mature stage is also accompanied by a migration of the dendritic cells from tissues to lymph nodes, where they can present the captured antigen to a large number of lymphocytes. The first evidence for reptilian dendritic cells was provided by the work of Zapata et al. (1981a) where “macrophage dendritic cells” were identified in the area between the red and white pulp (marginal zone) of the spleen of the Caspian turtle (Mauremys caspica). These observations were confirmed in the reports of Kroese and Van Rojigen (1983) and Kroese et al. (1985) investigating the antigen-trapping activity in the spleen of the red-eared slider and of the reticulated python (Python reticulatus). Using horseradish-peroxidase (HRP)–antiHRP immune complexes, the trapping of the antigen complexes by cells with dendritic cell features were detected in the periellipsoidal lymphocytic sheath (PELS) of splenic white pulp (see Section 2.3.2.3) of the red-eared slider (Kroese and Van Rojigen, 1983) and at the periphery of the white pulp in the reticulated python (Python reticulatus) (Kroese et al., 1985). These cells showed a limited (Kroese et al., 1985) or a lack (Kroese and Rojigen, 1983) of phagocytic activity using carbon particles delivered by injection, similar to what is observed in mature dendritic cells of mammals. More recently, in the Caspian turtle, Leceta and Zapata (1991) described the presence of dendritic cells in the inner zone of PELS, while macrophages were observed in the outer zone. 2.2.3.1.2 Heterophils   The heterophil is another phagocyte (see also Chapter 3, Section 3.3.1). These cells are involved in the inflammatory response of reptiles during microbial infections, parasitic diseases, and nonspecific inflammation (Campbell, 1996), and eventually in the formation of heterophilic granulomas (Montali, 1988). The involvement of these cells in the inflammatory response has been experimentally studied in American alligators injected subcutaneously with turpentine (Mateo et al., 1984b). At 4 hours after injection, heterophils were the first inflammatory cell seen to reach the target site, while other inflammatory cells, such as monocytes, required 24 hours. Phagocytic activity of heterophils was observed by Mateo et al. (1984a) in American alligators inoculated with Staphylococcus aureus, and by Efrati et al. (1970) in lizards using inert substances. There is little to no information concerning the metabolic events at the intracellular level leading to killing of phagocytosed microorganisms.

2.2.3.2 Phagocytic Activation   The phagocytic activity of macrophages and heterophils can also be influenced by acting either on the target antigen (such as opsonins) or directly on the

phagocyte (such as gamma interferon). In mammals, opsonins are primarily comprised of complement factors called C opsonins (Gasque, 2004), which bind to the complement receptors of phagocytes, and immunoglobulins, in which the fragment crystallizable portion (Fc) (see Section 2.3.3) binds to Fc receptors on phagocytes. Once foreign microorganisms are covered with one or more of these compounds, phagocytic cells show enhanced phagocytic uptake and intracellular processing activity. Pasmans et al. (2001) provided direct evidence of the opsonization effect in the red-eared slider where antibody-opsonized Salmonella enterica was able to induce a higher respiratory burst in macrophages than nonopsonized bacteria. Phagocytosis and subsequent intracellular events can be also influenced by other nonopsonin factors such as cytokines. Interferons (Schultz et al., 2004) have been shown to confer killing capacity to fish macrophages against microorganisms that are resistant to nonactivated macrophages (Ellis, 2001). Duck interferon gamma has been shown to induce nitrite secretion in chicken macrophages (Schultz et al., 2004), a factor that is known to be involved in the respiratory burst of macrophages. It is likely that similar mechanisms could occur in reptiles even if direct evidence is still lacking. In fact, the work of Mondal and Rai (2001; 2002a) suggests the existence in reptiles of a functional IL-1-like molecule that could be produced by phagocytosing macrophages. IL-1 is known to be required in the IL-12-mediated interferon gamma release (Murtaugh and Foss, 2002), suggesting the possibility of enhanced activity of macrophages.

2.2.3.3 Natural Cytotoxic Cells (NCC)   Natural cytotoxic cells or natural killer cells (NK) are a subset of lymphocytes characterized by the innate ability to kill infected cells without first being primed and activated by APCs or by other immune cells. Natural killer cells engage infected cells via their Fc receptors, which bind to antibodies coating infected cells (see Section 2.3.3). This engagement starts the whole cascade of processes that will eventually end with the killing of the infected cells. This process is called antibody-dependent cell-mediated cytotoxicity (ADCC). The indirect evidence of the existence of these specific cell subsets or of a functionally similar group of cells in reptiles was initially found by Jurd and Doritis (1977) in the European green lizard (Lacerta viridis) and then better characterized by Sherif and El Ridi (1992) in the African beauty racer (Psammophis sibilans), and by Munoz et al. (2000) and Munoz and De La Fuente (2001a, b) in the Caspian turtle. The NK activity was found to be higher in winter and summer (winter and spring for thymic cells from males) than in autumn and spring in the Caspian turtle (Munoz and De La Fuente, 2001a, b; Munoz et al., 2000), while a stronger NK activity was detected in the African beauty racer in spring and autumn (Sherif and El Ridi, 1992). This suggested a seasonal variation in the number of cellular subsets that are involved in the NK activity in these two species or possibly a different cellular composition of the NK armory. Sherif and El Ridi (1992) suggested that NK activity in

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the African beauty snake might be associated with B- and T-like lymphocytes because their seasonal variation in cell numbers was directly proportional to the NK activity in this species. In contrast, Munoz and De La Fuente (2001a, b) and Munoz et al. (2000) suggested that the enhancement of NK-mediated cytotoxic activity, which occurs at the same time as lymphoid tissue involution in the Caspian turtle, could be an evolutionary adaptation to this seasonal physiologic immunosuppression.

2.2.4 Monocytes, Azurophilic Monocytes, Eosinophils, Basophils Monocytes, azurophilic monocytes, eosinophils, and basophils are additional inflammatory cells that have been described in the blood of reptiles (see also Chapter 3) (Campbell, 1996; Montali, 1988). Unfortunately, their specific functions have been poorly investigated. While monocytes have been described as an active component of granuloma formation in response to intracellular pathogens (Campbell, 1996; Montali, 1988), functional differences between monocytes and azurophilic monocytes remain unknown. More details on monocytes and azurophilic monocytes can be found in Chapter 3. Eosinophils from infected snapping turtles (Chelydra serpentina) have been reported to be able to phagocytize immunocomplexes (Mead and Borysenko, 1984), and eosinophils from a healthy young American alligator showed phagocytic and microbicidal capacity against Staphylococcus aureus (Mateo et al., 1984a). Basophils have been reported to be involved in histamine release upon stimulation (Mead et al., 1983; Sypek et al., 1984; Sypek and Borysenko, 1988).

2.2.5 Chemical Mediators of Inflammation Montali (1988) has given a thorough description of the histological features of reptilian inflammation. Unfortunately, in the reptilian host, little is known about the subcellular and intercellular aspects of this process. The chemical mediators of inflammation have been widely studied in mammals. These molecules consist of vasoactive amines (histamine, serotonin), plasma proteases (the complement system, the kinin system, the clotting system), arachidonic acid metabolites (prostaglandins, leukotrienes, and lipoxins), platelet-activating factor (PAF), chemokines and cytokines, nitric oxide, the liposomal constituents of leukocytes, oxygen-derived free radicals, and neuropeptides (Collins, 1999). Unfortunately, very little is known about the presence or activity of these chemicals in reptiles, and whether they are functionally similar to those in mammals.

2.3 Specific Defense Mechanisms 2.3.1 Lymphocytes Lymphocytes are components of adaptive immunity. They are traditionally divided into two groups, B- and T-lympho-

cytes. The B and T letters are derived from the organs where these two lymphocytes subsets mature, the bursa of Fabricius (birds) and the bone marrow (mammals) for the B-cells and the thymus for the T-cells (mammals). In actuality, both B- and T-cells originate in the bone marrow (mammals), and only later do T-cells migrate to the thymus where they replicate and complete their maturation process. At the end of this period, both B- and T-cells leave their maturation sites and enter the blood stream and reach the peripheral lymphoid organs and tissues (lymph nodes, gut-associated lymphoid tissue [GALT], bronchial-associated lymphoid tissue [BALT], spleen), where they are more likely to encounter their specific antigen. Lymphocytes keep recirculating from peripheral lymphoid organs to the blood until they encounter their specific antigen. This event leads to activation of B- or T-cells. T-cells are divided into two major subsets: cytotoxic Tcells (TC) and helper T-cells (T H). These two cell populations are also characterized in mammals by the presence of two distinctive cell surface markers named CD4+ (T H) and CD8+ (TC) T-cells. CD4+ T-cells are a very specialized group of cells that have a number of very important functions, such as the activation of naive B-cells, CD8+ T-cells, and macrophages. Additionally, CD4+ T-cells are involved in regulation of the adaptive immune response. The CD4+ T-cell population is comprised of two additional subpopulations: T H1 and T H2 CD4+ T-cells. As a general rule, T H1 cells are critical for enhancing the immune response against intracellular pathogens, while for extracellular pathogens, T H2 are key components of the immune response. TC (CD8+) recognize peptides of a pathogen that are presented externally by MHC-I. Viral proteins produced in an infected cell are degraded by a multicatalitic protease complex called the proteosome. The resulting peptides of the viral agent are then translocated into the endoplasmic reticulum where they are loaded on the MHC-I to be expressed on the cell surface. The foreign peptides embedded in a specialized groove on the MHC-I can be detected by the T-cell receptor (TCR) on the CD8+ T-cells. When an activated CD8+ T-cell encounters the specific target, it attaches to the target cell and releases a number of factors (mainly perforin and granzyme), which generate holes in the cell membrane, leading to cell death. CD4+ T-cells can also recognize antigenic peptides exposed in the context of MHC molecules as CD8+ T-cells do, but some differences exist. While CD8+ T-cells recognize the foreign peptide when it is bound to the MHC class I molecules, CD4+ T-cells recognize peptides bound to MHC class II molecules. B-cells are the lymphoid population orchestrating the humoral immune response. B-cells can sense the presence of antigens through immunoglobulins on their surface. These membrane-bound immunoglobulins are the antigen-committed B-cell receptors, the counterpart of the T-cell TCR. They have been shown to exist on reptilian B-like lymphocytes (Fiebig and Ambrosius, 1976; Mead and Borysenko, 1984; Sherif and El Ridi, 1992). Once a B-cell has encountered its

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specific antigen, it usually requires a second signal from a CD4+ T-helper-cell to become activated and able to produce antibodies. The activation of the B-cell leads to the clonal expansion of that cell to create a subset of B-cells specific for the antigen. These activated and expanded B-cells will further mature to plasma cells (see Chapter 3, Section 3.3.5), which will start to produce and release antibodies into the environment directed against the priming antigen. Following resolution of the infection, activated T- and B-cells that have been committed to the specific pathogen will form circulating memory cells, which will be reactivated when the host encounters the same antigen.

2.3.1.1 Functional B- and T-like Activity in Reptiles   The existence of different populations of lymphocytes (B- and Tlike) in reptiles has been indirectly documented in several studies, but has never been conclusively demonstrated. Nevertheless, several experimental studies strongly suggest the organization of the circulating pool of lymphocytes into two major populations of B- and T-like cells, similar to what is known for mammals and birds. Among others, Cuchens and Clem (1979a, b) assessed the existence of two distinct functional B- and T-like populations of lymphocytes in the American alligator. The differential response to known T- and B-cell mitogens was used to functionally distinguish between these two cell populations. Additionally, a rabbit polyclonal antibody directed against alligator immunoglobulins was used to more specifically identify and distinguish the immunoglobulin-producing cells (B-like) from other lymphocytes. Pillai and Muthukkaruppan (1977, 1982) characterized lymphocytes in the variable agama (Calotes versicolor) through the specific detection of rosette-forming cells (RFC) and plaque-forming cells (PFC), which are presumably T- and B-cell derived, respectively, following immunization with sheep erythrocytes (SRBC). Evidence of the existence of T-cell-like subsets of lymphocyte populations in reptiles has been reported by several authors. El Masri et al. (1995) discussed the existence of four subpopulations of T-like lymphocytes in the ocellated skink (Chalcides ocellatus), which were phenotypically distinguishable either by the presence or absence of two surface antigens (Theta antigen [Thy-1] and Peanut agglutinin receptor [PNA]). Other indirect evidence suggesting the existence of a subset of TH-like cells was given by El Ridi et al. (1981) during the evaluation of the effect of seasons on the immune system of the schokari sand racer (Psammophis schokari). While the lymphoid component of the spleen, thymus, and part of the GALT were well developed in autumn and spring, these tissues were markedly depleted of their lymphoid elements during the summer and the winter. The humoral response to different antigens, such as rat erythrocytes (RRBC), human serum albumin (HSA), or polyvinyl pyrrolidone (PVP), was strong during the spring and the autumn. In contrast, during the summer the humoral response to RRBC and HSA was weak. Interestingly, the humoral response was strong against

PVP, a known T-cell-independent antigen in mammals. These results were supportive of the existence of T-like helper cell activity, likely to be present in the autumn and the spring, but absent during the other seasons due to reduction or disappearance of this specific subset of T-like lymphocytes. More recently, indirect evidence of T-cell activity has also been documented in the tuatara (Sphenodon punctatus) (Burnham et al., 2005). The authors detected peripheral blood mononuclear cell (PBMC) proliferation following stimulation with T-cell mitogens (ConA, PHA). Evidence of different B-like subsets can be found in studies by Munoz and De la Fuente (2000; 2001a). The authors observed seasonal variability in cell adherence (B-cell related function) and cell proliferation to pokeweed mitogen (PKW; a B-cell mitogen) in the Caspian turtle. In contrast, there was a constant proliferation response year-round when lipopolysaccharide (LPS; another B-cell mitogen) was used. These results suggest the existence of a subset of B-like lymphocytes that undergo variation in their number during the year, different from a year-round stable (LPS-sensitive) subset.

2.3.2 Lymphoid Organs The lymphoid system of reptiles is comprised of major organs and structures such as the bone marrow, thymus, and spleen. Other accessory secondary lymphoid organs include the GALT, BALT, and other lymphoid aggregates.

2.3.2.1 Bone Marrow   The bone marrow is one of the major lymphopoietic and hemopoietic organs. In mammals, the bone marrow is the site where hemato-, myelo- and lymphopoiesis occur, and where B-lymphocytes mature. In reptiles, bone marrow is located in the marrow cavities in: (1) certain skull bones; (2) long bones of lizards, chelonians and crocodilians; and (3) in the marrow cavities of the ribs and vertebrae of snakes (Figures 2.1–2.3). Additionally, in chelonians it can be found in the plastron, carapace, and pelvis (Garner, 2006). The investigation of the hemopoietic activity of the reptilian bone marrow categorizes reptiles as a transitional group between amphibians, where the spleen is the major erythropoietic organ, and birds, where the bone marrow is practically the only blood-forming organ (Cooper et al., 1985; Zapata et al., 1981b). In the regal horned lizard (Phrynosoma solare), the spleen is the primary blood-forming organ, but in most lizards it is the bone marrow. In turtles, red cell formation is shared almost equally between spleen and bone marrow (Jordan, 1938; Zapata et al., 1981b). The bone marrow, obtained from the tibia and femur of the Spanish wall lizard (Podarcis hispanica), is the chief blood-forming organ of this reptile (Zapata et al., 1981b). A few primitive cells (hemopoietic stem cells), along with erythroid cells at different stages of maturation, granulocytic, and rare lymphoid cells were observed in the lizard bone marrow. Extravascular granulocytopoiesis and intravascular erythropoi-

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esis was described to occur in a stroma composed of reticular cells and venous sinuses. Mature and immature plasma cells were also detected in the bone marrow of the Spanish wall lizard, demonstrating plasmacytogenic capacity in the lizard at this site (Zapata et al., 1981 b). Electron and light microscopic investigation of the bone marrow of snakes (Bothrops jararaca, Bothrops jararacusu, Waglerophis merremii, Elaphe taeniura taeniura, Boa constrictor, and Python reticulatus) revealed the presence of hematopoietic marrow in the vertebrae (dorsal part of the vertebrae in the neural spine, both ends of the neural arch, ridge of the anterior wall of the vertebral canal) and in the ribs (Sano-Martins et al., 2002). Protrusions, filled with mature and immature blood cells, were directed toward the lumen of almost all these sinuses. The authors hypothesized that blood cells would be released from the extravascular space into the lumen of venous sinuses. Most of the new blood cells would enter into the systemic circulation following the rupture of these blood-cell-filled protrusions. Minor proportions of new blood cells would be released into the general circulation by transcytosis. In the desert tortoise (Gopherus agassizii), bone marrow samples were collected from the nuchal, anal, cranial, and caudal marginal and costal scutes of the carapace, and from the gular, bridge, femoral, and anal scutes of the plastron (Garner et al., 1996). Samples were also collected from the femur, humerus, and the pelvis. Marrow stroma was comprised of a fibrous network of reticular cells admixed with fat, arterioles, venules, nerves, and numerous thin-walled, endothelial-lined blood sinuses. Within the extravascular spaces, melanophores and melanocytes could be observed along with granulocytes with eosinophilic granules (heterophils and eosinophils could not be distinguished), granulocyte precursors, and mononuclear cells. In contrast, erythrocyte precursors were observed in blood sinuses. Granulocytes with eosinophilic granules were the most common leukocytes found in the marrow, while plasma cells accounted for less than 1% of all cells. Erythrocyte precursors could be observed either lining the internal surface of blood sinuses (early precursors), or within the lumina of the sinuses (later maturation stages). Basophils were rarely seen in extravascular regions.

2.3.2.2 Thymus   The thymus is unique to vertebrates (Bockman, 1970) and is the organ where T-lymphocyte cells mature to become fully competent. The thymus cellular components play a critical role in the functional maturation of T-cells in mammals. While it is unknown whether a similar process occurs in reptiles, the functional evidence of the existence of an MHC system in reptiles (Farag and El Ridi, 1985, 1990; Saad and El Ridi, 1984) analogous to that of mammals suggests that a similar chain of events should occur in reptiles. 2.3.2.2.1 Anatomy   The reptilian thymus has been investigated in the major groups of reptiles, and embryologic (Cooper et al., 1985) and morphologic (Bockman, 1970) differences have been seen.

In chelonians, the thymus is located cranial to the heart near the division of the subclavian and common carotid artery (Figures 1.263, 1.278, 2.4, 2.5, 4.39). The thymus of crocodilians and alligators is more similar to that of birds than to that of other reptilian groups (Bockman, 1970). An enlarged posterior extremity is located immediately cranial to the heart, while another narrower portion projects anterior in the cervical region, toward the base of the skull. The crocodilian thymus is in close proximity to the jugular vein, the vagus nerve, and the common carotid artery (Van Bemmelen, 1888). In lizards and snakes, the thymus is located immediately cranial to the heart, closely associated with the common carotid artery, jugular vein, and vagus nerve (Figure 1.274) (Cooper et al., 1985). The number of thymic lobes in lizards and snakes is variable (Bockman, 1970). The thymus of reptiles is surrounded by a capsule of dense connective tissue (Figure 1.281). Septa extend from the capsule and subdivide the thymus of chelonians into distinct lobules (Figures 1.281, 2.6). In chelonians, some species have lobules consisting of an outer cortex and inner medulla (Figures 2.6–2.7), while in others this division is not apparent or varies in appearance seasonally (Figures 2.8– 2.9) (Cooper et al., 1985). Lobulation is lacking in crocodilians, lizards, and snakes (Bockman, 1970). Thymocytes and epithelial cells are the most common cell types in the reptilian thymus (Figure 2.10). The epithelial cells are distributed in both the medulla and the cortex, and can occur singly or in aggregates (nests). Myoid cells are another cell type in the thymic parenchyma (Figures 2.11–2.12). Cross-striations resembling skeletal muscle may be seen in some myoid cells (Bockman, 1970). Granulocytes with eosinophilic granules and others with basophilic granules can also be seen in the thymus (Bockman, 1970). Thymic cysts (Bockman, 1970) or epithelial cysts are additional structures in the thymus (Figure 2.13) (Cooper et al., 1985). Cysts are always associated with epithelial cells and can be either intracellular or extracellular (Bockman, 1970). Their origin and function are unclear. 2.3.2.2.2 Thymic Involution   The thymus undergoes a cyclic, reversible seasonal involution associated with a severe depletion of lymphoid cells and an increase of connective tissue (Figure 2.14). This involution tends to progress with age, where the lymphoid population is constantly decreasing season after season, never reaching the density seen in the young individual (Bockman, 1970). Involution has also been described in starved or diseased reptiles. This kind of involution can revert once the source of stress is removed (Cooper et al., 1985) (also see Section 2.3.6.7.2).

2.3.2.3 Spleen   The spleen has been investigated in less then 30 of the more than 7,500 species of extant reptiles (Tanaka, 1998). The reptilian spleen can be oval, spherical, or have an elongated shape. It is located in the abdominal cavity in close association with the pancreas. In some species

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this association is so close that the two organs are identified as a single anatomical entity called the splenopancreas (Figures 1.127, 1.130–1.133, 2.15–2.16). The spleen consists of the following major anatomic components: a connective tissue capsule, trabeculae extending into the pulp (in most species), red pulp, and white pulp (Figures 2.17–2.19). The white pulp represents the immunologically active component of the spleen. It is well developed and undergoes seasonal variation (Tanaka, 1998). The general structure of the spleen is similar in chelonians, crocodilians, and lizards, with differences seen in snakes. The spleens of several chelonians have been investigated including the Chinese soft-shelled turtle (Pelodiscus sinensis) (Murata, 1959), Japanese box turtle (Mauremys japonica) (Murata, 1959; Tanaka, 1998), snapping turtle (Borysenko and Cooper, 1972; Borysenko, 1976a, b), red-eared slider (Kroese and Van Rooijen, 1982, 1983), Caspian turtle (Leceta and Zapata, 1985, 1991; Zapata et al., 1981a), and yellow-marginated box turtle (Cistoclemmys [Cuora] flavomarginata) (Tanaka, 1998). Trabeculae projecting from the capsule into the parenchyma may or may not be present (Tanaka, 1998). The white pulp is composed of two types of lymphoid aggregates that surround vascular elements. Arterioles in the white pulp are surrounded by a sheath of lymphocytes called periarteriolar lymphoid sheaths (PALS). These contain immunoglobulin-negative lymphoid cells, mature and immature plasma cells, and interdigitating cells (Leceta and Zapata, 1991). As central arterioles branch, they give rise to capillaries that are surrounded by a cuff of reticular tissue (ellipsoid), which is in turn surrounded by lymphoid tissue, the periellipsoidal lymphoid sheaths (PELS). Using hematoxylin and eosin staining, PALS and PELS are difficult or impossible to distinguish (Figure 2.19). However, at least in the red-eared slider (Kroese and Van Rooijen, 1982), they can be distinguished with a trichrome stain for smooth muscle cells and a silver impregnation stain for reticular fibers. Using a trichrome stain, smooth muscle cells can be seen in the wall of arterioles (PALS) but not in capillaries (PELS) (Figures 2.20–2.21). In red-eared sliders, the PALS are situated in a network of reticular fibers while in PELS this network is reduced or absent (Figures 2.22–2.23). While PALS and PELS can be distinguished by trichrome staining in desert tortoises, both have reticular fibers in the lymphoid sheaths, so silver staining is not useful for differentiation. In the Caspian turtle, PELS are separated into inner and outer zones by a discontinuous layer of reticular cell processes (Leceta and Zapata, 1991). Surface immunoglobulin-positive lymphocytes and dendritic cells are predominant in the inner zone, and macrophages, cytoplasmic immunoglobulin-positive cells, and Ig-negative lymphocytes are present in the outer zone (Leceta and Zapata, 1991). With light microscopy, the border between the white and red pulp is not well defined because many lymphocytes and granulocytes extend into the red pulp (Tanaka, 1998). No germinal centers have been observed in chelonian spleens (Leceta and Zapata, 1991), even following

antigenic stimulation (paratyphoid vaccine) (Kroese and Van Rooijen, 1982). The red pulp surrounds the white pulp and is composed of pulp sinuses and pulp cords (Figures 2.17– 2.19). Pulp sinuses are delicate structures consisting of flattened endothelial cells surrounded by a thin collagen layer (Tanaka, 1998). The pulp cords comprise the intervascular tissue and contain inflammatory cells such as lymphocytes, macrophages, plasma cells, granulocytes, and structural elements such as interstitial cells (Tanaka, 1998). The role of the spleen in erythropoietic activity is still controversial, even though some authors (Murata, 1959) seem to exclude this possibility. Among crocodilians, the anatomy of the spleen has been described for the American alligator (Dittman, 1969). As in chelonians, the spleen is surrounded by a connective tissue capsule, which in adults extends into the interior as connective tissue trabeculae. The white and red pulps are distinguishable, and the lymphoid tissue of the white pulp forms PALS and PELS (Figures 2.24–2.25) (Tanaka, 1998). Descriptions of the lizard spleen include those of the regal horned lizard (Jordan and Speidel, 1929), Japanese fivelined skink (Eumeces latiscutatus) (Kanesada, 1956; Murata, 1959), shingleback skink (Tiliqua rugosa) (Wetherall and Turner, 1972), variable agama (Kanakambika and Muthukkaruppan, 1973), Egyptian mastigure (Uromastyx aegypticus), rainbow skink (Mabuya quinquetaeniata) (Hussein et al., 1978b), sandfish (Scincus scincus) (Hussein et al., 1979b), ocellated skink (Chalcides ocellatus) (El Deeb et al., 1985; Hussein et al., 1978a; Saad and Bassiouni, 1993), starred lizard (Agama stellio) (Saad and Bassiouni, 1993), common agama (Agama agama), and Kishinouy’s skink (Eumeces kishinouyei) (Tanaka, 1998). A thin fibrous capsule surrounds the spleen (Figure 2.26), and fibrous trabeculae separate the parenchyma into lobules in some lizards such as the regal horned lizard (Jordan and Speidel, 1929), while others lack them (Tanaka, 1998). White pulp, which may have a nodular appearance, surrounds arteries (Hussein et al., 1978b; Murata, 1959; Wetherall and Turner, 1972) (Figures 2.26–2.27). Periarteriolar lymphoid sheaths (PALS) and PELS have also been described in lizard spleens (Tanaka, 1998). Nodular structures similar in appearance to germinal centers were observed in regal horned lizards (Jordan and Speidel, 1929) and in Kishinouy’s skink (Tanaka, 1998). The red pulp is present but it has been described as poorly developed in some lizards such as the green iguana (Figures 2.26–2.27) and the common agama (Tanaka, 1998). In the tuatara, the white and the red pulp of the spleen have been identified, but no structures resembling germinal centers have been described (Marchalonis et al., 1969). In general, the spleen of the tuatara resembles that of lizards (Tanaka, 1998). Descriptions of the spleens of snakes include those of the grass snake (Natrix natrix) (Hartmann, 1930), Japanese fourlined rat snake (Elaphe quadrivirgata) (Murata, 1959), diadem snake (Spalerosophis diadema) (Hussein et al., 1979a), scho-

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kari sand racer (El Ridi et al., 1981), reticulated python (Kroese et al., 1985), and Japanese rat snake (Elaphe climacophora) (Tanaka and Hirahara, 1995). As in other reptiles, a connective tissue capsule surrounds the spleen. The snake spleen is unique because it consists almost entirely of white pulp (Hartmann, 1930; Murata, 1959) (Figures 2.28–2.30). Within the white pulp are lymphocytes, reticular cells, and macrophages (Hussein et al., 1979a). In the Japanese rat snake, the red pulp is replaced by a perilymphoid fibrous zone (PLFZ) (Tanaka and Hirahara, 1995). The PLFZ has many small venous vessels (Tanaka and Hirahara, 1995) and is poorly separated from the white pulp because of the large number of infiltrating lymphocytes and interstitial cells (Tanaka, 1998) (Figure 2.28). No lymphoid structures resembling PALS appear around the septal arterioles. However, multiple lymphoid lobules, which are separated out by the septa and the PLZF, have been observed. These lobules contain round lighter zones resembling germinal centers (Tanaka, 1998) (Figure 2.28). Very limited information is available concerning T- and Blike lymphocyte distribution in the reptile spleen. Pitchappan and Muthukkaruppan (1977) showed that in both thymectomized and antithymocyte serum-treated variable agamas it was possible to obtain lymphoid cell depletion in PALS. Because PALS were repopulated by lymphocytes in sham-thymectomized control lizards only, and not in thymectomized lizards, these areas were considered to be thymus dependent. More recently, Leceta and Zapata (1991) reported the presence of immunoglobulin-negative lymphoid cells along with immature and mature plasma cells in the PALS of the Caspian turtle, with a few immunoglobulin-positive cells in the periphery. In contrast, immunoglobulin-positive cells were predominant in the inner region of PELS. 2.3.2.3.1 Spleen Function and Seasonal Variation   There are very few studies investigating reptilian splenic function following antigenic stimulation. Following antigenic stimulation with keyhole limpet hemocyanin (KLH), a strong proliferative response resulting in an increase in white pulp mass was reported in the snapping turtle. This active proliferation was first observed in the white pulp (8 to 10 days after immunization) and later in the red pulp (15 to 20 days after immunization) (Borysenko, 1976a, b). No histological changes could be observed in the spleen of either the snapping turtle or the variable agama after a second immunization (Kanakambika and Muthukkaruppan, 1973, Borysenko, 1976a, b). The red pulp of the spleen of the schokari sand racer is well developed in winter, while only a scarce number of lymphoid aggregates can be observed in the white pulp. In spring, the lymphoid aggregates increase their size and their number, becoming denser and almost confluent. By the end of June and through July, the splenic lymphoid tissue starts to regress slowly in size with further regression in August (El Ridi et al., 1981). The humoral response to different antigens (PVP, RRBC, HAS) is strong in spring and autumn, when the lymphoid tissue population is abundant, while it is

poor in winter and in summer, when the lymphoid population is reduced. However PVP, a T-cell-independent antigen, also evoked a strong response during the summer (but not in winter) (El Ridi et al., 1981).

2.3.2.4 Lymphoid Aggregations and Lymphoid Accessory Structures   Lymphoid tissues that are not part of the major lymphoid organs (bone marrow, thymus, and spleen) are grouped together. In mammals these aggregates include the gut-associated lymphoid tissue (GALT) (comprising the adenoids, the tonsils, and the Peyer’s patches) the bronchial associated lymphoid tissue (BALT), and the lymph nodes. These structures are in areas of the body where immune cells are most likely to encounter an antigen. Several of these lymphoid structures have also been described in reptiles. 2.3.2.4.1 GALT   The GALT is distributed along the digestive tract, and is the largest lymphoid structure in the entire body. In humans, circulating lymphocytes account for only 2 to 5% of the total population, with the large majority found in the gastrointestinal tract (Mehandru et al., 2004). In reptiles, GALT can be observed along the entire gastrointestinal tract, with major accumulations located in the esophagus, ileocaecal junction, colon, and cloaca. For the most part, these aggregates are generally small, nonencapsulated, and extend from the lamina propria to the submucosa. (Cooper et al., 1985). In boid snakes, unique esophageal tonsils have been described (Jacobson and Collins, 1980) (see Chapter 1; Figures 1.08– 1.110). Esophageal tonsils have an ellipsoid shape, are slightly raised, and have a central cleft where the mucosal epithelium invaginates into a submucosa having abundant blood vessels and lymphoid cells. In reptilians, GALT lymphoid cells and plasma cells are present (Zapata and Solas, 1979; Solas and Zapata, 1980), and unlike the lymphoid components of the spleen and thymus, GALT does not seem to undergo seasonal variation (Cooper et al., 1985). However, some exceptions have been reported (Hussein et al., 1978a,b). 2.3.2.4.2  Lymph Nodes and Other Lymphoid Structures   In reptiles there are no lymph nodes that are comparable to those seen in mammals. Nevertheless, lymph node–like structures have been described, along with ectopic lymphoid tissue, in different organs such as the lung, kidney, urinary bladder, pancreas, and testes (Cooper et al., 1985). Lymph node–like structures have also been described in perivascular areas of different reptiles (Cooper et al., 1985) suggesting a possible, but not proven, functional homology with lymph nodes of mammals.

2.3.3 Immunoglobulins (Antibodies) Immunoglobulins (or antibodies) are glycoglycolproteins secreted by B-cell lymphocytes in response to infection. They have a conserved structure that resembles a “Y.” In general immunoglobulins are composed of two pairs of identical light and heavy chains that are held together by non-covalent forces

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Diagram 2.1  Mammalian (human) immunoglobulin structure. A representation of the structure of mammalian (human) immunoglobulins. Constant and variable domains of light and heavy chains are depicted in different colors. IgM here is depicted as a monomer form.

Diagram 2.2  Avian immunoglobulin structure. A representation of the structure of the avian IgY and IgY∆Fc types is shown. Constant and variable domains of light and heavy chains are depicted in different colors. IgY∆Fc lacks the heavy chain constant domains 3 and 4 that are present in the long IgY form. Reptilian IgY and IgY∆Fc are likely to be similar in structure.

and di-sulfide bonds (Diagrams 2.1–2.4). They are joined by a flexible stretch of polypeptide chain known as the hinge region, which in mammalian immunoglobulins gives flexibility to the molecule. This flexibility allows the molecule to reach and then conform to specific antigenic determinants (Diagrams 2.1–2.2). Both light and heavy chains are composed of variable and constant domains. Variable domains are those interacting with the antigens, while constant domains are those that are structural. Light chains are composed of two domains (variable light [VL] and constant light [CL]) while the heavy chains are composed of 4 (human IgG, IgD, IgA) or 5 (human IgM and IgE) domains (a variable heavy [VH] and the additional 3 or 4 constant heavy [CH1, CH2, CH3, and CH4]) (Diagrams 2.1–2.2).

2.3.3.1 Immunoglobulins of Mammals   In mammals, immunoglobulins consist of 5 different classes of heavy

chains. These classes or isotypes are IgG, IgA, IgM, IgD, and IgE (Diagram 2.1). In contrast, there are only two different classes of light chains, kappa and lambda. Each immunoglobulin is characterized by a pair of identical heavy chains, which define its isotype, and a pair of identical light chains, either lambda or kappa. While the overall structure of immunoglobulins is very conserved, they have hypervariable regions involved in antigen binding (VH and VL), also known as complementary determining regions (CDRs). These regions each have a distinct amino acid sequence, which is the product of recombination events that occur in the B-cell during maturation. This process, occurring in each maturating Bcell, results in a large array of immunoglobulins, each having a specific binding affinity for one molecular conformation. Each B-cell carries one membrane-bound immunoglobulin (identical to the soluble immunoglobulins that the B-cell is committed to synthesize following activation) mounted on its surface, which may match the complementary molecular

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Diagram 2.3  Papain immunoglobulin degradation. The products of proteolytic activity of papain on immunoglobulin is shown. Papain proteolytic digestion determines separation of the different functional structure of the immunoglobulin: the fragment antigen binding (Fab) and the fragment crystallizable (Fc).

Diagram 2.4  Pepsin immunoglobulin degradation. The products of the proteolytic activity of pepsin on immunoglobulin is shown. Pepsin proteolytic digestion determines the separation of the two Fabs, still joined together [F(ab’)2], from the Fc that is further digested into smaller fragments.

pattern from a pathogen. When there is positive recognition, complementary stimulus is usually required and provided by T-helper cells, and the B-cell then becomes an antibodysecreting plasma cell. During production of immunoglobulins, the structure of the molecule undergoes a series of modifications. One of the most important of these changes is class switching, where the current heavy chain component is substituted with that of another class. Each immunoglobulin is first produced and released as an IgM. IgM is a pentameric molecule with high avidity. It can be detected in the plasma of recently infected individuals, and progressively disappears after the acute stage

of the disease. IgM can undergo class switching to another isotype such as IgA, a dimeric immunoglobulin generally confined to the mucosal surfaces, or to IgG, the most common circulating isotype (80%). IgM can also switch to IgE, a monomeric immunoglobulin involved in parasitic and allergic immune responses in mammals. The function of the fifth known isotype, IgD, is unknown. Antigen specificity is maintained during class switching. Following class switch, the antigen specificity is enhanced through two separate steps named somatic hypermutation and affinity maturation. Somatic hypermutation is characterized by a very high rate of point mutations selectively within the CDR. This process is influenced by evolutionary pressures favoring mutations that enhance antigen affinity. Affinity maturation selects and promotes mutations derived from the original B-cell clone with higher antigen affinity. In mammals, this process occurs within lymph nodes and germinative centers. These three key events, (1) class-switching recombination, (2) somatic hypermutation, and (3) affinity maturation are the cornerstones of the antibody-based (humoral) immune response. When mammalian immunoglobulins are digested using papain or pepsin (two proteases) it is possible to isolate their functional structures. Papain cuts mammal immunoglobulins into three portions, two fragments antigen binding (Fab) and the fragment crystallizable (Fc) (Diagram 2.3). The Fabs consist of a whole light chain (CL + VL) and part of the heavy chain containing VH and CH1. The Fc portion is composed of the remaining constant domains of the two heavy chains (CH2, CH3, and CH4 if present), including the hinge region. Fabs bind to the specific antigen, while Fc allows interaction with immune cells. In contrast, pepsin cuts the immunoglobulin molecule so that the two antigen-binding regions are still bound through the hinge [2x (CL + VL) + 2x (VH + CH1)]. This fragment is known as F(ab’)2. The rest of the immunoglobulin is further degraded by pepsin (Diagram 2.4). The Fc portion is in the stem of the Y shape of the immunoglobulin (Diagrams 2.1–2.4). It interacts with the immune system. The Fc portion is recognized by isotype-specific Fc receptors on immune cells. In mammals, IgG is recognized by Fc receptors on phagocytic cells, such as macrophages and neutrophils, which can bind and engulf pathogens coated with IgG (See Section 2.2.3.2). The Fc portion of IgE is recognized by the Fc receptors of mast cells and basophils, which respond by releasing inflammatory mediators. Another important function of the Fc portion is fixing the complement and priming the classical complement cascade, which is important for recruitment and activation of phagocytes (See Section 2.2.3.2). In mammals, the complement is primarily activated by two of the five isotypes, IgM and IgG. Because of its pentameric structure, IgM can offer multiple binding sites to complement protein C1q, resulting in better complement activation than IgG.

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Finally, the Fc portion of the immunoglobulin is critical for the transport of antibodies to places that they could not reach (mucous secretions including tears and milk, and the fetal blood circulation) autonomously.

2.3.3.2 Immunoglobulins of Nonmammalian, Nonreptilian Vertebrates   The best studied nonmammalian immunoglobulins are those of fish. In cartilaginous fish, there are three heavy chain classes, IgM, IgW, and IgNAR, and three light chain classes (Dooley and Flajnik, 2006). Five isotypes have been identified in teleost fish: IgM (analogous to mammalian IgM), IgD (analogous to mammalian IgD), IgZ, IgT, and an as yet unnamed isotype (Savan et al., 2005). The number of light chains in teleost fish varies from 1 to 4 (Saha et al., 2004; Ishikawa et al., 2004). In amphibians, there are three known heavy chain isotypes: IgM, similar to mammalian IgM; IgY, which is analogous to mammalian IgG and IgE; and IgX, which is preferentially expressed in the gut (Du Pasquier et al., 2000). There are also three light chain classes (Du Pasquier et al., 2000). Birds have three known heavy chain classes: IgM, analogous to mammalian IgM; IgA, analogous to mammalian IgA; and IgY, which is analogous to mammalian IgG and IgE (Lundqvist et al., 2006). The genes needed for recombination and somatic hypermutation of the immunoglobulin genes are present in all jawed vertebrates (Flajnik, 2002). While it is accepted that somatic hypermutation occurs in nonmammalian vertebrates including reptiles (Turchin and Hsu, 1996), it is not clear to what extent affinity maturation does. Affinity maturation is not as pronounced in amphibians as it is in mammals (Flajnik, 2002); however, there is good evidence for affinity maturation in nurse sharks (Diaz et al., 1998).

2.3.3.3 Immunoglobulins of Reptiles   Reptilian immunoglobulins have been poorly investigated, and much of the information available for these molecules has been derived from the homologous counterparts in other species. Supporting evidence that an affinity maturation–like process occurs in reptiles was provided by Ambrosius et al. (1972), Ambrosius and Fiebig (1972), and Fiebig (1972, 1973), who reported affinity maturation of antibodies from the Russian tortoise (Agrionemys [formerly Testudo] horsfieldii). Using 2,4-dinitrophenyl (DNP) as an antigen, the affinity of these antibodies increased fivefold in the primary response and 300-fold in the secondary response. In contrast with this observation, Grey (1963) reported a lack of change in the avidity of the antibodies in the painted turtle (Chrysemys picta) following immunization with KLH and bovine serum albumin (BSA). However, the data are questionable, as no secondary response was detected during this experiment. Affinity maturation was also not observed in the glass lizard (Ophisaurus sp.) immunized with DNP (Ambrosius and Fiebig, 1972; Fiebig, 1972, 1973). These observations suggest differences in the affinity maturation–like process within and between different groups of reptiles.  

There is very limited information on the structure of reptilian immunoglobulins. IgY is considered a relative of mammalian IgG and IgE (Leslie and Clem, 1969). Warr et al. (1995) refer to IgY as the low molecular weight serum antibody of birds, reptiles, amphibians, and probably lungfish. IgY has two heavy and two light chains, similar to the mammalian immunoglobulin prototype, and has a molecular mass of approximately 180 kDa. The heavy chains of IgY have one variable and four constant domains. Unlike mammalian IgG, IgY is missing the hinge region that gives flexibility to the molecule, suggesting that IgY is less flexible. A truncated IgY of approximately 120 kDa is found in anseriform birds, some reptiles, and lungfish (Warr et al., 1995). This truncated form lacks two constant domains of the heavy chains (Diagram 2.2). The molecular weights of the light and heavy chains of the truncated form of IgY of the snapping turtle are 22.5 and 38 kDa, respectively (Merz et al., 1975). Schumacher et al. (1993) determined the light and heavy chain weights of Gopherus agassizii IgY to be 27 and 65 kDa, respectively. In all orders of reptiles, at least two distinct isotypes have been detected: IgM and IgY, resembling mammalian IgM and IgG, respectively (Ambrosius, 1976). Other immunoglobulin types, which could either reflect different isotypes or different subclasses of the same isotype, seem to vary within the different reptilian groups. 2.3.3.3.1 Chelonia   Ambrosius (1976) conducted a 7-year immunization study in Testudo hermanni using pig serum proteins as the antigen. The first immunoglobulin type detected was a 19S high-molecular-weight, IgM-like immunoglobulin, followed a few months later by a 7S low-molecular-weight immunoglobulin. Both 19S and 7S antibodies were sensitive to 2-β mercaptoethanol (2-ME), a reducing agent. Following a second immunization, a second 7S low-molecular-weight immunoglobulin, which was 2-ME resistant, was detected, suggesting chemical-structural differences between the two 7S immunoglobulins. The production of the 2-ME-resistant 7S antibody increased following each additional immunization, while 19S immunoglobulin production decreased. Finally, a low-molecular-weight 5.7S 2-ME-sensitive antibody was detected after the fourth immunization. The 7S antibodies are likely to be homologous to mammalian IgG and avian IgY, and their respective sensitivity and resistance to 2-ME suggest either the existence of different isotypes or subclasses of the same isotype The 5.7S antibody is antigenically similar to the 7S antibody but lacks two C-terminal domains in the Fc region (Ambrosius et al., 1972; Warr et al., 1995). This extremely lowmolecular-weight immunoglobulin may be homologous to the truncated (approximately 120 kDa) IgY isoform of ducks (Warr et al., 1995). It is 110 to 120 amino acids shorter than the larger IgY (Cooper et al., 1985) (Diagram 2.2). Truncated IgY has agglutination activity, binds the complement, and is capable of passing into eggs (Chartrand et al., 1971). Similar antibody production in other chelonians has been observed (Cooper et al., 1985; Leceta and Zapata, 1986). More recently,

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the kinetics of antibody production was followed in green turtles using monoclonal antibodies directed against 19S, 7S, and 5.7S antibodies (Herbst and Klein, 1995). The earliest response detectable by enzyme-linked immunosorbent assay (ELISA) was IgY (7S), followed by truncated IgY (5.7 S), which required 3 to 4 or up to 8 months to be detected. Kinetics of the IgM production was more difficult to evaluate (Herbst and Klein, 1995). Immunoglobulin isotypes homologous to IgA (Vaerman et al., 1975), IgD, and IgE have not been identified yet in chelonians or other reptiles. Recently a J-chain protein, known also as a joining fragment (J), a peptide normally associated with polymeric immunoglobulins (IgA and IgM), has been identified and cloned from the red-eared slider (Iwata et al., 2002). The authors identified a 1934 base pair complementary-DNA (cDNA) clone with an open reading frame of 477 nucleotides, encoding 159 amino acids. The mature J-chain protein was determined to be composed of 137 amino acids, and the predicted amino acid sequence was highly homologous with the J-chain sequence from human (60%), mouse (61%), cow (60%), rabbit (60%), chicken (69%), brushtail possum (65%), bullfrog (47%), and African clawed frog (58%). Messenger RNA (mRNA) was found to be expressed in the lung, stomach, spleen, and intestine by northern blot (See Chapter 7), while J-chain-positive plasma cells were detected by immunohistochemistry in the intestine and spleen. This suggests the presence of a mucosal immune system utilizing J-chain-containing Igs in the turtle. The physicochemical analysis of both 7S and 17S Russian tortoise antibodies revealed a lower level of flexibility when compared to those of mammals. This lower flexibility reflects the lower freedom of intramolecular rotation of tortoise immunoglobulins in comparison with those of mammals (Zagyansky, 1973). 2.3.3.3.2 Crocodylia   High (19S) and low (7S) molecular weight antibody isotypes and two distinct light chains have been observed in crocodilians (Clem and Leslie, 1969; Saluk et al., 1970). 2.3.3.3.3 Rhynchocephalia   Two different immunoglobulin isotypes have been detected in the tuatara (Marchalonis et al., 1969). An 18S immunoglobulin resembling the mammalian IgM pentavalent structure, and a 7S molecule characterized by the same light chains of the 18S immunoglobulin but with different heavy chains (different heavy chain = different isotype) have been described. Despite the detection of two distinct antibody isotypes, functional activity was found only in the 18S Ig (Marchalonis et al., 1969). 2.3.3.3.4 Sauria   Ambrosius (1976) reported the existence of three different immunoglobulin types (isotypes) in lizards; a high-molecular-weight immunoglobulin, homologous to the IgM of mammals, along with two other low-molecular-weight immunoglobulins (7.3S and 6.8S) representing two distinct isotypes or two different subclasses within the same

isotype. Of the two, the 7.3S immunoglobulin may be homologous to the IgG of mammals and IgY of birds, while the 6.8S Ig appears to be unique to lizards (Ambrosius, 1976). Cooper et al. (1985) report that lizards possess an early high-molecular-weight antibody weighing 16 to 19S (2-ME sensitive) and a later low-molecular-weight antibody of approximately 7S, which may or may not be sensitive to 2-ME. 2.3.3.3.5 Ophidia   Snakes possess at least two distinct immunoglobulin isotypes. The high-molecular-weight immunoglobulin weighs 19.6 to 20.5S and the low-molecular-weight immunoglobulin weighs 7 to 9S (Salanitro and Minton, 1973; Portis and Coe, 1975). Transition from high to low molecular weight has been observed during the later stage of the immune response (Salanitro and Minton, 1973). Portis and Coe (1975) have also described a high-molecular-weight secretory immunoglobulin in the bile of the northwestern garter snake (Thamnophis ordinoides).

2.3.3.4 Valence and Affinity of Reptilian Antibodies   The binding properties of high- and low-molecular-weight antibodies produced against DNP conjugates by the Russian tortoise and the armored glass lizard (Ophisaurus apodus) have been investigated (Ambrosius et al., 1972; Fiebig, 1972, 1973). Functional evaluation of the IgM-like antibody of the Russian tortoise led to the determination of five binding sites for each molecule, while four to five functional binding sites were determined for the homologous molecule of the armored glass lizard (Ambrosius, 1976). The expected binding capacity of a mammalian IgM molecule is 10 binding sites (5 monomeric Ig with two binding sites each). The lower binding capacity could be interpreted either as the result of fewer binding sites or low affinity of the binding sites for the specific antigen (Ambrosius, 1976). A bivalent binding capacity has been determined for tortoise 7S and lizard 7.3S antibodies, which show low heterogeneity in binding properties (Ambrosius et al., 1972; Fiebig, 1972, 1973). The 6.8S antibody of the armored glass lizard appears to be functionally monovalent. However, the possibility of a second binding site with low affinity could not be excluded (Ambrosius, 1976). Investigation of the binding properties of chelonian 5.7S antibodies revealed bivalent binding capacity and strong binding site heterogeneity (Ambrosius, 1976). 2.3.3.5 Maturation of the Immune Response   In the Russian tortoise and the armored glass lizard, no affinity maturation was detected during IgM-like production after being injected with DNP conjugates (Ambrosius et al., 1972; Ambrosius and Fiebig, 1972; Fiebig, 1972, 1973). In contrast, while no affinity maturation was detected in the production of 7.3S and 6.8S antibodies of the armored glass lizard, a dramatic increase in the affinity of low-molecular-weight antibodies was observed in the Russian tortoise following the second immunization. Affinity of the low-molecular-weight Russian tortoise antibodies increased fivefold during the primary response

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and 300-fold during the secondary response (Ambrosius et al., 1972; Ambrosius and Fiebig, 1972; Fiebig 1972, 1973). The values were in the same range as those detected for anti-DNP antibodies from birds and mammals (fluorescence quenching experiments) (Eisen, 1964; Eisen and Siskind, 1964; Gallagher and Voss, 1969).

2.3.3.6 Functions of Reptile Immunoglobulins   The functions of antibodies include neutralization, agglutination, precipitation, opsonization, and complement fixation. Neutralization is the ability to block the interaction of viruses or toxins with their cognate receptor structures exposed on the host cell membrane. Agglutination occurs when antibodies bind to particulate antigens (whole bacteria, cells), the antigens become clumped, and consequently cannot diffuse into the organism. Agglutination induces the activation of phagocytosis that is very active on antibody-antigen complexes. Precipitation occurs when antibodies bind with soluble antigens, which then precipitate and become inactivated. The term opsonization is used to indicate the enhanced phagocytosis of antigens once opsonins (antibodies, complement fractions, and other molecules) are bound to them (see also Section 2.2.3.2). Antibody-dependent opsonization occurs through the interaction of the Fc of the antibodies with Fc- receptors, which are expressed on the cell surface of phagocytes. 2.3.3.6.1 Reptilian Antibody Neutralization   The neutralization activity of reptilian antibodies is used in the serum neutralization test (SNT; see Chapter 8) for determining exposure of chelonians to viruses (Marschang et al., 1997; Origgi et al., 2001). Serum neutralization antibodies required seven to nine weeks to be detected in the SNT setup for determining exposure to herpesvirus, which was two to five weeks longer than the time required to detect IgY by an ELISA test (Origgi et al., 2001). It is possible that the actual production of SN tortoise antibodies occurs earlier, but the detection threshold might require a higher amount of antibody, which can only be detected at later times. Another explanation is that neutralization activity is only present after antibody maturation, requiring several weeks. The ELISA detects all antiherpesvirus antibody production, which is likely to start soon after the infection, while the presence of antibodies capable of neutralization could require more time. 2.3.3.6.2 Reptilian Precipitating and Agglutinating Antibodies   The production of agglutinating and precipitating antibodies has been documented in reptiles (Ambrosius, 1976; Cooper et al., 1985). 2.3.3.6.3 Complement Activation and Opsonization of Particles  See Sections 2.2.3.2 and 2.2.2.4 of this chapter. 2.3.3.6.4 Hypersensitivity Response   No data is available for reptiles. It is documented that avian IgY can mediate anaphylactic reactions (Warr et al., 1995), but it is not known if the evolutionarily related reptilian IgY mediates the same reaction.

2.3.4 Antibody Response to Antigens 2.3.4.1 Chelonian   Pioneering studies include the work of Metchnikoff (1901), who evaluated the effects of temperature on antibody production in the European pond turtle (Emys orbicularis), and Noguchi (1903), who cross-injected the painted turtle, spotted turtle (Clemmys guttata), and Blanding’s turtle (Emydoidea blandingii) with xenogenic combinations of erythrocytes Using in vitro snapping turtle spleen explants, agglutinins against mouse erythrocytes were detected after two to eleven days, and agglutinins against sheep red blood cells were detected after four to twenty days (Sidky and Aurebach, 1968). Testudo graeca produced temperature-dependent agglutinins after immunization with Brucella abortus antigen (Maung, 1963). The highest agglutinin levels were reached 15 weeks after injection. Ambrosius and Lehman (1964, 1965) immunized Hermann’s tortoise subcutaneously with heatinactivated normal pig serum (NPS) using aluminum hydroxide as an adjuvant. An agglutination titer of 16 (agglutination units) was detected within 30 days of injection. The tortoises injected with NPS without adjuvant showed a delayed response. Tortoises kept at lower temperatures (20 to 21°C instead of 28°C) showed an even longer delay. Antibody production in Hermann’s and Russian tortoises was also assessed against DNP coupled to protein antigens. Both species produced antibodies reacting against the carrier and the DNP hapten. The maximum level of precipitating anti-DNP antibodies appeared later in the secondary immune response than that of antibodies detected by passive hemagglutination (Ambrosius and Frenzel, 1972).

2.3.4.2 Crocodylia   Metchnikoff (1901) evaluated the ability of American alligators to produce a neutralizing antibody to tetanus toxin. Lerch et al. (1967) evaluated the humoral response to KLH using ring agglutination and immunoelectrophoresis. The primary response was first detected 20 days after immunization.

2.3.4.3 Rhynchocephalia   In the tuatara, the highest antibody response to Salmonella flagellin antigen was detected 60 to 80 days after injection using a live bacteria immobilization assay (Marchalonis et al., 1969).

2.3.4.4 Sauria  The northern rock agama (Stellio caucasica) and the rapid racerunner (Eremias velox) can produce antibodies against tick-borne encephalitis virus in a temperature-sensitive manner (Vorob’eva, 1965). The absence of the production of both hemagglutinin and hemolysins in splenectomized variable agamas following SRBC inoculation (Kanakambika and Muthukkaruppan, 1972) suggests a critical role for the spleen in antibody production. There was limited but detectable antibody production found in the same lizards when immunized against the same antigen 7 days before splenectomy.

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The humoral response of the desert iguana and the common chuckwalla (Sauromalus ater [obesus]) against the H antigen of Salmonella (STH) was investigated by Evans and Cowles (1959), Evans (1963) and Evans et al. (1965). The authors detected agglutination titers ranging from 1:80 to 1:640 when the lizards were kept at 35°C. When the lizards were kept either at 25°C or 40°C, the humoral responses were reduced. The immune response of the desert iguana was also assessed by Wright and Shapiro (1973) using KLH as an antigen. The maximum titer after a single immunization, determined by a passive hemagglutination (PH) assay, was detected 15 days after injection when the lizards were maintained at 37°C. Cunningham’s skinks (Egernia cunninghami) (Tait, 1969) and shingleback skinks (Wetherall and Turner, 1972) immunized with either SRBC (Cunningham’s skinks) or salmonella, RRBC and BSA in complete Freund’s adjuvant (shingleback skinks) showed the best response when maintained at a body temperature of 30°C. Saad and El Ridi (1988) successfully induced a primary immune response in vitro by stimulating ocellated skink splenocytes using RRBC as the antigen. The response was easily detected at 10 days after stimulation when the splenocytes were maintained at 37°C.

2.3.4.5 Ophidia   There are several reports that support the presence of natural lysins and agglutinins in snake serum, but details about the nature and features of these reactions are still lacking (Cooper et al., 1985). The mole snake (Pseudaspis cana) has been shown to produce agglutinins against typhus endotoxin and meningococci in a dose-dependent manner (Grasset et al., 1935). Four weeks were necessary for the black racer (Coluber constrictor), black rat snake (Pantherophis [Elaphe] obsoletus), and western fox snake (P. vulpina) to develop antibodies directed against BSA and KLH in complete Freund’s adjuvant after a single immunization, with the titer remaining constant for three months (Salanitro and Minton, 1973). This is different from the ringneck spitting cobra (Hemachatus hemachatus), which required two additional boosters following primary immunization in order to produce an antityphus endotoxin agglutination titer of 1:600 (Grasset et al., 1935). This suggested either a low sensitivity of the snake to this antigen or procedural problems (Cooper et al., 1985). Low doses of antigen were also not efficacious in eliciting a detectable humoral response in the common night adder (Causus rhombeatus) (Grasset et al., 1935).

2.3.4.6 Recent Serologic Applications   Investigation of the reptilian humoral immune response has recently benefited from the development of novel serologic assays. The enzymelinked immunosorbent assay (ELISA), polyclonal antibodies, and hybridoma technology (Liddell and Cryer, 1991) for the production of monoclonal antibodies have been used since the early 1990s in several studies on reptiles. ELISA tests have been developed to evaluate the antibody production in the desert tortoise (Gopherus agassizii), gopher tortoise (Gopherus polyphemus) (Schumacher et al., 1993; Brown et

al., 1999), American alligator, broad-snouted caiman (Caiman latirostris), Siamese crocodile (Crocodylus siamensis) (Brown et al., 2001), green turtle (Coberley et al., 2001; Herbst and Klein, 1995; Work et al., 2000), Greek tortoise, Hermann’s tortoise (Origgi et al., 2001), and boa constrictor (Boa constrictor) (Kania et al., 2000; Lock et al., 2003). Seroconversion against the selected immunogen could be detected as early as 4 weeks post injection (PI) in the Greek tortoise (Origgi et al., 2001), gopher tortoise (Brown et al., 1999), desert tortoise (Schumacher et al., 1993), and boa constrictor (Lock et al., 2003), and 6 weeks in the American alligator, broad-snouted caiman, Siamese crocodile (Brown et al., 2001), and green turtle (Work et al., 2000). Other serological tests such as SNT and hemagglutination inhibition (HI) are used for the evaluation of reptilian immune responses. Serum neutralization has been used to evaluate the immune response of tortoises to pathogens (Marschang et al., 1997; Origgi et al., 2001) and American alligators to West Nile virus (Klenk et al., 2004), while HI has been used for evaluating snakes for exposure to paramyxovirus (Jacobson et al., 1981). For more information on serodiagnostics in reptile medicine, see Chapter 8 of this book.

2.3.5 Cell-Mediated Immune Responses The evaluation of the cell-mediated immune response in reptiles has been, for the most part, a functional evaluation. It has been based on the detection and measurement of three basic cell-mediated-based reactions: allograft and xenograft rejection, graft versus host rejection (GVHR), and mixed lymphocyte reaction (MLR). All three cell-mediated reactions have been observed in reptiles, indirectly suggesting the existence of allo-reactive T-cells and the reptilian homologues of the basic determinants and players of the cell-mediated immune response as seen in higher vertebrates.

2.3.5.1 Allografts and Xenografts   Temperature- and agedependent rejection of allografts were observed by Borysenko (1969 a, b) in the snapping turtle. Yntema (1970) investigated the influence of genetic distance and antigen sharing on xenotransplants. Tissues obtained from painted turtles and transplanted to embryonic snapping turtles were either accepted or partially rejected, while xenografts obtained from Florida soft-shelled turtles (Apalone [Trionyx] ferox) were always rejected. The shorter phylogenetic distance between snapping turtles and painted turtles in comparison to snapping turtles and soft-shelled turtles is one possible explanation for this finding (Cooper et al., 1985; Krenz et al., 2005). Transplantation experiments have also been performed in other reptiles such as the spectacled caiman (Caiman crocodilus) (Borysenko, 1970), green anole (Anolis carolinensis) (May, 1923), Mexican spiny-tailed iguana (Ctenosaura pectinata) (Cooper and Aponte, 1968), variable agama (Manickavel and Muthukkaruppan, 1969), desert night lizard (Xantusia vigilis) (Cooper, 1969), checkered whiptail lizard

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(Cnemidophorus tesselatus), six-lined racerunner (C. sexlineatus) (Maslin, 1967), desert grassland whiptail lizard (C. uniparens) (Cuellar, 1976), and garter snake (Thamnophis sirtalis) (Terebey 1970, 1972). Autografts healed while allografts were rejected at varying times unless obtained from related populations (parthenogenic lizards) (Maslin, 1967).

2.3.5.2 Graft versus Host Reaction (GVHR)   In vitro GVHR has been observed when allogenic spleen cells obtained from adult snapping turtles were mixed with spleen fragments of hatchlings (Sidky and Auerbach, 1968). Interestingly, the GVHR was observed only in spleen fragments obtained from individuals up to three months of age, and not in spleen explants obtained from turtles older than three months of age.

2.3.5.3 Mixed Lymphocyte Reaction (MLR)   Saad and El Ridi (1984) evaluated MLR in the ocellated skink along with GVHR and skin allograft rejection. Splenocytes responded with strong proliferation in one-way and two-way mixed leukocyte cultures, which was seen as evidence for the existence of MHCs. In vivo GVHR experiments confirmed the in vitro MLR findings. Intraperitoneal injections of splenocytes into newborn allogenic recipients were followed by splenomegaly and retarded growth, and resulted in mortality. Farag and El Ridi (1985) investigated MLR in the African beauty racer (Psammophis sibilans) using spleen cells obtained from outbred snakes. Proliferation observed in the mixed leukocyte cultures was interpreted as evidence of the presence of a subset(s) of lymphocytes capable of recognizing and responding to alloantigens. Stronger evidence came from analyzing the outcome of skin grafts, MLR, and cell-mediated lympholysis between random pairs of snakes. Significant positive correlations were found between MLR disparity, graft rejection, and cell-mediated lympholysis, indirectly suggesting the existence of a reptilian MHC (Farag and El Ridi, 1990).   A low level of cell proliferation was seen in a two-way MLR experiment in the tuatara (Burnham et al., 2005). The authors speculated that the minimal genetic differences at the histocompatibility loci of the two donors might have impacted the results.

2.3.6 Memory The most distinctive feature of adaptive immunity is probably the ability of the host immune system to react to a previously encountered antigen in a faster, stronger, and longer lasting manner than during the primary response. The existence of immunological memory is based on three distinctive elements of the secondary immune response: a shorter latency, a higher antibody titer, and a longer-lasting response. Immunological memory is based on the existence of a circulating pool of B- and T-lymphocytes produced during the primary response, which unlike the other members of the effector clones, do not die. These memory cells will prime the secondary response whenever they encounter the antigen

that evoked the primary response. It is not known how long a primed pool for an antigen will survive. It is possible to detect circulating antibodies several decades after exposure to the specific antigen, suggesting either self-sustained memory cell replication or renewed stimulation via chance encounters with the antigen. While there is some experimental evidence suggesting a lack of immunological memory in reptiles, it may have been biased by experimental conditions (Cooper et al., 1985). The immunosuppressive effects of stress may introduce bias into many reptile immunological studies. For example, while Cooper and Aponte (1968) described second set skin allografts surviving longer than first sets in the Mexican spiny-tailed iguanas, the low temperature at which the lizards were kept (25°C) may have influenced the outcome (Cooper et al., 1985). Similarly, Maung (1963) did not detect a secondary humoral immune response to Brucella abortus in immunized Greek tortoises kept at 15 to 30°C. No secondary response was detected in the tuatara (Marchalonis et al., 1969) immunized with Salmonella spp flagellin (Cooper et al., 1985). No secondary immune response was detected in desert iguanas and chuckwallas immunized with STH (Evans, 1963; Evans and Cowles, 1959; Evans et al., 1965), in contrast with the results of Wright and Shapiro (1973), who found higher antibody titers during the secondary response of the desert iguana to KLH. Hermann’s tortoises immunized with heat-inactivated pig serum and aluminum adjuvant showed a higher, faster titer increase in the secondary response than the primary response (Ambrosius and Lehmann, 1964, 1965). In Hermann’s tortoises and Russian tortoises immunized with DNP-protein, the maximum level of precipitating antibodies was produced later in the secondary response (Ambrosius and Frenzel, 1972). The most recent evidence of humoral immunological memory in reptiles is in a report by Origgi et al. (2004). In an experimental transmission study of a tortoise herpesvirus in Greek tortoises, a shorter lag phase and higher serum neutralization titers were seen in the infected tortoises following a second challenge with the antigen. However, it was not possible to detect a higher amount of total nonneutralizing antibodies directed against the virus. The duration of the secondary response peak was not determined due to study length. No secondary response was found in the rapid racerunner (Vorob’eva, 1965) following secondary exposure to tickborne encephalitis, while humoral memory was detected in the shingleback skink immunized with different antigens in complete Freund’s adjuvant (Wetherall and Turner, 1972). In snakes, evidence of the existence of a secondary response was found in the black racer, fox snake, and black rat snake immunized against BSA and KLH in complete Freund’s adjuvant, administered either subcutaneously or intraperitoneally (Salanitro and Minton, 1973). The secondary response was characterized by higher antibody titers, a shorter lag phase before the peak titer, and a duration of four months (Cooper et al., 1985). Finally, a secondary humoral

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immune response was also detected in American alligators immunized with KLH (Lerch et al., 1967).

2.3.7 Factors Affecting the Immune Response The immune response is a very complex process that can be influenced by many factors. Age, nutritional status, reproductive status, genetic background, antigen, the route used by the antigen to enter the host, temperature, time of day, season, psychological stress, and many other factors that can be grouped as intrinsic (host) and extrinsic (nonhost) factors, play a critical role in the immune response. Reptiles are ectotherms, and many of them brumate or aestivate during the cold and warm season, respectively. These factors need to be taken into account and evaluated when the overall immune response is considered.

2.3.7.1 Extrinsic Factors 2.3.7.1.1 Temperature   Reptiles are ectotherms and all their physiological activities are influenced by environmental temperature. Both the humoral and cell-mediated immune responses are heavily affected by temperature. Ambrosius (1976) and Cooper et al. (1985) reviewed this topic. Much work still needs to be done Hermann’s tortoises immunized with inactivated normal pig serum (NPS) show a delayed production of antibodies when kept at 20 to 21°C as compared to 28°C (Ambrosius and Lehman, 1964, 1965). Maung (1963) reported a suppressed humoral immune response to Brucella abortus in Greek tortoises kept below 10°C as compared to 15 to 30°C. In desert iguanas, upper (40°C) and lower (25°C) temperature limits were associated with a lower antibody production against STH compared to 35°C (Evans and Cowles, 1959; Evans, 1963; Evans et al., 1965). In the rapid racerunner and northern rock iguana, antibody response against tick-borne encephalitis virus was not detectable when the lizards were kept at 4°C, but was present at 37°C (Vorob’eva, 1965). No antibody response was observed in the Cunningham’s skink immunized with SRBC (Tait, 1969) when kept at 20°C, while a measurable response was present at higher temperatures, and maximal response was seen at 30°C. Similar temperature-dependent humoral responses were observed by Wetherall and Turner (1972) in the shingleback skink. The immune response of snakes has also been shown to be temperature dependent (Grasset et al., 1935). The outcomes of allografts and xenografts are temperature dependent in the snapping turtle (Borysenko, 1969a). Similarly, the outcome and the progress of GVHR reactions are markedly influenced by temperature in hatchling snapping turtles (Borysenko and Tulipan, 1973). 2.3.7.1.2 Seasonality and Hormones   There is a large body of literature showing a seasonal influence in the reptilian immune response. Zapata et al. (1992) reviewed this topic. Ambrosius (1966) reported large differences in the immune

response of Hermann’s tortoises immunized at different times of the year. The strongest immunological response was observed in tortoises immunized during the spring (April), while tortoises immunized in early October reacted weakly. Interestingly, tortoises immunized later in autumn (November) induced a humoral response slightly lower than spring immunization, but stronger than early autumn. Leceta and Zapata (1986) evaluated primary and secondary antibody responses and PFC responses of the Caspian turtle to SRBC in summer and autumn. Following primary immunization, both 2-ME-sensitive antibodies and splenic PFCs were seen in autumn but not summer. During the secondary response, 2-ME-resistant antibodies were detected both in summer and autumn, while the number of PFCs was significantly reduced during summert The reptilian cell-mediated immune response is also influenced by season. In the African beauty racer, the MLR was significant only in a few months of spring and autumn, with abrogation during the rest of the year (Farag and El Ridi, 1985). In the ocellated skink, MLR and skin graft rejection was abrogated in winter, and was significantly lower in spring through midsummer than midsummer to autumn (Saad and El Ridi, 1984). Munoz and De la Fuente (2001 a) investigated Caspian turtle splenocyte function (substrate adherence, chemotaxis, lymphoproliferative response to mitogens, antibody-dependent cellular cytotoxicity, and natural killer–like cell-mediated cytotoxicity) during different times of the year. Low substrate adherence, high chemotaxis, and high cytotoxic activity were observed in winter. During spring, high activity was recorded only for mitogen-induced lymphoproliferative responses. High chemotaxis and cytotoxicity were observed during the summer, while in autumn, only substrate adherence was enhanced. The same authors investigated seasonal influence on thymic cells of the Caspian turtle (Munoz and De la Fuente, 2001b). Chemotaxis, proliferative response to mitogens, and cytotoxicity were also affected differently by the seasonal cycle. In general, the lowest responses were seen in autumn for both sexes. Female thymic cells expressed the highest cytotoxicity and chemotaxis activity during the summer. Proliferative responses to mitogens peaked in spring for both sexes. The effect of season on the same functional parameters was also assessed for the peripheral lymphocytes of the Caspian turtle (Munoz et al., 2000). Chemotaxis, lymphoproliferative response to mitogens, and cytotoxicity were high in winter. Proliferative responses to mitogens were further increased in spring and then decreased during the summer, while cytotoxicity, adherence, and chemotaxis increased. Only substrate adherence showed high values in autumn. Sidky and Auerbach (1968) reported that spleen cells obtained from two hibernating snapping turtles were unable to induce GVHR when mixed with spleen fragments that were obtained from young turtles. Seasons also affect the structure and remodeling of the lymphoid tissues (see Sections 2.3.2.2.1, 2.3.2.3.1). In the Caspian turtle, both spleen and thymus show seasonal variation,

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although the cortex and medulla of the thymus (Figures 2.6– 2.9) and PALS and PELS (Figures 2.20–2.23) in the spleen white pulp are differently affected (Leceta and Zapata, 1985). Summer is the season when most thymic involution occurs, followed by some recovery in early autumn and a slower decrease in winter. In early spring, despite the increase in thymic volume, the cortex is not well developed. Thymic cortex and medulla are both well developed in late spring, when the thymus is further increased in size. The white pulp of the spleen also reaches its maximum development in late spring, followed in the summer by a marked reduction. The nonlymphoid components of the Caspian turtle thymus are also seasonally influenced (Leceta et al., 1989). In the rainbow skink and Egyptian mastigure, the thymus was highly involuted in winter and well developed in other seasons (Hussein et al., 1978b). Similarly, the splenic white pulp was severely depleted in winter but extensively developed in spring, summer, and autumn. In these seasons, development of the splenic white pulp of the rainbow skink almost obscured the red pulp. Maximal development of the white pulp was not as extensive in the Egyptian mastigure as in the rainbow skink. Mastigure splenic lymphoid tissue could be observed as periarteriolar aggregates, with the red pulp always well delineated. GALT was well represented in both lizards. In the Egyptian mastigure, intestinal lymphoid aggregates were reduced in winter, being found only in the caecum and colon, in contrast with distribution during the rest of the year throughout the gastrointestinal tract. Almost no seasonal variation was observed in the GALT distribution of the rainbow skink. In the ocellated skink, the lymphoid tissue of the thymus and spleen undergoes severe involution in the winter (Hussein et al., 1978a). Interestingly, the GALT appear to be influenced differently by season, with a reduction of the esophageal lymphoid nodules in the winter, followed by an increase in number and size in spring and summer, and a decrease again in autumn. In contrast, the GALT in the stomach and the small and large intestines are not influenced by season. Seasonal changes in lymphoid tissue development and humoral response were also studied in the sandfish (Hussein et al., 1979b). The antibody response to RRBC paralleled the developmental stage of the lymphoid tissues. A humoral response was not detected in winter, when the thymus of the sandfish was involuted and the splenic lymphoid tissue was markedly depleted. The response increased during the spring, and peaked in summer and autumn when the lymphoid organs showed full development. Seasonal changes in lymphoid tissues were also reported in the diadem snake (Hussein et al., 1979a) and Schokari sand racer (El Ridi et al., 1981). Seasonal changes in reptilian immune function appear to be correlated with systemic hormonal variations and structural changes in the lymphoid tissues. In the ocellated skink, four subpopulations (PNA+ Thy-1-, PNA+ Thy-1+, PNA- Thy1+, and PNA-Thy-1-) of T-lymphocytes were independently

affected by seasonal variation in endogenous steroid levels (El Masri et al., 1995). Lymphocytes were extracted from bone marrow, spleen, and thymus, and different populations of T-lymphocytes were identified. Each T-cell population was affected differently by endogenous steroid levels, and to some extent by organ distribution. The authors further supported their data with in vitro and in vivo experiments using exogenous hydrocortisone, testosterone propionate, and purified fractions of thymosin alpha. Saad and El Ridi (1988) studied the effect of seasonal variations of endogenous corticosteroid (CS) on immunological function in the ocellated skink. They found that white pulp development and strong immune response occurred from the spring through early autumn, and correlated with low CS levels. Severe lymphoid tissue involution and immune impairment were observed in autumn and winter, associated with high endogenous CS levels. Long-term administration of exogenous hydrocortisone acetate (HC) to ocellated skinks with fully developed lymphoid tissue in summer supported these observations. After HC treatment, the lizards showed a high and lasting elevation in blood CS levels associated with lymphoid involution and impairment of immune reactivity, mimicking the physiological status of lizards in autumn through winter. Furthermore, the treatment of the lizards with a CS synthesis antagonist (metyrapone) at the beginning of autumn interfered with seasonal-dependent immunosuppression. Similar results have been seen in the marine iguana (Amblyrhynchus cristatus) (Berger et al., 2005). Corticosterone was also shown to be responsible for thymocyte apoptosis in the Indian leaf-toed gecko. The cellular DNA fragmentation induced by corticosterone treatment appeared to be dose dependent and required 48 hours (Hareramadas et al., 2004). The thymus gland of the Indian leaf-toed gecko was shown to undergo profound seasonally dependent structural changes, most markedly in reticuloepithelial cells of the thymic cortex, which could be correlated to levels of testosterone (Hareramadas and Rai, 2001). Light and electron microscopy showed marked involution of the thymus in winter, when androgen is at a maximal level. Thymus regenerates in the spring to become fully developed in the summer, when testicular steroidogenic activity is minimal. The authors were able to positively correlate these findings with castration and testosterone replacement experiments. Similar results have been seen in the Caspian turtle (Varas et al., 1992). Hareramadas and Rai (2005) later described the direct inhibitory effect of dihydrotestosterone on thymic cell proliferation and its indirect effect of enhancing the caspase-dependent apoptotic process in thymocytes, mediated by thymic cortical reticuloepithelial cells, in the Indian leaf-toed gecko. Belliure et al. (2004) investigated the effects of testosterone on the T-cell mediated response to mitogens in two lacertid lizards, the Algerian sandrunner (Psammodromus algirus) and the European fringe-fingered lizard (Acanthodactylus erythrurus). Phytohemagglutinin (PHA) stimulation appeared to be

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strongly suppressed in testosterone-treated males of both species. A significant negative relationship between individual variability of T-cell-mediated responsiveness and plasma testosterone concentration was also observed. Similar results have been found in the ocellated skink (Saad et al., 1990). In female Indian leaf-toed geckos, estrogen directly inhibited thymic cells and indirectly controlled thymic cell apoptosis mediated by thymic cortical reticuloepithelial cells (Hareramadas and Rai, 2006). In the western fence lizard (Sceloporus occidentalis), 17-alpha-ethinylestradiol was found to decrease peripheral lymphocyte counts, but only to affect splenic lymphocyte numbers at high doses (Burnham et al., 2003). Enhanced splenic lymphocyte proliferation was seen in MLR in treated fence lizards. Finally, Saad and Shoukrey (1988) investigated the influence of sex hormones on the immune activity of the African beauty racer. The authors evaluated the immune response of male and female subjects on the basis of functional tests. Females were better responders than males in primary antiRRBC antibody production and response to mitogens. 2.3.7.1.3 Differential Response to Type and Amount of Antigen   In reptiles, similar to what is known in other vertebrates, the type of antigen appears to influence the dynamics of the immune response (Ambrosius, 1976). Wetherall and Turner (1972) showed a poor antibody response in shingleback skinks immunized with RRBC, while the responses to Salmonella spp. and to BSA were strong. Frenzel and Ambrosius (1971) and Ambrosius et al. (1972) found that tortoises immunized with protein antigens, such as Bacillus CalmetteGuerin (BCG) or serum proteins, developed high levels of antibodies for sustained periods after reaching peak titers. In contrast, following immunization with DNP-conjugatedproteins, the antibody titer decreased relatively fast after the peak response. Hemmerling (1971) observed that precipitating and nonprecipitating low-molecular-weight (probably 7.3S and 6.8S, respectively) antibodies were produced by the armored glass lizard, according to the protein antigen used. Pig serum induced both precipitating and nonprecipitating antibodies, whereas bovine IgG induced only nonprecipitating antibodies (Ambrosius, 1976). The affinity of reptilian antibodies to antigens can be affected by the antigen type (Fiebig, 1972). In the Russian tortoise, the different carriers of DNP were shown to influence both the isotype of the antibody produced and its affinity. Immunizing tortoises with DNP-poly-L-lysine (DNP-PLL), DNP-human serum albumin (DNP-HAS), DNP-BCG, or DNPBrucella abortus, resulted in a strong high-molecular-weight response only after DNP-Brucella abortus. The other three conjugates induced mostly the production of low-molecular-weight antibodies. Of these, DNP-PLL gave the weakest response, while DNP-HAS and DNP-BCG elicited a good humoral response and showed a typical anamnestic response. Nevertheless, the affinity of the antibodies remained almost the same throughout the course of the immune response.

Interestingly, DNP-Brucella immunization, despite the weak low-molecular-weight antibody response, showed a remarkable increase in affinity. The amount of antigen used for immunization has been reported to influence the strength of the immune response. In lizards, (Wetherall and Turner, 1972) while the maximum antibody titer produced following immunization with BSA was independent of the antigen amount (over a range of 1 to 200 mg), the response showed a tendency to occur earlier at higher doses. In the mole snake, agglutinins to typhus endotoxin and meningococci were produced in a dose-dependent fashion (Grasset et al., 1935). In the common night adder and ringneck spitting cobra, low doses of antigens did not induce antibody synthesis (Grasset et al., 1935). Grey (1963) reported no differences in the painted turtle immunized with or without adjuvant. In contrast, Ambrosius and Lehmann (1964, 1965) and Ambrosius (1967) observed that Hermann’s tortoises immunized with pig serum proteins showed a higher titer when an aluminum hydroxide adjuvant was used. An even stronger response was detected when incomplete Freund’s adjuvant was used. Interestingly, the presence of the adjuvant was reported to influence the antibody isotype production. 2-ME-sensitive antibodies were found in tortoises immunized without adjuvant and 2-MEresistant antibodies predominated in tortoises immunized with adjuvant during the secondary response. In lizards, Wetherall and Turner (1972) observed a strong effect of complete Freund’s adjuvant on shingleback skinks immunized with BSA. A lower antibody titer was detected when complete Freund’s adjuvant was not used. Jacobson et al. (1991) showed that eastern diamondback rattlesnakes could develop antibodies against inactivated paramyxovirus either with or without adjuvant. However, a uniform response was not seen. More recently, Marschang et al. (2001) observed no significant differences between tortoises immunized with inactivated tortoise herpesvirus antigen with or without adjuvant. 2.3.7.1.4 Route of Administration   Origgi et al. (2004) reported comparable immune responses in tortoises injected with live tortoise herpesvirus either IM or intranasally (IN). Detection of serum neutralizing and nonserum neutralizing antibodies revealed a similar kinetic antibody response independent of the injection schedule adopted (see also Section 2.3.11). Wetherall and Turner (1972) compared IP and IM immunization of shingleback skinks administered different antigens. Salmonella antigen gave a strong immune response to both injection routes, while BSA elicited a higher titer when injected IP.

2.3.7.2 Intrinsic Factors   Unfortunately, our knowledge concerning the immune composition of reptiles is limited. We have no information concerning reptilian MHC variability and other features that define the immunological identity of this class of vertebrates. This is one of many interesting and

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challenging aspects of reptilian immunology still waiting to be investigated.

2.3.8 The Immune Response during Bacterial Diseases See Chapter 8 for information.

2.3.9 The Immune Response during Viral Diseases See Chapter 8 for information.

2.3.10 The Immune Response during Parasitic Diseases See Chapter 8 for information.

2.3.11 Vaccination Very limited information on the existence of a true secondary immune response and of affinity maturation in all reptiles leave us without many arguments to sustain or reject the possibility and feasibility of an effective vaccine for a reptilian infectious disease. There are only two modern examples of vaccination experiments: Jacobson and colleagues (1991) and by Marschang and colleagues (2001), respectively. Jacobson et al. (1991) injected three groups of six paramyxovirus seronegative eastern diamondback rattlesnakes (Crotalus atrox) with an inactivated paramyxovirus suspension. The viral suspension was either adjuvanted (oil-emulsion or aluminum hydroxide) or not. Each of the groups received one of the three available formulations of the vaccine. Following vaccination, two of the snakes that received no adjuvant and three of those that received the oil-emulsion vaccine seroconverted. No seroconversion was observed in the snakes injected with the aluminum adjuvanted vaccine. At 296 days post vaccination, all snakes were seronegative with the exception of one of the snakes vaccinated with the oil-emulsionated viral suspension. Marschang and colleagues (2001) vaccinated a group of Mediterranean tortoises (Testudo spp) using inactivated tortoise herpesvirus with or without adjuvant (aluminum hydroxide). The tortoises were vaccinated three times at 45day intervals. No significant rise in antibody titers was noted in vaccinated animals, and antibody titers measured dropped below the cutoff level sporadically in all positive animals. Finally, no correlation was seen between titer increases and the type of vaccine administered. A third vaccination experiment was conducted by Origgi et al. (2001, 2004), who injected two groups of Greek tortoises with a live tortoise herpesvirus. Tortoises in one group were inoculated IN, while the second group was inoculated IM. An initial dose of 15,000 TCID50 was used, followed by a second dose of 150,000 TCID50, administered by same route 11 months later. All but one tortoise seroconverted. Antibod-

ies required a longer time to be detected by SN than ELISA. Antibodies were detectable for at least 10 months after a single administration. The presence of detectable SN did not prevent the occurrence of clinical signs following challenge with the second administration of virus. Features of a secondary immune response were observed after the second administration of the live virus.

2.3.12 Future In this chapter we have summarized findings of numerous immunological studies from the 1900s into the beginning of the 21st century. While these studies have provided valuable information on seasonal changes, morphology of the immune system, and humoral response to antigens, certain components of the immune system, such as cell-mediated immunity, remain virtually untouched. The advent of molecular medicine may provide the necessary data to more completely understand the workings of the reptile immune system. While the components of the immune system needed to efficiently protect reptiles from pathogens seem to be in place, we do not know how similar their function is to those in mammals, birds, amphibians, and fish. In-depth study of the effect of seasonality and temperature on the dynamics of the reptilian immune system may reveal mechanisms that have been lost by endothermic vertebrates. We believe that further investigation of reptilian immunology is necessary and will provide valuable insights into comparative immunology.

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Figure 2.1  Neotropical rattlesnake, Crotalus durissus. Viperidae. Photomicrograph of a vertebra. Numerous lymphoid and myeloid cells are seen within marrow spaces. H&E stain.

Figure 2.2  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Photomicrograph of a vertebra. Numerous lymphoid and myeloid cells are seen within marrow spaces. H&E stain.

Figure 2.3  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of periototic bone of the skull. Granulocytes and a multinucleated osteoclast are seen within a marrow space. H&E stain.

Figure 2.4  Green turtle, Chelonia mydas. Cheloniidae. The thymus (arrows) is paired and is located cranial to the heart.

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Figure 2.5  Green turtle, Chelonia mydas. Cheloniidae. Acetic acid digestion of connective tissue around the heart. The thymus (T) is seen cranial to the heart. Courtesy of Bobby R. Collins.

Figure 2.6  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of the thymus. A connective tissue capsule (CT) surrounds the thymus. Lobules are separated by connective tissue septa, with each lobule consisting of an outer cortex (CO) and inner medulla (ME). H&E stain.

Figure 2.7  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Higher magnification photomicrograph of the thymus in Figure 2.6. The outer darker cortex (CO) and inner paler medulla (ME) of a lobule are seen. H&E stain.

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Figure 2.8  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of a thymic lobule. A distinct outer cortex and medulla are not seen. H&E stain.

Figure 2.9  Desert tortoise, Gopherus agassizii. Testudinidae. Higher magnification photomicrograph of a thymic lobule in Figure 2.8. A distinct outer cortex and medulla are not seen. H&E stain.

Figure 2.10  Burmese python, Python molurus bivittatus. Pythonidae. Photomicrograph of an aggregate of epithelial cells (EP) surrounded by thymocytes (TH). H&E stain.

Figure 2.11  Burmese python, Python molurus bivittatus. Pythonidae. Photomicrograph of a thymic lobule. Numerous myoid cells, having an eosinophilic cytoplasm, are seen along with thymocytes. H&E stain.

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Figure 2.12  Burmese python, Python molurus bivittatus. Pythonidae. Higher magnification photomicrograph of a thymic lobule in Figure 2.11. Numerous myoid cells, having an eosinophilic cytoplasm, are seen along with thymocytes. H&E stain.

Figure 2.13  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Photomicrograph of the thymus. Numerous cysts (arrows) are seen throughout the section. H&E stain. Courtesy of John Roberts.

Figure 2.14  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of an involuted thymus. The lymphoid tissue is severely depleted (compare with Figures 2.6 and 2.7). A distinct cortex and medulla are not seen. H&E stain.

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Figure 2.15  Prairie rattlesnake, Crotalus viridis, Viperidae. Photomicrograph of the splenopancreas. The pancreas consists of the exocrine pancreas (EX) and scattered aggregates of epithelial cells that comprise the islets of Langerhans (IS). In this snake the spleen (SP) is contiguous with the pancreas. H&E stain.

Figure 2.16  Prairie rattlesnake, Crotalus viridis. Viperidae. Higher magnification photomicrograph of the splenopancreas in Figure 2.15. The exocrine pancreas (EX) and islets of Langerhans (IS) are contiguous with the spleen. H&E stain.

Figure 2.17  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of the spleen. A very limited amount of red pulp (RP) is seen within the more abundant white pulp. The white pulp consists of vessels (VE) surrounded by lymphoid tissue. H&E stain.

Figure 2.18  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of the spleen showing red pulp intermixed with white pulp. White pulp consists of vessels surrounded by lymphoid tissue. H&E stain.

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Figure 2.19  Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of the spleen showing red pulp intermixed with white pulp. White pulp consists of vessels surrounded by lymphoid tissue. With this stain, periarteriolar lymphoid sheaths (PALS) and the periellipsoidal lymphoid sheaths (PELS) cannot be distinguished. H&E stain.

Figure 2.20  Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of a periarteriolar lymphoid sheath (PALS). Muscle fibers in the arteriolar wall stain red. Masson’s trichrome stain. Courtesy of Allan Pessier.

Figure 2.21  Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of a periellipsoidal lymphoid sheath (PELS). Only collagen (blue) is seen in the vessel wall indicating this is a capillary. Masson’s trichrome stain. Courtesy of Allan Pessier.

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Figure 2.22  Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of a periarteriolar lymphoid sheath (PALS). Around the central arteriole, the lymphoid tissue is embedded in a mesh of silver impregnated (black) reticular fibers. Gordon and Sweet’s stain. Courtesy of Allan Pessier.

Figure 2.23  Red-eared slider, Trachemys scripta elegans. Emydidae. Photomicrograph of a periellipsoidal lymphoid sheath (PELS). Directly around the central capillary is a silver impregnated ellipsoid (arrow) of reticular fibers. Few reticular fibers are seen within the lymphoid tissue around the ellipsoid. Gordon and Sweet’s stain. Courtesy of Allan Pessier.

Figure 2.24  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of the spleen showing several sheathed vessels surrounded by abundant red pulp. H&E stain.

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Figure 2.25  American alligator, Alligator mississippiensis. Alligatoridae. Higher magnification photomicrograph of the spleen in Figure 2.24. Several vessels are surrounded by lymphoid tissue. H&E stain.

Figure 2.26  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a cross-section of the spleen. The surface is covered by a fibrous capsule. Sheathed vessels are not seen. The red pulp is not abundant. H&E stain.

Figure 2.27  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of the spleen. The white pulp is more abundant than the red pulp. H&E stain.

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Figure 2.28  Burmese python, Python molurus bivittatus. Pythonidae. Photomicrograph of the spleen. The white pulp (WP) has a lobular distribution, with fibrous septa separating adjacent lobules. The red pulp (RP), also known as the perilymphoid fibrous zone (PLFZ), is very limited. H&E stain.

Figure 2.29  Burmese python, Python molurus bivittatus. Pythonidae. Photomicrograph of the spleen. Red pulp and white pulp are distributed throughout the splenic parenchyma. Sheathed vessels are not present. H&E stain.

Figure 2.30  Prairie rattlesnake, Crotalus viridis. Viperidae. Photomicrograph of the spleen. The white pulp (WP) is more abundant than the red pulp (RP). H&E stain.

3 Circulating Inflammatory Cells Nicole I. Strik, A. Rick Alleman, and Kendal E. Harr

Contents

3.1 General Concepts of Hematology

3.1 General Concepts of Hematology.........................167 3.1.1 Collection and Handling of Blood   Samples.......................................................167 3.1.2 Hematology Procedures.............................170 3.1.3 General Considerations..............................176 3.2 Erythrocytes and Erythrocyte Responses   in Disease................................................................176 3.2.1 Normal Erythrocyte Morphology and   Function ....................................................176 3.2.2 Abnormalities in Erythrocytes.................. 177 3.2.3 Anemia and Polycythemia........................ 178 3.3 Leukocytes and Leukocyte Responses   in Disease............................................................... 179 3.3.1 Heterophils................................................ 179 3.3.2 Eosinophils................................................ 180 3.3.3 Basophils................................................... 180 3.3.4 Lymphocytes ............................................ 181 3.3.5 Plasma Cells ............................................. 181 3.3.6 Monocytes.................................................. 181 3.3.7 Azurophils................................................. 182 3.3.8 Tumors of Hematopoietic Tissue.............. 182 3.4 Thrombocytes and Thrombocyte Responses   in Disease............................................................... 182 3.5 Infectious Agents in the Peripheral Blood........... 183 3.5.1 Hemoparasites........................................... 183 3.5.2 Viral Inclusions in Blood Cells................. 185 3.5.3 Bacteria...................................................... 185 Acknowledgments............................................................ 186

Most infectious agents of reptiles elicit an inflammatory response in affected tissues. Many inflammatory and infectious conditions also result in significant and specific changes in the peripheral blood. Evaluation of the hemogram and the blood film provides rapid, valuable information for the assessment of the health status of reptiles as well as for the further identification of certain disease processes. In this chapter we review the proper collection and handling of reptilian blood specimens, laboratory procedures used to analyze these specimens, the normal morphologic features of each blood cell type, as well as changes in cell types that may be associated with various disease conditions. While species variation in cell morphology always presents a challenge to those who work with reptile blood, the information presented here may serve as a guideline to the evaluation of blood from chelonians (turtles and tortoises), crocodilians (alligators, caiman, crocodiles, gharial), and squamates (lizards and snakes).

References......................................................................... 186

3.1.1 Collection and Handling of Blood Samples Blood sampling represents an invasive procedure, and as such there is associated pain and the risk of bacterial infection at the sampling site, and subsequently systemically. As with any procedure, it is best to learn from a person with extensive experience sampling from the site to be used. An aseptic technique is necessary, with cleansing of the site prior to sampling. Repeated alternating applications (typically three) of organic iodine soap or 2% chlorhexidine and 10% isopropyl alcohol or 70% ethanol should be used. While the degree of pain associated with sampling from each site has not been scientifically assessed, veterinarians having expertise in reptile health assessment can make appropriate recommendations. Ultimately the investigator needs to select a method that causes the least amount of pain and suffering,

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especially if an animal is awake and manually restrained, when the sample is collected. The total amount of blood that can be obtained safely from a reptile depends upon the reptile’s size and health status. The total blood volume of reptiles varies among species, but as a generalization, is approximately 5 to 8% of total body weight (Lillywhite and Smits, 1984; Smits and Kozubowski, 1985). Thus, a 100-g snake has an estimated blood volume of 5 to 8 ml. Because clinically healthy reptiles can acutely lose 10% of their blood volume without detrimental consequences, approximately 0.7 ml of blood can be safely withdrawn from a snake weighing 100 g. Much larger volume percentages of blood can repeatedly be removed over an extended period of time (Lillywhite et al., 1983). However, this practice is limited to experimental animals under controlled laboratory conditions.

3.1.1.1 Chelonia  A variety of sites have been used by different clinicians and researchers to obtain blood from chelonians. These include the heart, jugular vein, brachial vein, ventral coccygeal vein, orbital sinus, cervical sinus, trimmed toe nails, and subcarapacial vein (Avery and Vitt, 1984; Dessauer, 1970; Gandal, 1958; Hernandez-Divers et al., 2002; Jacobson, 1987; Maxwell, 1979; McDonald, 1976; Nagy and Medica, 1986; Owens and Ruiz, 1980; Rosskopf, 1982; Stephens and Creekmore, 1983; Taylor and Jacobson, 1982). Each sampling method has certain advantages and disadvantages. While some methods, such as cardiac sampling, are more invasive than other methods, they may have value in certain situations, such as experimental studies where serial samples are being collected, and in neonates, where this may be the only site from which blood can be obtained. In young chelonians, before the shell has calcified, a needle can be inserted through the plastron into the heart (Figure 3.1). It is imperative to cleanse the plastron several times (as described above) before the needle is passed through the shell. Older tortoises with calcified shells require either drilling a hole through the plastron over the heart using a sterile drill bit, or using a spinal needle for percutaneous sampling through soft tissue in the axillary region at the base of the forelimbs. General anesthesia is required to create a hole in the shell. This technique is only recommended for laboratory-housed chelonians where they can be monitored on a daily basis. In all situations, a sterile technique is necessary because contamination of the pericardial sac with bacteria and other potential pathogens can lead to pericarditis and possibly death. After blood is collected, the hole is sealed with an appropriate sealant such as bone wax (Johnson and Johnson, Co., Sommerville, NJ) and a sterile methacrylate resin (Cyanoveneer, Ellman International Mfg., Inc., Hewlett, NY). In turtles and tortoises, orbital sinus sampling has been used for collecting small volumes of blood in capillary tubes (Nagy and Medica, 1986). However, in order to prevent damage to periocular tissues and possible trauma to the cornea, a significant amount of care must be taken when using this

technique. Another consideration is the extent of pain associated with this procedure. The periocular tissues are extremely sensitive to tactile stimulation, and before using this sampling site, the animal needs to be anesthetized (Jacobson, 1993). This is mandatory. A further problem with this technique is dilution of the sample with extravascular fluids and secretions, which may alter the composition of plasma and affect the accuracy of blood cell counts. Blood samples are also commonly obtained from the scapular vein, brachial vein, and brachial artery of chelonians (Avery and Vitt, 1984; Rosskopf, 1982). However, vessels associated with limbs can rarely be visualized through the skin, and sampling is usually blind. In addition, because lymphatics are well developed in chelonian forelimbs (Ottaviani and Tazzi, 1977), obtaining blood samples from these vessels may result in hemodilution with lymph. At times during venipuncture, pure lymph may be obtained, which appears as a clear fluid (Gottdenker and Jacobson, 1995). If a lymph-contaminated blood sample is used for a complete blood count and plasma biochemical determinations, the results of these assays will be erroneous. Blood sampling from trimmed toenails has been utilized by some investigators. Limited amounts of blood can be obtained from this site, and the quantities needed for specific testing may render this collection site unacceptable. Lymphatics are also found in the toenail bed, and lymph and interstitial fluid will dilute the sample and affect plasma biochemical and serologic values. Other considerations for using this site include pain and risk of subsequent infection. Along with vessels, the tissues within the toenail are also invested with nerve endings. Similar to sampling from the heart through the shell, bacteria may gain access to the vascular system if the exposed toenail vessel is not sealed (cauterized or covered with a surgical resin) and if the fibrin clot is subsequently dislodged when the animal ambulates or digs. Therefore, this technique should not be used in the field where it is difficult or impossible to prevent vessel exposure following release. Another blind site that has been utilized to collect blood from chelonians is a subcarapacial venipuncture site (Hernandez-Divers et al., 2002). This vessel is located in the angle where the cervical vertebrae join the shell and is formed by the junction of the common intercostals and the caudal cervical branch of the external jugular veins. Similarly, where the caudal (tail) vertebrae join the carapace there is a dorsal vein that can be collected. The size of the needle used in obtaining a sample from these sites will depend upon the size of the chelonian. In large chelonians, a spinal needle will be necessary. This venipuncture site is particularly useful in sampling from small chelonians. In tortoises and freshwater turtles, blood is readily obtained from a postoccipital venous plexus that is located dorsal to the cervical vertebrae, behind the occipital protuberance of the skull (Gottdenker and Jacobson, 1995). A needle is passed at right angles to the cervical vertebrae, and using gentle pressure on the barrel of the syringe, a sample

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can be collected. In sea turtles, the dorsal cervical sinus that originates from the postoccipital venous plexus is a commonly used venipuncture site (Owen and Ruiz, 1980). The needle is inserted perpendicular to the dorsal neck, lateral from either side of the midline and immediately cranial to the carapace. Depending upon the turtle’s size, the paired sinuses are found 0.5 to 3 cm deep. The head of the turtle is best maintained lower than the body (Figure 3.2). The use of a restraint board improves the stabilization of the turtle and may help to decrease operator handling. As with other sites in which vessels cannot be visualized, hemodilution with lymph can occur. The only peripheral blood vessels that can consistently be visualized in many small and moderately sized tortoises as well as some freshwater turtles are the jugular vein and carotid artery (Jacobson et al., 1991). If the head can be restrained and the jugular seen, this site is ideal for obtaining a pure blood sample from chelonians (Figure 3.3). The major disadvantage of sampling from these vessels is that manual extension and restraint of the head of the tortoise beyond the margins of the plastron is required, which at times may be difficult or impossible. Tactile stimulation of the rear limbs usually causes the tortoise to extend its head from the shell, allowing the sampler to restrain the head. Once grasped, the head is pulled out with one hand, and while sitting, the sampler positions the tortoise between the knees, with the tortoise’s head pointing toward the sampler’s body. Alternatively, restraint boards may be used to assist in stabilizing the animal. The jugular vein and carotid artery are well developed on both the right and left sides of the neck. Once the head is extended, the jugular vein can often be seen as a bulge through the cervical skin. The carotid artery is deeper and more difficult to visualize, and is located ventral and parallel to the jugular vein. Once either vessel is identified, the skin over the puncture site should be cleansed with 70% ethanol and a 23- or 25-gauge butterfly catheter can be used for obtaining the sample. With the cap removed from the end of the tube, blood will flow down the tube once the needle is inserted into the vessel. Once the blood is obtained, and as the catheter is removed, using gauze, pressure should be applied to the sampling site for at least 5 minutes to prevent hematoma formation. Myopathy may also result if excessive force is used to maintain the head in an extended position.

3.1.1.2 Crocodylia  In members of this order, blood samples can be obtained from the supravertebral vessel located caudal to the occiput and immediately dorsal to the spinal cord (Olson et al., 1975). This site is commonly associated with hemodilution, e.g., with cerebrospinal fluid, and care must be taken not to injure the spinal cord. Using manual restraint, a 3.75-cm, 22- or 23-gauge needle is inserted through the skin in the midline directly behind the occiput and is slowly advanced in a perpendicular direction (Figure 3.4). As the needle is advanced, gentle, negative pressure is placed on the plunger. If the needle is passed too deep, the spinal cord will

be pithed. Other commonly used sites of blood collection include the heart via cardiocentesis and the ventral coccygeal vein (Jacobson, 1984). The heart is located in the ventral midline, approximately 11 scale rows behind the forelimbs. In collecting blood from the coccygeal vein, the crocodilian is placed in dorsal recumbency and the needle is inserted through the skin toward the caudal vertebrae. This venipuncture site is recommended for smaller-sized crocodilians (Figure 3.5).

3.1.1.3 Sauria  In these reptiles, blood samples can be obtained from several sites. In large lizards, it is most convenient to obtain blood from the ventral tail vein (Figure 3.6) (Esra et al., 1975). In smaller lizards, which represent most of the pet lizards, venipuncture is generally difficult. Some investigators have collected blood in a microcapillary tube (Samour et al., 1984). However, as with chelonians, pain and risk of infection are common complications. For animals to be sampled and released in the field, this method is not recommended. Microcapillary tubes can also be used to obtain blood samples from the orbital sinus (LaPointe and Jacobson, 1974); however pain and potential injury to the cornea precludes the use of this site, unless the animal is anesthetized. A large ventral abdominal vein is present in certain lizards such as the green iguana (Iguana iguana), and blind samples can be collected from this site in a manually restrained iguana. 3.1.1.4 Ophidia  In snakes, blood can be collected from a variety of sites, including the palatine veins (Figure 3.7), ventral tail vein, and via cardiocentesis (Olson et al., 1975; Samour et al., 1984). Cardiocentesis is commonly used in snakes. As long as the heart is not excessively traumatized with multiple attempts at sampling, the procedure is safe and effective (Jacobson, 1993). Some clinicians believe that this method should be limited to those snakes over 300 grams (Jackson, 1981). Some muscular snakes, such as large boas and pythons, are difficult to manually restrain and bleed from the heart. Venomous snakes are also problematic, and the heart may not be the ideal sampling site unless anesthetized. In these instances, the tail vein may be the ideal venipuncture site (Figure 3.8). For cardiocentesis, the heart is located either by direct visualization of the cardiac pulse through ventral scales or by palpation (Figure 3.9). The heart is relatively moveable within the coelomic cavity, and is easy to manually relocate several scale rows both cranially and caudally. Placing a thumb at its apex and forefinger at its base stabilizes the heart once it is located. A 23- or 25-gauge needle attached to a 3- to 6-ml syringe is advanced under a ventral scale, starting at the apex and aiming toward the base. With gentle suction, a sample can be obtained. Sometimes a clear fluid is withdrawn, representing the pericardial fluid. In such cases, the needle should be withdrawn, a new syringe and needle secured, and the procedure repeated. In obtaining samples from this location, the clinician may notice that blood can be withdrawn with each beat of the heart. After three failed

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attempts at obtaining a blood sample from the heart, further sampling attempts should be discontinued (Jacobson, 1993).

3.1.1.5 Serial Blood Sampling  Certain conditions, such as drug pharmacokinetic studies, necessitate serial blood collection over a relatively short period of time. In some trials, one or more samples may need to be collected daily for a one-week period or more. In a serial collection study in ball pythons (Python regius), 139 blood samples were obtained by cardiocentesis over a 120-day period. The animals were subsequently monitored and no clinical or pathological problems were seen by 73 days after the last blood sample was taken (Isaza et al., 2004). For certain sized reptiles, vascular catheterization can be utilized for serial blood sampling. This procedure has been previously reported for tortoises (Prezant et al., 1994), lizards ([iguanas] Maxwell and Jacobson, 2004), and snakes (Jacobson and Hodge, 1980; Young et al., 1997). Vascular catheterization may not be possible in small (under 100 grams) reptiles. While serial blood samples can be collected from peripheral vessels and the heart of certain reptiles, ethical considerations may preclude serial sampling from the same site more than one or two times daily. Limits for serial cardiac sampling are not yet clearly defined.

3.1.2 Hematology Procedures Evaluation of complete blood counts (CBCs) include red blood cell (RBC) count, hemoglobin concentration (Hb), packed cell volume (PCV), white blood cell (WBC) count, differential leukocyte counts, and morphologic evaluation of blood cells. Normal blood cells of reptiles include erythrocytes, granulocytes (heterophils, eosinophils, basophils), mononuclear leukocytes (lymphocytes, monocytes, azurophils) and thrombocytes. Individual blood cell types will be reviewed in the following sections of this chapter. Reference intervals for various cell types in different reptile species are summarized in Tables 3.1 to 3.4. Further information can be obtained from the Reference Ranges for Physiological Values in Captive Wildlife (CD-ROM), produced by the International Species Information System (ISIS), www.isis.org. Blood should be collected into microtainer tubes containing lithium heparin (Becton-Dickinson, Rutherford, NJ), or in some species, dipotassium ethylenediaminetetraacetic acid (K-EDTA) as an anticoagulant. The use of EDTA should be used with caution because it can cause hemolysis in some species, especially chelonians (Jacobson, 1987; Muro et al., 1998). The use of EDTA has been reported suitable as an anticoagulant for snake blood (Dotson et al., 1995, Lamirande et al., 1999; Salakij et al., 2002). In some reptiles, such as the green iguana, white blood cell counts and differential leukocyte counts were found to be more similar to those of nonanticoagulated blood films in samples collected in EDTA versus those collected in heparin (Hanley et al., 2004). When heparin is used as an anticoagulant, a bluish tinge in the background of blood films may be observed (Hawkey and

Dennett, 1989), and leukocytes may not stain as intensely as blood that is collected in EDTA. Furthermore, leukocytes and thrombocytes generally clump more in heparinized blood than in EDTA-anticoagulated blood (Hawkey and Dennett, 1989). Cell clumping may adversely affect the cell counts and the accuracy of the blood film evaluation. In order to avoid these artifacts, blood with no anticoagulant may be used to perform a blood film immediately after collection. Other anticoagulants that can be used for CBCs include sodium heparin and ammonium heparin (Campbell, 1996a). Heparinization of the needle and syringe during slow blood collection may prevent clot formation. After removal of the blood volume needed for hematologic analysis, heparin-anticoagulated blood samples are suitable for biochemical analysis, which requires early separation of plasma from red cells in order to ensure appropriate measurements of plasma constituents such as potassium (Abou-Madi and Jacobson, 2003). With very small patients, only a single drop of blood may be obtained. This can still provide valuable diagnostic information if used to make a properly prepared blood film. The coverslip method of preparing a blood film is preferred over the slide-to-slide technique because it minimizes traumatic injury to the large and fragile reptilian blood cells, and provides a more even distribution of blood cells for estimating cell counts. For the coverslip method, two coverslips, a fine brush and a blood-filled microhematocrit tube are needed. The brush is used to remove glass or dust particles from the coverslip surfaces. The coverslips should only be touched at their edges. One coverslip is placed between the thumb and index finger of one hand, and a small drop of blood is then put in the center of the coverslip by using the microhematocrit tube. The second coverslip is placed over the first in a crosswise fashion without using any pressure (Figure 3.10). As the blood spreads between the coverslips, they are then quickly separated by holding the overlapping corners of the coverslips and sliding them apart using no pressure and staying in the horizontal plain. Rapid drying (warm air blow dryer) is preferred to avoid drying artifacts in red blood cells (Figure 3.11). The preferred hematologic stains for reptile blood films are Romanowsky-type stains, such as Wright-Giemsa, WrightLeishman’s, or May-Grünwald. Quick stains (e.g., Diff-Quik®, American Scientific Products, McGraw Park, IL) may also be used, but heterophil granules tend to coalesce with Diff-Quik (Muro et al., 1998). Therefore, heterophils may appear less eosinophilic and less distinct when stained with Diff-Quik (LeBlanc, 2001). Wright-Giemsa stains may be superior to rapid staining methods, but it may not be practical to maintain in a clinical laboratory setting. After collection and direct blood film preparation, the sample should be gently inverted several times to prevent clotting and allow a uniform mixing of blood cell components. Insufficient sample volume relative to the anticoagulant in the tube may erroneously decrease the hematocrit value as well as altering other aspects of the hemogram. Clots or fibrin

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Table 3.1  Hematologic values for tortoises. Mean of various Parameter speciesa RBC/µl 523,000 (154,166–980,000) Hg (g/dl) N/A PCV (%) 27 (23–35) MCV (fl) N/A MCHC (g/dl) N/A MCH (pg) N/A Thrombocytes/µl N/A Total protein (g/dl) N/A WBC/µl N/A Heterophils/µl N/A Heterophils (%) N/A Lymphocytes/µl N/A Lymphocytes (%) N/A Monocytes/µl N/A Monocytes (%) N/A Eosinophils/µl N/A Eosinophils (%) N/A Basophils/µl N/A Basophils (%) N/A Azurophils/µl N/A

Desert tortoises; winterb 590,000

Desert tortoises; summerb 635,000

Desert tortoisesc N/A

6.3 23.6 402 26 106 N/A 2.85 4,780 3,074 N/A 410 N/A 46 N/A 80 N/A 532 N/A 99

7 26 407 28 110 N/A 3.65 4885 2563 N/A 583 N/A 0 N/A 61 N/A 1,011 N/A 0

N/A N/A N/A N/A N/A N/A N/A N/A N/A 33 N/A 23 N/A 11 N/A 1 N/A 30 2

Frye FL. 1991. Hematology as applied to clinical reptile medicine, in Biomedical and Surgical Aspects of Captive Reptile Husbandry, 2nd ed, Vol. 1, Frye FL (Ed.), Krieger Publishing Co., Melbourne, FL, 209–277. b Gopherus agassizzi; Christopher MM, Berry KH, Wallis IR, Nagy KA, Henen BT, and Peterson CC. 1999. J Wildl Dis 35:212–238. c Gopherus agassizzi; Alleman AR, Jacobson ER, and Raskin RE. 1992. Am J Vet Res 53:1645–1651. a

Parameter RBC/µl Hg (g/dl) PCV (%) MCV (fl) MCHC (g/dl) MCH (pg) Thrombocytes/µl Total protein (g/dl) WBC/µl Heterophils/µl Heterophils (%) Lymphocytes/µl Lymphocytes (%) Monocytes/µl Monocytes (%) Eosinophils/µl Eosinophils (%) Basophils/µl Basophils (%) Azurophils/µl d e

Aldabra tortoisesd N/A N/A 21.1 N/A N/A N/A N/A 5.8 2,083 N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A

Burmese mountain tortoisesd N/A N/A 24.8 N/A N/A N/A N/A 6.2 4,822 N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A

Hermann’s tortoisese 490,000 6.14 24.44 514.8 26.32 122.64 N/A N/A 7,240 N/A 48.6 N/A 22.5 N/A 3 N/A 23.3 N/A 2 N/A

Geochelone gigantea and Manouria emys; Abou-Madi N and Jacobson ER. 2003. Vet Clin Pathol 32:61–66. Testudo Hermanni; Muro J, Cuenca R, Pastor J, Vinas L, and Lavin S. 1998. J Zoo Wildl Med 29:40–44; mean values from heparinized blood samples.

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Table 3.2   Hematologic values for turtles. Mean of various Parameter speciesa Hawaiian green turtlesb Sea turtlesc Freshwater turtlesc 523,000 RBC/µl (154,166–980,000) N/A 385,000 558,000 Hg (g/dl) N/A N/A N/A N/A PCV (%) 27 (23–35) 29 (17–35) 33.3 22.9 MCV (fl) N/A N/A N/A N/A MCHC (g/dl) N/A N/A N/A N/A MCH (pg) N/A N/A N/A N/A Thrombocytes/µl N/A N/A 36/100 WBC N/A Total protein (g/dl) N/A N/A 4.15 3 WBC/µl N/A 13,800 (5,900–23,600) 4,250 6,660 Heterophils/µl N/A 1,400 (300–3,200) N/A N/A Heterophils (%) N/A N/A N/A N/A Lymphocytes/µl N/A 10,000 (3,600–18,600) N/A N/A Lymphocytes (%) N/A N/A N/A N/A Monocytes/µl N/A 800 (100–1,900) N/A N/A Monocytes (%) N/A N/A N/A N/A Eosinophils/µl N/A 1,700 (700–3,200) N/A N/A Eosinophils (%) N/A N/A N/A N/A Basophils/µl N/A 0 (0–100) N/A N/A Basophils (%) N/A N/A N/A N/A Azurophils/µl N/A N/A N/A N/A a Frye FL. 1991. Hematology as applied to clinical reptile medicine, in Biomedical and Surgical Aspects of Captive Reptile Husbandry, 2nd ed, Vol. 1, Frye FL (Ed.), Krieger Publishing Co., Melbourne, FL, 209–277. b Chelonia mydas; Work TM, Raskin RE, Balazs GH, and Whittaker SD 1998. Am J Vet Res 59:1252–1257. c Moon PF and Hernandez-Divers SM. 2001. Reptiles: aquatic turtles (chelonians), in Zoological Restraint and Anesthesia, Heard D (Ed.), International Veterinary Information Service, Ithaca, NY, www.ivis.org. Table 3.3   Hematologic values for crocodilians. Mugger crocodiles, Mugger crocodiles, American alligatora,c juvenileb adultb Salt-water crocodiled RBC/µl 384,000;1,049,000 690,000(580,000–810,000) 800,000(640,000–990,000) 600,000–1,200,000 (618,000–1,480,000) Hg (g/dl) N/A 8.3 (5.2–12.7) 8.64 (6,600–10,100) 4.7–12.2 PCV (%) 27 (20–35) 24.9 (16–38) 25.4 (19–30) 17–41 MCV (fl) N/A 362.43 (239–520) 327.69 (241–448) 240–311 MCHC (g/dl) N/A 33.36 (30.4–34.4) 33.85 (32.9–34.7) N/A MCH (pg) N/A 120.68 (77.6–168) 110.78 (67.5–136) 72–92 Thrombocytes/µl 23,000 20,900 (12,000–29,000) 23,800 (13,000–32,000) 4,000–71,000 Total protein (g/dl) N/A N/A 3.19 (2.9–3.9) 4.1–7.0 WBC/µl 6,400 8,710 (5,060–15,400) 6,970 (4,400–15,600) 6,400–25,700 Heterophils/µl N/A 5,600 (3,120–9,550) 4,450 (2,240–6,700) 800–7,400 Heterophils (%) 54.7 N/A N/A N/A Lymphocytes/µl N/A 2,480 (1,160–4,930) 1,830 (790–3,100) 4,500–21,600 Lymphocytes (%) 23.9 N/A N/A N/A Monocytes/µl N/A 90 ( 0–320) 120 (0–300) 0–1,200 Monocytes (%) 0.7 N/A N/A N/A Eosinophils/µl N/A 530 (0–1,290) 560 (140–1,020) 0– 700 Eosinophils (%) 10.4 N/A N/A N/A Basophils/µl N/A 10 (0–100) 0 0– 400 Basophils (%) 12.7 N/A N/A N/A a Alligator mississippiensis; Mateo MR, Roberts ED, and Enright FM. 1984. Am J Vet Res 45:1046–1053. b Crocodylus palustris; Stacy BA and Whitaker N. 2000. J Zoo Wildl Med 31:339–347. c Frye FL. 1991. Hematology as applied to clinical reptile medicine, in Biomedical and Surgical Aspects of Captive Reptile Husbandry, 2nd ed, Vol. 1, Frye FL (Ed.), Krieger Publishing Co., Melbourne, FL, 209–277. d Crocodylus porosus; Dessauer H. 1970. Blood chemistry of reptiles: Physiological and evolutionary aspects, in Biology of the Reptilia, Vol. 3, Gans C, Parsons TS (Eds.), Morphology C, Academic Press, New York, 1–72. Parameter

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Table 3.4   Hematologic values for lizards. Green iguana, Green iguana, Green iguana, Inland bearded Mean of various Parameter speciesa malesb femalesb juvenilesb dragonc RBC/µl 867,500 1,000,000–1,700,000 1,200,000–1,800,000 1,300,000–1,600,000 N/A (466,000–2,050,000) Hg (g/dl) N/A 6.7–10.2 9.1–12.2 9.2–10.1 N/A PCV (%) 29 (10.8–35) 29.2–38.5 33– 44 30–47 17–50 MCV (fl) N/A 228–303 235–331 N/A N/A MCHC (g/dl) N/A 22.7–28 24.9–31.0 N/A N/A MCH (pg) N/A N/A N/A N/A N/A Thrombocytes/µl N/A N/A N/A N/A N/A Fibrinogen (mg/dl) N/A 100–200 100–300 100–300 N/A Total protein (g/dl) N/A 4.4–6.5 4.9–7.6 4.2–6.1 4.5–9.5 WBC/µl N/A 11,100–24,600 8,200–25,200 8,000–22,000 6,736–19,946 Heterophils/µl N/A 1,000–5,400 600–6,400 1,000–3,800 1,619–7,339 Heterophils (%) N/A N/A N/A N/A 17 - 43 Lymphocytes/µl N/A 5,000–16,500 5,200–14,400 6,200–17,200 4,012–12,033 Lymphocytes (%) N/A N/A N/A N/A 47–69 Monocytes/µl N/A 200–2,700 400–2,300 300 - 600 0–499 Monocytes (%) N/A N/A N/A N/A 0–4 Eosinophils/µl N/A 0–300 0–400 0–400 N/A Eosinophils (%) N/A N/A N/A N/A N/A Basophils/µl N/A 100–1,000 200–1,200 100–700 205–3,191 Basophils (%) N/A N/A N/A N/A 2–18 Azurophils/µl N/A N/A N/A N/A 0–1,085 Azurophils (%) N/A N/A N/A N/A 0–9 a Frye FL. 1991. Hematology as applied to clinical reptile medicine, in Biomedical and Surgical Aspects of Captive Reptile Husbandry, 2nd ed, Vol. 1, Frye FL (Ed.), Krieger Publishing Co., Melbourne, FL, 209–277. b Iguana iguana; Harr KE, Alleman AR, Dennis PM, Maxwell LK, Lock BA, Bennett RA, and Jacobson ER. 2001. J Am Vet Med Assoc 218:915–921; ranges of clinically normal green iguanas. c Pogona vitticeps; Eliman MM. 1997. Bull Assoc Rept Amphib Vet 7:1–3. Common Parameter chameleond RBC/µl 400,000–1,700,000 Hg (g/dl) N/A PCV (%) 24 MCV (fl) N/A MCHC (g/dl) N/A MCH (pg) N/A Thrombocytes/µl N/A Total protein (g/dl) 4.7 WBC/µl 31,200 Heterophils/µl N/A Heterophils (%) 66 Lymphocytes/µl N/A Lymphocytes (%) 25 Monocytes/µl N/A Monocytes (%) 9 Eosinophils/µl N/A Eosinophils (%) 0 Basophils/µl N/A Basophils (%) 0.5 Azurophils/µl N/A Azurophils (%) N/A d Chamaeleo chamaeleon; Cuadrado M, Diaz-Paniagua C, Quevedo MA, Aguilar JM, and Prescott IM. 2002. J Wildl Dis 38:395–401.

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strands in the sample render it unusable. The sample should be submitted to the laboratory as rapidly as possible. Blood films should be made immediately after collection and sent to the laboratory for evaluation along with the anticoagulated blood. Prolonged storage of blood in anticoagulant can alter cell numbers and morphology, e.g., degranulation of granulocytes, vacuolation of blood cells or cell lysis.

3.1.2.1 Erythrocytes  Erythrocyte evaluation includes determination of the packed cell volume (PCV), total red blood cell count (TRBC), hemoglobin (Hb) concentration, and evaluation of red blood cell morphology on the blood film. The microhematocrit method is used to obtain the PCV. A microhematocrit tube is filled to about 90% of its capacity and sealed with clay at one end. The tube is then spun at 12,000g for 5 minutes in a microhematocrit centrifuge. The spun microhematocrit tube provides PCV, total plasma protein, icterus index, and after heat precipitation, a rough estimation of the fibrinogen present in the sample. PCV can vary from 20 to 45% depending on the species (Sypek and Borysenko 1998). Plasma color should be clear to light yellow. Due to dietary pigments, orange to yellow plasma can be seen in herbivorous reptiles, and greenish-yellow plasma in snakes; green plasma has been observed in some lizards due to high concentrations of biliverdin (Campbell, 2004). The TRBC can be obtained by manual methods (hemocytometer) or automated cell counter. The two manual methods include the erythrocyte Unopette system and the Natt-Herrick’s solution. With the erythrocyte Unopette system (Becton-Dickinson, Rutherford, NJ), the provided pipette is used to mix the blood sample in the Unopette vial, making a 1:200 dilution. Both sides of the hemocytometer chamber are carefully filled and the chamber placed in a humidified petri dish. After allowing the cells to settle for at least 5 to 10 minutes, the total number of red cells over the four corners and large central square of the chamber are counted on low power (e.g., 20X magnification). The elliptical, large red blood cells stain weekly violet with a darker nucleus (Figure 3.12). The TRBC (cells/µl) is calculated by multiplying the number of red cells counted x 10,000. The Natt-Herrick’s solution, a methyl-violet-based stain, is used as a combination diluent and stain (Campbell, 1995; Natt and Herrick, 1952). First, a 1:100 dilution is prepared by mixing 990 µl Natt-Herrick’s stain and 10 µl of well-mixed blood sample. Then 100 µl of this dilution is mixed with 900 µl of saline (isotonic sodium chloride solution, 0.85%), which results in a 1:1,000 dilution of the initial blood sample. Both sides of a hemocytometer chamber are carefully filled and the cells counted as described for the Unopette system. The TRBC (cells/μl) is then calculated by multiplying the counted red blood cells × 1,000. Hemoglobin values are measured as for mammals by utilizing the cyanmethemoglobin method or a hemoglobinometer (Coulter Electronics, Hialeah, FL). For accurate spectrophoto-

metrical measurement of Hb with the cyanmethemoglobin method in reptiles, the removal of free nuclei from lysed RBC by centrifugation (5 minutes at 500 x g) is required before measuring the optical density (Campbell, 1996a) because the presence of free nuclei may falsely elevate the measured Hb value. Normal hemoglobin in different reptile species can vary from 5.5 to 12 g/dL (Sypek and Borysenko, 1988). The red cell indices (mean corpuscular volume [MCV], mean corpuscular hemoglobin [MCH], and mean corpuscular hemoglobin concentration [MCHC]) can then be calculated using the spun PCV, RBC, and Hb concentration using the following formulas: MCV (fl) = (PCV x 10)/RBC MCH (pg) = (Hb x 10)/RBC MCHC (mg/dl) = (Hb x 100)/PCV Lizards are known to have the smallest red cells of all reptile species, with increasing red cell size in ascending order in snakes, turtles, alligators, and tortoises (Sypek and Borysenko 1988, Campbell 1996a, Gottdenker and Jacobson 1995). There is an inverse relationship between red cell size and total RBC count, with those reptiles having small erythrocytes tending to have higher erythrocyte counts (Sypek and Borysenko 1988, Campbell 1996a). MCV and MCHC are used to characterize anemia in mammals and may be of benefit in determining the cause of anemia in reptiles and the evaluation of erythrocyte responses in disease (see Section 3.2.2).

3.1.2.2 Leukocytes   Leukocyte evaluation includes total white blood cell count (WBC), determination of the differential leukocyte count, and evaluation of leukocyte morphology on the blood film. The nucleated reptile erythrocytes and thrombocytes interfere with automated cell counters. Although the accuracy of these instruments is superior to manual methods when using mammalian blood, there have been no published validation studies in reptiles. Preliminary findings (Harr, unpublished observations) indicate that it may be possible to obtain a reliable total WBC count in reptiles using the Cell Dyn 3500 and the associated Veterinary Computer Package (Abbott Diagnostics, Abbott Park, IL). However, it appears that even using sophisticated analyzers, the variation among reptile species will require the operator to modify parameters and perform validation studies for each species tested. Although the precision of manual methods is less than desirable, they are relatively accurate and practical. However, the accuracy of the count obtained should be confirmed by performing a leukocyte estimation on the blood film. The two commonly used manual methods for total leukocyte counts in reptiles are the direct count using NattHerrick’s solution and the semidirect count using phloxine B solution, both using a hemocytometer (Campbell, 1996a; Schermer, 1967). The Natt-Herrick’s solution is prepared as described for the TRBC, and the round and dark violet–staining leukocytes

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in the 10 large squares of the hemocytometer are counted. The total number of WBC from the count is then multiplied by 100 and yields the total WBC (cells/µl). The advantage of the Natt-Herrick’s solution is that it can be used to perform total counts of erythrocytes, leukocytes, and thrombocytes. The disadvantage of this method is that differentiation between small lymphocytes and thrombocytes may often be difficult and may result in erroneous leukocyte counts. Lymphocytes appear smaller, round, and weakly violet, compared to the irregularly shaped, weakly violet thrombocytes (Figure 3.13). Another disadvantage of the Natt-Herrick’s solution is its potential for laboratory errors when using manually prepared dilutions. In reptiles with normally high numbers of circulating heterophils (e.g., iguanas and some tortoises), the semidirect phloxine B method can also be used (Campbell, 1995). The reddish-stained heterophils and eosinophils are easily identified in the hemocytometer chamber. A simplification of this method is the Eosinophil Unopette 5877® system (BectonDickinson, Rutherford, NJ), which was developed for determining the total number of eosinophils in mammalian blood (Costello, 1970). To determine the WBC count, a 1:32 dilution of blood is made using an Eosinophil Unopette that contains phloxine dye diluent. After loading the hemocytometer chamber, it is placed in a humified petri dish and allowed to settle for at least 5 to 10 minutes. The red-staining cells are then counted in all nine large squares on both sides of the hemocytometer. The resulting number is then mathematically adjusted to achieve the total leukocyte, heterophil, and eosinophil concentrations per cubic millimeter of blood: Heterophils + Eosinophils/µl = (cells counted x 10 x 32) / 18

Example Total count of eosinophils and heterophils in the hemocytometer: 350 Heterophils + Eosinophils/μl = (350 x 10 x 32) / 18 = 6,200/µl After performing the leukocyte differential, the total leukocyte (white blood cell) concentration (TWBC/µl) is calculated using the following formula: TWBC/ µl = [(Heterophils + Eosinophils/µl) x 100] / % Heterophils and Eosinophils (Campbell, 1996a).

Example The differential yielded heterophils 45 %; eosinophils 3 %; basophils 1 %; lymphocytes 39%; monocytes 12 % TWBC/µl = (6,200 x 100)/ 48 =   12,900/μl

If the sample volume is small, only enough to prepare a blood film, a leukocyte estimate can be performed, similar to that used in birds (Campbell and Coles, 1986). Given a properly prepared blood film with even distribution of blood cells, cell counts can be estimated using the following formula: # cells/µl = (average # of cells per field) x (objective power)2 The objective used to estimate leukocyte numbers should be one where approximately 5 to 10 leukocytes are seen per field. For example, the total number of leukocytes are counted under 50X magnification in 10 fields and yielded a total of 50 cells. The average of 5 is then multiplied by 2500, resulting in a total of 12,500 cells/µl. Because hemocytometer counts have an inherent degree of error, the authors recommend routinely estimating leukocyte numbers from the blood film for the purpose of quality control.

3.1.2.3 Thrombocytes  As in mammals, the reptilian thrombocytes tend to clump rapidly in vitro. Therefore, the presence of clumps have to be reported (Figure 3.14) and the number can be subjectively described as decreased, adequate, or increased. If the blood film contains representative areas with well-distributed cells, an estimate may be performed by counting the thrombocytes per 100 leukocytes. Normal numbers vary between 25 and 350 thrombocytes per 100 leukocytes (Sypek and Borysenko, 1988). Because this value depends on the total leukocyte count, erroneous reductions or elevations are likely. Manual thrombocyte counts may also be performed with the same hemocytometer chamber loaded with the Natt-Herrick’s solution for total erythrocyte and leukocyte counts. The numbers of thrombocytes in the central large square are counted on both sides of the hemocytometer. The total number of thrombocytes on both sides is then multiplied by 500 in order to obtain the number of thrombocytes per microliter. Because thrombocyte clumps are readily noted in the hemocytometer, the accuracy of this method is dubious.

3.1.2.4 Total Protein and Fibrinogen  Although total protein values are often determined utilizing a refractometer, in the authors’ opinion, this method is inaccurate for measuring plasma protein in reptiles. Proper methods for determining total protein are based on the Biuret method via spectophotometry (Kingsley, 1972). Normal plasma protein values can range from 3 to 8 g/dl (Campbell 1996a).  The fibrinogen content in a blood sample can be roughly estimated via the heat precipitation test. This test is done by measuring the difference in total plasma solids before and after heat precipitation of fibrinogen. Two filled microhematocrit tubes are centrifuged at 12,000g. The total plasma solids are determined by refractometer from one tube. The second tube is heated in a 56°C water bath for 3 minutes, followed by centrifugation for one minute at 12,000g. The total solids are

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then determined in the second tube. The difference in total solid measures is an estimate of fibrinogen. Values generally range between 100 and 200 mg/dL. The heat precipitation method is not sensitive enough to differentiate low-normal fibrinogen values from those that are below normal. Ocular micrometry and a coagulation-based assay are more accurate in determining fibrinogen values (Meyer and Harvey, 2004). High fibrinogen values are indicative of active inflammation and physiologic change in mammals, particularly large animals. At this time, little is known regarding the significance in reptile patients.

3.1.2.5 Reference Intervals  While blood values for a variety of reptiles have been published, relatively few studies have followed guidelines for establishing reference intervals (Walton, 2001). The best-studied reptile to date is the desert tortoise (Gopherus agassizii), where reference intervals have been determined for certain wild populations (Christopher et al., 1999). Reference intervals for hematocrit, plasma biochemicals, and plasma protein electrophoretogram fractions for green turtles (Chelonia mydas) and loggerhead sea turtles (Caretta caretta) in Florida are currently under investigation. These values are available online (Jacobson et al., 2007).

3.1.3 General Considerations Many factors influence the hemogram of reptilian patients. When interpreting a hemogram of a reptilian patient, external factors such as season, environmental conditions, venipuncture site, and laboratory methods have to be considered as well as individual variances such as age and gender. Such effects are discussed for each leukocyte type. Overall, the information of the hematologic findings may be helpful in the diagnosis of disease, evaluating prognosis, and monitoring patient response to therapy.

3.2 Erythrocytes and Erythrocyte Responses in Disease 3.2.1 Normal Erythrocyte Morphology and Function Unlike mammalian erythrocytes, reptilian, avian, amphibian, and piscine red blood cells (RBC) have nuclei. Nucleated RBCs are elliptical and larger than nonnucleated RBCs, with amphibian RBCs being the largest. In general, mature reptilian erythrocytes have nuclei that are irregularly round to oval, with dense coarse chromatin and homogenous eosinophilic (red) cytoplasm (Figure 3.15). Immature, polychromatophilic erythrocytes have a similar shape but are slightly rounder with light blue cytoplasm upon Romanowsky staining (Figures 3.16–3.17). Nuclei from polychromatophils contain clumped chromatin with obvious, pale euchromatin indicative of the active hemoglobin production occurring in

these cells. A more immature form of erythrocyte, the rubricyte, has a round, slightly irregular nucleus with clumped chromatin and a round, dark blue cytoplasm (Figure 3.18). Rubricytes are similar in their appearance to lymphocytes and must be distinguished in animals with a regenerative response. This stage of erythrocyte maturation is capable of replication. Therefore, mitotic figures of circulating erythrocytes may be seen in blood films (Figures 3.19–3.20), especially in patients with active regeneration. Mitotic activity in reptilian peripheral blood is not by itself indicative of a neoplastic process. Binucleated erythrocytes are considered indicative of abnormal erythropoiesis and have been associated with severe chronic inflammatory disease and neoplasia in birds (Campbell, 1995). As erythrocytes age, their nucleus rounds up to perfect spheres and appears pyknotic with dark, dense chromatin. These cells may be seen in low numbers on blood films of healthy reptiles (Figures 3.16, 3.21). Anucleated erythrocytes, erythroplastids, may occasionally be seen and have no pathologic significance in birds and reptiles (Campbell, 1995; Hawkey and Dennett, 1989) (Figure 3.22). Erythrocyte size, number, and hemoglobin content have been compared among 441 species of mammals, birds, and reptiles (Hawkey et al., 1991). Reptiles have lower total RBC counts, hemoglobin concentrations, and PCVs than do either mammals or birds. These findings indicate a greater oxygencarrying capacity of the blood in birds and mammals compared to ectothermic animals such as reptiles. Erythrocyte function is similar to that of mammals though differences exist within and among the different orders of the Reptilia. Nucleated RBCs contain hemoglobin tetramers that carry oxygen and carbon dioxide to and from the tissues, respectively. Hemoglobin structure appears to be relatively well conserved across the different species of reptiles (Coates, 1975). However, small changes in molecular structure result in significant variation in oxygen affinity (Rucknagel and Braunitzer, 1988). In general, lizards tend to have a significantly higher oxygen affinity while chelonia have decreased oxygen affinity (Johansen et. al., 1980; Torsoni, and Ogo, 1995). Two functionally different hemoglobin tetramers have been separated from the blood of adult red-eared sliders (Trachemys scripta elegans), which exhibit marked differences in oxygen affinity and in concentration of ATP (adenosine triphosphate) associated with hemoglobin (Frische et al., 2001). It is postulated that these two hemoglobin molecules exist in the same RBC, though this functional difference may be due to erythrocyte age as it is in mammals. Additionally, turtle erythrocytes have been proposed as a model for the evolutionary transition state between RBCs relying on aerobic metabolism and the anaerobically metabolizing mammalian RBCs, a transition that is homologous to that occurring in maturing mammalian RBCs (Mauro and Isaacks, 1997). Red blood cells are continuously produced by bone marrow elements and removed from the blood by phagocytes present in splenic tissue. Production of erythrocytes occurs predominantly in the extravascular space in bone

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marrow though erythroid precursors can also replicate in the peripheral blood. In neonates, the yolk sac is the primary site of erythropoiesis, which may continue through the first year of life (Vasse and Beaupain, 1981). The life span of mammalian red blood cells is proportional to size, and ranges from 2 to 5 months in domestic animals (Meyer and Harvey, 2004). In comparison to mammalian red cells, nucleated red cells have a life span up to 600 to 800 days which is likely associated with the low metabolic rate of reptiles (Frye, 1991). Nucleated red blood cells undergo programmed cell death and offer an excellent model for the study of apoptosis. Investigation in this area is just beginning (Miyamoto et al., 2005). Gender differences have been reported in several species of reptiles, but this appears to vary with species and potentially with the study. Male New Guinea snapping turtles (Elseya novaeguinae) and grass snakes (Natrix natrix) have been reported to have significantly higher hemoglobin, PCV, and bilirubin (a hemoglobin breakdown product) than females (Anderson et al., 1997; Wojtaszek, 1991). Nongravid and gravid female green iguanas were reported to have significantly higher hemoglobin concentrations, PCV, and MCHC than males (Harr et al., 2001). Elevated concentrations of methemoglobin, which would be considered pathologic in mammals, have been reported in healthy reptiles. However, more recent studies, in which techniques to prevent the oxidation of methemoglobin are instituted in study design, reveal that methemoglobin values are actually similar to mammalian values. Older literature reports apparently healthy snakes with methemoglobin percentage ranging from 6 to 28%, lizards with methemoglobin ranges of 2 to 5%, and turtles with methemoglobin percentages of a massive 5 to 60% in healthy animals (Prado, 1946; Pough, 1969; Sullivan and Riggs, 1964). This is in direct contrast to studies designed to compare methemoglobin across species where no statistically significant difference could be found among mammals, reptiles, and birds using the same methodology (Rodkey et al., 1979). Additionally, more recent studies in the red belly black snake (Pseudechis phorphyriacus) revealed a lower methemoglobin of 3%, which is more consistent with mammalian values; less than 2% methemoglobin was present in the saltwater crocodile (Crocodylus porosus), Johnston’s crocodile (C. johnstoni), common snakeneck turtle (Chelodina longicollis), and forest skink (Sphenomorphus quoyi); and less than 1% methemoglobin present in the yellow-footed tortoise (Geochelone denticulata) and red-footed tortoise (G. carbonaria) (Gruga and Grigg, 1980; Torsoni et al., 2002). This indicates that high methemoglobin concentrations and methodologies prior to 1975 are circumspect and values should be rechecked with contemporary experimentation prior to further quotation in the literature.

3.2.2 Abnormalities in Erythrocytes Characterization of disease processes associated with abnormal erythrocyte morphology has been limited in reptiles. Polychromasia (multiple colors) is the presence of bluish or immature RBCs on stained blood smears. It is observed with some frequency in moderately to severely anemic reptiles (Figures 3.16–3.20). This represents a regenerative response and an attempt by the animal to return to homeostasis. Low numbers (< 1% RBC number) of polychromatophils are considered normal for most reptiles. Reptilian erythroid regenerative response appears to be slower than that observed in mammals. When anemia was induced in the red-eared slider using phenylhydrazine hydrochloride, 30 days elapsed prior to any regenerative response, and the authors report up to 8 weeks prior to maximal regenerative response. Rabbits showed a regenerative response in 5 days in the same study (Sheeler and Barber, 1965). Decreased MCHC and decreased MCV have been documented to be associated with polychromasia in reptiles (Sheeler, 1965). Both mammalian and reptilian polychromatophils contain decreased quantities of hemoglobin, which is actively produced in these immature cells, resulting in decreased MCHC. In mammals, MCV generally increases during a regenerative response due to the slightly larger size of mammalian reticulocytes. However, reptilian polychromatophils are generally smaller in size than mature reptilian RBCs, resulting in decreased MCV. Poikilocytosis may appear as fusiform or teardrop-shaped erythrocytes that can be seen in a low number in healthy reptiles, and may be explained as preparation artifact (Frye, 1991). It has been speculated that an increased number of these cells may be seen in animals with septicemia, or others with severe chronic infection (Frye, 1991). Intracytoplasmic inclusions have been reported in normal sea turtles (Work et al., 1998) and tortoises (Alleman et al., 1992). The authors of this chapter have seen similar erythrocytic inclusions in other reptile species. These individual, small, basophilic punctate or clear ring-shaped inclusions may be present in a variable number of erythrocytes in a blood film with no known clinical significance (Figures 3.23–3.25). Ultrastructural investigations revealed that these inclusions are consistent with degenerate organelles (Alleman et al., 1992). Square to rectangular to occasionally hexagonal, pale, crystalline-like cytoplasmic inclusions consistent with hemoglobin crystals were initially investigated in rhinoceros iguanas (Cyclura cornuta and C. figgensi) using transmission electron microscopy (Simpson et al., 1980; Simpson et al., 1982). In the author’s experience, similar crystals are observed with some frequency in various species of lizards, snakes, and tortoises. These have been documented in the literature in the green iguana, and crystals may also be observed in the nucleus in this species (Harr et al., 2001) (Figures 3.26–3.27). The cause and significance of these inclusions are unknown.

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However, ultrastructural analysis of the hemoglobin protein reveals microtubules of polymerized hemoglobin, which are virtually identical to those observed in RBCs of deer and humans with sickle cell anemia (Simpson, 1982). These proteins are found in low percentages in normal animals (< 1%) and diseased rhinoceros iguanas (5 to 10%). It is unlikely that these low numbers of afflicted RBCs significantly impact oxygen transport. It should be noted that these crystals can be found in healthy, reproductively active animals, and so the clinical significance is unknown. Genetic influence should be investigated, especially in a captive breeding situation. Viral inclusions have been observed in erythrocyte cytoplasm (see Section 3.5.2.2). A Brazilian lancehead viper (Bothrops moojeni) that was evaluated for renal carcinoma was found to have two types of inclusions present concomitantly in the same RBC. One type of inclusion contained viral particles (small aggregates of granular eosinophilic material) and the other inclusion was crystalline, translucent, hexagonal, and contained an unknown protein (Johnsrude et al., 1997). The snake was markedly anemic and exhibited a marked regenerative response. Ultrastructural analysis revealed an iridovirus consistent with snake erythrocyte virus and crystalline structures that were different than typical hemoglobin crystals. The viral inclusions are similar to acidophilic inclusions in East African Chameleons (Chamaeleo dilepis) documented on electron microscopy to contain viral particles consistent with the family Iridoviridae (Telford and Jacobson, 1993). Erythroparasites are also present in RBC cytoplasm and may be confused with true inclusions. Some parasites may be associated with anemia and other pathologic disease states (see Section 3.5).

3.2.3 Anemia and Polycythemia Artifactual changes should be ruled out prior to interpretation of anemia in reptiles. Lymphatic vessels are present in close proximity to blood vessels in the tail, forelimb, and other regions of the body (Lopez-Olvera, 2003; Ottaviani and Tazzi, 1977). Dilution of the blood sample with lymph results in decreased PCV, hemoglobin concentration, etc. (Heard et al., 2004). If the sample has a decreased PCV with no evidence of regeneration and increased numbers of small lymphocytes (Figure 3.28), submission of a new sample should be requested to verify results. Upon confirmation that the sample is representative of the patient, anemia should be characterized and the following potential diagnoses should be ruled out. Anemia may be caused by increased RBC destruction, decreased RBC production, or blood loss. The anemia should be characterized based on polychromasia as either regenerative or nonregenerative. It should be noted that reptiles have increased time to regenerative response so one must consider the chronicity of the anemia. In general, if the anemia has persisted for more than one month with no significant response, it may be classified as nonregenerative. Many systemic inflamma-

tory diseases as well as liver and renal failure, may result in nonregenerative anemia. Viral infection has been documented to cause moderate to severe anemia. Compared to a population of normal green turtles, a decrease in mean PCV of approximately 35% was observed in green sea turtles afflicted with severe fibropapillomatosis, a disease seen in sea turtles around the world and thought to be caused by a herpesvirus (Herbst et al., 2004). This disease is also associated with decreased total protein and white blood cells including lymphocytes, basophils, and eosinophils (Work and Balazs, 1999). Thrombocyte changes associated with herpesvirus infection have not been documented. However, this is likely a result of study design and requires further investigation. The fact that multiple cell lines are decreased in herpetic infection indicates that there is likely a change in bone marrow microenvironment causing a component of decreased cell production in this disease. Further investigation is warranted. Evidence exists that toxin exposure may cause anemia. Concentrations of Sigma chlordanes in fat biopsies from loggerhead sea turtles were negatively correlated with red blood cell counts, hemoglobin, and hematocrit (Keller et al., 2004). Starvation results in decreased RBC numbers, PCV, and hemoglobin concentration in the checkered keelback (Xenochrophis piscator), an elapid snake (Pati and Thapliyal, 1984). Additionally, when exposed to human hormones, red cell mass in these starved snakes increased by hormone combinations including urinary erythropoietin and L-thyroxine. It appears that these hormones work synergistically to stimulate erythrocyte production. There may also be a circadian effect on erythrocyte production where exposure to sunlight is a positive synergistic factor (Pati and Gupta, 1991). During hibernation and dry periods, reptiles may not drink for weeks to months at a time. This results in a dehydrated state and potentially marked changes in blood values that should not be overly interpreted. Both hemoglobin and PCV have been documented to increase due to these seasonal and annual changes in desert tortoises (Dickinson et al., 2002). However, statistically significant (p < 0.01) decreased PCV and hemoglobin concentration have been documented in a laboratory setting in the Cunningham skink (Egernia cunninghami) when exposed to low temperatures. In this study, a decrease of 12°C over a 48-hour period resulted in a decrease of > 20% in PCV and hemoglobin concentration when compared to control lizards housed at optimal temperatures (Maclean et al., 1975). Additionally, chronic cold and submergence has been explored in the hibernating painted turtle (Chrysemys picta). When housed at cold temperatures equivalent to winter, hemoglobin in this species exhibited a significant right shift and increased oxygen affinity even with a concurrent decrease in pH. These observed changes in blood oxygen transport may facilitate oxygen loading during winter submergence, thereby allowing hibernation under water (Maginniss, et al., 1983; Maginniss and Hitzig, 1987; Rucknagel et al., 1988). In conclusion, knowledge of the indi-

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vidual species and exact environmental stressors is required for interpretation of blood changes in hibernating reptiles.

Abnormal heterophil morphology is commonly seen in response to inflammatory disease, particularly infectious diseases. Toxic changes are frequently observed morphologic alterations in heterophils and commonly accompany severe inflammation or infection. These changes occur in the bone 3.3 Leukocytes and Leukocyte marrow prior to release of leukocytes into the peripheral Responses in Disease blood. Toxic changes in heterophils should be reported both by the number of heterophils affected and the severity. The Leukocytes of reptiles include granulocytes (heterophils, most severe toxic changes are due to bacterial toxins, and eosinophils, basophils) and mononuclear cells (lymphocytes, are often related to enteric disease or other Gram-negative monocytes). There is a wide variation among different species bacterial infections. Toxicity is recognized by increased cytoin both cell size and morphology of cytoplasmic granules, and plasmic basophilia, degranulation, vacuolation, abnormal in the nuclei of circulating granulocytes. Lymphocytes and cytoplasmic granulation, and excessive nuclear lobation (Figmonocytes found in the peripheral blood of reptiles resemble ures 3.33–3.37). These toxic changes vary in intensity with those in mammals. However, there is variability of leukocyte severity of disease. Mild toxicity is associated with increased percentages within species and even genera (Alleman et al., cytoplasmic basophilia, with heterophil granules being nor1999). The cellular descriptions used in this text are based on mal in number and shape. Degranulation may be an artifact the leukocyte morphology in Romanowsky-type stains. from the blood film preparation or prolonged storage in anticoagulant (Figure 3.38). However, in the presence of other 3.3.1 Heterophils toxic changes, such as cytoplasmic basophilia, vacuolation, Heterophils generally are large, round cells with clear cyto- and abnormal granulation, it can be interpreted as an indicaplasm that contains numerous pink-orange, spindle-shaped tor of toxicity. Abnormally colored or shaped granules accomcytoplasmic granules, which may partially obscure the nucleus pany severe toxicity. The granules of toxic heterophils may (Figures 3.29–3.32). The shape of the nucleus depends on be pleomorphic, dark basophilic to purple, and may be larger the species studied and may vary from a single round to oval, than the normal. Nuclear lobation in species that usually have eccentrically placed nucleus (most snakes, chelonians, and monomorphic round nuclei can also be interpreted as a toxic crocodilians) to a nucleus with two or more lobes (lizards). change (Campbell, 1996a) (Figure 3.39). Left shifting is indiThe average size of reptilian heterophils ranges from 10 to cated by the presence of myelocytes and metamyelocytes. 23 µm, generally varying between the species and even in Compared to mature heterophils, these immature cells may the individual blood sample (Saint Girons, 1970). The num- contain an enlarged nucleus, more pleomorphic granules, ber of cytoplasmic granules also varies among different spe- and more basophilic cytoplasm, which may contain primary cies. Granules of chelonians and crocodilians are eosinophilic granules (Figures 3.40–3.41). This condition is commonly and fusiform (Figures 3.29–3.30), whereas heterophils of associated with an increased tissue demand of heterophils, squamata have eosinophilic, angular, or pleomorphic gran- such as severe infection. However, both phenomena, left ules in their cytoplasm and one round or multilobed nucleus shifting and toxicity, may occur concurrently and have to be (Figures 3.31–3.32) (Montali, 1988). Snake heterophils have interpreted in the light of leukocyte counts and clinical findabundant, poorly formed pleomorphic elongated granules, ings of the patient. Intracytoplasmic bacteria may be seen in which are frequently so dense that they appear as fused heter- the peripheral blood of severely septicemic animals, but is ogenous eosinophilic material (Figure 3.31). They typically rarely observed (Figure 3.42). Degranulated heterophils (Figure 3.43) are commonly have one eccentrically placed, monomorphic round nucleus. Snake heterophils have been described as two morphologi- seen in squamates (Alleman et al., 1999) and crocodilians. cally distinct forms with pleomorphic eosinophilic granules This phenomenon may be an artifact of sample handling, (irregularly, oval or elongated), which are hypothesized to prolonged storage in anticoagulant, or inadequate fixation. represent different stages of cell maturation (Alleman et al., However, it may also be part of the normal in vivo aging pro1999; Bounous et al., 1996; Salakij et al., 2002). Staining inten- cess of these cells, if nuclear pyknosis is also present (Allesity of heterophil granules may vary depending on their stage man et al., 1999). Based on cytochemical staining and ultrastructural studof development in the peripheral blood (Egami and Sasso, 1988). Lizards have spindle, rod-shaped, or pleomorphic ies, reptilian heterophils are assumed to be functionally cytoplasmic granules, and their nucleus may appear lobed in equivalent to mammalian neutrophils. The primary functions some species, as commonly seen in the green iguana (Fig- of these cells are phagocytosis and microbicidal activity, and a ure 3.32) (Montali, 1988). Crocodilians have larger but fewer heterophilic response is mainly associated with inflammatory granules compared to the numerous, smaller granules seen in disease (Azevedo and Lunardi, 2003; Duguy, 1970; Mateo et lizards (Frye, 1991). Crocodilian heterophils have round, oval, al., 1984; Montali, 1988; Sypek and Borysenko, 1988). Heteroor lenticular nuclei, and may rarely be binucleated (Mateo et phils from most reptile species stain negative for peroxidase enzyme, a recognized marker for mammalian neutrophils. al., 1984).

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However, heterophils of green iguanas stain positive with benzidine peroxidase and Sudan black B, indicating bactericidal capabilities and oxidative burst function in this and possible other lizard species (Harr et al., 2001). In the inland bearded dragon (Pogona vitticeps), heterophils have been shown to stain with Sudan black, an eosinophil and heterophil marker, which suggests that squamates may combine the function of heterophils and eosinophils into one cell (Eliman, 1997; Montali, 1988). Heterophils may account for between 30 to 45% of the total leukocyte count in healthy reptiles (Frye, 1991), and have been described as a predominant leukocyte type in most chelonians and crocodilians (Alleman et al., 1992; Christopher et al., 1999; Mateo et al., 1984; Montali, 1988). The mean heterophil count and total WBC count in crocodilians has been described as significantly higher in adult males than in adult females (Stacy and Whitaker, 2000). Highest heterophil numbers in reptiles are generally observed during the summer months and lowest during hibernation (Duguy, 1970). Heterophilia may be seen as a response to inflammation caused by infectious agents (microbes, parasites), tissue injury, or tissue necrosis. Gravidity, glucocorticoid administration, stress, neoplasia, and granulocytic leukemia have also been reported to cause increased numbers of heterophils (Campbell, 1996a; Cuadrado et al., 2002; Duguy, 1970). Heteropenia may be seen with acute, overwhelming infection, resulting in excessive tissue demand for heterophils. Other hematologic abnormalities, such as toxic changes and left shift of heterophils, will likely be observed concurrently.

3.3.2 Eosinophils The eosinophil is a large (9 to 20 µm), round cell, about equal in size to heterophils. They have a clear cytoplasm with distinct eosinophilic, spherical granules and one roundish elongated, lenticular or bilobed nucleus, which may be centrally or eccentrically placed (Figures 3.44–3.45) (Campbell, 1996a; Montali, 1988). Two types of eosinophils, namely small and large eosinophils, have been described in Hawaiian green turtles (Work et al., 1998). In some reptile species, such as iguanas, the eosinophilic granules may stain blue or bluegreen with Romanowsky-type stains (Hawkey and Dennett, 1989; Heard et al., 2004); they are referred to as green eosinophils (Figures 3.46–3.47). Eosinophil granules for most reptile species stain positive with benzidine peroxidase, which allows their differentiation from heterophils (Alleman et al., 1992; Sypek and Borysenko, 1988). However, this is not the case in the green iguana where heterophils stain peroxidase positive and green eosinophils are not known to stain with any cytochemical stains used thus far (Harr et al., 2001). The presence or absence of eosinophils is controversial in snakes. From the published literature, it appears that eosinophils may be present in some snake species, but not in others. Eosinophils in snakes have been described (Campbell, 1996a; Salakij et al., 2002; Sypek and Borysenko, 1988; Troiano et al.,

1997), but some authors suggest that they represent a granulocyte most consistent with a second heterophil type (Alleman et al., 1999; Dotson et al., 1995; Egami and Sasso, 1998; Montali, 1988). Eosinophil granules of king cobras have been described as variably sized, round pale basophilic granules that may obscure the nucleus, similar to those described in iguanas, or they are large spherical bulging, basophilic to purple granules (Salakij et al., 2002). Abnormal eosinophil morphology is rare. In the authors’ experience, severely left-shifted eosinophils contain intracytoplasmic pale to moderately basophilic, round primary granules admixed with spherical eosinophilic granules (Figure 3.48). Other eosinophil abnormalities, such as degranulation, are generally thought to be of no diagnostic value and usually an artifact of blood film preparation or inadequate staining. In healthy reptiles, eosinophil numbers vary from 7 to 20% (Frye, 1991), with higher prevalence in turtles but very low numbers in lizards (Sypek and Borysenko, 1988). The eosinophil numbers are influenced by seasonal factors, with lowest numbers reported in summer and highest numbers during the hibernation period (Duguy, 1970). Increased numbers of eosinophils may be associated with parasitism and nonspecific immune stimulation. Eosinophils are proven to participate in the immune response of chelonians and are found to phagocytize immune complexes (Mead and Borysenko, 1984). The significance of eosinopenia in reptiles is unknown.

3.3.3 Basophils Reptilian basophils are generally small cells with dark, round, metachromatic-staining granules that often obscure the dark purple, centrally located nucleus (Montali, 1988) (Figures 3.49–3.50). They range in size from 7 to 20 µm, with the smallest reported in lizards (Frye, 1991). When visible, the nucleus appears round and nonlobed, and is slightly eccentrically placed. Basophils may be degranulated and appear as small round cells with purple cytoplasm, which may show distinct, clear vacuoles (Figure 3.51). This may be the result of using water-based stains (Campbell, 1996a), but has also been noted by the authors when Romanowsky-type stains with alcohol fixation are used. The clinical significance of degranulation is unclear. The function of reptilian basophils appears to be similar to mammalian basophils because they are involved in the processing of surface immunoglobulins and releasing histamine (Mead et al., 1983; Sypek et al., 1984; Sypek and Borysenko, 1988). Basophil numbers are highly variable. They depend on species and possibly on influences from season, geographic region and age of the animal (Work et al., 1998). Healthy turtles and tortoises may have up to 40% basophils of the leukocyte differential (Alleman et al., 1992; Duguy, 1970; Sypek and Borysenko, 1988). Seasonal variations have been reported as minimal (Saint Girons, 1970), with lower baso-

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phil numbers during hibernation in desert tortoises (Christopher et al., 1999) and increased numbers with activity (Saint Girons, 1970). Basophilia may be associated with blood parasites (e.g., hemogregarines and trypanosomes), as well as with viral infection (e.g., iridovirus infection) (Sypek and Borysenko, 1988).

3.3.4 Lymphocytes Reptilian lymphocytes resemble those in mammals and also vary in size from small to large, with a diameter from 5 to 10 µm in small and up to 15 µm in large lymphocytes (Figures 3.52 and 3.53) (Saint Girons, 1970). The most critical aspect of the leukocyte differential in reptiles is the differentiation between small lymphocytes and thrombocytes (Figure 3.54). Compared to thrombocytes, lymphocytes are larger with distinct borders. They are round to oval in shape with scant, lightly to moderately basophilic cytoplasm. The lymphocyte nuclei are larger, round to oval in shape, and centrally or eccentrically located. Their chromatin is clumped and they typically exhibit a high nuclear-to-cytoplasmic ratio. Immature lymphocytes are large with a scant amount of moderately to intensely basophilic cytoplasm and a round to oval nucleus that contains smooth lighter purple-staining chromatin, which may contain a nucleolus (Figure 3.55). They may be present in very low numbers in healthy animals. Except for the American alligator, reptilian lymphocytes generally stain negative with all cytochemical stains, which is specifically helpful in the differentiation from periodic acid-Schiff (PAS)–positive thrombocytes (Alleman et al., 1992; Alleman et al., 1999; Mateo et al., 1984, Salakij et al., 2002). Lymphocytes of American alligators stain PAS-positive, whereas thrombocytes are PAS-negative (Mateo, 1984). Reactive lymphocytes are an indication of antigenic stimulation. They are typically larger than small, well-differentiated lymphocytes and generally have abundant, moderately to intensely basophilic cytoplasm that may contain a small amount of discrete, punctate vacuoles (Figure 3.56). Some reactive lymphocytes are plasmacytoid (plasma cell-like) in their appearance (Figures 3.57–3.58). Some cells may contain a variable number of small round or needle-like azurophilic granules (Figure 3.59–3.60). Very low numbers of immature lymphocytes, lymphoblasts, with smooth chromatin and one or two prominent nucleoli, may also be identified in animals with disease conditions that result in antigenic stimulation (Figures 3.55 and 3.61). As in mammals, reptilian lymphocytes are classified as B- and T-lymphocytes, whereby B-cells produce certain immunoglobulins and T-cells are responsible for an adequate cellular immune response (Sypek and Borysenko, 1988). Lymphocytes are the most prevalent leukocyte type in the peripheral blood and hematopoietic tissues of most reptile species, with percentages being as high as 80% (Alleman et al., 1999; Lamirande et al., 1999; Salakij et al., 2002; Sypek and Borysenko, 1988; Troiano et al., 1997; Work et al., 1998).

Lymphocyte numbers are influenced by seasonal and individual factors. They are reported to be lowest during winter or hibernation periods and highest during summer months as well as during ecdysis (Christopher et al., 1999; Wallach and Boever, 1983). The lower number of lymphocytes during these times has been related to a relative inability of some temperate species to produce a primary immune response during low temperatures or hibernation periods (Wright and Cooper, 1981). Furthermore, females tend to have higher lymphocyte numbers under identical conditions than males of the same species and age (Duguy, 1970; Sypek and Borysenko, 1988). Adult crocodilians have significantly lower lymphocyte counts and total WBC counts than juveniles and subadults (Stacy and Whitaker, 2000). Lymphocytosis may occur with wound healing and with infectious or inflammatory disease, most likely representing a chronic component. Viral diseases and certain parasitic diseases, such as anasakiasis, spirorchidiasis, or hematozoa, may also cause a peripheral lymphocytosis (Campbell, 1996a). Other causes of lymphocytosis include lymphoid leukemias and inclusion body disease (IBD) of boid snakes. In the early stage of IBD, leukocyte counts of greater than 100,000/µl have been documented, with most leukocytes being lymphocytes. Intracytoplasmic IBD inclusions were noted in some of the peripheral blood lymphocytes (see Section 3.5.2) (Jacobson, 1999). Lymphopenia has been reported with malnutrition (Campbell, 1996a) and with endogenous or exogenous corticosteroids.

3.3.5 Plasma Cells Plasma cells are round to oval with distinct borders and abundant intensely basophilic cytoplasm, which contains a pale perinuclear Golgi zone (Sypek and Borysenko, 1988) (Figure 3.62). They typically have an eccentrically placed nucleus with coarse chromatin, and have a lower nuclearto-cytoplasmic ratio compared to unstimulated lymphocytes. When immunologically stimulated, plasma cells may contain variable numbers of clear or pale basophilic vacuoles (Russell-bodies), using Romanowsky-type stains. These are referred to as Mott cells. Plasma cells may be observed in increased numbers in blood from reptiles with severe infectious or inflammatory disease. In healthy reptiles, plasma cells may comprise 0.2 to 0.5% of the leukocyte percentage (Frye, 1991).

3.3.6 Monocytes The largest leukocytes in the peripheral blood of reptiles are monocytes, with a size range reported to vary from 8 to 25 µm. They are similar to their counterparts in mammalians with regard to morphology and function (Sypek and Borysenko, 1988). They are round to oval with distinct borders and abundant grayish, pale to moderately basophilic cytoplasm

182 Circulating Inflammatory Cells

(Figures 3.63–3.64). Depending on their reactive stage, monocytes may contain variably sized, distinct, cytoplasmic vacuoles (Figures 3.65–3.66). Their pleomorphic nucleus may appear round, oval, indented, U-shaped, or lobulated and is comprised of finely clumped chromatin. Compared to well-differentiated lymphocytes, the nuclear chromatin stains paler purple in color. The authors have observed a leukocyte type distinctive from the normal monocyte in lizard species, including the monitor lizard, tegu lizard, and chameleon. Although morphologically different, the cell is most consistent with the monocyte cell line. These leukocytes are about the size of heterophils with light basophilic cytoplasm that contains very small, dust-like pink granules. Their eccentric nucleus is bilobed, band shaped, or trilobed (Figures 3.67–3.68). Monocyte numbers may account for up to 10% of leukocytes (Sypek and Borysenko, 1988). In some South African reptiles, monocytes have been reported at up to 20% of total leukocytes (Pienaar, 1962). Monocyte numbers change minimally with seasonal variation and their relative percentage remains fairly constant (Duguy, 1970; Sypek and Borysenko, 1988). However, high monocyte numbers have been described during hibernation of desert tortoises (Christopher et al., 1999) and in dystocic chameleons (Cuadrado et al., 2002). Monocytes increase with antigenic stimulation and are suggestive of a chronic infectious process. They are involved in granuloma and giant cell formation and are specifically associated with granulomatous responses to bacterial infections and to ova of spirorchid trematodes (Campbell, 1996a). A chlamydia-like organism and pox-like virus were identified by Jacobson and Telford (1990) in circulating monocytes of flap-necked chameleons (Chamaeleo dilepis) (Figure 3.69).

3.3.7 Azurophils This leukocyte type is commonly observed in squamates and crocodilians, but only occasionally in chelonians (Alleman et al., 1999; Dotson et al., 1995; Hawkey and Dennett, 1989). This unique leukocyte type combines morphologic features of monocytes and granulocytes in one cell (Montali, 1988). Azurophils are large round cells with fine, dustlike, azurophilic to purple cytoplasmic granules, and may contain a low number of clear, punctuate vacuoles. They typically have one round to oval, slightly eccentrically placed nucleus with clumped chromatin (Figures 3.70–3.71). Compared to mature azurophils, immature azurophils have a larger nucleus, which may be oval to pleomorphic in shape, and a higher nuclear-to-cytoplasmic ratio (Figure 3.72). Cytochemical staining functionally differentiates snake azurophils from other cells with azurophilic granules found in other reptile species. Snake azurophils stain positive with benzidine peroxidase, Sudan black B and PAS, similar to mammalian neutrophils (Alleman et al., 1999). They are phagocytic cells, which have been proven to mount an oxidative burst similar to the mammalian neutrophil (Heard et al., 2004). Azurophils are the second most frequently found leuko-

cyte type found in the peripheral blood of snakes (Alleman et al., 1999; Salakij et al., 2002). However, increased percentages of mature or immature azurophils are associated with inflammatory or infectious disease (i.e., bacterial infections) in snakes, particularly in acute stages of disease (Jacobson et al., 1997). In snakes, cells resembling mammalian monocytes may be seen, but are thought to be a reactive form of azurophils in response to inflammation or an infectious process (Dotson et al., 1995). Azurophils in lizards are benzidine peroxidase and Sudan black B negative, and are considered to be monocytoid in their origin (Harr et al., 2001). It is suggested that they are referred to as monocytes or azurophilic monocytes (Heard et al., 2004). They are found only in lower percentages and increased numbers are interpreted as for monocytes.

3.3.8 Tumors of Hematopoietic Tissue In reptiles, the most commonly reported hematopoietic tumors are lymphoid malignancies, predominantly seen in snakes and lizards (Garner et al., 2004). Lympho- and myeloproliferative diseases may be accompanied by varying degrees of leukopenia or leukocytosis, as reported in a lymphoid leukemia in an Aruba Island rattlesnake (Crotalus unicolor), with > 100,000 cells/µl (Lock et al., 2001). Atypical cells, such as blast cells (Figure 3.73) or haystack-like needle-shaped crystalline inclusions in granulocytic leukocytes may be identified in the peripheral blood, as reported in an iguana with myeloid leukemia (Frye, 1991). In lizards, leukemia of undetermined origin (Goldberg and Holshuh, 1991), myelogenous leukemia (Tocidlowski et al., 2001), chronic monocytic leukemia (Gregory et al., 2004) and lymphosarcoma (Schultze et al., 1999) have been reported. Various species of snakes have been diagnosed with acute lymphocytic leukemia (Frye and Carney, 1973), myelogenous leukemia (Hruban et al., 1992), lymphocytic leukemia with multicentric T-cell lymphoma (Raiti et al., 2002), or lymphosarcoma with lymphoid leukemia (Lock et al., 2001). Myeloproliferative disease has also been described in two turtles, a gecko, and a rattlesnake (Frye and Carney, 1972; Marcus 1973; Garner et al., 2004). Bone marrow examination, along with special cytochemical stains may be helpful in the assessment of these diseases.

3.4 Thrombocytes and Thrombocyte Responses in Disease Mammalian platelets are anucleated, cytoplasmic fragments of megakaryocytes. However, thrombocytes in nonmammalian species are nucleated cells, which originate from a distinct cell line in the bone marrow, with the thromboblast being the most likely immature precursor. Thrombocytes are elliptical to round with distinct borders, and have one centrally placed, round or ovoid, intensely violet-staining nucleus with dense chromatin (Figure 3.74).

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Inactivated thrombocytes have clear cytoplasm, which may occasionally show fine azurophilic granules (Figure 3.75). Thrombocytes generally measure 8 to 16 µm in length and 5 to 9 µm in width (Sypek and Borysenko 1988). During blood film preparation, they are easily ruptured or activated (Campbell, 1995). When ruptured, the thrombocytes appear as free nuclei with smooth margins (Figure 3.76). Like platelets, they undergo shape change, degranulation, and aggregation on activation. Therefore, thrombocytes are usually aggregated on blood films, which may help in their identification and differentiation from lymphocytes (Figure 3.77). Activated thrombocytes may form pseudopods with irregular cytoplasmic borders (Figure 3.78) or may have cytoplasmic vacuolation (Figure 3.79), or both. As previously mentioned, cytochemical staining with periodic acid-Schiff (PAS) is focally positive in reptilian thrombocytes and negative in lymphocytes, allowing differentiation between these two cell lines (Alleman et al., 1992; Alleman et al., 1999; Salakij et al., 2002). The finding is opposite in the American alligator (Mateo et al., 1984). Thrombocytes play a major role in thrombus formation and are part of wound healing (Sypek and Borysenko, 1988). They also may have limited phagocytic ability (Dieterlen-Lievre, 1988). Bacteria, tissue debris, senescent erythrocytes, and hemosiderin have been identified in the cytoplasm of reactive thrombocytes (Figure 3.80). Thrombocyte numbers have been reported ranging from 25 to 350 cells per 100 leukocytes (Pienaar, 1962). Immature thrombocytes are larger than the mature form and their cytoplasm has a bluish tinge. Increased numbers of immature thrombocytes, usually accompanied by a concurrent thrombocytosis, indicate a regenerative response with increased bone marrow output as a result of enhanced peripheral utilization or destruction. Binucleated thrombocytes may be observed in anemic patients (Frye, 1991) (Figure 3.81). In severe inflammatory disease, nuclei of thrombocytes may become polymorphic. This may also be seen in reptiles with conditions resulting in prolonged anorexia (Hawkey and Dennett, 1989). In animals with suspected thrombocytopenia, it should be first confirmed that the count is not falsely reduced as a result of collection, delay in sample processing, or laboratory error. As previously described, a common cause of pseudothrombocytopenia is thrombocyte clumping. Causes for thrombocytopenia are many and may include decreased platelet production as a result of infectious, toxic, or neoplastic bone marrow disease. Other causes include accelerated platelet destruction or use resulting from infectious, inflammatory, or immune-mediated disease.

3.5 Infectious Agents in the Peripheral Blood Blood parasites are commonly found in reptiles, particularly in wild-caught animals, and are usually considered an inci-

dental finding and nonpathogenic. However, some hemoparasites have the potential for causing clinical disease, such as anemia. Predisposing factors, such as stress of other pathogens or inadequate husbandry, may increase the potential for hemoparasites to cause clinical disease. Hemoparasites are identified within their target cells or free in the plasma and include the diverse group of hemoprotozoa, piroplasmids, and filarial worms. Other infectious diseases, which may be detected in the peripheral blood, include viral inclusions and bacterial agents.

3.5.1 Hemoparasites Hemogregarines, plasmodiids, and trypanosomes are common hemoprotozoa in reptiles (Figures 12.1–12.7). Generally, hemoparasitic protozoans require invertebrates as intermediate hosts, such as arthropod or annelid vectors (Telford, 1984). For detailed information regarding the biology of parasites, the reader is referred to Chapter 12, Section 12.2.

3.5.1.1 Hemogregarines  Four genera of intracellular parasites are included among the hemogregarines: Hemogregarina, Hepatozoon, Karyolysus, and Hemolivia. These genera cannot be accurately classified based on their appearance in blood cells alone (Telford, 1984). On Romanowsky-stained blood films, the gamonts of hemogregarines appear as sausage-shaped inclusions with pale to purple cytoplasm and one centrally to slightly eccentrically placed, darker purplestaining nucleus (Figures 3.44, 12.78–12.79), except in Hemogregarina infections where erythrocytic meronts may be present. These are unpigmented and are typically found in the cytoplasm of red, and sometimes in white blood cells (Jacobson, 1983). The gamonts may push the nucleus of the host cells to one side or surround it. The host cells may appear irregular in shape and size (Lane and Mader, 1996). Rarely, two or more organisms may be found in one erythrocyte, or the gamont may be found extracellularly. Because gamonts of different hemogregarines are morphologically indistinguishable in the peripheral blood, the general term hemogregarine is used to report their presence (Keymer, 1981). Hemogregarines belonging to the genus Hepatozoon are commonly found in terrestrial and aquatic snakes (Figures 3.82–3.83). Hemogregarine sporozoites are often transmitted by infected arthropods and leeches (Frye, 1991; Salakij et al., 2002). Freshwater turtles and alligators are usually infected with Hemogregarina.  Although hemogregarines are considered nonpathogenic, a correlation between the detection of hemogregarines in the peripheral blood with granulomas of Hepatozoon meronts in the liver was reported in a snake (Wozniak et al., 1998). Hemogregarines are well adapted to their natural host, but can cause significant clinical inflammatory disease in unnatural host species (Wozniak and Telford, 1991). Hemogregarines and Hepatozoon in snakes and lizards have been shown to have potential for congenital and oral transmission (Telford, 1984).

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3.5.1.2 Hemococcidia   Schellackia and Lainsonia occur in the peripheral blood of lizards. Lainsonia is commonly found in wild-caught Latin American green iguanas (Frye, 1991; Harr et al., 2001). Sporogony and merogony take place in the host, and the intermediate hosts serve as mechanical vectors only. The parasite is transmitted when mosquitoes or bloodsucking mites take a blood meal from an infected host and are then ingested by the reptile. Meronts are found in the intestinal epithelium and sporozoites can be found in leukocytes (primarily lymphocytes) and erythrocytes, where they form round, oval, or elongate inclusions (Keymer, 1981).

3.5.1.3 Plasmodium  Five genera of the family Plasmodiidae are reported: Plasmodium, Fallisia, Saurocytozoon, Haemocystidium, and Haemoproteus. Snakes and turtles may become infected with Hemoproteus, whereas Plasmodium, Fallisia, Saurocytozoon, and Haemocystidium infections have been reported in lizards (Lane and Mader, 1996). Over ninety species and subspecies of Plasmodium have been described in reptiles (Telford, 1984). These fairly common malarial parasites have been mostly identified in semiaquatic and terrestrial chelonians and many lizards, and occasionally in snakes (Telford, 1984). Sporogony occurs in the invertebrate host, which transmits infectious sporozoites via its next blood meal. Merogony and gametogony take place in the reptilian host (Telford, 1984). Therefore, gametocytes, meronts, and trophozoites can be found in their erythrocytes as well as extracellularly. Plasmodium gametocytes are round, oval, or elongate, pale eosinophilic, deep pink or bluish structures that often contain many golden brown to black, mostly refractile pigment granules (Figure 3.84). Pigment granules are considered to be from hemoglobin breakdown products, and can also be seen in Haemoproteus sp. and Haemocystidium sp. These pigment granules in Plasmodium and Haemoproteus help in differentiation from hemogregarines, which lack any pigment (Campbell, 1996b). Trophozoites of Plasmodium may appear as focal packets or signet-ring structures in the erythrocyte cytoplasm (Keymer, 1981). Saurocytozoon produces large round gametocytes in leukocytes and lack the pigment granules found in Plasmodium and Hemoproteus (Keymer, 1981) (Figure 3.85). In most instances, natural infections in members of the family Plasmodiidae are considered nonpathogenic, but cases of mild anemia have been reported (Lane and Mader, 1996). Chronic infections are thought to persist throughout life (Frye, 1991). However, cases of severe anemia have been described as well, and some forms of Plasmodium sp. are considered a potential cause for thrombosis in capillary beds (Frye, 1991).

3.5.1.4 Trypanosomes  These are large flagellate protozoa that possess a kinetoplast with a distinct or indistinct, undulating membrane (trypomastigote). These parasites may be found extracellularly in the peripheral blood of many reptile species (Figures 3.86, 12.7). They are transmitted by bloodsucking arthropods (phlebotomine sand flies, biting dipteran

flies) in terrestrial reptiles, and by leeches in aquatic reptiles (Frye, 1991; Keymer, 1981). Although trypanosomiasis can cause severe parasitemia, it is commonly associated with lifelong subclinical infections (Lane and Mader, 1996).

3.5.1.5 Piroplasmida  Members of this group include Aegyptianella (Tunetella) and Sauroplasma (Serpentoplasma, Chelonoplasma) and are rare blood parasites of chelonians, lizards, and snakes (Keymer, 1981). These protozoal hemoparasites appear as very small (1 to 2 µm in diameter) punctuate basophilic inclusions of erythrocytes (Figure 3.87), usually in small aggregates, surrounded by clear vacuoles (Frye, 1991).

3.5.1.6 Sauroleishmania  This protozoan is related to trypanosomes and has been reported in the peripheral blood of reptiles, primarily in lizards (Keymer, 1981). The amastigote form of the organism appears in the cytoplasm of blood cells, particularly erythrocytes. It appears singly or may be numerous, and may condense to round basophilic inclusions with a central hollow (Paperna et al., 2001) (Figure 3.88). In mammals, the organism is found in macrophages and has a characteristic, distinctive, bar-shaped extrachromosomal DNA fragment called a kinetoplast. The motile promastigote form is found extracellularly and possesses a flagellum (Frye, 1991). Because the organism is rarely identified in peripheral blood film preparations, culture techniques are primarily used (Novy, MacNeal and Nicolle´s [NNN] medium; Campbell, 1996b). In reptiles, Sauroleishmania is presumably transmitted by phlebotomine sandflies (Frye, 1991).

3.5.1.7 Microfilaria  Various genera of filarid worms can be found in reptiles, some of which are specific, such as: Macdonaldius (snakes, lizards), Saurositus (lacertid lizards), Foleyella (chameleons, lacertid lizards), Oswaldofilaria (crocodilians, lacertid lizards), and Cardianema (chelonians, lacertid lizards) (Lane and Mader, 1996). Microfilaremia is usually an incidental finding when found on Romanowsky-stained blood films from reptiles (Figures 3.89–3.90, 12.148). Filariasis is mostly subclinical, but with heavy infestation, thrombosis and blockage of blood vessels may occur resulting in edema. fibrosis, and/or necrosis in the affected area (IrizarryRovira et al., 2002). Microfilaria are transmitted by bloodsucking mosquitoes or ticks (Lane and Mader, 1996).

3.5.1.8 Spirorchiidae  These digenetic flukes develop in the circulatory system of reptiles, particularly chelonians. The cercariae stage penetrates skin or mucous membranes of the host and matures in its heart and blood vessels. The adult flukes release eggs, which penetrate the vessel wall or accumulate in terminal capillaries (Lane and Mader, 1996). Eggs may be found in the peripheral blood of heavily infected animals.

Circulating Inflammatory Cells  185

3.5.2 Viral Inclusions in Blood Cells

to be the causative agent of the disease (Marquardt and Yaeger, 1967). Based on ultrastructural studies, viral particles were 3.5.2.1 Inclusion Body Disease (IBD)  This disease is identified and the name snake erythrocyte virus (SEV; Johndescribed in boid snakes (i.e., boa constrictors and pythons) srude et al., 1997; Smith et al., 1994) was given to the virus. and is diagnosed by the finding of inclusions in a variety of The crystalline inclusions are probably comprised of cellular tissues, including circulating lymphocytes and rarely thrombo- and viral byproducts of lipids and proteins (Johnsrude et al., cytes and basophils. They may be identified in granulocytes 1997). Iridovirus infection has been reported to be pathogenic as well, since inclusions have been described in myeloid pre- in squamatans and may cause severe anemia (Johnsrude et cursors in the bone marrow (Garner and Raymond, 2004). A al., 1997; Telford, 1984). Recently, iridoviral inclusions have retrovirus has been proposed as the causative agent of IBD been identified in the leukocytes (monocytes, azurophils, (Jacobson et al., 2001). The origin of the inclusions is not and heterophils) of an eastern box turtle (Terrapene carolina) known thus far, but it has been shown that they are comprised (Allender et al 2006). These inclusions were single, round to of an IBDV-associated protein (Wozniak et al., 2000). Using oval, pink, and granular, measuring 3 to 7 µm in diameter. Romanowsky-type stains, these inclusions appear as oval to Some inclusions were found as multiple fragments. They were lentiform, homogenously pale basophilic intracytoplasmic observed in monocytes, azurophils, and heterophils (Figinclusions of variable sizes (Figures 3.91–3.96, 9.132). The ures 3.103–3.105). The isolated virus was identified as frog viral inclusions are distinctly outlined against the background virus 3 (FV3) (family Iridoviridae, genus Ranavirus). of scant cytoplasm, which they may nearly fill completely. Fre3.5.2.3 Poxvirus  Poxviral inclusions have been described quently, the inclusion pushes the nucleus of leukocytes aside, by Jacobson and Telford (1990) in the peripheral blood of a giving them a half-moon-shaped appearance. Hematoxylin flap-necked chameleon. These inclusions were identified as and eosin stain can also be used to identify these inclusions membrane-bound, pleomorphic, basophilic to purple incluon blood films. With this stain they appear pale to moderately sions in circulating monocytes (Figures 3.69, 9.61). eosinophilic. Identification of inclusions on the blood film is diagnostic. However, the absence of circulatory intracytoplas3.5.2.4 Nonviral Inclusions  Viral inclusions have to be difmic inclusions does not necessarily rule out IBD (Jacobson, ferentiated from hemoglobin crystals, degenerate organelles, 2002). If inclusion bodies are not identified in the peripheral cellular debris, phagocytized material, and hemoparasites. blood, the preferred diagnostic samples for an antemortem Healthy iguanas have been reported with clear symmetrical diagnosis of IBD are biopsies of liver, stomach, or esophahexagonal inclusions (Figures 3.26–3.27) that resembled geal tonsils, (Jacobson, 2002). Cytology of H&E-stained tissue hemoglobin crystals (Harr et al., 2001; Simpson et al., 1980). imprints may be helpful to provide more rapid information Small, basophilic punctate or clear ring-shaped inclusions than histology, but may cause false positive or false negative in red blood cells of the desert tortoise (Gopherus agassizii) results (Jacobson, 2002; Garner and Raymond, 2004). For furhave been identified as degenerate organelles (Alleman et al., ther information, the reader is referred to Chapter 9. 1992) (Figures 3.23–3.25). These inclusions may be present 3.5.2.2 Iridovirus  Iridoviral inclusions have been described in a variable number of erythrocytes in the peripheral blood of reptiles with no known clinical significance. Azurophils in blood cells from lizards, snakes, and turtles (Marquardt and or monocytes containing clear punctuate vacuoles of variYaeger, 1967; Allender et al 2006). Their morphology is hetable sizes probably contain lipid, which has to be differenerogenous among species. In lizards, these inclusions appear tiated from viral inclusions as well (Garner and Raymond, in erythrocytes as small acidophilic punctuate to oval inclu2004) (Figure. 3.106). Melanin-containing macrophages or sions that may be associated with rectangular albuminoid monocytes with phagocytized cells or cellular material can vacuoles (Telford and Jacobson, 1993). The identification of also be identified in the peripheral blood of healthy reptiles these inclusions was formerly described as pirhemocytonosis (Figures 3.107–3.110). (Daly et al., 1980). Based on transmission electron microscopy, these inclusions were identified as viral particles consistent 3.5.3 Bacteria with those of the family Iridoviridae. In chameleons, the virus was named lizard erythrocytic virus (LEV; Telford and Jacob- A variety of bacterial organisms has been described in the son, 1993) (Figures 3.97, 9.71–9.72). Iridoviral infections in peripheral blood of reptiles. An extracellular, heterogenous snakes may have two types of inclusions in erythrocytes, one group of bacteria may be seen associated with scales or kerativiral (small aggregates of granular eosinophilic material) (Fig- naceous material, and would indicate skin contamination. Stain ure 9.73) and one crystalline (translucent, hexagonal and flat) contamination can be another source of extracellular bacteria (Figures 3.98–3.102) (Johnsrude et al., 1997). As the infec- observed in the blood film. In cases of septicemia, intracellution progresses, the crystalline inclusions increase in size, with lar and extracellular monomorphic bacteria would be present typically a single inclusion per red blood cell. Two inclusions (Figure 3.111). Heterophils or monocytes with phagocytized may occasionally be noted. Before identification of iridovirus, bacteria, observed in septicemic patients, indicate a guarded a protozoan parasite formerly named Toddia was assumed prognosis (Figures 3.42, 3.112). In these cases, blood culture

186 Circulating Inflammatory Cells

and sensitivity are indicated. Septicemia, when observed during morphologic evaluation, warrants antibiotic therapy. Simpson et al. (1981) have described a spiral-shaped bacterium in the peripheral blood and bone marrow of a rhinoceros iguana (Cyclura cornuta) (Figures 3.113, 6.106). This bacterium shared similarities with members of the family Spirillaceae Chlamydia has also been reported in circulating monocytes of flap-necked chameleons (Jacobson and Telford, 1990) (Figures 3.69,  9.61). Chlamydophila pneumoniae inclusion bodies have been observed in up to 2% of circulating monocytes of emerald tree boas that showed repetitive regurgitation ([Corallus caninus] Jacobson et al., 2007) (Figure 3.114).

Acknowledgments The authors thank the Clinical Pathology Service, College of Veterinary Medicine, University of Florida, for their technical assistance in performing the blood films.

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Miyamoto M, Vidal BC, and Mello ML. 2005. Chromatin supraorganization, DNA fragmentation, and cell death in snake erythrocytes. Biochem Cell Biol 83:15–27. Montali RJ. 1988. Comparative pathology of inflammation in the higher vertebrates (reptiles, birds and mammals). J Comp Pathol 99:1–26. Moon PF and Hernandez-Divers SM. 2001. Reptiles: aquatic turtles (chelonians), in Zoological Restraint and Anesthesia, Heard D (Ed.), International Veterinary Information Service, Ithaca, NY, www.ivis.org. Muro J, Cuenca R, Pastor J, Vinas L, and Lavin S. 1998. Effects of lithium heparin and tripotassium EDTA on hematologic values of Hermann’s tortoises (Testudo hermanni). J Zoo Wildl Med 29:40–44. Nagy KA and Medica PA. 1986. Physiological ecology of desert tortoises in Southern Nevada. Herpetologica 42:73–92. Natt MP and Herrick CA. 1952. A new blood diluent for counting the erythrocytes and leukocytes of the chicken. Poult Sci 31:735–738. Olson GA, Hessler JR, and Faith RE. 1975. Techniques for blood collection and intravascular infusions of reptiles. Lab Anim Sci 25:783–786. Ottaviani G and Tazzi A. 1977. The lymphatic system, in Biology of the Reptilia, Gans C and Parsons TS (Eds.), Vol. 6, Morphology E. Academic Press, New York, 315–462. Owens DW and Ruiz GJ. 1980. New methods of obtaining blood and cerebrospinal fluid from marine turtles. Herpetologica 36:17–20. Paperna I, Boulard Y, Hering-Hagenbeck SH, and Landau I. 2001. Description and ultrastructure of Leishamnia zuckermani n.sp. amastigotes detected within the erythrocytes of the South African gecko Pachydactylus turneri Gray, 1864. Parasite 8:349–353. Pati AK and Gupta S. 1991. Circadian time dependence of erythropoietic and respiratory responses of Indian garden lizard, Calotes versicolor, to mammalian urinary erythropoietin and thyroxine. Gen Comp Endocrinol 82:345–354. Pati AK and Thapliyal JP. 1984. Erythropoietin, testosterone, and thyroxine in the erythropoietic response of the snake, Xenochropis piscator. Gen Comp Endocrionol 53:370–374. Pienaar, U de V. 1962. Haematology of Some South African Reptiles. Witwatersrand University Press, Johannesburg, South Africa, 1–299. Pough FH. 1969. Environmental adaptations in the blood of lizards. Comp Biochem Physiol 31:885–901. Prado JL. 1946. Inactive (non-oxygen-combining) hemoglobin in the blood of Ophidia and dogs. Science 103:406. Prezant RM, Isaza R, and Jacobson ER. 1994. Plasma concentrations and disposition kinetics of enrofloxacin in gopher tortoises (Gopherus polyphemus). J Zoo Wildl Med 25:82–87. Raiti P, Garner MM, and Wojcieszyn J. 2002. Lymphocytic leukemia and multicentric T-cell lymphoma in a diamond python, Morelia spilota spilota. J Herpetol Med Surg 12:26–29. Rodkey FL, Robertson RF, and Kim CK. 1979. Molar absorbance of cyanmethemoglobin from blood of different animals. Am J Vet Res 40 :887–888. Rosskopf WJ Jr. 1982. Normal hemogram and blood chemistry values for California desert tortoises. Vet Med Small Anim Clin 77:85–87. Rucknagel KP and Braunitzer G. 1988. Hemoglobins of reptiles. The primary structure of the major and minor hemoglobin component of adult Western Painted Turtle (Chrysemys picta bellii). Biol Chem Hoppe Seyler 369:123–131.

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Saint Girons MC. 1970. Morphology of the circulating blood cell, in Biology of the Reptilia, Vol. 3, Gans C and Parsons TC (Eds.), Academic Press, San Diego, CA, 73–91. Salakij C, Slakij J, Apibal S, Narkkong NA, Chanhome L, and Rochanapat N. 2002. Hematology, morphology, cytochemical staining, and ultrastructural characteristics of blood cells in king cobras (Ophiophagus hannah). Vet Clin Pathol 31:116–126. Samour HJ, Risley D, March T, Savage B, Nieva O, and Jones DM. 1984. Blood sampling techniques in reptiles. Vet Rec 114:472–478. Schermer S. 1967. The Blood Morphology of Laboratory Animals, 3rd ed, F.A. Davis, Philadelphia, PA, 137–169. Schultze AE, Mason GL, and Clyde VL. 1999. Lymphosarcoma with leukemic blood profile in a Savannah monitor lizard (Varanus exanthematicus). J Zoo Wildl Med 30:158–164. Sheeler P and Barber AA. 1965. Reticulocytosis and iron incorporation in the rabbit and turtle: a comparative study. Comp Biochem Physiol 16: 63–76. Simpson CF, Jacobson ER, and Harvey JW. 1980. Noncrystalline inclusions in erythrocytes of a rhinoceros iguana. Vet Clin Pathol 9:24–26. Simpson CF, Jacobson ER, and Harvey JW. 1981. Electron microscopy of a spiral-shaped bacterium in the blood and bone marrow of a rhinoceros iguana. Can J Comp Med 45:388–391. Simpson CF, Taylor WJ, and Jacobson ER. 1982. Sickling hemoglobin polymerization in iguana erythrocytes. Comp Biochem Physiol A 73:703–708. Smith TG, Desser SS, and Hong H. 1994. Morphology, ultrastructure, and taxonomic status of Toddia sp. in northern water snakes (Nerodia sipedon sipedon) from Ontario, Canada. J Wildl Dis 30:169–175. Smits AW and Kozubowski MM. 1985. Partitioning of body fluids and cardiovascular responses to circulatory hypovolaemia in the turtle Pseudemys scripta elegans. J Exp Biol 116: 237–250. Stacy BA and Whitaker N. 2000. Hematology and blood biochemistry of captive mugger crocodiles (Crocodylus palustris). J Zoo Wildl Med 31:339–347. Stephens GA and Creekmore JS. 1983. Blood collection by cardiac puncture in conscious turtles. Copeia 1983:522–523. Sullivan B and Riggs A. 1964. Haemoglobin: reversal of oxidation and polymerization in turtle red cell. Nature 204:1098–1099. Sypek J and Borysenko M. 1988. Reptiles, in Vertebrate Blood Cells, Rowley AF, Ratcliffe NA (Eds.), Cambridge University Press, Cambridge, U.K., 211–256. Sypek JP, Borysenko M, and Findlay SR. 1984. Anti-immunoglobulin induced histamine release from naturally abundant basophils in the snapping turtle, Chelydra serpentina. Dev Comp Immunol 8:359–366. Taylor RW Jr. and Jacobson ER. 1982. Hematology and serum chemistry of the gopher tortoise, Gopherus polyphemus. Comp Biochem Physiol A 72:425–428. Telford SR Jr. 1984. Haemoparasites in reptiles, in Diseases of Amphibians and Reptiles, Hoff GL, Frye FL, and Jacobson ER (Eds.), Plenum Publishing Corporation, New York, 385–517. Telford SR Jr. 1989. Reptilian Haemosporozoa: A perception of life cycle patterns, in Proceedings of the Third Colloquium on Pathology of Reptiles and Amphibians, Orlando, FL, 48–49.

Telford SR Jr and Campbell HW Jr. 1981. Parasites of the American alligator, their importance to husbandry and suggestions toward their prevention and control, in Proceedings of the First Annual Alligator Production Conference, Cardeilhac P, Lane T, and Larsen R (Eds.), College of Veterinary Medicine, University of Florida, Gainesville, FL. Telford SR Jr and Jacobson ER. 1993. Lizard erythrocytic virus in east African chameleons. J Wildl Dis 29:57–63. Tocidlowski ME, McNamara PL, and Wojcieszyn JW. 2001. Myelogenous leukemia in a bearded dragon (Acanthodraco vitticeps). J Zoo Wildl Med 32:90–95. Torsoni MA and Ogo SH. 1995. Oxygenation properties of hemoglobin from the turtle Geochelone carbonaria. Braz J Med Bil Res 28:1129–1131. Torsoni MA, Stoppa GR, Turra A, and Ofo SH. 2002. Functional behavior of tortoise hemoglobin Geochelone denticulata. Braz J Biol 62:725–733. Troiano JC, Vidal JC, Gould J, and Gould E. 1997. Haematological reference intervals of the South American rattlesnake (Crotalus durissus terrificus, Laurenti, 1768) in captivity. Comp Haematol Int 1:109–112. Vasse J and Beaupain D. 1981. Erythropoiesis and hemoglobin ontogeny in the turtle Emys orbicularis L. J Embryol Exp Morphol 62:129–138. Wallach JD and Boever WJ. 1983. Diseases of Exotic Animals, Medical and Surgical Management. WB Saunders Co, Philadelphia, PA, 983–987. Walton RM. 2001. Establishing reference intervals: health as a relative concept. Sem Av Exotic Pet Med 10:66–71. Wojtaszek JS. 1991. Hematology of the grass snake Natrix natrix natrix L. Comp Biochem Physiol A 100:805–812. Work TM and Balasz GH. 1999. Relating tumor score hematology in green turtles with fibropapillomatosis in Hawaii. J Wildl Dis 35:804–807. Work TM, Raskin RE, Balazs GH, and Whittaker SD 1998. Morphologic and cytochemical characteristics of blood cells from Hawaiian green turtles. Am J Vet Res 59:1252–1257. Wozniak EJ and Telford SR Jr. 1991. The fate of Hepatozoon species naturally infecting Florida black racers and watersnakes in potential mosquito and soft tick vectors, and histological evidence of pathogenicity in unnatural host species. Int J Parasitol 21:511–516. Wozniak EJ, Telford SR Jr, DeNardo DF, McLaughlin GL, and Butler JF. 1998. Granulomatous hepatitis associated with Hepatozoon sp. meronts in a southern water snake (Nerodia fasciata pictiventris). J Zoo Wildl Med 29:68–71. Wozniak E, McBride J, DeNardo D, Tarara R, Wong V, and Osburn B. 2000. Isolation and characterization of an antigenically distinct 68-kd protein from nonviral intracytoplasmic inclusions in boa constrictors chronically infected with the inclusion body disease virus (IBDV: Retroviridae). Vet Pathol 37:449–459. Wright RK and Cooper EL. 1981. Temperature effects on ectotherm immune responses. Dev Comp Immunol 5 Suppl 1:117–122. Young LA, Schumacher J, Papich MG, and Jacobson ER. 1997. Disposition of enrofloxacin and its metabolite ciprofloxacin after intramuscular injection in juvenile Burmese pythons (Python molurus bivittatus). J Zoo Wildl Med 28:71–79.

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Figure 3.1   Gopher tortoise, Gopherus polyphemus. Testudinidae. Cardiac puncture. Courtesy of Elliott Jacobson.

Figure 3.2  Loggerhead sea turtle, Caretta caretta. Cervical sinus venipuncture. Courtesy of Elliott Jacobson.

Figure 3.3  Desert tortoise, Gopherus agassizii. Testudinidae. Jugular venipuncture. Courtesy of Elliott Jacobson.

Figure 3.4  American alligator, Alligator mississippiensis. Alligatoridae. Blood collection from the supravertebral vessel. Courtesy of Darryl Heard.

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Figure 3.5  American alligator, Alligator mississippiensis. Alligatoridae. Coccygeal venipuncture. Courtesy of Brian Grossbard.

Figure 3.6  Green iguana, Iguana iguana. Varanidae. Ventral tail vein venipuncture. Courtesy of Elliott Jacobson.

Figure 3.7  Blood python, Python curtus. Pythonidae. Blood collection from the palatine veins. Courtesy of Elliott Jacobson.

Figure 3.8  Rhino viper, Bitis nasicornis. Viperidae. Tail vein venipuncture. Courtesy of Elliott Jacobson.

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Figure 3.9  Boa constrictor, Boa constrictor. Boidae. Cardiac puncture. Courtesy of Elliott Jacobson.

Figure 3.10  Preparation of two coverslips. For a detailed description, see Section 3.1.2.

Figure 3.11  Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of drying artifact in red blood cells, visible as refractile, shiny, and pleomorphic inclusions on surfaces of erythrocytes. WrightGiemsa stain.

Figure 3.12  Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of leukocytes (white arrowheads) and erythrocytes, as seen on the hemocytometer during quantitative analysis. Natt-Herrick’s stain.

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Figure 3.13  Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of leukocytes (arrowheads) and thrombocyte clumps with elliptically shaped, weakly violetstaining thrombocytes (arrow), as seen on the hemocytometer during quantitative analysis. Natt-Herrick’s stain.

Figure 3.14  Chaco tortoise, Geochelone chilensis. Testudinidae. Photomicrograph of thrombocyte clumps in a heparinized sample. Wright-Giemsa stain.

Figure 3.15  Tegu lizard, Tupinambis teguixin. Teiidae. Photomicrograph of mature erythrocytes. Wright-Giemsa stain.

Figure 3.16  Cottonmouth, Agkistrodon piscivorous. Viperidae. Figures 3.16 through 3.18 show photomicrographs of different stages of developing polychromatophils in a patient with marked regenerative anemia (PCV 9%). A single pyknotic erythrocyte is noted here (arrow). Wright-Giemsa stain.

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Figure 3.17  Cottonmouth, Agkistrodon piscivorous. Viperidae. Photomicrograph of stages of developing polychromatophils. Two thrombocytes are seen in bottom margin. Wright-Giemsa stain.

Figure 3.18  Cottonmouth, Agkistrodon piscivorous. Viperidae. Photomicrograph of a rubricyte with nucleolus (arrowhead), a lymphocyte (arrow), and two thrombocytes. Wright-Giemsa stain.

Figure 3.19  Pancake tortoise, Malacocherus tornieri. Testudinidae. Photomicrograph of mitosis in a polychromatophil next to a thrombocyte. The tortoise was severely anemic (PCV 6%). Wright-Giemsa stain.

Figure 3.20  Pancake tortoise, Malacocherus tornieri. Testudinidae. Photomicrograph of a recently divided polychromatophil and polychromasia. Wright-Giemsa stain.

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Figure 3.21  Yellow-footed tortoise, Geochelone carbonaria. Testudinidae. Photomicrograph of two erythrocytes with pyknotic nuclei (arrowheads), one polychromatophil (arrow) and two thrombocytes. Wright-Giemsa stain.

Figure 3.22  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of an erythroplastid (arrowhead) and a polychromatophil (arrow). Wright-Giemsa stain.

Figure 3.23  Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of mature erythrocytes with small, punctate basophilic inclusions, and a single thrombocyte in a healthy tortoise. Wright-Giemsa stain.

Figure 3.24  Ball python, Python regius. Pythonidae. Photomicrograph of small, punctate basophilic inclusions and variably sized, clear vacuoles in erythrocytes and two thrombocytes in a healthy snake. Wright-Giemsa stain.

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Figure 3.25  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of small punctate basophilic (arrows) and clear (arrow heads) intraerythrocytic inclusions in a healthy alligator. Two heterophils and a small lymphocyte also are seen. Wright-Giemsa stain.

Figure 3.26  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of rectangular and hexagonal, clear erythrocyte inclusions. Wright-Giemsa stain.

Figure 3.27  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of rectangular and hexagonal, clear erythrocyte inclusions. Wright-Giemsa stain.

Figure 3.28  Pancake tortoise, Malacocherus tornieri. Testudinidae. Photomicrograph of hemodilution with lymph during brachial venipuncture. Small well-differentiated lymphocytes predominate; a single intermediate lymphocyte, a single heterophil, and erythrocytes with drying artifact visible as shiny refractile and pleomorphic inclsions on surfaces of erythrocytes. Wright-Giemsa stain.

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Figure 3.29  Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a heterophil (arrowhead; spindleshaped granules) and a single eosinophil (spherical granules). Wright-Giemsa stain.

Figure 3.30  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of a heterophil, a small well-differentiated lymphocyte (arrowhead) and a medium-sized lymphocyte (arrow). Wright-Giemsa stain.

Figure 3.31  Ball python, Python regius. Pythonidae. Photomicrograph of a heterophil with dense fusiform eosinophilic granules and refractile drying artifact. Wright-Giemsa stain.

Figure 3.32  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of two bilobed heterophils. Wright-Giemsa stain.

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Figure 3.33  Chinese dragon, Physignathus cocincinus. Agamidae. Photomicrograph of three moderately toxic heterophils with degranulation and cytoplasmic basophilia. Wright-Giemsa stain.

Figure 3.34  Spectacled caiman, Caiman crocodilus. Alligatoridae. Photomicrograph of a single severely toxic, immature heterophil with degranulation, abnormal granulation, cytoplasmic basophilia, and cytoplasmic vacuolation; one thrombocyte (upper left margin). Wright-Giemsa stain.

Figure 3.35  Fischer’s chameleon, Chameleo fischeri. Chamaeleonidae. Photomicrograph of three severely toxic, left-shifted heterophils with cytoplasmic basophilia, degranulation, abnormal granulation, and excessive nuclear lobation; mature erythrocytes and three thrombocytes. Wright-Giemsa stain.

Figure 3.36  Spur-thigh tortoise, Testudo graeco ibera. Testudinidae. Photomicrograph of two severely toxic heterophils (arrowheads) with cytoplasmic basophilia, degranulation, vacuolation, abnormal granulation, and pleomorphic nuclei; one plasmacytoid, reactive lymphocyte (arrow), erythrocytes with small basophilic and clear inclusions. Wright-Giemsa stain.

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Figure 3.37  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of two severely toxic heterophils with cytoplasmic basophilia, vacuolation, degranulation, and abnormal granulation; erythrocytes with drying artifact. WrightGiemsa stain.

Figure 3.38  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of two degranulated heterophils (arrowheads), degranulated basophil (arrow) and a small lymphocyte. Wright-Giemsa stain.

Figure 3.39  Spur thigh tortoise, Testudo graeco ibera. Testudinidae. Photomicrograph of a single severely toxic, bilobed heterophil (arrowhead), a lymphocyte; erythrocytes with clear, punctate inclusions consistent with degenerate organelles. Wright-Giemsa stain.

Figure 3.40  Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a nontoxic immature heterophil with band-shaped nucleus and a few primary, purple-staining granules. Wright-Giemsa stain.

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Figure 3.41  Spur-thigh tortoise, Testudo graeco ibera. Testudinidae. Photomicrograph of a severely toxic, immature heterophil. Wright-Giemsa stain.

Figure 3.42  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of a severely toxic, degranulated heterophil with intracytoplasmic, rodshaped bacterium. Wright-Giemsa stain.

Figure 3.43  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of a degranulated heterophil with degranulated pink cytoplasm (arrowhead) and an eosinophil (arrow). Wright-Giemsa stain.

Figure 3.44  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of an eosinophil with hemogregarine gamont. Wright-Giemsa stain.

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Figure 3.45  American crocodile, Crocodylus acutus. Crocodylidae. Photomicrograph of an eosinophil (arrowhead) and a toxic heterophil (arrow). WrightGiemsa stain.

Figure 3.46  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a green eosinophil (arrowhead) and basophil (arrow). Wright-Giemsa stain.

Figure 3.47  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a green eosinophil, a lymphocyte, and an immature azurophilic monocyte. WrightGiemsa stain.

Figure 3.48  Flowerback box turtle, Cuora galbinifrons. Emydidae. Photomicrograph of an eosinophil (arrow) and a larger, left-shifted eosinophil with primary basophilic and secondary eosinophilic granules (arrowhead). Wright-Giemsa stain.

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Figure 3.49  Blood python, Python brongersmai. Pythonidae. Photomicrograph of a heterophil, a basophil (arrowhead), an azurophil (arrow), and thrombocytes. WrightGiemsa stain.

Figure 3.50  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a basophil. Wright-Giemsa stain.

Figure 3.51  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of a degranulated basophil (black arrowhead), a heterophil (black arrow), a lymphocyte (grey arrowhead), and a thrombocyte (grey arrow). WrightGiemsa stain.

Figure 3.52  Plateau spiny lizard, Sceloporus clarkii vallaris. Iguanidae. Photomicrograph of a large reactive lymphocyte (arrowhead), two small lymphocytes (arrow), two erythrocytes, and one polychromatophil. Wright-Giemsa stain.

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Figure 3.53  Monitor lizard, Varanus indicus. Varanidae. Photomicrograph of a small lymphocyte and an intermediate, granular lymphocyte (arrowhead). Wright-Giemsa stain.

Figure 3.54  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of a lymphocyte (arrowhead) and a thrombocyte. Wright-Giemsa stain.

Figure 3.55  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a lymphoblast with nucleolus (arrowhead) and a bilobed heterophil. WrightGiemsa stain.

Figure 3.56  African spur-thigh tortoise, Geochelone sulcata. Testudinidae. Photomicrograph of a large reactive lymphocyte with moderately basophilic cytoplasm and small, discrete vacuoles; erythrocytes with drying artifact as well as clear, punctate and small basophilic inclusions (degenerate organelles). Wright-Giemsa stain.

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Figure 3.57  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a plasmacytoid lymphocyte (arrowhead), a bilobed heterophil, and a thrombocyte. Wright-Giemsa stain.

Figure 3.58  Bearded dragon, Pogona vitticeps. Agamidae. Photomicrograph of a large, reactive, plasmacytoid lymphocyte; three thrombocytes and polychromasia. Wright-Giemsa stain.

Figure 3.59  Blood python, Python curtus. Pythonidae. Photomicrograph of a reactive lymphocyte with abundant pale basophilic cytoplasm that contains dust-like pink granules (center), a lymphocyte (left margin), a thrombocyte (arrowhead), and a rubricyte (arrow). WrightGiemsa stain.

Figure 3.60  African spur-thighed tortoise, Geochelone sulcata. Testudinidae. Photomicrograph of a reactive, granular lymphocyte. Wright-Giemsa stain.

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Figure 3.61  Ball python, Python regius. Pythonidae. Photomicrograph of a lymphoblast with nucleolus (arrowhead), a small lymphocyte (arrow) and a thrombocyte; erythrocytes with small basophilic and clear, punctate inclusions, consistent with degenerate organelles. WrightGiemsa stain.

Figure 3.62  Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph of a plasma cell or plasmacytoid lymphocyte. Wright-Giemsa stain.

Figure 3.63  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of a monocyte. Wright-Giemsa stain.

Figure 3.64  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of a monocyte. Wright-Giemsa stain.

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Figure 3.65  African spurred tortoise, Geochelone sulcata. Testudinidae. Photomicrograph of two reactive monocytes; erythrocytes with variably sized discrete, punctate vacuoles. Wright-Giemsa stain.

Figure 3.66  Aldabra tortoise, Dipsochelys dussumieri. Testudinidae. Photomicrograph of two reactive monocytes (arrowheads) and five large reactive lymphocytes. Wright-Giemsa stain.

Figure 3.67  Monitor lizard, Varanus indicus. Varanidae. Photomicrograph of heterophil, basophil, and monocytoid leukocyte; lymphocyte at top left and thrombocyte at bottom left margin. Wright-Giemsa stain. Courtesy of Dr. Heather Wamsley.

Figure 3.68  Red tegu lizard, Tupinambis rufescens. Teiidae. Photomicrograph of four monocytoid leukocytes, two reactive azurophils, two heterophils, and two lysed cells. Wright-Giemsa stain.

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Figure 3.69  Flap-necked chameleon, Chamaeleo dilepis. Chamaeleonidae. Photomicrograph of a monocyte (nucleus at position 9) with Chlamydia inclusion (at position 3) and poxvirus inclusion (at position 6). Wright-Giemsa-stain.

Figure 3.70  Indigo snake, Drymarchon corais couperi. Colubridae. Photomicrograph of an azurophil (arrowhead) and a heterophil with coalescing granules; thrombocyte. Wright-Giemsa stain.

Figure 3.71  Rainbow boa, Epicrates cenchria cenchria. Boidae. Photomicrograph of three reactive azurophils, a heterophil with fused granules that appear as heterogenous eosinophilic material, and a small lymphocyte in the upper right margin. WrightGiemsa stain.

Figure 3.72  Mangrove monitor lizard, Varanus indicus. Varanidae. Photomicrograph of two immature azurophils. Wright-Giemsa stain.

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Figure 3.73  Yellow-footed tortoise, Geochelone denticulata. Testudinidae. Photomicrograph of three lymphoblasts; the tortoise was diagnosed with lymphoma in the skeletal muscle of the ventral cervical region based on histopathology. WrightGiemsa stain.

Figure 3.74  Green iguana, Iguana iguana. Iguanidae. Photomicrograph of three thrombocytes. Wright-Giemsa stain.

Figure 3.75  Aldabra tortoise, Geochelone gigantia. Testudinidae. Photomicrograph of three thrombocytes with a few, dustlike, azurophilic cytoplasmic granules. Wright-Giemsa stain.

Figure 3.76  Tegu lizard, Tupinambis teguixin. Teiidae. Photomicrograph of two ruptured thrombocytes (arrowheads), a basophil (arrow), and a heterophil. WrightGiemsa stain.

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Figure 3.77  Monitor lizard, Varanus indicus. Varanidae. Photomicrograph of a single lymphocyte (arrowhead) in a thrombocyte clump. WrightGiemsa stain.

Figure 3.78  Rainbow boa, Epicrates cenchria crassus. Boidae. Photomicrograph of thrombocytes with pseudopods. Wright-Giemsa stain.

Figure 3.79  African spurthighed tortoise, Geochelone sulcata. Testudinidae. Photomicrograph of a vacuolated thrombocyte with few fine azurophilic granules. Wright-Giemsa stain.

Figure 3.80  Cottonmouth, Agkistrodon piscivorous. Viperidae. Photomicrograph of a phagocytic thrombocyte with intracytoplasmic material, which stained positive for hemosiderin (arrowhead), three normal thrombocytes, and one polychromatophil (arrow). Wright-Giemsa stain.

210 Circulating Inflammatory Cells

Figure 3.81  Blood python, Python brongersmai. Pythonidae. Photomicrograph of binucleated thrombocyte (arrowhead), two thrombocytes with pseudopods, and one heterophil. WrightGiemsa stain.

Figure 3.82  Eastern indigo snake, Drymarchon corais couperi. Colubridae. Photomicrograph of erythrocytes with gametocytes of Hepatozoon sp. Wright-Giemsa stain.

Figure 3.83  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of Hepatozoon fusifex. Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 3.84  Agama lizard, Agama agama. Agamidae. Photomicrograph of Plasmodium giganteum, female gametocyte in erythrocyte. Giemsa stain. Courtesy of Sam R. Telford Jr.

Circulating Inflammatory Cells  211

Figure 3.85  Common tegu lizard, Tupinambis teguixin. Teiidae. Photomicrograph of Saurocytozoon tupinambi; female gametocyte. Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 3.86  Crevice spiny lizard, Sceloporus poinsettii. Iguanidae. Photomicrograph of Trypanosoma poinsettii; Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 3.87  Japanese grass lizard, Takydromus tachydromoides. Lacertidae. Photomicrograph of Sauroplasma (Piroplasmida). Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 3.88  Tree gecko, Hemidactylus platycephalus, Gekkonidae. Photomicrograph of Sauroleishmania. Giemsa stain. Courtesy of Sam R. Telford Jr.

212 Circulating Inflammatory Cells

Figure 3.89  Gila monster, Heloderma suspectum. Helodermatidae. Photomicrograph of a filarid nematode in a peripheral blood film. Giemsa stain. Courtesy of James Jarchow and Carlos Reggiardo.

Figure 3.90  Monitor lizard, Varanus indicus. Varanidae. Photomicrograph of a filarid nematode in the feathered edge of a peripheral blood film. Wright-Giemsa stain.

Figure 3.91  Common boa constrictor, Boa constrictor imperator. Boidae. Photomicrograph of a lymphocyte with large homogenously basophilic IBD inclusion that pushes the nucleus aside; two thrombocytes. Wright-Giemsa stain.

Figure 3.92  Rainbow boa, Epicrates cenchria cenchria. Boidae. Photomicrograph of two lymphocytes with distinct, homogenously basophilic IBD inclusions indenting the lymphocyte nucleus. Wright-Giemsa stain.

Circulating Inflammatory Cells  213

Figure 3.93  Common boa constrictor, Boa constrictor imperator. Boidae. Photomicrograph of a lymphocyte with IBD inclusion (arrowhead), three normal lymphocytes (arrows), three thrombocytes, and two azurophils. Wright-Giemsa stain.

Figure 3.94  Common boa constrictor, Boa constrictor imperator. Boidae. Photomicrograph of a lymphocyte with crescent-shaped nucleus due to compression by an IBD inclusion, two normal lymphocytes (arrows), and three thrombocytes. WrightGiemsa stain.

Figure 3.95  Common boa constrictor, Boa constrictor imperator. Boidae. Photomicrograph of a basophil with an IBD inclusion between the nucleus and basophil granules. Polychromatophil and three thrombocytes. WrightGiemsa stain.

Figure 3.96  Jamaican boa, Epicrates subflavus. Boidae. Photomicrograph of a thrombocyte with intracytoplasmic IBD inclusion. Wright-Giemsa stain.

214 Circulating Inflammatory Cells

Figure 3.97  Japanese grass lizard, Takydromus tachydromoides. Lacertidae. Photomicrograph of LEV inclusions in erythrocytes. Giemsa stain. Courtesy of Dr. Sam R. Telford Jr.

Figure 3.98  Terciopelo, Bothrops asper. Viperidae. Photomicrograph of a polychromatophil with two hexagonal, crystalline inclusions and granular eosinophilic snake erythrocyte virus (SEV) inclusions (arrowhead), an immature erythroid precursor (arrow), an erythrocyte, and a partially lysed lymphocyte. Wright-Giemsa stain.

Figure 3.99  Terciopelo, Bothrops asper. Viperidae. Photomicrograph of an erythrocyte precursor with snake erythrocyte virus (SEV) inclusions (arrowheads), two lymphocytes (arrows), thrombocytes, and polychromatophils. WrightGiemsa stain.

Figure 3.100  Terciopelo, Bothrops asper. Viperidae. Photomicrograph of an erythrocyte precursor with snake erythrocyte virus (SEV) inclusion, a rubricyte (arrow), a mitotic figure, a thrombocyte (arrowhead), and a lymphocyte. Wright-Giemsa stain.

Circulating Inflammatory Cells  215

Figure 3.101  Terciopelo, Bothrops asper. Viperidae. Photomicrograph of two erythrocyte precursors with hexagonal, crystalline inclusions and granular eosinophilic viral particles (arrowhead), a rubricyte next to two mature erythrocytes, and a single azurophil (left margin). Wright-Giemsa stain.

Figure 3.102  Terciopelo, Bothrops asper. Viperidae. Photomicrograph of two erythrocyte precursors with hexagonal, crystalline inclusions and granular eosinophilic inclusion of snake erythrocyte virus (SEV) (arrowheads). Wright-Giemsa stain.

Figure 3.103  Eastern box turtle, Terrapene carolina carolina. Emydidae. Photomicrograph of pink, granular, round to oval iridoviral inclusions in leukocytes (frog virus 3, FV3, family Iridoviridae, genus Ranavirus). Wright stain. Courtesy of Michael M. Fry.

Figure 3.104  Eastern box turtle, Terrapene carolina carolina. Emydidae. Photomicrograph of pink, granular, round to oval iridoviral inclusions in leukocytes (frog virus 3, FV3, family Iridoviridae, genus Ranavirus). Wright stain. Courtesy of Michael M. Fry.

216 Circulating Inflammatory Cells

Figure 3.105  Eastern box turtle, Terrapene carolina carolina. Emydidae. Photomicrograph of pink, granular, round to oval iridoviral inclusions in leukocytes (frog virus 3, FV3, family Iridoviridae, genus Ranavirus). Wright stain. Courtesy of Michael M. Fry.

Figure 3.106  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of a monocyte with phagocytized material, suggestive of lipid. Wright-Giemsa stain.

Figure 3.107  Eastern indigo snake, Drymarchon corais couperi. Colubridae. Photomicrograph of a monocyte with melanin granules and a heterophil. Wright-Giemsa stain.

Figure 3.108  Aldabra tortoise, Dipsochelys dussumieri. Testudinidae. Photomicrograph of a monocyte with melanin granules. Wright-Giemsa stain.

Circulating Inflammatory Cells  217

Figure 3.109  Common boa constrictor, Boa constrictor imperator. Boidae. Photomicrograph of a monocyte with phagocytized material and erythrocytes with small, basophilic inclusions. Wright-Giemsa stain.

Figure 3.110  Fischer’s chameleon. Chamaeleo fischeri. Boidae. Photomicrograph of a monocyte with phagocytized erythrocyte (erythrophagia); erythrocytes with drying artifact. Wright-Giemsa stain.

Figure 3.111  Common boa constrictor, Boa constrictor imperator. Boidae. Photomicrograph of two extracellular small rod-shaped bacteria (arrowhead); blood culture revealed significant growth of Morganella morganii and Clostridium sp.; three thrombocytes and polychromasia, next to a mitotic figure (arrow). WrightGiemsa stain.

218 Circulating Inflammatory Cells

Figure 3.112  Fischer’s chameleon, Chameleo fischeri. Boidae. Photomicrograph of a presumably degranulated heterophil with phagocytized rodshaped bacteria. Wright-Giemsa.

Figure 3.113  Rhinoceros iguana, Cyclura cornuta. Iguanidae. Photomicrograph of spiralshaped bacteria phagocytized in a monocyte (M) and free in the peripheral blood. Wright-Giemsa stain.

Figure 3.114  Emerald tree boa, Corallus caninus. Boidae. Photomicrograph of Chlamydophila pneumoniae inclusion in a circulating monocyte (arrowhead). Infection was confirmed by PCR. Wright-Giemsa stain.

4 Reptile Necropsy Techniques Scott P. Terrell and Brian A. Stacy

Contents

4.1 Introduction

4.1 Introduction............................................................219 4.1.1 Why Do a Necropsy?..................................219 4.1.2 Objectives of a Necropsy.......................... 220 4.2 Necropsy Basics..................................................... 220 4.2.1 Costs Associated with a Necropsy........... 220 4.2.2 Carcass Preservation, Shipping and   Disposal..................................................... 221 4.3 Equipment ............................................................ 222 4.4 Components of a Necropsy.................................. 223 4.4.1 Data Gathering ......................................... 223 4.4.2 Documentation and Description.............. 223 4.4.3 External Examination .............................. 224 4.4.4 Dissection and Internal Examination....... 224 4.5 Sample Storage and Submission........................... 228 4.5.1 Cytology..................................................... 228 4.5.2 Light Microscopy....................................... 229 4.5.3 Transmission Electron Microscopy........... 231 4.5.4 Toxicology................................................. 231 4.5.5 Microbiology.............................................. 231 4.5.6 Molecular Diagnostic Tests....................... 231 4.5.7 Handling Tissues for Diagnosis of   Specific Pathogens..................................... 231 4.6 Necropsy Precautions and Zoonotic   Disease Concerns.................................................. 233 4.7 After the Necropsy................................................ 233 4.7.1 Cleanup Considerations............................ 233 4.7.2 Electronic Storage, Archiving, and   Retrieval of Reports................................... 233 4.7.3 Tissue Archives.......................................... 233 4.8 Conclusion............................................................. 235 References......................................................................... 235 Appendix 4.1.................................................................... 236

Whereas the postmortem examination of a human is referred to as an autopsy, the postmortem examination of an animal is referred to as a necropsy. The necropsy is an essential component of any quality veterinary medical practice and may be an essential skill for field biologists and other scientists as well. In captive animal populations, the necropsy is an opportunity to learn from the death of an animal so that futures illnesses in other animals may be prevented or more easily diagnosed and treated. In wild animal populations, the necropsy may provide vital data about population declines, habitat changes, effects of human or other animal populations, and causes of catastrophic die-offs. A necropsy may not provide all the answers, but it is an important first step (and sometimes last step) in the investigation of animal and environmental health issues.

4.1.1 Why Do a Necropsy? The reasons for doing a necropsy are as varied as the specimens that may be examined. Basically, necropsy is an opportunity to learn. Necropsy is an opportunity to learn from one’s mistakes, to learn about an interesting or unusual case, to learn about the biology and anatomy of a particular species, to learn the cause of illness or injury in an individual animal or population of animals, and perhaps to learn about the animal’s environment. No matter how you look at it, the death of an animal is unfortunate. What is truly unfortunate, however, is failure to take the opportunity to learn from the death of that animal. That is the value of a necropsy. Probably the most common reason a necropsy is performed is for diagnostic purposes (i.e., to determine the cause of death or illness). Before beginning a diagnostic necropsy, one must understand that necropsy examination is only one part of a complete diagnostic assessment or workup. Compo-

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220  Reptile Necropsy Techniques

nents of a complete diagnostic workup often include collection of medical history, microscopic examination of tissues (via light and electron microscopy), bacterial culture, fungal culture, virus isolation, and a variety of additional ancillary tests. A complete diagnostic workup from beginning to end typically is only performed by trained veterinary pathologists and it is the pathologist that makes the final diagnosis in most cases. A person not specially trained in pathology should not approach a necropsy with the idea that a diagnosis will be made during the postmortem examination. Rather, the prosector (the person performing the necropsy) should concentrate on the primary goals of the necropsy (objective data collection, detailed description of findings, complete sample collection) so that a pathologist or other specialist involved in identifying the specific cause of death or disease can use the data and samples collected. It is very rewarding if the cause of death or disease is immediately apparent at necropsy, but in most cases a confirmed diagnosis will be made only after tissues are further analyzed and examined by trained personnel. Many different diagnostic tests can be used to detect or monitor prior or current infection or clinical illness. The results of any single diagnostic test must be interpreted in the context of the entire clinical picture of the animal, including the history and pattern of disease in the population, clinical signs, results of other tests, as well as the postmortem data, including gross and histopathologic findings. Detection or isolation of an infectious agent or detection of antibodies to that agent provides only partial information in an investigation of a morbidity or mortality problem. In some instances, findings may be completely incidental to the actual cause of disease or death.

4.1.2 Objectives of a Necropsy Specific projects or procedures may place emphasis on certain components of the necropsy examination. For example, a necropsy performed for disease diagnosis may emphasize complete tissue collection into formalin, whereas a necropsy performed as a part of an environmental contaminant study may concentrate effort on the collection of specific tissues for toxicologic analysis. Regardless of the project design or overall objectives, the most important goals when performing the actual necropsy include: (1) accurate and objective data collection, (2) detailed description of findings, and (3) complete sample collection. Each of these goals will be discussed in detail later in this chapter. Data collection, including clinical or field observations, is a vital component of the procedure. Accurate and detailed description of findings during the necropsy is essential. Finally, a complete necropsy should include collection and archiving of fixed and frozen samples from all tissues so that necessary materials are available for immediate use, as well as important retrospective studies and research (e.g., toxicologic studies, nutrient analysis, virus isolation, transmission studies, immunodiagnostics,

genetic studies, and molecular diagnostic tests). The prosector can accomplish these goals by using a systematic approach to the necropsy procedure in every case. Practice and repetition are essential skills to develop as a prosector. Dissection technique and knowledge of anatomy are learned skills that will increase with every specimen examined.

4.2 Necropsy Basics A properly performed necropsy can yield vast amounts of diagnostic and biological data, whereas a poorly performed necropsy can result in frustration, lost data, misinterpretation of findings, or a missed diagnosis. In most cases, necropsy technique is easily learned and the procedure can be performed with a limited amount of equipment. The quality of the necropsy will depend upon the background and training of the person doing the examination. Ideally, the person should have some experience with dissection and knowledge of anatomy. Necropsies should be done in a cool indoor facility whenever possible so that samples can be properly collected for microbial isolation; however, in many cases, necropsies are performed in the field. Field conditions are less than ideal for some aspects of the necropsy, but valuable data can still be collected.

4.2.1 Costs Associated with a Necropsy Costs associated with necropsy vary depending on whether a veterinarian, pathologist, or biologist performs the examination, and whether whole animals or tissues are sent to a diagnostic laboratory. This section presents the basic costs involved with performing necropsies and fees are current at the time of this writing. Undoubtedly, costs will change over time, thus receiving laboratories should be contacted for current fees. In the United States and Canada, most states and provinces have a veterinary diagnostic laboratory that will accept specimens for necropsy. Similarly, most veterinary schools will accept submissions. Several private veterinary pathology services also exist, some with expertise in reptiles and other exotic species. In the authors’ experience, state diagnostic laboratories tend to be less expensive than veterinary college diagnostic services. A summary of basic laboratory fees for 4 representative state diagnostic labs is provided in Table 4.1. State laboratories may charge $25 to $90 per specimen for necropsy and histopathologic evaluation with additional costs depending on needs for toxicology, microbiology, and so on. The same service at a veterinary college or private laboratory may cost $150 to $300 or more. Local laboratories or veterinary colleges should be consulted prior to submission of specimens to discuss specific expectations and costs. Although cost is a consideration, experience with reptiles should be the primary criterion for selecting a diagnostic laboratory.

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Table 4.1   Fees for diagnostic and pathology services at regional diagnostic labs (some fees only for valid in-state submissions). These labs were chosen as examples; most states have a diagnostic lab to which samples may be submitted. Prices are listed in U.S. dollars. Illinois State Veterinary Diagnostic Laba

University of Georgia College of Veterinary Medicine, Athens Labb

Colorado State University Veterinary Diagnostic Laboratoryc

Florida Animal Health Diagnostic Labd

Necropsy plus histology

$90

$35

$60

$25

Surgical biopsy or individual tissue histologic evaluation

37

25

30

30

Bacterial culture

18

8

15

10

Toxicology (lead evaluation used as example test)

17

15

10

30

Parasitology

9

7

15

5

Source: http://www.cvm.uiuc.edu/vdl/FEE_SCHEDULE_2003.html December 2004. Source: http://hospital.vet.uga.edu/dlab/fees.php May 2006. c Source: http://www.dlab.colostate.edu/security2/test_price.cfm May 2006. d Source: Personal communication, Florida Animal Health Diagnostic Lab, April 2006. a b

It is not always possible or practical to send a complete carcass to a diagnostic laboratory for necropsy. In many cases, the prosector performs the necropsy and sends tissues to laboratories for histopathologic evaluation, microbiology, toxicology, and so on. Histopathology is a fundamental component of necropsies and disease investigation, and is responsible for the bulk of the cost associated with tissue submission. Processing fees for histopathology vary among laboratories, thus a set of 8 tissues (typical minimum) will cost anywhere from $65 to $80 or more. In the authors’ experience, toxicologic testing may cost $10 to $35 or more, depending on the type of compound. Bacterial cultures typically cost approximately $8 to $20 per sample, and additional fees are incurred for special techniques such as anaerobic culture and antimicrobial sensitivity determination. Parasite identification costs range from $5 to $15. Electron microscopy costs vary widely depending on the laboratory.

4.2.2 Carcass Preservation, Shipping and Disposal A basic rule of necropsy is that “fresh is best.” Ideally, the necropsy should be performed as soon as possible after death. Decomposition or autolysis is a common problem in reptiles because many species are housed in heated enclosures or aquaria, which accelerates decomposition. If the necropsy is delayed, carcasses should be cooled on ice or refrigerated. Smaller specimens (less than 10.0 kg) are better preserved by refrigeration than larger animals. Under optimal refrigeration conditions, adequate preservation for histopathological examination of most major organs is retained for around 72 hours if the initial postmortem condition of the carcass is good.

Freezing should be avoided and is used only as a last resort to prevent severe decomposition. Freezing renders the carcass useless for many diagnostic purposes, as histologic architecture is distorted or destroyed and many bacterial and fungal pathogens do not survive the freeze and thaw process. If necropsy is not performed immediately after death, the overall appearance of the specimen will dictate whether or not a complete necropsy (i.e., including histopathology) will be worthwhile. If decomposition is advanced, as indicated by bloating, skin discoloration, and sloughing of skin, scales, or shell scutes, collection of tissues for histopathologic evaluation will often be unrewarding. However, even rotted tissues may be valuable for conducting toxicologic, molecular, or genetic studies, as well as detecting major grossly apparent abnormalities. Procedures for shipment of carcasses or samples depend on the destination and ultimate purpose(s) of the specimens. It is important to contact the laboratory prior to shipment to discuss proper and specific recommendations for carcass or sample handling and shipment. General considerations for shipping biological materials include: (1) prevention of leakage of package contents, (2) prevention of decomposition of samples, (3) proper labeling of submissions, and (4) inclusion of data sheets or appropriate submission forms. All receptacles containing formalin or other liquids should be sealed with tape or a paraffin product (such as Parafilm®) and then placed in a second container such as a screw top container or sealed plastic bag. The best containers for shipping tissues in formalin are screw-top containers; many of these are available and are specifically designated for this purpose. Avoid using glass jars as shipping containers due to the possibility of breakage. Flip-top formalin containers

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have a tendency to leak and should always be double sealed. Recycled plastic drug bottles and glass baby food containers are commonly used for formalin tissue storage, but these containers do not seal well and some sort of secondary containment is absolutely necessary. A useful alternative to shipping containers of formalin is to ship previously fixed tissues in formalin-soaked paper towels or gauze. Formalin is poured off of the tissues after 24 to 48 hours of fixation, and tissues are then wrapped in formalin-soaked paper or gauze and shipped in double-sealed plastic bags. This technique works very well and reduces the chance of formalin spillage. All of the appropriate paperwork should be packaged with the samples and all containers should be clearly labeled with proper identification, especially if samples are from multiple individuals. Paperwork submitted with formalin tissues should be sealed in a separate plastic bag because leaked formalin or other fluids will smear ink and render data illegible. Decomposition of fresh specimens or thawing of frozen specimens during shipment is an important consideration. Commercially available cold packs should be used for carcasses or samples that must remain cold but not frozen during shipment. Ice (wet ice) should be avoided as it will melt and tends to leak, regardless of how well the package is sealed. Cold packs should be placed along the inside of Styrofoamlined boxes or coolers and below the carcass or sample. To prevent freezing, do not place cold packs directly on top of a sample and do not use dry ice. Dry ice, however, is ideal for samples that should remain frozen during shipment. Alternatively, multiple cold packs can be used and should be placed on top of a sample to maintain the lowest possible temperature. After completion of the necropsy, the carcass should be disposed of in a manner compliant with federal, state, and local agencies. Regulations regarding carcass disposal vary widely among states and even within states depending on location and county, thus the appropriate authority should be consulted. Burial may or may not be an option in some areas. Some states and counties allow carcasses to be dumped at local landfills. Another option is a commercial disposal service that disposes of carcasses for a fee. Also, most diagnostic laboratories provide incineration or disposal services as part of the necropsy examination.

4.3 Equipment Equipment needs vary depending on the species, size of the animal, and the specific goals of the necropsy. A basic equipment and tool list is provided in Table 4.2. The most rudimentary necropsy of snakes, lizards, crocodilians, and even sea turtles can be performed with a knife, scissors, formalin container, and freezer bags (Figures 4.1–4.2). A pair of garden shears or hedge clippers may be valuable for cutting ribs and other bones in larger reptiles. Removal of the brain from large specimens and removal of the plastron of

Table 4.2   Equipment list for necropsy of reptiles. Necropsy form Coveralls or other appropriate clothing Rubber boots or shoe covers  Rubber or latex gloves Surgical mask  Camera (digital is preferred) String, labels, assorted bottles, waterproof pen Container for formalin and appropriate volume of 10% neutral buffered formalin Forceps — several sizes Tissue cutting board Necropsy knives and sharpener Scalpel blades (#20 and #10) and handle Scissors — several sizes Postmortem shears (garden shears, hedge cutter) Alcohol lamp or butane burner  Matches or lighter 70% alcohol Fixative for electron microscopy such as Trumps solution (should be kept chilled) Sterile whirl-pack bags (zip-type bags can suffice) Cryotubes Microbial culturette swabs Microbial transport media Dry ice and ice chest or cooler Scale Stryker saw or Dremel tool Calipers Rulers Microscope slides

tortoises and turtles may require a cutting instrument, such as a Dremel® tool (Racine, WI) or Stryker ® saw (Kalamazoo, MI) (Figure 4.3). Chelonian biologists often use a caliper to measure the dimensions of a chelonian’s shell. All equipment should be thoroughly cleaned and disinfected following each necropsy (see Section 4.7.1). Ideally, protective clothing and equipment should be worn while performing a necropsy. These items include some sort of washable outer clothing or apron and proper rubber boots or protective shoe covering. Rubber or latex gloves are essential and a facemask is recommended, especially if power tools are used. In some situations (i.e., field procedures), protective clothing may not be available or practical, but the use of protective clothing and equipment should be stressed in any procedure involving the potential for infectious disease transmission or the illness or death of multiple animals in an outbreak situation.

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4.4 Components of a Necropsy The necropsy can be divided into several components to make the process systematic and simple to perform. These components include: (1) data gathering, (2) documentation and description, (3) external examination, (4) dissection and internal examination, and (5) sample storage and submission.

4.4.1 Data Gathering Data collected prior to the necropsy includes a thorough medical history, field observations, and vital data such as weights and measurements. The importance of the medical history and field observations cannot be overemphasized. The prosector or pathologist often has no idea what has occurred in the animal’s past, in the population, or in the field environment. These historical events and information may greatly influence the choice of diagnostic tests (i.e., specific toxicant testing), additional tissues to be collected, or conclusions at the completion of the necropsy.

4.4.1.1 Medical History  The medical history is one of the most important pieces of the puzzle in regards to the postmortem examination. Any and all details that pertain to a particular animal or group of animals may provide vital clues to the cause of disease or death. The information that is generally provided includes:

a. Details of recent illnesses b. Time course of disease c. Treatments administered prior to death d. Behavioral changes or clinical signs prior to death e. Dietary changes or changes in food availability e. Details of illnesses in other animals f. Details of disease in similar species g. Evidence of deaths in multiple animals h. Possibility of traumatic injury or accidental or intentional poisoning i. Clinicopathologic data (i.e., blood values, urinalysis, etc.) j. Ancillary diagnostic test data (i.e., radiographs) In addition to these objective data, the authors also request that the submitting party (veterinarians, field biologists, private individuals) provide a subjective evaluation of the animal or mortality event. Theories provided by veterinarians, biologists, or by the general public, although unusual in some cases, are often based on reality and may provide a unique perspective on an animal death or mortality event.

4.4.1.2 Field Observations  During a wildlife mortality event, careful and complete description of the health problem by the field biologist is a critical step in arriving at a diagnosis. Basic biological, environmental, habitat, and weather data may all provide clues to the cause of the mortality event. Biologi-

cal information includes specific species identification, age, sex and breeding activity, as well as the size and weights of affected animals. Environmental and habitat data may include habitat types, vegetation characteristics, water level changes, food sources, presence of other species or domesticated animals, and human presence in the area. Weather information such as temperature, rainfall, storm data, lightning strikes, and unusual weather events can be important, especially in areas where severe weather may contribute to mortality.

4.4.1.3 Biological Measurements  Body weight is an important measurement for any medical procedure and should be recorded in every case. Standard biological length measurements are also recorded. In most reptiles, the total length (measured from tip of snout to tip of tail) and snoutvent length (measured from tip of snout to cloaca) are the most common values recorded (Figure 4.4). Shell measurements are collected in chelonians and include straight carapace length, curved carapace length, straight width, curved width, and height. Tree calipers (used for measuring tree widths) are used to measure medium to large-sized chelonians, such as sea turtles. More detailed measurements may be taken when performing specific biological studies or when examining specific species. One measure collected by some researchers is the body condition index, which was conceived as an objective method of assessing nutritional condition. The body condition index is calculated as a ratio of body weight to estimated body volume. Methods for calculating the body condition index have been described for some chelonian species, such as the desert tortoise (Gopherus agassizii) (Nagy et al., 2002). There are two commonly used methods for estimating body volume and calculating the body condition index in chelonians. The first method calculates the ratio of weight (in grams) to body volume, which is estimated by multiplying carapace length, width, and height (in centimeters) (Nagy et al., 2002). The second method estimates body volume by cubing the midline carapace length (MCL). These ratios may naturally vary by MCL due to confounding changes in body proportions, thus it is possible that comparisons between animals of different sizes will be inaccurate. Therefore, practical use of the body condition index requires comparison with a species database that includes values from healthy animals of different sizes. Such databases exist only for a small number of species and may be valid for only specific size classes. Many prosectors record individual organ weights as part of the routine necropsy procedure, although the authors do not do this routinely. The weight of some organs, such as the liver, will be influenced by physiological conditions such as reproductive activity and alterations associated with season.

4.4.2 Documentation and Description Documentation of necropsy findings is facilitated by using a well-organized necropsy data form. A data form also ensures

224  Reptile Necropsy Techniques

that components of the exam are not forgotten or missed. Necropsy report sheets vary between various institutions and have not been standardized. A sample of a standard necropsy form used by the author for multiple species is available at the end of this chapter (Appendix 4.1). A necropsy form designed specifically for field personnel and veterinarians examining sea turtles is available on the world wide web (http:// www.vetmed.ufl.edu/sacs/wildlife/seaturtletechniques/necropsyreport1.htm). This form was developed in the mid-1990s (Stamper et al., 1997) and has subsequently gone through multiple revisions by various pathologists and biologists. Unfortunately, such detailed necropsy protocols may not be practical for field use, especially when assistance is limited; thus examination and sample collection must be prioritized. Whenever samples are collected, all information that relates to the sample must be thoroughly described and entered into either a field notebook, necropsy data sheet, or other forms developed for this purpose. This information should be as detailed as possible. Excess information can always be eliminated, but information that is forgotten or missed is lost forever. As described in the introduction to this chapter, final determination of cause of death or disease will often require the expertise of a veterinary pathologist or other specialists. Description of lesions seen at necropsy is an important skill. The prosector must serve as the eyes for the pathologist and other specialists not present at the postmortem examination. Lesions should be described by location in or on the body, distribution (focal, multifocal, diffuse), size, color, texture, and even smell. Lesion size should be described as objectively as possible using a ruler rather than references to common items such as marbles, tennis balls, fruits, or other food items. The task of providing a detailed description has been made easier by increased access to digital photography. Lesions should be described and photographed if possible so that images can be sent to the pathologist or other specialists along with documentation and samples from the case. Specimens should be photographed next to a size reference (small ruler or other standard reference). Ideally, the lesion should be photographed in situ (in the body), as well as dissected out of the body on a neutral background (i.e., cutting board).

4.4.3 External Examination The initial step of any necropsy is a thorough external examination, which is similar in approach to clinical examination of a live animal. There are important external indicators of nutrition condition and hydration status that should be evaluated. The eyes often become sunken in severely dehydrated or malnourished animals. Muscle condition is assessed by examining prominence of skeletal structures, especially vertebral processes, which are unapparent or barely palpable in a healthy animal. Malnourished chelonians often feel notably light relative to their size when they are lifted. The girth of the

tail base of lizards and crocodilians is also a good indicator of nutritional status. Important areas that should be examined include the integument, shell, oral cavity, eyes, ears, nares, and cloacal opening. The carcass should be carefully palpated for evidence of trauma, fractures or other anomalies. Any abnormalities, such as swelling around joint spaces, missing digits, and cutaneous or subcutaneous masses are recorded. Some species have common problems or areas that should receive additional attention. For example, the amount and distribution of epibiota coverage on sea turtles can be an indicator of general health and should be documented (Figure 4.5). Line drawings of reptile specimens, including dorsal and ventral views, can be used to document the location of lesions. Scrapings can be collected from skin lesions, such as areas of shell disease, hyperkeratotic skin surfaces, or other sites prior to beginning the internal examination.

4.4.4 Dissection and Internal Examination The class Reptilia includes a vast array of species with great diversity in individual body types. Despite this diversity, the internal anatomy is fairly well conserved across all species with the exception of some key differences. Dissection techniques will be described for each of the major reptile body types. The location of major organs presented in this section can also be found in Chapter 1. Reptiles may exhibit prolonged involuntary movement of skeletal muscle and cardiac contractions following death, which can be unsettling to prosectors and observers. Other animals thought to be dead may in actuality still be alive. Determining when some reptiles are truly dead is not as easy as it is with a mammal and bird. Large tortoises can be especially challenging. Some prosectors choose to decapitate freshly dead or euthanized reptiles to ensure that the animal is definitively dead prior to dissection.

4.4.4.1 Lizards and Crocodilians  Lizards and crocodilians have a similar body shape and may be approached in much the same way despite some differences in anatomy. Precautions should be taken when examining venomous lizard species, which include Gila monsters and beaded lizards (Heloderma spp.) (see Section 4.4.4.2). Dissection of the carcass is begun by placing the animal in dorsal recumbency (lying on its back). An incision is made with a scalpel or scissors along the ventral midline from the cloaca cranially to the intermandibular space (Figure 4.6). The skin is reflected laterally along the length of the incision to expose the underlying subcutis and muscle. Muscle sections can be collected at this time. In lizards, the ceolomic cavity is entered via a midline incision in the muscles caudal to the rib cage. In larger lizards and crocodilians, it may be necessary to incise the skin along the lateral aspects of the ventrum over the costochondral junctions. The rib cage is then removed by cutting the ribs on either side at the costochondral junctions (Figure 4.7). Alternatively, the ventral skin, underlying muscle, and ribs of

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crocodilians may be removed in toto as an entire plate. At this stage, the entire ceolomic cavity is exposed (Figures 4.8, 4.9). Laterally compressed species of lizards, such as chameleons, are best approached by removing one complete side of the coelomic wall and rib cage (Figure 4.10). The fat bodies of lizards can be robust and may cover much of the viscera in healthy or obese animals. Fat bodies are reflected caudally or excised to view the organs (Figure 4.8). After the coelomic cavity is opened, the next step is to free the upper aeropharyngeal tract in preparation for removal of the visceral organs. The skin around the ventral mandible is incised and reflected to expose the tongue and allow examination of the oral cavity, glottis, and pharynx (Figure 4.11). In crocodilians, the gular valve and glottal region are common sites of pathological lesions and should be closely examined. Other important tissues to collect from the cervical or neck region are the thymus, thyroid gland, and parathyroid glands, which are distributed from the caudal mandible to just cranial to the heart. In lizards, the paired thymus is located ventral and medial to the internal carotids and jugular vein. Crocodilians have an elongated thymus that may extend from the base of the heart along the neck (Chiasson, 1962). The lizard and crocodilian thyroid gland may be single, bilobed, or paired depending on the species (Lynn, 1970) (Figures 4.12, 4.13). Lizards may have one or two pairs of parathyroid glands. In green iguanas (Iguana iguana), the caudal pair is located at the origins of the internal and external carotid arteries and the anterior glands lie medial to the rami of the mandible (Figures 4.13, 4.14). Crocodilians may have one or two pairs of parathyroid glands located near the common carotid artery (Clark, 1970). In the event that the parathyroid glands and thymus cannot be identified, a large section of the connective tissue cranial to the heart may be collected, placed in formalin, and later serially sectioned to locate these structures. The next step of the necropsy is examination of the cardiovascular system. The pericardial sac is incised and the heart is removed by cutting the great vessels at the heart base. Examination of the heart is illustrated in the snake necropsy section (see Section 4.4.4.2). Lizards have a three-chambered heart and a right and left aorta (Figure 4.13) (Webb et al., 1971; White, 1968). Crocodilians are the only reptiles to have a complete interventricular septum, and thus a four-chambered heart with two atria, two ventricles, as well as a right and left aorta (Akers, 1996; Webb et al., 1971; White, 1968). In addition, many reptiles, including crocodilian, lizard, and chelonian species have a fibrous gubernaculum cordis, which attaches the apex of the ventricle to the pericardial sac (see Section 4.4.4.3). This structure should not be confused with an adhesion. Once the heart is removed, there are two basic options for removal of the remaining organs. The trachea, esophagus, lungs, and other viscera may be removed intact or the gastrointestinal viscera and liver removed separately by incising the esophagus cranial to the liver, leaving the trachea, esophagus,

and lungs within the carcass. The latter technique may be easier in larger specimens such as crocodilians. The organs are removed by applying gentle traction, lifting caudally, and cutting connective tissue attachments as needed. The colon is cut after tying it off to prevent spillage, and the entire viscera is laid out on the dissection table for examination (Figure 4.15). The liver is the largest organ and is normally a mahogany color, but may be light brown or tan due to lipid accumulation or dark brown to black if atrophied. The gallbladder is adjacent to the liver in all crocodilians and most lizards; however it is located caudal to the liver in some lizard species. Crocodilians have a single intracoelomic fat body that is located within the right caudal quadrant of the ceolomic cavity (Figures 1.9–1.10, 4.9). This structure is commonly mistaken for other organs such as the liver or pancreas and should be identified and assessed as an indicator of nutritional condition. The pancreas is located near the duodenal loop (Figures 1.128, 4.15). The spleen is an ovoid, dark red structure and is closely associated with the pancreas and near the stomach (Figures 1.128, 4.15–4.16). The close association of the spleen and pancreas is most notable in some lizards, such as varanids, where areas of the endocrine pancreas are observed within the spleen and may be confused with pathologic lesions. The gastrointestinal tract of lizards and crocodilians is a simple structure comprised of the stomach, small intestine, and large intestine (Figure 4.15). The intestinal tract serosa, as well as the serosal lining of the ceolomic cavity of some lizard species, is pigmented black (not to be confused with pathologic change) (Figure 4.10). The small intestine of crocodilians is normally very thick and muscular. The small and large intestine can be difficult to differentiate during postmortem examination and multiple sections representative of the different regions should be collected. All that remains in the carcass at this stage are the trachea, esophagus, and lungs (if removed separately), as well as the adrenal gland, gonads, kidneys, and urinary bladder. To examine the respiratory tract, open the pharynx and trachea and continue the incision into the major airways of the lungs (see Section 4.4.4.2 for illustration). The extrapulmonary bronchi are extensive in adult crocodiles and gharial and should be carefully examined for lesions and exudate (Figure 4.17). Carefully palpate the lungs for areas of firmness, nodules, or other abnormalities and open smaller airways to look for exudate and parasites. The adrenal glands and genitourinary tract lie in the caudodorsal aspect of the coelom. The kidneys are extremely caudal in most lizards and often sit in a retrocoelomic space underneath the pelvic brim and require cutting of the pelvis for complete examination and removal (Figures 1.187, 4.18–4.19). Testes often are mistakenly collected as kidneys when the prosector fails to cut through the pelvis. Enlargement of the kidneys will result in protrusion of the kidneys beyond the pelvis and is a useful marker for renal disease in some lizard species (Figure 4.19). Also, sexually active male lizards may exhibit formation of sex segments in the

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kidneys (see Chapter 1), which can be very prominent grossly as areas of white discoloration (Figure 4.10). Crocodilians and some lizard species lack a urinary bladder and ureters empty directly into the urodeum (Chiasson, 1962). The adrenal glands are typically paired elongate structures, and in lizards are incorporated into the mesenteric connective tissues medial to the gonad in both males and females (Gabe, 1970). If not immediately visualized, adrenal glands may be located by gently palpating the cranial aspects of the kidneys, supportive tissues of the reproductive tract, and associated structures. Adrenal tissue is typically yellowish or pale tan. Adrenal glands and gonads are often collected together unless the ovaries of a female are enlarged due to active folliculogenesis. After all ceolomic organs have been removed from the carcass, samples of muscle, bone, and nerve can be collected. In animals less than 10.0 kg, it is easy to collect sections of femur, bone marrow, surrounding muscle, and sciatic nerve by cutting a section from the distal aspect of the femur and collecting it with all of the surrounding tissues. Bone samples from larger specimens are collected with shears or saws. To examine the brain, the animal is decapitated at the atlantooccipital junction (if this was not done at the beginning of the necropsy). This step can usually be accomplished without the use of a saw or bone shears by incising into the atlantooccipital joint and transecting the spinal cord with a knife or scalpel. The skin overlying the skull of most lizard species can be reflected to allow access to the skull bones for brain removal. In crocodilians, the skin and skull are intimately associated and cuts for brain removal are made through skin and bone concurrently. For small to mediumsized lizards and small crocodilians, the skull is incised with rongeurs or a Dremel tool in a roughly rectangular or trapezoidal pattern extending from the foramen magnum to the level of the midorbit (Figures 4.20A–C). In larger crocodilians, a similar approach is made using a handsaw or Stryker saw to cut through the thick bone (Figure 4.21). The brain is removed and collected whole into formalin or bisected longitudinally for multiple studies (i.e., half for formalin, half for viral isolation). Eyes can be collected from the skull before or after brain removal. A short section of spinal cord can be collected from the cranial aspect of the cervical spine in medium and large specimens. Methods for examination and removal of the spinal cord are described in Section 4.4.4.2. In smaller specimens, a section of vertebrae with spinal cord intact may be collected whole into formalin for later decalcification and histologic examination.

4.4.4.2 Snakes  Prior to examination of a snake, there are special precautions that must be considered if working with a venomous species. The same precautions apply to venomous lizard species. As the first step, the head should be secured with the mouth closed by taping the jaws shut or by inserting the head into a hard cylindrical container (Figure 4.22). An

empty syringe case works well to secure the head, depending on the size of the specimen being examined. The head should then be removed and placed into formalin to deactivate the venom prior to collection of the brain or any other samples from the head. Most, if not all, venom components are destroyed by formalin fixation; however, it is unclear if some compounds may remain toxic in some instances, especially if fixation of deep tissues is incomplete. Therefore, the heads of venomous species should always be handled with extreme care by experienced personnel and eye protection should be used when incising venom glands. Also, be aware of the location of the fangs when handling both venomous and nonvenomous species to avoid personal injury.   Dissection of the snake is begun by placing the animal in dorsal recumbency. An incision is made with a scalpel or scissors along the ventral midline from the cloaca cranially to the intermandibular space. In larger snakes, it may be necessary to make the initial skin incision at the lateral edge of the thick ventral scales. Once the skin incision has been made, the skin is reflected laterally along the length of the incision to expose the underlying subcutis and muscle (Figure 4.23). Muscle sections can be collected at this time. The temporomandibular junction is then incised to facilitate detailed inspection of the oral cavity. The tongue, glottis, proximal esophagus, and trachea can be examined and collected. The coelomic cavity is entered via a midline incision and the entire length of the cavity is visualized at this time (Figure 4.23). Snakes in fair or good nutritional condition have prominent fat bodies in the coelomic cavity that extend cranially to the middle region of the body. As in other species, the thymus and endocrine organs cranial to the heart are identified and collected before dissection continues. The thymus in snakes is a paired structure with cranial and caudal lobes located cranial to the heart. The snake thyroid gland is a single structure (Lynn, 1970) (Figures 1.271, 1.277–1.279, 4.24). Snakes possess two pairs of parathyroid glands; one pair is often located between the anterior and posterior lobes of the thymus, the second pair located at the bifurcation of the carotid artery (Clark, 1970). The coelomic viscera of the snake can often be removed in toto. The esophagus and trachea are transected caudal to the pharynx (Figure 4.25). With gentle caudal traction on the free ends of the esophagus and trachea, the ceolomic viscera can be lifted caudally from the carcass using blunt dissection techniques or occasional sharp dissection to sever connective tissue attachments (Figure 4.25). The viscera are removed caudally to and including the cloaca and are placed on the dissection table for examination and sampling (Figure 4.26). Be aware of the scent glands in the cloacal region, which will emit a strong odor if punctured. Snakes have a three-chambered heart comprised of two atria and one ventricle (Figure 4.27). An incision is made through the apex and continued into the atria and major vessels to visualize the endocardial surfaces and valves. Next, the trachea, bronchi, and lung(s) are examined. In many

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snakes, the left lung is significantly smaller than the right or completely absent. Boids are among the species with a well-developed left lung. The trachea should be opened and the incision continued into the axial chamber to allow complete examination (Figure 4.28). The snake lung contains a large axial chamber that terminates into a long air sac that may extend into the most caudal aspects of the coelom (Figures 1.129, 1.171, 1.180, 4.29). Multiple samples should be collected from different areas of the lung. The liver is an elongate brown organ. The gallbladder in most snakes is located distal to the liver and is connected by a long common bile duct (Figures 1.131–1.133, 4.25, 4.30). The spleen is often located distal to the stomach within the mesenteric connective tissues and is a small reddish round structure (Figures 1.131–1.133, 4.30). The pancreas is a smooth or multilobular, pale tan organ located caudal to the spleen near the duodenum (Figures 1.131–1.133, 4.30). Some snake species possess a fused spleen and pancreas (a.k.a. splenopancreas) (see Chapter 1). As demonstrated in Figure 4.30, a useful technique for locating the spleen and pancreas is to first locate the gallbladder and then examine the surrounding tissues for these organs. The gastrointestinal tract of snakes is a simple structure comprised of the stomach, small intestine, and large intestine. The small and large intestines are difficult to distinguish during postmortem exam and multiple representative sections should be collected. The adrenal glands of snakes are thin, elongated structures located within the connective tissues that support the gonads (mesorchium and mesovarium) in both males and females (Gabe, 1970) (Figures 1.291–1.292, 4.31). The adrenal glands can often be recognized by their yellowishtan color and are typically collected with the gonads. Snakes lack a urinary bladder. The kidneys of snakes are multilobular and are located cranial to the cloaca. The kidneys are usually dark brown (Figures 1.182–1.183), but will turn light tan in reproductively active males due to sex segment formation (Figures 1.184, 4.32). At this stage of the necropsy, all that remains in the carcass is the spinal column, associated musculature, skin, and head. If not previously sampled, various muscle groups can be examined and collected at this time. It is a good practice to palpate the ventral surfaces of the vertebrae of snakes for irregularities. Vertebral disease may be examined by collecting cross sections of vertebrae whole into formalin (less than 1 cm in thickness) for later decalcification and histologic examination. The head is removed at the atlantooccipital junction. For small snakes (those with heads measuring less than 2 cm in length), the head may be collected whole into formalin, later decalcified, and then serially sectioned. For larger snakes or cases requiring detailed examination of the nervous system, the brain should be removed with small bone-cutting shears or a Dremel tool (Figures 4.33A–D).

There are two basic methods for removing the spinal cord. The first method involves separating segments of the vertebral column and extracting the spinal cord from each segment (Figures 4.34A–C). This technique is useful for larger reptiles, including crocodilians, and is an alternative if a Stryker saw or Dremel tool is not available. The second method exposes the spinal cord by performing a dorsal laminectomy using a Stryker saw or Dremel tool and rongeurs (Figures 4.35A–B). This method requires some experience to perform without damaging the spinal cord, but allows more careful examination of the spinal cord and potentially produces less histologic artifact. Removal of the eyes of snakes requires special consideration due to the presence of the spectacle, which is a fused eyelid complete with a thin dermis and epidermis. It is important that the orientation of the spectacle and cornea remain preserved, especially for evaluation of ocular disease. The entire globe and associated spectacle are removed by cutting a square in the periorbital skin and dissecting around the globe, severing extraocular muscles and other attachments (Figures 4.36A–B).

4.4.4.3 Turtles and Tortoises  An excellent description of sea turtle anatomy and necropsy techniques is available (Wyneken, 2001). As mentioned previously, a specific marine turtle necropsy form is available to help guide the prosector. As with other reptiles, chelonians are necropsied in dorsal recumbency (plastron up). The plastron is removed intact by cutting through the skin around the plastron edges and through the attachment to the carapace on both sides at the marginal bridges (Figure 4.37). In sea turtles, the marginal bridge can be easily cut with a knife (a saw is not required) by inserting the blade lateral to the inframarginal scutes. In other turtles and tortoises, the marginal bridge must be cut with a saw or bone shears (Figures 4.38A–B). Attachments of the pectoral and pelvic muscles to the plastron are cut close to the plastron as it is lifted from the body. The gular area of the lower jaw is incised just medial to and along the edges of the mandible. This incision is extended into the oropharynx to expose the tongue, glottis, and proximal trachea, which are lifted and exteriorized. At this stage, the oral cavity is visualized and tissues are sampled as needed, including portions of tongue and glottis for histology. The trachea and esophagus are severed immediately cranial to the base of the forelimbs and removed from the carcass as a unit. Next, the forelimbs, hindlimbs and associated skeletal girdles are removed by incising the musculature and cutting through or disarticulating bones as needed, thus exposing the entire coelomic cavity.  Before proceeding with examination and removal of visceral organs, it is easiest to remove the thyroid gland and thymus before anatomic orientation is obscured. The thymus in most chelonians is paired and located adjacent to the carotid or subclavian arteries (Figures 2.4–2.5, 4.39). The single thyroid gland is located at the base of the right and left aortas

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(Lynn, 1970) (Figures 1.276, 4.40). The parathyroid glands may be difficult to identify, but are located adjacent to or within lobules of thymus (Clark, 1970) (Figure 1.281). Following identification of the endocrine organs above, the pericardium is incised and the heart is removed and sampled. Turtles and tortoises have a three-chambered heart with two atria and one ventricle. The apex of the ventricle is attached to the pericardium by the gubernaculum cordis (Figure 4.41). The pulmonary arteries of sea turtles contain sphincters that appear as segmental accordion-like circumferential ridges on the endothelial (luminal) surface. The large bilobed liver is carefully dissected free of its attachments to the stomach and other connective tissues. The gallbladder is typically green in color and may be collapsed or tightly distended. The gastrointestinal tract and associated organs (spleen and pancreas) are removed by lifting the cut free end of the esophagus and bluntly or sharply dissecting the connective tissue attachments of the stomach and intestinal tract away from the carapace and underlying tissues. Be aware that the esophagus has an abrupt leftward bend in most species at the level of the base of the neck before it enters the stomach. The stomach, small intestine, and colon can be removed in toto, including the cloaca. Near the distal colon and cloaca, the urinary bladder is located ventral to the rectum and is removed and examined separately from the intestinal tract. The pancreas is located near the pyloric sphincter and duodenum within the mesenteric connective tissue of the intestinal tract. As in other reptiles, the spleen and pancreas are closely associated (Figure 4.42). After the liver and gastrointestinal tract are removed, the lungs, gonads, kidneys, and adrenal glands are easily visualized (Figure 4.43). In addition, the right and left aortas and dorsal aorta typically are left in the carcass and can be opened at this time (Figure 4.43). The lungs are adhered to the carapace and can be examined within the carcass or removed for closer inspection. To remove the lungs, lift gently on the cut end of the trachea and sharply dissect the delicate lung tissue from the peritoneum covering the inner surface of the carapace. The adrenal glands, gonads and kidneys are located caudal to the lungs (Figures 1.167, 1.186, 1.226, 1.288, 4.44). The adrenal glands are often flattened, orange-yellow in color, and lie ventral to the kidneys along the midline. In some species the adrenal glands are fused together. The kidneys are located in a retrocoelomic space (a thin membrane separates the kidneys from the coelomic cavity) and lie against the plastron. The gonads, including the testis and vas deferens in the male or the ovary and oviduct in the female, are attached via supportive tissues along the lateral aspect of the caudal coelom. Numerous follicles of various sizes may be present in reproductively active females (Figures 1.227, 1.233, 4.45). The head should be removed from the cervical spine at the atlantooccipital junction. In small turtles and tortoises (those with a head less than 2 cm in length), the head can

be collected whole for later decalcification. This technique allows the head to be serially sectioned, which is helpful for examining the middle ear in cases of aural abscess formation. In larger animals, the brain must be extracted using a Stryker saw or Dremel tool. Removal of the brain from sea turtles is illustrated in Figures 4.46A–H. This method also is applicable to other large chelonians. Cervical vertebral segments and the cervical spinal cord can be examined in smaller specimens by collecting crosssections into formalin for later decalcification. For larger animals such as sea turtles, a dorsal laminectomy can be easily performed using rongeurs to expose and collect the cervical cord. When indicated, spinal cord segments within the vertebrae of the carapace are collected using bone shears or a saw. The ventral aspects of the vertebral bodies are cut away using a Stryker or handsaw to expose the ventral aspect of the spinal cord, which is then carefully removed. Necropsy sample collection is completed with removal of the eyes and any other special tissues that may be required.

4.5 Sample Storage and Submission A variety of chemicals are used to preserve the diverse tissues collected during a necropsy. It is important to remember that multiple tissue types and sample types often are necessary to arrive at a definitive final diagnosis. A sampling protocol should be followed to ensure that the most valuable samples and information are collected in a systematic fashion. Be prepared to collect the following samples: (1) tissues for cytology and histopathology, (2) tissues for electron microscopy, (3) samples for microbiology, (4) tissues for toxicology, and (5) parasites. This section outlines the relevant techniques and materials for collecting these samples.

4.5.1 Cytology Examination of touch impressions and wet mounts of various lesions is extremely helpful for diagnosing disease problems in reptiles. Cytologic samples can be processed easily and rapidly, thus important information can be gathered soon after or during the necropsy, often before histopathology results are available. Furthermore, some infectious agents, such as chlamydiae and blood parasites, are more easily observed in cytologic preparations and blood smears, as are important structures of protozoa that are necessary for identification. Although cytology is a valuable clinical tool, this section will focus on applications relevant to necropsy. The appropriate cytologic method will depend on the nature and consistency of the lesion to be examined. A common problem with cytologic samples is that samples are smeared or applied too thickly on slides to accurately assess them by microscopy. Generally, a sample is appropriately distributed on a slide if one can easily see through the 

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material after it is applied. There are several methods for creating preparations that will yield valuable diagnostic information. The most common are fine-needle aspiration, squash preparations, touch preparations, and skin or shell scrapings. Lesions containing fluid or purulent material or firm masses are easily sampled using fine-needle aspiration. If the lesion is on the skin surface, it should be cleaned with a small amount of 70% ethanol and allowed to dry prior to sampling. Aspirates are discharged onto a slide and direct smears are prepared using standard techniques. Alternatively, exudates or material can be collected with a swab and gently rolled onto a microscope slide to obtain the appropriate thickness. Some organisms, such as protozoa and nematode larvae, are more easily observed in a wet mount (a small amount of saline is added to the sample and a coverslip is applied) than in a stained preparation. Aspirates containing tenacious material with thick cellular fragments should be prepared using a squash technique, whereby a second slide or coverslip is used to flatten the material and both slides are quickly pulled apart. Touch impressions of tissue samples is another valuable cytologic technique and commonly are collected during necropsy. The cut surface of a specimen is blotted onto a paper towel or other absorbent surface to remove blood and then is applied to a microscopic slide to exfoliate cells and material (Figures 4.47A–B). For solid or firm tissues, the surface may be scratched with a needle or blotted on gauze to aid exfoliation. Scrapings can be collected from cutaneous lesions, such as shell lesions of chelonians or hyperkeratotic surfaces. Scrapings are spread on the slide surface using a scalpel blade or coverslip, allowed to dry, and stained by routine methods. Pathogens within keratinized material can be visualized with this method. In addition, potassium hydroxide can be used to digest keratinized material and improve examination for pathogens. Depending upon the suspected disease or presence of pathogenic organisms, a variety of staining techniques can be used to evaluate smears and detect infectious agents. It is a good practice to make at least three replicates of all slide preparations or smears so that ample material will be available for special stains, such as Gram stain for bacteria, acid fast stain for mycobacteria, and fungal stains such as new methylene blue and gomori methamine silver (GMS). The method of fixation of the sample to the slide will depend upon the staining procedure used. Wright-Giemsa stain and rapid commercial staining kits such as Dif-Quick® are most commonly used for initial evaluation of cytologic samples. For Wright-Giemsa stain, the smear is fixed in absolute methanol for approximately 10 seconds. Commercial kits offer rapid fixation and staining of samples in minutes; thus cytology is easily incorporated into examination of necropsy samples.

4.5.2 Light Microscopy The most common preservative used for diagnostic samples collected at necropsy is 10% neutral phosphate buffered formalin (NBF). Tissues collected in formalin are used for histopathology, special stains for infectious agents, and some molecular tests. The two key considerations for preserving tissues in formalin are: (1) samples must be of the proper thickness and (2) an adequate amount of formalin must be used. To achieve adequate fixation, tissue samples generally must be around 1.0 cm or less in thickness (Figure 4.48). Formalin can penetrate only 0.5 cm of tissue in 24 hours; therefore, sections that are too thick will decompose (autolyze) despite being immersed in formalin preservative. Among the most common reasons for improper sizes of tissue samples are inadequate cutting instruments and cutting surfaces; therefore, be prepared with a sharp knife, scalpel, or razor blade and have a cutting board available. The ratio of 10% NBF to tissue should be 10:1 (i.e., 10 times as much liquid as tissue present in the container). Failure to fix tissues in the appropriate amount of formalin will similarly result in autolysis of the tissues after collection. Very small tissues (< 5.0 mm in size) can be placed into plastic cassettes to ensure they are not lost during submission (Figure 4.49). Hard tissues, such as bone, should be fixed in a separate formalin container to allow adequate penetration and fixation prior to decalcification. The eyes from larger specimens will also require decalcification due to the presence of scleral ossicles (bones). The brain from cases with evidence of neurologic disease can be specially fixed in high-concentration formalin. First, the brain is placed whole into a separate container with enough 37% formaldehyde to just cover the brain tissue. Next, 10% NBF is added to the container until the brain floats neutrally buoyant. Fixation of the brain tissue is complete when the brain sinks to the bottom of the container, usually after approximately 24 hours. Other fixatives used for light microscopy are Bouin’s fixative and Davidson’s solution. Bouin’s fixative was historically used for fixation of eyes and some reproductive tissues, and is still used by some specialists. This fixative is no longer popular with most pathologists due to the highly explosive picric acid ingredient. Formalin is an adequate preservative for eyes, but the lens becomes very hard during fixation and it can be very difficult to obtain quality sections of the globe. Davidson’s solution is an alternative fixative for eyes and does not contain picric acid. Representative sections from all organs should be collected. A list of complete tissue and sampling sites is provided in Table 4.3. Tissue specimens from organs that appear normal should be saved in addition to obvious lesions. Specimens from lesions should be representative of the entire lesion and should be large enough to include adjacent normal tissue, which facilitates comparison with diseased areas. Furthermore, active processes and primary etiologic agents are often found at the edges of lesions.

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Table 4.3   Routine tissue samples and site collected for histopathologic examination.a Skin: One or more sections is collected from various body sites. Muscle: Muscle sections are typically collected from the hindlimbs of chelonians, lizards, and crocodilians, and from the dorsal musculature of snakes. Bone and bone marrow: Ribs are collected from snakes and larger chelonians, lizards, and crocodilians. The whole femur or humerus can be collected from smaller animals. The femur, humerus, or other long bone should be cracked or cut to allow penetration of formalin into the marrow space. Plastron and carapace: A thin section of each can be taken from any aspect, but typically is easiest at the lateral margins. Thyroid gland: Collect whole if less than 1 cm or collect a representative section if larger. Thymus: A 1.0 cm section is collected with surrounding connective tissue. Parathyroid glands: Collect whole with surrounding connective tissue. Often the internal or external carotid arteries are attached. Oral cavity: Any lesions are collected with adjacent normal tissue. Tongue: The tip of the tongue is collected. Esophagus: A cross-section through the mid-portion is collected. Trachea: A complete tracheal ring is collected anywhere along the length. Heart: A section from one atrium and the ventricle, including atrioventricular valve is collected. Sections of aorta and great vessels are also collected. Lung: In species with two distinct lungs, at least one section from each lung is collected to include portions of larger cartilaginous airways. Preferably, multiple sections from different areas are collected. Liver: Collect at least two sections from different lobes. If the gallbladder is present within or adjacent to the liver, liver and gallbladder sections can be collected together in one piece. Gallbladder: Collect as part of the liver section or separately as a small portion of the gallbladder wall. Pancreas: If pancreas and spleen are separate organs, the pancreas should be collected with a small portion of adjacent duodenum to facilitate identification of the tissue. If the pancreas and spleen are fused, a single section can be collected to include both splenic and pancreatic tissue. Spleen: If separate from the pancreas, collected intact (after collection of any frozen samples). For larger reptiles, a complete crosssection is collected. Stomach: Sections are collected near the esophageal sphincter, within the body and from the pyloric region. In smaller specimens, one long section may be collected that includes all three sites. Small and large intestine: Tubular sections of intestinal tract (i.e., unopened) are collected from proximal, middle, and distal portions. A small incision can be made in one end of the collected segment to allow formalin penetration into the lumen, especially for larger specimens. Care should be taken to avoid damaging the mucosal surface of the intestinal tract with instruments or by rough handling (touching, scraping, etc.). Cloaca: A small portion of the cloacal wall, including the mucocutaneous junction, is collected. Adrenal gland: Collect either whole or as part of the adjacent gonad. Both left and right glands are sampled. Gonad: If possible, sections of gonad should include adjacent oviduct or vas deferens on both left and right sides. In larger animals, separately collect sections of the tubular portions of the reproductive tract. Kidney: Sections from cranial and caudal poles of both left and right kidneys are collected and should include both cortical and medullary regions. The ureter may also be collected if visible. Bladder: A small portion of the wall (0.5 cm2) is collected, taking care not to destroy the mucosal surface by rough handling. Spinal cord: Thin segments of vertebrae with spinal cord (< 1.0 cm) can be collected for decalcification. If the spinal cord is removed from the vertebral column, almost any length can be collected due to the overall thin cross-sectional diameter of the cord. Extract the cord very carefully, avoiding bending and stretching. Peripheral nerves: A length of sciatic nerve is most commonly collected with adjacent hind limb muscle. Ideally, the ends of the nerve should be fixed to a rigid surface (e.g., cardboard or tongue depressor) prior to fixation. Brain: If the brain is removed from the skull, it is placed whole into formalin or split longitudinally into halves, one for formalin and one for freezing or other uses. Eyes: The entire eye is immersed in formalin or other fixative without puncturing the globe. a

These recommendations assume that gross lesions are not present, and that any lesions that are present are collected as part of the routine sample.

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4.5.3 Transmission Electron Microscopy Please see Chapter 6 for a detailed description of electron microscopy fixation techniques.

4.5.4 Toxicology In most instances, only fresh or frozen tissues are suitable for toxicologic analyses. Considerations regarding sample type, collection procedures, packaging, and storage are different for specific toxicants or compounds and will require consultation with the receiving diagnostic laboratory or toxicologist. Samples for heavy metal analysis typically include liver, kidney, and skeletal muscle, which are placed in separate plastic bags and frozen on dry ice or in an ultrafreezer until submitted. Samples for pesticide analysis, including fat, liver, kidney, and skeletal muscle should be wrapped in aluminum foil, placed in plastic bags, and similarly frozen. Approximately 200 grams of tissue should be the minimal amount collected if possible.

4.5.5 Microbiology A thorough diagnostic workup of suspected viral, bacterial, fungal, and parasitic diseases includes attempts to isolate and identify the pathogen. Specimens must be collected and transported in a way that preserves pathogen viability, minimizes contamination by commensal and incidental microorganisms, and minimizes overgrowth of the specimen by fast-growing species. To prevent contamination, blood, tissue fluids, exudates, and tissue biopsies obtained for microbial culture must be collected under aseptic conditions using sterile instruments and appropriate techniques. Samples that are not collected, handled properly, or delivered to an experienced clinical microbiological laboratory in a timely manner often yield spurious or confounding results. Instruments can be “sterilized” for the collection of microbiologic samples in the laboratory or in the field by using alcohol and a flame source. Instruments (typically forceps and scissors) are allowed to soak in 90% to 100% ethanol for at least 5 minutes. The flame source is used to burn the ethanol off the instruments prior to touching the tissues to be cultured. The instruments should be allowed to cool before sampling tissues as excess heat will kill infectious agents, especially if the sample is relatively small. Tissue samples should be collected in a sterile container and transported to the laboratory as soon as possible. Additional considerations for tissue samples are discussed in Section 4.5.7. A variety of culture swabs or culturettes are commercially available for both aerobic and anaerobic culture, and may be used to swab lesions or body cavities, or as an alternative to collection of tissue sections for culture. A common method of obtaining bacterial cultures during a postmortem examination is referred to as the sear and stab technique (Figures 4.50A–C). A scalpel blade is heated by a flame source and the flat side is applied (while hot) to the organ or tissue to be cultured, thereby sear-

ing the outside and killing any surface organisms. The scalpel is then stabbed into the tissue through the sear site and a culture is collected through the incision. Contact should be made with the receiving laboratory well in advance so that they can advise the field worker on the laboratory’s capabilities, submission deadlines, proper collection materials, and preferred transport media. Although many species of bacteria and fungi can be cultured using standard media and procedures, many important microorganisms, such as Mycoplasma species and Mycobacterium species, require specialized culture media and conditions.

4.5.6 Molecular Diagnostic Tests Detection and characterization of pathogenic organisms has made tremendous advances with the development of molecular techniques, such as nucleic acid hybridization (Southern and northern blotting, in situ hybridization) and amplification techniques (polymerase chain reaction [PCR]). Although molecular diagnostic tests exist for many bacteria and fungi shared between reptiles and other vertebrates, those for pathogenic organisms unique to reptiles are in the relatively early stages of development and are routinely available for a very limited number of infectious agents (Chapter 7). Furthermore, most of these tests are offered only by a few research laboratories. Nevertheless, prosectors should anticipate the eventual availability of additional tests and collect the appropriate specimens. The preferred samples for most diagnostic molecular techniques are frozen samples, stored in either a conventional freezer (−20°C) or ultrafreezer (< −70°C). Ultrafreezing offers better preservation and increases the chances of subsequent microbial or viral isolation. Immediate freezing and storage of fresh tissue samples in liquid nitrogen is preferable for research requiring nondegraded DNA and RNA. Tissue samples or parasites preserved in ethanol are also suitable for molecular methods such as PCR. Paraffin-embedded, formalin-fixed tissues (tissue blocks) may be used for molecular detection; however nucleic acid degrades over time and yield will greatly depend on the duration of fixation in formalin and the requirements of specific assays. For many applications, ability to obtain reliable results using molecular techniques significantly declines after about 48 hours in formalin; thus tissues should be processed into paraffin blocks as soon as possible after complete fixation. Some techniques are useful for obtaining results from suboptimal samples; thus specialists or laboratory personnel should be consulted for special circumstances.

4.5.7 Handling Tissues for Diagnosis of Specific Pathogens 4.5.7.1 Viruses 

Preliminary diagnosis of viral disease is usually made by histopathologic examination of fixed tissues

232  Reptile Necropsy Techniques

obtained by biopsy or at necropsy (see Chapter 9). Coupled with history and clinical signs, the occurrence of characteristic cytopathologic changes, such as cell degeneration (swelling and lysis), syncytia formation (fusion of adjacent cells), and intranuclear or intracytoplasmic inclusion bodies, provide the first clue that a viral agent may be involved. Electron microscopic examination of these fixed tissues may confirm the presence of virus-like particles within the lesion and provide preliminary identification of the agent. Complete diagnosis is achieved by virus isolation from fresh or frozen (< −70°C) samples in an appropriate tissue culture system, followed by immunological and molecular characterization of the isolate. In cases where an appropriate cell culture system has not been developed for the agent, further identification may be achieved by agent-specific immunohistochemical techniques using agent-specific antibodies or by molecular techniques using oligonucleotide probes and primers (Chapter 7).  In suspected cases of viral disease, the minimum samples collected should include a complete tissue set preserved in 10% NBF and frozen samples of all major organs and lesions. Obviously clinical symptoms, such as respiratory or neurological disease, will prioritize sample collection from affected systems. Ideally, viral isolation is attempted using fresh (not previously frozen) tissues, as some viruses, such as herpesviruses, are compromised by the freezing process (see Chapter 13). However, viral culture services, which require maintenance of appropriate cell lines, are not offered on a routine diagnostic basis for reptiles. Some research laboratories, however, maintain cell lines suitable for isolation of reptile viruses. In an outbreak or special interest situation, some researchers will attempt viral isolation and should be consulted for sample requirements. Fresh tissue samples are placed in virus transport media (serum-free cell culture media containing antibiotics and antifungals) and shipped on ice to the laboratory as soon as possible. More often, attempted virus isolation is delayed and comparatively more laboratories offer molecular tests, thus the standard approach to sample collection for diagnosis of viral disease in most situations is collection of frozen tissues, which is extremely important for cases of suspected viral disease. Samples are best stored at −70°C or preferably in liquid nitrogen, which increases the chances for future virus isolation. Many of even the most environmentally sensitive viruses usually retain some infectivity if they are rapidly frozen and stored below −70°C. If unavailable, samples preserved in a standard freezer (−20°C) are still adequate for molecular detection (such as by PCR) and genetic studies. Electron microscopy (EM) can be performed on NBFfixed tissues, as well as tissues that have been processed and embedded in paraffin for routine histopathology. However, special fixatives for EM greatly improve the ability to detect and characterize agents and ultrastructural pathologic changes (see Chapter 6).

4.5.7.2 Bacteria and Fungi  Gross and histologic examination of lesions or cytology usually provide the first evidence of bacterial or fungal disease (see Chapters 10 and 11). In addition to routine hematoxylin and eosin, special tissue stains such as tissue Gram stains (Brown and Brenn), silver impregnation stains (Warthin-Starry, Gomori methamine silver), and acid fast stains (Zeihl Nielson and Fites) can help narrow the range of possible agents. Smear preparations of lesion exudates or impression smears of affected tissues can be made, stained, and examined in the field. Submission of specimens for bacterial and fungal culture should follow the guidelines previously discussed (microbiology; see Chapter 13). Also, immunodiagnostic and molecular diagnostic techniques can be applied to fixed or frozen tissues or to culture isolates. 

4.5.7.3 Protozoan Parasites  To date, most protozoal diseases documented in reptiles are pathogens of the gastrointestinal tract (see Chapter 12). Fecal analysis (direct smears, flotation) performed on a fresh postmortem fecal sample (or cytologic preparations of the gastric or enteric mucosa in the case of Cryptosporidium) can aid in detection. Many gastrointestinal protozoa, however, are commensal organisms; therefore, detection of organisms within characteristic histopathologic lesions is often the first indication that a species is pathogenic. Systemic protozoal infections, such as microsporidiosis, also are commonly diagnosed by histologic examination (see Chapter 12). Although histopathology is invaluable, subcellular structures that are critical to positive identification of organisms may be obscured or unapparent in histologic section. Therefore, cytologic specimens, electron microscopy, and molecular studies may be necessary for definitive identification, and are especially important for characterizing novel protozoan pathogens. Samples for these purposes should be collected as previously described.

4.5.7.4 Metazoan Parasites   Metazoan or multicellular parasites include endoparasites, such as nematodes (roundworms), trematodes (flukes), and cestodes (tapeworms), as well as ectoparasites such as ticks and mites. Endoparasites are found in virtually all organ systems of reptiles (see Chapter 12). Parasites are commonly encountered in free-ranging or wild-caught animals or in reptiles maintained on natural substrate or other conditions where parasite intermediate hosts or parasites of naturally infected reptiles may be encountered. Special techniques are available for the collection of parasites, such as the use of sieves to examine gastrointestinal contents, but these are usually performed only as part of research protocols. The more common scenario encountered during necropsies is the collection of organisms observed with the naked eye or in cytologic specimens. The most commonly used and widely available preservative for parasites collected during necropsy is 70% ethanol, which is suitable for preserving both endoparasites and ectoparasites. Flatworms (trematodes and cestodes) should be relaxed in

Reptile Necropsy Techniques  233

water for a few hours prior to fixation. Also, acanthocephalans must be left in water until the proboscis is extended if specimens are to be identified later. Specialty preservatives, such as alcohol-formalin-acetic acid solution (AFA) for trematodes, may be provided by consulting parasitologists and should be used appropriately if available.

4.6 Necropsy Precautions and Zoonotic Disease Concerns The phylogenetic distance and physiological differences separating reptiles from humans lowers the risk of disease transmission from reptiles to man. However, reptiles may harbor a number of bacterial species that are known human pathogens or are opportunistic pathogens in a wide range of vertebrate species. The primary agents of concern are bacterial species of the genera Mycobacterium, Salmonella, Vibrio, and Chlamydiophila. Unfortunately, we do not have enough information about many infectious agents in reptiles to be certain of the risks. Although it is unlikely that field personnel will use universal precautions in handling reptiles, they should realize that risks exist and have appropriate materials available for disinfecting wounds received while handling these animals. Personnel should seek medical attention if wounds become infected or if they become systemically ill after working with reptiles. Gloves should always be worn when performing necropsies. 

4.7 After the Necropsy 4.7.1 Cleanup Considerations Disinfection of clothing and equipment following a necropsy procedure is essential to reduce the risk of contaminating future specimens and to reduce the likelihood of transmitting disease to other animals that come into contact with the prosector, equipment, or surfaces. Latex gloves, masks, scalpel blades, needles, and other disposable items should be discarded after each necropsy and appropriate biohazard containers should be used. Aprons, boots, instruments, and any other materials that come into contact with a carcass, tissues, or fluids should be cleaned thoroughly with hot water and detergent, followed by a good-quality disinfectant. Decontamination techniques should be adapted to the field whenever possible. Dilute sodium hypochlorite (10% bleach solution, 1 part bleach to 10 parts water) is an excellent and inexpensive disinfectant, but it is corrosive and is rapidly deactivated by organic debris; therefore, washing before application and thorough rinsing are necessary. Glutaraldehyde solutions or formalin are also powerful disinfectants but are very toxic. Chlorhexidine solutions and povidone iodine solutions are effective and less toxic alternatives. If the presence of myco-

bacteria is suspected, phenol derivatives are more effective for killing organisms on instruments and surfaces. Alcohol is not an effective disinfectant on surfaces or instruments unless the instrument is flamed or left in contact with alcohol for very long periods of time. Metal instruments should be allowed to air dry or should be autoclaved following cleaning.

4.7.2 Electronic Storage, Archiving, and Retrieval of Reports The days of storing necropsy reports solely as hard copy paper reports are nearly gone as biologists, veterinarians, and pathologists are quickly moving to electronic data storage. Most electronic necropsy reports are stored as either a word processing document (Microsoft Word®, Corel WordPerfect®, or similar program) or as a file in a zoological record-keeping program such as Medical Animal Records Keeping System (MedARKS©) offered by the International Species Inventory System (ISIS©) (Eagan, Minnesota). Other database programs, such as Microsoft Access® also may be used. Actual reports are generated as a Word file and stored in digital PDF format with hard copy backups as a precaution. The search feature of most word processing programs allows for searching of multiple records for key words or specific diagnoses. In addition, most pathologists will code diagnoses by one of several standard systems. Traditional coding systems include the systemized nomenclature of veterinary medicine (SNOVET) and the systemized nomenclature of medicine (SNOMED), whereby individual diagnoses are assigned a code according to topography (location where the lesion occurred in the body), morphology (the character of the lesion), and etiology (the cause of the lesion) (Palotay, 1983). One author uses a simple coding system adapted from a private diagnostic service (Dr. Michael Garner, Northwest Zoopath, Snomish, Washington). Under this system, diagnoses are coded by etiology and body site (Table 4.4) and codes are easily searched using the search features of Microsoft Excel® or Word.

4.7.3 Tissue Archives With proper storage, tissues collected at necropsy can be as valuable in 20 years as they were at the time of collection. Tissues collected at necropsy are typically stored long term in three forms: as wet tissues in bags of formalin or alcohol, as frozen tissues and as paraffin blocks. Tissues can be stored for years as wet tissues in containers of formalin, but formalin will cross-link proteins and degrade DNA over time (a few days to a few weeks) and will render many molecular-based ancillary tests (PCR, in situ hybridization, etc.) impossible. The long-term effect of alcohol on fixed tissues is unknown, and storage in alcohol following fixation is being used to delay protein and nucleic acid degradation. Other problems with long-term formalin storage include evaporation and air quality. Heat-sealed thick plastic containers or pouches will allow some evaporation to occur and will introduce forma-

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Table 4.4   Coding system for electronic archival of necropsy diagnoses. (Adapted with permission from M. Garner, Northwest Zoopath, Snomish, Washington.) M — Mammal

10. Autoimmune and immunosuppressive disease

B — Bird

11. Traumatic disease and environmental exposure (and drowning)

R — Reptile

12. Toxic disease

A — Amphibian

13. Reproductive and perinatal disease

F — Fish

14. Euthanasia

Disease category 1. Noninflammatory cardiopulmonary and vascular disease 2. Degenerative disease

15. No pathologic diagnosis made/diagnosis unknown 16. Necropsy not performed (scavenging, autolysis, lost) 17. Normal tissue

3. Deposition and storage diseases

18. Whole body (for photo index)

a. Amyloid

19. Hyperplasia and hypertrophy

b. Mineral

20. Environmental exposure (heat, cold, water quality)

c. Nonamyloid glomerular deposits

Body site

d. Pneumoconiosus

L — Integumentary system

e. Lipid

N — Cardiovascular system

f. Iron

O— Liver

g. Melanin

P— Gastrointestinal system – non liver

h. Miscellaneous

Q— Hematopoietic system (spleen, lymph node, thymus)

i. Urate deposition / gout

S — Endocrine system

j. Yolk material

T — Musculoskeletal system

4. Anomalies and defects

U— Nervous system

5. Anesthetic/drug/surgical complications

V — Respiratory system

6. Metabolic and endocrine disorders

W— Renal system

7. Nutritional disease

X — Reproductive system

8. Neoplastic disease/proliferative

Y — Special senses (eyes, ears)

9. Infectious and inflammatory disease

Z — Multisystemic disease

a. Bacterial b. Mycobacterial c. Fungal d. Viral e. Parasitic f. Inflammation of unknown cause

Examples Diagnosis

Code

Traumatic skull fracture in a reptile

R, 11T (fracture)

Mycobacterial septicemia in a reptile

R, 9bZ (Mycobacterium chelonae)

Fungal skin and liver disease in a reptile

R, 9cL, 9cO (Mucor sp.)

Fatty liver in a reptile

R, 3eO (hepatic lipidosis)

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lin, which is a carcinogen, into the environment. If long-term storage of tissues in formalin is considered, one should store tissues in a well-ventilated area and plan to periodically refill and re-bag valuable samples every few years. Paraffin blocks are small, stable, and easily stored. To solve the problems associated with formalin evaporation and air quality, some institutions have turned to paraffin blocks as an alternative. Large sections of tissue are trimmed into paraffin blocks for the sole purpose of long-term storage. These tissues do not degrade nor does formalin need to be replenished. This process is, however, time consuming and the large numbers of paraffin blocks must be stored in a climate-controlled environment to prevent melting in summer heat. While most pathologists and diagnostic facilities permanently retain paraffin blocks, one should still inquire of the specific lab to determine policies regarding paraffin block retention and disposal. If your laboratory periodically disposes of paraffin blocks, request that blocks be sent to you for long-term storage. Ultrafreezers are the most common way of retaining tissues to be utilized for a variety of medical and biological studies. Tissues can be stored at −40, −70, or −80°C and optimal conditions depend on the types of tests that will be performed. Ultrafreezers are expensive and may require periodic maintenance to ensure valuable tissue and blood samples are not lost. An alarm system, preferably one with the capability of calling a pager or phone when there is a temperature increase, is highly recommended. Samples placed in an ultrafreezer should be accurately indexed using a logbook or electronic system to avoid tissue loss, and so that time spent locating and removing samples is minimized to reduce thawing and refreezing.

4.8 Conclusion Postmortem examination or necropsy of dead reptiles is an opportunity for veterinarians, biologists, and field personnel to learn about individual, animals, animal populations, and the environment. The necropsy should be systematically conducted with goals in mind that include accurate and objective data collection, detailed description of findings, and complete sample collection. Valuable historical data and necropsy findings should be recorded on necropsy data sheets and supplemented with photographs whenever possible. Complete

tissue sets should be collected and additional samples will depend on the overall objectives of the necropsy and circumstances of death. The diverse anatomical characteristics of reptiles provide a challenge for any prosector that can be overcome only by practice and systematic techniques. With proper equipment, training, and knowledge of dissection and sampling techniques, postmortem examination of reptiles may provide vast amounts of information now and in the future.

References Akers TK. 1966. Some circulatory characteristics of Alligator mississippiensis. Copeia. 1966:552–555. Ashley LM. 1955. Laboratory Anatomy of the Turtle. William. C. Brown Company. Dubuque, IA. Chiasson RB. 1962. Laboratory Anatomy of the Alligator. William. C. Brown Company. Dubuque, IA. Clark, NB. 1970. The parathyroid, in Biology of the Reptilia, Vol 3, Gans C and Parsons TS (Eds.), Academic Press, New York, 235–261. Davies PM. 1981. Anatomy and physiology, in Diseases of the Reptilia. Cooper JE and Jackson OF (Eds.), Academic Press, New York, 9–73. Gabe, M. 1970. The adrenal, in Biology of the Reptilia, Vol 3, Gans C and Parsons TS (Eds.), Academic Press, New York, 236–318. Lynn WG. 1970. The thyroid, in Biology of the Reptilia, Vol 3, Gans C and Parsons TS (Eds.), Academic Press, New York, 201–234. Nagy KA, Henen BT, Vyas DB, and Wallis, IR. 2002. A condition index for the desert tortoise (Gopherus agassizii). Chelon Conserv Biol 4:425–429. Palotay J.L. (1983) SNOMED-SNOVET: an information system for comparative medicine. Med. Inform., London, 8:17–21. Stamper MA, Cornish T, Lewbart G, Epperly SP, Boettcyer R, Braun J, Levine JF, Correa M, Miller R, Moeller R, Driscoll C, and Colbert A. 1997. Cooperative efforts between veterinary diagnostic facilities and government agencies in assessing two sea turtle stranding episodes, in Proceedings of the 17th Annual Sea Turtle Symposium, Orlando, Florida, 4–8. Webb G, Heatwole H, and DeBavay J. 1971. Comparative cardiac anatomy of the Reptilia. I. The chambers and septae of the varanid ventricle. J Morphol. 134:335–350. White FN. 1968. Functional anatomy of the heart in reptiles. Am Zool. 8:211–219. Wyneken, J. 2001. The Anatomy of Sea Turtles. U.S. Department of Commerce, NOAA Technical Memorandum NMFS-SEFSC-470.

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Appendix 4.1 Standard necropsy form used for multiple reptile species. Necropsy #:__________________ Date of necropsy:_ ___________________________ Prosector: _______________________ Species:______________________ Date of death:_ ______________________________ Identifier#:___________________ Age:________________________________________Sex:_____________________________ Medical history / field observations (continue on bottom or back of page): Weight:______________________ Other measurements:__________ Body condition: ______________ poor moderate good obese Carcass condition:_____________ autolyzed good fresh Check boxes for tissues stored in: General exam (external, skin, subcutis, tattoos, bands, etc): Palpate skeletal system for abnormalities: Oral cavity: Ears / eyes: SubQ / peritoneal / ceolomic fat: Musculoskeletal system Bone marrow Ceolomic cavity: Trachea: Thyroid / parathyroid: Thymus: Heart / major vessels / aorta: Esophagus: Stomach: Small intestine: Large intestine: Cloaca and bursa: Pancreas: Spleen: Liver: Lungs: Bladder: Gonads: Repro tract: Adrenal glands: Kidneys / ureters: Brain: Spinal cord: Peripheral nerves:

Formalin

Frozen

Other

Continuation space: Photos? Tissues submitted for culture (list): Tissues frozen (standard set = brain, liver, spleen, heart, lung, kidney, skeletal muscle, intestine): Other diagnostics (list tissue and test): Formalin tissues sent to: (list pathologist, date, tissues): Preliminary diagnoses, comments, or impressions:

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Figure 4.1  Necropsy instruments. Basic instruments required for necropsy include a knife, scissors, forceps, bonecutting instruments (rongeurs and shears), and a cutting board.

Figure 4.2   Sample containers. A variety of containers are needed for collection of samples. Basic requirements include containers of 10% neutral buffered formalin for tissue fixation, tissue cassettes for smaller samples, and tubes and plastic bags for fresh or frozen specimens. All containers should be clearly labeled with appropriate identification.

Figure 4.3   Power instruments. Electric cutting tools are useful for examining the brain and spinal cord, removing the plastrons from chelonians and collecting bone samples. Commonly used instruments are Stryker saws and Dremel tools.

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Figure 4.4   Schneider’s dwarf caiman, Paleosuchus trigonatus. Alligatoridae. Common biological measurements of reptiles include snout-vent length, which is measured from the rostrum or end of the maxilla to the cloacal opening or vent.

Figure 4.5   Loggerhead sea turtle, Caretta caretta. Cheloniidae. Epibiota accumulation on the integument of the appendages and head can be an indicator of underlying disease and poor general health in sea turtles. Numerous small barnacles are adhered to the hind flipper and there is associated dermatitis in this example.

Reptile Necropsy Techniques  239

Figure 4.6   Black and white tegu, Tupinambis merianae. Teiidae. To begin the internal examination, the animal is placed in dorsal recumbency and the first incision is made from the cloacal opening cranially to the intermandibular space (blue dashed line). Courtesy of Philippe Labelle.

Figure 4.7   Black and white tegu, Tupinambis merianae. Teiidae. The rib cage is removed by incising through the costochondral junctions or by cutting the ribs with bone cutters (blue arrows), thus exposing the heart and liver. Courtesy of Philippe Labelle.

Figure 4.8   Black and white tegu, Tupinambis merianae. Teiidae. The entire coelomic cavity is exposed following removal of the rib cage and incision through the coelomic wall. The coelomic fat bodies, which are robust in this specimen, have been reflected caudally over the pelvic region. Courtesy of Philippe Labelle.

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Figure 4.9   Nile crocodile, Crocodylus niloticus. Crocodylidae. As in Figure 4.9, the entire coelomic cavity is exposed. Note the unpaired intracoelomic fat body (blue arrow) unique to crocodilians.

Figure 4.10   Blue panther chameleon, Furcifer pardalis. Chamaeleonidae. Laterally flattened species, such as chameleons, are necropsied by removing the rib cage and coelomic wall on one side, thus exposing all of the visceral organs. Note the black pigmentation of the serosa (normal coloration) in this species. The kidneys (arrow) are light in color due to sex segment formation in this male.

Figure 4.11   Cuban crocodile, Crocodylus rhombifer. Crocodylidae. The intermandibular skin and soft tissues are incised to reflect the tongue and expose the gular valve, glottis, and pharynx for examination.

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Figure 4.12   Monitor lizard, Varanus spp. Varanidae. The paired thyroid glands are located within the ventral cervical (neck region) (white arrows).

Figure 4.13   Green iguana, Iguana iguana. Iguanidae. The paired thyroid glands (TG) in this species are joined by a narrow bridge of tissue. The posterior parathyroid glands typically are located at the bifurcation of the internal carotid (IC) and external carotid (EC) arteries. This area (black arrows) can be collected and serially sectioned to locate the glands. In addition, note the right and left aortic arches (RAA and LAA, respectively), pulmonary artery (PA), atria (A) and ventricle (V). Courtesy of Tanja Zabka.

Figure 4.14   Green iguana, Iguana iguana. Iguanidae. The anterior parathyroid glands are located along the inner (medial) surface of the rami (R) of the mandible and require careful dissection to locate them. One anterior parathyroid gland (black arrow) is visible dorsolateral to the trachea. Courtesy of Tanja Zabka.

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Figure 4.15   Cuban crocodile, Crocodylus rhombifer. Crocodylidae. All visceral organs have been removed and arranged for examination and sampling. Note the coelomic fat body (FB). The heart and genitourinary tract typically are removed separately, but were left attached for demonstration purposes. The lungs (L), heart (H), liver (LV), gallbladder (GB), stomach (SP), spleen (P), pancreas (P), small intestine (SI), large intestine (LI), cloaca (CL), kidneys (K), ovaries (O), and oviducts (OV) are visible.

Figure 4.16   Black and white tegu, Tupinambis merianae. Teiidae. Splenopancreas. The spleen (SP) and pancreas (P) are fused or closely associated in many reptiles.

Figure 4.17   Mugger crocodile, Crocodylus palustris. Crocodylidae. The trachea of adult crocodiles and gharial forms a prominent bend and extrapulmonary bronchi are extensive. Note the thick white pericardial sac immediately caudal to the bronchi.

Figure 4.18   Nile crocodile, Crocodylus niloticus. Crocodylidae. After the gastrointestinal tract is removed, the kidneys and reproductive tract can be examined. This specimen is an immature female and the ovaries (O) and tubular oviducts (OV) are visible on the ventral surface of the kidneys (K).

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Figure 4.19   Green iguana, Iguana iguana. Iguanidae. In many lizard species, the pelvic symphysis must be split to observe the kidneys, which lie far caudally within the retrocoelomic space. Compare the normal kidneys (right) with those of an animal with renomegaly (enlargement of the kidneys) (left).

Figures 4.20A−C   Schneider’s dwarf caiman. Paleosuchus trigonatus. Alligatoridae. The brain is removed from crocodilians by first incising through the parietal and squamosal bones of the skull from the foramen magnum to approximately the level of the mid-orbit (Figure A). The bone is removed by forceps or rongeurs (Figure B) to expose the brain (Figure C). The opening may be widened using rongeurs to facilitate removal of the brain.

Figure 4.21   Chinese alligator, Alligator sinensis. Alligatoridae. The method for removing the brains of larger crocodilians is similar to that illustrated in Figure 4.20. A handsaw or Stryker saw is required to cut through the thick bone.

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Figure 4.22   Black-tailed rattlesnake, Crotalus molossus. Viperidae. The heads of venomous snakes are placed into a protective covering, such as a syringe case as seen here, and held in place with tape as a safety precaution. Preferably, the head is removed and placed into formalin before proceeding with the necropsy.

Figure 4.23   Black-tailed rattlesnake, Crotalus molossus. Viperidae. The coelomic cavity is opened by making an incision from the cloaca to the intermandibular space, thus exposing all of the visceral organs.

Figure 4.24   Gopher snake, Pituophis melanoleuces. Colubridae. The thyroid gland of snakes is a single (unpaired) structure (white arrow) located immediately cranial to the heart base.

Figure 4.25   Black-tailed rattlesnake, Crotalus molossus. Viperidae. The trachea and esophagus are transected caudal to the pharynx and larynx and the carcass is completely eviscerated by applying firm traction in the caudoventral direction and severing attachments as necessary. The cranial aspect of the esophagus may require extra dissection to remove it intact.

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Figure 4.26   Black-tailed rattlesnake, Crotalus molossus. Viperidae. The viscera is placed on the necropsy surface for locating, examining and sampling individual organs. Heart (H), lung (L), liver (LV), esophagus (E), stomach (S), gallbladder (GB), splenopancreas (SP), intestine (I), testes (T), vas deferens (VD), kidneys (K), cloaca (CL) and coelomic fat bodies (FB) are visible in this photograph.

Figure 4.27   Burmese python, Python molurus bivittatus. Pythonidae. The heart is opened by slicing through the apex of the ventricle and continuing the incision into the atria and major vessels.

Figure 4.28   Burmese python, Python molurus bivittatus. Pythonidae. To examine the respiratory tract, the trachea is opened using a pair of scissors and the incision is continued into the bronchi and axial chamber. Note the presence of a welldeveloped left lung, which is a normal anatomic feature of boid snakes.

Figure 4.29   Ball python, Python regius. Pythonidae. The lung is an elongate structure and terminates caudally into a thin-walled air sac. Image courtesy of Philippe Labelle.

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Figure 4.30   Burmese python, Python molurus bivittatus. Pythonidae. The gallbladder (GB), spleen (SP), and pancreas (P) are located in close proximity to one another; thus the gallbladder is a good landmark for locating these organs. Note the location of the stomach (S) and duodenum (D).

Figure 4.31   Burmese python, Python molurus bivittatus. Pythonidae. The adrenal glands (AG) are located adjacent to the gonads (testes [T] in this example) and are easily sampled with the gonadal tissue. Adrenal glands are often light tan as compared to the surrounding tissues and may be located by gentle palpation. Distal colon (C), kidney, and fat body also are visible in this photograph.

Figure 4.32   Western rattlesnake, Crotalus viridis ssp., Viperidae. The kidneys of snakes are elongated, multilobular, and are normally dark brown. The light tan color of these specimens is due to sex segment formation in a reproductively active male.

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Figures 4.33A–D   Burmese python, Python molurus bivittatus. Pythonidae. The first step in removing the brain from larger snakes is to make a dorsal midline incision and reflect the muscles of the mandible to expose the skull (Figure A). Using either a Dremel tool or rongeurs, the dorsal aspect of the skull is cut away from the foramen magnum to the level of the rostral orbit (Figure B). The brain is exposed and the opening is widened as necessary to facilitate removal (Figure C). The rostral-most aspect or olfactory region of the brain is severed with a scalpel, and cranial nerves are cut as necessary to free the brain as it is extracted (Figure D). During removal of the brain, it is easiest to tilt the head up (caudodorsal direction) so that the brain falls away from the skull under its own weight. Avoid handling the brain with instruments as this destroys the nervous tissue and creates histological artifact.

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Figures 4.34A–C   Burmese python, Python molurus bivittatus. Pythonidae. Segments of the spinal cord may be collected by this simple technique. The intercostal and hypaxial muscles are cut with a knife on either side of the segment(s) to be sampled. A saw is used to make a transverse cut into the intervertebral space to the level of, but not breaching, the spinal canal (Figure A). The dorsal side of the cut segment is placed on the edge of a table or other surface and downward pressure is applied to disarticulate the vertebrae and expose a segment of spinal cord (Figure B). Using a scalpel, transect the spinal cord and nerve roots to remove the section (Figure C). Avoid pulling or bending the spinal cord and do not inadvertently squeeze the tissue with fingers or instruments.

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Figures 4.35A–B   Burmese python, Python molurus bivittatus. Pythonidae. A second method for spinal cord removal is to perform a dorsal laminectomy using a Stryker saw or Dremel tool. The epaxial muscles are reflected laterally to expose the dorsal aspects of the vertebrae. Cuts are made on either side of the dorsal processes to the level of the spinal canal (Figure A). These cuts are angled slightly toward the midline. Rongeurs are used to cut away the dorsal aspect of the vertebrae, thus exposing the spinal cord (Figure B). The spinal nerves are severed and the cord is carefully removed.

Figures 4.36A–B   Burmese python, Python molurus bivittatus. Pythonidae. To remove the globe and preserve the overlying spectacle, cuts are made in the periorbital skin, as shown (A). Carefully dissect around the eye, severing extraocular muscles, the optic nerve, and associated soft tissue (B). Place the specimen in fixative and avoid excessive handling after removal.

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Figure 4.37   Green turtle, Chelonia mydas. Cheloniidae. The coelomic cavity is exposed by incising the skin around the entire plastron and cutting through the marginal bridges. In sea turtles, the marginal bridges are not ossified and are easily cut with a knife, as shown. The pectoral musculature is reflected cranially and the ventral pelvis is removed to expose the caudal coelom. The trachea (T), esophagus (E), heart (H), both liver lobes (LV), stomach (S), intestine (I), urinary bladder (UB), and cloaca (CL) are visualized.

Figure 4.38A–B   Forest turtle, Heosemys spinosa. Emydidae. For most hard-shelled turtles and tortoises, the marginal bridges, which attach the plastron to the carapace, must be cut with a saw. A Stryker saw is commonly used for this purpose and cuts are made as indicated (blue dashes) (Figure A). As in Figure 4.37, the musculature of the forelimbs, hindlimbs, and associated skeletal girdles are reflected or excised to view the coelomic cavity (Figure B). The trachea (T), esophagus (E), heart (H), liver (LV), stomach (S), intestine (I), and cloaca (CL) are visible.

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Figure 4.39   Loggerhead sea turtle, Caretta caretta. Cheloniidae. The paired thymus (lobules of yellow or white tissue) is located in the cervical (neck) region and is associated with the carotid arteries.

Figure 4.40   Loggerhead sea turtle, Caretta caretta. Cheloniidae. Chelonians have a single (unpaired) thyroid gland (black arrow) located cranial to the base of the right and left aortas.

Figure 4.41   Loggerhead sea turtle, Caretta caretta. Cheloniidae. The apex of the ventricle is attached to the pericardium by the gubernaculum cordis (black arrow).

Figure 4.42   Loggerhead sea turtle, Caretta caretta. Cheloniidae. As in other reptiles, the spleen (S) and pancreas (P) typically are closely associated. Nodules of accessory splenic tissue may be seen within the pancreas (black arrows).

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Figure 4.43   Green turtle, Chelonia mydas. Cheloniidae. After the liver and gastrointestinal tract are removed. The lungs (L), right aorta (RA), left aorta (LA), dorsal aorta (DA), adrenal glands (AG), gonads (immature ovaries [O] in this example), and kidneys (K) remain for examination and sampling.

Figure 4.44   Loggerhead sea turtle, Caretta caretta. Cheloniidae. The kidneys (K) are within the retrocoelomic space and are covered by a thick white layer of connective tissue and perirenal adipose (fat). The reproductive tract lies ventral to the kidneys. This example is an immature female (ovaries [O]). The adrenal glands (AG) are ventromedial to the cranial poles of the kidneys and are fused in this species.

Figure 4.45   Loggerhead sea turtle, Caretta caretta. Cheloniidae. A hierarchy of developing and regressing follicles are present in this ovary and include large yellow vitellogenic follicles (VF), shrunken vitellogenic follicles undergoing atresia (AF), white developing follicles of various sizes (DF), and postovulatory follicles (PO) from the most recent clutch.

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Figure 4.46A–D   Loggerhead sea turtle, Caretta caretta. Cheloniidae. To remove the brain of a large chelonian, remove the eyes and cut away the muscles of mastication using a knife or scalpel (Figure A). Cut through the dorsal bones of the skull, as shown, using a cutting instrument such as a Stryker saw or Dremel tool (Figure B). Make a second cut adjacent to midline. To make this cut, angle the blade away from midline as demonstrated (i.e., do not cut perpendicular to the skull) (Figure C). Using a rigid flat instrument, such as a screw driver, remove the cut section (Figure D).

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Figure 4.46E–H   Next, make an additional cut through the junction of the supraoccipital and parietal bones to remove the second half of the cut section of skull, which is still firmly attached. Cut parallel to the surface of the skull as shown (Figure E) and remove the second half of the cut section (Figure F). Rongeurs may be required for this step. At this stage, the fibrous dura mater (blue arrow) covering the olfactory region of the brain is exposed. Next, cut off the caudal portion of the supraoccipital bone (black arrows) (Figure F). Observe that this portion of the bone is cut away in Figure G. As demonstrated in Figure G, make a cut from the foramen magnum to the rostral aspect of the brain case. Repeat this cut on the other side. Be careful when making these cuts as it is easy to accidentally cut into the cranial vault and damage the brain. Remove the cut section of skull using rongeurs and a rigid flat instrument as needed to expose the brain. Carefully cut through and reflect the dura mater to expose the olfactory nerves (ON), cerebrum (CE), cerebellum (CB), and brainstem (BS) (Figure H). Note the location of the salt glands (SG) in sea turtles. To remove the brain, cut the olfactory nerves with a scalpel and tilt the skull back on its base so that gravity lifts the brain from the skull. Sever the cranial nerves, beginning with the optic nerves, to free the brain as it is extracted.

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Figure 4.47A–B   Making touch impressions. Grasp the tissue to be sampled and blot the cut surface onto tissue until there is no imprint left behind (Figure A). Hold the tissue cut side up and gently touch the glass slide to the cut surface several times to make rows of individual impressions (Figure B). Make a minimum of three slides. Fix and stain accordingly.

Figure 4.48   Proper tissue collection for formalin fixation. Samples to be preserved in formalin should be 0.5 cm or less in thickness to achieve proper fixation. Large organs, such as the liver (shown here), are serially sectioned with a knife or scalpel to allow complete inspection of the tissue and to obtain samples of the appropriate thickness. A sharp blade and good cutting surface are essential.

Figure 4.49   Tissue cassettes. To avoid losing small samples, tissues may be placed into tissue cassettes prior to being placed into a formalin container. Sponges (center) or microcassettes (right) are available for very small tissues. Do not place large sections of tissue into cassettes as they will be damaged and may not fix adequately. There should be plenty of space around the sample and the lid should close easily without contacting the sample (left).

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Figures 4.50A–C   Sear and stab technique. First, a new scalpel blade is heated under a flame source. The flat surface of the blade is then applied to the surface of the organ or tissue to be cultured, thus searing the outside (Figure A). Next, the blade is stabbed through the seared area (light brown) into the target tissue (Figure B). A sterile culturette is inserted deep into the incision, avoiding contact with the surface tissue (Figure C). Always handle scalpel blades and culturettes in a sterile manner.

5 Host Response to Infectious Agents and Identification of Pathogens in Tissue Section Brian A. Stacy and Allan P. Pessier

Contents

5.1 Introduction

5.1 Introduction........................................................... 257 5.2 Reptilian Leukocytes and Macrophages............... 258 5.2.1 Acidophils.................................................. 258 5.2.2 Heterophils................................................ 258 5.2.3 Eosinophils................................................ 258 5.2.4 Basophils and Mast Cells.......................... 259 5.2.5 Monocytes, Macrophages, Azurophils....... 259 5.3 Distribution of Lymphoid and Hematopoietic   Tissue .................................................................... 260 5.4 The Inflammatory Response................................. 260 5.4.1 Gross Appearance of Exudates ............... 260 5.4.2 Time Course of Inflammatory Processes...260 5.4.3 Temperature Effects on Inflammatory   Responses.................................................. 261 5.4.4 Granuloma Formation............................... 261 5.5 Proliferative Host Responses ............................... 262 5.5.1 Cryptosporidiosis....................................... 262 5.5.2 Viral Diseases and Proliferative Lesions ... 262 5.5.3 Proliferative Osteoarthritis and   Osteoarthrosis in Squamates.................... 263 5.6 Lesions and Tissue Deposits Associated with   Inflammation.......................................................... 263 5.6.1 Tissue Responses Secondary to   Inflammation............................................. 263 5.6.2 Splendore-Hoeppli Reaction..................... 264 5.6.3 Amyloid-like Material................................ 264 5.6.4 Immune Complex-Associated   Glomerulonephritis................................... 264 5.7 Detection of Infectious Agents in Tissue Section...264 5.7.1 Viral Infections.......................................... 264 5.7.2 Bacterial Infections.................................... 265 5.7.3 Fungal Infections....................................... 266 5.7.4 Parasites in Tissue Section........................ 266 5.8 Immunohistochemistry/In Situ Hybridization...... 266 References......................................................................... 267

The knowledge base concerning the immune system and inflammatory responses of reptiles is dwarfed by that of fish, birds, and even amphibians. Current information on reptilian inflammatory responses is based on a small number of experimental studies, as well as observational studies of naturally occurring disease (Huchzermeyer and Cooper, 2000; Mateo et al., 1984a; Montali, 1988; Smith et al., 1988a; Smith et al., 1988b; Soldati et al., 2004; Tucunduva et al., 2001). While components of both innate and adaptive immune systems have been identified, much of the information is morphologic or descriptive and few mechanistic studies have been published (Cooper et al., 1985; Mead and Borysenko, 1984; Pasmans et al., 2001; Ramaglia et al., 2004; Sypek and Borysenko, 1988; Sypek et al., 1984; Zarkadis et al., 2001;). Detailed information on acute phase reactants and cytokines are increasingly available for fish and birds and perhaps will stimulate similar studies for reptiles (Bayne and Gerwick, 2001; Kaiser et al., 2004; Laing and Secombes, 2004; Qureshi et al., 2000; Schultz et al., 2004; Secombes et al., 2001). The understanding of inflammatory responses in reptiles has been hampered by incomplete and sometimes contradictory published information. For instance, the function of important circulating leukocytes such as the azurophil and eosinophil are unknown, although there is often a presumption of homologous function to morphologically similar cells in other vertebrate classes. Furthermore, the different perspectives and terminology of zoologists, hematologists, and veterinarians and the dangers inherent in broad generalizations applied to a diverse class of animals can lead to confusion and misinterpretation. Thus, critical reading and interpretation of the literature are essential. For more detailed information on circulating inflammatory cells, see Chapter 3.

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The emphasis of this chapter is on reptilian host responses to infectious agents as observed by the diagnostic pathologist in histologic section. It is intended to provide a review of the published literature supplemented by the experience of the authors and their colleagues and to serve as a basis for more specific investigation of inflammatory processes. Immunology and hematology are discussed in greater detail in Chapters 2 and 3, respectively.

5.2 Reptilian Leukocytes and Macrophages 5.2.1 Acidophils Acidophil is a collective term used for leukocytes with prominent eosinophilic cytoplasmic granules and includes both heterophils and eosinophils (Montali, 1988). In some publications, reptilian acidophils are described as a single lineage and termed type I and type II eosinophils on the basis of their resemblance to mammalian eosinophils (Azevedo and Lunardi, 2003; Montali, 1988; Sypek and Borysenko, 1988). This terminology has led to confusion in the literature as information on heterophils and eosinophils is reported interchangeably (Sypek and Borysenko, 1988). At present, morphologic and cytochemical evidence suggests that heterophils and eosinophils represent separate distinct cell lines (Alleman et al., 1992; Azevedo and Lunardi, 2003; Garner et al., 1996; Harr et al., 2001; Mateo et al., 1984(b); Montali, 1988; Sypek and Borysenko, 1988). In blood films, heterophils have abundant fusiform cytoplasmic granules that can usually be distinguished from the round cytoplasmic granules of eosinophils. Differentiation based on morphology alone can be complicated by variability in heterophil granule morphology, especially in snakes. Because processing for histology distorts the morphology of cytoplasmic granules, differentiation of heterophils and eosinophils is usually possible only in stained blood films and cytological preparations of bone marrow or exudates. They cannot be reliably distinguished in histologic section (Garner et al., 1996; Montali, 1988). Thus by convention, most pathologists refer to acidophils in histologic section as heterophils unless proven otherwise. In birds, techniques for cytochemical differentiation of eosinophils from heterophils in histologic section are described (Lam, 2001; Maxwell, 1984). These methods are not validated for reptiles and may be difficult to apply because of peroxidase activity in some squamate heterophils and azurophils (Harr et al 2001; Montali, 1988; Tucunduva et al., 2001).

5.2.2 Heterophils The heterophil is the functional equivalent of the mammalian neutrophil and is identified histologically in a wide variety of reptilian inflammatory reactions. Heterophils are among the first cells recognized at sites of inflammation and

have been demonstrated to be phagocytic for bacteria and foreign materials (Montali, 1988; Mateo et al., 1984a; Sypek and Borysenko, 1988; Jacobson et al., 1997). Cytochemically, heterophils are negative for benzidine peroxidase in chelonians, crocodilians, and some squamates (Alleman et al., 1992; Alleman et al., 1999; Mateo et al., 1984b). In other squamates, heterophils are positive for benzidine peroxidase (Harr et al., 2001; Montali, 1988; Tucunduva et al., 2001). It has been suggested that this peroxidase activity represents combined actions of heterophils and eosinophils into one cell, especially because the eosinophil is either absent or rare in many squamates, particularly snakes (Montali, 1988). Other authors hypothesize that peroxidase activity in heterophils of the green iguana (Iguana iguana) represents bactericidal capabilities and oxidative responses comparable to mammalian neutrophils, rather than that of avian or other reptilian heterophils (Harr et al., 2001). Nonoxidative mechanisms of antimicrobial activity, such as beta-defensins found in avian heterophil granules, have not yet been described for reptilian heterophils (Harmon, 1998). In histologic section, nondegenerate heterophils have abundant brightly eosinophilic cytoplasmic granules and a round to lobated nucleus (Figure 5.1). Occasionally, acidophils (presumptive heterophils) with brown or golden cytoplasmic granules that can resemble hemosiderin pigment are observed (Montali, 1988) (Figure 5.2). In our experience, these cells may be seen adjacent to heterophils with typical orange-red (i.e., eosinophilic) tinctorial properties, and are cytochemically negative for iron and melanin pigments. The reason for the tinctorial differences in these golden heterophils is unknown. Degenerate and degranulated heterophils can resemble macrophages and care should be taken to distinguish these cells in acute inflammatory reactions (Figure 5.3) (Montali, 1988).

5.2.3 Eosinophils Eosinophils are commonly observed in peripheral blood films from chelonians and crocodilians, but their occurrence is variable in squamates (Montali, 1988; Azevedo and Lunardi, 2003; Mateo et al., 1984b; Alleman et al., 1992; Alleman et al., 1999; Raskin, 2000). Among the squamates, eosinophils are most frequently observed in some lizard species and only rarely in snakes (Harr et al., 2001, Raskin, 2000, Salakij et al., 2002). Identification of eosinophils in peripheral blood is often based on morphologic criteria, especially round eosinophilic to blue-green cytoplasmic granules, and to a lesser extent, pale blue cytoplasmic coloration (Montali, 1988; Raskin, 2000; Hawkey and Dennett, 1989). With the exception of some lizards (Harr et al., 2001), eosinophils of most species are cytochemically positive for benzidine peroxidase (Azevedo and Lunardi, 2003; Mateo, 1984b; Alleman et al., 1992). As noted previously, eosinophils cannot be reliably differentiated from heterophils in histologic section because tissue processing distorts granule morphology.

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In mammals, the presence of eosinophils in inflammatory lesions is usually associated with parasitic infections or TypeI hypersensitivity reactions. Although these associations have been extrapolated to birds and reptiles (Frye, 1991; Mader, 2000), experimental evidence and well-documented accounts of natural disease are inconclusive regarding the function of the eosinophil in nonmammalian vertebrates (Montali, 1988; Fudge and Joseph, 2000). Development of techniques to detect reptilian eosinophils in histologic section may help clarify the role of this cell in inflammatory reactions.

5.2.4 Basophils and Mast Cells Basophils are observed in the peripheral blood from all of the major reptilian groups and can be the predominant circulating leukocyte in some chelonians (Raskin, 2000; Sypek and Borysenko, 1988) (Figure 5.4). Morphologically, reptilian basophils and mast cells are similar to those of other species and are characterized by purple metachromatic granules that fill the cytoplasm and can obscure the nucleus. Typically, mast cell granules are much smaller that those of basophils. Mast cells are frequently observed in connective tissues of the mesentery, tongue, gastrointestinal tract, heart, and skeletal muscle (Sottovia-Filho, 1973) (Figure 5.5). Giemsa and toluidine blue stain mast cell granules dark purple and can be useful for detecting these cells. Although the function of reptilian basophils and mast cells is unknown, studies in snapping turtles (Chelydra serpentina) have shown that basophils have surface immunoglobulins and release histamine upon degranulation in a manner similar to that described in mammals (Sypek et al., 1984).

5.2.5 Monocytes, Macrophages, and Azurophils Monocytes resembling those observed in mammals and birds can be observed in all major reptilian groups (Alleman et al., 1999; Raskin, 2000; Tucunduva et al., 2001). In snakes, the predominant circulating cell of presumed monocytic origin is the azurophil, which is morphologically characterized by numerous fine cytoplasmic granules. Monocytes without cytoplasmic granules can also occur in snakes; however when present, these cells are a minor component of the differential cell count (Dotson et al., 1995; Alleman et al., 1999; Lamirande et al., 1999; Tucunduva et al., 2001). Conversely, monocytes with small numbers of cytoplasmic granules (azurophils; azurophilic monocytes) are observed in a variety of species, such as the desert tortoise and green iguana (Alleman et al., 1992; Harr et al., 2001), but are distinguished from azurophils of snakes by the absence of benzidine peroxidase activity (Heard et al., 2004). Granuloma formation is extremely common in reptiles (Montali, 1988); thus macrophages are a prominent component of most inflammatory reactions studied in histologic section. Morphologically, reptilian macrophages are similar

to those in other vertebrate classes, and while commonly mononuclear (Figure 5.6), they also form multinucleated giant cells (Figure 5.7). Macrophages are phagocytic and in turtles [red-eared slider (Trachemys scripta)], peritoneal macrophages exhibit bacteriocidal respiratory burst activity (Pasmans et al., 2001). As compared to other vertebrates, there is little specific information on reptilian macrophages in terms of cytokine production and immune system regulation (Klasing, 1998; Qureshi et al., 2000).

5.2.5.1 Azurophils  Azurophils are circulating leukocytes unique to the squamates, especially snakes (Montali, 1988). They are mononuclear, and as their name implies, possess a myriad of fine azurophilic cytoplasmic granules. As with other reptilian leukocytes, there is confusion regarding the origin of the azurophil. Azurophils are strongly benzidine peroxidase positive, which has drawn cytochemical comparisons to both mammalian neutrophils and monocytes (Alleman et al., 1999). In older publications, some authors classify azurophils as granulocytes akin to the mammalian neutrophils, or as different stages of the reptilian heterophil (Cooper et al., 1985). Currently, most authors consider the azurophil to be of the monocyte lineage and functionally and morphologically distinct from granulocytes, such as heterophils. The function of azurophils in inflammatory reactions requires clarification at the tissue level. Although azurophilia is frequently observed in association with inflammatory conditions, azurophils usually are not identifiable in histologic section. An experimental study in Brazilian boas (Boa constrictor constrictor) demonstrated heterophils, azurophils, and benzidine peroxidase–negative monocytes in peripheral blood, but only heterophils and macrophages were observed on tissue-implanted coverslips processed in a manner similar to blood films (Tucunduva et al., 2001). If azurophils are functioning as monocytes, tissue fixation and processing may obscure azurophil cytoplasmic granules, and, subsequently, only cells identifiable as macrophages are recognized in histologic section. Alternatively, azurophils may undergo morphologic changes during tissue migration with loss of cytoplasmic granules. Additional studies are needed to determine the origin of azurophils and investigate possible transformation to tissue macrophages. 5.2.5.2 Melanomacrophages  Melanomacrophages are unique melanin-producing cells of apparent macrophage origin found in fish, amphibians, and reptiles (Agius and Roberts, 2003; Gyimesi and Howerth, 2004). These cells are components of the systemic mononuclear phagocyte system and their size and number are variable among reptile species. In reptiles, melanomacrophages are most prominent as discrete aggregates (melanomacrophage centers) in the liver and to a lesser degree in the spleen, kidney, and at sites of chronic inflammation (Figures 1.45, 1.46, 1.48–1.51, 5.8, 6.36–6.38). Melanomacrophages are most abundant in chelonians and can be scant in some squamates. Most of the

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published information available on these cells originates from work performed in fish and amphibians.   Melanomacrophages synthesize melanin (Scalia et al., 1988) and frequently contain other pigments such as hemosiderin, lipofuscin, and ceroid (Agius and Roberts, 2003). The melanin pigments in melanomacrophages may potentiate or facilitate neutralization of free radicals, which are produced during catabolic states or may be involved in the synthesis of bacteriocidal compounds. The hypothesized role of melanin as a free radical scavenger is supported by the inverse relationship between hepatic melanin content and superoxide dismutase (SOD) activity in reptiles (Sichel et al., 1987). The SOD activity of chelonian livers, which typically have the greatest amount of melanin, is much lower than that of examined lizard species. In some cases, melanomacrophages have been shown to function at low temperatures and this may be of adaptive value for host defense in heterothermic animals (Johnson et al., 1999). Melanomacrophages are phagocytic for erythrocytes, microorganisms, and other debris, and should be closely examined for infectious agents. It has been suggested that melanomacrophage centers may be involved in antigen processing in a fashion analogous to germinal centers in the lymph nodes of mammals (Ferguson, 1976). An increase in the size of melanomacrophage centers (melanomacrophage hypertrophy) is a frequent nonspecific finding in reptilian diagnostic pathology (Figure 5.9). This change is often referred to or diagnosed as melanomacrophage hyperplasia; however, it is unclear whether size increase represents an increase in individual melanomacrophage size or an increase in melanomacrophage number (hyperplasia) (Gyimesi and Howerth, 2004). Evaluation of melanomacrophages is subjective and accurate assessment requires comparison with a healthy animal of the same species, sex, and physiologic state. In some cases, melanomacrophage hypertrophy can be marked and associated with significant hepatocellular atrophy. In these cases, black discoloration of the liver may be apparent on gross examination. Factors that may contribute to melanomacrophage hypertrophy include seasonal variation, debilitation, emaciation, stress and chronic inflammation (Agius and Roberts, 2003; Christiansen et al., 1996). Finally, melanomacrophage centers may be extensively involved in chronic bacterial diseases, such as mycobacteriosis. In these cases, melanomacrophages exhibit hypertrophy with increasing proportions of lightly or nonpigmented cells (Figure 5.10).

5.3 Distribution of Lymphoid and Hematopoietic Tissue The principle lymphoid organs of most reptiles are the thymus and spleen (Cooper et al., 1985; Sypek and Borysenko, 1988) (see Chapter 2). Reptiles lack lymph nodes, but aggregates of lymphoid cells are common in a variety of tissues, especially

the gastrointestinal tract and to a lesser degree, the lungs, urinary bladder, kidney, and pancreas (Figure 5.11). Hematopoiesis principally occurs in the bone marrow (Figures 2.1–2.3), but significant extramedullary hematopoiesis (EMH) can be observed in locations such as the liver (portal and subcapsular regions), spleen, and thymus. Acidophilic cell lines (heterophil and eosinophil) are often seen as prominent foci in areas of extramedullary hematopoiesis and can resemble inflammatory heterophilic infiltrates (Figure 5.12). Care should be taken not to misinterpret small focal lymphoid or hematopoietic cell aggregates in a variety of tissues as an inflammatory process. Other criteria for inflammatory processes such as tissue injury, presence of a vascular response and extensive infiltration of tissue structures may assist in distinguishing EMH and normal lymphoid aggregates from inflammation.

5.4 The Inflammatory Response 5.4.1 Gross Appearance of Exudates In contrast to the liquefied or creamy suppurative exudate (pus) derived from neutrophils in mammals, inflammatory exudates derived from heterophils in reptiles form solid aggregates or clumps (Figures 5.13–5.14) (Montali, 1988; Huchzermeyer and Cooper, 2000). These exudates, especially in subcutaneous abscesses, can be distinctive on section and are characterized by concentric layers of solid material (Figure 5.15). Differences in the gross appearance of exudates between reptiles and mammals have been attributed to differences in hydrolytic enzyme activity or lack of proteases in heterophils (Montali, 1988). Solid exudates must be differentiated from areas of tissue necrosis or organized fibrin. Accumulations of keratin, such as that observed with squamous metaplasia of the ophthalmic glands and middle ear (aural abscesses) of turtles with hypovitaminosis A, must also be differentiated from solid exudates (Brown et al., 2004; Elkan and Zwart, 1967) (Figure 5.16).

5.4.2 Time Course of Inflammatory Processes Initial manifestations of inflammatory processes in reptiles are similar to those observed in mammals and include congestion of venules and capillaries as well as fibrin exudation. The initial inflammatory cell observed both experimentally and in cases of spontaneous disease is the heterophil (Figure 5.17) followed by monocytes or macrophages. A small number of experimental studies in reptiles provide some information about the temporal progression of inflammatory lesions. In Brazilian boas (Boa constrictor constrictor) surgically implanted with cotton thread and American alligators (Alligator mississippiensis) injected with turpentine, heterophils were present as early as 4 hours following tissue injury (Mateo et al., 1984a; Tucunduva et al., 2001). By histologic examination, significant numbers of both heterophils and macrophages were present

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within one day. By seven days, inflammatory reactions were organized into discrete concentrically arranged aggregates (granulomas), which in American alligators were characterized by central, densely packed necrotic heterophils surrounded by macrophages. In both Brazilian boas and American alligators, multinucleated giant cells were observed as early as 7 days. Neovascularization and fibroplasia were observed at 14 days in American alligators, but not at 15 days in Brazilian boas. In garter snakes (Thamnophis sirtalis) with experimental skin wounds held at 30°C ambient temperature, active fibroblasts could be observed as early as 5 days following injury and were common by 10 days (Smith et al., 1988b).

5.4.3 Temperature Effects on Inflammatory Responses As with other important physiologic processes (Chapter 1), inflammation and immune responses in reptiles may be affected by ambient temperature (see Chapter 2 for temperature effects on immune response). Behavioral fever is when sick ectotherms increase their body temperature by seeking external heat sources, presumably in order to increase efficiency of the host response to infectious agents (Vaughn et al., 1974; Muchlinski et al., 1999). This behavior may effectively mimic fever induced by endogenous inflammatory mediators in homeotherms. Evidence for endogenous pyrogens has also been described (Bernheim and Kluger, 1977). Temperature may influence the speed, composition, and duration of the inflammatory response (see Chapter 2). In desert iguanas (Dipsosaurus dorsalis) experimentally inoculated with Aeromonas hydrophila, tissue migration of granulocytes into injection sites was increased in animals held at 41°C compared to those held at 35°C and 38°C (Bernheim, 1978). In another study, lower temperatures (13.5°C and 21°C) did not affect the character and intensity of the inflammatory response at 2 days in garter snakes (Thamnophis sirtalis) with experimental skin wounds (Smith et al., 1988b). Snakes held at higher temperatures (30°C), however, had earlier resolution of regional inflammation by 3 to 6 weeks. In the same study, active fibroblasts were observed in wounds as early as 5 days in snakes housed at higher temperatures, but were delayed to 3 weeks or more in snakes held at 13.5°C. There is some limited evidence that leukocyte function is influenced by temperature. The respiratory burst of peritoneal macrophages of red-eared sliders (Trachemys scripta) was suppressed at lower temperatures (Pasmans et al., 2001). Also, histamine release from the basophils of the snapping turtle (Chelydra serpentina), in contrast to mammals, occurred over a wide temperature range of 10°C to 27°C and increased with rising temperature (Sypek et al., 1984).

5.4.4 Granuloma Formation The hallmark of many inflammatory reactions in reptilian species is granuloma formation, which is a response to

a wide variety of bacterial, fungal, and parasitic infections (Figures 5.18–5.19). There are two principle types of granuloma, the heterophilic granuloma and the histiocytic granuloma (Figures 5.20–5.21), and each has a different proposed pathogenesis (Montali, 1988). Morphologically, the granuloma types are distinguished by the composition of the necrotic center, which consists of degranulated and necrotic heterophils in the heterophilic granuloma and degenerate macrophages in the histiocytic granuloma (Figures 5.22A–C). Differentiation of the granuloma type can sometimes be helpful diagnostically and provides a useful model for understanding host responses to infectious agents. A third type of granuloma, the chronic granuloma, represents an end-stage lesion formed initially from either a heterophilic or histiocytic granuloma (Montali, 1988; Soldati et al., 2004) (Figure 5.23).

5.4.4.1 Heterophilic Granulomas  Heterophilic granulomas are inflammatory lesions associated with extracellular pathogens, including most bacterial and fungal infections, or with tissue injuries that result in infiltrates of large numbers of heterophils. Formation of a heterophilic granuloma (Figures 5.24A–C) begins with a localized accumulation of heterophils at a site of infection or injury (heterophilic abscess) (Montali, 1988). Heterophils at the center of the abscess degranulate and become necrotic with a transition to viable intact heterophils at the edges of the lesion (Figure 5.25). Subsequently, macrophages accumulate at the periphery, thus forming the heterophilic granuloma (Figures 5.26A–D). In American alligators with experimentally induced lesions, structures consistent with heterophilic granulomas were formed as early as 7 days following tissue injury and included features such as small numbers of multinucleated giant cells (Mateo, 1984a). Lymphoid cell infiltrates and peripheral fibrosis, which are common in mammalian granulomas, are features of chronicity and usually are not present unless the heterophilic granuloma is transitioning into a chronic granuloma (Montali, 1988). In some instances, macrophages can be numerous in heterophilic granulomas, making distinction of granuloma types difficult (Figure 5.27). If detectable, the inciting agents of heterophilic granulomas, such as bacterial colonies or fungal hyphae, are usually observed within the central aggregates of necrotic heterophils. The formation of heterophilic granulomas is not limited to solid tissues, but can also occur within the lumen of hollow structures such as airways, or at the surface of the skin or mucous membranes (Figures 5.28–5.29). The mechanism of heterophilic granuloma formation has been likened to a foreign body reaction in which the degenerate heterophils or components of heterophil granules incite the granulomatous response (Montali. 1988). A similar mechanism has been discussed for the formation of eosinophilic granulomas in mammals (Fairley, 1991). Because reptilian exudates do not liquefy, it has been suggested that heterophilic granuloma formation may be a means of dispersing these lesions.

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5.4.4.2 Histiocytic Granulomas  In contrast to heterophilic granulomas, histiocytic granulomas are usually associated with obligate intracellular bacteria such as mycobacteria or chlamydiae, but may also be observed with fungal infections (Jacobson et al., 2002; Montali, 1988; Soldati et al., 2004). Formation of histiocytic granulomas (Figures 5.30A–C) begins with accumulation and organization of macrophages at the site of infection. As the lesions progress or enlarge, there may be necrosis of the central macrophages with formation of a necrotic core (Figures 5.31A–C). Frequently, small to moderate numbers of heterophils may be observed infiltrating between peripheral macrophages and within the necrotic core (Figure 5.32). In most instances, the number of heterophils and amount of central necrotic debris are less than that observed in heterophilic granulomas. 5.4.4.3 Chronic Granulomas  The persistence of either heterophilic or histiocytic granulomas eventually leads to transition into chronic granulomas. Macrophages, which are frequently epithelioid, predominate in chronic granulomas and are encompassed by fibrous connective tissue and variable numbers of lymphocytes and plasma cells (Figure 5.33). Chronic granulomas frequently contain central amorphous debris, which can have a prominent laminated appearance and in some circumstances may become mineralized (Figures 5.34–5.35). It is often difficult or impossible to determine the original mechanism of granuloma formation in longstanding chronic granulomas. A recent review of naturally occurring granulomatous inflammation in reptiles determined that the majority (58/90) of observed granulomas were the chronic form (Soldati et al., 2004).

5.5 Proliferative Host Responses While granuloma formation is the most common response to infectious agents in reptiles, some important pathogens incite proliferative lesions and diffuse inflammatory cell infiltrates. This section describes the most common infectious diseases associated with these types of responses.

5.5.1 Cryptosporidiosis A classic example of a proliferative lesion in reptiles is hypertrophic gastritis of snakes with cryptosporidiosis. This condition is characterized by marked hyperplasia of gastric surface epithelium and mucous neck cells with variable submucosal infiltrates of lymphocytes, plasma cells, and heterophils (Brownstein et al., 1977) (Figures 5.36–5.37; Figures 12.49, 12.65–12.66, 12.69). Epithelial proliferation is also observed with Cryptosporidium sp. infection associated with aural polyps in green iguanas and enteritis in leopard geckos (Eublepharis macularis) (Terrell et al., 2003; Uhl et al., 2001). Interestingly, gastric cryptosporidiosis in lizards can present either as a proliferative lesion similar to snakes or as an atrophic condition with atrophy of granular cells (Frost et al., 1994; Oros et al., 1998).

5.5.2 Viral Diseases and Proliferative Lesions 5.5.2.1 Paramyxoviruses and Reoviruses  Paramyxovirus (PMV) and reovirus infections are an important cause of proliferative pneumonia in squamates. These viruses are associated with pronounced hyperplasia of respiratory epithelial cells (Type-II pneumocytes) and variable diffuse interstitial infiltrates of heterophils, lymphocytes, plasma cells and macrophages (Jacobson et al., 1992; Jacobson et al., 1997; Jacobson et al., 2001; Lamirande et al., 1999) (Figures 5.38– 5.39, 9.86–9.90). Epithelial proliferation may be the predominant finding in some cases with minimal inflammation. In addition, paramyxovirus infection in some snakes is associated with pancreatitis and florid ductular hyperplasia resembling pancreatic carcinoma; however, the role of PMV in the pathogenesis of this lesion has not been definitively established (Jacobson et al., 1992; Ratcliffe, 1943; Rebecca Papendick personal communication, 2004) (Figures 5.40, 9.91, 9.134– 9.136). It should be noted that respiratory epithelial proliferation, while suggestive of viral infection, is not pathognomonic, and other differentials, including chronic nonviral pneumonia (bacterial and parasitic) and toxic injury should be considered (Homer et al., 1995) (Figures 5.41–5.42, 12.36). Confirmation of viral infection requires molecular diagnostics, such as PCR or in situ hybridization (see Chapter 7), or electron microscopy (see Chapter 6). Other documented causes of proliferative pneumonia in reptiles include inclusion body disease (IBD) in snakes, respiratory tract chlamydophilosis in Burmese pythons (Bodetti et al., 2002), upper respiratory tract mycoplasmosis in tortoises caused by Mycoplasma agassizii (Brown et al., 1999; McLaughlin et al., 2000) and pulmonary intranuclear coccidiosis in chelonians (Garner et al., 2006). 5.5.2.2 Herpesviruses  Some herpesviruses of reptiles are associated with epithelial or fibroepithelial proliferation. Perhaps the most well-known example is fibropapillomatosis of sea turtles associated with chelonid fibropapilloma–associated herpesvirus (C-FP-HV). Tumors associated with C-FP-HV have various proportions of epithelial and fibrous components; however, epithelial proliferation is usually present to some degree in most cases at the histologic level (Herbst et al., 1999). Proliferation of fibroblasts of the papillary dermis is a consistent finding in cutaneous tumors, and acanthosis, orthokeratotic hyperkeratosis, epithelial degeneration and basilar cleft formation are common epithelial changes (Figures 5.43A–B, 9.17). Visceral tumors associated with CFP-HV typically arise from the fibroblastic component. Viral inclusion bodies are only present in a minority of cases. Viral particles resembling herpesvirus have also been observed in cutaneous papillomas of European green lizards (Lacerta viridis) (Cooper et al., 1982). (Figures 9.37–9.38). Another example of a proliferative lesion associated with herpesviral infection is stomatitis associated with Varanid herpesvirus-1, which was documented in four green tree monitors (Varanus prasinus) (Wellehan et al., 2005) (Figure 9.41). Lesions

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were described as chronic proliferative gingivitis. Epithelial cells were tightly packed or formed villous projections and there was involvement of associated salivary ducts. Interestingly, two animals were also diagnosed with, or subsequently developed, oral squamous cell carcinomas. An example of epidermal proliferation associated with a nonviral agent is dermatophilosis caused by Dermatophilus congolensis (Figures 10.30–10.43).

5.5.3 Proliferative Osteoarthritis and Osteoarthrosis in Squamates Proliferative lesions of the vertebral column and ribs characterized by new bone formation and proliferation of fibrocartilaginous-osseous tissue or matrix are common in snakes. Three general forms are encountered: localized new bone formation secondary to osteomyelitis and segmental vertebral lesions with distortion of bone architecture, degenerative joint changes, and ankylosis of adjacent vertebra with or without infection and inflammation (Figures 5.44, 10.126–10.130). The latter two forms have distinctive radiographical and pathological features in snakes and warrant further discussion and investigation of pathogenesis. The clinical, pathological, and microbiological findings in 15 snakes with segmental vertebral lesions have been reviewed (Isaza et al., 2000). Histologic lesions were categorized as active bacterial osteoarthritis, noninflammatory osteoarthrosis with foci of chronic inflammation, and noninflammatory osteoarthrosis without concurrent inflammation (Figures 10.128–10.130). Bone and blood culture from cases of noninflammatory osteoarthrosis were negative, and cases of active bacterial osteoarthritis yielded Salmonella spp. in most instances. It was hypothesized that these categories represent a progression of bacterial vertebral osteomyelitis that ultimately results in ankylosis and vertebral remodeling with resolution of inflammation. In another study of Arizona ridgenose rattlesnakes (Crotalus willardi), a high incidence of osteomyelitis was associated with infection by S. arizonae serotype 56:Z4,Z23 (Ramsay et al., 2002). In this cohort, bone lesions were inflammatory and often progressive, and where characterized by granulomatous osteomyelitis, sequestra formation, and pathologic fractures. From these studies, many proliferative vertebral lesions of snakes are associated with osteomyelitis and positive bone or blood cultures for Salmonella sp. or, less commonly, other bacteria, including Gram-positive species (Isaza et al., 2000; Ramsay et al., 2002). Other cases have florid bone or fibrocartilaginous-osseous proliferation with minimal or no proximate inflammation. The latter finding suggests that other factors, such as primary degenerative disease, changes in the blood flow, and circulating inflammatory mediators, may be involved. Although there have been excellent observational studies of these lesions, experimental or mechanistic studies are needed to investigate pathogenesis and progression of proliferative vertebral disease in snakes.

Vertebral lesions in snakes have been reported as or compared to osteitis deformans or Paget’s bone disease (PBD) in humans by some investigators (Frye and Charney, 1974; Kiel, 1977). This term has disseminated widely among reptile enthusiasts and veterinarians and is broadly applied to a variety of vertebral lesions. Paget’s bone disease in humans, however, has specific clinical, radiographical, and pathological diagnostic criteria, and presents in three phases of progression: the initial lytic phase, the mixed lytic and sclerotic phase, and the final sclerotic phase (Rosenburg, 1999). In reptiles, there are no reports of such a progression or other evidence to support any substantial similarity between vertebral lesions of snakes and PBD of humans. Some investigators have focused on the appearance of a mosaic pattern of cement or reversal lines, which is a feature of human pagetoid bone during the later phases (Frye and Charney, 1974; Kiel, 1977). However, reversal lines, which are the result of bone resorption and deposition, are prominent in some healthy snakes and may appear mosaic in areas of normal bone remodeling (Figure 5.45). In addition, vertebral ankylosis, which is rare in human PBD cases, is very common in snakes (Saifuddin and Hassan, 2003). In the authors’ opinion, the use of the term Paget’s bone disease in reptiles is a misnomer that confuses the clinical and pathological picture and should be avoided. Vertebral lesions of lizards are not well studied, and thus any similarities to the pathogenesis of, or associated findings in, similar lesions of snakes are not known. Segmental vertebral lesions of lizards that resemble those of snakes are seemingly less common. However, proliferative osteoarthropathy with ankylosis is relatively common in the tails and lumbosacral vertebrae of large saurians, especially green iguanas (Iguana iguana). The pathogenesis of these lesions requires further study. Possible underlying causes of lizard vertebral lesions encountered by the authors include repetitive trauma, degenerative joint disease, and underlying metabolic disease.

5.6 Lesions and Tissue Deposits Associated with Inflammation 5.6.1 Tissue Responses Secondary to Inflammation It is important to be cautious when extrapolating information about inflammation and host response across animal taxa; however, there are many examples of tissue responses in reptiles that are akin to well-known responses in birds and mammals. Metaplastic, hyperplastic, and fibrotic changes secondary to inflammation are frequently observed in reptiles and should be recognized as such. One key example is secondary squamous metaplasia of mucosal epithelia, which is a response to chronic irritation and inflammation  (Figure 5.46). In the presence of inflammation, it is impor-

264  Host Response to Infectious Agents and Identification of Pathogens in Tissue Section

tant to differentiate secondary squamous metaplasia from that of hypovitaminosis A (Elkan and Zwart, 1967).

5.6.2 Splendore-Hoeppli Reaction In mammals, some types of bacterial and fungal infections result in deposition of characteristic eosinophilic, clubshaped material that radiates around the infectious agent. This response is known as the Splendore-Hoeppli reaction and the deposits may be comprised of precipitated immunoglobulins. Splendore-Hoeppli material comprises the socalled sulfur granules, which are a gross feature of infections that elicit this reaction. In reptiles, there is a single report of material morphologically resembling the Splendore-Hoeppli reaction in association with Neisseria sp. infections in rhinoceros iguanas (Cyclura cornuta) and green iguanas (see Chapter 10) (Plowman et al., 1987) (Figure 10.17).

5.6.3 Amyloid-like Material Amyloid is a heterogeneous insoluble proteinaceous substance composed of insoluble fibrils that appear in tissues as deposits of eosinophilic hyaline material (amyloidosis). In animals, the most common form of amyloidosis is secondary amyloidosis associated with chronic inflammation, whereby the tissue deposits are derived from the acute phase protein serum amyloid A (SAA). Serum amyloid A has been identified in salmonid fish (Jorgensen et al., 2000), but little is known of this protein in reptiles. Although common in mammals and birds, there are only rare reports of amyloidosis in reptiles or other lower vertebrates (Cosgrove and Anderson, 1984; Cowan, 1968; Mashima et al., 1997). Published descriptions of amyloidosis-like conditions in reptiles are brief and do not describe traditional diagnostic criteria for the detection of amyloid, such as birefringence with Congo Red stain (CR), florescence with thioflavin-T, or observation of typical amyloid fibrils by transmission electron microscopy. Thus, the occurrence of amyloid in reptiles, as defined for other vertebrates, is not established in the peer-reviewed literature. Tissue deposits of substances that resemble amyloid on routine hematoxylin and eosin–stained sections that do not react with CR have been observed in a variety of reptiles and termed paramyloid (Richard J. Montali, personal communication). The authors have examined a single case of paramyloid by transmission electron microscopy and determined the tissue deposit to consist of collagen fibrils (Figures 5.47A–B). Careful examination of other putative cases of amyloidosis in reptiles is warranted.

5.6.4 Immune Complex–Associated Glomerulonephritis Glomerulonephritis associated with the deposition of immune complexes within glomerular capillary loops is a well-recognized lesion in mammalian pathology. Lesions consist of

variable degrees of capillary loop thickening and glomerular hypercellularity. Characterization of these glomerular changes frequently requires techniques such as special histochemical stains, transmission electron microscopy, and immunofluorescence to differentiate them from changes caused by diseases without an immunologic basis such as diabetes mellitus. Although a variety of histologic changes has been described in reptilian glomeruli (Cosgrove and Anderson, 1984; Cowan, 1968; Zwart, 1964), none to our knowledge has been investigated beyond light microscopy (Figure 5.48).

5.7 Detection of Infectious Agents in Tissue Section 5.7.1 Viral Infections Morphologic features of viruses visible using electron microscopy and viral diseases of reptiles are discussed in much greater detail in Chapters 6 and 9, respectively. Molecular tools for identifying viruses in tissues and cell culture are discussed in Chapter 7. The host response to viral infection can range from necrotizing as observed with some adenovirus and herpesvirus infections to proliferative as observed for ophidian paramyxovirus infection (Jacobson et al., 1997), herpesvirusassociated fibropapillomas in green sea turtles (Herbst et al., 1999), and stomatitis in green tree monitors (Wellehan et al., 2005). The primary lesions induced by viruses can often be obscured by intense inflammatory responses associated with secondary bacterial infections. In histologic section, the presence of characteristic inclusion bodies is one indicator of some types of viral infection. Viral inclusions may be intranuclear, as with herpesvirus or adenovirus infections, (Figures 5.49–5.50, 9.2, 9.8, 9.10, 9.23–9.24, 9.26–9.27, 9.29, 9.32, 9.35, 9.39, 9.42–9.43, 9.45–9.46, 9.48, 9.50) or intracytoplasmic as with paramyxovirus, poxvirus, or iridovirus infections (Jacobson et al., 1981; Marschang et al., 1999; Stauber and Gogolewski, 1990) (Figures 5.51, 9.56, 9.62, 9.64, 9.69–9.74). While useful for diagnosis, the presence of inclusion bodies can be variable and depends on the type of virus, the presence of secondary bacterial infection, and the duration of illness. For instance, cytoplasmic inclusion bodies are not consistently present in cases of respiratory paramyxovirus infection of snakes and iridovirus infections of turtles (Jacobson et al., 1992; Marschang et al., 1999; DeVoe et al., 2004), and are observed only in some stages of herpesvirus-associated papillomatosis of sea turtles (Herbst et al., 1999) (Figure 9.24). Some cellular components can resemble and must be distinguished from viral inclusion bodies. Common structures that may be mistaken for inclusion bodies include cytoplasmic invaginations, large nucleoli in cells of hyperplastic or regenerative tissues and large eosinophilic cytoplasmic inclusions composed of intermediate filaments (Figures 5.52–5.53, 6.14–6.25). The latter are most often observed in the cyto-

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plasm of cutaneous epithelial cells. Also, the inclusions associated with intranuclear coccidiosis in tortoises may be easily confused with the intranuclear inclusions of herpesviruses (Figures 5.54, 12.34–12.38). In these cases, transmission electron microscopy or other techniques may be required for identification.

5.7.2 Bacterial Infections Bacterial infections in reptiles are reviewed in Chapter 10. Bacterial infections are usually characterized by heterophilic or chronic granulomas in histologic section. Exceptions include infections with organisms that have an intracellular component to their pathogenesis such as Mycobacterium sp. and Chlamydophila sp., and the diffuse inflammatory responses associated with organisms such as Mycoplasma sp. and Helicobacter sp. (Busch et al., 2002; McLaughlin et al., 2000) (Figures 5.55–5.56, 10.57–10.66). In chronic granulomas associated with bacterial infections, the organism number may be very low and the etiologic agent difficult to detect in histologic section. Tissue Gram stains may be useful in detecting or better characterizing intralesional bacteria (Figures 5.57–5.58). Common tissue Gram stains include the Brown and Brenn (BB), Brown and Hopps (BH), and Goodpasture’s methods. The BB stain favors Gram-positive organisms whereas the BH favors Gram-negative organisms. When interpreting tissue Gram stains, care should be taken not to mistake the blue staining of intralesional heterophil granules for Gram-positive bacteria (Figure 5.57). Stains such as Giemsa and a variety of silver-based stains (e.g., Steiner’s and Warthin-Starry) are nonspecific and can be useful for screening histologic sections for bacteria or for demonstrating unusual organisms such as Helicobacter sp.

5.9.2.1 Mycobacterium  Mycobacterial infections in reptiles are most often characterized by the formation of histiocytic or chronic granulomas (Montali, 1988; Soldati et al., 2004). However, the histologic appearance of these infections can vary widely. Mycobacterial lesions range from distinct granulomas to diffuse histiocytic infiltrates and occasionally manifest as heterophilic granulomas (Figures 5.59–5.62, 10.25–10.27). In a series of 23 reptilian granulomas associated with mycobacteria, 14 granulomas were classified as chronic, 8 as histiocytic, and 1 as heterophilic (Soldati et al., 2004). It is not known why mycobacteria elicit heterophilic granuloma formation in some cases. The authors have observed heterophilic granulomas in cases with extracellular organisms, similar to some cases of atypical mycobacteriosis in mammals (Appleyard and Clark, 2002; Raymond et al., 2000). Differences in the distribution of organisms and the character of the inflammatory response may be influenced by host factors or the species of mycobacteria. This occasional variation in granuloma morphology may limit solely using

granuloma type as a guide in differential diagnosis (Soldati et al., 2004).  In most instances, an acid-fast stain is required to visualize mycobacteria in histologic section, and usually a ZiehlNeelson acid-fast stain is sufficient. Occasionally, the species of mycobacteria causing reptilian mycobacteriosis are better demonstrated with alternative techniques such as Fite’s acid-fast stain (Figure 5.63). Infrequently, and more often in cases with high numbers of organisms, mycobacteria can be observed in hematoxylin and eosin-stained sections or as Gram-positive bacilli (Appleyard and Clark, 2002) (Figure 5.63). The number of organisms observed in reptilian mycobacterial granulomas can vary from scant, requiring careful and prolonged examination of tissue sections, to abundant, resembling classic cases of avian mycobacteriosis (Figures 5.64–5.65, 10.26–10.27). In impression smears and cytologic preparations, a characteristic appearance of mycobacteria is negative staining of bacilli within the cytoplasm of macrophages (Figure 5.66). Differentials for acidfast bacteria in histologic section and cytologic preparations include Nocardia sp., Rhodococcus sp., and Legionella sp. (Bacciarini et al., 1999; Bentz et al., 2000; Echeverri et al., 2001). Occasionally cellular debris and cellular products such as lipofuscin need to be differentiated from acid-fast bacilli in histologic section.

5.9.2.2 Chlamydophila  Chlamydophila spp. are obligate intracellular bacterial pathogens and are associated with either histiocytic or chronic granulomas in snakes (Jacobson et al., 2002; Soldati et al., 2004). Also, some gastrointestinal tract lesions in emerald tree boas (Corallus caninus) consisted of mixed infiltrates of lymphocytes, plasma cells, heterophils, and macrophages (Jacobson et al., 2002). In contrast, chlamydiosis in green sea turtles (Chelonia mydas) is primarily associated with necrotizing lesions and diffuse inflammatory infiltrates of acidophils, macrophages and lymphocytes (Bodetti et al., 2002; Homer et al., 1994) (Figures 10.68– 10.69). Similarly, acute or subacute chlamydiosis in juvenile Nile crocodiles (Crocodylus niloticus) results in hepatitis with necrosis and lymphoplasmacytic infiltrates (Huchzermeyer, 2003). Blepharo-conjuctivitis is observed in crocodiles with chronic chlamydial infections (Huchzermeyer, 2003). Chlamydial organisms can be difficult to visualize or unapparent on hematoxylin and eosin–stained sections. In some cases, granulomas may contain central amphophilic to basophilic granular inclusions (Jacobson et al., 2002; Bodetti et al., 2002) that correspond to developmental stages of Chlamydophila (Figures 5.67, 10.76). A modified Macchiavello or Gimenez stain may help demonstrate organisms in histologic section; however, care should be taken to distinguish putative organisms from acidophil granules or artifactual staining (Homer et al., 1994). Immunohistochemistry using a monoclonal antibody that recognizes chlamydial lipopolysaccharide (Homer et al., 1994; Jacobson et al., 2002; Bodetti et al.,

266  Host Response to Infectious Agents and Identification of Pathogens in Tissue Section

2002; Soldati et al., 2004) (Figures 10.77, 10.81–10.83) is a more specific approach to diagnosis in histologic section.

5.7.3 Fungal Infections For detailed information on fungal (mycotic) diseases of reptiles see Chapter 11. As with bacterial infections, the early response to most fungal infections involves heterophils with subsequent formation of heterophilic granulomas (Figure 5.68). In some instances, histiocytic or chronic granulomas are observed (Figure 5.69). Significant necrosis is associated with some fungal lesions, especially with cutaneous infections (Jacobson, 1980 and Nichols et al., 1999). Necrosis can also be a feature in fungal infections with vascular invasion and subsequent thrombosis. Fungi can be observed in hematoxylin and eosin–stained sections, but are easily overlooked when numbers are small or if large amounts of necrotic debris are present. A variety of special stains are available for visualizing fungi in histologic section with the most common being periodic acid-schiff (PAS) and Gomori methenamine silver (GMS) (Figures 5.70–5.71). Although the species of fungus cannot be definitively determined on the basis of morphology in histologic section, there are frequently morphologic clues that can aid in narrowing differential diagnoses or assessing the significance of fungal culture results (Chandler et al., 1980). Ascomycete fungi, such as Aspergillus sp. and Fusarium sp., have hyphae that are septate with parallel walls and dichotomous branching (Figure 5.72). In contrast, Zygomycete fungi, such as Mucor sp. or Basidiobolus sp., have hyphae that are aseptate (or infrequently septate) and broad or bulbous with variable hyphal diameters (Figure 5.73). Other fungi have distinctive forms that can be recognized in tissue section. For instance, the fungi that cause chromomycosis or phaeohyphomycosis have brown pigmented sclerotic bodies or hyphae that usually are easily recognized in section. The Chrysosporium anamorph of Nannizziopsis vriesii, a recently recognized cutaneous pathogen of lizards, snakes and crocodilians, produces distinctive arthroconidia (Nichols et al., 1999; Pare and Sigler, 2002) (see Chapter 11). Fungi that classically cause systemic infections in mammals, including Cryptococcus neoformans and Coccidioides immitis, are rare in reptiles and have characteristic features, such as distinctive thick mucopolysaccharide coats (C. neoformans) or endosporulation (C. immitis) (Figure 5.74) (McNamara et al., 1994; Timm et al., 1988).

5.7.4 Parasites in Tissue Section Chapter 11 reviews parasitic infections and infestations of reptiles and provides some diagnostic features of parasites in tissue section. The host response to parasites is widely variable depending on the type of organism, location within the host, and immune status. As noted earlier, the role of eosinophils in reptilian inflammatory responses to metazoan parasite infections is still unclear, although acidophils (heterophils or eosin-

ophils) are a component of many early host responses. For encysted cestodes or nematodes, lesions may be composed of relatively inactive chronic granulomas lined by attenuated (flattened) macrophages (Figures 5.75–5.76). Spirorchiid fluke eggs in aquatic turtles are often surrounded by layers of multinucleated giant cells with fewer numbers of other inflammatory cells, and form within vessels or perivascular tissue (Gordon et al., 1998) (Figures 5.77, 12.114–12.116). Some metazoan parasites, such as ascarids and acanthocephalans, can cause extensive necrotizing lesions due to parasite migration or secondary bacterial infection. In certain cases others, such as pentastomids, may not elicit a significant response by the host. Infections with protozoal organisms such as microsporidia or Hepatozoon are associated with areas of necrosis (Figures 12.82, 12.85), predominantly macrophage infiltrates, or with histiocytic granulomas (Jacobson et al., 1998 and Wozniak et al., 1998) (Figure 5.78). Infections with coccidian parasites, including Cryptosporidium sp., intranuclear coccidia (Garner et al., 2006), and enteric coccidia frequently cause epithelial proliferation or necrosis and regeneration of injured epithelial cells (Figure 5.79). Inflammatory infiltrates in response to coccidian parasites are usually mixtures of lymphocytes, plasma cells, and macrophages, and have a diffuse distribution. Infections with other protozoa, such as the amoeba Entamoeba invadens, cause predominantly necrotizing lesions (Kojimoto et al., 2001) (Figures 5.80–5.81, 12.12–12.21). Recognition of metazoan parasites in tissue section is relatively straightforward, whereas protozoa can be difficult to identify and distinguish from host structures or other pathogens. For example, mucus blebs or extrusion of small cells can occasionally mimic Cryptosporidium sp. (Figure 5.82) and trophozoites of Entamoeba invadens can be confused with macrophages (Figure 5.81). A variety of histologic stains can assist in diagnosis of protozoal infections (Gardiner et al., 1988). Gram’s stains will stain microsporidia Gram-positive (Figures 5.83, 12.85). The periodic acid-Schiff (PAS) stain will detect polar granules of microsporidia and can help highlight other protozoa, such as trophozoites of Entamoeba invadens (Figures 5.81, 12.18, 12.22). Acid-fast stains will stain some mature spores of microsporidia and myxozoa, as well as the sporozoites of Sarcocystis species (Figure 5.83). Although modified acid-fast stains are commonly used for detecting Cryptosporidium oocysts in fecal smears, the use of acid-fast stains in histologic section for detection of Cryptosporidium sp. is rarely successful in our experience.

5.8 Immunohistochemistry and In Situ Hybridization Immunohistochemistry (IHC) and in situ hybridization (ISH) are techniques that demonstrate antigens (IHC) or nucleic acid sequences (ISH) of specific infectious agents in paraf-

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fin-embedded histologic sections (see Chapter 7 for further details). These techniques use either polyclonal or monoclonal antibodies (IHC) or DNA probes (ISH) linked to enzyme-based chromagens that enable localization of the signal to individual cells or regions within affected tissues (Figure 5.84). When applied and interpreted correctly, the major advantages of IHC and ISH are rapid, specific, and sensitive detection of infectious agents using only histologic sections, and the ability to directly associate infectious agents with histologic lesions. The use of histologic sections is especially important in reptilian diagnostics because in many instances only paraffin-embedded tissues are available for examination. Reagents specific for some reptilian pathogens, such as respiratory paramyxovirus or chelonian herpesvirus (Homer et al., 1995; Origgi et al., 2003; Sand et al., 2004), have been described. However, some antibodies are available only in a few research laboratories and are not widely accessible for general use as diagnostic tools. For other pathogens, such as Chlamydophila (Homer et al., 1994; Jacobson et al., 2002) or Cryptosporidium sp., genus-specific reagents are widely available in veterinary diagnostic laboratories for use in domestic animals and are easily applied to reptilian diagnostics. Interpretation of immunohistochemistry for infectious agents in reptilian samples is usually straightforward, but some nonspecific background staining can be observed because of endogenous peroxidase activity in reptilian leukocytes or spurious binding to other tissue components (Homer et al., 1994; Homer et al., 1995; Jacobson et al., 2002). Tissue fixation and processing, such as prolonged formalin fixation, can affect the outcome of IHC and ISH and should be considered when interpreting results. IHC and ISH are used increasingly in the diagnosis of disease in reptiles; however accurate interpretation requires experience and familiarity with histology and particular antibodies and probes.

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Terrell SP, Uhl EW, and Funk RS. 2003. Proliferative enteritis in leopard geckos (Eublepharis macularius) associated with Cryptosporidium sp. J Zoo Wildl Med 34: 69–75. Timm KI, Sonn RJ, and Hultgren BD. 1988. Coccidioidomycosis in a Sonoran gopher snake, Pituophis melanoleucus affinis. J Med Vet Mycol 26:101–104. Tucunduva M, Borelli P, and Silva JRMC. 2001. Experimental study of induced inflammation in the Brazilian boa (Boa constrictor constrictor). J Comp Path 125:174–181. Uhl EW, Jacobson E, Bartick TE, and Micinilio J, Schmidt R. 2001. Aural-pharyngeal polyps associated with Cryptosporidium infection in three iguanas (Iguana iguana). Vet Pathol 38:239–242. Vaughn LK, Bernheim HA, and Kluger MJ. 1974. Fever in the lizard Dipsosaurus dorsalis. Nature 252:473–474. Wellehan JFX, Johnson AJ, Latimer KS, Whiteside DP, Crawshaw GJ, Detrisac CJ, Terrell SP, Heard DJ, Childress A, and Jacobson ER. 2005. Varanid herpesvirus 1: a novel herpesvirus associated with proliferative stomatitis in green tree monitors (Varanus prasinus). Vet Micro 105:83–92.

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Figure 5.1  Carpet python, Morelia spilota. Pythonidae. Photomicrograph of bone marrow. Numerous heterophils comprise an area of granulopoiesis. Note the abundant eosinophilic granules. H&E stain.

Figure 5.2  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of lung with acute heterophilic pneumonia. Typical heterophils with eosinophilic granules are intermixed with “golden” heterophils characterized by light brown granules. H&E stain.

Figure 5.3  Malaysian giant turtle, Orlitia borneensis. Bataguridae. Photomicrograph of lung with acute heterophilic pneumonia. Degenerate heterophils have lost their granules and may be mistaken for macrophages. Note that a few cells contain some remaining granules (arrow), which identify them as heterophils. H&E stain.

Figure 5.4  Spiny turtle, Heosemys spinosa. Emydidae. Photomicrograph of spleen with circulating granulocytosis. Moderate numbers of basophils (purple granules) and heterophils (red granules) are circulating through the spleen. Giemsa stain.

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Figure 5.5  Dumeril’s boa, Acrantophis dumerili. Boidae. Photomicrograph of liver. Mast cells (arrow) are within connective tissvue surrounding a bile duct. Note the fine basophilic (purple) granules. Giemsa stain.

Figure 5.6  Mugger crocodile, Crocodylus palustris. Crocodylidae. Photomicrograph of lung with chronic bacterial bronchopneumonia. Abundant macrophages are aggregated within an airway and are interspersed with fewer heterophils. H&E stain.

Figure 5.7  Mugger crocodile, Crocodylus palustris. Crocodylidae. Photomicrograph of lungs with chronic bacterial pneumonia and granuloma formation. Multinucleated giant cells are forming at the margin of a chronic granuloma. H&E stain.

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Figure 5.8  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of normal melanomacrophage centers within the liver. H&E stain.

Figure 5.9  Desert tortoise, Gopherus agassizii. Testudinidae. Liver: Melanomacrophage hyperplasia and hepatocellular pigment accumulation. Note the enlargement of the melanomacrophage centers as compared with Figure 5.8. H&E stain.

Figure 5.10  Fork-nosed chameleon, Furcifer minor. Chamaeleonidae. Photomicrograph of the liver showing melanomacrophage hyperplasia (mycobacteriosis). Note the progressive increase in sparsely or nonpigmented cells within the melanomacrophage centers. H&E stain.

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Figure 5.11  Western rattlesnake, Crotalus viridis. Viperidae. Photomicrograph of normal esophageal tonsil. H&E stain.

Figure 5.12  Western rattlesnake, Crotalus viridis. Viperidae. Photomicrograph of the spleen showing granulopoiesis. This area of granulopoiesis (granulocyte or heterophil production) is distinguished from true inflammation by the absence of other features of inflammation, such as infiltration of normal structures, cellular injury or degeneration, or vascular response (congestion, edema, fibrin deposition). H&E stain.

Figure 5.13  Wood turtle, Glyptemys (formerly Clemmys) insculpta. Emydidae. Transverse section of the head through the middle ear revealing chronic, unilateral, exudative otitis media (aural abscess). The left middle ear cavity is dilated and the wall is thickened by white fibrous connective tissue. Note the plug of tan exudate within the lumen and compare the affected side with normal right ear.

Figure 5.14  Caiman lizard, Dracaena guianensis. Teiidae. Exudative and proliferative pneumonia caused by a paramyxovirus infection. The exudate within the airways forms clumps and small aggregates rather than the liquefied suppurative material formed by mammals.

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Figure 5.15  American alligator, Alligator mississippiensis. Alligatoridae. Pedal abscess. The typical laminated appearance of reptile exudates is demonstrated in this chronic abscess on the palmar surface of the foot.

Figure 5.16  Wood turtle, Glyptemys (formerly Clemmys) insculpta. Emydidae. Photomicrograph of the middle ear seen in Figure 5.13. There is chronic, unilateral, exudative otitis media (aural abscess). This illustrates the laminated or multilayered appearance of the exudate. The exudate is a mixture of degenerate heterophils and exfoliated keratin (inset). The latter was produced by squamous metaplasia of the middle ear lining. H&E stain.

Figure 5.17  Water monitor, Varanus salvator. Varanidae. Photomicrograph of the lung showing bacterial embolus with acute heterophilic pneumonia (acute septicemia). Heterophils infiltrate the pulmonary septum and surround an embolus of bacteria (central purple material) in this example of an acute inflammatory response. H&E stain.

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Figure 5.18  Blood python, Python curtus. Pythonidae. Small intestine with chronic granulomatous enteritis (unknown etiology). A common appearance of chronic granulomas is a single tan or gray nodule or multinodular cluster as seen in this image. Note the extension of the granulomas through the wall and mucosa. Courtesy of Rebecca Papendick.

Figure 5.19  California red-sided garter snake, Thamnophis sirtalis infernalis. Colubridae. Chronic granulomatous dermatitis and myositis (Gram-positive bacteria) of the tail. The dermis and underlying tissue are expanded and effaced by multiple coalescing chronic granulomas. The granulomas are comprised of central tan necrotic material surrounded by lighter bands of fibrous connective tissue.

Figure 5.20  Caiman lizard, Dracaena guianensis. Teiidae. Photomicrograph the lung with heterophilic granuloma formation (mycobacteriosis). A heterophilic granuloma is comprised of a central zone of intact and degenerate heterophils surrounded by variable numbers of macrophages and infiltrating heterophils. H&E stain.

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Figure 5.21  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of the liver with histiocytic granuloma formation (mycobacteriosis). Histiocytic granulomas are comprised of aggregates of macrophages with minimal or no infiltration by heterophils. In this case, the hepatocytes around the granuloma have intracytoplasmic inclusion bodies (boid inclusion body disease). H&E stain.

Figure 5.22A–C  Photomicrographs of different types of granulomas. The center of the heterophilic granuloma (Figure A) is comprised of intact and degenerate heterophils. In contrast, the histiocytic granuloma (Figure B) is comprised of macrophages throughout with occasional heterophils, degenerate cells, and a small central focus of mineralization. The chronic granuloma (Figure C) consists of dense, sparsely cellular material and is the end-stage progression of both heterophilic and histiocytic types. H&E stain.

Figure 5.23  Viperid species. Viperidae. Photomicrograph of a chronic granuloma (etiology unknown) within the small intestine. Chronic granulomas are characterized by central eosinophilic, sparsely cellular, or acellular necrotic material surrounded by a layer of macrophages. H&E stain. Courtesy Rolando Quesada.

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Figure 5.24A–D.   Heterophilic granuloma formation. Heterophilic granulomas begin as aggregations of heterophils in response to an inflammatory stimulus, often an extracellular pathogen (Figure A). Heterophils undergo degeneration (Figure B) and become surrounded by macrophages, thus forming a heterophilic granuloma (Figure C). As the granuloma becomes chronic, the central heterophils become increasingly degenerate and leave behind a central mass of eosionophilic necrotic material (Figure D). (Adapted from Montali RJ. 1988. J Comp Path 99:1–26. With permission.)

Figure 5.25  Nile crocodile, Crocodylus niloticus. Crocodylidae. Photomicrograph of a heterophilic granuloma within the thymus. Note the transition from intact heterophils at the periphery of the granuloma to degenerate heterophils within the center. The macrophages in this example have a prominent vacuolated appearance. H&E stain.

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Figures 5.26A–D  Dumeril’s boa, Acrantophis dumerili. Boidae. Photomicrograph of the kidney with chronic interstitial nephritis and heterophilic granuloma formation (presumptive septicemia). Heterophils aggregate within the renal interstitium (Figure A) and macrophages accumulate as the lesions progress (Figure B) forming multinucleated giant cells (Figure C). In more advanced granulomas, the central heterophils become progressively degenerate (Figure D). H&E stain.

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Figure 5.27  California red-sided garter snake, Thamnophis sirtalis infernalis. Colubridae. Photomicrograph of chronic granulomatous dermatitis and myositis (Gram-positive bacteria) within the body wall. This is an example of histiocyte- or macrophage-rich heterophilic granuloma. Note bacterial colonies (purple) within the central zone of heterophils. H&E stain.

Figure 5.28  Mugger crocodile, Crocodylus palustris. Crocodylidae. Photomicrograph of the lung with a heterophilic granuloma due to a mixed bacterial infection. This granuloma is forming within an airway. Note the central bacterial colonies (purple) surrounded by heterophils and multinucleated giant cells. H&E stain.

Figure 5.29  Komodo dragon, Varanus komodoensis. Varanidae. Photomicrograph showing chronic heterophilic and histiocytic coelomitis with multinucleated giant cell formation (septic coelomitis secondary to yolk leakage). The heterophilic exudate along the body wall is bordered by a band of palisading multinucleated giant cells. Note the similarity of the cellular response and organization to that of heterophilic granulomas formed within tissue. H&E stain.

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Figure 5.30A–C  Progression of a histiocytic granuloma. Macrophages (histiocytes) aggregate in response to an inflammatory stimulus, often an intracellular pathogen (Figure A). Macrophages within the center of the aggregate undergo necrosis (Figure B), which progresses to central areas of caseation (Figure C). (Adapted from Montali RJ. 1988. J Comp Path 99:1–26. With permission.)

Figures 5.31A–C  McGregor’s tree viper, Trimeresurus mcgregori. Viperidae. Photomicrograph of the spleen. Histiocytic granuloma formation (mycobacteriosis) is seen. The histiocytes form small aggregates (Figure A) and accumulate into distinct granulomas (Figure B). The central cells in the more advanced granulomas are undergoing necrosis and mineralization (Figure C). H&E stain.

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Figure 5.32  Kenya horned viper, Bitis worthingtoni. Viperidae. Photomicrograph of a histiocytic granuloma (mycobacteriosis) within the liver. Small numbers of infiltrating heterophils, as seen in this image, may be observed in histiocytic granulomas. H&E stain.

Figure 5.33  Emerald tree boa, Corallus caninus. Boidae. Photomicrograph of a chronic granuloma (unknown etiology) within the small intestine. This chronic granuloma is surrounded by dense bands of fibrous connective tissue and small numbers of lymphocytes. H&E stain.

Figure 5.34  Blood python, Python curtus. Pythonidae. Photomicrograph of a chronic granuloma (unknown etiology) within the small intestine. The central necrotic material has a prominent laminated appearance. H&E stain.

Figure 5.35  Emerald tree boa, Corallus caninus. Boidae. Photomicrograph of a chronic granuloma with mineralization (unknown etiology) within the small intestine. Mineralization is occasionally observed in the center of some reptile granulomas. H&E stain.

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Figure 5.36  Western rattlesnake, Crotalus viridis. Viperidae. Photomicrograph of normal stomach: Compare to Figure 5.37. H&E stain.

Figure 5.37  Gopher snake, Pituophis melanoleucus. Colubridae. Photomicrograph of the stomach with proliferative gastritis with mucous neck cell hyperplasia and surface cryptosporidia. Compare this image with Figure 5.36. Note the abundance of the mucosal neck cells and the diminution of gastric glands. H&E stain.

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Figures 5.38A–B  Western rattlesnake, Crotalus viridis. Viperidae. Photomicrograph of normal lung; compare to Figures 5.39A–B. H&E stain.

Figures 5.39A–B  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Photomicrograph of lung with proliferative and exudative pneumonia caused by ophidian paramyxovirus. The respiratory epithelium is severely hyperplastic and the faveolar air spaces are filled with inflammatory cells and exfoliated epithelium, which includes multinucleated syncytial cells. Compare to Figures 5.38A–B. H&E stain.

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Figure 5.40  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Photomicrograph of a splenopancreas. Pancreatitis with epithelial hyperplasia, syncytial cell formation, and intracytoplasmic inclusion bodies associated with ophidian paramyxovirus are seen. Note the multinucleated syncytia formation, anisocytosis, and anisokaryosis of the pancreatic epithelial cells. H&E stain.

Figure 5.41  Radiated tortoise, Geochelone radiata. Testudinidae. Photomicrograph of a lung revealing proliferative pneumonia with intranuclear protozoa (intranuclear coccidiosis). Protozoal zoites are visible within epithelial nuclei (inset). H&E stain.

Figure 5.42  Viperid species. Viperidae. Photomicrograph of a lung. Chronic exudative and proliferative pneumonia with intrafaveolar nematodes (Rhabdias sp.) (arrow) is seen. The pulmonary epithelium in this nonviral pneumonia is plump and cuboidal rather than the flat squamous epithelium of a normal lung. H&E stain.

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Figures 5.43A–B  Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph of a cutaneous fibropapilloma. Consistent features of herpesvirus-associated fibropapillomas are proliferation of the papillary dermis (Figure A) and degeneration and necrosis of cells within the basal epithelium (Figure B). H&E stain.

Figure 5.44  Carpet python, Morelia spilota. Pythonidae. Photomicrograph of a vertebral articular facet. Periosteal new bone formation and degeneration of articular cartilage are seen. This section is from a snake with exuberant new bone formation and spondylosis. Note the joint space (arrow) with abnormal cartilage in the lower half of the image. The new bone consists of a central core of woven bone that has been remodeled and replaced on the outer surface by eosinophilic parallel-fibered bone (inset). H&E stain. Figure 5.45  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of a normal vertebra. Prominent reversal lines (basophilic lines) (arrows) are a normal feature of reptilian vertebral bone. H&E stain.

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Figure 5.46  Wood turtle, Glyptemys (formerly Clemmys) insculpta. Emydidae. Photomicrograph of a middle ear. Normal mucosa (upper) and squamous metaplasia (lower) are seen. The squamous metaplasia in this case is unilateral and associated with a chronic inflammation (aural abscess), not hypovitaminosis A. H&E stain.

Figures 5.47A–B  Green anaconda, Eunectes murinus. Boidae. Photomicrograph of spleen. Atypical collagen deposition (amyloid-like appearance) is seen. The spleen contains coalescing deposits of homogenous eosinophilic material that resembles amyloid of mammals and birds (Figure A) (H&E stain.). The deposits stain intensely blue with Masson’s trichrome stain, which is consistent with collagen (inset). Transmission electron microscopy reveals that the deposits are comprised of atypical collagen fibers and are not consistent with amyloid (Figure B). Uranyl acetate and lead citrate stain. Courtesy of Dalen Agnew.

Figure 5.48  Wagler’s viper, Tropidolaemus wagleri. Viperidae. Photomicrograph of a kidney with glomerulonephritis (unknown etiology). The glomerular mesangium is expanded and hypercellular, and there is infiltration by leukocytes. The uriniferous space is dilated, and the surrounding tubules are separated by interstitial edema. H&E stain.

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Figure 5.49  Greek spurthighed tortoise, Testudo graeca ibera. Testudinidae. Photomicrograph of the tongue. Epithelial cells with intranuclear inclusion bodies (herpesvirus) and bacterial colonies are seen. H&E stain.

Figure 5.50  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of the small intestine with epithelial intranuclear inclusion bodies (consistent with adenovirus) (arrows). H&E stain.

Figure 5.51  Eastern box turtle, Terrepene carolina carolina. Emydidae. Photomicrograph of oral mucosa. Amphophilic intracytoplasmic inclusion bodies in epithelial cells (iridovirus) (arrows) are seen. H&E stain. Courtesy of April Johnson.

Figure 5.52  American alligator, A. mississippiensis. Alligatoridae. Photomicrograph of a bile duct within the liver. The round pale-staining structures (arrows) within the nuclei of the bile duct epithelium are invaginations of cytoplasm and may be mistaken for inclusion bodies. H&E stain.

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Figure 5.53  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of the trachea. Epithelial hyperplasia, intracellular edema, and necrosis secondary to fungal tracheitis are seen. This section is adjacent to an active inflammatory lesion. The nucleoli of some cells are very prominent (arrows) and may be mistaken for inclusion bodies. H&E stain.

Figure 5.54  Malagasy spider tortoise, Pyxis arachnoides brygooi. Testudinidae. Photomicrograph of pancreas. Necrotizing pancreatitis with intranuclear protozoa (intranuclear coccidia) are seen. Multiple pancreatic acinar cells have eosinophilic intranuclear inclusions (arrows) identified as protozoal zoites. These inclusions should be distinguished from those caused by herpesvirus or other viruses. H&E stain.

Figure 5.55  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of pericardial surface showing heterophilic and fibrinous pericarditis (Mycoplasma alligatoris). The pericardial surface (upper left) is covered by exudate comprised of fibrin and heterophils. H&E stain.

Figure 5.56  American alligator, Alligator mississippiensis. Crocodylidae. Photomicrograph of synovium. Heterophilic and histiocytic synovitis (Mycoplasma alligatoris) are seen. The synovial vessels are congested and the stroma is extensively infiltrated by heterophils and macrophages. Fibrin deposits are present on the surface. H&E stain.

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Figure 5.57  Viper species. Viperidae. Photomicrograph of stomach. Heterophilic gastritis with intralesional Gram-negative bacilli are seen. Note the red staining of the clusters of bacteria (arrow) within the center of the image. The granules of the surrounding heterophils stain purple and should not be mistaken for Gram-positive organisms. Granules of the gastric granular cells stain red (lower left). Brown and Brenn stain. Courtesy Rolando Quesada.

Figure 5.58  Dumeril’s boa, Acrantophis dumerili. Boidae. Photomicrograph of yolk sac. Yolk sacculitis with intralesional Grampositive bacilli (Clostridium sp.) are seen. Brown and Brenn stain.

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Figure 5.59  Rattlesnake, Crotalus sp. Viperidae. Photomicrograph of fat body. Heterophilic and histiocytic steatitis with intralesional acid-fast bacilli (mycobacteriosis) are seen. Heterophils predominate in this subacute mycobacterial lesion. H&E stain and Fite’s acid-fast (inset).

Figure 5.60  Mugger crocodile, C. palustris. Crocodylidae. Photomicrograph of lung with a histiocytic granuloma due to mycobacteriosis. H&E stain.

Figure 5.61  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of lung. There is diffuse histiocytic and lymphocytic pneumonia with intralesional acid-fast bacilli (mycobacteriosis). Contrast the diffuse character of the infiltrate with the formation of distinct granulomas in Figure 5.60. H&E stain and Fite’s acid-fast (inset).

Figure 5.62  Caiman lizard, Dracaena guianensis. Teiidae. Photomicrograph of lung with a heterophilic granuloma (mycobacteriosis). A heterophilic granuloma is comprised of a central zone of intact and degenerate heterophils surrounded by varying numbers of macrophages and infiltrating heterophils. H&E stain.

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Figure 5.63  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of lung. Histiocytic pneumonia with intracellular acid-fast; Grampositive bacilli (mycobacteriosis) are seen. The mycobacteria stain Gram-positive (Brown and Brenn, left image) and the acid-fast (Fite’s acid-fast, right image).

Figure 5.64  Kenya horned viper, Bitis worthingtoni. Viperidae. Photomicrograph of splenopancreas. There is a granulomatous splenitis with rare intrahistiocytic acid-fast bacilli (mycobacteriosis). This is an example of a paucimicrobial lesion. Fite’s acidfast stain.

Figure 5.65  McGregor’s tree viper, Trimeresurus mcgregori. Viperidae. Photomicrograph of splenopancreas. A chronic granuloma with intralesional acid-fast bacilli (mycobacteriosis) is seen. Numerous mycobacteria are within the central necrotic debris. Fite’s acid-fast stain.

Figure 5.66  Twin spotted rattlesnake, Crotalus pricei pricei. Viperidae. Photomicrograph of an impression smear of a granuloma from the heart. Macrophages with intracellular, negative-staining bacilli (mycobacteriosis). Wright-Giemsa stain. Courtesy of Rebecca Papendick.

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Figure 5.67  Emerald tree boa, Corallus caninus. Boidae. Photomicrograph of liver. A histiocytic granuloma with intralesional basophilic inclusions (chlamydophilosis) is seen. The large basophilic inclusions within macrophages can be a diagnostic feature of chlamydial granulomas. H&E stain.

Figure 5.68  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of trachea. A heterophilic tracheitis with intralesional fungal hyphae is seen. Heterophils are the predominant cell present and are interspersed with faint fungal hyphae (arrows). H&E stain.

Figure 5.69  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Photomicrograph of liver. A chronic granuloma with intralesional fungal hyphae is seen. Fungal elements are present within the central necrotic debris as well as the within the peripheral zone of intact macrophages. H&E stain.

Figure 5.70  Bushmaster, Lachesis muta muta. Viperidae. Photomicrograph of skin with a granulomatous dermatitis. Rare intralesional fungal hyphae (Fusarium sp.) are seen. Fungal elements can be rare and detection may require special stains and careful examination. H&E stain and Gomori methenamine silver stain (GMS) (inset).

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Figure 5.71  Cottonmouth, Agkistrodon piscivorus. Viperidae. Photomicrograph of a salivary gland. Granulomatous periadenitis with intralesional fungal hyphae is seen. Note the multinucleated cell formation in this histiocytic granuloma and the magenta color of the hyphae stained with periodic acid-Schiff (PAS) stain (inset). H&E stain. Courtesy of John Roberts.

Figure 5.72  Bushmaster, Lachesis muta muta. Viperidae. Photomicrograph of skin. Dermatitis with intralesional fungal hyphae (Fusarium sp.) is seen. The fungal hyphae have parallel walls and are septate with dichotomous branching. Gomori methenamine silver stain (GMS).

Figure 5.73  Common boa, Boa constrictor. Boidae. Photomicrograph of intestine. There is an ulcerative enteritis with intralesional fungal hyphae. The branching hyphae have nonparallel walls and rare septa. H&E stain.

Figure 5.74  Gopher snake, Pituophis melanoleucus. Colubridae. Photomicrograph of liver. Granulomatous hepatitis with intralesional fungal spherules of Coccidioides immitis is seen. The double contoured wall and endosporulation are characteristic features of C. immitis. H&E stain.

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Figure 5.75  Western rattlesnake, Crotalus viridis. Viperidae. Photomicrograph of mesentery. There is a chronic granuloma with an intralesional larval cestode. The wall is comprised of a narrow band of macrophages, fewer granulocytes, and a thin inner layer of fibrin. H&E stain.

Figure 5.76  Water monitor, Varanus salvator. Varanidae. Photomicrograph of lung. Bacterial embolus and a parasitic granuloma are seen. A bacterial embolus with surrounding heterophils is adjacent to a histiocytic granuloma with an intralesional nematode larva. H&E stain.

Figure 5.77  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of thymus. Intravascular spirorchiid eggs with granuloma formation are seen. H&E stain.

Figure 5.78  Bearded dragon, Pogona vitticeps. Agamidae. Photomicrograph of liver. Histiocytic hepatitis with intralesional protozoa (microsporidiosis) is seen. There is extensive infiltration of the liver by macrophages and fewer heterophils. A cluster of small pale-staining spores (arrow) is present in the center of the image. H&E stain.

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Figure 5.79  Nile crocodile, Crocodylus niloticus. Crocodylidae. Photomicrograph of small intestine. There is enteritis with epithelial hyperplasia, fusion of mucosal folds, and intralesional coccidia (inset). H&E stain.

Figure 5.80  Russell’s viper, Vipera russelli. Viperidae. Photomicrograph of liver. There is necrotizing hepatitis with intralesional trophozoites (Entamoeba invadens). An extensive area of necrosis borders viable hepatocytes (upper right). Rare trophozoites are present at the margins of the necrotic tissue. H&E stain. Courtesy John Roberts.

Figure 5.81  Russell’s viper, Vipera russelli. Viperidae. Photomicrographs of liver. There is necrotizing and histiocytic enteritis with intralesional trophozoites (Entamoeba invadens). Trophozoites can sometimes be mistaken for macrophages in tissue section. Note the typical appearance of a trophozoite with its small endosome (left). H&E stain. Periodic acid-Schiff (PAS) stains the trophozoites bright magenta and aids in their detection (right). Courtesy John Roberts.

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Figure 5.82  Nile crocodile, Crocodylus niloticus. Crocodylidae. Photomicrograph of an esophageal tonsil. Cell extrusion is seen within this normal esophagus. Cell extrusion sites and mucus blebs can resemble Cryptosporidium and other protozoa. H&E stain.

Figure 5.83  Bearded dragon, Pogona vitticeps. Agamidae. Photomicrograph of liver. There is a histiocytic hepatitis with intralesional protozoa (microsporidiosis). Microsporidia spores are acid fast (left) and Gram-positive (right). Fite’s acid fast and Brown and Brenn stains.

Figure 5.84  Sidewinder, Crotalus cerastes. Viperidae. Photomicrograph of stomach. Cryptosporidia line the mucosal surface and are detected by a specific monoclonal antibody (inset; Avidin biotin peroxidase complex method).

6 Identifying Reptile Pathogens Using Electron Microscopy Elliott R. Jacobson and Don A. Samuelson

Contents

6.1 General Comments

6.1 General Comments................................................ 299 6.2 Historical Perspectives........................................... 300 6.3 Electron Microscopy.............................................. 300 6.3.1 Positive Staining Transmission Electron   Microscopy (PSEM)................................... 300 6.3.2 Negative Staining Transmission Electron   Microscopy (NSEM)................................... 303 6.3.3 Scanning Electron Microscopy (SEM)...... 304 6.4 Collection of Samples............................................ 304 6.4.1 Blood Cells and Cell Cultures................... 304 6.4.2 Biopsies...................................................... 304 6.4.3 Postmortem Specimens............................. 305 6.4.4 Paraffin-Embedded Tissues...................... 305 6.4.5 Feces, Aspirates, and Washings for   Negative Staining Electron Microscopy.... 306 6.5 Identifying Pathogens in Cells and Tissues......... 306 6.5.1 Understanding Ultrastructure of   Normal Cells.............................................. 307 6.5.2 Viruses....................................................... 307 6.5.3 Bacteria.......................................................311 6.5.4 Parasites..................................................... 312 Acknowledgments............................................................ 313 References..........................................................................314

While certain pathogens can be identified or categorized into major groups using routine hematoxylin and eosin (H&E) staining or special staining of tissue sections, others require higher levels of magnification than provided by light microscopy. Electron microscopy (EM), particularly transmission electron microscopy (TEM), has increased magnification, superior cell preservation, and greater spatial resolution (1.5 to 2.0 nm) than light microscopy (Gondos et al., 1978; Hayat, 2000). Whereas TEM was originally developed to identify subcellular structures of normal tissues, in the 1960s it became a tool for use in basic virology and for understanding the pathogenesis of diseases caused by viruses (Wills, 1983). Electron microscopy has become an important diagnostic tool to identify cellular and subcellular abnormalities as well as certain pathogens in man and animals that could not be visualized using light microscopy (Gibbs et al., 1980). TEM has been particularly useful in identifying viruses, certain bacteria, and certain protozoan parasites in ultrathin sections of tissue. Using both gold-labeled monoclonal and polyclonal antibodies produced against specific organisms, a more accurate diagnosis can be made. Negative staining electron microscopy (NSEM) also has become a valuable diagnostic tool, particularly when looking for viruses and protozoa in fecal specimens of an animal being screened. While scanning electron microscopy (SEM) has some use in diagnosing infectious agents in animals, it has more value in demonstrating the location of certain pathogens that frequent the cell surfaces of tissues. Electron microscopic examination of tissue specimens from live and dead reptiles for the presence of pathogens can be a tedious and costly process. Using TEM and NSEM, searching for pathogens such as viruses in cell culture is far

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easier and quicker than in tissues, especially if cytopathic changes in cell culture indicate the presence of an infectious agent. Compared to light microscopic sections (generally 5µm to 7 µm; 90 mm2), much smaller sections (30 to 60 nm; 1000 µm2) and far fewer cells are seen in an ultrathin section. So to be successful, either the pathogen has to be abundant and evenly dispersed throughout the tissues, or the pathologist needs to be able to focus on very small areas of tissue where the best chance to observe the pathogen exists. Prior to looking at ultrathin sections, screening of semithin sections stained with toluidine blue is recommended to ensure that the search with the electron microscope will be rewarding. In doing so, the investigator needs to be able to recognize the structures of interest in semithin sections stained with toluidine blue and under the electron microscope the investigator needs to be familiar with normal cellular structures, such as cellular vesicles, secretory material, and cross-sections of tubules, microvilli and cilia, which may be confused with viruses (Dalton and Haguenau, 1973). The images obtained using electron microscopy may not necessarily represent the actual appearance of the cells, organelles, or pathogens in the normal state. Each time a specimen is collected and processed, the final appearance with the electron microscope has been altered in some way. Understanding the best methods for collection and preservation of biologic material is essential for producing quality images with the least alteration possible. In this chapter we will discuss various techniques for collecting the best specimens for examination, how to handle and fix these specimens, and how to look for specific pathogens with the electron microscope. From the initial collection of the sample to sectioning of the resin-embedded sample, there are multiple steps where poor technique will result in low-quality micrographs. Pitfalls and problems in making specific identifications also will be discussed. Transmission electron photomicrographs of a variety of reptile pathogens seen at the University of Florida and elsewhere over the last 25 years, in both natural and experimental infections, are included to demonstrate the value of this diagnostic tool and the appearance of these agents under the electron microscope. More detailed information regarding EM can be found elsewhere (Bozzola and Russell, 1999; Hayat, 2000; Maunsbach and Afzelius, 1999).

6.2 Historical Perspectives While the light microscope can trace its roots to primitive compound and simple microscopes that were invented in the late 1500s and the following century, the first electron microscope was developed in the early 1930s with continued improvements in the 1940s. At this time, the focus was on the technical aspects of the scopes. It was not until the 1950s when EM was first applied to define the subcellular nature of biological systems. Two types of electron microscopes were

developed at about the same time: transmission electron microscopes and scanning electron microscopes. A hybrid between the two, the scanning transmission microscope, is primarily an analytical tool that can map the atomic composition of the specimen. The transmission electron microscope resulted in magnification around 1000 times over the light microscope. With the increase in magnification of the specimen, there was also a dramatic increase in resolution of microscopic structures. Structures along the cytoplasmic membrane, those structures involved in cell-to-cell interaction, and various subcellular structures were observed for the first time. Additionally, pathogens such as viruses and other microbes not recognizable by the light microscope could be identified. While the era of descriptive EM for most basic biological structures common to cells has passed, the use of the electron microscope as an analytical tool will go on for some time to come. There are also endless opportunities for identifying new pathogens in the tissues of ill reptiles and other nondomestic species. Up to the early to mid-1990s, images of structures visualized under the electron microscope were captured on film as negatives and ultimately as black and white photomicrographs. More recently, most modern EM facilities have converted to digital imaging. The quality of digital images is comparable to those obtained on negatives and can be stored and posted on a web page allowing access by multiple investigators. Digital images are easily sent as attachments to e-mail messages. Most scientific journals will now accept electronic manuscripts and digital images, thus making the need for submission of hard copies unnecessary. However, there is generally a limit to how many high-resolution images can be e-mailed per message. However, a file transfer protocol (FTP) site can be created on a server for uploading large files from a computer that can be accessed and downloaded by other computers. The ability to electronically transmit and submit digital files has revolutionized the field of scientific publications.

6.3 Electron Microscopy 6.3.1 Positive Staining Transmission Electron Microscopy (PSEM) In PSEM, the specimens undergo fixation (primary and secondary), followed by embedding, curing, and sectioning. Subsequently, ultrathin sections are stained with heavy metal ions (lead and uranyl), which bind to organelles and macromolecules, a process that increases their density. As the electron beam passes through the specimen, differential staining of cellular structures results in increased contrast. The image can be captured on negative film, which can be converted into a positive. In most modern laboratories, digital images can be captured and stored on a computer, resulting in electronic access.

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6.3.1.1 Fixatives and Fixation  The purpose of fixation is to prevent changes in tissues (cells) that are collected for evaluation. The methods used in preparing samples for PSEM are fairly harsh, and optimum fixation is needed to stabilize the samples to prevent changes (such as swelling and shrinkage) that would take place in otherwise unfixed samples. The ideal fixative is one that results in preservation of the structure of the tissue (cells, organelles, macromolecules, pathogens) as close to that of the normal state as possible. The fixative should prevent artifactual changes that may occur as the specimen is processed, sectioned, stained, and examined as an electron beam passes through it under the electron microscope. Different fixation protocols have been developed for use in preserving specific cellular and subcellular structures. Fixation of tissues can be performed either in the animal using vascular perfusion, or tissues can be removed and placed into the fixative (at 4°C), and delicately sectioned into 1-mm3 portions. Vascular perfusion is the best method of achieving uniform fixation of tissue. Even under the best of conditions, uniform fixation of all components of a sample rarely occurs. When using vascular perfusion, the major artery supplying the organ of interest is cannulated and the corresponding vein draining that organ is severed. Using gentle pressure, the organ is completely perfused with cold (4°C) fixative, thereby draining all the blood from the organ. By flushing the organ at specific time intervals with cold buffer, the duration of fixation can be controlled.  Because the size of the specimen will directly affect uniform and proper fixation, tissue specimens need to be gently cut into small specimens. A handheld razor blade is often used to cut specimens into small cubes, with 0.5 mm3 being an ideal size (Hayat, 2000). When employed, the tissues need to be swiftly cut in one motion to avoid mechanical damage. 6.3.1.1.1 ˙Glutaraldehyde and Osmium Tetroxide  Whereas neutral buffered 10% formalin (NBF) is the universal fixative for light microscopy, and many other fixatives have been developed for better preservation and staining of specific structures at a light microscopic level, since its development over 40 years ago (Sabatini et al., 1963), sequential fixation with glutaraldehyde (primary fixative) and osmium tetroxide (secondary or post fixation) at 4°C or room temperature has become the universal method of fixing tissues in PSEM. These two fixatives are capable of stabilizing a wide variety of different types of molecules. Glutaraldehyde functions as a fixative by cross-linking protein, while osmium tetroxide reacts primarily with lipids. In the process of reacting with lipids, osmium tetroxide is reduced, adding density and contrast to the tissue. Because of this, osmium tetroxide also functions as a stain. While glutaraldehyde penetrates tissues rather slowly (1 mm per h), osmium tetroxide is even slower (0.5 mm per h). For tissues previously fixed in NBF for light microscopy, portions can be transferred to a modified Karnovsky’s solution (1% paraformaldehyde and 2% glutaraldehyde in 0.1 M of sodium cacodylate with 0.001 M of calcium chloride at pH 7.4) when

PSEM is to be used for determining the presence of viruses in the specimen. However, structural preservation will be less than if glutaraldehyde and osmium tetroxide were used initially. The art of fixation is to determine the best fixation time for the tissue being evaluated for both primary and secondary fixatives. Most EM laboratories have generic protocols that can be used for a wide variety of tissues. Modifications can be made with both primary and secondary fixatives to provide better preservation of specific tissues and structures (Hayat, 1981). Until specifically determined, reptile tissues should be fixed in glutaraldehyde for 2 to 4 h, followed by postfixation for 0.5 to 2 h in osmium tetroxide. For preservation of tissue and cell structures, a buffer needs to be added to the primary fixative. The most common buffers used are cacodylate and phosphate. The amount used will depend on the most appropriate pH needed for the tissue being fixed. An ideal pH range for mammalian tissues, 7.2 to 7.4, is probably an appropriate range for reptile tissues. Attention is also needed in determining the total osmolality of the fixative. In mammals, 320 milliosmoles is considered an ideal osmolality. In reptiles, an ideal osmolality has not been established. Given that reptiles can vary widely in their plasma osmolality, a single ideal osmolality may not be achievable. Studies are needed to understand the effect of osmolality on the preservation of different reptile tissues processed for electron microscopy. Glutaraldehyde comes in different grades and concentrations. Electron microscopy–grade glutaraldehyde in a 50% concentration is recommended. Glutaraldehyde is generally sold in ampoules and once opened must be used quickly. It degrades rapidly when not refrigerated. Using 50% glutaraldehyde, 6 ml added to 94 ml of buffer will result in a 3% solution. In contrast to glutaraldehyde, osmium tetroxide (often available in ampules) is stable for long periods of time if the ampoule is properly sealed with Parafilm and placed in a zip-type bag. As with the primary fixative, a working solution of osmium tetroxide (1 to 2%) is prepared in buffer. Extreme caution is important when handling any fixative, and this is especially true for fixatives used for PSEM. Because they are extremely toxic, always wear gloves, work under a hood, and use protective eyewear. 6.3.1.1.2 Trump’s Solution  A fixative the author routinely uses for doing light and PSEM is Trump’s solution (TS) (McDowell and Trump, 1976). This solution is a combination of 4% formaldehyde and 1% glutaraldehyde. The value in using TS is that the formaldehyde will penetrate tissues more deeply and quickly than glutaraldehyde, and once prepared, TS is stable under refrigeration for prolonged periods of time. Additionally, once tissues are fixed in TS, they can remain in it for weeks prior to processing. The author has examined field-collected reptile blood cells that were stored in TS for extended periods of time before being processed for PSEM. Trumps solution is often used in diagnostic pathology where collected tissues may or may not be submitted for PSEM.

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When preparing TS, it is important to use only EM-grade formaldehyde, which can be purchased or made fresh from paraformaldehyde powder. When possible, do not use histological-grade formaldehyde because it contains contaminants that can compromise preservation. 6.3.1.1.3 Neutral Buffered 10% Formalin  Neutral buffered 10% formalin is the fixative most commonly used for evaluating tissues for diagnostic purposes by light microcopy. While this fixative is not ideal for biological ultrastructural studies, it does work well enough for diagnostic purposes (Wills, 1983). Tissues fixed in NBF can be postfixed in osmium tetroxide and then processed for EM. Viral ultrastructure tends to be preserved very well in NBF-fixed tissues. Neutral buffered 10% formalin is usually prepared from a stock aqueous solution of 37–40% formaldehyde (the trade name is formalin or formol). A fixative sold as 10% formalin is actually a 3.7 to 4% solution of stock formaldehyde. This is an example of histological jargon that has persisted through time. 6.3.1.1.4 Fixation for Immunostaining  In immunocytochemistry a fixative needs to be used that allows the antigen to be recognized and bound by its corresponding antibody (Skepper, 2000). Some antibodies may identify only their corresponding antigen in unfixed frozen sections (cryoimmobilization). For those that survive fixation, the duration and the strength of the fixative needs to be established for each antigen that the investigator will attempt to label. While some antigens may survive the processing used in routine PSEM, many will not. For immunostaining, the two most common fixatives used are 4% formaldehyde and 1% glutaraldehyde (Skepper 2000); subsequent fixation in osmium tetroxide may or may not be used. These fixatives are prepared in either 0.1 M phosphate, HEPES (N-2-hydroxyethylpiperazine-N’-2ethane sulfonic acid) or PIPES (piperazine-N-N’-bis 2-ethane sulfonic acid) buffer. A typical fixative would be 0.5% glutaraldehyde and 4% formaldehyde in 0.07 M of phosphate buffer for one hour at room temperature. Some investigators recommend adding calcium chloride to the fixative to enhance cellular detail (Stirling 1990). If there is a plan to add calcium chloride, then phosphate buffers need to be avoided because the addition of calcium chloride will result in a calcium phosphate precipitate. Tissues should be fixed at 4°C and for most studies a fixation time of 2 h can be used initially to assess the quality of the fixation for the antigen being labeled. As with routine positive staining TEM, fixation of tissues and organs can be done using vascular perfusion.

6.3.1.2 Infiltration and Embedding  Tissues for PSEM are first infiltrated (replacement of the dehydration agent) with an epoxy medium. This is followed by embedding, a process in which the entire tissue is completely impregnated with the resin. The resin is used to both embed the tissue and to attach it to the block that is placed in the ultramicrotome for sectioning. The epoxy medium consists of a resin, hardener, and

accelerator. Some of the currently used resins are Epoxy Resin 812, Araldite, and Spurr’s resin. For immunocytochemistry, LR White is a commonly used resin. The advantages of this resin are that it is hydrophilic, which allows good penetration of aqueous antibody-containing solutions, and that it is not necessary to completely dehydrate the tissue before transferring it to the resin. Also, it can be polymerized with ultraviolet light at low temperatures avoiding the denaturing effects of heating to 60°C that occur during routine embedding. A major disadvantage is the resulting low contrast of the sections. Reducing the temperature of thermal curing of the resin may also be necessary to preserve the antigen to be labeled.

6.3.1.3 Sectioning, Staining, and Labeling  Tissue sections in diagnostic light microscopy generally range from 5 to 7 µm, with sections cut from paraffin-embedded tissues on a microtome using stainless steel blades. In PSEM, the sections need to be much thinner to allow penetration of the electron beam. The ideal section needs to be between 30 and 60 nm. These thin sections are about 100 times thinner than a paraffin section. Ultramicrotomes are used in producing thin sections and glass or diamond knives are used to cut the sections. Cutting sections from paraffin-embedded blocks is far easier and takes less time than producing quality sections for PSEM. The cutting surface (face) of a resin-embedded block is considerably smaller than that of a paraffin block (Figure 6.1). The number of cells visualized in a section for PSEM is considerably less than that for light microscopy.  Prior to cutting a thin section, a thick or semithin section (0.5 to 1 µm) should be cut from the epoxy block, stained with toluidine blue, and examined using a light microscope. This will allow the investigator to identify the best portion of the specimen to examine by PSEM. Extraneous material can be removed from the block prior to cutting the thin section by careful trimming with a razor blade or glass knife. The investigator should become familiar with the appearance of cells and structures of interest with toluidine blue compared to H&E (Figures 6.2–6.5). In routine diagnostic pathology, H&E is the stain of choice when initially screening tissues. Changes that may be obvious with H&E may not be so obvious using toluidine blue. Because only one stain is used, the contrast is far less than with H&E. However, there is less shrinkage and more detail can be observed. In light microscopy the paraffin sections are placed on a glass slide for eventual staining. In TEM a fine mesh nonmagnetic metal (often copper) grid is commonly used as the support matrix for the sections. The diameter for most grids is approximately 3 mm, with each grid capable of supporting several ultrathin sections. Mesh grids can be selected with different numbers of bars per mm. Depending upon the specimen, additional support may be needed. For instance, in negative staining electron microscopy (NSEM), plastic-coated grids are needed to support liquid specimens. As mentioned earlier, compared to light microscopic sections, semithin and ultrathin sections are much smaller sections and have far fewer cells (Figure 6.6).

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In PSEM, the two most common stains used are 2% uranyl acetate and 2.6% lead citrate. The staining with uranyl acetate can be either in preembedded samples or postembedded sections on mesh grids. Subsequently, sections on copper grids can be stained with lead citrate. In immunocytochemistry, a primary antibody (monoclonal or polyclonal antibody produced against the antigen) is first used to bind the antigen. Because antigens can be modified when tissues or cells are fixed, dehydrated, embedded, and cured, the ability of the antibody to recognize the antigen should first be assessed on unfixed frozen sections. Immunofluorescence is often used in initial screening of the affinity of the antibody to the antigen in frozen sections. The labeling can be done either prior to embedding or after embedding. Pre-embedding labeling is generally used for antigens on the surface of the cell. It is also used when the antigen is extremely sensitive to fixation. In such material the antigen can be labeled prior to fixation in cryosections mounted on a glass slide. For antigens within the cell, postembedded material is generally used. If samples are treated with osmium tetroxide, removal in resin sections may be necessary prior to labeling. This can be achieved by pretreating resin sections with the following oxidizing agents: 4% sodium metaperiodate (Bendayan and Zollinger, 1983) (Figures 6.7, 6.8) or 1% periodic acid (Storm-Mathisen and Ottersen, 1990). These oxidizing agents can also be used sequentially (Skepper 2000). After the primary antibody has been added to the sample, labeling is accomplished using a species-specific secondary antibody that is conjugated with ferritin, peroxidase, or colloidal gold (5 nm to 15 nm). For instance, if a mouse monoclonal antibody is used to determine the presence of its corresponding antigen, then a conjugated antimouse antibody (such as rabbit or goat antimouse antibody) would be used as the secondary antibody. Faulk and Taylor (1971) were the first to use immunogold for antigen labeling. A silver enhancement technique was subsequently developed for detecting small amounts of gold in tissue section (Lackie et al., 1985). If labeling occurs in cryosections and not in ultrathin plastic-embedded sections under the electron microscope, then the antigen-altering step needs to be identified and modified. In immunocytochemistry, either nickel or gold grids are employed instead of copper grids because the copper reacts with the antibody-containing solutions.

6.3.2 Negative Staining Transmission Electron Microscopy (NSEM) In NSEM, the background area surrounding structures of interest such as cells, organelles released by lysed cells, macromolecules, and various pathogens is surrounded by heavy metal atoms, which act as an electron stain (Almeida, 1980). The electron beam penetrates the structure of interest and not the background since the area immediately around the structure of interest is denser than the structures themselves. Thus the structures appear lighter in contrast to a dark surrounding back-

ground. Negative staining is not used on sectioned material, but instead the stain is either mixed with the specimen being evaluated or added onto the specimen once it is placed on the grid. The process is much simpler and quicker than for PSEM.

6.3.2.1 Processing  The most commonly used negative stains are potassium phosphotungstate (PTA; 0.5 to 4.0%) and uranyl acetate (0.5 to 1.0%). Potassium phosphotungstate was the first negative stain to be reported (Brenner and Horne, 1959). In cases where a viral agent is suspected, this technique may be used in a variety of specimens for determining their presence (Figures 6.9–6.11). The optimum pH of the stain will vary from alkaline to acidic depending on the group of viruses thought to be present. The ability to identify the agent will be affected by the concentration of the pathogen in the sample and the extent of background stained material. For the majority of viruses, a concentration of 106 virions/ml of starting material is needed in order to identify virus (both infectious and noninfectious) in the specimen (Almeida, 1980). Depending on the nature of the tissue, different methods of processing the sample are required to detect viral particles. Fluid from vesicles can be obtained with a sterile pipette and may be placed directly on a Formvar-coated copper grid (200- to 400-mesh), while large amounts of fluid (serum, urine, lung washings) require centrifugation for clarification. In these cases the supernatant, after low-speed centrifugation (1500g) or the diluted pellet after high-speed centrifugation (15,000g), is placed on the grid for staining. Fecal material requires suspension and concentration and will be placed in distilled water or phosphate buffered saline (PBS). Water is often preferable because it results in lysis of cells and release of their contents. A very useful method is to mix fecal material with water or PBS to give a 20% suspension in a final volume of 5 ml. After centrifugation, a drop of the supernatant is placed on a grid and negatively stained. Alternatively one drop of a 1:1 mixture of supernatant and stain is placed directly on a grid for examination. Excess water on the grid can be removed with filter paper. If the grids will be stored or cannot be examined immediately, the stain of choice is uranyl acetate because it will not have long-term adverse effects.  Tissues such as liver, kidney, and spleen require grinding. A 10% homogenate in water is a good starting point. If cell cultures are examined for the presence of a virus, the cells should be lysed using either sonication or freezing and thawing. Immunolabeling also can be performed on negatively stained samples. The primary and secondary antibodies are added prior to adding the negative stain. Uranyl acetate (1%), PTA (2 %), and ammonium molybdate (2%) are most commonly used. For visualizing the virus, the primary antibody against a specific virus can be added alone. This will cause the virus to aggregate and thus make it easier for identification.

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6.3.3 Scanning Electron Microscopy (SEM)

vein of chelonians, the ventral tail vein and the supravertebral vessel of crocodilians, the ventral tail vein of lizards, and Compared to TEM, SEM can scan much thicker specimens, the palatine veins, ventral tail vein, and via cardiocentesis with the image having a three-dimensional appearance. The in snakes (Jacobson, 1992). Blood is first collected in a tube surface of cell monolayers, tissues, and multicellular organisms containing an anticoagulant such as lithium heparin. Blood can be viewed in three dimensions (Figure 6.12–6.13). films prepared on coverslips (see Chapter 3) and stained with Wright-Giemsa stain should be viewed under a light micro6.3.3.1 Fixation and Processing  Prior to fixation, it may scope before submitting for TEM. The tube containing the be important to rinse the surface of the specimen with physisample can be placed on crushed ice or in a refrigerator until ologic buffer to clean off any material that may obscure visua decision is made about processing for TEM. If red blood alization. Fixatives and buffers similar to those used in TEM cells are to be examined, whole blood can be centrifuged can be used for SEM. Because it is the surface of the speciand the plasma discarded. A portion of the pellet (0.1 ml) can men that is generally viewed in SEM, penetration of the fixabe removed and placed in a plastic tube containing the fixative into the tissue is generally not an issue. Thus tissues can tive. The fixative used will depend upon the specific electron be thicker than those processed in TEM. Following fixation microscopic technique that will be utilized. Unfortunately, the specimen is rinsed and either freeze dried, chemically optimum pH and osmolality of fixatives routinely used in TEM dehydrated, or critical point dried. have not been determined for reptile blood. The tube should 6.3.3.2 Mounting  In TEM, copper mesh grids are used to be inverted several times to suspend the cells in the fixative, support the specimen. In SEM, the specimen is mounted on and the tube should subsequently be placed in crushed ice a metallic stub that is often aluminum. Specimens are either until fixation is complete (generally 1 to 2 h for cells). This is directly attached to the stub using an adhesive or if the speci- generally followed by secondary fixation in osmium tetroxmen is mounted on another substrate, such as monolayers ide. After washing in buffer, the method most often used for embedding is to add fixed cells to 1.5 to 2% bacteriologicalgrowing on coverslips, the substrate is adhered to the stub. grade heated dissolved agar and to centrifuge the mixture in a heated tube (Taniguchi et al., 1994). Once the agar solidifies, it 6.3.3.3 Coating  After specimens have been mounted, they can be removed from the tube and embedded in the resin. If are coated with a conductive metal such as gold. This prethe focus of the study is white blood cells, then the buffy coat vents a buildup of charge on the surface of the specimen that of the blood can be removed with a pipette following centrifucan interfere with the electron beam used in visualizing the gation and then processed according to the method for whole specimen. Gold is most commonly applied using the sputter blood. Certain white cells are directly below the margins of coating technique. Gold sputtering devices are commercially the buffy coat, so a narrow layer of cells directly below the available. buffy coat layer should also be collected. For cell cultures, cells grown in plastic flasks can be detached using trypsin or a cell culture scraper. Next, the cells 6.4 Collection of Samples are processed according to the procedure for whole blood. A novel cell culture technique for TEM utilized the pyramidal Images of samples evaluated with the electron microscope portion of a Beam capsule (Wang et al., 1993). Untreated, the will be only as good as the methods used in collecting and Beam capsule cap, which is polyethylene, does not provide a processing them. Artifactual changes in cellular components suitable surface for cell growth. The authors washed the capand changes in the tertiary structure in antigens to be labeled sule’s inner surfaces with 5% hydrochloride and then coated it using immunocytochemistry can occur at any stage in the prowith CR-human extracellular matrix. Cells are then added and cessing of the sample. The most common diagnostic samples once established they can be fixed in glutaraldehyde, postcollected include: (1) cell cultures and blood cells, (2) biopsies fixed in osmium tetroxide, and embedded in situ in resin. of organs and tissues, (3) portions of organs and tissues from postmortem specimens, (4) paraffin-embedded tissues, and 6.4.2 Biopsies (5) feces, aspirates, and washings for NSEM. Methods of collection are discussed below. While light microscopy is generally considered the gold standard for evaluating biopsy specimens, TEM is being used more frequently in the evaluation of biopsy specimens because of its 6.4.1 Blood Cells and Cell Cultures accuracy and cost effectiveness (Dabbs and Silverman, 1988). Blood cells are often examined under the electron microscope In one study, TEM played a major role in the final diagnosis in to determine the nature of cellular inclusions, certain intra27.1% of the cases examined (Dardick et al., 1991). While light cytoplasmic bacteria, and identification of protozoan paramicroscopic immunohistochemical techniques are commonly sites. Blood from reptiles can be collected from a variety of used for demonstrating the presence of proteins of certain sites (see Chapter 3) including the heart, jugular vein, brachial pathogens such as viruses, chlamydia, and mycoplasma, direct vein, ventral coccygeal vein, orbital sinus, and subcarapacial

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visualization of these pathogens can only be made under the electron microscope. In diagnosing diseases in reptiles, biopsies are routinely collected. Diseases of the integument are common in reptiles, and the integumentary system is the easiest to biopsy. For biopsy of the integument, a 2% xylocaine block is satisfactory and can be infiltrated around the biopsy site and the skin cleaned with 70% ethanol and allowed to dry. A biopsy punch or scalpel blade can be used for collecting the sample. Following biopsy, the skin may require a single suture for closure. The chelonian shell, for most species, is an extremely hard structure and a rotary power saw (Dremel Mototool, Dremel Mfg. Co., Racine, Wisconsin) is an excellent tool for collecting a biopsy specimen. For such procedures, a general anesthetic is required. Biopsies of internal organs can be collected using ultrasound-guided techniques, endoscopy, and excision biopsies during surgery. Because most adult reptiles are small (less than 60 grams in adult body weight), collecting biopsies from internal organs will often not be practical for animals of this size and smaller. Several biopsy specimens should be collected and processed for cytology, histopathology, and EM. The most rewarding samples will be those obtained from a grossly observable lesion. In some diseases, the periphery of the lesion, where abnormal tissue interfaces with normal-appearing tissue, will be the best area to sample. In other diseases, such as those causing granulomatous lesions, the central portion of the lesion will be most rewarding. Biopsies obtained using ultrasound-guided techniques and automated biopsy devices, and those obtained by endoscopy, are generally small (1 mm wide and a few mm in length). Because of this, it is easy to miss the best area harboring the pathogen. Biopsies collected during surgery such as wedge specimens collected with a scalpel blade, are often larger and may have a better diagnostic value compared to those collected using biopsy devices. Because most pathogens are not uniformly dispersed within a tissue, due to the relatively small size of specimens obtained using an endoscope, multiple samples should be collected. Because of the cost and time necessary to prepare and evaluate the specimen, light microscopy will generally be used to screen the sample for its suitability prior to processing and examination using EM. As soon as the biopsies are collected, those to be submitted for EM should be placed in the appropriate fixative, which is chilled and kept on crushed ice. This is followed by fixation in osmium tetroxide and then transfer to buffer where it can be kept until a decision is made to process for EM. The value of the biopsy specimen for ultrastructural studies will depend upon the technique used in collecting and processing the specimen. It is easy to induce crush artifacts, and because of this, the sample needs to be handled with the utmost care. If biopsies are collected at surgery, avoid collecting the sample with forceps. The biopsy can be collected and moved to the fixative using one side of a scalpel blade. Hypodermic needles can be used to move the sample from the blade to the fixative. Needles can also be used to remove a sample from the jaws of an endoscopic biopsy device.

6.4.3 Postmortem Specimens With certain infectious diseases, necropsies will provide the best opportunity for ultimate identification of the causative agent. When an epizootic occurs in a collection of reptiles, the most rewarding cases to workup are those evaluated early in the course of the outbreak. As the outbreak proceeds, secondary invaders may mask the primary pathogen. For many viruses, the replication phase is the best time for identification of the pathogen to be made. Inclusions seen in certain viral diseases, such as herpesvirus and adenovirus, may be prominent only during a relatively short period in the disease process. At necropsy there is an opportunity to collect tissues from all major organ systems. Not all lesions are apparent at a gross level, and their presence will be appreciated only under the light microscope. When gross lesions or changes in tissues are visualized, small pieces from the lesion can be collected, placed in a Petri dish containing chilled TS, and then cut into small portions. With large organs such as the liver, multiple slices can be obtained and each slice can be subdivided into a portion for light microscopy and a portion for EM. With an obvious lesion, we have collected up to 10 slices from a particular organ. Once the slice with the best lesion is identified, the corresponding portion in TS can be submitted for EM. The best way to fix tissues in animals that are severely ill and will be euthanized, is to anesthetize the animal, cannulate an artery supplying the organ of interest, partially sever the vein draining the organ, and then perfuse the organ under gentle pressure. With moderate-sized reptiles, such as bearded dragons and iguanas, the entire animal can be initially perfused with formalin. Subsequently, selected tissues can be collected and additionally fixed in glutaraldehyde followed by subsequent fixation in osmium tetroxide.

6.4.4 Paraffin-Embedded Tissues When fresh glutaraldehyde fixed tissue is not available, wellpreserved paraffin-embedded tissue can be valuable material for diagnostic TEM (Johannessen, 1977). In a TEM study of 15 cytological specimens, in 13 the paraffin-embedded tissue was adequate for ultrastructural evaluation, and it clarified or extended the diagnosis in seven of these cases (Young et al., 1993). When handled properly, most pathogens in paraffinembedded tissue maintain their ultrastructural morphology. Viruses will generally maintain morphologic characteristics, even in less than ideal material. The great value of paraffin-embedded tissue is the ability to localize a specific area to remove and submit for TEM. The histological section on the microscopic slide can be used for orientation. Under the light microscope, the area of interest is identified, circled with a wax pen, and then the slide is placed on the face of the  paraffin-embedded tissue. When the area of interest is identified in the block, and depending upon the size of the tissue in the block, a biopsy punch or a number 11 scalpel blade can be used to remove the portion of interest. The tissue is then deparaffinized, cut into 0.5 mm3 pieces, rehydrated,

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and processed for TEM including fixation in glutaraldehyde and postfixation in osmium tetroxide. In a more rapid procedure, small portions of paraffin-embedded tissue are cut from the block and placed directly in a combination of xylene/ osmium tetroxide followed by embedding in resin (Chien et al., 1982). If paraffin-embedded tissue is no longer available, several methods have been described for removing and processing histological sections on microscopic slides for TEM (Kraft et al., 1983; Chien et al., 1982; Yunis et al., 1977). The coverslip is removed in xylene, the section is rehydrated, and the section is subsequently stained with osmium tetroxide. The section is dehydrated and then the section is covered with a 1:1 acetone and resin mixture followed by replacement with pure resin. A BEEM capsule containing pure resin is applied face down over the section and allowed to polymerize in an oven. In an alternative method, once the coverslip is removed in xylene, the slide is rinsed several times in a xylene and propylene oxide mixture. The section can be covered with a thin layer of the resin (Epon-araldite) and placed in the oven. Once hardened, a razor blade is used to lift the resin-coated section from the slide. Under a dissecting microscope, the area of interest is removed with a scalpel blade and attached to the top of a BEEM capsule block of the resin and processed for EM. Much skill and experience is needed to be successful in removing and processing histological sections on microscopic slides for TEM. This is not a straightforward and easy procedure to do.

6.4.5 Feces, Aspirates, and Washings for Negative Staining Electron Microscopy Feces, aspirates of lesions, washings of body cavities, and washings of systems such as the pulmonary system can provide very valuable information when an infectious disease is suspected. When examining feces, samples collected daily for several days may be needed to rule in or rule out the presence of a pathogen using NSEM. Pathogens may not be continuously shed into the lumen of the digestive tract. Lung washes should be routinely collected from reptiles with respiratory disease. While samples can be collected from some reptiles using manual restraint, other patients will have to be sedated or anesthetized. The jaws of the reptile are held apart and a sterile catheter is guided through the glottis into the lung field. The location of the lung field will vary among the major groups of reptiles, and with snakes will differ even among members of the same family. The clinician needs to know the location of the lung(s) before collecting a lung wash. With a syringe attached to the catheter, sterile saline (1 ml for a 200-gram snake) can be introduced into the lung field and aspirated several times. Material collected can be used for cytological evaluation, microbiological culture, and NSEM. If the reptile is large enough, samples can be collected via bronchoscopy. Bronchoscopy has an added advantage of permitting the lower respiratory tract to be examined directly.

The author prefers the patient to be anesthetized when performing bronchoscopy with the flexible fiber-optic bronchoscope passed through the endotracheal tube. A t-tube connected to the endotracheal tube can be used to allow the technique to be performed while the patient remains connected to the gas anesthesia machine. Utilizing this technique, the tracheobronchial system can be methodically examined and sampling procedures such as lavage, culture, brushing, and transbronchial biopsy can be performed on specific areas of the respiratory tract. When collecting samples for culture, the plugged telescoping catheter brush system is the method of choice (Schaer et al., 1989). Fluid from body cavities can be obtained using a spinal needle and an ultrasound-guided technique. Fluid from vesicles can be collected using a 23- to 25-gauge needle. Aspirates of vesicles, coelomic washes, and lung washes of low cellularity can be concentrated utilizing a number of techniques. As discussed above, cells can be concentrated by simple centrifugation in a plastic-capped tube followed by removal of the pelleted sediment for NSEM.

6.5 Identifying Pathogens in Cells and Tissues In the late 1960s and early 1970s, EM became an important tool for visualizing viral morphogenesis in tissue culture and pathogenesis in experimental animals (Wills, 1983). Subsequently, EM became an important tool for diagnosing pathogens, such as viruses and protozoan parasites, in antemortem and postmortem specimens. Many structures and bodies within red blood cells of reptiles, originally thought to be parasites, were found by EM to be either degenerating cytoplasmic organelles (Alleman et al., 1992) or viruses (Smith et al., 1994; Telford and Jacobson, 1993). The era of describing new pathogens utilizing EM continues today. While EM can be used only to categorize these pathogens to particular groups, the information can be used to select molecular tests (such as polymerase chain reaction; see Chapter 7) that can make a more specific diagnosis and to develop initial strategies for managing epizootics and treating individual animals. Transmission electron microscopy has been commonly used to identify the nature of intracytoplasmic and intranuclear inclusions observed in H&E-stained sections under the light microscope. While intranuclear inclusions may indicate the presence of certain viruses such as adenovirus, herpesvirus, papovavirus, and paramyxovirus, and intracytoplasmic inclusions can indicate the presence of poxvirus, iridovirus, and paramyxovirus, nonviral inclusions also exist. Transmission electron microscopy is the best tool for determining their nature. For example, nonviral intracytoplasmic and intranuclear inclusions seen in histological material from a rat snake were found to represent some type of storage material that was probably lysosomal in origin (Jacobson et al., 1979b)

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(Figures 6.14–6.15). Intranuclear inclusions seen in lymphocytes of a king snake with lymphoma consisted of nonviral, electron-dense granular structures (Jacobson et al., 1980a) (Figure 6.16–6.17). In histological section, eosinophilic intracytoplasmic inclusions seen in pancreatic acinar cells of a Nile crocodile (Figure 6.18) were found to be accumulations of protein and concentric whirls of rough endoplasmic reticulum (Jacobson, 1989) (Figure 6.19). Intracytoplasmic inclusions seen in H&E-stained sections of lung of a desert tortoise with a proliferative pneumonia (Figure 6.20) were found by TEM to be composed of nonviral electrondense material (Figure 6.21). On a light microscopic level, while macrophage-engulfed hemoglobin (Figure 6.22) may be confused with viral inclusions, an accurate diagnosis can be made using TEM (Figure 6.23). Eosinophilic globules (Figure 6.24) seen in hepatocytes of a female diamondback terrapin (Malaclemys terrapin) were found by TEM to consist of nonviral membrane-bound flocculent material (Figure 6.25).

6.5.1 Understanding Ultrastructure of Normal Cells Understanding the ultrastructure of normal cells is essential in order to identify pathogens under the electron microscope. Cross-sections and tangential sections of pores, secretion granules, pinocytotic vesicles, proliferations within the endoplasmic reticulum, keratohyalin granules in epidermal cells, microvesicles from microvesicular bodies, and cross-sections of microvilli and microtubules can be confused with certain viruses. Pathologic changes in cells may add to the confusion (Wills, 1983). Additional images of pseudoviruses can be found elsewhere (Dalton and Haguenau, 1973). Cellular organelles and cell membrane structures in reptiles are similar in ultrastructural appearance to those in mammals. The nucleus is often round and can be euchromatic with a distinct nucleolus (Figure 6.26) or heterochromatic with a less obvious nucleolus. Apical modifications, including cilia and microvilli, are essentially the same as those found in mammals. Microvilli can contain a prominent core of actin, as in absorbing cells along the intestinal tract, or contain fewer microfilaments as seen in areas of secretion, such as along the canaliculi of adjoining hepatocytes (Figure 6.27). Mitochondria (Figure 6.28) possess the classic bi-membrane structure with well-formed cristae and are typically round in most cells, but can elongate to some extent in others. The Golgi apparatus is present in cells involved in packaging and secretion, being recognized by vesicles arising from individual cisternae of dictysomes (Figure 6.29). The endoplasmic reticulum is most ubiquitous, being well represented in most cells both as its protein-secreting form, rough endoplasmic reticulum (rER) (Figure 6.30), and its nonribosomal form, smooth endoplasmic reticulum (sER) (Figure 6.31). Smooth ER is usually associated with a variety of functions such as hormone synthesis, carbohydrate production, detoxification, and cation, principally

calcium, sequestration. Lysosomes (Figure 6.32) are less frequently encountered, being found in cells actively defending the body, such as macrophages and granulocytes, or in cells that experience considerable oxidative activity. Fluid containing proteins, carbohydrates, and other substances are ingested by pinocytotic vesicles with the exchange of metabolites among many cells, (Figure 6.33). To ensure this exchange, junctional complexes (Figure 6.34), consisting of tight junctions, desmosomes, and zonula occludentes, are formed so that substances simply cannot bypass these cells. In older cells, including liver, brain, and muscle cells, as components of the cell are being replaced, remnants of recycled organelles may remain as residual bodies, forming lipofuscin bodies (Figure 6.35). Hepatic melanomacrophages (Figure 6.36), commonly seen in many species of reptiles and other lower vertebrates, are absent in the liver of mammals. Both iron and melanin can be demonstrated in the melanomacrophages using a Perl’s iron stain (Figure 6.37) and a Fontana stain (Figure 6.38), respectively. In kidney of chronically ill reptiles, golden brown pigment (with H&E stain) is commonly seen in renal epithelial cells (Figure 6.39). Monocytes containing this material are seen in the renal interstitium (Figure 6.40). Using a Fontana stain, this material (in renal epithelial cells and monocytes) stains positive for melanin (Figure 6.41). Using TEM, this pigment is compatible with melanin or melanin by-products (Figure 6.42).

6.5.2 Viruses Light microscopic findings, such as intracytoplasmic or intranuclear inclusions, may suggest the presence of a viral pathogen. However, as previously mentioned, not all inclusions are viral in nature, and not all viral infections form inclusions in tissues or cell culture. Certain changes, such as proliferation of the epithelium lining air passageways and perivascular cuffing and round cell infiltrates in the central nervous system, may suggest a viral infection. In such cases, tissues with lesions or cells in culture showing cytopathic effects may be submitted for electron microscopic evaluation. Size, site of replication, morphogenesis, and presence or absence of an envelope are often used to initially categorize a virus to family. A more specific identification can be made using molecular techniques such as polymerase chain reaction (PCR; see Chapter 7). Here I will present examples of viral pathogens identified in tissues of ill reptiles and isolated in cell culture. More detailed information on viral diseases of reptiles can be found in Chapter 9.

6.5.2.1

Herpesviridae  Herpesviruses are enveloped viruses with a double-stranded DNA core surrounded by 162 icosahedrally arranged capsomeres. Infections with herpesvirus have been reported in chelonians (Jacobson et al., 1982a; Jacobson et al., 1985a; Origgi et al., 2004), lizards (Wellehan et al., 2004), snakes (Simpson et al., 1979), and more recently, crocodilians (McCowan et al., 2004; Govett et al., 2005). Her-

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pesviruses are a particular problem in tortoises where severe oral glossitis, pharyngitis, and systemic disease have been reported. Tortoises in the genus Testudo [Greek tortoise (T. graeca), Hermann’s tortoise (T. hermanni), and Russian tortoise (Agrionemys [formerly Testudo] horsfieldii)] are particularly prone to infection. In tortoises, eosinophilic intranuclear inclusions are commonly seen in epithelial cells of affected tissues stained with H&E (Jacobson et al., 1985a) (Figures 6.2, 6.43, 9.35). Hypertrophic nucleoli can also be eosinophilic in H&E stained sections and may be confused with inclusions. Herpesvirus inclusions have been seen in lung, liver, and kidney of emydine turtles (Jacobson et al., 1982a) (Figures 9.26–9.27), in lung and trachea of green turtles with respiratory disease (Jacobson et al., 1986) (Figures 9.8, 9.10), and in cutaneous fibropapillomas of green turtles (Chelonia mydas) (Jacobson et al., 1991b) (Figure 6.44). In semithin sections stained with toluidine blue, inclusions stain blue (Figure 6.3). Using TEM, virions can be identified both in the nucleus and cytoplasm of infected cells; they may accumulate in intercellular spaces. In reptiles, as in other vertebrates, intranuclear particles generally lack an envelope (Figure 6.45) and have either radio-lucent or radio-dense cores (Figure 6.46). Inclusions may not be seen when nuclei contain a small number of virions. Immature particles in the nucleus obtain their primary envelope from the nuclear membrane (Figure 6.47), with enveloped particles found in the cytoplasm (Figures 6.48–6.49). Envelopes also may be obtained at the Golgi, endoplasmic reticulum, and at the cytoplasmic membrane. Glycoprotein-containing projections occur on the surface of the envelope. In reptiles, enveloped particles range from 120 to 140 nm. The envelope is readily seen in negatively stained material. A herpesvirus has been identified in venom glands of cobras (Simpson et al., 1979) and using NSEM, the author has found particles in the venom of Mojave rattlesnakes (Crotalus scutulatus) producing poor quality venom (Figure 6.50). Using TEM, herpesvirus may be confused with adenovirus, which also replicates within the nucleus. However, adenovirus lacks an envelope, and thus mature particles are smaller (65 to 85 nm). Subclinical infections are commonly seen in certain herpesvirus infections, and while DNA sequences of herpesvirus can be identified in infected cells using molecular techniques, intact mature virus may not be found using TEM.

6.5.2.2 Adenoviridae  Adenoviruses are nonenveloped viruses with a double-stranded DNA core surrounded by 252 icosahedrally arranged capsomeres. Virions range in size from 65 to 85 nm. In reptiles, infections with adenovirus have been reported in crocodilians (Jacobson et al., 1984), lizards (Jacobson and Gardiner, 1990; Jacobson et al., 1996), and snakes (Jacobson et al., 1985b; Schumacher et al., 1994b). The only report of an adenovirus in a chelonian is the isolation of an adenovirus (along with a herpesvirus) from a leopard tortoise with biliverdinuria, wasting, and episodes of hemorrhage (McArthur et al., 2004). Using light microscopy,

adenoviral infections are often indicated by the formation of intranuclear inclusions. Using H&E staining, the inclusions are commonly basophilic, and often result in karyomegaly (Figure 6.51) (Jacobson et al., 1996). However, eosinophilic inclusions were reported in a chameleon with inclusions in tracheal epithelial cells (Jacobson and Gardner, 1990). Inclusions may be seen only during discrete periods of the virus replication cycle. Inclusions in adenovirus infections tend to be larger and using TEM, more densely packed with virions (Figure 6.52) than with herpesvirus infection. Virions have hexagonal outlines, an electron-dense core, lack an envelope (Figure 6.53), and may be arranged into crystalline arrays (Figure 6.54). Virions are released with lyses of the cell.

6.5.2.3 Poxviridae  Poxviruses are enveloped viruses with a double stranded DNA core. They are the largest of all viruses, measuring approximately 250 to 400 nm × 150 × 250 nm. Replication is in the cytoplasm. In reptiles, infections with poxvirus have been reported in crocodilians (Buenviaje et al., 1992; Horner, 1988; Huchzermeyer et al., 1991; Jacobson et al., 1979; Pandey et al., 1990; Penrith, 1991) and lizards (Jacobson and Telford, 1990; Stauber and Gogolewski, 1990). The major system affected is the integumentary system. Using light microscopy, poxvirus infections are often indicated by the formation of intracytoplasmic inclusions. Using H&E staining, the inclusions are typically eosinophilic (Figure 6.55). Nonviral intracytoplasmic inclusions consisting of protein (keratohyalin granules) and superficially resembling poxvirus, have been seen in skin lesions of some reptiles with vesicular skin lesions. It is easy to identify poxvirus in cytoplasmic inclusions (Figure 6.56) using TEM. Virions initially develop adjacent to inclusions, in areas of granular viroplasm (Figure 6.57). Immature particles are round and as they mature they give rise to the inclusion seen at the light microscopic level. Mature virions are found within the inclusion and have a biconcave dumbbell-shaped nucleoid, with paired lateral bodies on either side of the nucleoid (Figure 6.58). Jacobson and Telford (1990) reported a dual infection of chlamydia and poxvirus in the cytoplasm of circulating monocytes of a flapneck chameleon (Chameleo dilepis) (Figures 6.59–6.61). Intracytoplasmic inclusions were also seen in macrophages in the spleen (Figure 9.62). 6.5.2.4 Iridoviridae  Iridoviruses may or may not have an envelope, have a double-stranded DNA core, range in size from 120 nm to 300 nm, and replicate in the cytoplasm. Infections with iridovirus virus have been reported in chelonians (Chen et al., 1999; Heldstab and Bestetti, 1982; Mao et al., 1997; Marschang et al., 1999; Muller et al., 1988; Westhouse et al., 1996), lizards (Drury et al., 2002; Telford and Jacobson, 1993), and snakes (Hyatt et al., 2002; Johnsrude et al., 1997; Smith et al., 1994). There are no reports in crocodilians. When examined by TEM, intracytoplasmic inclusions seen in WrightGiemsa-stained peripheral blood films of chameleons (Figure 6.62) were observed to consist of an albuminoid body

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(Figure 6.63), with mature virus in the surrounding cytoplasm (Telford and Jacobson, 1993) (Figure 6.64). Mature virus was enveloped and had sharp icosahedral outlines, a trilaminar structure, and a very electron-dense staining. In tortoises and box turtles (Terrapene carolina) inclusions have be seen in oral and visceral epithelial cells (Johnson et al., 2004; Marschang et al., 1999). In some cases inclusions are abundant and easy to identify, while in others relatively few may be seen despite an intensive search of the section. In a gopher tortoise with tracheitis and pneumonia, intracytoplasmic inclusions were seen in epithelial cells lining the respiratory tract, and using TEM, inclusions consisted of viral particles compatible with an iridovirus (Westhouse et al., 1996) (Figures 6.65–6.66). In green tree pythons with systemic Ranavirus infection, icosahedral viruses (142 nm) were identified in intracytoplasmic inclusions in the liver and infected cultured cells (Hyatt et al., 2002) (Figure 6.67). Immuno-electron microscopy was performed using an antibody raised against epizootic hematopoietic necrosis virus of fish (a Ranavirus) and labeled with protein A-gold. Positive labeling was seen (Figure 6.68) and was considered further proof of viral identity.

(Heldstab and Bestetti, 1984). Basophilic inclusions within enterocytes of 6-wk-old California mountain kingsnakes consisted of viral particles consistent in size and morphology to adenovirus and aggregates of smaller non-enveloped particles consistent with Dependovirus (Wozniak et al., 2000a). Parvoviruses were isolated from a corn snake (Elaphe guttata) (Ahne and Scheinert, 1989) (Figure 9.77) and a boa constrictor and royal python (Ogawa et al., 1992; Farkas et al., 2004). The isolate from the royal python was sequenced and identified as serpentine adeno-associated virus (SAAV) in the genus Dependovirus (Farkas et al., 2004).

6.5.2.7 Circoviridae  The family Circoviridae consists of

small (10 to 20 nm) nonenveloped virions that have icosahedral symmetry. The genome consists of circular, singlestranded DNA. There are two genera within the family. The genus Gyrovirus includes chicken anemia virus, having a negative-sense genome. The genus Circovirus has an ambisense genome and includes porcine circoviruses, duck circovirus, goose circovirus, and psittacine beak and feather disease virus (Phenix et al., 2001; Kloet and Kloet, 2004). While the virus replicates in the nucleus, viral inclusions can be found 6.5.2.5 Papillomaviridae  Members of this family are in both the nuclei and the cytoplasm of infected cells. A capsmall nonenveloped double stranded DNA viruses having an tive painted turtle (Chrysemys picta) that was necropsied after icosahedrally arranged capsid with a diameter of 40 to 55 being found dead in its tank had multiple foci of necrosis in nm. The virus replicates in nuclei, with or without the for- the spleen and liver, with macrophages containing multiple mation of light microscopic inclusions. In reptiles, there are intracyroplasmic inclusions. Using TEM, inclusions consisted only a few reports of papillomavirus infection. In a Bolivian of small 10- to 20-nm virions (Figure 6.71) that were comside-necked turtle with mild proliferative skin lesions, TEM patible with members of the family Circoviridae. revealed intranuclear particles arranged in crystalline arrays, and consistent with papillomavirus (Jacobson et al., 1982b) 6.5.2.8 Paramyxoviridae  Members of the family Para(Figure 6.69). Raynaud and Adrian (1976) reported on papil- myxoviridae contain single-stranded RNA, are 100 to 300 lomas in European green lizards (Lacreta viridis), and while nm in diameter, and are enveloped. Virus replication occurs herpesvirus appears to be the most important virus in these mainly in the cytoplasm with maturation by budding from the cell membrane. The family includes the 4 genera: (1) lesions, papovavirus also was identified using TEM. paramyxovirus (parainfluenza), (2) morbillivirus (measles 6.5.2.6 Parvoviridae  Members of the family Parvoviridae distemper-rinderpest group), (3) pneumovirus (respiratory are small, unenveloped single-stranded DNA viruses with a syncytial virus and pneumonia virus of mice), and (4) rubudiameter ranging from 15 to 22 nm. Because they have a very lavirus (mumps). Paramyxoviruses have been isolated from small genome, they are dependent on host cellular activity numerous species of snakes from several different families in order to replicate. Rapidly dividing cells, including those (see Chapter 9). Paramyxovirus has also been isolated from in the gastrointestinal tract and bone marrow, are targets for lizards (Jacobson et al., 2001a; Marschang et al., 2002). Recent parvovirus. In mammals, intranuclear inclusions are some- comparative analyses of partial gene sequences for the large times seen.  (L) protein and hemagglutinin neuraminidase (HN) protein of The genus Dependovirus was constructed for defective 16 reptilian paramyxoviruses recovered from multiple species parvoviruses, which are seen in adenovirus-infected cells of snakes from different families indicated that there were at and need adenovirus for their replication. Using TEM, they least 2 distinct subgroups of isolates and several intermedihave been seen in the nuclei of bearded dragons that were ate isolates (Ahne et al., 1999). The complete RNA genome co-infected with adenovirus (Figure 6.70) (Jacobson et al., sequence of the archetype reptilian paramyxovirus, Fer-De1996; Kim et al., 2002). Particles have round to hexagonal Lance virus (FDLV) was determined to be 15,378 nucleotides outlines, both electron-dense and electron-lucent cores, and in length and consisting of 7 nonoverlapping genes (Kurath can be arranged into arrays. Nuclear lysis may be seen in et al., 2004). Comparisons made with other paramyxoviruses infected cells, with virus released into the cytoplasm. Using also resulted in a recommendation that a new genus be creTEM, a virus resembling Dependovirus was seen in snakes ated for this virus. It was suggested that the new genus be infected with adenovirus and having gastrointestinal disease named Ferlavirus, with FDLV as the type species.

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The TEM appearance of reptile paramyxovirus is similar to those infecting mammals and birds. In NSEM, the envelope can be seen (Figure 6.72), and when lysed, the innercoiled helical herringbone-appearing nucleocapsid strand is released (Figure 6.73). In TEM, both spherical and filamentous forms can be seen enveloping from the cell membrane (Figure 6.74–6.75). Strands of nucleocapsid material are generally seen within the cytoplasm of infected cells (Figure 6.76). Using a polyclonal antibody produced in rabbits, sites of cytoplasmic nucleocapsid strands were labeled with gold (Richter et al., 1996). In a recently described paramyxovirus in a Boelen’s python with encephalitis (West et al., 2001), intranuclear inclusions were seen, and using an in situ hybridization technique with a probe developed for avian paramyxovirus, paramyxovirus nucleic acid was identified within nuclei. Similar intranuclear inclusions and nucleocapsid material have been seen in diamond pythons and carpet pythons with encephalitis (Boyer et al., 2000).

6.5.2.9 Reoviridae  Members of this family are nonenveloped double-stranded RNA viruses, range in size from 60 to 80 nm, and have two or three shells, each with icosahedral symmetry. The family consists of 9 genera, with each genus having its own morphology and physiochemical characteristics. In reptiles, infection with reovirus has been reported in chelonians, lizards, and snakes (Ahne et al., 1987; Blahak, 1994; Blahak et al., 1995; Lamirande et al., 1999; Marschang, 2000; Vieler et al., 1994; Wellehan et al., 2005). They appear to be distinct from those of mammals and birds and may eventually be placed in their own genus. Regarding pathology and TEM and NSEM characteristics, the best-studied reovirus from a reptile was isolated from juvenile Moellendorff’s rat snakes (Elaphe moellendorffi) and beauty snakes (Elaphe taenuris) that died soon after importation into the U.S. and were subsequently found to have pneumonia (Lamirande et al 1999). Transmission electron microscopy of viper heart (VH2) cells infected with the rat snake reovirus revealed intracytoplasmic spherical to icosahedral particles measuring 70 to 85 nm (Figure 6.77). NSEM revealed a double capsid layer (Figure 6.78). The size, location, and morphogenesis of the particles were consistent with members of the family Reoviridae.

6.5.2.10 Flaviviridae  Members of this family are spherical, enveloped, positive-sense, single-stranded RNA viruses that are approximately 45 to 60 nm in diameter. Japanese encephalitis virus was isolated from Chinese rat snakes (Elaphe rufodorsata) in Korea (Lee et al., 1972). A flavivirus-like agent was isolated from a leopard tortoise (Geochelone pardalis) with a wasting disease (Drury et al., 2001). Recently, mortality-associated outbreaks of West Nile virus (WNV) infection were reported in farmed American alligators (Alligator mississippiensis) in Georgia (Miller et al., 2003) and Florida (Jacobson et al., 2005), and farmed Nile crocodiles (Crocodylus niloticus) in Israel (Steinman et al., 2003). Vero cells are used for isolating WNV from alligators (Figures 6.79–6.80).

6.5.2.11 Togaviridae   Members of the family Togaviridae are enveloped, intracytoplasmic, positive-sense, single-stranded RNA viruses that are approximately 70 nm in diameter. While some reptiles are known to be susceptible to infection with western equine encephalomyelitis virus (Thomas and Eklund, 1962; Gebhardt et al., 1973; Bowen, 1977), no TEM photomicrographs could be found to show viral morphology in reptile host tissues.

6.5.2.12 Retroviridae  Members of the family Retroviridae are spherical, enveloped, single-stranded RNA viruses that measure 80 to 100 nm in diameter. The icosahedrally arranged capsid contains a helical nucleocapsid. Virions replicate in the cytoplasm with envelopment from cytoplasmic and cell membranes. Retroviruses have been identified in all orders of reptiles, many of which were considered endogenous (Martin et al., 1999; Herniou et al., 1998). A retrovirus has been identified in boid snakes with a disease named inclusion body disease (IBD) (Schumacher et al., 1994; see Chapter 9). Intracytoplasmic inclusions seen in visceral epithelial cell inclusions (Figures 6.81–6.82) and nerve cell bodies in the central nervous system commence as polyribosome-derived clusters of small round subunits (Figures 6.83). As additional subunits are deposited on the periphery, the inclusions enlarge (Figure 6.84–6.85). In some sections the inclusions have concentric profiles with subunits on the surface (Figure 6.86). There can be considerable variation in size of inclusions within tissues of a snake with IBD. Inclusions contain a 68-kd protein band (Wozniak et al., 2000), and while in some cases the subunits have an ultrastructural appearance resembling viral particles (Figure 6.87), the current findings indicate that the inclusions are nonviral in composition. While a retrovirus has been observed in cells containing inclusions, it takes much searching under the electron microscope to find mature particles.   Using TEM, morphogenesis has been described for three retroviral isolates that were obtained from boa constrictors having inclusion body disease (Jacobson et al., 2001b). Evaluation of infected VH2 cells revealed intracytoplasmic and extracellular virions (Figures 6.88–6.89). Aggregates of intracytoplasmic C type retroviral particles were found in approximately 95% of infected VH2 cells. Immature budding and mature particles were detected within cytoplasmic vacuoles and phagolysosomes (Figure 6.90). In what we believed to be senescent cells, particles were also seen budding from rough endoplasmic reticulum (Figure 6.91). Particles were pleomorphic; many, particularly budding forms, had a unique morphology. In some budding forms, the nucleic acid crescent was asymmetrically arranged (Figure 6.92). In immature particles, no intermediate layer could be discerned between the electron-dense crescent and the overlying cell membrane. The electron-dense core of the mature virus was typically central, with a loosely applied core shell or capsid (Figure 6.93). Variations in structure of mature virus included eccentric cores with a closely applied core shell (a B-type

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characteristic), eccentric cores in a cylindrical or conical core shell (a Lentivirus characteristic), bar shaped cores (a D-type characteristic), particles with double cores and core shells, and particles with empty core shells and separate cores. Mature particles measured 80 to 90 nm in diameter. Primary cultured kidney cells from boa constrictors with IBD commonly contained vacuoles with electron-dense material (Figure 6.7). Viral particles were more readily identified in these vacuoles following metaperiodate and antigen retrieval treatment for immunogold labeling (Figures 6.8, 6.88) rather than using routine processing for TEM (Figure 6.94). Extracellular C-type retroviral particles were associated with approximately 90% of infected primary kidney cells, whereas intracytoplasmic C-type retroviral particles were found in approximately 80% of these cells. Viral particles in kidney cells were pleomorphic. Periodically, double nucleoid virions were seen. Mature particles measured 80 to 90 nm in diameter. Retroviruses were also demonstrated in the following neoplasms in Burmese pythons (Python molurus bivittaus): (1) round cell tumor, (2) mucinous colonic adenocarcinoma, (3) transitional cell carcinoma of the kidney, and (4) fibrosarcoma (Chandra et al., 2001). Portions of the four neoplasms were examined by TEM and revealed both extracellular and intracellular type C particles within cytoplasmic vacuoles, budding into vacuoles, free within the cytoplasm, and within phagolysosomes (Figure 6.95). Particles ranged in size from 93 to 96 nm. C-type retroviruses were also identified using TEM in renal epithelial cells of lance-headed vipers (Bothrops moojeni) having renal adenocarcinoma (Hoge et al., 1995) (Figures 6.96– 6.99). Mature virus was also found in the lumen of tubules and is apparently eliminated with the urine.

6.5.3 Bacteria Transmission electron microscopy is an important diagnostic tool when trying to identify the presence of certain bacteria that are either difficult to identify directly in tissue section or difficult to culture. Mycoplasma has surfaced as an important pathogen in tortoises and crocodilians, and TEM has been important in its initial identification in tissue section. Chlamydophila has also surfaced as an important pathogen in reptiles and the early reports were based on the characteristic morphology of its developmental stages. Transmission electron microscopy has also been used to identify miscellaneous bacteria seen in reptiles that were not successfully cultured.

6.5.3.1 Mycoplasma  Mycoplasmas have emerged as important pathogens in reptiles (see Chapter 10). In 1988, a chronic upper respiratory tract disease (URTD) was recognized in desert tortoises (Gopherus agassizii) in the Desert Tortoise Natural Area, Kern County, CA. The disease was characterized clinically by serous, mucous, or purulent nasal and ocular discharge, conjunctivitis and palpebral edema. At a light micro-

scopic level there is infiltration of the nasal cavity mucosa and submucosa with inflammatory cells accompanied by hyperplasia and degeneration of upper respiratory tract epithelium (Jacobson et al., 1991a; Brown et al., 1994; Jacobson et al., 1995). Similar signs of disease have been seen in free-ranging gopher tortoises (Gopherus polyphemus) (McLaughlin et al., 2000). Transmission electron microscopic examination of the upper respiratory tract mucosa of free-ranging desert and gopher tortoises with URTD demonstrated the presence of a bacteria with features compatible with Mycoplasma including: (1) lack of a cell wall, (2) pleomorphism, and (3) size range of 400 to 800 nm (Figures 6.100–6.101). A previously undescribed species of mycoplasma, Mycoplasma agassizii, was subsequently cultured from nasal lavages of affected tortoises. Pasteurella testudinis, another potential pathogen implicated in the etiology of this disease (Snipes et al. 1980, Snipes and Bieberstein 1982), also was cultured from affected tortoises. See Chapter 10.17 for details on Mycoplasma in tortoises.  Crocodilians (alligators and crocodiles) are susceptible to infection with a pathogenic mycoplasma. Mycoplasma crocodyli causes polyarthritis, pneumonia, and death in Nile crocodiles (Kirchhoff et al., 1997; Mohan et al., 1995). In 1995, an outbreak of mycoplasmosis caused the death or euthanasia of 60 American alligators from a population of 74 captive bull alligators (Clippinger et al., 2000). On gross necropsy and histopathologic examination, pneumonia, pericarditis, and multifocal arthritis were seen. Using TEM, mycoplasma was identified in synovial tissue (Figure 6.102). Using TEM, a mycoplasma was observed adhered to the epithelium lining of the pulmonary tissue and trachea of a Burmese python with proliferative pneumonia and tracheitis (Penner et al., 1997) (Figure 6.103).

6.5.3.2 Chlamydia  The order Chlamydiales consists of 4 families. Of these, the family Chlamydiacea contains members that are known to be pathogens in humans and other animals. A reclassification of Chlamydiacea has resulted in the recognition of nine species within the following two genera: Chlamydia (C. trachomatis, C. suis, C. muridarum) and Chlamydophila (C. psittaci, C. pneumonia, C. felis, C. pecorum, C. abortus, and C. caviae) (Everett et al., 1999). Chlamydiosis has been reported in several species of reptiles including puff adders (Bitis arietans) (Jacobson et al., 1989), a flap-necked chameleon (Jacobson and Telford, 1990), green turtles in aquaculture (Homer et al., 1994), Nile crocodiles (Huchzermeyer et al., 1994), green iguanas (Iguana iguana) (Bodetti et al., 2002), Burmese pythons (Bodetti et al., 2002), and emerald tree boas (Corallus caninus) (Jacobson et al., 2002). Chlamydia are obligate intracellular pathogens with a unique developmental cycle that involves the interconversion among an extracellular survival form, the elementary body (EB) and an intracellular replicating form, the reticulate body (RB). Because of the difficulty of culturing reptile chlamydia (the author is not aware of any isolates), TEM was instrumental in making an initial diagnosis of chlamydiosis in the above

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reptile cases. The first case reported involved puff adders with granulomatous lesions in visceral organs and the heart. H&E staining of affected tissues demonstrated basophilic inclusions in the center of the granulomas (Jacobson et al., 1989a). TEM demonstrated that the unique developmental stages of chlamydia were within the inclusions (Figure 6.104). Large reticulate (initial) bodies had a granular cytoplasm and measured 400 to 800 nm in diameter. Small elementary bodies had an eccentric electron-dense core and varied from 200 to 340 nm. Intermediate bodies measured 360 to 500 nm with a central dense core and granular cytoplasm. In a  flap-necked chameleon, Wright-Giemsa-stained peripheral blood films revealed two different inclusions within circulating monocytes (Jacobson and Telford, 1990). TEM demonstrated that one of the inclusions consisted of mature poxvirus, while the second inclusion consisted of the following developmental stages of chlamydia (Figures 6.60–6.61): (1) large oval, 800- to 900-nm reticulate bodies; (2) contracted 440- to 680nm intermediate bodies having an electron dense center; and (3) small, round, dense 400- to 440-nm elementary bodies. In green turtles with myocardial necrosis, and no evidence of inclusions (Homer et al., 1994), TEM was initially undertaken to determine if a virus was present. Again, developmental stages of chlamydia (Figure 6.105) were seen with the following measurements: (1) large oval to round, 600- to 880-nm retriculate bodies; (2) oval to round, 390- to 650-nm intermediate bodies having an electron dense center; and (3) small oval to round, 270- to 380-nm bodies. Differences in the size (diameter) range for the different stages in these cases may be a reflection of the manner in which tissues were collected, processed, and embedded. TEM will continue to be a valuable tool in initially identifying the presence of chlamydia in reptile tissues. Molecular tools such as PCR can then be used to determine the nucleic acid sequences of certain genes for categorization at the specific level (Bodetti et al., 2002).

6.5.3.3 Miscellaneous Bacteria  For the most part, bacteria are isolated from lesions in reptiles by culture in specific microbiologic broths and media, and identification is often made using biochemical profiles such as those used in the API system (Analytab Products, Plainview, NY). However, many reptile bacteria cannot be easily categorized using this approach, with many (if not most) yet to be categorized on a specific level. The limitations of the methods used in routine categorization of bacteria isolated from humans and domestic animals need to be realized when interpreting the results of bacteria identifications from diagnostic samples.  Transmission electron microscopy has value in identifying bacteria that have very special requirements and are difficult to isolate. As previously discussed, Mycoplasma agassizii was first identified in the nasal cavity of desert tortoises using TEM before it was isolated (Jacobson et al., 1991). Subsequently, it was isolated in a special broth or media (SP4) used routinely in Mycoplasma isolation attempts. In a rhinoceros iguana with a bacteremia, a spiral-shaped microorganism was

seen in a blood film (Figure 6.106), and following the iguana’s death and necropsy, it was seen using light microscopy in blood vessels and sinusoids of multiple tissues (Jacobson et al., 1980). A subsequent TEM study of the organism in tissues revealed that it had approximately 14 flagella at each polar extremity, was covered by an electron-lucent enveloping sheath, had a discrete electron-dense cell wall, and had blebs on the cell surface (Figure 6.107). These features had some similarity to those members of the bacterial family Spirillaceae. In a map turtle (Graptemys barbouri) that died with cutaneous edema, light microscopic examination of tissues stained with hematoxylin and eosin revealed numerous intracytoplasmic basophilic bodies within Kupffer cells in the liver and macrophages in multiple tissues (Jacobson et al., 1989). Transmission electron microscopy demonstrated the bodies as cytosomes that contained numerous nonflagellated bacteria (Figure 6.108). Subsequently, a bacteria was cultured from the iguana that was identified as Elizabethkingia meningoseptica (formerly Flavobacterium meningosepticum).

6.5.4 Parasites In mammals, TEM has been a useful diagnostic tool for identifying stages of replication and cellular pathogenic effects of certain protozoal parasites such as Trypanosoma, Leishmania, Babesia, Theileria, Cytoauxzoon, Plasmodium, Toxoplasma gondii, Sarcocystis, Cryptosporidium, and members of the phylum Microspora. In reptiles, ultrastructural studies have been performed on a variety of protozoal parasites including Leishmania (Lewis, 1975), coccidia (Gardiner et al., 1986; Paperna, 2003; Paperna and Lainson, 1999; Paperna and Lainson, 2000), haemogregarines (Paperna and Smallridge, 2001; Ramadan et al., 1995; Smallridge and Paperna, 2000; Stehbens and Johnston, 1967) (Figure 6.109), Haemoproteus (Sterling, 1972; Sterling and DeGiusti, 1972), Hepatozoon (Figure 6.110) (Smith and Desser, 1997), Karyolysus (Cubero Sanchez and Alvarez Calvo, 1987), Schellackia (Klein et al., 1992; Ostrovska and Paperna, 1987; Sinden and Moore, 1974), Plasmodium (Aikawa, 1971; Aikawa and Jordan, 1968; Boulard et al., 1983; Klein et al., 1988; Moore and Sinden, 1973; Scorza, 1971), Sarcocystis (Zaman and Colley, 1975) and Trypanosoma (Ranque, 1970). Some intracytoplasmic bodies originally thought to be parasites were found by TEM to represent pools of virus (Smith et al., 1994; Stehbens and Johnson, 1966). Of the various protozoal parasites known to infect and cause disease in reptiles, TEM has been useful in identifying the presence of Cryptosporidium, a pathogenic intranuclear coccidian present in several species of tortoises, and microsporidian infection of bearded dragons.

6.5.4.1 Cryptosporidium  O’Donoghue (1995) reported Cryptosporidium in over 57 different species of reptiles including 40 species of snakes, 15 species of lizards, and 2 species of tortoises. Oocysts of Cryptosporidium were subsequently reported in the feces of wild green turtles in the Hawaiian

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Islands (Graczyk et al., 1997). It has been associated with hypertrophic gastritis and other gastrointestinal lesions in numerous species of snakes and lizards (Brownstein et al., 1977). In an Egyptian tortoise that died with clinical signs of enteritis, Cryptosporidium was identified on the luminal surface of over 80% of enterocytes in the intestine (Graczyk et al., 1998). In leopard geckoes (Eublepharis macularius) with weight loss and anorexia, light microscopy revealed organisms compatible with Cryptosporidium associated with hyperplasia and mononuclear cell infiltrates in the small intestine (Terrell et al., 2003). Using TEM, Cryptosporidium was identified on the apical cell surface of villous enterocytes (Figure 6.111). Cryptosporidium also was identified in aural and pharyngeal polyps in green iguanas (Figure 6.112) (Fitzgerald et al., 1998; Uhl et al., 2001). A presumptive diagnosis is often made using light microscopy, with ovoid to spherical organisms measuring 1.6 to 3.0 µm adherent to the luminal cell surface. Using light microscopy, organisms are best seen in plastic-embedded, thick sections stained with toluidine blue. Developmental stages are best appreciated using TEM and include oocysts, trophozoites, meronts, macrogametocytes, and microgametocytes. Parasites are typically within a parasitophorous vacuole, which is of host origin and is continuous with the host cell membrane. Organisms are attached to the surface of the host cell by an undulant feeder organelle.

6.5.4.2 Intranuclear Coccidia  Intranuclear coccidiosis was identified in radiated tortoises (Geochelone radiata) presented with clinicopathologic findings supportive of renal failure (Jacobson et al., 1994). Light microscopic examination revealed intranuclear protozoa in renal epithelial cells, hepatocytes, pancreatic acinar cells, and duodenal epithelial cells (Figure 6.113). A portion of the small intestine was removed from paraffin and processed for TEM. Compared to sections from paraffin-embedded tissue, developmental stages were better appreciated in resin-embedded thick sections stained with toluidine blue (Figure 6.114). Using TEM, infected nuclei were more numerous closer to the lumen of the intestinal tract. Infected nuclei were hypertrophic and contained one or more organisms. Trophozoites measured 2 to 4 µm in diameter, and meronts were up to 15 µm in diameter and contained up to 16 merozoites. Merozoites budded from a spherical residium in a sunburst array (Figure 6.115). Both microgametocytes and macrogametocytes were seen. Oocysts were uncommon; they were unsporulated and measured up to 12 µm in diameter. Additional cases of intranuclear coccidiosis in tortoises were subsequently reported (Garner et al., 1998; Garner et al., 2006). Molecular tools (PCR) are now being used to categorize these coccidians.

6.5.4.3 Microsporidia  The phylum Microspora consists of obligate intracellular unicellular protozoans that are termed collectively the microsporidia. Microsporidia have an unusual life cycle. Infection begins with injection of sporoplasm into the host cell followed by a proliferating merogonic phase. Eventu-

ally a sporogonic phase begins in which meronts of simple structure transform into sporonts of relative complex structure. It is the morphology, internal and external, of both stages that are used to distinguish microsporidia. While over 100 genera and almost 1000 species have been reported in a wide variety of invertebrates and all classes of vertebrates, relatively few have been described in reptiles. Systemic microsporidiosis was reported in three captive bearded dragons showing nonspecific signs of illness (Jacobson et al., 1998). Light microscopic examination of H&E-stained tissue sections revealed severe hepatic necrosis with clusters of light basophilic intracytoplasmic microorganisms packing and distending hepatocytes and free in areas of necrosis, and within cytoplasmic vacuoles in distended renal epithelial cells, pulmonary epithelial cells, gastric mucosal epithelial cells, enterocytes, capillary endothelial cells, and ventricular ependymal cells in the brain. The microorganism was Gram-positive, acid-fast, and had a small polar granule that stained using the periodic acid-Schiff reaction. Electron microscopic examination of deparaffinized liver revealed merogonic and sporogonic stages of an organism compatible with members of the phylum Microspora. Developmental stages were free in the cytoplasm and were not surrounded by a membrane. Presporulation stages (meronts), approximately 3 µm in diameter, had a diffusely granular, lightly radio-dense cytoplasm with a denser single nucleus. Some presporulation stages were binucleate during division into uninucleate forms. The outer cytoplasmic membrane was smooth. The sporulation stage consisted of sporonts, sporoblasts, primary spores, and secondary spores. Sporoblasts had a granular deposit on their surface, which condensed during spore formation into a thin electron-dense layer forming the exospore. Primary spores were oval, had broadly oval poles, a thin exospore and a thicker endospore, and polar filaments that consisted of 6 pairs of coils in cross-section (Figure 6.116). The anchoring disc of the filament was constricted, elongated, bulbous, and covered the end of the filament. A polar sac did not cover the polaroplast. The filament was isofilar, short, approximately 0.1 µm in diameter, and coiled in six turns. A single nucleus was located in the center of the spore. In the posterior end of the spore, tubular coils of the Golgi system were seen. Many primary spores had germinated, as seen by an absence of contents. In germinated spores, the everted polar tube was anchored subapically, at an angle to the longitudinal axis of the spore (Figure 6.117). Twelve spores were measured and found to be 2.0 to 2.5 µm by 1.0 to 1.1 µm. Secondary spores had a thin exospore and endospore, which was thicker than that of primary spores (Figure 6.118). The polar filament was coiled in 6 turns. Because of a radiodense interior, no further details were discernible.

Acknowledgments The authors thank Hank Adams and Michael M. Garner for reviewing this chapter and providing comments.

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Figure 6.1  Cutting surface of tissues in a paraffin block compared to the surface of a tissue embedded in resin (arrow).

Figure 6.2  Desert tortoise, Gopherus agassizii. Testudinidae. Herpesvirus infection. Photomicrograph of tongue with eosinophilic intranuclear inclusions (arrows). H&E stain.

Figure 6.3  Desert tortoise, Gopherus agassizii. Testudinidae. Herpesvirus infection. Photomicrograph of tongue with basophilic intranuclear inclusions (arrows). Toluidine blue stain.

Figure 6.4  Carpet python, Morelia spilota. Pythonidae. Intranuclear inclusions. Photomicrograph of brain with eosinophilic intranuclear inclusions (arrows). H&E stain. Courtesy of Shelly Newman.

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Figure 6.5  Carpet python, Morelia spilota. Pythonidae. Photomicrograph of brain with pale basophilic intranuclear inclusions (arrows). Toluidine blue stain. Courtesy of Shelly Newman.

Figure 6.6  The sectioning surface of resin-embedded tissue (R) results in much smaller toluidine blue stained sections (TB) than in paraffin-embedded tissue (P) and corresponding H&E-stained sections. The surface of the resin block is trimmed to an even smaller tip when ultrathin sections are prepared for electron microscopy. Several sections can be mounted on each copper grid (G).

Figure 6.7  Boa constrictor, Boa constrictor. Boidae. Retrovirus infection. Transmission electron photomicrograph of primary cultured kidney cells from a snake with inclusion body disease prior to metaperiodate treatment. Vacuoles (*) with electron-dense material are seen in the cytoplasm. Viral particles (arrow) can be seen in one vacuole that is less dense. Uranyl acetate and lead citrate stain. (From Jacobson ER et al. 2001. Amer J Vet Res 62:217– 224. With permission.)

Figure 6.8  Boa constrictor, Boa constrictor. Boidae. Retrovirus infection. Transmission electron photomicrograph of primary cultured kidney cells from a snake with inclusion body disease following metaperiodate treatment. Gold-labeled viral particles (arrow) are readily seen within a cytoplasmic vacuole. Uranyl acetate and lead citrate stain. (From Jacobson ER et al. 2001. Amer J Vet Res 62:217–224. With permission.)

320  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.9  Caiman lizard, Draecena guianensis. Teiidae. Paramyxovirus infection. Transmission electron photomicrograph of viper heart cells inoculated with lung homogenate from a dead caiman lizard. Within culture media of infected cells, negatively stained filamentous nucleocapsid strands with a herringbone structure characteristic of the Paramyxoviridae are seen. Phosphotungstic acid stain. (From Jacobson ER et al. 2001. J Vet Diag Investig 13:143–151. With permission.)

Figure 6.10  Hermann’s tortoise, Testudo hermanni. Testudinidae. Herpesvirus infection. Transmission electron photomicrograph of negatively stained particles compatible with herpesvirus are seen within culture media of Terrapene heart cells infected with tissue homogenates from a Hermann’s tortoise. Phosphotungstic acid stain.

Figure 6.11  Box turtle, Terrapene carolina. Emydidae. Ranavirus infection. Transmission electron photomicrograph of negatively stained particles compatible with Ranavirus are seen within culture media of Terrapene heart cells infected with a tongue homogenate from a dead box turtle. Phosphotungstic acid stain.

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Figure 6.12  Gopher tortoise, Gopherus polyphemus. Testudinidae. Mycoplasma infection. Color-enhanced scanning electron photomicrograph showing colonies of Mycoplasma agassizii on the surface of experimentally infected cultured tracheal cells of a gopher tortoise. Image courtesy of Hank Adams.

Figure 6.13  Desert tortoise, Gopherus agassizii. Testudinidae. Mycoplasma infection. Scanning electron photomicrograph of nasal cavity showing numerous microorganisms scattered over the nasal mucosa of a tortoise with upper respiratory tract disease. Mycoplasma was isolated from the upper respiratory tract of this tortoise. (From Jacobson ER et al., 1991. J Wildl Dis 27:296–316. With permission.)

Figure 6.14  Deckert’s rat snake, Elaphe obsolete deckerti. Colubridae. Nonviral inclusions. Photomicrograph of hepatocytes with vacuolar changes and deeply eosinophilic to lightly basophilic intracytoplasmic (IC) granules. An intranuclear inclusion (IN) with margination of the chromatin material is also seen. H&E stain. (From Jacobson ER et al., 1979. J Wildl Dis 27:296– 316. With permission.)

Figure 6.15  Deckert’s rat snake, Elaphe obsolete deckerti. Colubridae. Nonviral inclusions. Transmission electron photomicrograph showing granules of variable size within several hepatocytes. One large granule is intracytoplasmic (IC) and appears to be bulging into a nucleus while another one is intranuclear (IN). Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1979. J Wildl Dis 27:296–316. With permission.)

322  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.16  King snake, Lampropeltis getula. Colubridae. Nonviral inclusions. Photomicrograph of spleen. There is a lymphocytic aggregate with a large Cowdry type-A intranuclear inclusion (arrowhead) within a lymphoblastic cell of a snake with lymphoma. Small intranuclear inclusions (arrows) also are seen. H&E stain. (From Jacobson ER et al., 1980. J Natl Cancer Inst 6:577–583. With permission.)

Figure 6.17  King snake, Lampropeltis getula. Colubridae. Nonviral inclusion. Transmission electron photomicrograph of spleen showing a lymphoid cell with an electron-dense finely granular intranuclear inclusion surrounded by a less dense area. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1980. J Natl Cancer Inst 6:577–583. With permission.)

Figure 6.18  Nile crocodile, Crocodylus niloticus. Crocodylidae. Nonviral inclusions. Photomicrograph of pancreas. Eosinophilic intracytoplasmic inclusions (arrows) are seen within acinar cells of a juvenile “runt” crocodile from a farm in Zimbabwe. H&E stain. Courtesy of Chris Foggin.

Figure 6.19  Nile crocodile, Crocodylus niloticus. Crocodylidae. Nonviral inclusion. Transmission electron photomicrograph of the pancreas seen in Figure 6.18. Swirls of endoplasmic reticulum correspond to the intracytoplasmic inclusions seen within acinar cells. Uranyl acetate and lead citrate. Courtesy of Chris Foggin.

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Figure 6.20  Desert tortoise, Gopherus agassizii. Testudinidae. Nonviral inclusions. Photomicrograph of respiratory tract. Eosinophilic globules (arrows) are seen within the cytoplasm of ciliated epithelial cells. H&E stain. Courtesy of Michael M. Garner.

Figure 6.21  Desert tortoise, Gopherus agassizii. Testudinidae. Nonviral inclusions. Transmission electron photomicrograph of respiratory tract seen in Figure 6.20. Intracytoplasmic eosinophilic globules consist of an inner core of radio-dense material surrounded by somewhat less dense material. Uranyl acetate and lead citrate. Courtesy of Michael M. Garner.

Figure 6.22  Nile crocodile, Crocodylus niloticus. Crocodylidae. Erythrophagocytosis. Photomicrograph of spleen showing engulfed portions of red blood cells (H) within macrophages. H&E stain. Courtesy of Chris Foggin.

Figure 6.23  Nile crocodile, Crocodylus niloticus. Crocodylidae. Erythrophagocytosis. Transmission electron photomicrograph of the spleen seen in Figure 6.22. Macrophages contain engulfed red blood cells. Uranyl acetate and lead citrate stain. Courtesy of Chris Foggin.

324  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.24  Diamondback terrapin, Malaclemmys terrapin. Emydidae. Photomicrograph of the liver. Eosinophilic globules (arrows) are seen within hepatocytes. H&E stain.

Figure 6.25  Diamondback terrapin, Malaclemmys terrapin. Emydidae. Transmission electron photomicrograph of the liver seen in Figure 6.24. The globules seen by light microcopy consist of membrane-bound electron-dense flocculent material. No virus was identified within this material. Uranyl acetate and lead citrate stain.

Figure 6.26  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Nucleus. Transmission electron photomicrograph of normal kidney. The nucleus of a renal epithelial cell is seen. Uranyl acetate and lead citrate stain.

Figure 6.27  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Canaliculi. Transmission electron photomicrograph of a normal liver. Adjoining hepatocytes with canaliculi are seen. Uranyl acetate and lead citrate stain.

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Figure 6.28  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Mitochondria. Transmission electron photomicrograph of a normal liver. Mitochondria with well-formed cristae are seen within hepatocytes. Uranyl acetate and lead citrate stain.

Figure 6.29  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Golgi apparatus. Transmission electron photomicrograph of a normal liver. The Golgi apparatus (arrows) is seen within a hepatocyte. Uranyl acetate and lead citrate stain.

Figure 6.30  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Rough endoplasmic reticulum. Transmission electron photomicrograph of a normal liver. Rough endoplasmic reticulum is seen within a hepatocyte. Uranyl acetate and lead citrate stain.

Figure 6.31  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Smooth endoplasmic reticulum. Transmission electron photomicrograph of a normal liver. Smooth endoplasmic reticulum is seen within a hepatocyte. Uranyl acetate and lead citrate stain.

326  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.32  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Lysosomes. Transmission electron photomicrograph of normal kidney. Lysosomes are seen within a renal epithelial cell. Uranyl acetate and lead citrate stain.

Figure 6.33  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Pinocytotic vesicles. Transmission electron photomicrograph of a normal liver. Pinocytotic vesicles are seen within a hepatocyte. Uranyl acetate and lead citrate stain.

Figure 6.34  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Junctional complexes. Transmission electron photomicrograph of the liver. Junctional complexes (arrows) are seen between hepatocytes. Uranyl acetate and lead citrate stain.

Figure 6.35   Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Lipofuscin granules. Transmission electron photomicrograph of a normal liver. Lipofuscin granules are seen within a hepatocyte. Uranyl acetate and lead citrate stain.

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Figure 6.36  Desert tortoise, Gopherus agassizii. Testudinidae. Melanomacrophages. Photomicrograph of liver. Melanomacrophages are scattered within the hepatic tissue. H&E stain.

Figure 6.37  Desert tortoise, Gopherus agassizii. Testudinidae. Melanomacrophages. Photomicrograph of the liver. Melanomacrophages stain blue, indicating presence of iron. Granules of iron are also seen in adjacent hepatocytes. Perl’s iron stain.

Figure 6.38  Desert tortoise, Gopherus agassizii. Testudinidae. Melanomacrophages. Photomicrograph of the liver. Melanomacrophages stain black, indicating the presence of melanin. Fontana stain.

Figure 6.39  Desert tortoise, Gopherus agassizii. Testudinidae. Cytoplasmic granules. Photomicrograph of kidney. Golden brown granules are seen within renal epithelial cells. H&E stain.

328  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.40  Desert tortoise, Gopherus agassizii. Testudinidae. Cytoplasmic granules. Photomicrograph of the kidney. Monocytes containing golden brown granules are within the renal interstitium, adjacent to renal tubular epithelial cells. H&E stain.

Figure 6.41  Desert tortoise, Gopherus agassizii. Testudinidae. Melanin granules. Photomicrograph of the kidney. Granules within renal epithelial cells stain black, indicating the presence of melanin or melanin byproducts. Fontana stain.

Figure 6.42  Desert tortoise, Gopherus agassizii. Testudinidae. Melanin granules. Photomicrograph of the kidney. Transmission electron photomicrograph demonstrating melanin or melanin byproducts in the cytoplasm of renal epithelial cells. Uranyl acetate and lead citrate stain.

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Figure 6.43  Argentine tortoise, Geochelone chilensis. Testudinidae. Herpesvirus infection. Photomicrograph of the oral cavity. Desquamated epithelial cells contain eosinophilic intranuclear inclusions (arrows). H&E stain. (From Jacobson ER et al., 1985. J Amer Vet Med Assoc 187:1227–1229. With permission.)

Figure 6.44  Green turtle, Chelonia mydas. Cheloniidae. Herpesvirus infection. Photomicrograph of eosinophilic intranuclear inclusions (arrows) within epithelial cells of a cutaneous fibropapillomas. H&E stain. (From Jacobson ER et al., 1991. Dis Aq Org 12:1–6. With permission.)

Figure 6.45  Green turtle, Chelonia mydas. Cheloniidae. Herpesvirus infection. Transmission electron photomicrograph of a cultured primary kidney cell infected with lung and tracheal homogenates from green turtles with lung, eye, and trachea (LET) disease. Nonenveloped virions compatible with herpesvirus are within the nucleus. Uranyl acetate and lead citrate stain.

Figure 6.46  False map turtle, Graptemys pseudogeographica. Emydidae. Herpesvirus infection. Transmission electron photomicrograph of the liver. Immature nonenveloped herpesvirus particles, with electron-lucent and electron-dense cores, are seen within the nucleus of a hepatocyte. Uranyl acetate and lead citrate stain.

330  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.47  False map turtle, Graptemys pseudogeographica. Emydidae. Herpesvirus infection. Transmission electron photomicrograph of the liver. Enveloped particles are seen in the paranuclear cytoplasm of a hepatocyte. A single virion is in the process of enveloping (arrow) from the nuclear membrane. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1982. J Amer Vet Med Assoc 181:1322–1324. With permission.)

Figure 6.48  Argentine tortoise, Geochelone chilensis. Oral cavity. Testudinidae. Herpesvirus infection. Transmission electron photomicrograph demonstrating mature enveloped herpesvirus particles within desquamated epithelial cells. Uranyl acetate and lead citrate stain.

Figure 6.49  Green turtle, Chelonia mydas. Cheloniidae. Herpesvirus infection. Transmission electron photomicrograph of a cutaneous fibropapilloma demonstrating mature enveloped herpesvirus particles within the cytoplasm of an epidermal cell. Intranuclear inclusions were seen in this area of the tumor. Uranyl acetate and lead citrate stain.

Figure 6.50  Mojave rattlesnake, Crotalus scutulatus. Viperidae. Herpesvirus infection. Transmission electron photomicrograph. Within extracted venom, negatively stained particles compatible with herpesvirus are seen. Phosphotungstic acid stain.

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Figure 6.51  Bearded dragon, Pogona vitticeps. Agamidae. Adenovirus infection. Photomicrograph of the liver. There is karyomegaly of hepatocytes that contain basophilic intranuclear inclusions (arrows). H&E stain.

Figure 6.52  Bearded dragon, Pogona vitticeps. Agamidae. Adenovirus infection. Transmission electron photomicrograph of the liver. A nucleus packed with nonenveloped adenoviral particles is seen. Uranyl acetate and lead citrate stain.

Figure 6.53  Argentine boa constrictor, Boa constrictor occidentalis. Boidae. Adenovirus infection. Transmission electron photomicrograph of the liver. Nonenveloped adenoviral particles with hexagonal outlines, and both electron-lucent and electron-dense cores, are seen within a hepatocyte nucleus. Uranyl acetate and lead citrate stain.

Figure 6.54  Jackson’s chameleon, Chameleo jacksoni. Adenovirus infection. Transmission electron photomicrograph of the trachea. An epithelial cell nucleus contains adenovirus arranged in crystalline arrays (CA). Uranyl acetate and lead citrate stain. (From Jacobson ER and Gardiner CH, 1990. Vet Path 27:210–212. With permission.)

332  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.55  Spectacled caiman, Caiman crocodilus. Alligatoridae. Poxvirus infection. Photomicrograph of skin. There is cytomegaly and karyomegaly of several epidermal cells containing eosinophilic intranuclear inclusions (arrows) indicative of poxvirus infection. Large inclusions (IN) are seen accumulating in the overlying keratin. H&E stain. (From Jacobson ER et al. 1979. J Amer Vet Med Assoc 175:937-940. With permission.)

Figure 6.56  Spectacled caiman, Caiman crocodiles. Alligatoridae. Poxvirus infection. Transmission electron photomicrograph of the skin. An infected epithelial cell contains several inclusion bodies filled with viral particles. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1979. J Amer Vet Med Assoc 175:937–940. With permission.)

Figure 6.57  Spectacled caiman, Caiman crocodiles. Alligatoridae. Poxvirus infection. Transmission electron photomicrograph of skin. An infected epithelial cell shows viral progression from immature spherical particles in the cytoplasmic matrix to mature oval particles within a cytoplasmic inclusion. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1979. J Amer Vet Med Assoc 175:937–940. With permission.)

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Figure 6.58  Spectacled caiman, Caiman crocodilus. Alligatoridae. Poxvirus infection. Transmission electron photomicrograph of skin. A mature poxvirus particle is elliptical and has a dumbbell-shaped nucleoid (N), lateral bodies (L) and an outer membrane (O). Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1979. J Amer Vet Med Assoc 175:937–940. With permission.)

Figure 6.59  Flap-neck chameleon, Chamaeleo dilepis. Chamaeleonidae. Poxvirus and chlamydial infections. Photomicrograph of a circulating monocyte in a blood film. In addition to the nucleus (N), two inclusions (C and P) are seen within the cytoplasm. This lizard had a monocytemia and all monocytes contained either one or both inclusions. Wright-Giemsa stain.

Figure 6.60  Flap-neck chameleon, Chamaeleo dilepis. Chamaeleonidae. Poxvirus and chlamydial infections. Transmission electron photomicrograph of a circulating monocyte seen in Figure 6.59. Two populations of pathogens corresponding to the two inclusions (C and P) are seen. The pathogens were identified as chlamydia (C) and poxvirus (P). Uranyl acetate and lead citrate.

Figure 6.61  Flap-neck chameleon, Chamaeleo dilepis. Chamaeleonidae. Poxvirus and chlamydial infections. Higher magnification transmission electron photomicrograph of a circulating monocyte seen in Figure 6.60. One population (P) is membrane bound and consists of numerous particles of poxvirus. The membrane of the second population (C) has ruptured, releasing its organisms (chlamydia) into the cytoplasm. Uranyl acetate and lead citrate. (From Jacobson ER and Telford SR., 1990. J Wildl Dis 26:572–577. With permission.)

334  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.62  Fischer’s chameleon, Bradypodion fischeri. Chamaeleonidae. Lizard erythrocyte virus infection. Numerous eosinophilic intracytoplasmic inclusions can be seen in red blood cells of a peripheral blood film. Wright-Giemsa stain.

Figure 6.63  Flap-neck chameleon, Chamaeleo dilepis. Chamaeleonidae. Lizard erythrocyte virus infection. Transmission electron photomicrograph of a red blood cell infected with lizard erythrocyte virus (iridovirus). The host cell nucleus (n), albuminoid vacuole (g), and viral particles are seen within the cytoplasm. Bar = 650 nm. Uranyl acetate and lead citrate. (From Telford SR and Jacobson ER., 1993. J Wildl Dis 29:57–63. With permission.)

Figure 6.64  Flap-neck chameleon, Chamaeleo dilepis. Chamaeleonidae. Lizard erythrocyte virus infection. Transmission electron photomicrograph of a red blood cell infected with lizard erythrocyte virus (iridovirus). Enveloped particles with an electron-dense core and icosahedral outlines are adjacent to an intracytoplasmic assembly pool containing developing virus. Bar = 160 nm. Uranyl acetate and lead citrate. (From Telford SR and Jacobson ER., 1993. J Wildl Dis 29:57–63. With permission.)

Figure 6.65  Gopher tortoise, Gopherus polyphemus. Testudinidae. Iridovirus inclusion. Photomicrograph of a desquamated tracheal epithelial cell. An eosinophilic intracytoplasmic inclusion (arrow) is seen adjacent to a pyknotic nucleus (n). H&E stain.

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Figure 6.66  Gopher tortoise, Gopherus polyphemus. Testudinidae. Iridovirus infection. Transmission electron photomicrograph of a tracheal epithelial cell containing numerous iridoviral particles within the cytoplasm and fewer in the extracellular space. Uranyl acetate and lead citrate. (From Westhouse RA et al., 1996. J Wildl Dis 32:682–686. With permission.)

Figure 6.67  Green tree python, Chondropython (Morelia) viridis. Boidae. Ranavirus infection. Transmission electron photomicrograph demonstrating Ranavirus within the cytoplasm of viper heart cells infected with tissue homogenates from a dead green tree python. Uranyl acetate and lead citrate stain. Courtesy of Alex Hyatt. (From Hyatt AD et al., 2004. J Wildl Dis 38:239–252. With permission.)

Figure 6.68  Green tree python, Chondropython (Morelia) viridis. Boidae. Ranavirus infection. Transmission electron micrograph identifying Ranavirus isolated in viper heart cells from a green tree python and labeled using an immunogold technique. Uranyl acetate and lead citrate stain. Courtesy of Alex Hyatt. (From Hyatt AD et al., 2004. J Wildl Dis 38:239–252. With permission.)

336  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.69  Bolivian side-neck turtle, Platemys platycephala. Chelidae. Papillomavirus infection. Transmission electron photomicrograph showing intranuclear papillomavirus particles arranged in a crystalline array. Bar = 1.8 µm. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1982. J Amer Vet Med Assoc 181:1325– 1328. With permission.)

Figure 6.70  Bearded dragon, Pogona vitticeps. Agamidae. Dependovirus infection. Transmission electron photomicrograph of the liver. A hepatocytic nucleus contains adenovirus (larger particles) and arrays of Dependovirus (smaller particles). Bar = 266 nm. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1996. Vet Path 33:343–346. With permission.)

Figure 6.71  Painted turtle, Chrysemys picta. Emydidae. Circovirus infection. Transmission electron photomicrograph of a hepatocyte with multiple inclusions consisting of arrays of a small virus compatible with a member of the family Circoviridae. Uranyl acetate and lead citrate stain. Courtesy of Francisco Uzal.

Figure 6.72  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Transmission electron photomicrograph of a negatively stained virion purified from cultured viper heart cells infected with an Aruba Island rattlesnake isolate of paramyxovirus. Phosphotungstic acid stain. (From Richter GA et al., 1996. Virus Res 43:77–83. With permission.)

Identifying Reptile Pathogens Using Electron Microscopy  337

Figure 6.73  Neotropical rattlesnake, Crotalus durissus. Viperidae. Paramyxovirus infection. Transmission electron photomicrograph of a negatively stained helical nucleocapsid strand released from a ruptured virion purified from viper heart cells infected with a neotropical rattlesnake isolate of paramyxovirus. The herringbone pattern can be seen. Phosphotungstic acid stain.

Figure 6.74  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Transmission electron photomicrograph of Vero cells infected with an Aruba Island rattlesnake isolate of paramyxovirus. Virus can be seen enveloping from cell membranes (arrows) and spheroidal particles are seen in the intercellular space. An intracytoplasmic inclusion (IC) consists of nucleocapsid material. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1997. Vet Pathol 34:450–459. With permission.)

Figure 6.75  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Transmission electron photomicrograph of Vero cells infected with an Aruba Island rattlesnake isolate of paramyxovirus. Filamentous forms (arrows) are seen in the intercellular space. Uranyl acetate and lead citrate. Courtesy of Thomas Geisbert.

Figure 6.76  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Transmission electron photomicrograph of the lung from a snake in an experimental transmission study. Nucleocapsid strands (NC) of paramyxovirus are seen within the cytoplasm of an epithelial cell lining an air passageway. Uranyl acetate and lead citrate stain. Courtesy of Thomas Geisbert.

338  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.77  Viper heart 2 cell (VH2). Reovirus infection. Transmission electron photomicrograph of a VH2 cell infected with Elaphe reovirus. Viral particles (arrows) are present within the cytoplasm. Bar = 200 nm. Uranyl acetate and lead citrate. Courtesy of Elaine Lamirande. (From Lamirande EW et al., 1999. Virus Res 63:135–141. With permission.)

Figure 6.78  Reovirus. Transmission electron photomicrograph of negatively stained Elaphe reovirus purified from cell culture. Note the double capsid structure. Phosphotungstic acid stain. Bar = 100 nm. Courtesy of Elaine Lamirande. (From Lamirande EW et al., 1999. Virus Res 63:135–141. With permission.)

Figure 6.79  American alligator, Alligator mississippiensis. Alligatoridae. West Nile virus infection. Transmission electron photomicrograph of a Vero cell infected with West Nile virus isolated from an American alligator. Intracytoplasmic viral particles (arrows) are seen adjacent to the nucleus (NU). Uranyl acetate and lead citrate stain. Courtesy of Lillian Stark.

Figure 6.80  American alligator, Alligator mississippiensis. Alligatoridae. West Nile virus infection. Higher magnification transmission electron photomicrograph of the West Nile virus infected Vero cell seen in Figure 6.79. Viral particles (arrows) are seen in the cytoplasm adjacent to the nucleus (NU). Uranyl acetate and lead citrate stain. Courtesy of Lillian Stark.

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Figure 6.81  Boa constrictor, Boa constrictor. Boidae. Inclusion body disease. Photomicrograph of the pancreas of a snake with inclusion body disease. Eosinophilic intracytoplasmic inclusions are seen within pancreatic acinar cells. H&E stain.

Figure 6.82  Boa constrictor, Boa constrictor. Boidae. Inclusion body disease. Transmission electron photomicrograph of the pancreas of a boa constrictor with inclusion body disease. Electron-dense inclusions (IN) are seen within the cytoplasm of two cells. Uranyl acetate and lead citrate stain.

Figure 6.83  Boa constrictor, Boa constrictor. Boidae. Inclusion body disease. Transmission electron photomicrograph of enterocytes in the small intestine of a snake with inclusion body disease. During the initial stage of inclusion formation, protein subunits from polyribosomes start accumulating in the adjacent cytoplasm. Uranyl acetate and lead citrate stain.

Figure 6.84  Boa constrictor, Boa constrictor. Boidae. Inclusion body disease. Transmission electron photomicrograph of a hepatocyte. Protein subunits are being deposited at the periphery of an inclusion. Bar = 5 µm. Uranyl acetate and lead citrate stain.

340  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.85   Boa constrictor, Boa constrictor. Boidae. Inclusion body disease. Transmission electron photomicrograph of an inclusion within a hepatocyte. The inclusion consists of protein subunits. Bar = 700 nm. Uranyl acetate and lead citrate stain.

Figure 6.86  Boa constrictor, Boa constrictor. Boidae. Inclusion body disease. Transmission electron micrograph of an inclusion within an enterocyte. In some profiles the inclusion has a concentric pattern with subunits on the surface. Bar = 300 nm. Uranyl acetate and lead citrate stain.

Figure 6.87  Boa constrictor, Boa constrictor. Boidae. Inclusion body disease. Transmission electron photomicrograph of an inclusion in an enterocyte. Deposited protein subunits have a virus-like appearance. Bar = 5 µm. Uranyl acetate and lead citrate stain.

Figure 6.88  Boa constrictor, Boa constrictor. Boidae. Retrovirus infection. Transmission electron photomicrograph of primary kidney cells cultured from a boa constrictor with inclusion body disease. The material was treated with metaperiodate and was subsequently labeled with gold. Gold-labeled particles are seen on the surface of retroviral particles within cytoplasmic vacuoles. Bar = 300 nm. Uranyl acetate and lead citrate stain.

Identifying Reptile Pathogens Using Electron Microscopy  341

Figure 6.89  Boa constrictor, Boa constrictor. Boidae. Retrovirus infection. Transmission electron photomicrograph of primary kidney cells from a snake with inclusion body disease. Extracellular retroviral particles are seen on the surface of kidney cells. Uranyl acetate and lead citrate.

Figure 6.90  Boa constrictor, Boa constrictor. Boidae. Retrovirus infection. Transmission micrograph of lymphocytes of a boa constrictor with inclusion body disease cocultured with viper heart cells. An intracytoplasmic vacuole has clusters of budding mature and immature particles (arrowheads) and particles are seen budding from the plasma membrane (arrow). Bar = 40 µm. Uranyl acetate and lead citrate. (From Jacobson ER et al., 2001. Amer J Vet Res 62:217–224. With permission.)

Figure 6.91  Boa constrictor, Boa constrictor. Boidae. Retrovirus infection. Transmission electron photomicrograph of lymphocytes of a boa constrictor with inclusion body disease cultured with viper heart cells. Viral particles are evident budding from rough endoplasmic reticulum (arrows) of a senescent cell. Bar = 500 µm. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 2001. Amer J Vet Res 62:217–224. With permission.)

Figure 6.92  Boa constrictor, Boa constrictor. Boidae. Retrovirus infection. Transmission electron photomicrograph of lymphocytes of a boa constrictor with inclusion body disease cultured with viper heart cells. A unique budding form of a type-C virus particle is seen, in which the nucleic acid crescent is asymmetrically arranged (arrow). Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 2001. Amer J Vet Res 62:217–224. With permission.)

342  Identifying Reptile Pathogens Using Electron Microscopy

Figure 6.93  Boa constrictor, Boa constrictor. Boidae. Retrovirus infection. Transmission electron photomicrograph of lymphocytes of a boa constrictor with inclusion body disease cultured with viper heart cells. Mature type-C virus particles are seen within a cytoplasmic vacuole. Uranyl acetate and lead citrate stain.

Figure 6.94  Boa constrictor, Boa constrictor. Retrovirus infection. Transmission electron photomicrograph of routinely processed primary kidney cells of a boa constrictor with inclusion body disease. While viral particles are labeled with gold particles, the structure of the virus is not as evident as with prior metaperiodate and antigen retrieval treatment (see Figures 6.8 and 6.88). Uranyl acetate and lead citrate stain.

Figure 6.95  Burmese python, Python molurus bivittatus. Pythonidae. Retrovirus infection. Transmission electron photomicrograph of the spleen of a snake with a neoplasm suggestive of lymphosarcoma. Intracellular type C–like particles are seen in a neoplastic cell. Bar = 0.7 µm. Inset: Viral particles with electron-dense cores and distinct bilaminar external membranes. Bar = 0.3 µm. Uranyl acetate and lead citrate stain. Courtesy of Sundeep Chandra. (From Chandra AMF et al., 2001. Vet Pathol 38:561–564. With permission.)

Identifying Reptile Pathogens Using Electron Microscopy  343

Figure 6.96  Lance-headed viper, Bothrops moojeni. Viperidae. Retrovirus infection. Transmission electron photomicrograph of a renal adenocarcinoma. Renal epithelial cells with extracellular retroviral particles are seen. Uranyl acetate and lead citrate stain. Courtesy of Alma Hoge.

Figure 6.97  Lance-headed viper, Bothrops moojeni. Viperidae. Retrovirus infection. Transmission electron photomicrograph of a renal adenocarcinoma. Mature retroviral particles are seen in a neoplastic renal epithelial cell. Uranyl acetate and lead citrate stain. Courtesy of Alma Hoge.

Figure 6.98  Lance-headed viper, Bothrops moojeni. Viperidae. Retrovirus infection. Transmission electron photomicrograph of the spleen of a lance-headed viper with renal adenocarcinoma cultured with viper heart cells. Retroviral particles are seen budding (arrows) from cytoplasmic membranes. Uranyl acetate and lead citrate stain. Courtesy of Alma Hoge.

Figure 6.99  Lance-headed viper, Bothrops moojeni. Viperidae. Retrovirus infection. Transmission electron photomicrograph of the spleen of a lance-headed viper with renal adenocarcinoma cocultured with viper heart cells. Retroviral particles are seen budding from cytoplasmic membranes adjacent to vacuoles. A few particles are seen within vacuoles. Uranyl acetate and lead citrate stain. Courtesy of Alma Hoge.

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Figure 6.100  Desert tortoise, Gopherus agassizii. Testudinidae. Mycoplasma agassizii infection. Transmission electron photomicrograph of nasal cavity mucosa showing Mycoplasma agassizii closely associated with the plasma membrane. Uranyl acetate and lead citrate stain.

Figure 6.101  Gopher tortoise, Gopherus polyphemus. Testudinidae. Mycoplasma agassizii infection. Transmission electron photomicrograph of the nasal mucosa showing Mycoplasma agassizii (arrows) closely associated with the plasma membrane. Uranyl acetate and lead citrate stain. (From McLaughlin GS et al., 2000. J Wildl Dis 36:272–283. With permission.)

Figure 6.102  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasma alligatoris infection. Transmission electron photomicrograph of the synovium of an alligator with arthritis. Mycoplasma is seen within the connective tissue. Uranyl acetate and lead citrate stain.

Figure 6.103  Burmese python, Python molurus bivittatus. Boidae. Mycoplasma infection. Transmission electron photomicrograph of the lung. Mycoplasma is closely associated with the luminal surface of epithelial cells lining the air passageway of a python with pneumonia. There are areas of increased electron density (arrowheads) where Mycoplasma contacts the epithelium. Uranyl acetate and lead citrate stain. (From Penner JD et al., 1997. J Comp Path 17:283–288. With permission.)

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Figure 6.104  Puff adder, Bitis arietans. Viperidae. Chlamydia infection. Transmission electron micrograph of a granuloma in the liver of a puff adder showing the following developmental stages of chlamydia: initial bodies (I), intermediate bodies (IB), and elementary bodies (EB). Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1989. J Zoo Wildl Med 20:364–369. With permission.)

Figure 6.105  Green turtle, Chelonia mydas. Cheloniidae. Chlamydia infection. Transmission electron photomicrograph of the heart of a green turtle with myocarditis showing the following developmental stages of chlamydia: initial bodies (I), intermediate bodies (IB), and elementary bodies (EB). Uranyl acetate and lead citrate stain. (From Homer BL et al., 1994. Vet Path 31:1–7. With permission.)

Figure 6.106  Rhinoceros iguana, Cyclura cornuta. Iguanidae. Spiral-shaped bacterium. Photomicrograph of a blood film. A spiral-shaped bacterium is seen free in the blood and within circulating monocytes (M). Wright-Giemsa stain. (From Jacobson ER et al., 1980. J Amer Vet Med Assoc 177:918– 921. With permission.)

Figure 6.107  Rhinoceros iguana, Cyclura cornuta. Iguanidae. Spiral-shaped bacterium. Transmission electron photomicrograph of a spiral-shaped bacterium. Flagella (F) are present at both poles, along with an absence of organelles in the polar region (O), and blebs (A) on the cell surface. Uranyl acetate and lead citrate stain.

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Figure 6.108  Barbour’s map turtle, Graptemys barbouri. Emydidae. Elizabethkingia meningoseptica (formerly Flavobacterium meningosepticum) infection. Transmission electron photomicrograph of the liver showing bacteria (arrows) within cytoplasmic vacuoles of a Kupffer cell. Elizabethkingia meningoseptica (formerly Flavobacterium meningosepticum) was isolated from the liver. Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1989. J Zoo Wildl Med 20:474–477. With permission.)

Figure 6.109  American alligator, Alligator mississippiensis. Alligatoridae. Hemogregarina crocodilinorum infection. Transmission electron photomicrograph of blood. Hemogregarina crocodilinorum is seen within the cytoplasm of a red blood cell. Uranyl acetate and lead citrate stain. Courtesy of John Harvey.

Figure 6.110  Prehensile-tailed skink, Carucia zebrata. Scincidae. Hepatozoon infection. Transmission electron photomicrograph of the liver. Zoites of Hepatozoon are seen within a capillary. Uranyl acetate and lead citrate stain.

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Figure 6.111  Leopard gecko, Eublepharis macularius. Eublepharidae. Cryptosporidium infection. Transmission electron photomicrograph of the small intestine. A cryptosporidial organism is seen on the apical surface of an enterocyte. The organism is surrounded by a host plasma membrane. Uranyl acetate and lead citrate. Courtesy of Scott Terrell.

Figure 6.112  Green iguana, Iguana iguana. Iguanidae. Cryptosporidium infection. Transmission electron photomicrograph of an aural polyp. Cryptosporidial organisms are seen on the surface of epithelial cells. Uranyl acetate and lead citrate. Courtesy of Elizabeth Uhl.

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Figure 6.113  Radiated tortoise, Geochelone radiata. Testudinidae. Intranuclear coccidiosis. Photomicrograph of the colon. Stages of a coccidian are seen (arrows) within nuclei of enterocytes. H&E stain.

Figure 6.114  Radiated tortoise, Geochelone radiata. Testudinidae. Intranuclear coccidiosis. Meront of an intranuclear coccidian is seen (arrows) within the nucleus of an enterocyte. Toluidine blue stain. (From Jacobson ER et al., 1994. J Zoo Wildl Med 25:95–102. With permission.)

Figure 6.115  Radiated tortoise, Geochelone radiata. Testdudinidae. Intranuclear coccidiosis. Transmission electron photomicrograph of the colon. Two coccidial organisms are seen within the nucleus (N). The meront contains zoites (Z) that are budding from the residuum (R) within the nucleus of an enterocyte. Uranyl acetate and lead citrate. (From Jacobson ER et al., 1994. J Zoo Wildl Med 25:95–102. With permission.)

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Figure 6.116  Bearded dragon, Pogona vitticeps. Agamidae. Microsporidium infection. Transmission electron photomicrograph of a primary microsporidian spore. The wall consists of an exospore (arrow) and endospore (en). Cross section of the polar filaments is seen as six pairs of coils. Uranyl acetate and lead citrate and stain. Bar = 0.5 µm. (From Jacobson ER, et al., 1998. J Zoo Wildl Med 29:315323. With permission.)

Figure 6.11­7  Bearded dragon, Pogona vitticeps. Agamidae. Microsporidium. Transmission electron photomicrograph of the liver showing a germinated microsporidian spore. There is an absence of contents and the polar tube is everted (arrow). Uranyl acetate and lead citrate stain. (From Jacobson ER et al., 1998. J Zoo Wildl Med 29:315–323. With permission.)

Figure 6.118  Bearded dragon, Pogona vitticeps. Agamidae. Microsporidium infection. Transmission electron photomicrograph of the liver showing a secondary microsporidian spore. The wall is thick, consisting of an endospore (en) and exospore (ex). Cross sections of the polar filament are seen as six pairs of coils. Uranyl acetate and lead citrate stain. (From Jacobson ER, et al., 1998. J. Zoo Wildl Med 29:315–323. With permission.)

7 Molecular Diagnostics April J. Johnson, Francesco C. Origgi, and James F.X. Wellehan, Jr.

Contents

7.1 General Comments

7.1 General Comments.................................................351 7.2 Blotting Techniques................................................351 7.2.1 General Considerations..............................351 7.2.2 Southern Blotting...................................... 352 7.2.3 Northern Blotting...................................... 352 7.2.4 Western Blotting........................................ 354 7.3 Polymerase Chain Reaction................................... 356 7.3.1 Introduction............................................... 356 7.3.2 Reagents..................................................... 356 7.3.3 Method....................................................... 357 7.3.4 Results........................................................ 359 7.3.5 Variations of PCR....................................... 359 7.3.6 Reverse Transcription–PCR (RT-PCR)....... 359 7.3.7 Interpretation of Results............................ 360 7.3.8 Future Directions....................................... 361 7.4 Molecular Phylogeny............................................. 361 7.4.1 Introduction to Molecular Phylogeny....... 361 7.4.2 Molecular Evolution.................................. 362 7.4.3 Sequence Selection.................................... 364 7.4.4 Sequence Alignment................................. 364 7.4.5 Tree-Building Methods.............................. 365 7.4.6 Methods of Measuring Confidence.......... 368 7.4.7 Further Reading......................................... 370 7.5 In Situ Hybridization............................................. 370 7.5.1 Introduction............................................... 370 7.5.2 Probes........................................................ 371 7.5.3 Tissue Preparation..................................... 372 7.5.4 Hybridization............................................. 372 7.5.5 Signal Detection........................................ 372 7.5.6 Limitations................................................. 372 7.6 2D-PAGE................................................................ 373 7.6.1 Introduction............................................... 373 7.6.2 The Procedure........................................... 373 7.6.3 Interpretation of Results, Pitfalls,   and Limitations...........................................374 7.6.4 Advantages and Disadvantages.................374 7.7 Arrays......................................................................374 7.7.1 Introduction................................................374 7.7.2 Gene-Expression Arrays.............................374 7.7.3 Protein Arrays............................................ 375 Glossary of Terms .......................................................... 376 References......................................................................... 376

The advent of molecular-based diagnostics has resulted in rapid advancements in investigation and diagnosis of infectious diseases in reptiles. Use of molecular techniques has resulted in a rapid proliferation of the number of reptile pathogens that can be identified, and a dramatic increase in the sensitivity and specificity of testing for them. As with any technique, the reliability of molecular diagnostics depends on the care and skill of the operator. In order to properly evaluate molecular diagnostic testing, it is crucial to understand the fundamentals of the various tests and assays. This chapter is designed to familiarize veterinarians with the basics of the molecular-based diagnostics currently employed in reptile medicine. The nuts and bolts of the most common diagnostic tests will be discussed, including their advantages and disadvantages, as well as pitfalls to consider when evaluating test results. We hope the information provided will aid clinicians and pathologists in the evaluation and interpretation of molecular-based diagnostic tests.

7.2 Blotting Techniques 7.2.1 General Considerations Detection of nucleic acids using blotting techniques began in the 1970s (Southern, 1975) with the development of the Southern blot. This procedure allowed for the identification of a specific DNA target among billions of nucleotides of genomic DNA. The power and the relative simplicity of this procedure led to the development of similar blot-based techniques for detection of specific RNAs (northern blot) and proteins (western blot).

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7.2.2 Southern Blotting 7.2.2.1 Introduction  The Southern blot analysis was developed in 1975 by E.M. Southern. Using this technique, a specific DNA fragment located on a solid support could be identified using a complementary strand of DNA as a probe. This diagnostic tool was later applied in experimental, clinical, and forensic settings. The versatility and specificity of this test has not been overlooked in the reptile arena, and several examples of its use exist in the literature, particularly within the last decade (Quackenbush et al., 1998; Martin et al., 1997; Turchin and Hsu, 1996; Radtkey et al., 1996).

7.2.2.2 The Procedure  The first step of Southern blot analysis involves enzymatic digestion of the DNA to be analyzed (generally either plasmid or genomic DNA) using one or more restriction endonucleases that recognize and cut specific short DNA sequences. This results in a discrete number of DNA fragments of different sizes (Diagram 7.1A). During the second step of the process, the DNA fragments that have been obtained by restriction digestion are separated by electrophoresis in an agarose gel (Diagram 7.1A). The migration rate of the fragments is a function of their size and the concentration of agarose in the gel. Larger DNA fragments will migrate at a slower pace than smaller fragments. DNA is negatively charged and thus migrates from the negative to the positive pole of the electrophoretic field. Following separation, the DNA fragments are denatured and transferred to a solid support such as a nitrocellulose or a nylon membrane, using capillary action or electrophoresis (Diagram 7.1B). The DNA fragments are then tightly bound and fixed to the membrane, typically by UV irradiation or heat, and then hybridized with one or more probes (Sambrook et al., 1989) (Diagram 7.1C). The probes that are used in Southern blot analyses can be either radioactive or enzyme labeled. Several commercial kits and standard protocols are available to prepare probes for Southern blot analysis. Originally, radioactive probes were more sensitive, but enzyme-labeled probes systems are now comparable. For hybridization, a prehybridization step is performed to block nonspecific binding and reduce the background. Following prehybridization, the probe(s) is then applied to the membrane. The selected probe is generally a DNA (or RNA) fragment with a specific nucleotide sequence complementary to the sequence that needs to be detected (target sequence). Different standardized protocols are available for the hybridization step, which often need to be slightly modified, according to the specific target sequence. The hybridization is followed by several washes, which are performed with buffers of different stringency. The degree of specificity of binding can be adjusted by the choice of buffers and temperature. Following hybridization, the membrane is either exposed to a film (radioactive or chemiluminescent detection) or immersed into the specific substrate solution required by

the enzyme detection system selected. The target sequence should then appear as dark blots on the nitrocellulose or nylon membrane (nonradioactive, nonchemiluminescent) or on the radiographic film (radioactive or chemiluminescent probe), and should correspond to the expected molecular size of the DNA fragment.

7.2.2.3 Interpretation of Results, Pitfalls, and Limitations  Southern blot analysis allows the detection of unique DNA sequences among genomes consisting of several billion nucleotides. It is comparable to finding a needle in a haystack. A reliable Southern blot application is principally a function of a successful hybridization, which depends on a number of factors including the amount of target DNA, the size of the probe, its specific activity, and the amount of DNA on the membrane. In ideal conditions, the method is sufficiently sensitive to detect less than 0.1 pg of DNA (radiographic exposure) (Sambrook et al., 1989). There are many factors that can influence sensitivity and specificity of Southern hybridization, including probe length, efficacy of probe labeling, label type, secondary structure, probe degeneracy, and G/C bias of the target sequence. Appropriate negative and positive controls need to be run to check for probe specificity and system efficiency. If a radioactive detection system is selected, specific training for handling of radioactive materials is required.

7.2.2.4 Advantages and Disadvantages  The Southern blot is a relatively simple and powerful tool to detect specific DNA sequences. With the development of the polymerase chain reaction (PCR; see Section 7.3), the value and the meaning of the Southern was revised to some extent. The typical question for the clinician concerns whether the target sequence is in the DNA being analyzed. This is answered most efficiently with a PCR reaction. For the researcher, there are a number of ongoing useful applications for the Southern blot. As an example, the Southern blot is able to map the position of the target sequence in the context of a genomic or plasmid DNA. When we are analyzing different replication intermediates of DNA viruses or different DNA fragments containing an identical sequence (the target), it is possible that Southern blotting can give very specific answers that PCR analysis cannot. The Southern blot has not become outdated with the advent of PCR; these two procedures can give complementary but different answers to the same question or to interdependent questions.

7.2.3 Northern Blotting 7.2.3.1 Introduction  The great potential of Southern blotting was soon applied to RNA with the development of the northern blot (Alwine et al., 1977, 1979), a technique in which RNA sequences are transferred to a membrane for detection with probes. As in Southern blotting, the main goal of north-

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$ Diagram 7.1 (A, B, C)  A schematic representation of the Southern blot procedure is shown. The extracted DNA is first cleaved by restriction digestion and then resolved on an agarose gel (A). The DNA fragments are then transferred to a solid substrate (nitrocellulose or nylon membrane) either by capillarity or electrophoretic transfer (B). Once the DNA fragments have been transferred and fixed on the membrane, one or more probes are used for the final hybridization and detection (development) steps (C).

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ern blotting is to be able to detect a specific target sequence from the test material (tissue, cell culture, in vitro transcription) transferred to a membrane for hybridization with one or more specific, labeled, RNA or DNA probes.   Northern blotting is useful for detection of specific messenger RNA (mRNA) transcripts and for their quantification, a technique that is very helpful in studying the expression of a gene. Research in reptile medicine and biology has benefited from application of this technique during the last few years (Custodia-Lora et al., 2004; Fujimi et al., 2004; Kurath et al., 2004; Vandenplas et al., 1985; Whitworth et al., 2000). RNA handling is more difficult because of the low stability of RNA compared to DNA. Working with RNA requires a very clean and meticulous approach at every step. RNAses, the enzymes that degrade RNA, are widely present in the environment and are very stable. All glassware needs to be autoclaved and handled with gloves. For RNA procedures nothing should be touched with bare hands.

7.2.3.2 The Procedure  Once the RNA of interest has been isolated, it needs to be resolved on a denaturing gel, which typically contains formaldehyde. Because of the toxicity of formaldehyde, it is important to pour these gels under a chemical hood. The target RNA is separated according to size on the denaturing gel. Larger RNA transcripts or fragments will run slower than smaller ones.   The gel is then transferred to a solid substrate, typically a nitrocellulose or nylon membrane as in Southern blotting. The RNA is then fixed on the membrane by heating (nitrocellulose) or by exposure to ultraviolet light (nylon). Hybridization and detection of the probe with the RNA fixed on the membrane is as previously described for Southern blotting (Diagram 7.1A, B, C). The appropriate detection system will be then adopted.

7.2.3.3 Interpretation of Results, Pitfalls, and Limitations  Most of what has been mentioned for Southern blotting can also be applied to northern blotting. The main difference is how easily RNA is degraded (RNA is less stable than DNA). Consequently, it is very important to check for RNA quality and integrity before starting the procedure. Loading a gel with degraded RNA will result in failure. Suitable controls need to be used throughout the procedure.

7.2.3.4 Advantages and Disadvantages  Northern blotting allows the operator to detect a unique RNA sequence in a mixture of RNAs. A single probe can detect mRNA transcripts of different lengths, which is useful for determining the half-life of a specific mRNA and its synthesis and degradation patterns. The main disadvantage of this technique is that it requires expertise in the proper handling of RNA and a considerable amount of time.

7.2.4 Western Blotting 7.2.4.1 Introduction  Western blotting (Towbin et al., 1979; Burnett, 1981) is to proteins what Southern blotting and northern blotting are to DNA and RNA, respectively. In western blotting, proteins obtained from tissues, cells, or supernatants are resolved in an SDS-polyacrylamide (SDS-PAGE) gel and then transferred to a solid support such as a nitrocellulose membrane. Unlike Southern and northern blotting, the probe is an antibody directed against the target protein. The antibody used for detection can be either a monoclonal antibody or a polyclonal antibody, and may be directly labeled or detected with a secondary labeled antibody (directed against the primary antibody). The proteins can be either in native or denatured status, and this will have direct implications in the detection of the target protein. The choice of the antibody is critical and needs to be done very carefully.   Western blot analysis has frequently been used in reptile medicine research (Coberley et al., 2001a, b; Daniels et al., 2003; De Oliveira et al., 2003; Duggan et al., 2002; Jacobson et al., 2001; Lock et al., 2003; Origgi et al., 2001; Piscopo et al., 2004; Schumacher et al., 1993; Wozniak et al., 2000) and we expect further application of this technique in the field.

7.2.4.2 The Procedure  The first step consists of the preparation of samples to be run in the gel and transferred to the membrane. The cell cultures or tissues (source of proteins) are treated with lysis buffers to solubilize proteins. Several lysis buffers are commonly available and the choice is dependent on the material from which proteins need to be extracted and solubilized. Tissues, cells, bacteria, yeasts, or other materials may require different buffers. Critical components of lysis buffers are protease inhibitors, which prevent degradation of the solubilized proteins. To reduce protein degradation, lysis should be done on ice. Once the solubilization is complete, the protein mixture is centrifuged at high speed to precipitate the particulate material. The supernatant is then harvested and the protein content is determined. Typically, protein electrophoresis is carried out in polyacrylamide gel under denaturing conditions (sodium dodecyl sulfate — SDS) with the presence of a reducing agent such as 2-mercaptoethanol. Anions of SDS-PAGE bind to proteins and, as a result, impart a negative charge to them. Because SDS binds proportionally to the molecular weight of the protein, independently of the amino-acid sequence, SDS–polypeptide complexes migrate through polyacrylamide gels according to their molecular weight (Diagram 7.2A) and not their charge. The effective range of separation of SDS-polyacrylamide depends on the concentration of the polyacrylamide gel and the amount of cross-linking. Consequently, the concentration of polyacrylamide is selected according the expected size of the proteins to be resolved in the gel. Once the proteins have been resolved by electrophoresis on the SDS-polyacrylamide gel, they are transferred to a solid support, such as a nitrocellulose membrane, by capillary

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Diagram 7.2 (A, B, C)  A schematic representation of the western blot technique is shown. The lysated proteins are denatured and loaded on an SDSPAGE acrylamide gel (A). The resolved proteins are then transferred to a nitrocellulose membrane by electrophoresis (B). The investigated protein is then detected using a primary unlabeled and a secondary labeled antibody coupled with the appropriate developing reagent (C).

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action or electrophoresis (Diagram 7.2B). The membrane is then treated with a blocking reagent to avoid nonspecific binding of the antibodies used in the detection step. The first step of detection of the western blot is the binding of the primary antibody to the target protein. Polyclonal or monoclonal antibodies can be used. The primary antibody is resuspended in blocking solution. The membrane is then immersed in this suspension, incubated for a variable amount of time, and washed several times to remove the unbound antibody. The membrane is then incubated in a different solution containing the secondary antibody. The secondary antibody is directed against the primary antibody. The secondary antibody is conjugated with an enzyme that can catalyze a colorimetric (horseradish peroxidase [HRP], alkaline phosphatase [AP], or others) or chemiluminescent reaction with a specific substrate (Diagram 7.2C). After washing away the unbound secondary antibody, the membrane is placed into a developing solution with the substrate for the selected enzyme. The presence of the target protein should appear (when present) as a dark blot on the membrane. If a chemiluminescent agent is chosen, the presence of the target protein will result in the appearance of a dark spot or band on the film.

7.2.4.3 Interpretation of Results, Pitfalls, and Limitations  Western blot analysis allows the detection of a target protein in a mixture of proteins. The detection of the target protein depends on each of the steps described above. The preparation of the sample may be the most critical step of the procedure. A degraded or partially degraded protein can result in a false negative finding. The choices of the appropriate lysis buffer and the use of protease inhibitors are very critical for protein integrity. The lysis should be performed on ice or in a refrigerated environment.  The migration of the proteins in the SDS-polyacrylamide gel is influenced by the acrylamide concentration and by the size of the target protein. It is important to select the appropriate acrylamide concentration according to the expected size of the target protein. Eight to twelve percent acrylamide gels are among those more commonly used for a wide range of protein sizes, and gels with a continuous density gradient are also commonly used. The selection of the appropriate antibody is another crucial step. The specificity of the primary antibody for the selected target protein needs to be confirmed using appropriate controls. The proteins present in the mixture have been denatured, so it is important that the selected antibody will recognize the target protein in its denatured conformation. An antibody that is specific for the native protein may not bind to the denatured protein. Another choice regarding the antibody is whether to use a monoclonal or polyclonal antibody. As a rule of thumb, monoclonal antibodies tend to be more specific but less sensitive. Polyclonal antibodies are generally more sensitive but less specific resulting in the appearance of secondary nonspecific bands. The best approach is to test all available antibodies or reagents to determine which one is best. The specificity of the selected

secondary antibody for the primary antibody that is used also needs to be assessed. As usual, appropriate controls are a must for the correct interpretation of results.

7.2.4.4 Advantages and Disadvantages  Western blot analysis is a very efficient system for protein detection, identification, and quantification. It is a relatively simple procedure that can be very powerful for protein studies or investigation. The main disadvantage of this technique is that it requires expertise in the proper handling of proteins and antibodies.

7.3 Polymerase Chain Reaction 7.3.1 Introduction The polymerase chain reaction (PCR) is a molecular-based technique that allows the exponential amplification of a targeted sequence of deoxyribonucleic acid (DNA) (Diagram 7.3). The concept was first conceived in 1983 by Kary Mullis (1990), who later received a Nobel Prize for its discovery in 1993. The invention of the PCR technique represents a milestone in molecular biology due to its wide range of applications. Among these countless applications, PCR is currently being used to detect viral (Origgi et al., 2004; Teifke et al., 2000; Wellehan et al., 2004), bacterial (Brown et al., 2004), fungal (Milde et al., 2000), and parasitic (Xiao et al., 2004) infections in reptiles. The PCR is a very sensitive test, capable of detecting femtograms (10 −15g) of the target DNA sequence in a large mixture approaching up to a microgram (10 −6 g) of nontarget DNA. Samples routinely used for reptile PCR include tissue from biopsies or necropsies, blood, plasma, oral, cloacal, nasal or tracheal washes, and oral or cloacal swabs. DNA can either be extracted from biological samples using commercial kits or extracted following a conventional phenol-chloroform extraction protocol (Sambrook et al., 1989) and added to a mixture of oligonucleotide primers (forward and reverse), free deoxyribonucleoside triphosphates (dNTPs), a thermally stable DNA polymerase, and a reaction buffer. The reaction mixture is then cyclically heated and cooled in a thermal cycler, repeatedly amplifying the target sequence. The DNA obtained at the end of the reaction can be used for a variety of applications, including sequencing, cloning, or as a probe for in situ hybridization (see Section 7.5) after additional modification.

7.3.2 Reagents The oligonucleotide primers (sense and antisense, or forward and reverse) are designed to be complementary to a specific target DNA sequence of interest (Diagram 7.3) and often span 18 to 24 bases in length (Sachse, 2003). DNA is a doublestranded chain made of a sugar and phosphate backbone with nitrogen-containing bases that bond to the complementary base on the second strand of the DNA (Diagram 7.4). A

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Diagram 7.3  A schematic representation of the polymerase chain reaction (PCR) is shown. The double stranded DNA is denatured. Primers bind to their complementary bases and extension fills in free dNTPs creating the complementary strands of DNA. The procedure is repeated, doubling the target DNA copies with each cycle.

complement of a base is the base to which it binds. Cytosine (C), thymine (T), and uracil (U) are six-sided pyrimidine rings, while guanine (G) and adenine (A) are purine compounds, which have a five-sided ring attached to a pyrimidine ring. A pyrimidine always bonds to a purine. C is always the complement to G, and A is always the complement of T (in DNA) or U (in RNA). The sense primer will bind to the antisense strand of the DNA target while the antisense primer will bind to the sense strand of the same DNA target (Diagram 7.5). The length of the amplified fragment is a function of the distance between the binding sites of the forward and the reverse primers. Primers can be designed to be specific if the sequence of the target DNA is known (Diagram 7.6), or they can be degenerate, with a mixture of primers that vary at one or more specific positions. Degenerate primers permit amplification of a target sequence in which some of the bases may vary between strains or species (Diagram 7.7). One example of degenerate primers being used to amplify a variety of herpesviruses in different species was performed by VanDevanter et al. (1996), where a portion of 22 herpesviruses were amplified with one set of primers. Fourteen of these viruses were in animals and 8 were from humans. This is also called a consensus PCR because the PCR is designed around portions of DNA that are conserved among multiple species of a group of organisms.

Diagram 7.4  The hydrogen bonding between bases of a double strand of DNA is shown.

The dNTPs are free bases (A, G, C, and T), which will be incorporated into the newly synthesized strands. The DNA polymerase is a thermostable enzyme that is needed to initiate the synthesis of the novel strand of DNA, while the reaction buffer contains a mixture of mineral salts that are required for the appropriate activity of the DNA polymerase.

7.3.3 Method The tube containing the DNA, primers, and other reagents is placed into a thermal cycler, which is programmed to undergo a series of heating and cooling cycles that are repeated at specific intervals. Each amplification cycle is characterized by three steps: denaturation, annealing, and extension (Diagram 7.3). The denaturation step consists of the separation of complementary DNA strands at a higher temperature, often between 92° and 94°C (Bagasra and Hansen, 1997). Denaturation is followed by a cooling phase that allows the annealing of the oligonucleotide primers to the target DNA. The annealing temperature is selected as a function of the melting temperature of the primers, which is the temperature at which 50% of the primers are bound to the complementary DNA strand. The melting temperature is dependent on the length of the primer and the G+C content. Because guanine and cytosine are linked by three hydrogen bonds, where adenine and thymine form only two hydrogen bonds (Diagram 7.4), more heat is required to dissociate the G-C bond than A-T bonds. The optimal G+C content of primers has been reported to vary between 40 and 60% (Sachse, 2003), while optimal annealing temperature is usually 2 to 4°C below melting temperature (Bagasra and Hansen, 1997). The molar ratio between primers and DNA is high in order to give advantage to the primers binding to the displaced DNA strands over reannealing of the strands to themselves. Extension occurs at a higher temperature, often around 72°C, the optimal tem-

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Diagram 7.5   A denatured strand of double-stranded DNA is shown with a specific sense primer bonding to the complementary bases of the antisense strand of DNA.

Diagram 7.6  A double strand of DNA is shown with complementary bases aligned. Specific primers are shown in red, and the arrows depict the direction of strand elongation that will occur in a PCR with the shown primers. The primers shown are shorter than would actually be used.

Diagram 7.7  Design of a degenerate primer. Sequences from different species are aligned. Primer base is the complement of conserved bases, or a degenerate base of nonconserved bases. Degenerate bases can be produced that bond to any combination of 2, 3, or 4 bases.

Diagram 7.8  Chromatogram of a portion of a DNA sequence. Each peak represents a base in the sequence, and each of the four colors corresponds to a base: green to adenine, blue to cytosine, red to thymine, and black to guanine. The letters listed above correlate to the peak and represents the computer-generated expected sequence.

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perature for most thermostable DNA polymerases. The polymerase incorporates free dNTPs to form new complementary strands of DNA. Each cycle doubles the number of the target DNA fragments (Diagram 7.3). Theoretically, the cycles can be repeated as many times as necessary to attain the desired number of target copies, but the reaction is often stopped after 35 to 40 cycles.

7.3.4 Results At completion of the program, the amplified DNA can be visualized. The product is resolved in an agarose gel by electrophoresis, similar to that previously described for the Southern blot (Diagram 7.1A). The gel is infused with ethidium bromide, a dye that binds to DNA and enables it to be visualized under ultraviolet light (Figure 7.1). A reference ladder, which consists of DNA fragments of known sizes, is run simultaneously so that the size of the DNA product can be visually determined and compared to the expected amplification size based on the primer design. The product can be purified and its identity confirmed by methods such as sequencing, restriction fragment length polymorphism (RFLP), or hybridization, as discussed below. DNA sequencing is usually performed on automated DNA sequencing machines. First, the DNA fragment is amplified with a mixture of normal dNTPs as well as terminal dideoxynucleotides (ddNTPs) linked to fluorescent dyes. The polymerase will not add on to a ddNTP, and so when a ddNTP is incorporated, the chain ends. Each of the four bases has a ddNTP linked to a different color dye. A set of DNA fragments of different lengths will be obtained, each with a basespecific terminal ddNTP. The synthesized DNA fragments are separated according to size using gel electrophoresis, and the fluorescent dyes are detected. A computer records the dyes running off the gel. The computer then generates a chromatogram (Diagram 7.8) and a sequence file. The sequence is then edited by comparing the chromatogram to the computer-generated sequence to ensure accuracy. Occasionally the dyes can create false readings that need to be manually edited. The edited sequence can then be used to search for similar sequences in the DNA databases. GenBank, the European Bioinformatics Institute (EBI), and the DNA Data Bank of Japan (DDBJ) are three DNA and protein databases that are linked and made available to the public free of charge via the Internet. These databases contain DNA and protein sequences that have been submitted by individual investigators. The Basic Local Alignment Search Tool (BLAST) is a program used to search sequences similar to the unknown sequence. There are several ways to search, the most common being BLASTN and TBLASTX. BLASTN searches by comparing nucleotide sequences. TBLASTX is used for nucleotide sequences that are expected to encode proteins, and is likely to pick up similarities in more distantly related genes than a BLASTN search will. The program translates the unknown sequence into each possible protein sequence in both direc-

tions and compares it to known protein sequences to search for the best match. Three bases (one codon) code for one amino acid and amino acids are the subunits of proteins. When the program translates the sequence, it converts the bases into amino acids. Translation is started from nucleotide 1 and this is defined as frame +1. Starting the translation from nucleotide 2 and 3 of the same strand will lead to the translation of frames +2 and +3 respectively. The process is then repeated for the opposite strand in order to determine the −1, −2, and −3 frames. These six frames include all the different possible combinations of amino acid sequences for that strand of DNA. Results of a BLAST search provide the closest matches (among those available), the degree of similarity of each of the matches, and the frame in which they matched (for TBLASTX), along with links to supplementary information about each match, including the original article(s) that was associated with it, the species it was sequenced from, and its taxonomic classification.

7.3.5 Variations of PCR Many variations have been added or made to the original PCR protocol by Mullis (1990). These modifications have led to the improvement of the performance of the reaction, and have largely enhanced its versatility. Among these modified protocols, the nested PCR has proven to be critical in enhancing the reaction sensitivity. The nested PCR consists of two sequential PCR amplifications. The first round amplifies a larger section of DNA. The product is then used as the template for a second PCR amplification using different primers internal to the first set, which amplifies a smaller fragment of DNA within the first round product. The nested PCR is more sensitive than conventional single-round qualitative PCR. Other nonstandard PCR protocols include the multiplex PCR, which amplifies more than one gene from more than one pathogen in a single reaction by incorporating multiple primer sets in the same reaction. Real-time PCR is a quantitative method of PCR that incorporates a fluorescent signal whose emission is proportional to the amount of product detected in each round of amplification. A laser scanner detects the fluorescence emitted by each of the PCR amplifications in real time. This technique is often abbreviated as RT-PCR, which can be confusing as the same abbreviation is often used for reverse transcription PCR, which is discussed below.

7.3.6 Reverse Transcription–PCR (RT-PCR) 7.3.6.1 Introduction  The direct amplification of RNA by the PCR system is not possible because of the inability of the DNA-dependent polymerase to prime a DNA synthesis reaction starting from an RNA template. This problem is overcome by the reverse transcription PCR (RT-PCR). RNA is first converted into a complementary DNA (cDNA) strand by an RNA-dependent DNA polymerase, the reverse transcriptase

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enzyme (RT) reaction. The cDNA product obtained can then be used as a template in a conventional PCR reaction (Diagram 7.9). RT-PCR has been used to diagnose the presence of RNA viruses, such as paramyxoviruses (Franke et al., 2001, Kindermann et al., 2001), and caliciviruses (Reid et al., 1999), and has also been used to look at gene expression by identifying messenger RNA (mRNA) in tissues (Lu et al., 2003, Herbst et al., 2001, Origgi et al., 2004).

7.3.6.2 Method  RNA is extracted from the biological sample. Next, either short random primers (Diagram 7.9), specific primers, or poly-T primers (Diagram 7.10) are added to the reaction mixture along with the extracted RNA template, a reaction buffer, dNTP mix, reverse transcriptase, and often an RNAse inhibitor to reduce possible RNA degradation during the reaction. The reaction mixture is typically placed in the thermocycler at 42°C for 45 minutes to 1 hour, but slight changes can be adopted according to the template and the kind of reverse transcriptase selected. When messenger RNA (mRNA) is targeted, a poly-T primer annealing to the poly-A tail of the terminal portion of most eukaryotic mRNA transcripts (Diagram 7.10) can be used in the RT reaction. The reverse transcriptase starts the synthesis of the complementary DNA (cDNA) from one of the hybridized primers. An RNA-DNA hybrid is produced. The reaction is then heated to inactivate the reverse transcriptase. For small transcripts, the DNA-RNA hybrid that is obtained at the end of the RT reaction does not generally interfere with the PCR reaction. However, when long transcripts are converted into cDNA, the degradation of the RNA strand may be necessary and can be performed by adding RNAse-H to the mixture. A PCR is then performed on the cDNA with primers designed to amplify the segment of interest.

7.3.6.3 Limitations  RNA is much less stable than DNA,

Contamination can occur at any step of the procedure, including handling of tissue, extraction of the DNA, sample loading prior to running the reaction, or handling of any component of the reaction mix. Nested PCR requires even more attention because it has an additional amplification step where contamination can occur. It is recommended that rooms be dedicated to setting up PCR reactions that are separate from those where DNA is extracted. Samples should be collected in as sterile a manner as possible. The knowledge of the nature of the suspected pathogens could be helpful in ruling out false positive results. For example, while some systemic pathogens may be found throughout the body, others will be location specific. The detection of a pathogen where it is unlikely

Diagram 7.9  A schematic depiction of reverse transcription PCR (RTPCR) using random primers to generate the first cDNA-RNA hybrid is shown. This targets all single-stranded RNA for creating cDNA.

as previously described in Section 7.2.3, and this must be considered when the investigator is planning to extract the sample. RNA is subject to degradation by RNAses, which are ubiquitous. Gloves must always be used when extracting and working with RNA, and available reagents should be used to eliminate RNAses on surface tops, gloves, and pipettes to help avoid degradation of RNA. Because of the unstable nature of RNA, false negatives are more likely with RT-PCR than with PCR, and appropriate positive controls should be incorporated where possible. Difficulties with interpreting results are the same as those listed above for PCR.

7.3.7 Interpretation of Results PCR is a very powerful tool, but care needs to be taken when interpreting results. Its high sensitivity makes contamination a serious confounding factor. Theoretically, one copy of a gene is sufficient to obtain more than one billion copies after 35 cycles of amplification. This implies that any contamination with target DNA is likely to result in a false positive result.

Diagram 7.10  A schematic depiction of RT-PCR using a poly-T primer to generate the cDNA-RNA hybrid is shown. This targets all messenger RNA that have poly-A tails for creating cDNA.

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to be found could suggest a possible contamination event. Samples should be transported in airtight containers on ice packs or dry ice. Any conditions that degrade DNA, such as hot temperatures or use of fixative agents such as formalin, should be avoided. A laboratory that performs PCR should aid clients with the interpretation of a positive or negative result, and should discuss what was done to confirm the result, if anything. There are several ways that results can be reported. The presence of a band on an agarose gel consistent in size with the desired product can be interpreted as a positive result. If nonspecific binding and amplification result in a band of the same size, this can result in a false positive. One way to discriminate whether a positive result is a true positive is the enzymatic digestion of the PCR product using restriction endonucleases. Endonucleases that are specific for one or more target nucleotide sequences within the PCR product can be readily identified and used to produce a specific fingerprint pattern of the digested product. Briefly, the product from a PCR is incubated with endonucleases, which cut DNA at specific sites. The product of the digestion is then resolved in an agarose gel, and the pattern obtained is compared with the pattern determined according to the known sequence of the expected PCR product. If the obtained pattern does not match the expected one, it is generally considered a false positive. The exception would be if mutations occur in the target sequence of the restriction endonucleases, abolishing the restriction site in a true positive sequence. This approach should only be used on products in which the sequence is known and not expected to differ between samples. This restriction endonuclease approach is more definitive than just basing results on PCR product size, but could still potentially result in false positives or negatives as described above. Another way to confirm a positive result is by hybridizing the product with a probe that is specific for the desired target. This is more specific than endonuclease restriction analysis as it uses a labeled complement of the target. There is a higher chance that a wrong sequence could have a specific cut site from an endonuclease than that a labeled complement will bind to a wrong sequence. The most definitive way to confirm a PCR result is by sequencing the product and comparing the results to known sequences in databanks. If the product has been previously sequenced and submitted to the database, a BLAST search will demonstrate the sequence to be most closely related to the previously published sequence. If the amplicon has not been previously identified, it will still show a match with the phylogenetically closest organism of the same taxonomic group. Unfortunately, sequencing is not always performed because of costs. The discriminative power of sequencing is unsurpassed by any other method, making it the method of choice when an absolute, unambiguous result is needed. False negative results should also be considered. These can occur when there is not enough DNA present in the sample or when DNA degradation has occurred. PCR requires

intact DNA, and degradation over time or some treatments that biological samples can undergo, such as formalin fixation, can compromise the DNA integrity. Formalin degrades DNA, and the longer tissues are stored in formalin, the less likely they will be useful for PCR (Tokuda et al., 1990). Tissues that have been embedded in paraffin can be used for PCR, but are better used for amplification of shorter sequences. Time also affects the integrity of DNA, depending on how DNA is stored. Storing samples at −80°C helps preserve longterm viability of DNA. The presence of an internal control for the amplification, such as the target of the PCR reaction, either spiked into one or more samples (inhibition test) or in a separate tube (reaction efficiency control), can control for false negative results as a consequence of degraded DNA or for inhibition of the PCR reaction due to the presence of agents, such as heparin, which can strongly reduce the PCR efficiency. Primers designed to amplify a portion of the animal’s genomic DNA can be used to determine that DNA is present and intact.

7.3.8 Future Directions PCR is constantly increasing in use, as more and more sequences become available through sequence databases. It is a useful tool for detecting known sequences and identifying new ones, and it is easy and inexpensive to perform. Ongoing technological innovations for PCR will continue to make the technique more sensitive and versatile.

7.4 Molecular Phylogeny 7.4.1 Introduction to Molecular Phylogeny With the exponential increase of molecular sequence data now available, the challenge is how to interpret new data in light of what is already available. Accurate analysis and proper interpretation of data can provide valuable information to the reptile clinician and pathologist in many areas, including the identification of pathogens, identification of virulence-associated strains, tracing the spread of epidemics, identification of proper disease models, and host–pathogen evolution. When infectious agents were first identified, we were very limited in our means for speciation and further strain identification. As an example, bacterial identification was typically accomplished by performing a limited number of biochemical tests, and strain identification was accomplished by serotyping. Given that most of the classical biochemical tests used have two possible outcomes, and approximately 20 tests are used for clinical identification of most bacteria, the classical biochemical system is able to discriminate 220 different bacteria, theoretically. The most commonly used gene for bacterial identification, the 16S ribosomal RNA gene (16S rRNA), is roughly 1,500 base pairs in length (Clarridge, 2004). There are four possible nucleotides at each site, so sequenc-

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ing of this gene is able to discriminate approximately 41500, or 21501 different bacteria. Not all of these combinations are biologically viable, but hypothetically speaking, use of this gene sequence has 21481 times the discriminating power of traditional biochemical methods of speciation. Sequencing of additional genes could provide even greater resolution. The use of molecular methods has corrected numerous misidentification errors due to traditional methods (Petti et al., 2005). The enhanced ability to determine relatedness has also resulted in the ongoing revision of bacterial taxonomy. Viral taxonomy has also been revised as a result of this new molecular-based speciation approach (Wellehan et al., 2003). Sequence-based identification is most well established in virology, and viral taxonomy is now primarily based on gene phylogeny and content. Parasitic and fungal taxonomy have been slower to adopt molecular techniques, although the sequence-based approach to phylogeny has already revealed many errors in the existing system (Morrison et al., 2004; Kurtzman and Robnett, 2003). Knowledge of correct organismal phylogeny is critical for the identification of appropriate disease models (Pumphrey and Grey, 1995; Lanford et al., 1998) and of epidemiologically relevant events such as when a pathogen crosses to a new host species (Wellehan et al., 2004). For the epidemiological characterization of pathogen strains, sequence data has also proven to be very useful. Serotyping typically depends on an individual protein being expressed, which may be detected by an antibody. Genetic data can provide much greater resolution than serotyping, and has repeatedly shown that serotyping may incorrectly identify relatedness (Tu et al., 2001; Amonsin et al., 2002). Unlike serotyping, phylogenies can be reliably inferred from sequence data, and this can be very useful to follow the spread of epidemics, especially with rapidly evolving pathogens such as RNA viruses (Towner et al., 2004; Troyer et al., 2004). The use of molecular sequence data is blossoming rapidly, and provides vital information on infectious disease outbreaks to the reptile clinician and pathologist. However, as with any data, misinterpretation of sequence phylogeny may lead to erroneous results. There are many potential pitfalls that can result from use of inappropriate methodology (Anderson and Swofford, 2004; Bollback, 2002), and anyone utilizing molecular phylogenetic data should have a basic understanding of the methods used and be aware of any potential resultant error.

7.4.2 Molecular Evolution Through errors in replication, failure to repair nucleic acid damage, and recombination, nucleotide sequences may accumulate changes over time. These changes may be divided into three categories: substitutions, indels (insertions and deletions), and recombination (Diagram 7.11). A substitution is the replacement of a nucleotide with another nucleotide. These are the most useful changes for establishing phylogeny of related organisms. If sequences

Diagram 7.11  Depiction of substitution, indel, and mutations.

recombination

share a common nucleotide from a common ancestor, that site is said to be homologous. As nucleic acids are composed of only four nucleotides, there are only three options for change of a nucleotide. The possibility of that nucleotide changing back to the original sequence exists, and is a serious confounding factor in inferring phylogenies. The term for nucleotides that are shared by two sequences that are not due to a direct common ancestor is homoplasy. We typically do not have access to ancestor sequences, and must rely on current sequences to infer phylogeny. As can be seen in Diagram 7.12, the presence of homoplasy might lead to incorrectly clustering the three sequences on the left, when ancestry actually clusters the two sequences on the left and the two on the right. Substitutions are often further subdivided into two groups: transitions (purine ↔ purine or pyrimidine ↔ pyrimidine, e.g., G↔A or C↔T) and transversions (purine ↔ pyrimidine, e.g., A↔C, A↔T, G↔C, or G↔T). Due to many reasons, including greater steric compatibility, the relatively high frequency of deamination of cytosine ultimately resulting in a C→T mutation, and the lower chance of a resultant amino acid change, transitions are typically more common than transversions (Lanave et al., 1986). An indel involves either the addition or removal of one or multiple nucleotides. While the presence of specific indels within a lineage may aid in grouping the sequences together, indels increase the difficulty of correctly aligning

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Diagram 7.12  A contrast between homology, where synonymous sites are due to common ancestry, and homoplasy, where they are not.

gene sequences, and any information that was present in a deletion is lost. Recombination involves the splicing of a gene of one lineage with a homologous gene of another lineage. This obviously creates great difficulty in interpreting gene history. When the results of phylogenetic analyses of different genes or regions of genes are incompatible, the occurrence of a recombination event should be considered. Further discussion of recombination is beyond the scope of this text. Most genes encode for proteins. Three nucleotides (a triplet codon) encode for each amino acid, and there is more than one possible set of three nucleotides coding for most amino acids (Diagram 7.13). The variability is typically found in the third nucleotide of the codon. It is therefore possible for a substitution in the nucleotide sequence not to alter the amino acid sequence, which is the functional product. These changes are known as silent mutations, and are not significantly influenced by selective evolutionary forces. Some genes encode for products that are functional as RNA molecules rather than as proteins, such as 16S ribosomal RNA (rRNA) in the bacterial ribosome. Any mutation in these genes will alter the final gene product, and there are no silent mutations. Other changes result in alteration of the functional molecule, and are subject to selection by evolutionary forces. Most of these changes at many sites will be deleterious to the functioning of the gene product, and this may result in failure of the organism to survive and reproduce. Nucleotides at such positions are said to be under forces of negative selection, and these nucleotides may be expected to change at a slower rate over time. It is also possible for evolutionary forces to promote change at a site. For example, if a protein of a pathogen is recognized by the immune system of a host, and that protein changes in such a manner that it is no longer recognized by the immune system, then change provides a selective advantage to the pathogen. Nucleotides at such positions are said to be under forces of positive selection, and these nucleotides may be expected to change at a faster rate over time. Overall, mutations that are strongly deleterious under the evolutionary forces working over a given period on an

Diagram 7.13  Triplet codons encoding amino acids. More than one codon may encode the same amino acid, and the variability is typically present in the third nucleotide of the codon.

organism are rapidly selected against and disappear from populations. Mutations that are advantageous are much less common, but if they offer a significant advantage, they may overtake a population. This means that most of the mutations that remain in the genetic record will offer very little selective advantage or disadvantage, and can be called nearly neutral. These mutations accumulate at a relatively constant rate, and can be used as a “molecular clock.” There may be significant variations in the rate of this clock for different genes and different organisms, however. There are markedly different rates of evolution among viruses. Large DNA viruses, such as adenoviruses and herpesviruses, mutate very slowly. Large DNA viruses often evolve so slowly that they may be seen to coevolve with their hosts (Benkö and Harrach, 2003; McGeoch and Davison, 1999). It is estimated that branching of the alphaherpesviruses from the beta- and gammaherpesviruses occurred between 374 and 413 million years ago (McGeoch and Gatherer, 2005). RNA viruses typically lack proofreading and evolve several orders of magnitude more rapidly. Even with generous models of evolution, it is hard to follow the evolution of most RNA viruses back farther than 100,000 years, and more conventional models of evolution often place this number at 50,000 years (Holmes, 2003). In multicellular organisms, it is only the mutations in germ cell lines (eggs and sperm) that are passed to later generations. One implication of this is that species with long generation times are expected to accumulate mutations at a slower rate than those with more rapid generation times, due to less frequent divisions of germ cell lines (Martin and Palumbi, 1993). Therefore chelonians, which have a long reproductive life span (Congdon et al., 2001), may be expected to accumulate mutations at a slower rate than some species of lizards that are shorter-lived (Dunham and Overall, 1994). Other factors may also play roles in rates of evolution in reptile hosts

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(Brondham, 2002). With the exception of some parasites, most reptile infectious diseases are not multicellular, and do not have separate germ lines of their own, so this would not apply directly. However in viruses capable of latency, such as herpesviruses and retroviruses, the host generation time may impact the rate of evolution of the virus.

7.4.3 Sequence Selection An important first step when undertaking a molecular phylogenetic study is the choice of the target sequence. The evolutionary distance that is being considered and the mutation rate of the organism must be taken into account. To follow an epidemic that is rapidly spreading through a population, a rapidly mutating gene, perhaps one under positive selection such as an antigenic coat protein, should be selected. This would be an optimal area for finding differences among closely related strains. However, if looking at evolutionary relationships of more distantly related organisms, the continued accrual of mutations resulting in homoplasy would diminish the ability to correctly resolve phylogeny, making a rapidly mutating gene a poor choice. A gene for which there is strong negative selection will have fewer nucleotides with a history of multiple changes, making it a better choice for resolving phylogeny over greater distances. Genes that are critical for basic organismal functions and are not under immune selection are often highly conserved. For these reasons, the 16S rRNA gene, which is essential for ribosomal function, has become a standard for bacterial phylogeny (Clarridge, 2004), 18S rRNA is a good choice for protozoa (Morrison et al., 2004), and viral polymerases are often good choices for long-range phylogeny (Attoui, 2002; Gonzalez et al., 2003; Knopf, 1998). Use of multiple genes may add additional power of resolution. Another factor that plays a role in gene selection is the number of available sequences for comparison. The sequence databases GenBank (National Center for Biotechnology Information, Bethesda, Maryland, www.ncbi.nlm.nih.gov), EMBL (Cambridge, United Kingdom), and Data Bank of Japan (Mishima, Shiuoka, Japan) may be easily accessed via the Internet, and search tools are available at these sites. The availability of an appropriate set of evenly related sequences is important for correctly establishing phylogeny; if some sequences are more distantly related than others, the phenomenon of long-branch attraction (which is discussed later) may occur. When an appropriate set of sequences is not available, the investigator should seek to obtain additional sequences from appropriately related organisms if possible.

nucleotides or peptides sharing ancestry can be compared. It has been shown that methods of nucleotide sequence alignment may have an even greater effect on results than methods of tree building (Morrison and Ellis, 1997). If all mutations were substitutions, alignment would be relatively straightforward. The presence of indels presents a problem — we need to insert gaps into some sequences to line up the nucleotides or peptides that share ancestry. As ancestor sequences are typically not available, we must base the alignment on the modern sequences that are available. The most reliable way to do this may involve looking at the biological function of the molecule. Sequences encoding proteins are often converted to amino acid sequences before alignment. Functional regions of a protein can be aligned manually, and it is reasonable to expect that regions of a protein sharing the same function share ancestry. Use of rRNA is even more straightforward in determining the history of nucleotides. Because the RNA is the functional molecule, every nucleotide is directly involved in the final gene product. There is extensive conserved secondary structure present, which is important for function and can be used for alignment. With some sequences, it may not be possible to create an alignment based on conserved function, and a mathematical algorithm is typically used. Alignment of a single pair of sequences is relatively straightforward. Sequence alignment can be visualized using a dot plot (Diagram 7.14). Sequences are aligned on the sides of a grid, and matches between the

7.4.4 Sequence Alignment Once a target sequence is obtained, it needs to be compared to other homologous sequences. A phylogeny can be built based on the mutations that have occurred at each nucleotide position. We therefore need to line up the sequences so that

Diagram 7.14  A dot plot of two sequences. The possible alignment indicated by the arrows is depicted beneath the dot plot.

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two sequences are indicated by a dot. Starting from the upper left-hand corner, an alignment may progress by moving one square to the right, introducing a gap in the lower sequence, one square down, introducing a gap in the upper sequence, or one square diagonally down and to the right, lining up two nucleotides without introducing a gap. The alignment indicated by the path marked by the arrows is given below the plot. Most alignment algorithms score the alignment by setting a cost for gaps, a cost for extending a gap, and a cost for mismatches. Costs for gap extension may vary depending on gap length. Cost for mismatches may also vary; some scoring algorithms look at one cost for transitions and another for transversions, and some assign different values to each possible change based on the prevalence of each nucleotide in the sequence. As there are a greater number of amino acids than nucleotides, scoring of protein sequences is more complex. Amino acids are more likely to be replaced by other amino acids that have similar chemical and physical features; matrices have been developed for mismatch scoring based on replacement rates of amino acids seen in alignments of conserved protein families. Each possible path across the dot plot may be scored, and the lowest score is considered the optimal alignment. While scoring all possible paths for two sequences across a two-dimensional grid is not too computationally intensive, the addition of a third sequence adds a third dimension to the grid, and additional dimensions are added with each additional sequence. The number of possible paths expands exponentially and rapidly becomes so large that it becomes untenable. As doing an exhaustive search (looking at all paths) is not feasible, heuristic methods, which try to find the optimal path without looking at all possible options, are used. Many methods first look at distances between each pair, and then form a guide tree by pairing the most closely related sequences and then adding additional sequences to the tree in order of distance. CLUSTAL, an early alignment algorithm that was commonly used, builds the guide tree as a template and aligns pairs of sequences at the end of branches (Higgins and Sharp, 1988) (Diagram 7.15). These alignments are used to form consensus sequences, which are then each used in alignments with the next node on the branch, and the process is repeated until all sequences have been added. Other programs have incorporated a number of improvements to alignment algorithms, amongst them differing gap penalties in conserved areas, procedures for building guide trees that assess the possibility of more than one subtree, weighting of sequences, comparison of trees built from varying guide trees, iterative alignments, and building of larger trees from smaller exact local alignments. Several comparative assessments of alignment programs have been published (Morrison and Ellis, 1997; Lassmann and Sonnhammer, 2002; Hickson et al., 2000; Thompson et al., 1999), and further discussion of alignment algorithm specifics is beyond the scope of this text. References for further reading are given at the end of this chapter. Sequence alignment is probably the area of molecular phylogenetic analysis that has the greatest need for further development.

Diagram 7.15  Depiction of a heuristic method of alignment of three sequences using a guide tree.

7.4.5 Tree-Building Methods Once a sequence has been aligned so that homologous nucleotides or amino acids can be compared, a phylogenetic tree can be built. There are a number of different methods for building trees from an alignment. Methods are often classified into two groups: distance methods and discrete methods. Distance methods score the distance between each pair of sequences in the multiple sequence alignment, much like that discussed for construction of guide trees for the alignment except that the pair is scored without further alignment rearrangements. The tree is then built from these distances. Discrete methods consider each nucleotide or amino acid site directly.

7.4.5.1 Distance-Based Trees  Two common distancebased methods include the unweighted pair group matching using averages algorithm (UPGMA) and the neighbor-joining algorithm (NJ). UPGMA is the simplest algorithm; first the closest pair of sequences is put together, and then the distances to the other sequences from each of the paired sequences are averaged (Diagram 7.16). The next closest pair of sequences is then put together, and so on. If branches contain differing numbers of sequences (for example, one

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Diagram 7.16  Construction of a distance-based tree by the unweighted pair group matching using averages algorithm (UPGMA) method.

Diagram 7.17  Construction of a distance-based tree by the neighborjoining (NJ) method.

branch contains three sequences and the other contains one), then the average takes into account the number of end nodes (i.e., one branch would count three times toward the average, the other one). Trees constructed using UPGMA have all sequences equidistant from the root (known as ultrametric), implying that all sequences have evolved at an equal rate. This does not appear to be the case in nature, and for this and other reasons, many researchers no longer favor UPGMA trees. The most favored distance-based method, NJ, is capable of producing branches of differing lengths. The NJ algorithm first calculates an r-value for each sequence, which is the sum of the distances to the other sequences. In the example given, the r-value for sequence W is 5 + 4 + 7 = 16. A second matrix of transformed distance values (shown in the lower half of the grid in Diagram 7.17) is then calculated. The transformed distance is the original distance value minus the average of the r-values of the sequences. In the example, the transformed value from sequence W to sequence X is 5 − [(16 + 18) / 2] = −12. The pair with the lowest transformed distance value is then combined. In this example, W-Y has the lowest transformed value (−14).

The distance of each sequence to the node of intersection is the untransformed distance plus the difference between each r-value divided by the number of sequences minus two. This number is then divided by two. In the example given, the distance from W to the W-Y node is the original distance (4) plus the difference between the r-values divided by the number of sequences minus two, (16/ [4 − 2]) – (20 / [−2]) = −2, giving a distance of (4 + [−2]) / 2 = 1. The distance from Y to the W-Y node is then (4 + [20 / {4 − 2} – 16 / {4 − 2}]) / 2 = 3. Although the lengths of the two branches are different, their sum equals the distance between the two sequences. The distance matrix then needs to be recalculated. The distance from the created node to the other sequences is the average of the distances to the sequences beyond the node minus the branch lengths from the nodes to the sequences. In the example given, the distance from the W − Y node to sequence X is the average of the distance from X to W minus the distance from W to the W-Y node (5 − 1 = 4) and the distance from X to Y minus the distance from Y to the W-Y node (10 − 3 = 7). The distance from W-Y node to X is then (4 + 7) / 2 = 5.5. The rest of the distance matrix is calculated, new

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r-values and transformed distances are calculated, and the process is then repeated. Note that when the final two nodes are combined, there is no inherent root node by this method. A root may be added by other methods if desired. There are other distance-based tree-building algorithms, but further discussion is beyond the scope of this text. The reader is directed to further references at the end of this chapter. A major advantage of the use of distance-based trees is their computational simplicity. The tree-building algorithm is simple enough that trees can be built from relatively large data sets in a short period of time with a personal computer. The major disadvantage of distance-based methods is loss of information. As sequences are converted into pair-based distances, information regarding evolution of individual sites across multiple sequences is not taken into consideration and is lost. Due to homoplasy, observed differences between sequences may not be accurate reflections of the evolutionary distance between them, and this error increases with distance. It is also much more difficult to incorporate variation in rates of change of different sites using distance-based methods. It is possible to evaluate all possible trees that meet the criteria of having at least the distances observed between all sequences and having no negative branch lengths. The tree meeting these criteria that has the lowest total sum of branch lengths is known as the minimum evolution tree. The distance methods discussed here, UPGMA and NJ, use an algorithm to attempt to find the minimum evolution tree. The tree resulting from the algorithm is the only tree that is evaluated. Other methods may also evaluate related trees without evaluating all possible trees. When more trees are evaluated, the chance of finding the optimal tree increases, but so does computational time. Use of algorithms to find the optimal solution without evaluating all possible solutions is known as a heuristic search. It is possible for a heuristic search to miss the minimum evolution tree. Exhaustive methods that use a criterion to evaluate all possible trees are available. While more rigorous than heuristic methods, these lose the advantage of computational speed, and keep the disadvantage of loss of information from individual sites.

7.4.5.2 Maximum Parsimony  Parsimony methods evaluate the number of changes for each nucleotide or amino acid. The maximum parsimony tree has the fewest number of changes, and is thus the simplest explanation of evolution given the data. This is a discrete rather than distance-based method. One way to look at the number of changes in a tree is to use the Fitch algorithm (Fitch, 1971) (Diagram 7.18). Each ancestor node is made from the data immediately above it. If the data above does not overlap, then a set is made from the combination of the data and a change is counted. If the data above overlaps, then a set is made from the intersection of the data and no change is counted. At the top of Diagram 7.18, there is no overlap of the data in either pair, so new sets are made from the combinations, and two changes

Diagram 7.18  Ancestry of nucleotides at the same site using the Fitch method of parsimony evaluation on a given tree. This is used to assess the number of changes at the site, which is minimized in a maximum parsimony tree.

are counted. In the middle of Diagram 7.18, one change is counted similarly. At the bottom of Diagram 7.18, the sets [A,T,G] and [A,G] overlap, so the new data set is [A,G], and no change is counted. Thus, there is a minimum of three changes, given this tree. There are a number of modifications of parsimony, including different scoring for different transitions or transversions and site weighting. There are also a large number of heuristic algorithms available to search for optimal trees without evaluating all possible trees. One of the biggest criticisms of maximum parsimony methods is the susceptibility to inferring incorrect trees due to homoplasy. In a phenomenon known as long-branch attraction, data sets containing sequences that are unevenly related may result in incorrect trees (Anderson and Swofford, 2004). More distantly related sequences will have accumulated more homoplasic changes, and may thus appear to be more closely related to each other by parsimony than they actually are, and cluster together. While this should not be a problem with data sets in which a large number of closely related sequences are available for comparison, real data sets without gaps are not often available. The data available at this time for most reptile pathogens is sparse, and most known reptile pathogens do not have other closely related sequences available for comparison.

7.4.5.3 Maximum Likelihood  Maximum likelihood methods look at the likelihood of the data turning out the way that it is found, given a tree and model of evolution. If we have a hypothesis of a tree including branchings and branch lengths, and we have a model of expected rate of evolution and the probability of changes of one nucleotide or amino acid to another, then we can calculate the likelihood of the data turn-

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ing out the way it is found. The tree that is most likely to result in the data found is the maximum likelihood tree. Obviously, correct outcome is very dependent on having a correct model of evolution, and incorrect models will result in erroneous trees. The likelihood of different model parameters can be evaluated as well. Homoplasy may occur in maximum likelihood evolutionary modeling, and thus, although homoplasy is not phylogenetically informative, maximum likelihood methods are not as easily misled by homoplasy. There are many attractive features of maximum likelihood methods of phylogeny, and a large number of molecular phylogeneticists consider them to be the best methods currently available. However, maximum likelihood methods are mathematically more complex, and as a result, much more computationally intensive. While there are heuristic methods of searching for the optimal tree, some of the heuristic methods applied to maximum parsimony are not as easily applied to maximum likelihood. As a result, maximum likelihood methods tend to take a lot of computational time, and evaluation of large numbers of longer sequences can be computationally daunting. For further details on the methodology of maximum likelihood phylogeny, the reader is referred to further references given at the end of this section.

7.4.5.4 Bayesian Methods  Bayesian methods have similarities to maximum likelihood methods, but Bayesian methods are built around posterior probabilities. A posterior probability takes into account the prior probability or what the results are expected to be before the data is analyzed. The posterior probability of a tree is the likelihood of the tree multiplied by the prior probability of the tree, divided by the sum of the likelihood of all trees multiplied by the prior probabilities of all trees. To find the highest posterior probability, this can be integrated. Parameters such as tree topology, branch lengths, and substitution parameters can all be modeled as probability distributions. Bayesian methods are mathematically complex, and it is not feasible to calculate them directly. More rapid heuristic methods are used. The method used for approximating the value of the integral is the Metropolis-coupled Monte Carlo Markov chain. The simplest way to visualize this is to look at a rooted tree with two sequences and a set invariant model of evolution. The only parameters that can vary are the two distances from the root node to the sequences. Possible values for the length of each branch can be graphed, with one branch length on the x-axis and the other on the y-axis. Each point will have a different posterior probability, which can be thought of as topography on the z-axis. The goal is to find the “high point.” A Metropolis-coupled Monte Carlo Markov chain starts at one point and calculates the posterior probability at that point. A nearby point is then randomly selected, and the ratio of the posterior probability of the new point and the posterior probability of the old point is calculated. If the ratio is greater than one, then the new point is more probable (which may be thought of as “uphill” on the topology) and becomes the new reference point used as the

base for selecting a new point. If the new point is less probable (“downhill”), then the ratio of the posterior probability of the new point and the posterior probability of the old point is the chance that the new point will become the new reference point. Thus if the new point is uphill, the chain moves there, but if the new point is downhill the chain may move there, but the steeper it is, the less likely it is to move. If the algorithm does not move, a new point is selected near the old point. This algorithm therefore tends to wander uphill, but can go downhill as well, which is useful when more than one peak is present. A large number of iterations are run. Early iterations are considered a burn-in period and discarded. The path may be expected to spend the most time in areas of highest posterior probability. The fraction of time that the chain is at any particular tree is an approximation of the posterior probability of the tree. The process is often repeated several times with different starting points. Any real tree to be evaluated would have more than two variable parameters, and thus more dimensions, but the concept is the same. The computational advantage of this algorithm is that when calculating the ratio of the posterior probability of the new point and the posterior probability of the old point, the denominator in both terms (the sum of the likelihood of all trees multiplied by the prior probabilities of all trees) cancels out. This denominator is the most computationally intensive part of posterior probability calculation. Bayesian methods provide the advantage of being able to use complex evolutionary models, and can be calculated more rapidly than maximum likelihood methods. The most controversial part of Bayesian methods is the incorporation of prior probability. From one perspective, it seems reasonable to incorporate aspects of expectations based on data not included in the set, and it is not unreasonable to expect an iguana to be more closely related to a chuckwalla than to Escherichia coli. On the other hand, subjectivity is potentially inserted into the data, and, philosophically, many feel that the data should stand alone. One way of dealing with this is to use flat prior probabilities, where all trees are given equal values for prior probability. As more data is added to a Bayesian analysis, the influence of prior probabilities decreases (Huelsenbeck et al., 2002). Other concerns about Bayesian methods include a lack of information on how far apart one point on a Markov chain should be from the next and the number of iterations that are needed. For further details on Bayesian methods of inferring phylogeny, the reader should consult the references given at the end of this section.

7.4.6 Methods of Measuring Confidence For any inference made from a set of data points, it is important to have confidence measures of the inference. For example, the data sets (100, 99, 100, 102, 99) and (46, 125, 129) both average 100, and if the average is the only number reported, then the person looking at the report would not be able to distinguish how reasonable it is to expect that the result of

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each data set would not change with further information. For sets of numbers, we often use standard deviations. Given the results of the data sets as 100 +/− 1.2 and 100 +/− 47, it is more reasonable to expect that the first data set has given the stronger prediction. Similarly, if a phylogenetic tree is given without any confidence measures, we have no way to evaluate the strength of that prediction. It is unfortunately common in much of the literature to present trees with no confidence measures. A number of methods have been developed to give confidence measures. It is important to note that current commonly used methods of measuring confidence intervals only consider the uncertainty in tree building after the alignment is made. The uncertainty in the alignment is not included, and so errors of alignment may result in false confidence intervals. Initial efforts have been made to incorporate the estimation of uncertainty in both alignment and tree building, but incorporate possibly unrealistic assumptions, such as assuming indels are of unit length (Lunter et al., 2005) or that indels are equally likely to occur on branches of any length (Redelings and Suchard, 2005).

7.4.6.1 Bootstrapping  Bootstrap methods look at how often phylogenetic information is replicated in subsets of the data. A number (often 100 or 1000) of random subsets of the sequence alignment are created (Diagram 7.19). Trees are then created from these subsets, and the number of trees built on these subsets in which a group of sequences clusters together is determined. The bootstrap values may be expressed as a percentage of subsets or as absolute numbers of subsets. If 100 subsets are used, the number is the same. Bootstrap values do not correspond directly to the probability of a clustering being real, and are considered to be a conservative measurement. Values less than 50% are considered insignificant. There is less consensus on values that are considered highly significant; bootstrap values of 95% or greater are considered highly significant by some. However, others have found that a bootstrap value of 70% correlates to a 95% probability of a clade being real (Hillis and Bull, 1993). A low bootstrap value means that a grouping is sensitive to the combination of sites that is evaluated, implying that more data may alter the grouping. In the tree displayed in Diagram 7.20, reptile virus A and reptile virus B cluster together in 98% of the trees made from the subsets of data, and there is strong bootstrap support. Reptile viruses A, B, and C cluster together only 65% of the time, so support for this grouping is less strong. Bootstrapping is probably the most commonly used measure of confidence in trees. One caveat when interpreting bootstrap values is that if an inappropriate tree-building method or data set is used, then bias present in the whole set may be reflected in the subsets. For example, if several more distantly related sequences are present in a data set, then maximum parsimony evaluation of subsets is likely to provide support for incorrect trees due to long-branch attraction.

Diagram 7.19  Creation of subsets of data for bootstrapping from a sequence alignment.

Additionally, bootstrapping assumes that each site evaluated is independent and identically distributed. Real molecular sequence data sets are typically neither. Rates of change differ at different sites, and pair bonding of nucleotides in RNA or interactions of amino acids in proteins mean that a change in one site may not be independent of other sites. The major drawback to bootstrapping is that it is computationally intensive. While initial calculation of a tree involves one data set, bootstrapping involves calculating trees for hundreds or thousands of data subsets. This may not be a problem with more rapid methods of tree building, but can be a large demand when more complex methods, such as maximum likelihood,

Diagram 7.20  A phylogenetic tree displaying bootstrap values. Reptile virus A and reptile virus B cluster together in 98% of the trees made from the subsets of data, and there is strong bootstrap support. Very strong support is present for A, B, C, and D grouping together. Reptile viruses A, B, and C cluster together only 65% of the time, so support for this grouping is less strong, and the reader should not completely exclude the possibility that C branches off from A and B before D does.

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are used. The number of replicates used in bootstrapping typically still ends up being less than the number of links in a Metropolis-coupled Monte Carlo Markov chain used with Bayesian methods, which often number in the millions.

7.4.6.2 Bayesian Posterior Probabilities  Construction of trees using Bayesian methods inherently provides a measure for confidence intervals. Posterior probabilities used represent an estimation of the probability that a clade is real given the prior probability, model, and data. Posterior probabilities tend to be higher than bootstrap values (Huelsenbeck et al., 2002; Alfaro et al., 2003; Douady et al., 2003; Erixon et al., 2003). Bayesian posterior probabilities are a better estimator of accuracy than bootstrap values for support values greater than 50%, so a posterior probability of 95% is more likely to correspond to an accuracy of 95%, whereas a bootstrap value is likely to be lower (Alfaro et al., 2003). Error rates of posterior probabilities increase when an incorrect model is used (Erixon et al., 2003). The greatest disparity between bootstrap values and Bayesian posterior probabilities is found on short internodes, and posterior probabilities are likely to be much higher on short internal branches (Alfaro et al., 2003). The risk of error using Bayesian methods on these short internodes is therefore higher, and those interpreting trees should be aware of this. Bootstrap values and posterior probabilities do not convey the same information; a high posterior probability implies that a branch is likely correct given that the data is representative, whereas a bootstrap value is suggestive of how susceptible results are to being altered by further data. Bayesian methods tend to be more rapid than maximum likelihood bootstrapping, and have been reported to run 80 times faster using popular phylogenetic programs (Douady et al., 2003).

al., 1969). In situ hybridization is based on the annealing of a labeled nucleic acid probe to complementary sequences in fixed tissues. This allows localization of the target DNA or RNA within tissue sections or cells (Diagram 7.21). When in situ hybridization was first being performed, a radioactive labeled probe was used and autoradiography was used to visualize the target, but fluorescent and colorimetric techniques are now being more commonly and safely used. There are many different methods and protocols for performing in

7.4.7 Further Reading For a more in-depth introduction to molecular evolution, the most easily read text we have found that does not require the reader to have prior background is Page and Holmes (1998). However, this text does not include some more recent developments and contains no information on Bayesian methods. An excellent reference text is Felsenstein (2004), although this text assumes that the reader has a background in the subject and is not organized for a novice reader. This author is not a big advocate for Bayesian methods, although they are presented in the text. For a good overview of Bayesian methods of phylogeny, see Huelsenbeck et al. (2002).

7.5 In Situ Hybridization 7.5.1 Introduction In situ hybridization (ISH) was first performed in the late 1960s and is based on the formation and detection of nucleic acid (DNA and RNA) hybrids (Gall and Pardue, 1969; John et

Diagram 7.21 (A, B)  A schematic depiction of in situ hybridization is shown. (A) shows hybridization to a cytoplasmic target and (B) shows hybridization to an intranuclear target.

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situ hybridization and each of them requires optimization before it can be considered reliable. In situ hybridization has been used in reptiles for chromosome mapping and gene expression since the early 1990s (Valleley et al., 1994; Young et al., 1994), but only in recent years has it been used for pathogen identification in tissue sections. In situ hybridization has been used in identifying adenovirus DNA in snakes (Ramis et al., 2000; Perkins et al., 2001; Raymond et al., 2003), herpesvirus DNA in tortoises (Teifke et al., 2000) and monitors (Wellehan et al., 2005), and paramyxovirus RNA in snakes (Sand et al., 2004). In situ hybridization is likely to become more commonly used for pathogen detection as more sequences of pathogens become available. The basic steps of ISH are (1) probe synthesis, (2) tissue preparation, (3) pretreatment of tissue to make it permeable to the probe, (4) hybridization of the probe to the target, and (5) detection of the annealed labeled probe (Diagram 7.22). In situ hybridization protocols vary by the type of probe, the type of label to incorporate onto the probe, and the method of visualization.

7.5.2 Probes The probe is the reverse complement of the target sequence, so the probe and target sequence will bind once they come into contact. It can be double-stranded DNA, single-stranded DNA, single-stranded RNA, or synthesized oligonucleotides (Wilkinson, 1998). The stringency of the reaction will determine the quality and the specificity of the signal. If the probe matches the target at 100%, the probe is likely to bind optimally and produce an intense signal. However, when hybridizing under stringent conditions, slightly mismatching related sequences will likely remain undetected. This problem can be overcome by designing a degenerate probe, which may cross-hybridize

Diagram 7.22  A schematic representation of the in situ hybridization procedure is shown.

with similar target sequences found in related species. Binding of degenerate probes may not be as stable, creating a weaker signal. The use of degenerate probes may also be associated with false positive results or with masking of true positives because of high background. The more conserved the nucleotide sequence, the better the chances of a probe hybridizing with different strains of the pathogen. The length of the probe also influences the stringency of the probe binding. Longer probes will have a more incorporated label, which will create a stronger signal and bind more specifically to the target. However if the probe is too long, it may not penetrate as well into the tissue to reach the target. The ideal length depends on the type of tissue, the fixative used, and the pretreatment of the tissue prior to hybridization. However, an average of 300 base pairs has been considered optimal (Wilkinson, 1998). The probe label allows the detection of the targeted nucleic acid. There are two categories of labels: radioactive and nonradioactive labels. Radioactive labels are more hazardous to use, but may have higher sensitivity, allowing the detection of the target when it is present in a lower copy number. The most commonly used isotopes are 3H, 32 P, 33P, and 35 S. Nonradioactive labels are more convenient, stable, safer to use, and less time consuming. The most commonly used nonradioactive labels are biotin, digoxigenin, and fluorescein. Double-stranded DNA probes can be labeled using nick translation, random priming, or by PCR. Large probes are often labeled by nick translation. Nick translation utilizes deoxyribonuclease I to create smaller fragments by creating little nicks in the DNA. DNA polymerase is then added, which will remove nucleotides from the 5’ end and add labeled nucleotides to the 3’ end. Random primer labeling can also be performed. In this case, double-stranded DNA is denatured to separate the two strands. A mixture of short random primers is added, which will bind where they find complementary sequences. A polymerase is added that will fill in the missing bases with labeled bases, creating a second strand of labeled DNA. The original strand remains unlabeled. In PCR labeling, some of the free dNTPs are labeled and incorporated into the newly synthesized strands during amplification. The advantage of this procedure is that the product can be made in large quantities with only a small amount of original template. Single-stranded DNA probes can be labeled by primer extension on single-stranded templates, which is not typically performed, or by asymmetric PCR, which is similar to the technique for PCR labeling. PCR labeling creates short segments of double-stranded DNA in which both strands are labeled. Asymmetric PCR uses only one primer, and thus labels only one of the strands of DNA. This strand is the reverse complement of the target sequence and will be highly specific. A disadvantage is that amplification of the labeled sequence will not be exponential but will be determined by the number of amplification cycles, so fewer copies will be made. Oligonucleotide probes can be synthesized with labels. The advantages of synthesized probes are that they can be created from

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a published sequence without having a copy of the template, and the procedure is commercially available. Single-stranded RNA probes are labeled by in vitro transcription and are most often used to detect RNA. To produce probes, DNA is cloned into RNA transcription plasmid vectors. Vectors are circular pieces of extrachromosomal DNA that are designed so that foreign pieces of DNA can be inserted at specific locations and replicated. The DNA is inserted downstream of an RNA polymerase promoter site, which signals the polymerase to start transcription at that site in the direction of the inserted DNA. RNA polymerase promoters can be placed on one or both sides of the inserted DNA, so that one or both directions can be transcribed: the target sequence, which can act as a negative control to detect nonspecific binding, and the reverse complement of the target sequence, which will hybridize to the target sequence and produce a signal.

7.5.3 Tissue Preparation Tissue preservation is critical for successful in situ hybridization. Similar to PCR, the DNA needs to remain intact. However, it also needs to be prepared in a way that can be detected. Cross-linking fixatives, such as formaldehyde, paraformaldehyde, and glutaraldehyde provide good tissue preservation, but can be degrading to DNA if fixation time is extended. Precipitating fixatives, such as alcohol and methanol, are better for maintaining nucleic acid intact, but may increase nonspecific binding, which will result in high background (Morel and Cavalier, 2001). Ideally, tissue should be fixed and embedded into paraffin as soon as possible after necropsy or biopsy. Sections can be cut from the paraffin and dewaxed prior to pretreatment. Pretreatment of tissues is performed to help make the target sequence more accessible to the probe and to decrease nonspecific binding of the probe, reducing background signal. Common pretreatment steps include permeabilization, deproteinization, acetylation, and denaturation of the probe and target nucleic acid. Ethanol or methanol can be added to remove lipid membranes to make the target more accessible; however if tissues are embedded into paraffin, this step is performed when removing the wax. Protease digestion will partially remove proteins that may inhibit access of the probe and could increase background. Acetylation can help prevent nonspecific binding of the probe to certain types of slides, remove endogenous biotin, and inactivate the proteases (Schwarzacher and Heslop-Harrison, 2000). Denaturation of the probe and target nucleic acid is necessary to separate the strands of DNA so that the probes can bind to the target sequences.

7.5.4 Hybridization Hybridization is the process of the probe annealing to the target sequence and should be carried out at the appropriate stringency. Stringency refers to the stability of the hybrid. For example, at 65% stringency, a hybrid where 65% of the bases match with the complementary base, and 35% of the bases

are mismatched, will remain stable. This would be considered a low stringency. A higher stringency would be at 90% where only hybrids with 90% of the bases matched and 10% of the bases mismatched will remain stable. By altering the stringency conditions, it is possible to set the specificity of the hybridization reaction. With specific probes, the hybridization reaction can be run under more stringent conditions, whereas with less specific probes, use of less stringent hybridization conditions should be expected. Stringency is therefore controlled by temperature, base composition, specificity and length of the probes, type of hybrid being produced, the composition of the hybridization solution, monovalent cation concentration, and formamide concentration (Schwarzacher and Heslop-Harrison, 2000). Typically, the temperature and hybridization solutions are altered (most often by varying concentrations of sodium ions and formamide) according to the predetermined stringency of the reaction. With higher concentrations of ions, DNA is more stable, where at lower concentrations it is less stable. Formamide also destabilizes DNA. Stringent washing after hybridization removes excess probe that is either not bound or weakly bound so as to reduce background signal. Washing at appropriate stringency conditions is critical to get the optimal signal and is usually either just above or just below the stringency conditions used for hybridization. If temperatures are not appropriate, the label can be removed or there may be an increase in background signal.

7.5.5 Signal Detection Detection will depend on the type of label used. Radioactive labels will require one of two types of detection. The slide can be exposed directly to radiographic film, which gives an idea of hybridization at the tissue level, or a photographic emulsion, which localizes the signal at the cellular level, can cover the slide. Once tissues are covered in emulsion, they are stored in darkness for a variable length of time depending on the isotope used. These emulsions capture the radiation being emitted from the hybrid. The radiation changes the silver salts of the emulsion into metallic silver grains that can be visualized by light microscopy. Detection of nonradioactive labeled probes takes a different approach. Probes labeled with digoxigenin must be bound to an antibody specific for the hapten, or to avidin when biotin labeling is used. These antibodies are generally directly conjugated either to fluorophores, which can be visualized under fluorescence microscopy, or to enzymes, which require a specific substrate for a colorimetric reaction to occur (Figure 7.2). For imaging using electron microscopy, the probe or secondary antibody can be conjugated to metals such as gold (see Chapter 6).

7.5.6 Limitations In situ hybridization allows the detection of pathogens in tissue sections. However, much care needs to be taken when

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interpreting both positive and negative results. Similar to PCR and RT-PCR, ISH results are dependent on the integrity of DNA or RNA in the tissues, in addition to how tissues and slides are treated prior to hybridization. Results will also depend on the stringency at which hybridization and washing is performed. Optimization of the test is essential prior to using ISH as a diagnostic tool. Samples should always be run with known negative and positive controls to distinguish signal from background hybridization. Samples should also be compared with routine H&E stained sections to look for natural pigmentation in tissues that could appear as signal on ISH.

7.6 2D-PAGE 7.6.1 Introduction Two-dimensional polyacrylamide gel electrophoresis (2DPAGE) is a technique used to separate proteins by both their isoelectric point (pI) (first dimension) and by their mass (second dimension). This technique has been known since the 1970s (O’Farrell, 1975), but it is only in recent years that it has become more popular with the advent of proteomics (Wilkins et al., 1996). The 2D-PAGE is far more versatile and informative than conventional one-dimensional protein gel electrophoresis (1D-PAGE). In addition to the ability to separate out approximately 10,000 proteins starting from a single protein pool, 2D-PAGE also allows the investigation of posttranslational modifications that proteins undergo in the cell. This feature has made the 2D-PAGE a very attractive tool for comparison of protein modifications in healthy and diseased organisms. The coupling of 2D-PAGE with mass spectrometry (MS) has further enhanced the power of this technique, allow-

ing specific identification of the proteins that are resolved by 2D-PAGE.

7.6.2 The Procedure The same procedure that was described for sample preparation in the western blot is appropriate for 2D-PAGE. Isoelectric focusing is performed using an acrylamide gel with a pH gradient. In the gel region characterized by an acidic pH, the carboxylic groups of the proteins have no net charge, while the basic groups tend to be positively charged. In the basicpH portion of the gel there is the opposite situation, where the carboxylic groups of the proteins are negatively charged and the basic groups are not charged because they are not dissociated. The net positive or negative charge makes the proteins move along the gel in the (relative) opposite direction until they reach the region of the gel with a specific pH where the proteins lose their net charge. The proteins are therefore separated according to their isoelectric point (first dimension) (Diagram 7.23). In the next step, the proteins are further resolved by their mass in a conventional SDS-PAGE in a direction perpendicular to the isoelectric focusing. The final gel is stained with Coomassie blue or other staining detection procedures. Analysis of thousands of protein spots on 2D-PAGE is greatly aided by computerization. It is possible to compare gels that have been obtained from samples derived from the same tissue, cell, or other material under different conditions. The statistical analysis of the results provides additional insights to the research investigation. 2D-PAGE can also be used to identify specific or unknown proteins. Any of the resolved spots can be cut out of the gel and analyzed by MS (Diagram 7.23). It is possible to then compare the results to those of available databases and 

Diagram 7.23  A schematic representation of the 2D-PAGE procedure is shown. Proteins extracted from tissues (conditions) A and B are first isoelectrically focused (first dimension) and then resolved in a SDS-PAGE (second dimension). The SDS-gels are then stained and the spots of interest are excised for MS analysis.

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identify the protein that is being investigated. When no match is available, the protein can be further analyzed by a tandem MS system that allows the fine separations of different peptide fragments, which are then sequenced. The amino acid sequence that is obtained is eventually compared to those of the available databases for the definitive identification of the novel protein.

7.6.3 Interpretation of Results, Pitfalls, and Limitations 2D-PAGE is a very powerful technique, but requires experience and expensive equipment for the analysis and MS. Several core laboratories are currently present in many institutions performing this technique.

7.6.4 Advantages and Disadvantages Two-dimensional gel analyses can be very expensive, especially when protein identification by MS is involved, but can give information that cannot be obtained otherwise. In contrast, when only a comparative study is needed, then the cost is lower and affordable for many laboratories. 2D-PAGE analysis could play an important role in research on diseases of reptiles. Unfortunately, limited funding opportunities in this field have not allowed wide application of this technique to date. Nevertheless, a first example of the usefulness of this technique was recently published (Coberley et al., 2002) on the identification of a sea turtle herpesvirus protein recognized by the hyperimmune plasma of exposed sea turtles. We expect further proteomic-based research in the future.

7.7 Arrays 7.7.1 Introduction In the late 1990s, Southern, northern, and western blotting technologies were miniaturized and applied to create array technology. Using the principles and the knowledge that was previously acquired with the blot technologies, a whole new way of approaching the investigation of gene and protein expression was born. In array technology, the membrane used in blot is replaced by a glass slide or a chip where thousands of unique nucleotide sequences or peptides can be bound and used to screen for the possible target on a scale previously unknown using conventional blot technology.

7.7.2 Gene-Expression Arrays Gene-expression microarrays consist of rows and columns of thousands of different DNA sequences (or features) either spotted on a glass surface (oligonucleotide and cDNA microarray) or directly synthesized on-chip (GeneChip®, Affymetrix, Santa Clara, CA). There are three types of microarrays cur-

rently used in research: The GeneChip® array, the cDNA array, and the oligonucleotide array. The GeneChip microarray consists of several thousand oligonucleotides of 25 nucleotides in length, which are directly printed on the chip surface using photolithographic solid-phase chemistry. In the cDNA microarray, the DNA sequences consist of PCR-generated cDNA clones spotted on a glass surface by a robot. An oligonucleotide microarray consists of a collection of oligonucleotide sequences directly spotted on a glass surface using an ink-jet system. While multiple sequences are generally present for a single mRNA transcript on the GeneChip; cDNA microarrays typically have one feature for each gene sequence tested. It is currently possible to place up to 19,000 DNA sequences on a single glass slide (Hegde et al., 2000), but the number can vary according the equipment used to spot the cDNA. The number of features that can be printed on a Gene-chip can be in the range of the hundreds of thousands thanks to its special chemistry. Additionally, on GeneChip microarrays there is a set of perfect match sequences and the corresponding mismatch sequences (each having a central mismatched base) for cross-hybridization subtraction.

7.7.2.1 The Procedure  The DNA, either spotted on the glass slide or directly synthesized on the chip, represents the target for the cDNA (or cRNA) probes. The probes are prepared from the RNA previously extracted from the tissue or the cells to be tested, using conventional technologies. These probes are labeled with fluorescent dyes (Hegde et al., 2000) and are expected to hybridize with the correspondent complementary sequence of the DNA target(s) that have been spotted (or printed) on the array. After hybridization, the array is scanned under a confocal laser to measure fluorescence intensities, which allows the simultaneous determination of the relative expression levels of all the genes represented in the array (Hegde et al., 2000). Because this technique is frequently used to detect differences between a treated and an untreated population, it is common to use different dyes to differentiate probes from the different populations, and to use a mixture of the two to hybridize with the DNA sequences spotted on the array (Diagram 7.24). The result of the hybridization will give rise to a series of spots with different colors (mixture of the two dyes in different proportions) and intensity. Colorimetric and intensity differences are then transformed into numeric values that reflect the ratio with which each probe hybridizes to an individual array element (Armstrong and Van de Wiel, 2004; Hegde et al., 2000; Van der Spek et al., 2003). Mixed hybridization is not possible with the GeneChip microarray, but this problem can theoretically be overcome through determination of the absolute amount of expression of each gene tested.

7.7.2.2 Interpretation of Results, Pitfalls, and Limitations  Array technology is a powerful technique and can be a great tool once the features of each variant of this tech-

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Diagram 7.24  A schematic representation of a gene-expression array is shown. The total RNA that was extracted from two tissues (conditions) is first reverse transcribed and labeled with one of two specific dyes (one for each condition). The labeled cDNA (or cRNA) probes are then hybridized with the complementary DNA sequences, which have been previously immobilized on the array. The different probes (from tissues A and B) will hybridize with different kinetics, giving rise to a different color for each DNA spot of the array. These different color reactions will then be scanned by a confocal laser apparatus and analyzed with specific software programs.

nology are understood. Possible limitations and pitfalls differ according to the type of array that is used. Each of these systems has potential limits. The results that are obtained with cDNA arrays can be influenced by the variation in the concentration of the DNA material spotted on a slide, differences in the length of each spotted sequence, pin-to-pin variation, and spot-size variation. Additionally, sequence mismatching can be problematic in GeneChip arrays. The quality of the reagents and of the RNA used is always critical. For optimal results and reproducibility, 50 to 100 µg of total RNA from each sample is needed (Hegde et al., 2000). Data normalization and analysis are two other important steps. Data normalization removes any source of error that could cause variation in the results, such as unequal labeling efficiency of the dyes, inappropriate background correction, and variation in spot size or intensity. Several software applications are available for data analysis.

7.7.2.3 Advantages and Disadvantages  Array technology is becoming more and more popular and shows great potential. Currently, application of this technology in reptile medical research is hampered by the lack of available reptile gene sequences. With so few gene sequences currently available, there is not a sufficient database for reptile array development. This will change as more sequences become available. Sequencing of the first reptile genome, Anolis carolinensis, is underway, and additional reptile genomes may be expected in the future. Another serious problem is the cost of this technique.

7.7.3 Protein Arrays Protein arrays are to proteins what gene arrays are to DNA. A protein microarray consists of antibodies, proteins, protein fragments, and peptides in rows and columns, often on a glass slide. The immobilized proteins are used to screen for protein–protein interaction with samples containing a mixture of known or unknown proteins that have been obtained from samples such as tissues, cells, or culture supernatants. Currently, protein arrays include the sandwich antibody microarray, antibody microarray, reverse array, and peptide arrays.

7.7.3.1 The Procedure  Procedures for protein arrays vary according to the array selected. The procedure adopted for the sandwich antibody microarray is based on that used in a sandwich ELISA (see Chapter 8). A collection of antibodies (capture antibodies) is attached in rows and columns to the glass slide surface. Each of the glass-bound antibodies recognizes a single protein determinant of a possible protein target. The test sample, which consists of a pool of proteins, is then overlaid on the glass slide. A protein–protein interaction is then supposed to occur between (theoretically) any of the proteins in the sample and one of the antibodies on the glass surface. Following protein binding, a second set of labeled antibodies paired with the first set of glass-bound antibodies is added for detection. Each of the antibodies of this second set is expected to bind to a different epitope of the protein than the cognate (paired) glass-bound antibody. The signal emitted by the positive sample is then detected with specialized equipment (Nielsen and Geierstanger, 2004). 

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If paired antibodies for pools of proteins are not available, antibody arrays can be an alternative to sandwich antibody arrays. Here, the glass slide is covered with a set of capture antibodies, and the sample containing a pool of labeled proteins is added to the array. The specific antibody–protein interaction is then detected and identified. In reverse arrays and peptide arrays, proteins are directly attached to the glass surface. In reverse arrays, multiple proteins are attached to the glass surface and can be probed with one labeled antibody, while on peptide arrays several small peptides from one or more proteins are fixed on the glass surface. Peptide arrays are useful to profile a specific protein activity or map a specific protein.

7.7.3.2 Interpretation of Results, Pitfalls, and Limitations  Many problems can occur with protein arrays, starting with the challenge of array production itself (Nielsen and Geierstanger, 2004). The limited availability of antibodies and of paired antibodies for the desired pool of target proteins, and the time-consuming selection of paired antibodies are serious problems. The main challenge remains the handling of the many reagents in the microarray (Nielsen and Geierstanger, 2004). Using many different capture antibodies, each directed against a different protein to be bound and captured, can be challenging. Further, optimal binding conditions for a specific antibody often differ from those of another antibody. Additionally, the shelf life of antigens and antibodies is limited, making it difficult to ensure that all the reagents are performing optimally in every experiment (Nielsen and Geierstanger, 2004). Finally, concerns about protein quality and antibody specificity discussed in the western blot section also apply to protein microarrays.

7.7.3.3 Advantages and Disadvantages  Protein microarrays have great potential for reptile medical research. Unfortunately, there is a lack of data on protein sequence and availability for most if not all the reptile proteins of potential interest. A reptile protein database needs to be built first, and this is likely to be a lengthy and expensive task.

Glossary of Terms Bayesian: :  A school of statistical thought that incorporates prior probabilities to look at posterior probabilities. bootstrapping:   A test of rigorousness that involves evaluating multiple subsets of data. discrete:   Tree-building methods that evaluate each individual nucleotide or amino acid, as opposed to distance-based methods. Examples include maximum likelihood and maximum parsimony. heuristic:   Using an algorithm to find a shortcut to optimal conditions, rather than evaluating all possible conditions for optimality.

homology:   Traits that are shared due to a common ancestor. homoplasy:   Traits that are shared that are not due to a common ancestor. This is a confounding factor when assessing phylogeny. indel:   An insertion or deletion in a sequence. These are a confounding factor when aligning sequences. likelihood:   The chance that a model will result in a given data set. Note that the likelihoods of all models resulting in a given data set do not sum to one, unlike probabilities. minimum evolution:   The distance-based tree consistent with the data that has the shortest possible total branch lengths. monoclonal antibodies:   Any of the highly specific antibodies produced in large quantity by the clones of a single hybrid cell formed in the laboratory by the fusion of a B-cell with a tumor cell. neighbor joining (NJ):   A heuristic distance-based treebuilding method. polyclonal antibodies:   A mixture of antibodies of multiple subclasses that recognizes multiple antigenic determinants as a result of an in vivo immune response to antigenic stimulation. posterior probability:   The probability of a tree, given the data. This is calculated using the likelihood of the tree multiplied by the prior probability of the tree, divided by the sum of the likelihood of all trees multiplied by the prior probabilities of all trees. transition:   A nucleotide change from a purine to a purine or a pyrimidine to a pyrimidine (e.g., G↔A or C↔T). transversion:  A nucleotide change from a purine to pyrimidine or a pyrimidine to a purine (e.g., A↔C, A↔T, G↔C, or G↔T). ultrametric:   A tree that assumes all sequences are equidistant from the root. This implies that all sequences have evolved at the same rate, which often does not appear to be the case in nature. unweighted pair group matching using averages (UPGMA):  An ultrametric heuristic distance-based tree-building method.

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Figure 7.1  500-base pair PCR products resolved in a 1% agarose gel by electrophoresis. Lane 1 shows a 100-base pair (bp) ladder to allow visual determination of the size of the PCR product. The bright band in the ladder is 500 bp and each band is 100 bp larger (closer to well) or smaller (further from well). PCR bands have migrated through the gel at a similar rate as the known 500-bp DNA fragment in the ladder.

Figure 7.2  A photomicrograph of in situ hybridization performed targeting herpesvirus DNA in a green tree monitor (Varanus prasinus). The dark coloration represents the herpesvirus antigen. Courtesy of Kenneth S. Latimer.

8 Serodiagnostics Elliott R. Jacobson and Francesco C. Origgi

Contents

8.1 General Comments

8.1 General Comments................................................ 381 8.2 Types of Serological Assays.................................. 382 8.2.1 Serum Neutralization Test......................... 382 8.2.2 Hemagglutination and   Hemagglutination Inhibition.................... 383 8.2.3 Enzyme-Linked Immunosorbent   Assay (ELISA)............................................. 383 8.2.4 Immunofluorescence Test......................... 384 8.2.5 Immunoperoxidase................................... 384 8.2.6 Western Blot Assay.................................... 384 8.3 Titer, Titrations, Quantitations, and Pitfalls.......... 385 8.4 Establishing Test Cutoffs....................................... 385 8.5 Sensitivity, Specificity, Positive Predictive   Values, and Negative Predictive Values................ 386 8.6 Collection and Handling of Blood Samples ....... 386 8.7 Serology for Viral Exposure.................................. 387 8.7.1 Herpesviruses............................................ 387 8.7.2 Iridovirus Infection of Chelonians........... 388 8.7.3 Paramyxovirus........................................... 388 8.7.4 Reovirus..................................................... 389 8.7.5 Arboviruses................................................ 389 8.7.6 Inclusion Body Disease of   Boid Snakes............................................... 389 8.8 Serology for Bacterial Exposure........................... 390 8.8.1 Mycoplasmosis........................................... 390 8.8.2 Leptospira.................................................. 391 8.8.3 Coxiella...................................................... 391 8.9 Serology for Parasite Exposure............................. 391 8.9.1 Cryptosporidiosis....................................... 391 8.9.2 Spirorchidiasis............................................ 391 8.10 Factors Affecting the Immune Response   in Reptiles.............................................................. 392 Acknowledgments............................................................ 392 References......................................................................... 392

Serology is based on the principle that most foreign molecules are capable of eliciting a distinct humoral immune response in vertebrates (except Agnatha) that can be assayed. In serology, antigen–antibody complexes can be measured and the findings used to aid in diagnosing the basis for illness and disease in animals. Typically, in order to serologically confirm the presence of a specific disease, paired samples (one obtained during the acute phase and one obtained during the convalescent phase) are needed to show seroconversion. For viruses, a fourfold or greater rise in antibody titer between acute and convalescent samples is indicative of recent infections (Murphy et al., 1999). Serologic assays are powerful tools for both diagnosing and screening individuals and populations of domestic animals and free-ranging wildlife. Assays may either measure the presence of the antibody itself directly by specific binding to the antigen or indirectly by measuring or determining changes in the antigen such as what occurs when antigen clumps or precipitates from solution. Originally, antibodies were measured in the serum of immunized humans and because of this, these assays were called serological assays. Today, plasma, lymph, cerebrospinal fluid, and other fluids can also be used as samples to be assayed serologically. For purposes of this chapter, serum and plasma are considered as the same type of sample. Serology is a powerful tool that allows animals or humans to be screened for exposure to an almost infinite array of foreign proteins. Often the result of a serological test is used to make a decision that might have an important impact either from a clinical or conservational point of view. Following the development of a hemagglutination inhibition test to determine exposure of snakes to a paramyxovirus (Jacobson et al., 1981), additional serologic tests became available to assess the humoral response of reptiles to new and important pathogens All tests were developed in research or governmental laboratories where investigators were interested

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in diseases of captive and free-ranging reptiles. Because of a somewhat limited need, none of these tests is commercially available. Tests have been developed to determine exposure of snakes and lizards to paramyxovirus, herpesvirus infection of marine turtles and tortoises, iridovirus of tortoises and box turtles, West Nile virus of crocodilians, mycoplasma of tortoises and crocodilians, cryptosporidium of snakes, and spirorchiid trematodes of sea turtles. These tests will be reviewed in this chapter. Of the various serologic tests, the enzyme-linked immunosorbent assay (ELISA) is becoming the serologic test with the most wide-ranging application. While ELISA is relatively simple and has certain positive attributes, such as high sensitivity, in the indirect format it does require specific antireptile immunoglobulins that will recognize and bind to the antibody being assayed. It takes about six months to one year and approximately $7,000.00 to $10,000 to purify, produce mono- and polyclonal antibodies, and validate their ability to bind to a specific antibody. In reptiles, the major classes of antibodies are IgM and IgY(IgG-like). This is discussed in detail in Chapter 2. As in mammals and birds, following exposure to an antigen, IgM appears first, with IgY developing several weeks after exposure (Ambrosius, 1976). While the time sequence for this response has been determined for certain antigens used in assessing the immune response of reptiles, little information is available for the time sequence following pathogen exposure. This chapter represents an update of a chapter previously published on this topic (Jacobson and Origgi, 2002). In this chapter we will review: (1) types of serologic assays to determine the exposure of reptiles to specific pathogens; (2) review concepts of titer, titrations, quantitations and associated pitfalls; (3) discuss concepts of sensitivity, specificity, positive predictive values, and negative predictive values; (4) collection and handling of blood samples; (5) assays available for determining exposure of reptiles to specific pathogens; and (6) factors affecting the immune response of reptiles.

8.2 Types of Serological Assays Of the various serological tests available for assessing the exposure of animals to pathogens, only a few have been developed for use in determining exposure of reptiles to specific pathogens. Serum neutralization, hemagglutination inhibition, and ELISA are the most widely used semiquantitative serologic assays that have been applied in studying reptile disease. Another test, western blot analysis, can be used to demonstrate the presence of antibodies against specific proteins of pathogens, which can be separated from each other by electrophoresis. Ideally, when a new test is validated, the findings should be compared to those of an independent test that is considered the gold standard. Transmission studies with the pathogen being investigated can also serve in validating the serological test by demonstrating seroconversion

following challenge. Once validated, positive and negative controls should be used every time new samples are tested. Additionally, when new batches of reagents need to be used, it is necessary to run a sample of the new and old batches in parallel to assess if they are comparable. Here we briefly review the mechanics of these assays. More detailed information on general principles of serology can be found elsewhere (Murphy et al., 1999; Turgeon 2003).

8.2.1 Serum Neutralization Test The serum neutralization test (SNT) is generally considered the gold standard when comparing different serologic tests used in determining the exposure of an animal to a virus (Murphy et al., 1999). It is based on the property of a specific antibody subtype (serum-neutralizing antibody) neutralizing a virus. A neutralized viral particle is totally incapable of interacting productively with the cellular surface receptors that normally would mediate its fusion with the membrane of the target cells. Serum neutralizing antibodies bind to the viral ligands (mostly glycoproteins) preventing their binding to the cellular receptor. According to this principle, serum of an animal exposed to a specific virus is likely to neutralize a higher amount of viral particles than the serum of naive animals. The SNT measures the neutralization strength of serum against a specific virus. Briefly, serial dilutions of an inactivated serum sample (heated at 56°C for 30 minutes) are mixed with a standardized amount of virus. The mixture is then left for 1 to 2 hours at room temperature or in an incubator at an optimum temperature. The incubation temperature will vary according to the specific properties of the virus and of the host. During incubation, serum-neutralizing antibodies, if present, will bind to the antigens exposed on the surface virions. The virus–serum mixture is then tested for the residual infectivity on a biological substrate, which could be cells in culture, embryonated eggs, or experimental animals. The final titer of the serum-neutralizing antibodies is determined by end point titration, and it corresponds to the highest serum dilution at which no residual infectivity is detected (according to the selected detection system). Serum neutralization offers a strong confirmation of viral exposure and requires simple instrumentation. The disadvantages of this test are the laborious procedure itself, that is, the length of time (up to 10 to 14 days) required to obtain a final result, the need for using live virus in the assay, and the necessity of the production of a detectable amount of serum-neutralizing antibodies by the host. It is important that the operator be aware of nonspecific neutralization activity, which might be present in some serum samples. If necessary, appropriate measures need to be taken to avoid misleading results influenced by these nonspecific factors.

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8.2.2 Hemagglutination and Hemagglutination Inhibition The hemagglutination and the hemagglutination inhibition tests are based on the same principle. Surface antigens of certain viruses (and other pathogens) can interact with the surface receptors of red blood cells of certain animals, resulting in the formation of bridges between the viral particle and red blood cells. Theoretically, a viral particle could agglutinate two red blood cells. When a large number of viral particles are mixed with a proportional number of red blood cells, the hemagglutinating properties of the virus causes a web of viral particles and red blood cells interacting with each other. Visually, hemagglutination is seen as a diffuse red mat covering the entire bottom of the microtiter well used in the test. When the reaction does not occur, the red blood cells tend to concentrate at the center of the well as a button. The hemagglutination test can be carried out using a sample suspected of containing the virus and red blood cells obtained from an appropriate host. In the hemagglutination inhibition test, the presence of antihemagglutinating antibodies directed against the target virus is assayed. In this case, serial dilutions of inactivated (heated at 56°C for 30 minutes) serum from the suspected infected animal is mixed and incubated with a standardized amount of virus. During the incubation, the antihemagglutinating antibodies are expected to interact with the virus and to block the ligands exposed on the surface of the virus itself. A standardized amount of red blood cells are then added. The hemagglutinating activity of the virus in the assay is expected to be reduced or totally eliminated by the presence of a specific (antivirus) antibody when the serum sample comes from an infected animal that has the circulating antibody. When the serum sample comes from an animal that was not exposed to the pathogen, the hemagglutinating activity of the virus should remain unchanged. The test is considered positive (confirming the exposure to the suspected virus) when the red blood cells are not agglutinated (button forms at the center of the well) in one or more wells containing the serially diluted sample from the suspect animal. The test is considered negative for the presence of antihemagglutinating antibodies when hemagglutination is detectable in all the wells containing the serially diluted sample. The titer of the sample is expressed as the highest dilution at which hemagglutination is still inhibited.

8.2.3 Enzyme-Linked Immunosorbent Assay (ELISA)

tally stored and statistically analyzed. The major start-up costs for performing this test include an ELISA reader and plate washer. These two pieces of equipment can be purchased for less than $10,000.

8.2.3.1 Direct ELISA  In the simplest format, the direct ELISA, the antigen is generally adsorbed on the surface of the wells of a microtiter plate and the antibody specific for the antigen is conjugated with an enzyme (generally alkaline phosphatase [AP] or horseradish peroxidase [HRP]). Once the antibody is bound to the antigen, the chromogen substrate is added (generally p-nitrophenyl phosphate disodium [PPNP] or diaminobenzidine [DAB] for AP and HRP, respectively) and the colorimetric reaction develops.

8.2.3.2 Indirect ELISA  The indirect format is most commonly used to determine the presence of a specific antibody. In the indirect ELISA, the primary antibody that binds to the antigen becomes the target of the secondary antibody conjugated to the enzyme. The secondary antibody can be either a polyclonal antibody (PAB) or monoclonal antibody (MAB) that recognizes and binds to the tested animal’s immunoglobulin (see Chapter 7). PABs are produced by many B cell clones in an immunized animal and thus react with one or more epitopes in the same or closely related antigens. They are produced by multiple immunizations of a suitable animal with a specific antigen. Rabbits are commonly used since they can produce an adequate quantity of antibody that will last (depending upon the amount used) for several years. The problem with PABs is that those derived from a single animal will eventually be consumed in the testing and a new rabbit will eventually be needed for production of more antibodies. MABs represent a homogenous population of antibodies that are derived from a single clone and, using hybridoma technology, are produced by fusing B-lymphocytes with an immortalized cell line. MABs typically react with one epitope on the antigen. They may not necessarily react with closely or distantly related molecules and thus may not have as wide an application as PABs. However, if the clones are stored properly, they will produce a specific MAB that has the potential to last forever. For reptiles, no secondary antibodies are commercially available and are generally produced by individual investigators. PABs and MABs have been produced at the University of Florida against immunoglobulins of the following species of reptiles: green turtle (Chelonia myda), Chinese box turtle (Cuora galbinifrons), desert tortoise (Gopherus agassizii), Mediterranean tortoises (Testudo graeca and T. hermanni), American alligator (Alligator mississippiensis), and boa constrictor (Boa constrictor). No PABs or MABs, have been produced against a lizard immunoglobulin. These reagents, especially the PABs, are cross-reactive against antibodies of multiple species of reptiles.

Among the various serodiagnostic tests available for the detection of antibodies to specific pathogens, the ELISA is one of the most popular and widely used. It is an extremely versatile assay because it can be formatted in several different ways. It is a relatively simple test to run, is sensitive, quantifiable, can be automated, and has a quick turnaround time. A 8.2.3.3 Competitive ELISA  The competitive ELISA is a parcolor change is converted to an optical density value using a ticular format that can be used to detect the presence of an spectrophotometer (ELISA reader) and the values can be digi-

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antibody or antigen. An antigen or antibody can be quantified by its ability to interfere with a pretitrated system. In the direct format, a known antigen or antibody is adsorbed to microtiter wells. If antigen is coated to the wells, then a mixture of a known amount of enzyme-labeled antibody that recognizes the coated antigen is added, along with the unknown plasma sample obtained from the suspect exposed animal. According to the level of the colorimetric reaction obtained with the test, it is possible to estimate the amount of the specific antibody present in the assayed sample. If the color is the same as in the pretitrated format, then all known labeled antibodies are binding to the antigen and the suspect sample does not contain the antibody that recognizes the antigen. If plasma from the suspect animal is added and binds to the coated antigen, then there will be a proportional reduction in the color change. In a 100% competition, no color development takes place.

8.2.3.4 Capture ELISA  In the capture (“sandwich”) antigen ELISA, the main goal is to detect the presence of a specific antigen instead of the antibodies developed to the antigen itself. This ELISA format requires the adsorption of the primary antibody on the microtiter plate wells. The sample to be tested is then added and the specific antigen, if present, is supposed to be captured by the antibodies bound to the plate. The secondary antibody, directed against the same antigen, conjugated with the enzyme, is then added. The chromogen substrate is finally added and the development of the colorimetric reaction will show the presence, and to a certain extent the amount, of antigen present in the sample tested. The capture ELISA can also be quantitative when a standard curve is obtained using predetermined amounts of antigen. In this ELISA format it would be advisable to use two different antibodies directed against two distinct epitopes of the same antigen in order to avoid ambiguous results due to the limited availability of binding sites.

8.2.4 Immunofluorescence Test It is known that some substances observed under UV (ultraviolet) lights can emit fluorescent light. These substances are called fluorochromes. The immunofluorescence test uses this simple principle for the detection of antigens of various pathogens in tissue sections or in cell culture. A monoclonal or polyclonal antibody directed against the specific antigen is conjugated with the fluorochrome. Immunofluorescence might be direct or indirect depending upon which antibody used in the reaction is labeled with the fluorochrome. When the antivirus antibody is labeled with the fluorochrome, the technique is considered direct. This test can also be used for determining the presence of an antibody in an exposed animal. The antibody in the exposed animal will bind to the specific antigen and a labeled secondary antibody is used to bind to the primary antibody. This format is similar to that used in the indirect ELISA. The fluorochromes are light sensi-

tive and will rapidly quench. A photographic or digital image archive is needed for long-term documentation of the results. Fluorochromes that produce different colors are commercially available and can be used to identify different antigens in the same tissue section. Special counterstaining enhances the antigen detection.

8.2.5 Immunoperoxidase The immunoperoxidase technique uses the same approach as the immunofluorescence test. The major difference consists of the use of an enzymatic labeling of the antibody and of a chromogen substrate in place of a fluorochrome to reveal the presence of the antigen (generally a specific protein). The most common enzymatic substrates (same as with ELISA) are alkaline phosphatase (AP) and horseradish peroxidase (HRP). Each requires different chromogen substrates. The immunoperoxidase technique can be used in a direct format to identify the presence of a specific antigen when polyclonal or monoclonal antibodies that specifically recognize the antigen are available. Immunoperoxidase also can be adapted for use as a serologic assay for determining the presence of a specific antibody that develops in response to a specific antigen. In this indirect format, a polyclonal or monoclonal antibody that recognizes immunoglobulin of the exposed animal is needed. First, plasma from the exposed animal is applied to the tissue section or cell culture containing the antigen. Next, a secondary labeled specific antireptile immunoglobulin that is known to bind to the antibody of the target species is added. Once the chromogen substrate is added (diaminobenzidine [DAB] or naphthol AS-MX phosphate for HRP and AP, respectively), it reacts with the enzyme, producing a colorimetric reaction. The immunoperoxidase technique offers a higher stability and more versatility than the immunofluorescence. In contrast with immunofluorescence, no special optical or light instruments are necessary to detect the immunoperoxidase reaction. The disadvantage compared to immunofluorescence is lower sensitivity. The counterstaining is critical and plays an important role in enhancing the quality of the results.

8.2.6 Western Blot Assay The western blot assay is used to determine the presence of antibodies against proteins of a pathogen. For additional details see Chapter 7. It is very similar to the immunoperoxidase assay with respect to the basic principles, but its format is radically different. This technique consists of three stages. In the first stage, the samples (generally a mixture of unpurified or purified proteins) are separated electrophoretically on an agarose gel. In the second stage the proteins are electrophoretically transferred to a nitrocellulose membrane. The third and last stage of the assay consists of the development stage, when a specific antibody directed against the target protein(s) is used to reveal its presence (or absence) on the membrane. As in the immunoperoxidase assay, the antibody

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is labeled with AP or HRP. The complementary chromogen substrate is then added (see above and Chapter 7). When the test is positive, a chromatic reaction detects the target protein as a visible band. For confirmation, molecular weight markers are run on the gel and used to determine the molecular weight of the recognized protein itself. This technique, theoretically very simple, is prone to artifact and technical problems. Extensive fine tuning of each of the different steps of the technique is often required. The presence of good quality protein in the sample and knowledge of basic protein chemistry is extremely important.

8.3 Titer, Titrations, Quantitations, and Pitfalls The primary goal of all the tests described above is to detect antibodies to a pathogen, and when possible to quantify them. The amount of antibodies detected is generally expressed as a titer. It is very important for the clinician to understand that the titer is not a quantitative measure of the antibodies present in the tested samples, but at most represents an ordinal measure of a specific antibody (Tyler and Cullor, 1989). What we measure is the intensity of a colorimetric reaction (ELISA and immunoperoxidase), the efficacy of neutralizing a virus (serum neutralization) or one of its biological properties (hemagglutination or hemagglutination inhibition). This result is not only dependent on and influenced by the amount of specific antibody present in the sample tested, but is also influenced by several other factors, such as the antibody affinity for the antigen, the integrity of the antigens used in the binding of antibody, the availability of the antibody present in the samples, and the proper preservation of the samples. A serum neutralizing titer of 1/64 does not necessarily mean that the sample contains more antibody than a sample with a titer of 1/8. While the amount of antibody could be very similar in absolute terms, a higher affinity of the antibodies for the antigen in one sample could result in quite different titers. A plasma sample that has not been properly collected, preserved, or shipped will result in erroneous antibody titers. The preparation of the antigen itself needs to be standardized and constantly upgraded. The perfect test does not exist and the ideal conditions are not always available in the real world. Each serodiagnostic test provides somewhat different information, and the test selected will be based upon the availability and properties of the antigen used in the test and the availability of specific reagents needed in the assay. The authors strongly suggest contacting laboratories and personnel experienced in running or developing the test of choice. Positive and negative controls are essential and the best plasma samples used as positive controls are those obtained from animals specifically immunized with the antigen used in the assay.

8.4 Establishing Test Cutoffs Many serologic tests by their nature are quantitative (see above), having results that can exist at any point on a scale (Crofts et al., 1988). Thus, cutoffs are needed to make a distinction between positive (those above the cutoff; exposed or infected) individuals and those that are negative (below the cutoff; uninfected or unexposed). When using cutoffs, quantitative test results become qualitative because they are changed from a numerical score to positive or negative. Establishing these cutoffs can be done in several different ways. Ideally, positive samples used in establishing cutoffs should come from animals in which the particular disease is known to occur, and negative samples from animals known to be free of disease. The best gold standard for categorizing an animal as infected is to isolate the pathogen from the animal or to use light or electron microscopy, which will demonstrate highly specific lesions characteristic of the disease or demonstrating the agent. However, no test is 100% sensitive or specific (see below). The failure to culture a pathogen from an animal does not necessarily mean the animal is negative because some pathogens are either difficult or impossible to culture. The clinician must always realize that isolating an organism from an animal with a certain disease may not by itself indicate that it is the cause of the disease. The isolated or identified organism may represent a secondary or opportunistic invader or an endogenous agent of no pathogenic potential, as is the case for many endogenous retroviruses. Several diagnostic tests should be used to confirm the presence or absence of a pathogen in an animal. Using the gold standard criteria given above, reference sera from animals (populations) known to be disease (pathogen) free and from those known to be infected with a specific pathogen are used to establish the cutoffs for a serologic test. In viral serology, SNT is considered the gold standard for detection and quantification of virus-specific antibodies (Murphy et al., 1999) and serum samples classified as positive and negative for exposure to a specific virus using SNT can be used for validating other tests. When the individual data for the positive and negative samples are plotted on a graph with the frequency on the y-axis and the quantitative results of the assay on the x-axis, a separation can be seen between values for negative animals and those for positive animals. In most situations there will be an overlap between these populations, and within this overlap the cutoff is often established. This means that when an animal’s serologic result falls within this overlap, there is a greater probability that an animal classified as positive or negative may actually be falsely classified. Transmission studies are very helpful in establishing cutoffs because prechallenged serum samples can serve as negative controls compared to postchallenged samples showing seroconversion. However, by their very nature, transmission studies are costly and large numbers of animals are seldom

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used in such studies. Because of this they have certain statistical limitations. When working with quantifiable serologic assays such as ELISA, a statistical method for determining the cutoff is to use the mean value plus either 2X (95% confidence interval) or 3X (99% confidence interval) the standard deviation (SD) of the group. In arboviral serology, the positive/negative (P/N) value of each sample is calculated by dividing the mean optical density (OD) of each duplicate pair of positive antigencontaining wells by the OD reading of the wells containing negative (uninfected) cell lysate (Beaty et al., 1995; Martin et al., 2000). Samples with a P/N value > 2 are considered positive for an antiarboviral antibody and samples with a P/N value < 2 are considered negative for an antiarboviral antibody. This has been used to classify alligators as having the antibody against West Nile virus and those that are antibody free (Jacobson et al., 2005b).

8.5 Sensitivity, Specificity, Positive Predictive Values, and Negative Predictive Values All diagnostic tests are expected to give an answer to the specific question: Has the subject been exposed to the pathogen? Besides the positive or negative answer that can be obtained from the test, it is crucial to know to what extent we can trust the test itself. An unreliable assay is likely to lead a clinician in the wrong direction on the diagnostic pathway. In the best of all possible worlds, a serological test should have a sensitivity and a specificity equal to 100%. The sensitivity is defined as the ability of the test to detect as positive those samples that have been collected from individuals actually exposed to a specific pathogen. The specificity is defined as the ability of the test to detect as negative those samples collected from individuals that have never been exposed to the suspected pathogen. It is very uncommon for a serological test to be characterized by a sensitivity and specificity of 100%. The two parameters are interdependent to some extent. In fact, if we are interested in detecting the presence of all the possible exposed animals to a pathogen, it is likely that our cutoff value for the positive samples will be brought down to the lowest value. In a similar scenario we are likely to obtain a very high sensitivity value, but at the same time we are running the risk of considering as positive several individuals that actually are not (low specificity). In contrast, if we would like to identify only the truly positive animals, we could choose the upper end of the cutoff value. As a consequence we are likely to miss several positive samples that would test as negative. In this last scenario, the specificity of the test would be very high, but the sensitivity would be low. Choosing the cutoff value for the positive sample becomes critical for the reliability of the test. When possible, the serological test in development needs to be compared

with a gold standard test whose sensitivity and specificity parameters are already established. Critical in test validation is the availability of experimentally immunized and nonimmunized animals to serve as known positive and negative controls. Once the cutoff is determined, the operator’s judgment will be used to choose the appropriate upper or lower end of the cutoff to apply to the test. For example, when a population of animals, supposedly disease free, is to be surveyed, it would be advisable to use a low cutoff so that all the positive animals are likely to be detected. When the mean value of the negative control plus 2XSD is used as the cutoff, the sensitivity is greater and the specificity less than when 3XSD is used. The area of overlap between negative animals and positive animals may be interpreted in some laboratories as a suspect. Two additional epidemiological parameters that are used to evaluate the reliability of a diagnostic test are the positive predictive value (PPV) and the negative predictive value (NPV). The PPV is the probability that the subject has the disease given a positive test result, while the NPV is the probability that the subject is disease free given a negative test result (Baldock, 1988). The PPV, NPV, sensitivity, and specificity are also interdependent because the PPV tends to be lower when sensitivity is high (high number of false positive samples) and the NPV tends to be lower when specificity is high (high number of false negative samples). Similar to the sensitivity and specificity parameters, the PPV and NPV should be as close as possible to 100% for the test to be considered reliable.

8.6 Collection and Handling of Blood Samples For details on this subject, see Chapters 3 and 13. Remember, the reptile’s size and health status will determine the total amount of blood that can be safely withdrawn. As previously stated, the total blood volume of reptiles varies between species, but as a generalization is approximately 5 to 8% of total body weight (Lillywhite and Smits, 1984; Smits and Kozubowski, 1985). The most important points to remember when submitting plasma or serum samples for serology are:

1. Try to utilize the same blood collection technique at all times. 2. Handle the blood in a consistent fashion. Plasma is preferred and lithium heparin is the recommended anticoagulant. 3. Centrifuge the blood immediately following collection and remove the plasma immediately following centrifugation. 4. Freeze the sample following collection, preferably on dry ice, in liquid nitrogen, or in an ultracold freezer at −70°C. 5. The sample should be transported frozen to the laboratory, preferably on dry ice.

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8.7 Serology for Viral Exposure Serologic tests are available for screening certain reptiles for exposure to several viruses, bacteria, and parasites. For the most part these tests were outcomes of research projects designed to understand their pathogenicity in reptiles. Some are available for screening private and zoologic collections of reptiles. In some cases where the “same” test is being run in different laboratories to determine exposure to the same pathogen, results are not easily compared because techniques and cutoffs vary among different laboratories. Here we review those pathogens for which serologic tests to determine exposure have been developed.

8.7.1 Herpesviruses 8.7.1.1 Tortoise Herpesvirus  The first serological test used to detect exposure to herpesvirus in tortoises was SNT (Marschang et al., 1997; Frost and Schmidt, 1997). In this assay, 25 to 100 microliters of plasma is required, varying with the desired numbers of replicates to be performed. Serum neutralization, in general, is considered the gold standard for serodiagnosis of viral infection (Murphy et al., 1999). Overall, SNT is an excellent diagnostic test that can be performed in every laboratory once the necessary parameters are well established and standardized. It is critical to rely on a standardized virus stock and protocol. It is also extremely important that the same person read the plates each time in order to avoid inconsistent interpretation. A positive result is considered a strong proof of exposure to the virus. It is a relatively easy test to perform, but the major disadvantage is the long time required for completion. According to our experience, a seronegative tortoise should be retested after 5 to 9 weeks before interpreted as negative. It appears that the lowest SNT titer might not be detected for several weeks following exposure. Other problems with this test include an absence of defined sensitivity and specificity, and an absence of a universally recognized reference strain.   The limitations of SNT were an incentive for the development of an indirect ELISA for detecting herpesvirus antibody in tortoises (Origgi et al., 2001). This assay was determined to be statistically as reliable as SNT, easier to perform, and was able to detect seroconversion of experimentally infected tortoises 2 to 5 weeks earlier than SNT. Consistent with SNT, the indirect ELISA could not distinguish between different herpesvirus isolates of the same serotype (Marschang et al., 2001; Origgi et al., 2001). The sensitivity and the specificity for this test were 97 and 98%, respectively. The positive and negative predictive values were 92 and 99%, respectively, demonstrating the reliability of this assay. A negative feature of the current assay is that whole virus is needed as the antigen in this test. This makes the procedure somewhat costly and

laborious because large amounts of virus must be harvested and purified. Recombinant viral proteins may eventually be used as surrogates for the whole viral antigen. This would reduce the cost of the test and allow better standardization of the antigen. This ELISA has been validated for Greek and Hermann’s tortoises and is being standardized for other species of tortoises. Along with SNT and ELISA, direct (DIP) and indirect (IIP) immunoperoxidase-based tests were developed at the University of Florida (Origgi et al., 2003). The IIP test allows detection of the antibody to tortoise herpesvirus, while the DIP can detect the presence of the herpesvirus antigen in paraffin-embedded, formalin-fixed tissue. The DIP and IIP serve as complementary tests for SNT and ELISA, and can serve as another level of validation for exposure and infection with herpesvirus.

8.7.1.2 Marine Turtle Herpesviruses  Immunoglobulin was purified from green turtles, and mouse monoclonal antibodies were produced against these antibodies for use in an indirect ELISA test (Herbst and Klein, 1995). These reagents were also adapted for use in an indirect immunoperoxidase-based serologic test to determine the presence of the antiherpesvirus antibody in the plasma of green turtles with fibropapilloma (Herbst et al., 1998). In this test, sections of tumors containing herpesvirus intranuclear inclusions were used as the source of the antigen in the assay.   An indirect ELISA was developed to determine the exposure of green turtles having a respiratory disease (lung-eyetrachea [LET] disease) to an associated herpesvirus (LETV) (Coberley et al, 2002a). In one study, plasma samples from wild juvenile green turtles from three sites over of 3-year period (1997 through 1999) in Florida were tested by ELISA to determine the presence of the antibody against this virus and changes in the antibody over time (Coberley et al., 2002b). While seropositive turtles were found at all 3 study sites, no changes were seen over the 3-year sampling period. In a concurrent study, the anti-LETV antibody status of turtles with fibropapillomas (FP) was compared with those free of FP. No correlation was seen between the severity of FP and the presence or absence of the anti-LETV antibody. When assays were performed, plasma from green turtles before and after immunization with LETV served as negative and positive controls. Western blotting was used as an additional confirmation of the presence of antibody in those turtles found seropositive by ELISA. The antibody was also demonstrated in the plasma of nesting green and loggerhead (Caretta caretta) sea turtles in Florida. The original isolate of LETV was from ill green turtles at Cayman Turtle Farm, Grand Cayman, BWI (Jacobson et al., 1986) and the survey in Florida was the first study to show exposure of wild green and loggerhead sea turtles to LETV.

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8.7.2 Iridovirus Infection of Chelonians Iridoviruses are emerging as important pathogens of chelonians. In our laboratory at the University of Florida, iridovirus (probably Ranavirus) was isolated from wild box turtles (Terrapene carolina), wild gopher tortoises (Gopherus polyphemus), and a Burmese star tortoise (Geochelone platynota). An indirect ELISA was validated to determine exposure of chelonians to this virus (April Johnson, personal communication). In this ELISA, an anti–desert tortoise monoclonal antibody (which cross-reacts with immunoglobulins of different chelonians) was used as the secondary antibody in the assay. Serological surveys will be performed on populations of wild box turtles and gopher tortoises in the southeastern United States.

8.7.3 Paramyxovirus A hemagglutination inhibition (HI) assay has been developed to determine the presence of the antibody against paramyxoviruses isolated from snakes (Jacobson et al., 1981; Jacobson et al., 1992; Richter et al., 1996). These isolates were collectively termed ophidian paramyxoviruses. More recently, paramyxoviruses have also been isolated from lizards (see Chapter 9). The HI assay has been most useful because of its relative simplicity and rapid turnaround time. Briefly, serum samples collected by heart puncture or tail venipuncture are diluted 1:10 in sterile physiological saline at 56oC for 30 minutes to inactivate the complement, then absorbed with washed and pelleted chicken erythrocytes to remove nonspecific agglutinins (12 hrs. at 5oC). Using microtiter methodology, serial doubling serum dilutions are made (1:10, 1:20, 1:40, 1:80, etc.) with 0.05 ml volumes of phosphate-buffered physiological saline containing 0.1% bovine serum albumin. The latter minimizes autoagglutination of the erythrocyte suspension used later to indicate whether the active virus is present or not. To each serum dilution is added an ophidian paramyxovirus suspension diluted to contain 8 hemagglutination units/0.05 ml. This has previously been determined by titration, taking advantage of the fact that the virus causes chicken erythrocytes to bind together (hemagglutinate). After allowing the virus–serum mixtures to interact for one (1) hour at room temperature, 0.1 ml of a 0.3% suspension of washed chicken erythrocytes is added to each well of the microtiter plate. The plates are placed in a refrigerator for 2 to 3 hours to permit settling of the erythrocytes. If antibodies are present in a particular serum dilution, they will bind to viral particles, which will prevent them from hemagglutinating the erythrocytes (hemagglutination inhibition). Where the antibody is present, the red cells settle into a “button” at the bottom of the microtiter well rather than forming a “mat” due to the hemagglutinating property of the viruses. The serum antibody “titer” is read as the reciprocal of the highest serum dilution that still causes hemagglutination inhibition. An HI titer < 20 is considered negative, 40 to 80 suspect, and > 80 positive, indicating expo-

sure to the virus. Snakes that survive paramyxovirus infections may have HI antibody titers exceeding 10,240. A single sample is only indicative of the exposure status at the time the sample was collected. In order to demonstrate an active infection, samples should be collected at 2- to 4week intervals. A fourfold increase in titer is indicative of a recent infection. In a zoological collection of snakes experiencing an outbreak of paramyxovirus, snakes sampled at 5 months following the death of the first snake showed elevated antibody titers; three months later, many snakes sampled had antibody titers < 1:80 (Jacobson et al., 1992). While the HI antibody does not remain elevated for prolonged periods in most exposed snakes, in others it may remain elevated for long periods of time. There is only a single report of a survey for anti-paramyxovirus antibodies in wild snakes. Of 21 eastern massasaugas (Sistrurus catenatus catenatus) collected in Illinois, there was sufficient blood to test 20 for antibodies against paramyxovirus (Allender et al., 2006). Using an HI test, all 20 snakes were found to be positive. However, these may represent false positives. These findings need to be validated using another assay such as western blot analysis or serum neutralization. In lizards, paramyxovirus infection was reported in caiman lizards (Draecena guianensis) in three different collections (Jacobson et al., 2001). Seven months following the outbreak in one collection, blood samples were collected from 17 surviving lizards to determine the presence of the antibody against an isolate from a dead caiman lizard. Of the 17 lizards, 7 had titers of < 1:20 and 10 had titers of > 1:20 and < 1:80. None were considered seropositive. In another study, blood samples were obtained from 35 free-ranging healthy spiny-tailed iguanas (Ctenosaura bakeri, C. similis) and 14 green iguanas (Iguana iguana rhinolopha) collected on the Honduran Islands of Utila and Roatan (Gravendyck et al., 1998). With SNT, 20 (41%) had antibodies against a reptilian paramyxovirus and 3 (9%) were positive using HI. Using HI, four of 23 samples obtained from wild collected lizards (Xenosaurus and Abronia) from Mexico had antibodies against a paramyxovirus isolated from a lizard in this study (Marschang et al., 2002). Few studies have been conducted looking at cross-reactivity between the various isolates of paramyxovirus. When two isolates of paramyxovirus having slightly different electrophoretic protein mobilities were used as coating antigens in an HI test, differences in titers for the same plasma samples were seen (Elliott Jacobson, unpublished findings). Still, most snakes categorized as positive were positive using both isolates, despite titer differences. In another study, a comparison of 60 snake samples showed considerable variation in reactivity when two different snake paramyxoviruses were used in the assay (Kania et al., 2000). In order to have an assay that is more sensitive and specific than HI, an indirect ELISA was developed to measure antiparamyxovirus antibody in snakes (Kania et al., 2000).

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Using ammonium sulfate precipitation and subsequent elution on DEAE Sephadex, the authors attempted to recover immunoglobulin from boa constrictor serum. A protein fraction corresponding to the gamma fraction of snake serum was eventually identified and injected into rabbits for production of PABs. When assessed against the plasmas of multiple species, the PABs only had strong reactivity against plasmas from boid snakes. While a strong correlation was found between samples positive by ELISA and HI, this study needed further validation to prove that boa constrictor immunoglobulin was actually present in the eluted protein fraction used for production of PABs. PABs were possibly produced against a protein in boa constrictor plasma other than immunoglobulin. Interpreting serologic findings and how they are to be used in managing a collection is not straightforward. In mammals, persistent paramyxovirus infections are rare. Measles can persist in some human patients in the brain. In a persistent infection, the virus continues to replicate, but at a much lower level than during the initial phase of infection and replication. Regarding viral shedding, we need to be able to distinguish between slow or residual infection and persistence. In slow or residual infection the virus is no longer replicating, but continues to be eliminated by the animal from replication sites. Thus an animal that has survived an active infection may continue to shed virus for some time following the end of the infection. For reptile paramyxoviruses, we do not know how long virus may continue to be shed following an active infection. Generally when neutralizing antibodies are being produced, viral replication eventually ceases. While we are measuring HI antibodies and not neutralizing antibodies, more than likely neutralizing antibodies are also being produced. This has been demonstrated in serological surveys conducted in Germany. While we do not know if snakes can have a persistent infection, there is no evidence to support this. If a persistent infection is uncommon in snakes or does not occur, then those snakes with high antibody titers that survive infection may be immune if exposed to the same virus. More research is needed to make recommendations on the disposition of seropositive snakes.

8.7.4 Reovirus Blood samples were obtained from 35 free-ranging healthy spiny-tailed iguanas (Ctenosaura bakeri, C. similis) and 14 green iguanas (Iguana iguana rhinolopha) collected on the Honduran Islands of Utila and Roatan (Gravendyck et al., 1998). Of 49 samples, 23 (47%) tested positive for exposure to reoviruses using SNT. Using SNT, 3 wild collected lizards (Abronia grandis) from Mexico had antibodies against a reptilian (green iguana) reovirus (Marschang et al., 2002).

8.7.5 Arboviruses Several arthropod-transmitted viruses are known to infect reptiles. Regarding members of the family Togaviridae, there is

evidence of the antibody to eastern equine encephalomyelitis virus (EEEV) in the blood of wild American alligators (Alligator mississippiensis) (Karstad, 1961), the presence of the antibody to western equine encephalomyelitis virus in the blood of Texas tortoises (Gopherus berlandieri) (Bowen, 1977) and several species of snakes (Thomas and Eklund, 1962; Gebhardt et al., 1973), and the antibody to EEEV and Venezuelan equine encephalomyelitis virus in the blood of tegu lizards (Tupinambis nigropunctatus) (Walder et al., 1984). Evidence of infection of reptiles with members of the family Flaviviridae included the isolation of Japanese encephalitis virus from Chinese rat snakes (Elaphe rufodorsata) (Lee et al., 1972) and the presence of antibodies to St. Louis encephalitis virus in a rattlesnake in California (Reeves, 1990). Mortality associated with another flavivirus, West Nile virus (WNV), was identified in farmed American alligators (Alligator mississippiensis) in Georgia (Miller et al., 2003) and Florida (Jacobson et al., 2005a). Antibodies were detected in Nile crocodiles (Crocodylus niloticus) in a commercial farming operation in Israel (Steinman et al., 2003) and an ELISA was developed to measure anti-WNV antibody in alligators (Jacobson et al., 2005b). These are discussed below.

8.7.5.1 West Nile Virus Infection of Crocodilians  In October 2002, WNV was identified in farmed American alligators (Alligator mississippiensis) in Florida showing clinical signs and having microscopic lesions indicative of central nervous system disease (Jacobson et al., 2005a). To perform seroepidemiologic studies, an indirect ELISA was developed to determine exposure of captive and wild alligators to WNV (Jacobson et al., 2005b). To validate the test, a group of WNV seropositive and seronegative alligators were identified at the affected farm using HI and the plaque reduction neutralization test (PRNT). The indirect ELISA utilized a rabbit antialligator immunoglobulin polyclonal antibody as the secondary antibody, and inactivated WNV-infected Vero cell lysate was used as the coating antigen. For all samples (n = 58), the results of the ELISA were consistent with the HI and PRNT findings. Plasma was collected from 669 wild alligators from 21 sites across Florida in April and October 2003. Four samples collected in April and six in October were found positive using HI, PRNT, and the indirect ELISA. This indicated that wild alligators in Florida have been exposed to WNV. These findings can be used as a baseline for future surveys. While no deaths were reported, antibodies to WNV were found in Nile crocodiles (Crocodylus niloticus) in a commercial farming operation in Israel (Steinman et al., 2003).

8.7.6 Inclusion Body Disease of Boid Snakes While the causative agent of inclusion body disease (IBD) of boid snakes has not been conclusively identified or isolated, a serologic assay needs to be in place once the causative agent is identified. This need resulted in the development

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and validation of an indirect ELISA for use in boa constrictors, Boa constrictor (Lock et al., 2003). Two boa constrictors were immunized with 2,4 dinitrophenylated bovine serum albumin (DNP-BSA). Pre-immune blood samples were collected and serial samples were subsequently collected. Using affinity chromatography, the anti-DNP antibody was purified and used to produce antiboa rabbit polyclonal and mouse monoclonal antibodies. An indirect ELISA and western blots were used to detect the anti-DNP antibody in immunized boa constrictors. In a subsequent study, 58 plasma samples from wild boa constrictors in Brazil were evaluated for exposure to a retrovirus isolate from captive boa constrictors with IBD (Lock and Jacobson, 2005). The values from this study can serve as a baseline reference range when assaying plasma samples from boa constrictors with IBD. A 68-KDa protein has been identified in the inclusions of snakes with IBD (Wozniak et al., 2000). This protein is currently being isolated and sequenced. If snakes having IBD produce antibodies against this protein, and snakes free of inclusions do not, then it may be possible to develop a serologic test that is an indicator of the presence of the 68-KDa protein. For further details, see Chapter 9.

8.8 Serology for Bacterial Exposure 8.8.1 Mycoplasmosis 8.8.1.1 Chelonian Mycoplasmosis  An indirect ELISA was developed to detect the presence of anti–M. agassizii antibodies in tortoise plasma (Schumacher et al., 1993). In the indirect ELISA assay, M. agassizii whole-cell lysate antigens are coated onto the bottom of a sample well. Tortoise plasma is added to the well, and in the presence of anti–M. agassizii antibodies in the plasma they will specifically bind to certain M. agassizii antigens. After washing, the secondary antibody (antitortoise mouse monoclonal antibody) is added, which in the presence of the tortoise antibody, will specifically bind to the tortoise antibodies, ultimately inducing a chromogenic reaction. In the absence of anti–M. agassizii antibodies in the plasma sample, no color develops. The intensity of the color development (absorbance or OD) is measured photometrically and compared to that of positive control plasma from infected tortoises and negative control plasma from unexposed tortoises.  Results of the ELISA are analyzed quantitatively but interpreted categorically (negative, suspect, or positive). The detection limit is the background OD value (i.e., wells containing all of the reagents except tortoise plasma). In the original assay, the ratio of the sample OD to negative control OD was used to categorize a tortoise. If the ratio was < 2, the sample was considered negative for anti–M. agassizii antibodies. If the ratio of sample OD to negative control OD was > 3, the sample was considered positive for anti–M. agassizii antibodies. If the ratio

of ODs was between 2 and 3, the sample was considered suspect. In an early transmission study (Brown et al. 1994), seroconversion was defined as either a change to an ELISA ratio > 2, or an increase > 0.1 unit compared to a previous sample. The cutoffs were chosen subjectively and conservatively, with the specific objective of minimizing the chance of false negative results. The cutoffs can be revised as new ELISA values of tortoises from populations with a defined prevalence of upper respiratory tract disease (URTD) become available. Currently the Mycoplasma Research Laboratory, Department of Infectious Diseases and Pathology at the University of Florida, uses titers to determine the serologic status of tortoises rather than using OD ratios (Brown et al., 2002). Using titers, 64. Two kinds of monoclonal antibodies have been developed for use as the second antibody in the indirect M. agassizii ELISA. One (HL1163) is a mouse antibody against tortoise immunoglobulin IgM, the class of antibodies that a tortoise produces first in response to infection by M. agassizii. The second (HL673) is a mouse antibody against the light chain of both IgM and IgY tortoise immunoglobulins. The latter antibody cross-reacts with the antibodies from multiple species of chelonians. Antibodies derived from maternal blood and transmitted to hatchling blood through the egg yolk may persist in the hatchlings for up to 1 year (Schumacher et al., 1999). From hatchlings derived from seropositive females, plasma samples should be taken at least 3 months apart to detect a change in antibody titer. More than a twofold increase in antibody titer indicates an active infection. Paired sampling provides more information than single sampling, but increases expense and turnaround time for testing. The advantages of diagnosing infection by ELISA include sensitivity and the ability to detect the stage of infection (early first time vs. chronic) by using different secondary antibodies. A disadvantage of ELISA testing is the consumption of plasma in the test. A positive test result proves past exposure but not current infection. There may be false positives caused by crossreaction of tortoise antibodies to other bacteria with similar antigens. There also may be false negatives due to the presence of other mycoplasma that do not cross-react with M agassizii. During routine screening of desert tortoises with URTD, a second genetically distinct mycoplasma, represented by tortoise isolate H3110 (American Type Culture Collection accession 700618), was cultured from affected tortoises (Brown et al., 1995). This was distinguished from previously described mycoplasmas using serology and comparing 16S rRNA gene sequences, and was named M. testudineum (Brown et al., 2004). Another complicating factor is the well-documented immune periodicity seen in reptiles. There is some evidence of seasonal fluctuation in M. agassizii–specific antibody concentrations. Application of diagnostic tests for mycoplasmal infections of desert and gopher tortoises, with management recommendations, has been published (Brown et al., 2002). For additional information on other aspects of this disease, see Wendland et al. (2006).

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8.8.1.2 Crocodilian Mycoplasmosis  In 1995 an epidemic of pneumonia with fibrinous polyserositis and multifocal arthritis was reported in a captive group of American alligators (Alligator mississippiensis) in Florida (Clippinger et al., 2000). A Mycoplasma sp. was cultured from ill and dead alligators and was subsequently named M. alligatoris (Brown et al., 2001a). Alligator immunoglobulin was purified from alligator plasma and a polyclonal antibody was raised in rabbits against the purified immunoglobulin. Polyclonal antialligator antibodies was purified and conjugated with biotin. This was used as the secondary antibody in an indirect ELISA that was developed for detecting exposure of alligators to M. alligatoris (Brown et al., 2001b). In a challenge study in American alligators, broad-nosed caiman (Caiman latirostris), and Siamese crocodiles (Crocodylus siamensis), the indirect ELISA detected seroconversion in all three species beginning six weeks after inoculation (Brown et al., 2001c). A seroprevalence study in wild alligators in Florida indicated that 5.4% were seropositive for anti–M. alligatoris antibodies (Brown et al., 2005). Seropositive alligators were found in 12 of 20 sites sampled. Mycoplasmosis also is known to occur in Nile crocodiles in Zimbabwe (Mohan et al., 1995). The clinical signs are similar to those seen in alligators with mycoplasmosis. The causative agent is M. crocodyli (Kirchoff et al., 1997). Recently, an indirect ELISA test with high sensitivity and specificity was developed to determine exposure of Nile crocodiles to M. crocodyli (Dawo and Mohan, 2007). This may help in understanding transmission of M. crocodyli among Nile crocodiles at farms in Zimbabwe.

8.8.2 Leptospira Leptospira has been isolated from turtles (Glosser et al., 1974; Hoeden, 1968), lizards (Plesko et al., 1962), and snakes (Ferris et al., 1961; Hyakutake et al., 1980). Garter snakes (Thamnophis sirtalis) inoculated with L. interrogans developed a leptospiremia and serum agglutinins (Abdulla and Karstad, 1962). In the same study, while the turtles Emys orbicularis, Emydoidea blandingi, and Chelydra serpentina also were inoculated, only E. blandingi developed a leptospiremia. Antibodies against Leptospira were detected in caiman (Rossetti et al., 2003) and snakes (Stanchi et al., 1986) in Argentina. The standard serologic test for Leptospira is the microscopic agglutination test.

8.8.3 Coxiella Coxiella burnetii agglutinins were detected in sera of 13 of 53 snakes (water snakes [Natrix natrix], rat snakes [Ptyas korros], cobra [Naja naja], and pythons [Python molurus]) and 2 of 16 tortoises (Kachuga sp.). A capillary agglutination test was used to detect anti-Coxiella antibodies in this study (Yadav and Sethi, 1979).

8.9 Serology for Parasite Exposure 8.9.1 Cryptosporidiosis An indirect ELISA was developed to determine exposure of snakes to Cryptosporidium serpentis (Graczyk and Cranfield, 1997). However, the assay did not detect an antibody response in certain reptiles (such as leopard geckos [Eublepharis macularius]) that are infected with cryptosporidium. This suggested that more than one cryptosporidium exists in reptiles and that the C. serpentis antigen used in the ELISA does not cross-react with all reptile cryptosporidium. In a recent report, several different species of Cryptosporidium were identified in reptiles including two genotypes of C. serpentis (Xiao et al., 2004). For details on cryptosporidium in reptiles see Chapter 12.

8.9.2 Spirorchiidiasis An ELISA was developed using the surface glycocalyx crude antigen of the adult blood trematodes Carettacola hawaiiensis, Haplotrema dorsopora, and Learedius learedi for detecting antibodies in blood of Hawaiian green turtles naturally infected with these parasites (Graczyk et al., 1995). For antigen preparation, all three species of trematodes were pooled. A direct ELISA using antireptilian/amphibian phosphatase-labeled IgG-recognized green turtle immunoglobulin at a dilution of 1/12,800. Next, an indirect ELISA was used with the antireptilian/amphibian phosphatase labeled IgG serving as the secondary antibody. The antibody to blood flukes was detected up to a dilution of 1/3,200. Of 59 samples collected from turtles at five sites, 47 (80%) were seropositive. To look at the relationship of spirorchiid infections and green turtle fibropapilloma, an indirect ELISA was developed using adult antigens of either Laeredius sp. or Haplotrema sp. and biotinylated monoclonal antibody HL857, which is specific for green turtle 7S IgY heavy chain (Herbst et al., 1998). Two wild green turtles, one with Learedius and one infected with Hapalotrema were serologically evaluated. Both turtles with spirorchiid infections had antispirorchiid titers. Because each turtle was infected with only one of these parasites, considerable antigenic cross-reactivity was found between these two species of spirorchiids. The ELISA was sensitive enough to detect exposure of green turtles to spirorchiids even though adult parasites could not be found in seropositive turtles at necropsy. Either there were too few parasites to find at necropsy or there was clearance of infection with persistence of the antibody. It is unknown how long antibodies continue to circulate once the infection has cleared.

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8.10 Factors Affecting the Immune Response in Reptiles Seroconversion of reptiles following experimental immunization or natural immunization can be influenced by a number of factors. The nature of the antigen, environmental conditions, and biology of the specific reptile are all variables that can affect the immune response. Not all foreign proteins result in an immune response. In such cases adjuvants may be used when maximizing the response of experimentally immunized reptiles. The route of infection, such as oral versus parenteral administration, will affect the hosts’ ability to respond immunologically. The timing of immunization is critical because the ability of a reptile to become immunocompetent following birth varies among species (El Deeb and Saad, 1990). Some reptiles have a competent immune system at birth while others need several months for the immune response to mature. Maternal transfer of the antibody needs to be considered when interpreting neonate serology. In the desert tortoise, the maternal antibody is transferred through the egg and can be measured in the neonate (Schumacher et al., 1999). As pointed out previously, environmental temperature affects the body temperature of reptiles and with it, the animal’s immune response to antigens. The immune response appears to respond maximally when the reptile has a body temperature within its thermal optimum zone. Reduced responses occur when a reptile is maintained at temperatures above and below this zone (Cone and Marchalonis, 1972; Tait, 1969). Seasonal cycles of lymphoid follicular involution and recrudescence occur in many reptiles, particularly those living in temperate regions. These cycles may correlate better with mating periods rather than changes in ambient temperature. During these periods, antibody levels may be low and corticosteroids and sex steroids high (Leceta and Zapata, 1986; Saad and El Deeb, 1990; Zapata et al., 1992). Sex-related differences have also been reported with males having lower antibody production during periods of high testosterone levels compared to females. Age, body temperature, season, and hormones (sex) all have to be considered when interpreting serologic findings. For further details on factors affecting the immune system of reptiles, see Chapter 2.

Acknowledgments This chapter represents an update of a chapter previously published on this topic: Jacobson ER, Origgi F. 2002. Use of serology in reptile medicine, in Seminars in Avian and Exotic Pet Medicine, Serodiagnostics, Fudge AM and Raidal S (Eds.), 11:33–45.

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Coberley SS, Herbst LH, Brown DR, Ehrhart LM, Bagley DA, Schaf S, Moretti R, Jacobson ER, and Klein PA. 2002a. Detection of antibodies to a disease-associated Herpesvirus of the green Turtle, Chelonia mydas. J Clin Micro 39:3572–3577. Coberley SS, Herbst LH, Ehrhart LM, Bagley DA, Hirama S, Jacobson ER, and Klein PA. 2002b. Survey of Florida green turtles for exposure to a disease-associated Herpesvirus. Dis Aq Org 47:159–167. Cone RE and Marchalonis JJ. 1972. Cellular and humoral aspects of the influence of environmental temperature on the immune response of poikilothermic vertebrates. J Immunol 108:952–957. Crofts N, Maskill W, and Gust ID. 1988. Evaluation of enzyme-linked immunosorbent assays: a method of data analysis. J Virolog Meth 22:51-59. Dawo F, and Mohan K. 2007. Development and application of an indirect ELISA test for the detection of antibodies to Mycoplasma crocodyli infection in crocodiles (Crocodylus niloticus). Vet Mircobiol 119:283-289. El Deeb SO and Saad AHM. 1990. Ontogenic maturation of the immune system in reptiles. Dev Comp Immun l14:151–159. Ferris DH, Rhoades HE, Hanson LE, Galton M, and Mansfield MR. 1961. Research into the nidality of Leptospira ballum in campestral hosts including the hog-nosed snake (Heterodon platyrhinos). Cornell Vet 51:405–418. Frost JW and Schmidt A. 1997. Serological evidence of susceptibility of various species of tortoises to infection by herpesviruses. Verh Ber Erkrg Zootiere 38:25–27. Gebhardt LP, St. Jeor SC, Stanton GJ, and Strigfellow DA. 1973. Ecology of western encephalitis virus. Proc Soc Exp Biol Med 142:731–733. Glosser JW, Sulzer CR, Ebergart M, and Winkler WG. 1974. Cultural and serological evidence of Leptospira interrogans serotype tarassovi infection in turtles. J Wildl Dis 10:429–435. Graczyk TK and Cranfield MR. 1997. Detection of Cryptosporidiumspecific immunoglobulins in captive snakes by a polyclonal antibody in the indirect ELISA. Vet Res 29:187–195. Graczyk TK, Aguirre AA, and Balazs GH. 1995. Detection by ELISA of circulating anti-blood fluke (Carettacola, Haplotrema, Laeredius) immunoglobulins in Hawaiian green turtles (Chelonia mydas). J Parasitol 8:416–421. Gravendyck M, Ammermann P, Marschang RE, and Kaleta EF. 1998. Paramyxoviral and reoviral infections of iguanas on Honduran islands. J Wildl Dis 34:33–38. Herbst LH and Klein PA. 1995. Monoclonal antibodies for the measurement of class-specific antibody responses in the green turtle, Chelonia mydas. Vet Immuno Immunopath 46:317–335. Herbst LH, Jacobson ER, Moretti R, Brown T, Sundberg JP, and Klein PA. 1995. Experimental transmission of green turtle fibropapillomatosis using cell-free extracts. Dis Aq Org 22:1–12. Herbst LH, Greiner EC, Ehrhart LM, et al. 1998. Serological association between spirorchidiasis, herpesvirus infection, and fibropapillomatosis in green turtles from Florida. J Wildl Dis 34:496 507. Hoeden J van der. 1968. Agglutination of Leptospirae in sera of fresh water turtles. Antonie van Leeuwenhoek 34:458–464. Hyakutake S, Biasi PD, Belluomini HE, and Santa Rosa CA. 1980. Leptospirosis in Brazilian snakes. Int J Zoonoses 7:73–77. Jackson OF. 1981. Clinical aspects of diagnosis and treatment, in Diseases of the Reptilia, Vol 2, Cooper JE and Jackson OF (Eds.), Academic Press, London, 507–534.

Jacobson ER and Origgi F. 2002. Use of serology in reptile medicine. Sem Av Exot Pet Med Serodiagnost, 11:33–45. Jacobson ER, Gaskin JM, Page D, Iverson WO, Johnson JW. 1981. Illness associated with paramyxo-like virus infection in a zoologic collection of snakes. J Amer Vet Med Assoc 179:1227-1230. Jacobson ER, Gaskin JM, Roelke M, Greiner EC, Allen J. 1986. Conjunctivitis, tracheitis, and pneumonia associated with herpesvirus infection in green sea turtles. J Amer Vet Med Assoc 189:1020-1023. Jacobson ER, Gaskin JM, Wells S, Bowler K, and Schumacher J. 1992. Epizootic of ophidian paramyxovirus in a zoological collection: pathological, microbiological, and serological findings. J Zoo Wildl Med 23:318–327. Jacobson ER, Origgi F, Pessier AP, Lamirande EW, Walker I, Whitaker B, Stalis IH, Nordhausen R, Owens JW, Nichols DK, Heard D, and Homer B. 2000. Paramyxo-like virus infection in caiman lizards, Dracaena guianensis. J Vet Diag Investig 13:143–151. Jacobson ER, Ginn PE, Troutman JM, Farina L, Stark,L, Klenk K, Burkhalter KL, and Komar N. 2005a. West Nile Virus infection in farmed American alligators (Alligator mississippiensis) in Florida. J Wildl Dis 41:96–106. Jacobson ER, Johnson AJ, Hernandez JA, Tucker SJ, Dupuis AP II, Stevens R, Carbonneau D, and Stark L. 2005b. Use of an indirect enzyme linked immunosorbent assay for detection of antibodies to West Nile Virus in American alligators (Alligator mississippiensis). J Wildl Dis 41:107–114. Kania SA, Kennedy MA, Nowlin N, Diderrich VR, Ramsay E. 2000. Development of an enzyme linked immunosorbent assay (ELISA) for the diagnosis of ophidian paramyxovirus. Infect Dis Rev 2:213-217. Karstad, L. 1961. Reptiles as possible reservoir hosts for eastern encephalitis virus, in Transactions of the 26th North American Wildlife and National Resources Conference, Wildlife Management Institute, Washington, DC, 185–202. Kirchoff H, Mohan K, Schmidt R, Runge M, Brown DR, Brown MB, Foggin CM, Muvavariwa P, Lehmann H, Flossdorf J. 1997. Mycoplasma crocodyli sp. nov., a new species from crocodiles. Int J Syst Bacteriol 47:742-746. Leceta J and Zapata A. 1986. Seasonal variation in the immune response of the tortoise Mauremys caspica. Immunol 57:483–487. Lee HW, Min BW, and Lim YW. 1972. Isolation and serologic studies of Japanese encephalitis virus from snakes in Korea. J Korean Med Assoc 15:69–74. Lillywhite HB and Smits AN. 1984. Lability of blood volume in snakes and its relationship to activity and hypertension. J Exp Biol 110:267–274. Lock BA and Jacobson ER. 2005. Use of an ELISA to survey exposure of wild caught boa constrictors, Boa constrictor, to retroviruses isolated from boid snakes with inclusion body disease. J Herp Med Surg 15:4–8. Lock BA, Green LG, Jacobson ER, and Klein PA. 2003. Enzymelinked immunosorbent assay for detecting the antibody response in Argentine boa constrictors (Boa constrictor occidentalis). Amer J Vet Res 64:388–395. Marschang RE, Gravendyck M, and Kaleta EF. 1997. Investigation into virus isolation and the treatment of viral stomatitis in T. hermanni and T. graeca. J Vet Med Series B. 44:385–394. Marschang, RE, Frost JW, Gravendyck M, and Kaleta EF. 2001. Comparison of 16 chelonid herpesviruses by virus neutralization tests and restriction endonuclease digestion of viral DNA. J Vet Med B48:393–399.

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Marschang RE, Donahoe S, Manvell R, and Lemos-Espinal J. 2002. Paramyxovirus and reovirus infections in wild-caught Mexican lizards (Xenosaurus and Abronia sp.). J Zoo Wildl Med 33:317–321. Martin DA, Muth DA, Brown T, Johnson, AJ, Karabatsos N, and Roehrig JT. 2000. Standardization of immunoglobulin M capture enzyme-linked immunosorbent assays for routine diagnosis of arboviral infections. J Clin Microbiol 38:1823–1826. Miller DL, Mauel MJ, Baldwin C, Burtle G, Ingram D, Hines II ME, and Frazier KS. 2003. West Nile virus in farmed alligators. Emerg Inf Dis 9:794–799. Mohan K, Foggin CM, Muvavariwa P, Honywill J, Pawandiwa A. 1995. Mycoplasma-associated polyarthritis in farmed crocodiles (Crocodylus niloticus) in Zimbabwe. Onderstepoort J Vet Res 62:45–49. Murphy FA, Gibbs EPJ, Horzinek MC, and Studdert MJ. 1999. Laboratory diagnosis of viral diseases, in Veterinary Virology, 3rd edition, Academic Press, New York, 193–224. Origgi FC, Klein PA, Mathes K, Blahak S, Marschang RE, Tucker SJ, and Jacobson ER. 2001. An enzyme linked immunosorbent assay (ELISA) for detecting herpesvirus exposure in Mediterranean tortoises (Spur-Thighed tortoise [Testudo graeca] and Hermann’s tortoise [Testudo hermanni]). J Clin Microbiol 39:3156–3163. Origgi FC, Klein PA, Tucker SJ, and Jacobson ER. 2003. Application of immunoperoxidase based techniques to detect Herpesvirus infection in tortoises. J Vet Diagn Investig 15:133–140 Plesko L, Janovicova E, and Lac J. 1962. Beitrag zur Bedeutung von Kaltblutern fur die Zirkulation der Leptospiren in der Natur. Zentralbl Bakteriol Parasitenkd Hyg Abt I Orig A 192:482–484. Reeves WC. 1990. Overwintering of arboviruses, in Epidemiology and Control of Mosquito-Borne Arboviruses in California, 1943-1987, Reeves WC, Asman SM, Hardy JL, Milby MM, Reisen WK (Eds.), California Mosquito Control Association, Sacramento, CA, pp. 357-382. R ichter GA, Homer BL, Moyer SA, Williams DS, Scherba G, Tucker SJ, Hall BJ, Pedersen JC, and Jacobson ER. 1996. Characterization of paramyxoviruses isolated from three snakes. Virus Res 43:77–83. Rossetti CA, Uhart M, Romero GN, and Prado W. 2003. Detection of leptospiral antibodies in caimans from the Argentinean Chaco. Vet Rec 153:632–633. Saad AH and El Deeb S. 1990. Immunological changes during pregnancy in the viviparous lizard, Chalcides ocellatus. Vet Immunol Immunopathol 25:279–286. Schumacher IM, Brown MB, Jacobson ER, Collins BR, and Klein PA. 1993. Detection of antibodies to a pathogenic mycoplasma in desert tortoises (Gopherus agassizii) with upper respiratory tract disease. J Clin Microbiol 31:1454–1460.

Schumacher IM, Rostal DC, Yates R, Brown DR, Jacobson ER, and Klein PA. 1999. Persistence of maternal antibodies against Mycoplasma agassizii in desert tortoise hatchlings. Amer J Vet Res 60:826–831. Smits AW and Kozubowski MM. 1985. Partitioning of body fluids and cardiovascular responses to circulatory hypovolemia in the turtle Pseudemys scripta elegans. J Exp Biol 116:237–250. Stanchi NO, Grisolia CS, Martino PE, and Peluso FO. 1986. Presence of antileptospira antibodies in ophidia in Argentina. Rev Argent Microbiol 18:127–130. Steinman A, Banet-Noach C, Tal S, Levi 0, Simanov L, Perk S, Malkinson M, and Shpigel N. 2003. West Nile virus infection in crocodiles. Emerg Infect Dis 9:887–889. Tait NN. 1969. The effect of temperature on the immune response in cold-blooded vertebrates. Physiol Zool 42:29–35. Taylor RW and Jacobson ER. 1981.Hematology and serum chemistry of the gopher tortoise, Gopherus polyphemus. Comparative Biochemistry and Physiology 72A:425–428. Thomas LA and Eklund CM. 1962. Overwintering of western equine encephalomyelitis virus in garter snakes experimentally infected by Culex tarsalis. Proc Soc Exp Biol Med 109:421–424. Turgeon ML. 2003. Immunology and Serology in Laboratory Medicine. 3rd edition. Mosby, St. Louis, MO. Tyler JW and Cullor JS.1989. Titers, tests and truism: rational interpretation of diagnostic serologic testing. J Amer Vet Med Assoc 194:1550–1557. Walder R, Suarez OM, and Calisher CH. 1984. Arbovirus studies in the Guajira region of Venezuela: activities of eastern equine encephalitis and Venezuelan equine encephalitis viruses during an interepizootic period. Am J Trop Med Hyg 33:699–707. Wendland LD, Brown DR, Klein PA, and Brown MB. 2006. Upper respiratory tract disease (mycoplasmosis) in tortoises, in Reptile Medicine and Surgery, 2nd edition, Mader DR (Ed.), Saunders, St. Louis, MO, 931–938. Wozniak E, McBride J, DeNardo D, Tarara R, Wong V, and Osburn B. 2000. Isolation and characterization of an antigenically distinct 68-kd protein from nonviral intracytoplasmic inclusions in boa constrictors chronically infected with the inclusion body disease virus (IBDV: Retroviridae). Vet Pathol 37:449–459. Xiao L, Ryan UM, Graczyk TK, Limor J, Li L, Kombert M, Junge R, Sulaiman IM, Zhou L, Arrowood MJ, Koudela B, Modry D, and Lal AA. 2004. Genetic diversity of Cryptosporidium spp. in captive reptiles. Appl Environ Microbiol 70:891–899. Yadav MF, Sethi MS. 1979. Poikilotherms as reservoirs of Q-fever (Coxiella burnetii) in Uttar Pradesh. J Wildl Dis 15:15–17. Zapata AG, Varas A, and Torroba M. 1992. Seasonal variations in the immune system of lower vertebrates. Immunol Today 13:142–147.

9 Viruses and Viral Diseases of Reptiles Elliott R. Jacobson

Contents 9.1 General Comments................................................ 396 9.2 Herpesviridae......................................................... 396 9.2.1 General Characteristics............................. 396 9.2.2 Chelonia..................................................... 396 9.2.3 Crocodylia.................................................. 399 9.2.4 Sauria......................................................... 399 9.2.5 Ophidia...................................................... 400 9.3 Adenoviridae.......................................................... 401 9.3.1 General Characteristics............................. 401 9.3.2 Chelonia..................................................... 401 9.3.3 Crocodylia.................................................. 401 9.3.4 Sauria......................................................... 401 9.3.5 Ophidia...................................................... 402 9.4 Poxviridae.............................................................. 403 9.4.1 General Characteristics............................. 403 9.4.2 Chelonia..................................................... 403 9.4.3 Crocodylia.................................................. 403 9.4.4 Sauria......................................................... 403 9.5 Iridoviridae............................................................. 404 9.5.1 General Characteristics............................. 404 9.5.2 Chelonia..................................................... 404 9.5.3 Sauria......................................................... 405 9.5.4 Ophidia .................................................... 405 9.6 Papillomaviridae.................................................... 406 9.6.1 General Characteristics............................. 406 9.6.2 Chelonia..................................................... 406 9.6.3 Sauria......................................................... 406 9.7 Parvoviridae .......................................................... 406 9.7.1 General Characteristics............................. 406 9.7.2 Sauria......................................................... 406 9.7.3 Ophidia...................................................... 406 9.8 Circoviridae............................................................ 406 9.8.1 General Characteristics............................. 406 9.8.2 Chelonia..................................................... 407 9.9 Paramyxoviridae.................................................... 407 9.9.1 General Characteristics............................. 407 9.9.2 Chelonia..................................................... 407 9.9.3 Crocodylia.................................................. 407 9.9.4 Sauria......................................................... 407 9.9.5 Ophidia...................................................... 407

9.10 Retroviridae............................................................ 409 9.10.1 General Characteristics............................. 409 9.10.2 Chelonia..................................................... 409 9.10.3 Crocodylia.................................................. 410 9.10.4 Rhynchocephalia....................................... 410 9.10.5 Sauria......................................................... 410 9.10.6 Ophidia...................................................... 410 9.11 Reoviridae ............................................................. 412 9.11.1 General Characteristics............................. 412 9.11.2 Chelonia..................................................... 412 9.11.3 Sauria ........................................................ 412 9.11.4 Ophidia...................................................... 412 9.12 Togaviridae............................................................. 412 9.12.1 General Characteristics............................. 412 9.12.2 Chelonia..................................................... 413 9.12.3 Crocodylia.................................................. 413 9.12.4 Sauria......................................................... 413 9.12.5 Ophidia...................................................... 413 9.13 Flaviviridae ........................................................... 413 9.13.1 General Characteristics............................. 413 9.13.2 Chelonia..................................................... 413 9.13.3 Crocodylia.................................................. 413 9.13.4 Sauria..........................................................414 9.13.5 Ophidia.......................................................414 9.14 Rhabdoviridae.........................................................414 9.14.1 General Characteristics..............................414 9.14.2 Chelonia......................................................414 9.14.3 Sauria..........................................................414 9.14.4 Ophidia.......................................................414 9.15 Caliciviridae............................................................414 9.15.1 General Characteristics..............................414 9.15.2 Ophidia.......................................................414 9.16 Picornaviridae.........................................................414 9.16.1 General Characteristics..............................414 9.16.2 Ophidia.......................................................414 9.17 Coronaviridae ........................................................415 9.17.1 General Comments.....................................415 9.17.2 Crocodylia...................................................415 9.18 Miscellaneous Viruses ...........................................415 Acknowledgments.............................................................415 References..........................................................................415

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9.1 General Comments Since 1976 an array of viruses have been either identified in tissue section using electron microscopy or isolated in cell culture from tissues of members of the orders Chelonia, Crocodylia, and Squamata (Essbauer and Ahne, 2001; Wellehan and Johnson, 2005). Except for a single report of a retroviral gene sequence (Tristem et al., 1995), there are no reports of viral infection of the tuatara (Sphenodon spp.), the only member in the order Rhynchocephalia. The majority of viruses identified in reptiles have been only circumstantially incriminated as causes of disease, with few studies fulfilling Koch’s postulates. In many of the older reports, viruses were identified using either electron microscopy or viral isolation. Several reptile viruses await isolation. Immunohistochemical staining has been used to identify viral antigens in tissue section. More recently, molecular techniques such as polymerase chain reaction (PCR) and in situ hybridization (see Chapter 7) are being used with increased frequency to identify viruses in tissues of reptiles or confirm identification of those isolated in tissue culture. This has revolutionized the ability to diagnose viral infections using small amounts of tissue or other biological samples such as mouth swabs and fecal samples. Many retroviruses have been identified in tumors of reptiles, but again, transmission studies have not been reported to confirm a causal relationship. In some situations, such as with western equine encephalitis virus, reptiles may serve as a reservoir host, with the virus being non-pathogenic in the infected reptile. Two species of turtles (Mauremys [formerly Clemmys] mutica and Chinemys [formerly Geoclemys] reevesii) were found to be infected with hepatitis B virus following inoculation with human sera that were positive for certain antigens of hepatitis B (Lee and Yoo, 1989), but natural infection has not been reported. Lastly, viruses of unknown disease potential have also been identified in reptiles. In this chapter, viral infections and viral diseases of reptiles will be discussed. DNA viruses (Herpesviridae, Adenoviridae, Poxviridae, Iridoviridae, Papillomaviridae, Parvoviridae, and Circoviridae) will be discussed first followed by RNA viruses (Paramyxoviridae, Retroviridae, Reoviridae, Togaviridae, Flaviviridae, Rhabdoviridae, Caliciviridae, and Picornaviridae).

9.2 Herpesviridae 9.2.1 General Characteristics Herpesviridae is a diverse family of enveloped, doublestranded DNA viruses with an icosahedral nucleocapsid and having a diameter generally ranging from 150 to 200 nm. Replication is within the nucleus of host cells. During stages of replication, when examining hematoxylin and eosin–stained (H&E) tissue sections, eosinophilic- to amphophilic-staining intranuclear inclusions may be seen. Inclusions will not be seen when the virus is in a latent state. They are found in many different orders of vertebrates including fish, amphib-

ians, reptiles, birds, and mammals. Herpesviruses have been further divided into the subfamilies Alphaherpesvirinae, Betaherpesvirinae, and Gammaherpesvirinae, initially based on properties of infection, host range, and behavior in culture. Phylogenetic relationships of herpesviruses are now formally based on genetic content, as defined by the homology of nucleic acid sequences and identification of particular genes unique to a virus subset. To date, published analyses of sequences indicate that all reptilian herpesviruses that have been analyzed belong to the subfamily Alphaherpesvirinae (McGeoch and Gatherer, 2005).

9.2.2 Chelonia 9.2.2.1 Herpesvirus Infection of Sea Turtles  A virus with the morphological appearance of herpesvirus has been shown to be the causative agent of epizootics of skin lesions termed grey patch disease in young green turtles (Chelonia mydas) between 56 and 90 days after hatching in aquaculture (Rebell et al., 1975). Skin lesions commenced as small circular papular lesions that coalesced into spreading patches (Jacobson, 1981) (Figure 9.1) containing epidermal cells and amphophilic intranuclear inclusions (Figure 9.2). Electron microscopy revealed inclusions to consist of viral particles having an electron-dense core. Particles were found enveloping from nuclear membranes, and mature enveloped particles found in the cytoplasm measured 160 to 180 nm. The most severe epizootics occurred in the summer, under stressful environmental conditions of high water temperatures (> 30°C), crowding, and organic pollution.  An epizootic characterized by pneumonia and caseous material covering the globes, within the oral cavity cranial to the glottis and within the trachea (Figures 9.3–9.5) was seen in green sea turtles more than one year old at Cayman Turtle Farm, Grand Cayman, British West Indies (Jacobson et al., 1986). This was named lung, eye, and trachea (LET) disease. Affected turtles were often seen with their mouths opened at the water surface, with harsh respiratory sounds. The disease spread quickly through a tank of turtles and generally had a clinical course of 2 to 3 weeks. Histologically, there were focal to diffuse areas of necrosis adjacent to the glottis (Figure 9.6) and extending into the trachea (Figure 9.7). In some areas there was ballooning degeneration and necrosis of tracheal mucosal epithelial cells, with amphophilic intranuclear inclusions (Figure 9.8). There was an interstitial and bronchopneumonia, with necrotic debris and inflammatory cells filling area passageways (Figure 9.9). In certain areas, desquamated epithelial cells contained eosinophilic intranuclear inclusions (Figure 9.10). Using transmission electron microscopy (TEM), a herpesvirus was identified in tracheal epithelial cells. Green turtle kidney cells that were cultured and inoculated with lung and trachea of affected turtles produced giant cells (Figure 9.11), with subsequent cytolysis. Transmission electron microscopy revealed herpesvirus in

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infected cells (Figure 9.12). This virus was named lung, eye, and tracheal virus (LETV) of green turtles (Jacobson et al., 1986). Nucleic acid sequencing of portions of the UL26, DNA polymerase, and glycoprotein B genes was performed (Coberley, 2002; Coberley et al., 2002), and, based on this and subsequent analyses (McGeoch and Gatherer, 2005), there was strong evidence for placing these sea turtle herpesviruses in the subfamily Alphaherpesvirinae. Fibropapillomatosis (FP) was first reported in sea turtles over 60 years ago when tumors were identified in green turtles from the Florida Keys (Lucke, 1938; Smith and Coates, 1938). Based on visual observation alone or histological evaluation of lesions, FP appears to be present in several other species of marine turtles, including loggerhead (Caretta caretta), hawksbill (Eretmochelys imbricata), and olive ridley (Lepidochelys olivacea) sea turtles. Tumors are seen as papillary, arborizing masses on the body surface (Figures 9.13–9.16). In an early report, a nodule was found in the lung of one turtle, and was composed of cells similar to those in the dermal portion of the cutaneous tumors (Schlumberger and Lucke, 1948). From November 12, 1985 to June 11, 1986, 30 of 53 turtles (57 %) of green turtles captured in the Indian River Lagoon system of the east central Florida coast had FP. A series of green turtles with FP were collected from the same population and multiple biopsies were obtained from each in an effort to better understand the light and electron microscopic changes in affected tissues and to identify associated potential pathogens (Jacobson et al., 1989). Histologically, fibropapillomas consisted of a slightly to moderately hyperplastic epidermis overlying a thickened hypercellular dermis (Figure 5.43A). In the earliest lesions, ballooning degeneration was present predominantly in the stratum basale (Figure 5.43B), where rete ridges extended into the dermis; aggregates of mixed inflammatory cells were present around dermal vessels. As the lesions matured, they developed an arborizing, papillary pattern (Figure 9.17). The pathogenesis of spontaneous and experimentally induced fibropapillomas was subsequently studied in green turtles (Herbst et al., 1999). In turtles with cutaneous FP, internal masses have been seen in multiple visceral structures including the kidney, liver, lung, heart, and gastrointestinal tract (Herbst, 1994; Norton et al., 1990) (Figures 9.18–9.21). Histologically the masses represent fibromas (Figure 9.22). In 1991, two green turtles from Key West, FL had herpesvirus-like intranuclear inclusions within FP epidermal cells (Jacobson et al., 1991) (Figures 9.23–9.24). Using transmission electron microscopy, particles compatible with herpesvirus were seen in the inclusions (Figure 9.25). In experimental transmission studies of FP in green turtles using cell-free tumor extracts, both filtered and unfiltered tumor extracts successfully induced tumor development (Herbst et al., 1995). Experimental tumors were histologically indistinguishable from spontaneous tumors found in free-living turtles, but lacked spirorchiid trematode eggs, which are commonly seen in tumors in wild turtles. In this study, viruslike particles compatible with herpesvirus were identified in

tumors. PCR has been used to identify herpesvirus genes in fibropapillomas of green and loggerhead sea turtles (Lackovich et al., 1999; Quackenbush et al., 1998). Tumor cells from terminally ill turtles had from 2 to 20 copies of viral DNA per cell (Quackenbush et al., 2001). Marine leeches (Ozobranchus spp.) were found to carry very high herpesvirus DNA loads, with some approaching 10 million copies per leech (Greenblatt et al., 2004a). This finding implicated the marine leech as a mechanical vector for the FP-associated turtle herpesvirus. In a recent report, a 43,843-bp sequence of the chelonid fibropapilloma-associated herpesvirus (C-FP-HV), including a novel 4-kb segment, showed that C-FP-HV is related to other alphaherpesviruses (Herbst et al., 2004). Five fragments encompassing 6801 bp of the viral genome that were amplified from tumors indicated that there were five viral variants. Of these, three from Florida (A, B, and C) were nearly identical; variant D, isolated from loggerhead turtles in Florida and North Carolina, differed by 5.6% from the others, and the Hawaiian variant differed from the Florida variants A, B, and C by only 2.2%. The findings provided evidence that C-FPHVs have been associated with sea turtle hosts for millions of years, and the authors believe that their findings provided evidence that environmental or ecological factors underlie the current panzootic. Similar findings were reported in another genomic study with FP-associated turtle herpesvirus (Greenblatt et al., 2004b).

9.2.2.2 Herpesvirus Infection of Freshwater Turtles  Two captive Pacific pond turtles (Clemmys marmorata) that died of a fatal systemic disease had intranuclear hepatic inclusions associated with areas of necrosis (Frye et al., 1977). Inclusions were also seen in renal tubular epithelial cells and splenic cells. Electron microscopic evaluation of the liver revealed that intranuclear inclusions consisted of particles containing an electron-dense core and measured 100 nm in diameter. Intracytoplasmic particles were enveloped and measured approximately 140 nm. Based on size, conformation, and structure, the particles most closely resembled herpesvirus.  An adult male painted turtle (Chrysemys picta) that died 6 days after being treated for an abscess at the Metropolitan Toronto Zoo, was found on necropsy to have a friable liver with an inspissated gallbladder; pulmonary edema was also seen (Cox et al., 1980). Histopathology revealed randomly scattered microfoci of coagulation necrosis throughout the liver, and with the presence of large intranuclear inclusions. Similar appearing inclusions were seen in metaplastic epithelial cells in the lungs. Electron microscopic examination of the liver revealed numerous hexagonal capsids measuring 85 to 115 nm in size. The virus particles were presumptively categorized as herpesvirus. A die-off of captive map turtles (Graptemys spp.) was attributed to a virus that also morphologically resembled herpesvirus (Jacobson et al., 1982a). The examined turtles showed severe hepatic necrosis with eosinophilic to amphophilic intranuclear inclusions present in the liver (Figures 9.26–9.27),

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kidney, and pancreas. Typical herpesvirus particles were identified by electron microscopy in sections of liver. The die-off followed the introduction of western painted turtles (Chrysemys picta) into a long-term captive group of map turtles.

9.2.2.3 Herpesvirus Infection of Tortoises  The first report of a herpesvirus-like agent associated with a lesion in a tortoise is that involving a desert tortoise (Gopherus agassizii) (Harper et al., 1982). A 6-year-old cachectic desert tortoise in captivity since hatching was found to have a pharyngeal abscess. On histologic examination, intranuclear inclusions were observed in superficial epithelial cells of the palatine mucosa. Electron microscopy demonstrated various developmental stages of a virus morphologically compatible with herpesvirus. Other papers concerning herpesvirus in desert tortoises followed (Pettan-Brewer et al., 1996; Martinez-Silvestre et al., 1999). In a recent report (Johnson et al., 2005a), a captive desert tortoise from California with a severe pharyngitis (Figure 9.28) was found by light microscopy to have herpesvirus-like intranuclear inclusion bodies in mucosal epithelial cells (Figure 9.29). Using TEM, herpesvirus particles were detected in the mucosal epithelium, and herpesvirus nucleic acid sequences were amplified using PCR. This herpesvirus was determined to be distinct from the previously described (see below) tortoise herpesvirus 1 (THV-1). Similar to THV-1, this novel herpesvirus, tortoise herpesvirus-2 (THV-2), also clustered with the alphaherpesviruses (Johnson et al., 2005a).  In another report, twelve hundred of 2200 recently imported Argentine tortoises (Geochelone chilensis) died over a 3-month period; red-footed tortoises (Geochelone carbonaria) imported with the Argentine tortoises and housed with them remained clinically healthy (Jacobson et al., 1985a). At necropsy, there was necrotic debris in the nasal passageways, on the surface of the tongue, and scattered across the pharyngeal mucosa (Figure 9.30). By light microscopy, there was diffuse mucosal surface of the oral cavity (Figure 9.31) and desquamated degenerating epithelial cells contained eosinophilic intranuclear inclusions (Figure 9.32). Electron microscopy demonstrated that the inclusions consisted of viral particles containing an electron-dense core. Particles consistent with herpesvirus were seen enveloping from cell membranes, and mature enveloped particles measuring approximately 125 nm were seen in the cytoplasm (Figure 9.33). Viral isolation attempts in green sea turtle embryo fibroblasts were negative. There are several reports of herpesvirus infection of tortoises in Europe, with stomatis-glossitis representing the major gross lesion (Figure 9.34), and eosinophilic to amphophilic intranuclear inclusions present in oral mucosal epithelial cells (Figures 5.49, 9.35). Of 13 Greek (Mediterranean spurthighed) tortoises (Testudo graeca) from two private colonies in the United Kingdom, herpes-like particles were detected by electron microscopy in two animals with stomatitis (Cooper et al., 1988). Initially, while swabs taken from the oral lesions

resulted in the isolation of a variety of microorganisms, treatment with several systemic and local antibiotics had no effect on the course of the disease. Eventually, viral particles were demonstrated by electron microscopy within bronchial and palatine mucosal epithelium. Treatment of subsequent cases with 5% acyclovir ointment was encouraging. This was followed by other cases of herpesvirus in tortoises in Europe (Braune et al., 1989; Drury et al., 1998). In a study in the United Kingdom, oral or choanal swabs were collected at multiple sites from the following species of tortoises: Greek, Hermann (T. hermanni), Russian (Agrionemys [formerly Testudo] horsfieldii), Asia-minor spur-thighed tortoise (T. g. Iberia), marginated tortoise (T. marginata), and unidentified Testudo (Soares et al., 2004). PCR was used to test samples from 146 tortoises to determine DNA sequences of Mycoplasma agassizii and herpesvirus. Of the samples tested, 15.8% were positive for M. agassizii and 8.4% for chelonian herpesvirus. Mixed infections with both pathogens were present in only two tortoises (1.37%). Russian tortoises were more likely to be infected with M. agassizii than other tortoises, and herpesvirus was found more often in both marginated tortoises and Asia Minor spurthighed tortoises than other species. Herpesvirus is not confined to oral lesions of tortoises. By electron microscopy, herpesvirus-like particles have also been seen in the intestinal contents of a Hermann’s tortoise, several of which had caseous material in the upper digestive tract, hepatomegaly, and enteritis (Lange et al., 1989). Herpesvirus was identified in Hermann’s tortoises with meningoencephalitis (Heldstab and Bestetti, 1989). In 16 Hermann’s tortoises and 8 spur-thighed tortoises with necrotizing glossitis and stomatitis, intranuclear inclusions were found in epithelial cells in the tongue, trachea, bronchi, alveolae, endothelial cells of capillaries of the glomeruli, and within neurons and glial cells in the medulla oblongata and diencephalons (Muller et al., 1990). Electron microscopic examination of the liver and trachea demonstrated enveloped virions that were morphologically consistent with herpesvirus. The authors considered imported tortoises to be latent carriers of this virus. Hepatitis associated with herpesvirus infection was reported in the Russian tortoise (Hervas et al., 2002). The first isolate of a tortoise herpesvirus was from a Hermann’s tortoise in a collection of tortoises with stomatitis and rhinitis (Biermann and Blahak, 1994). Five tortoises were submitted for postmortem evaluation. Using light microscopy, characteristic eosinophilic intranuclear inclusions were seen in the epithelial cells of the esophagus, trachea, and tongue of all tortoises. Samples of brain, lung, trachea, intestine, and liver from different tortoises were prepared and inoculated onto Terrapene heart cells (TH-1). Herpesvirus was isolated from all the tissue samples. More than one strain of tortoise herpesvirus has been identified (Marschang, 1999; Marschang et al., 2001a). There are conflicting reports of differential morbidity and mortality of Mediterranean tortoises infected with herpesvirus. In the spring of 1995, a Greek tortoise was added to a

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collection of 21 Hermann’s tortoises and 2 Greek tortoises (Marschang et al., 1997). At the end of the summer of the same year, several Hermann’s tortoises started showing clinical signs of stomatitis and rhinitis with nasal and ocular discharge, necrotic lesions of the tongue, swelling of the cervical region and the lower jaw, anorexia, and lethargy. As of the mid-autumn of the same year, 13 of the original 21 Hermann’s tortoises had died. Several tortoises were submitted for necropsy, and although no inclusion bodies were observed in any tissues using light microscopy, herpesvirus was isolated from 10 of the 11 pharyngeal swabs collected from live tortoises in the collection, from the buffy coat of one live tortoise, and from several tissue samples obtained from necropsied tortoises. While blood samples were collected from tortoises in the collection, none of the Hermann’s tortoises had titers to herpesvirus. In contrast with these observations, in an epidemic of chronic rhinitis in 1990 in a private collection of Mediterranean tortoises in Spain consisting of 50 adult Greek tortoises, 12 Hermann’s tortoises, and 3 four-toed tortoises, only the Greek tortoises (n = 33) were affected (Muro et al., 1998). None of the Hermann’s or four-toed (Russian) tortoises showed clinical signs of disease. Of the ill Greek tortoises, 8 died and 12 were euthanatized during the same year. By light microscopy, lesions were limited to the oral cavity and the respiratory system, varying in severity among different tortoises. In the most severe cases, eosinophilic intranuclear inclusions were detected in the epithelial cells of the upper respiratory tract and mucosa of the tongue. Electron microscopy revealed particles of size, morphology, and structure consistent with those of herpesvirus. In Japan, an outbreak of herpesvirus was reported in recently imported pancake tortoises (Malacochersus tornieri) and four-toed (Russian) tortoises (Une et al., 1999). Degenerate primers directed to anneal with a conserved portion of the nucleotide sequence of the herpesviral DNA polymerase gene confirmed the presence of herpesvirus and indicated that it might belong to the subfamily Alphaherpesvirinae (Une et al., 2000). A further step in the molecular diagnostics of tortoise herpesvirus was the development of an in situ hybridization technique for the detection of tortoise herpesvirus nucleotide sequences in formalin-fixed, paraffin-embedded tissues (Teifke et al., 2000). In this report the ability of the DNA probe to hybridize with allegedly different herpesvirus strains from distant geographical locations suggested the possibility of a high degree of conservation in the nucleotide sequence of the helicase gene among the tortoise herpesviruses. Phylogenetic analysis based on the partial sequence of the helicase gene suggested the positioning of tortoise herpesvirus within the alpha subfamily. A causal relationship between tortoise herpesvirus infection and stomatitis-rhinitis of tortoises was recently fulfilled (Origgi et al., 2004). Greek tortoises that were experimentally infected (inoculated by intranasal and intramuscular routes) with two tortoise isolates, developed productive infections. Clinical signs of illness developed, which included conjunc-

tivitis, oral plaques, and oral discharge (Figure 9.36). Using PCR and reverse transcription PCR analyses, herpesvirus DNA sequences were found at multiple visceral sites. Additionally, 20 of the 40 positive samples were obtained from the cranial nerves and brains of the infected tortoises. The inability to isolate virus from tissues having viral DNA sequences suggested that viral latency is probably a feature of infection in tortoises.

9.2.3 Crocodylia There are only two reports of herpesvirus infection of crocodilians. In the first, 1 of 20 six-month-old saltwater crocodiles (Crocodylus porosus) that were used in an experimental stress study in Australia developed a surface crust on the abdominal skin (McCowan et al., 2004). Using light microscopy, H&Estained sections revealed large, amphophilic intranuclear inclusions. Using TEM, the inclusions consisted of viral particles consistent with herpesvirus. Recently, at a farm in North Carolina, American alligators (Alligator mississippiensis) with hemorrhagic lymphoid follicles in the cloaca were biopsied, and using consensus primers that targeted a conserved region of the polymerase gene were tested by PCR for the presence of herpesvirus gene sequences. An amplicon was produced and direct sequencing in both directions indicated that it was most closely related to tortoise herpesvirus-1 (Johnson et al., 2005b; Govett et al., 2005).

9.2.4 Sauria 9.2.4.1 Lacertid Lizard Papillomas  Papillomas are commonly seen in the European green lizard (Lacerta viridis) (Figure 9.37). Raynaud and Adrian (1976) presented a detailed report describing the distribution of papillomas in European green lizards maintained as a breeding colony. The papillomas ranged in diameter from 2 to 20 mm, and numbered 2 to 25 per individual. In females, the papillomas were most commonly found in the caudal lumbar area of the body, in the vicinity of the tail base; papillomas were rarely found around the head. In males, the papillomas had a dorsocranial distribution, around the base of the head. Neither males nor females had papillomas associated with ventral scales. The authors associated the distribution to the reproductive behavior whereby males inflict bite wounds on females at the base of the tail, and combative behavior between males during which males inflict bites at the base of the neck of other males. Histopathologic examination of papillomas showed hyperkeratosis and hyperplasia of epidermal cells (Figure 9.38). The nucleus of epidermal cells was often hypertrophied with margination of chromatin material and intranuclear inclusions. By electron microscopy, viral particles compatible with those of herpesvirus were identified. In addition, particles morphologically similar to papova- and reovirus were demonstrated.

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9.2.4.2 Iguana Herpesvirus  A herpesvirus was isolated from heart cell cultures of a green iguana (Iguana iguana) (Clark and Karzon, 1972; Zeigel and Clark, 1972). No lesions were seen in the iguana. Inoculation of 12 young iguanas produced no consistent pattern of lesions, and although there was a higher mortality rate in the inoculated lizards, no causal relationship was established. Herpesvirus was subsequently identified in iguanas having histiocytic lymphoid infiltration of the liver, spleen, myocardium, and bone marrow (Frye et al., 1977). 9.2.4.3 Herpesvirus-Associated Visceral Necrosis  All 9 wild-caught red-headed agamas (Agama agama) purchased by a zoological institution died over a 6-month period (Watson, 1993). In one of the agamids there were areas of necrosis in the liver and spleen, with intranuclear inclusions seen in these areas. A second agamid had inclusions in the lung. Using electron microscopy, a virus consistent with herpesvirus was found in the tissues from these lizards. A San Esteban Chuckwalla (Sauromalus varius) that died without showing any previous signs of illness was found by light microscopy to have diffuse hepatic necrosis, with many hepatocytes having intranuclear inclusions (Figure 9.39); the inflammatory response was not significant (Wellehan et al., 2003). The appearance of viral particles on electron microscopy was consistent with a herpesvirus (Figure 9.40). Degenerate PCR primers targeting a conserved region of herpesvirus DNAdependent DNA polymerase were used to amplify products from liver tissue. Nucleotide sequencing of the PCR product showed that the sequence from this lizard was unique. This virus was termed iguanid herpesvirus 2 and phylogenetic and comparative sequence analysis of this study suggested that this virus was a novel member of the subfamily Alphaherpesvirinae. However, it was the opinion of the authors of another study (McGeoch and Gatherer, 2005), focusing on the phylogeny of the reptilian herpesviruses, that more sequence data was needed before categorization in the Alphaherpesvirinae. 9.2.4.4 Herpesvirus-Associated Oral Lesions in Lizards  Herpesviruses have been found in several lizards with oral lesions. A herpesvirus was seen associated with stomatitis in two species of plated lizards (Gerrhosaurus spp.) and specific identification was made using PCR and sequencing (Wellehan et al., 2004a). Gingival samples from four green tree monitor lizards (Varanus prasinus) (from two different collections) with proliferative stomatitis (Figure 9.41) were examined using PCR primers targeting a conserved region of herpesvirus DNA-dependent DNA polymerase (Wellehan et al., 2005a). DNA was extracted from tissue and amplicons were obtained from three of the four cases. The amplicons were sequenced and comparative sequence analysis supported these viruses as novel members of the subfamily Alphaherpesvirinae; this was named varanid herpesvirus 1. DNA in situ hybridization of tissues from three lizards was positive for herpesvirus in the oral basal epithelium mucosa of all three lizards and the brain of two lizards.

9.2.5 Ophidia 9.2.5.1 Venom Gland Herpesvirus  Padgett and Levine (1966), in attempting to elucidate the substructure of murine Rauscher leukemia virus, incubated venoms of the Indian cobra (Naja naja) and the banded krait (Bungarus fasciatus) with mouse oncornavirus. Using electron microscopy, particles similar to capsids of herpesvirus were observed. In a subsequent electron microscopic study of venoms of Indian cobras and banded kraits, similar appearing particles were demonstrated (Monroe et al., 1968). These particles were from 100 to 125 nm in diameter and possessed a central electron-dense core typical of herpesvirus. Light microscopic examination of venom glands of two of 25 Siamese cobras (N. n. kaouthia) with a history of low-grade venom production revealed degeneration and necrosis of patches of columnar glandular epithelium with infiltration of the subepithelium with mixed inflammatory cells (Simpson et al., 1979). By electron microscopy, naked and enveloped herpesvirus-like particles were seen in necrotic and ruptured cells. The author of this chapter has seen particles of herpesvirus (Figure 6.50) in venom of Mojave rattlesnakes (Crotalus scutulatus). 9.2.5.2 Herpesvirus Infection of Boa Constrictors  In a series of 16 neonatal boa constrictors (Boa constrictor) that were captive-born in a zoologic park, nine were stillborn and six died within the first year of life (Hauser et al., 1983). Histopathologic evaluation revealed that two of the dead boa constrictors had hepatic necrosis with presence of amphophilic intranuclear inclusions. One snake also had inclusions in the pancreas and adrenal cortex, while the other snake had inclusions in the kidneys associated with an acute exudative glomerulonephritis. By electron microscopy, the intranuclear inclusions consisted predominantly of nucleocapsids usually arranged in a crystalline array. Unenveloped and enveloped particles were seen in the cytoplasm, with the latter measuring 115 to 125 nm. 9.2.5.3 Mixed Viral-Associated Gastrointestinal Diseases of Snakes  Gastrointestinal disease in a four-lined rat snake (Elaphe quatuorlineata), a boa constrictor, an Aesculapian snake (Elaphe longissima), and a gaboon viper (Bitis gabonica) were associated with a variety of viruses including adeno-, parvo-, picorna-, and herpesvirus-like viruses (Heldstab and Bestetti, 1984). Clinical signs prior to death were a necrotizing stomatitis and suspected intestinal disease in the rat snake, signs of central nervous system (CNS) disease in the boa constrictor, and loss of appetite and yellow gelatinous feces in the Aesculapian snake; the gaboon viper did not exhibit any signs of illness preceding death. Isolation attempts were not reported and a causal relationship was not established.

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9.3 Adenoviridae 9.3.1 General Characteristics Members of the family Adenoviridae are nonenveloped, linear, double-stranded DNA viruses with an icosahedral nucleocapsid and having a diameter generally ranging from 80 to 110 nm. Replication is within the nucleus of host cells. During stages of replication, when examining H&E-stained tissue sections, intranuclear inclusions may be seen. With H&E staining, while inclusions are typically basophilic, eosinophilic inclusions also have been seen. The virus is released when the cell lyses. Adenoviruses are categorized into the following genera: Mastadenovirus (mammals), Aviadenovirus (birds), and two recently accepted genera, Atadenovirus (ruminants, birds, snakes, lizards, and a marsupial) and Siadenovirus (frog, poultry). A fifth genus is proposed for a sturgeon adenovirus.

9.3.2 Chelonia The only report of an adenovirus in a chelonian is the isolation of an adenovirus (along with a herpesvirus) from a leopard tortoise (Geochelone pardalis) with biliverdinuria, wasting, and episodes of hemorrhage (McArthur et al., 2004).

9.3.3 Crocodylia 9.3.3.1 Nile Crocodile Adenovirus  Light microscopic findings in two 8-month-old Nile crocodiles (Crocodylus niloticus) from a crocodile farm in Zimbabwe, included multifocal to diffuse areas of hepatic necrosis with basophilic intranuclear inclusions (Figure 9.42) in one crocodile, and necrotizing enteritis containing similar staining intranuclear inclusions in crypt epithelial cells (Figure 9.43) in the second crocodile (Jacobson et al., 1984). Other reports followed (Foggin 1987, 1992). By electron microscopy, inclusions in the small intestine and liver were found to be composed of crystalline arrays of viral particles (Figure 9.44). Viral particles ranged in diameter from 75 to 80 nm, had hexagonal outlines, an electron-dense core, and were not enveloped. All of these features were compatible with adenovirus. Using negative staining electron microscopy, particles consistent with adenovirus were seen in feces of three Nile crocodiles originating from a farm in Mozambique (Huchzermeyer et al., 1994).

9.3.4 Sauria 9.3.4.1 Adenovirus Infection of Chameleons  A 6-monthold, 15-gram captive-born Jackson’s chameleon (Chamaeleo jacksoni) became anorexic and died 3 days later (Jacobson and Gardiner, 1990). A necropsy was performed and histopathology revealed a proliferation of the mucosal epithelium lining the esophagus and trachea. In H&E-stained tissue sections, eosinophilic intranuclear inclusions were seen within many of

the epithelial cells of both the esophagus and trachea. Based upon size, morphologic characteristics, and location, the viral particles seen were most compatible with those of adenovirus. Light microscopic examination of tissues of a wild-collected adult male mountain chameleon (Chamaeleo montium) that died after a 28-day history of anorexia revealed moderate numbers of round to ellipsoid, basophilic, intranuclear inclusions within enterocytes in the small intestine (Kinsel et al., 1997). Ultrastructurally, the inclusions consisted of crystalline arrays of hexagonal viral particles measuring 67 to 76 nm in diameter with electron-dense cores, which were consistent with an adenovirus. Except for the presence of inclusions, no additional light microscopic changes were seen.

9.3.4.2 Adenovirus-Associated Hepatic Necrosis of Lizards  A captive bearded dragon (Amphibolurus barbatus) in New Zealand that died following intermittent periods of inappetence was found on histological examination of tissues to have numerous foci of coagulative necrosis of the liver, with hepatocytes containing eosinophilic intranuclear inclusions (Julian and Durham, 1985). Electron microscopy demonstrated the inclusions to be composed of viral particles that were morphologically consistent with those of adenovirus. A similar virus was reported in a group of genetically related Rankin’s dragon lizards (Pogona henrylawsoni) (Frye et al., 1994). Seven of nine lizards had large basophilic intranuclear inclusions within hepatocytes. In a subsequent report, of four neonate bearded dragons (Pogona vitticeps) from two collections that became ill and died, one had basophilic intranuclear inclusions within areas of hepatic necrosis (Figure 9.45), and three had similar inclusions within enterocytes scattered throughout the small intestine (Jacobson et al., 1996). Using electron microscopy, particles consistent with adenovirus (Figure 6.52) and Dependovirus (Figure 6.70) were observed. Similar lesions and viral particles were seen in bearded dragons dying with signs of neurologic disease (Kim et al., 2002). In a savannah monitor (Varanus exanthematicus) that died without premonitory signs, light microscopy revealed multifocal areas of hepatic necrosis with degenerating hepatocytes containing basophilic intranuclear inclusions (Jacobson and Kollias, 1986). The myocardium also contained multifocal areas of necrosis, and endothelial cells contained similar basophilic intranuclear inclusions. Electron microscopic findings confirmed the inclusions to be composed of adenoviruslike viral particles.   PCR was recently used to taxonomically categorize adenoviruses from samples collected from the following lizards: fat-tail geckos (Hemitheconyx caudicinctus) with a lymphoplasmacytic enteritis and intranuclear inclusion bodies; leopard geckos (Eublepharis macularius) with weight loss; a tokay gecko (Gekko gecko) that had a severe proliferative gastritis with intralesional protozoa consistent with Cryptosporidium spp; a Gila monster (Heloderma suspectum) with signs of regurgitation; a blue-tongued skink (Tiliqua scincoides intermedia) with numerous intranu­clear basophilic inclusions in

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the small intestine; a bearded dragon that had intranuclear inclusions in the intestinal mucosa, hepatocytes, and bile ducts; and a mountain chameleon with adenoviral inclusions within enterocytes (Wellehan et al., 2004b). Sequencing results indicated that these lizards were infected with adenoviruses that were sufficiently different to be con­sidered distinct species. All were grouped within the genus Atadenovirus.

9.3.5 Ophidia 9.3.5.1 Adenovirus-Associated Hepatic Necrosis in Boa Constrictors  A 4-kg adult boa constrictor submitted for postmortem evaluation was found to have a grossly enlarged liver, with swollen borders and pale areas scattered throughout (Jacobson et al., 1985b). The most significant light microscopic lesion was severe diffuse hepatic necrosis with an infiltrate of heterophils and small mononuclear cells. Numerous large basophilic intranuclear inclusions, resulting in ballooning of nuclei and margination of chromatin material, were seen scattered throughout the liver (Figure 9.46); electron microscopy revealed that inclusions consisted of adenovirus (Figure 9.47). Intranuclear inclusions can be found in the small intestine (Figure 5.50). A neonatal boa constrictor injected with liver suspensions of this snake was found dead in its cage at 14 days following inoculation; histopathologic evaluation of multiple organs demonstrated multifocal areas of hepatic necrosis, and a morphologically identical virus was isolated in cultures of viper heart cells grown at 25°C. In two rosy boas (Lichanura trivirgata) that died with an adenovirus-like infection, one snake had hepatic necrosis with basophilic inclusions in hepatocytes, renal tubular epithelial cells, and endocardium, and in the second snake, inclusions within hepatocytes and heterophils within sinusoids of the liver were seen (Schumacher et al., 1994a). Using an aviadenovirus-specific oligoprobe and an in situ hybridization technique, adenoviral DNA was identified in inclusion-bearing nuclei of hepatocytes of one boa constrictor with hepatic necrosis (Ramis et al., 2000) and within inclusion-bearing hepatocytes, Kupffer cells, and endothelial cells lining sinusoids of the liver of another boa constrictor with concomitant inclusion body disease and amoebiasis (Perkins et al., 2001). A PCR primer designed to amplify a 1700-bp segment of Atadenovirus was used to amplify a fragment of an adenovirus isolated from a boa constrictor (Marschang et al., 2003). The sequence was identical to a DNA fragment sequenced from a corn snake (Elaphe guttata guttata) that died with pneumonia (Farkas et al., 2002).

blunting and dilatation, and mucosal necrosis and hyperplasia of the small intestine. With H&E staining, numerous enterocytes had basophilic intranuclear inclusions (Figure 9.48). Similar inclusions were seen in gastric epithelial cells. Electron microscopy demonstrated that the inclusions consisted of viral particles consistent in size and morphology to adenovirus. Aggregates of smaller nonenveloped parvovirus-like (Dependovirus) particles were seen around the larger particles. In another report, a 30-year-old coastal mountain kingsnake (L. z. multifasciata) that developed signs of neurological disease, had foci of gliosis within the caudal cerebrum, brainstem, and cervical spinal cord (Raymond et al., 2003). With H&E staining, magenta-staining intranuclear inclusions were observed in glial cells and endothelial cells. Electron microscopy revealed that inclusions consisted of viral-like particles consistent with adenovirus, and a DNA in situ hybridization technique using adenovirus-specific oligonucleotide probes resulted in positive staining of inclusion-bearing nuclei.   The author of this chapter has seen several species of colubrid snakes (Arizona mountain kingsnakes [Lampropeltis pyromelena], corn snakes, and pine snakes [Pituophis melanoleucus]) with severe enteritis having adenoviral intranuclear inclusions in enterocytes of the small tract (Figures 9.49–9.50).

9.3.5.3 Adenovirus Infection in Viperid Snakes  Using light microscopy, a Mojave rattlesnake (Crotalus scutulatus) with an abdominal mass was found to have severe multifocal hemorrhage and ulceration of the intestinal tract with intralesional amoebae and basophilic intranuclear inclusion bodies within enterocytes (Perkins et al., 2001). Electron microscopy revealed that inclusions consisted of viral-like particles consistent with adenovirus, and a DNA in situ hybridization technique using adenovirus-specific oligonucleotide probes, resulted in positive staining of inclusion-bearing nuclei.  An adult captive-born palm viper (Bothriechis marchi), which was found dead in its enclosure and subsequently necropsied, was found to have esophagitis and stomatitis (Raymond et al., 2002). In hematoxylin and eosin tissue sections, eosinophilic intracytoplasmic inclusion bodies consistent with those seen in inclusion body disease of boid snakes were seen in multiple tissues, and basophilic intranuclear inclusions were seen in mucosal epithelial cells lining the oral cavity and esophagus. Transmission electron microscopy demonstrated that the intranuclear inclusions were composed of viral particles consistent with those of adenovirus. 9.3.5.4 Mixed Viral-Associated Gastrointestinal Diseases of Snakes  As mentioned previously, gastrointestinal disease

9.3.5.2 Adenovirus Infection in Colubrid Snakes  A in a four-lined rat snake, a boa constrictor, an Aesculapian

group of seven 6-week-old Sierra mountain kingsnakes (Lampropeltis zonata multicincta), started regurgitating and suddenly died (Wozniak et al., 2000a). Necropsies were performed and tissues were collected from three snakes for histopathology. Light microscopy revealed segmental villus

snake, and a gaboon viper were associated with a variety of viruses including adeno-, parvo-, picorna-, and herpesviruslike viruses (Heldstab and Bestetti, 1984).

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9.3.5.5 Isolation of Adenoviruses from Tissues of Moribund Snakes  An adenovirus (SnAdV-1) was isolated in iguana heart

9.4.3 Crocodylia

(IgH-2) cells and terrapene heart (TH-1) cells from the internal organs of a moribund corn snake exhibiting clinical signs of pneumonia (Juhasz and Ahne, 1992). Highest infectivity was obtained in IgH-2 cells at 25°C. Infected IgH-2 cells showed the development of three morphologically different intranuclear inclusion bodies. During viral assembly, the particles formed typical crystalline arrays in the nucleus. Approximately 60% of the genome of this virus was cloned and sequenced (Farkas et al., 2002). Homology searches showed that genes (most importantly p32K) of SnAdV-1 were closest to those of members of the genus Atadenovirus. Results of this study suggested a reptilian origin for Atadenovirus.  Another adenovirus-like agent was isolated from a moribund royal python (Python regius) (Ogawa et al., 1992). The virus replicated in IgH2-cells at 30°C forming eosinophilic intranuclear inclusion bodies. The virus proved to be stabile to treatment with chloroform, pH 3 and pH 12 but it was labile to heat (56°C).

9.4.3.1 Caiman Poxvirus  The first report of a virus-associ-

9.4 Poxviridae 9.4.1 General Characteristics Poxviruses are large (220 to 450 nm long), ovoid to brick-shaped double-stranded DNA viruses that replicate and assemble within cytoplasmic viroplasm of infected cells. They are enveloped and the core is cylindrical or biconcave. The family is divided into the two subfamilies: Chordopoxvirinae (vertebrate poxviruses) and Entomopoxvirinae (invertebrate poxviruses). There are multiple genera within these families. Histologically, using H&E staining, eosinophilic intracytoplasmic inclusions are often seen in cells where a virus is replicating.

9.4.2 Chelonia There is a single report of poxvirus infection in a chelonian. A Hermann’s tortoise developed small white-yellow papular lesions of the lower eyelids of both eyes and the rostrum near the right eye (Oros et al., 1998a). The tortoise subsequently showed respiratory distress, developed a nasal discharge, and died two weeks later. At necropsy the skin lesions had increased in size. Light microscopy revealed that deep epidermal cells within the skin lesions contained eosinophilic intracytoplasmic inclusions. The turtle also had a bronchopneumonia with bacterial colonies and inflammatory cells within the lumen of the bronchi. Transmission electron microscopy of the skin lesions revealed particles consistent with those of poxvirus.

ated disease in a crocodilian is that of a poxvirus demonstrated in skin lesions of captive caimans (Caiman crocodilus) (Jacobson et al., 1979). Several captive juvenile caimans were submitted with gray-white circular skin lesions scattered over the head, mid-body surface, and tail (Figures 9.51– 9.52). While in some caiman there were severe lesions that resulted in sloughing of digits, other individuals only had focal lesions, often involving the palpebrae (Figure 9.53). Light microscopic evaluation of skin lesions showed large eosinophilic intracytoplasmic inclusions within hypertrophied epithelial cells (Figures 9.55– 9.56). Numerous inclusions were present in an extremely thickened keratin layer. Electron microscopy demonstrated that the inclusions consisted of myriads of viral particles that were morphologically typical of poxvirus (Figures 6.56–6.58). The size of 200 x 100 nm was smaller than previously reported poxviruses of vertebrates and insects. Subsequently, caiman pox was identified in Hungary (Vetesi et al., 1981), South Africa (Penrith et al., 1991), Brazil (Cubas, 1996; Matushima and Ramos, 1995), and Colombia (Villafane et al., 1996).

9.4.3.2 Crocodile Poxvirus  A poxvirus was identified in juvenile farm-reared Nile crocodiles (Crocodylus niloticus) in Zimbabwe in 1982 (Foggin, 1987). This was followed by reports at other Nile crocodile farms in Africa (Buoro, 1992; Horner, 1988; Huchzermeyer et al., 1991; Pandey et al., 1990) and saltwater and freshwater (Crocodylus johnstoni) crocodile farms in Australia (Buenviaje et al., 1992, 1998). Crusty brown lesions were seen in the oral cavity, head, eyelids, palmar surface of feet, and ventral and lateral body surfaces (Figures 9.57–9.59). Skin lesions were raised and ulcerated. The outbreaks occurred in hatchlings and juveniles less then 2 years of age. As with caiman pox, while morbidity may be high, mortality was generally low, with lesions spontaneously regressing. Histologically there was hyperkeratosis and parakeratosis with hypertrophic epithelial cells containing intracytoplasmic inclusions (Figure 9.60). Using electron microscopy, numerous dumbbell shaped virions were seen. Based on their electron microscopic morphology, Gerdes (1991) tentatively classified this virus as a Parapoxvirus.

9.4.4 Sauria 9.4.4.1 Poxvirus Infection of a Tegu  Poxviral dermatitis was reported in a captive tegu (Tupinambis teguixin) (Stauber and Gogolewski, 1990). Multiple brown-colored papules were observed on the integument and histologic examination revealed that epidermal cells contained large intracytoplasmic eosinophilic inclusions. Electron microscopy demonstrated that inclusions contained viral particles morphologically compatible with those of poxvirus. The remaining lesions healed spontaneously in 3 to 4 months after they were first noticed.

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9.4.4.2 Poxvirus Infection of a Flap-Necked Chameleon  Of blood films examined from 170 specimens of 15

9.5.2 Chelonia

Chamaeleo spp. in Tanzania, 3 of 50 flap-necked chameleons (C. dilepis) were found to have intracytoplasmic inclusions within circulating monocytes (Jacobson and Telford, 1990). One of the chameleons was maintained in captivity and was sequentially bled. At 46 days, a second type of inclusion was occasionally seen within monocytes. The lizard became ill and was euthanatized on day 55. All circulating monocytes were found to have either one or both types of inclusions (Figure 9.61). Histologic examination of multiple tissues revealed similar inclusions within macrophages in the spleen (Figure 9.62) and liver. By TEM, the first inclusion seen was composed of poxvirus while the second, less commonly seen inclusion, contained developmental stages of chlamydia (Figures 6.60–6.61). The first inclusion was similar in appearance to inclusions described for jewelled chameleons (C. lateralis) from Madagascar (Brygoo, 1963), and although these were considered to be viral inclusions, ultrastructural studies were not done.

9.5.2.1 Iridovirus Infection of European Tortoises  The

9.5 Iridoviridae 9.5.1 General Characteristics Iridoviruses are large (120 to 200 nm) double-stranded cytoplasmic DNA viruses. Iridoviruses can occur as enveloped and nonenveloped forms; both are infectious. Replication occurs in two stages; the first stage is in the nucleus and second stage is in the cytoplasm. The family Iridoviridae consists of four genera. Two genera, Chloriridovirus and Iridovirus, infect insects. Lymphocystivirus infects fish, and Ranavirus has been shown to be capable of infecting fish, amphibians, and reptiles (Mao et al., 1997). Frog virus 3 (FV3), the type species for the genus Ranavirus, was first isolated in 1966 from a renal carcinoma in a leopard frog (Granoff et al., 1965), although it was subsequently determined that there was no association of the virus with the tumor (Granoff et al., 1966). Another Ranavirus, which was originally named tadpole edema virus (TEV), was recovered from bullfrog tadpoles manifesting a syndrome characterized by edema and visceral necrosis (Wolf et al., 1968). Significant research with amphibian iridoviruses did not make much progress until the early 1990s when worldwide declines in amphibians brought new interest regarding the role of these viruses in amphibian mortality events. In a study of 64 amphibian mortality and morbidity events, iridovirus was the most common cause of mortality (Green et al., 2002). While iridoviruses have been reported as pathogens in reptiles, they have not received as much attention as those in amphibians. In vertebrates, eosinophilic to basophilic intracytoplasmic inclusions may be seen in a variety of tissues processed for light microscopy and stained with hematoxylin and eosin. Inclusions are generally not numerous. Inclusions may also be seen in red blood cells in a peripheral blood film stained with Wright-Giemsa stain. Some reptiles have been seen with numerous iridoviral inclusions in circulating red blood cells without any apparent adverse effects.

first report of iridovirus infection of a tortoise involved a spur-tailed (Hermann’s) tortoise found on necropsy to have multiple grey spots disseminated throughout the liver; the spleen was congested and small white foci were seen on the cut surface (Heldstab and Bestetti, 1982). Histopathology revealed multiple areas of hepatic necrosis with hepatocytes adjacent to necrotic areas containing strongly basophilic intracytoplasmic inclusions; inclusions were also seen in mucosal epithelial cells of the intestine. By TEM, intracytoplasmic inclusions consisted of accumulations of hexagonal virus particles located around electron-dense material. A few free virions were also observed in nuclei. The virions were composed of an electron-dense nucleocapsid enclosed in a hexagonal envelope containing two or more distinct layers. Mean viral diameter varied between 140 and 160 nm. Based upon size and morphologic properties, the agent was considered to be an iridovirus. Several years later, an epidemic in a captive group of Hermann’s tortoises was reported (Muller et al., 1988). While iridovirus was identified in a Horsfield’s (Russian) tortoise in the United States, no disease or pathology was mentioned (Mao et al., 1997). Recent isolates from two of seven Hermann’s tortoises that died in a zoo in Switzerland were found by PCR to have major capsid protein sequences closely related to the Ranavirus FV3 (Marschang et al., 1999). This was the first report linking a reptile iridovirus with that of an amphibian.

9.5.2.2 Iridovirus Infection of Tortoises and Turtles in the United States  The first report of iridovirus infection in a wild tortoise involved a gopher tortoise (Gopherus polyphemus) in Florida that had signs of upper respiratory tract disease (Westhouse et al., 1996). Necropsy revealed an ulcerative tracheitis (Figure 9.63), pneumonia, and ulcerative pharyngitis and esophagitis. By light microscopy, basophilic intracytoplasmic inclusions were seen within epithelial cells of the oral cavity, esophagus, and respiratory tract (Figures 5.51, 9.64). Electron microscopy revealed that inclusions consisted of particles compatible with iridovirus (Figure 9.65). There was no attempt at virus isolation and no molecular studies were performed. Starting in August 2003, iridovirus (Ranavirus) infection was identified in additional gopher tortoises in Florida, a dead captive Burmese star tortoise (Geochelone platynota) in a zoological collection in Georgia, from multiple eastern box turtles (Terrapene carolina carolina) at a nature sanctuary in Pennsylvania, and wild box turtles in Georgia and Florida (Johnson et al., 2004). All tortoises and box turtles had  overlapping signs, including cervical edema, palpebral edema, rhinitis, and stomatitis-glossitis (Figures 9.66–9.68). Basophilic intracytoplasmic inclusions were occasionally seen in visceral epithelial cells (Figures 9.69–9.70). While iridovirus was identified in a box turtle in another report in the United States, no disease or pathology was mentioned (Mao et al.,

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1997). Five of seven box turtles in a large collection in North Carolina died with fibrinoid vasculitis of multiple organs as the predominant lesion (De Voe et al., 2004). While inclusions were not seen within cells of any tissues, an iridovirus was isolated from two turtles. Sequence analysis of a portion of the major capsid protein resulted in the categorization as Ranavirus.

9.5.2.3 Iridovirus Infection of Soft-Shelled Turtles in China  In 1997, an outbreak of a disease called red neck disease occurred at a farming operation of Chinese soft-shelled turtles (Pelodiscus [formerly Trionyx] sinensis) in the Shenzhen area of China (Chen et al., 1999). A virus consistent with iridovirus was isolated in several fish cell lines. In experimental transmission studies, the isolated virus caused mortality in young turtles with typical clinical signs of the disease including neck swelling and ventral hemorrhage.

9.5.3 Sauria An iridovirus represents the first reptilian viral infection, although initially this intraerythrocytic agent was considered to be a protozoan and originally named Pirhemocyton tarentolae (Chatton and Blanc, 1914). The organism was seen in circulating red blood cells as intracytoplasmic inclusions. The viral nature of the inclusions was elucidated in subsequent studies (Stehbens and Johnston, 1966) in which TEM examination of red blood cells of an infected gecko (Gehyra variegata) revealed that the inclusions represented virus assembly pools or factory areas. This was presumptively considered to be an iridovirus based upon structure and morphology. Similar intracytoplasmic acidophilic-staining erythrocytic inclusions were seen in Giemsa-stained peripheral blood films in flap-necked chameleons (Chamaeleo dilepis) and a Fischer’s chameleon (Bradypodion fischeri) in Tanzania (Figures 9.71–9.72) (Telford and Jacobson, 1993). Using TEM, iridoviral particles were seen in red blood cells (Figures 6.63–6.64). The virus was named lizard erythrocyte virus (LEV). An iridovirus was isolated from feces and oral swabs of chameleons (four-horned chameleon [Chamaeleo quadricornis] and high-casqued chameleon [Chamaeleo hoehnelii]) in a captive breeding group of chameleons in the United Kingdom (Drury et al., 2002a). Negative staining electron microscopy also demonstrated a virus compatible with iridovirus in feces of a four-horned chameleon in the same colony. The pathogenicity of this virus was not determined. Iridoviruses were isolated from the lung, liver, and intestine of two bearded dragons, a four-horned chameleon, and the skin of a frill-necked lizard (Chlamydosaurus kingii) (Just et al., 2001). While a PCR primer designed to amplify the partial gene for the major capsid protein (MCP) of FV3 failed to amplify this sequence, primers designed to amplify this sequence for an invertebrate iridovirus (Chilo iridescent virus) resulted in 500-bp product with 97% identity to the nucleotide sequence of this virus and

100% identity to the sequences of Gryllus bimaculatus iridescent virus, another invertebrate iridovirus. The authors concluded that invertebrate iridoviruses might be transmitted to reptiles through the insects on which they feed. No histologic findings were reported on the cases, so it is unknown whether these viruses were pathogenic in their hosts. Iridoviruses were isolated from tissues of a deceased leaf-tailed gecko (Uroplatus fimbriatus) and a four-horned chameleon, and from a skin sample obtained from a live green iguana with skin disease (Marschang et al., 2002a). Sequencing of a large portion of the major capsid protein gene indicated that the isolate from the leaf-tailed gecko was related to FV3, the type species of the genus Ranavirus (Marschang et al., 2005).

9.5.4 Ophidia 9.5.4.1 Snake Erythrocyte Virus  The genus Toddia was created by França (1911) for intracytoplasmic eosinophilic inclusions associated with crystalloid bodies first seen by Dutton et al. (1907) in red blood cells of amphibians in tropical Africa and then by França (1911) in red blood cells of the square-marked toad (Bufo regularis) from Portuguese Guinea. Based on its cytological characteristics it was considered a protozoan. Of 163 cottonmouths (Agkistrodon piscivorus leucostoma) from Louisiana, Marquardt and Yaeger (1967) found Toddia in blood films of four. They described the formation of associated intracytoplasmic inclusions that progressed from a spheroid to crystalloid-square and believed that Toddia and Pirhemocyton of lizards to be taxonomically related. Based on the ultrastructural study of Stehbens and Johnston (1966), Marquardt and Yaeger (1967) considered both to be some type of DNA virus. Infection of red blood cells with Toddia with formation of intracytoplasmic crystalloid bodies was also described from the northern water snake (Nerodia sipedon sipedon) (Brooker and Yongue, 1982; Smith et al., 1994). Three types of inclusions were seen in infected cells of water snakes in Canada: translucent inclusions, small acidophilic bodies, and square-shaped crystalloids (Smith et al., 1994). Ultrastructural studies revealed that the smaller inclusion represented sites of membrane-bound viral assembly sites that were distinct from that seen in LEV. The authors recommended that the etiologic agent of Toddia infections of North American snakes should be renamed Snake Erythrocytic Virus (SEV). Johnsrude et al. (1997) described two types of inclusions (one viral and the other crystalline) in red blood cells of a severely anemic lancehead viper (Bothrops moojeni) that had SEV. The author of this chapter identified SEV inclusions in red blood cells of wild-collected, apparently healthy plains garter snakes (Thamnophis radix) (Figure 9.73) and a wild-collected eastern ribbon snake (T. sauritus) (Figure 9.74).

9.5.4.2 Ranavirus Infection of Green Tree Pythons  Histologic examination of two green tree pythons (Chondropython viridis) that died after illegal importation into Australia

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revealed ulceration of the nasal mucosa, necrotizing pharyngitis, and hepatic necrosis (Hyatt et al., 2002). The virus was isolated and characterized using immunohistochemistry, electron microscopy, polyacrylamide gel electrophoresis, enzymelinked immunosorbent assay, PCR, restriction endonuclease digestion, and DNA hybridization. The nucleotide sequence of the entire major coat protein of a green tree python isolate was 98% homologous with epizootic hematopoietic necrosis virus of fish and Bohle iridovirus of the burrowing frog (Limnodynastes ornatus), and 97% homologous with FV3. These findings supported the categorizing of the green tree python iridovirus as a new Ranavirus. The virus was provisionally named Wamena virus after the location in Irian Jaya where the snakes originated.

have icosahedral symmetry. Three genera infect vertebrates (Parvovirus, Erythrovirus, and Dependovirus) and three infect insects (Densovirus, Iteravirus, and Contravirus). Replication is within nuclei, and depending on the stage of infection, inclusions may or may not be seen. Because of the small genome, the virus is dependent upon host cellular function for replication. They generally replicate in actively dividing cells such as those in the intestinal tract and bone marrow.

9.6 Papillomaviridae

9.7.3 Ophidia

9.6.1 General Characteristics Members of the family Papillomaviridae are small (45 to 55 nm in diameter), nonenveloped, double-stranded DNA viruses that have icosahedral symmetry and replicate within host nuclei. Intranuclear inclusions may or may not be observable using light microscopy.

9.6.2 Chelonia Recently imported Bolivian side-neck turtles (Platemys platycephala) were submitted with circular papular skin lesions, which in some animals coalesced into patches (Jacobson et al., 1982b) (Figure 9.75). In some cases the lesions progressed to large areas of necrosis. Light microscopic evaluation of skin lesions revealed hyperkeratosis and hyperplasia with acanthosis; no inclusions were noted (Figure 9.76). Electron microscopic examination of skin biopsies revealed intranuclear crystalline arrays of hexagonal particles measuring approximately 42 nm (Figure 6.69). These aggregates were morphologically consistent with papillomavirus. However, the lesions never progressed into typical papillomas supported by fibrovascular stalks. Papillomavirus-like particles (49 mm in diameter) were identified using negative staining electron microscopy in a lung washing of a Horsfield’s Russian tortoise (Drury et al., 1998).

9.7.2 Sauria In two reports, bearded dragons with adenovirus infection of the intestinal tract and liver also had particles consistent with Dependovirus intermixed with adenovirus in infected nuclei (Jacobson et al., 1996; Kim et al., 2002) (Figure 6.70).

9.7.3.1 Viral-Associated Gastrointestinal Diseases of Snakes  Gastrointestinal disease in a four-lined rat snake (Elaphe quatuorlineata), a boa constrictor (Constrictor constrictor), an Aesculapian snake (Elaphe longissima), and a gaboon viper (Bitis gabonica) were associated with a variety of viruses including adeno-, parvo-, picorna- and herpesvirus-  like viruses (Heldstab and Bestetti, 1984). Clinical signs prior to death were a necrotizing stomatitis and suspected intestinal disturbances in the rat snake, signs of CNS disease in the boa constrictor, and loss of appetite and yellow gelatinous feces in the Aesculapian snake; the gaboon viper did not exhibit any signs of illness preceding death. Isolation attempts were not reported and a causal relationship was not established.

9.7.3.2 Dependovirus Infection  As previously mentioned, basophilic inclusions within enterocytes of 6-week-old Sierra mountain kingsnakes consisted of viral particles consistent in size and morphology with adenovirus and aggregates of smaller nonenveloped particles consistent with Dependovirus (Figure 9.77) (Wozniak et al., 2000a). Parvoviruses were isolated from a corn snake (Elaphe guttata) (Ahne and Scheinert, 1989) and a boa constrictor and royal (ball) python (Ogawa et al., 1992; Farkas et al., 2004). The isolate from the royal (ball) python was sequenced and identified as serpentine adeno-associated virus (SAAV) in the genus Dependovirus (Farkas et al., 2004).

9.6.3 Sauria Using electron microscopy, papillomas in European green lizard contained particles morphologically similar to herpesvirus, papovavirus, and reovirus (Raynaud and Adrian, 1976).

9.7 Parvoviridae 9.7.1 General Characteristics Members of the family Parvoviridae are small (18 to 26 nm in diameter), nonenveloped, single-stranded DNA viruses that

9.8 Circoviridae 9.8.1 General Characteristics The family Circoviridae consists of small (10 to 20 nm), nonenveloped virions that have icosahedral symmetry. The genome consists of circular, single-stranded DNA. There are two genera within the family. The genus Gyrovirus includes chicken anemia virus, having a negative-sense genome. The genus Circovirus has an ambisense genome and includes porcine

Viruses and Viral Diseases of Reptiles  407

circoviruses, duck circovirus, goose circovirus, and psittacine beak and feather disease virus (De Kloet and De Kloet, 2004; Phenix et al., 2001). While the virus replicates in the nucleus, viral inclusions are found both within nuclei and the cytoplasm of infected cells.

9.8.2 Chelonia

in feces of Nile crocodiles fed chickens from a farm having an outbreak of Newcastle disease (Huchzermeyer et al., 1994). In the same report, a paramyxovirus was also seen in the feces of a crocodile that was not fed poultry.

9.9.4 Sauria

The family Paramyxoviridae consists of single-stranded negative-sense RNA viruses that range in diameter from 150 to 300 nm and can exist in spheroidal or filamentous forms. Mature virus undergoes envelopment at the cytoplasmic membrane. Inclusions consisting of nucleocapsid strands can be found in either the nucleus or cytoplasm. Virions contain 10 to 12 proteins. The family Paramyxoviridae consists of the subfamilies Paramyxovirinae and Pneumovirinae. Within the Paramyxovirinae are the following genera: (1) Respirovirus (Sendai virus, human parainfluenza viruses 1 and 3); (2) Morbillivirus (measles, distemper-rinderpest group); (3) Rubulavirus (mumps, human parainfluenza 2 and 4); (4) Henipavirus (Hendra virus, Nipah virus); and (5) Avulavirus (Newcastle disease virus, avian paramyxoviruses). Within the subfamily Pneumovirinae are the genera Pneumovirus (human respiratory syncytial virus) and Metapneumovirus (avian metapneumovirus, turkey rhinotracheitis virus).

There are few reports of PMV infection of lizards. A paramyxolike virus was isolated from a false tegu (Callopistes maculatus) (Ahne and Neubert, 1991). Using partial sequences for the L and HN genes, comparisons were made to 16 snake isolates. The false tegu isolate was considered to be in an intermediate group. The only reported mortality event in lizards with a PMV involved three separate epidemics involving caiman lizards (Dracaena guianensis) that were imported into the United States (Jacobson et al., 2001a). Lizards showed few clinical signs; they were either found dead or were anorexic prior to death. Necropsies were performed on several lizards and indicated a severe pneumonia with caseous material in the lungs (Figure 5.14). Light microscopy revealed a proliferative heterophilic and histiocytic pneumonia (Figures 9.79–9.80). Using TEM, a filamentous virus was seen budding from cell membranes and intracytoplasmic nucleocapsid filaments compatible with PMV were seen in the cytoplasm of alveolar type II cells lining air passageways (Figure 9.81). An immunoperoxidase-staining technique using a rabbit polyclonal antibody raised against a snake paramyxovirus resulted in positive staining of a viral antigen in lung tissue sections. Paramyxovirus was isolated from lung homogenates inoculated into TH1 and viper heart cells. In another study, a virus consistent with a PMV was isolated from a cloacal swab obtained from a wild-caught Mexican lizard, Xenosaurus platyceps, which was housed in a zoological collection (Marschang et al., 2002b). A virus morphologically consistent with PMV was identified budding from epithelial cells lining the respiratory tract of a water dragon (Physignathus concinus) with a proliferative interstitial pneumonia (Boyer et al., 2005).

9.9.2 Chelonia

9.9.5 Ophidia

A captive painted turtle (Chrysemys sp.) that was found dead in its tank was necropsied and multiple tissues were collected for histologic evaluation. There were multifocal areas of necrosis in the spleen and liver, and macrophages contained multiple intracytoplasmic inclusions (Figure 9.78). Using TEM, inclusions consisted of small virions (10 to 20 nm) (Figure 6.71) that were compatible with members of the family Circoviridae.

9.9 Paramyxoviridae 9.9.1 General Characteristics

There is a single report indicating the presence of a paramyxovirus (PMV) in a chelonian. Mediterranean tortoises imported into Switzerland from Turkey arrived with dermatitis (Zangger et al., 1991). Using light microscopy, there was hyperkeratosis, parakeratosis, acanthosis, and ballooning degeneration of epidermal cells. Intracytoplasmic inclusions were seen in the stratum germinativum. Electron microscopy demonstrated pleomorphic particles (140 to 500 nm) having nucleocapsids measuring 12 nm in diameter. The particles were similar in size and appearance to paramyxovirus.

9.9.3 Crocodylia Using negative staining electron microscopy, PMV particles considered to be those of Newcastle disease virus, were seen

In 1976 a respiratory epizootic spread through a collection of vipers (first reported as occurring in fer-de-lance [Bothrops atrox] but subsequently the affected snakes were identified as lancehead vipers [Bothrops moojeni]) at a snake farm in Switzerland (Foelsch and Leloup, 1976). Although Pseudomonas and Aeromonas were isolated from the respiratory tracts of dead snakes, a virus (Fer-de-Lance virus [FDLV]) with ultrastructural properties similar to the myxoviruses was identified (Clark and Lunger, 1981; Clark et al., 1979). That FDLV is a PMV as indicated by demonstrating it possessed a single-stranded RNA genome and had a sedimentation value of 50S. In the first reported die-off in the United States from which a PMV was recovered, 8 to 9 rock rattlesnakes (Crotalus lepidus) died with clinical signs of CNS disease (Figure 9.82) (Jacobson et al., 1980). In a sub-

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sequent report, 8% of the total viperid collection in a zoo in Louisiana died during a 2- to 3-month period; the affected genera were Bitis, Bothrops, Crotalus, Trimeresurus, and Vipera, (Jacobson et al., 1981). Isolates were obtained from a neotropical rattlesnake (Crotalus durissus) and a bush viper (Atheris squamiger). A paramyxovirus was recovered from the lungs of two dead Ottoman vipers (Vipera xanthena xanthena) from a zoological collection (Potgieter et al., 1987). Subsequently, PMV-associated die-offs were identified (either by virus isolation or characteristic histopathologic changes) in a variety of private and zoologic collections in the United States. In some collections, although multiple species of viperid snakes were maintained, only one or two species per collection were severely affected. In one collection having at least 12 crotalid species, the major species affected was the Mexican west coast rattlesnake (Crotalus basiliscus). In another die-off, a long-term collection of 30 to 40 north Pacific rattlesnakes (Crotalus viridis oreganus) died over a 1- to 2-month period. In 1987 a major die-off of Mojave rattlesnakes in a research collection was attributed to PMV infection. In 1987, an epizootic of PMV occurred in snakes following the opening of a new reptile house at a zoo in the United States (Jacobson et al., 1992). Of 42 viperid and nonviperid snakes that died, 26 had pathological changes consistent with PMV. The virus was isolated from tissues of five snakes. In 1988 a paramyxovirus was isolated from a black mamba (Dendroaspis polylepis) in a serpentarium experiencing a die-off of viperids, elapids, boids, and colubrids. In the Federal Republic of Germany, a myxovirus-like agent was recovered from a red-tailed rat snake (Elaphe oxycephala) (Ahne et al., 1987). Isolates in Germany have been obtained from snakes in the families Boidae, Elapidae, Colubridae, Viperidae, and Crotalidae (Essbauer and Ahne, 2001; Franke et al., 2001). In the Canary Islands, immunohistochemical labeling with a polyclonal antibody identified paramyxovirus antigen in tissues of the following snakes, which died in several different collections: western diamondback rattlesnake (Crotalus atrox), rhinoceros viper (Bitis nasicornis), king snake (Lampropeltis getulus), black rat snake (Elaphe obsoleta), ball python and Burmese python (Python molurus bivittatus) (Oros et al., 2001). A paramyxovirus was isolated from the sputum of a captive Neotropical rattlesnake in a serpentarium in Brazil that had clinical signs of pneumonia; PCR and nested-PCR were used for its identification (Nogueira et al., 2002). An epizootic of PMV was reported in a colony of vipers in Brazil that are known as urutu (Bothrops alternatus) (Kolesnikovas et al., 2006). Proliferative pneumonia was seen in three of four snakes that were euthanized, and a virus consistent with PMV was isolated from pooled tissues of two snakes, and from lung, liver, and pancreas of a third snake. It is the respiratory tract that appears to be targeted by PMV in snakes. Gross changes range from diffuse hemorrhage to multifocal to diffuse accumulation of caseous

necrotic debris within the lumen of the air passageways of the lung and air sac system (Figure 9.83). The lung tissue is often thickened and edematous. Free blood may be found in the coelomic cavity. Terminally, many snakes may expel blood or caseopurulent material from the glottis, which may fill the nasal passageways, cover the oral mucosa (Figure 9.84), and may be partially swallowed into the upper esophagus. Blood may be seen on cage walls (Figure 9.85). Although no consistent changes are seen in other organs, in rattlesnakes an enlarged pancreas may be seen. Histologic examination of lung tissue from snakes that die of natural (Jacobson et al., 1992) or experimentally induced (Jacobson et al., 1997) PMV infection generally reveal a moderate to profuse amount of cellular debris and exudate filling both major and minor air passageways (Figure 9.86). In early stages of infection in Aruba Island rattlesnakes (Crotalus unicolor) the proliferation of lining epithelium was minimal (Jacobson et al., 1997) (Figure 9.87). Gram-negative microorganisms are often seen within this material. As the infection progresses, alveolar cells lining the primitive alveoli undergo hypertrophy, hyperplasia, and metaplasia. Characteristically, hyperplastic alveolar cells completely proliferate over capillary beds that are normally at the surface of the primitive alveoli (Figures 9.88–9.89). Intracytoplasmic inclusions are uncommon within alveolar cells. The pulmonary septae are often thickened with edema fluid. While in some cases there are relatively few inflammatory cells in the interstitium, in others there are infiltrates of mixed inflammatory cells (Figure 9.90). In some snakes, pancreatitis and pancreatic necrosis may also be seen (Figure 9.91) in addition to the pulmonary changes. In some cases, pancreatic necrosis may be the primary change. Occasionally a snake suffering from PMV infection may exhibit signs of CNS disease (Figure 9.82). Just prior to death, when snakes are in an agonal state, they may twist around and appear to have neurological disease. This does not represent true neurological disease. Still, some cases of snakes with neurological disease have been seen. In a rock rattlesnake, glial nodules, perivascular cuffing (Figure 9.92), demyelination, and some degeneration of axon fibers with moderate ballooning of axon sheaths were seen in the brain, brainstem, and upper spinal cord (Jacobson et al., 1980). An adult male Boelen’s python (Morelia boeleni) presented with acute neurologic disease was found on histologic examination to have a nonsuppurative meningoencephalitis (West et al., 2001). Occasional eosinophilic intracytoplasmic inclusions were noted in glial cells. Electron microscopy revealed that the inclusions contained stacks of filaments 13 to 14 nm wide. With the use of a generic paramyxovirus cDNA probe, an in situ hybridization technique demonstrated positive staining in sections of brain and esophageal ganglion. Paramyxovirus was the likely cause of the encephalomyelitis in this python, and this virus should be included in the differential diagnosis of pythons exhibiting CNS disease.

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In the United States, Boyer et al. (2000) reported a neurological disease in captive diamond (Morelia spilota spilota) and carpet pythons (Morelia spilota variegata). In some of these cases, eosinophilic intranuclear inclusions (Figure 9.93) were seen in H&E-stained sections of the brain. Using TEM, the inclusions were found to be composed of filamentous structures resembling nucleocapsid strands of paramyxovirus. In a diamond python there were particles with morphologic features of a retrovirus in one area of the brain. Historically, an identifying virus in infected tissue was used to make a presumptive diagnosis of OPMV (Figure 9.94). Several cell lines have been used to isolate paramyxovirus including commercially available viper heart cells and Vero cells. Infected cells fuse, resulting in multinucleated giant cells (Figure 9.95). Once cytopathic effects are seen, the cells can be scraped from the flask, pelleted, placed in agar, and processed for TEM. The virus can be visualized enveloping from the cell membrane (Figure 9.96–9.97). Using a rabbit-derived polyclonal antibody raised against an isolate of PMV, an immunofluorescence technique was used to label a virus in cell culture (Figure 9.98) and in frozen and paraffin-embedded tissue section (Figure 9.99). Immunoperoxidase staining, using the same polyclonal antibody raised against an isolate of ophidian paramyxovirus, was used to demonstrate antigen in tissue section (Figure 9.100) (Homer et al., 1995; Oros et al., 2001). A cDNA:RNA in situ hybridization was used to localize OPMV in formalin-fixed, paraffin-embedded tissue sections (Sand et al., 2004). Reverse transcriptase-PCR assay was used to detect a 153-bp region of OPMV genome in the total RNA extracted from either paraffin-embedded tissue from suspect cases or cell cultures infected with OPMV (Sand et al., 2004). The morphologic appearance, physico- and biochemical properties, and cytopathologic effects of three snake isolates were consistent with members of the family Paramyxoviridae (Richter et al., 1996). Comparative sequence analysis of 16 reptilian paramyxoviruses resulted in the splitting of these viruses into two subgroups (a and b) and those that were intermediate (Ahne et al., 1999). The authors believed that the two subgroups represented distinct viral species containing multiple strains. The five intermediate isolates may represent additional distinct species. The results of another study, in which the partial sequence for the large (L) and hemagglutinin-neuraminidase (HN) genes for isolates from a neotropical rattlesnake and a bush viper, supported this splitting of the ophidian paramyxoviruses into two main groups, with bridging intermediate isolates (Kindermann et al., 2001). The neotropical rattlesnake virus clustered within an intermediate group and the bush viper isolate clustered within group b. Based on the sequences for the partial L gene, the division of the viruses into these two groups was supported by the maximum bootstrap value of 100%. However, using the sequences for the partial HN gene, the same division was supported by a Bootstrap value of only 64%. Still, the authors believed that splitting these viruses into two groups was valid and that members of the two groups

may represent a different viral species. Partial cDNA coding for a protein compatible with the paramyxovirus fusion (F) protein was characterized from the jararaca (Bothrops jararaca), a viper native to Brazil (Junqueira de Azevedo et al., 2001). Compared to fusion proteins of other paramyxoviruses, it had the highest sequence similarity (37 to 39%) to those of human parainfluenza 3 and Sendai virus. Based on sequences for the L and F genes from 18 and 16 snake paramyxovirus isolates, respectively (in Germany), and comparisons with those of mammal paramyxoviruses, a new genus was proposed for the snake paramyxoviruses within the subfamily Paramyxovirinae (Franke et al., 2001). The complete RNA genome sequence of FDLV was determined to be 15,378 nucleotides in length and consisted of 7 nonoverlapping genes (Kurath et al., 2004). Comparisons made with other paramyxoviruses also resulted in a recommendation that a new genus be created for these viruses. It was suggested that the new genus be named Ferlavirus, with FDLV as the type species.

9.10 Retroviridae 9.10.1 General Characteristics Members of the family Retroviridae are spherical, enveloped, positive-sense, single-stranded RNA viruses measuring from 80 to 100 nm. They have glycoprotein surface projections, contain an icosahedral capsid, and have a helical nucleocapsid. The family consists of Alpharetrovirus, Betaretrovirus, Gammaretrovirus, Deltaretrovirus, Epsilonretrovirus, Lentivirus, and Spumavirus. The nucleocapsid varies in shape among genera and is eccentric in type B virions, concentric in type C (Spumavirus) and human T-cell leukemia related viruses (HTLV)/bovine leukemia virus (BLV), and rod- or cone-shaped in Lentivirus. RNA replication is unique, starting with reverse transcription of viral RNA into cDNA, and positive-sense cDNA synthesis with cDNA integrated into host chromosomal DNA, which is used for transcription of viral RNA. Replication is intracytoplasmic with envelopment at cytoplasmic membranes. While many retroviruses are associated with diseases, there also are many endogenous retroviruses with no apparent pathogenic effects.

9.10.2 Chelonia Endogenous retroviral sequences were obtained from an isolate from a red-eared slider (Trachemys [formerly Chrysemys] scripta) (Herniou et al., 1998). No disease was reported to be associated with this partially sequenced virus. Fibropapillomas and skin from asymptomatic green turtles were positive for polymerase-enhanced reverse transcriptase and electron microscopic evaluation of sucrose gradient purification fractions of a fibroma from the heart revealed particles consistent with retrovirus (Casey et al., 1997). However, a chelonid fibropapilloma-associated herpesvirus is probably the causative agent of this disease (Herbst et al., 2004).

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9.10.3 Crocodylia A novel group of retroviruses was identified in the following crocodilians: Chinese alligator (Alligator sinensis), smoothfronted caiman (Paleosuchus palpebrosus), broad-nosed caiman (Caiman latirostris), Nile crocodile, Orinoco crocodile (Crocodylus intermedius), and gharial (Gavialis gangeticus) (Herniou et al., 1998; Martin et al., 2002). No disease was reported to be associated with this partially sequenced virus.

9.10.4 Rhynchocephalia Highly divergent endogenous retroviral sequences were obtained from a tuatara (Sphenodon punctatus) (Tristem et al., 1995). No disease was reported to be associated with this partially sequenced virus.

9.10.5 Sauria Endogenous retroviral sequences were identified in samples from a Komodo dragon (Varanus komodoensis) (Martin et al., 1997). No disease was reported to be associated with this partially sequenced virus.

9.10.6 Ophidia 9.10.6.1 Retroviruses Associated with Neoplasms  The first oncornavirus associated with reptilian neoplastic cells was a C-type virus (105 to 110 nm) that was demonstrated by electron microscopy to be budding from spleen tissue cultures in the 48th and 52nd passage from a Russell’s viper (Vipera russelli) with a precardial myxofibroma (Zeigel and Clark, 1969). The authors suggested that the virus-producing cell line originated in cells of the tumor that metastasized to the spleen’s vascular compartment. Structures resembling Atype particles were observed in paranuclear inclusions in this C-type particle-producing viper cell line (Lunger et al., 1974). Electron microscopic examination of an embryonal rhabdomyosarcoma from a corn snake revealed particles resembling C-type viruses (Lunger et al., 1974). This represented the first report describing the morphogenesis of C-type virus particles within the primary tumor tissue of a poikilothermic. Using electron microscopy, retroviral particles were found to be associated with leukemia in two boa constrictors (Ippen et al., 1978) and a yellow rat snake (Elaphe obsoleta quadrivittata) (Zschiesche et al., 1988). A-type particles were described in a metastatic intestinal adenocarcinoma in an emerald tree boa (Corallus caninus) (Oros et al., 2004). No transmission studies were performed with these viruses to determine their oncogenic potential.  Renal cell carcinomas (Figure 9.101) were reported in a colony of lancehead vipers in Brazil (Hoge, 1997; Hoge et al., 1995). Using TEM, retroviral particles (Figures 6.96–6.99) were seen in these tumors and tumor cells that were grown

in flasks. Tumors from four Burmese pythons (undifferentiated malignant round cell tumor [Figures 9.102–9.103], colonic adenocarcinoma, transitional cell carcinoma of the kidney, intermandibular carcinoma) that were evaluated by TEM and found to be infected with extracellular and intracellular C-type-like retroviral particles (Chandra et al., 2001) (Figures 6.95, 9.104). The relationship of these particles with the tumors was not determined.

9.10.6.2 Retroviruses in Clinically Healthy Snakes  Retrovirus-like particles were identified in the venom gland of the jararacussu (Bothrops jararacussu), a viper from Brazil (Carneiro et al., 1992). Sequence data is available for an endogenous retrovirus of the jararaca (Martin et al., 1997; Herniou et al., 1998), another viper from Brazil. Van Regenmortel et al. (2000) categorized a viper retrovirus as a member of the genus Gammaretrovirus.

9.10.6.3 Retrovirus-like Particles Associated with Red Blood Cell Inclusions in a Water Snake  In a report describing intracytoplasmic inclusions in red blood cells of a yellow-bellied water snake (Nerodia erythrogaster flavigaster) that were similar to those seen with LEV and SEV (iridoviruses), inclusions in this snake consisted of particles that were similar in appearance to those of A-type and C-type retroviruses (Daly et al., 1980).

9.10.6.4 Inclusion Body Disease of Boid Snakes  Starting in the 1970s in the United States, a disease named inclusion body disease (IBD) was first appreciated in private and zoologic collections of boid snakes around the world, including the boa constrictor, green anaconda (Eunectes murinus), Haitian boa (Epicrates striatus), Burmese python, Indian python (P. m. molurus), reticulated python (P. reticulatus), and ball python (Schumacher et al., 1994b). In 1998, IBD was reported in Australia in captive native carpet and diamond pythons (Carlisle-Nowak et al., 1998) and in captive boa constrictors in the Canary Islands, Spain (Oros et al., 1998b), and three cases in Belgium (Vancraeynest et al., 2006). In addition, a disease resembling IBD was diagnosed in an eastern king snake (Lampropeltis getulus) that was housed with boa constrictors (Jacobson et al., 2001b), and in palm vipers (Botriechis marchi) (Raymond et al., 2001).   Starting in the late 1970s and extending into the mid1980s, Burmese pythons were the most common boid snake seen with IBD, with head tilts, disequilibrium, and opisthotonos as the most common clinical signs. Starting in the early 1990s, more cases have been reported in boa constrictors as compared to Burmese and other pythons. In boa constrictors, in addition to signs of CNS (central nervous system) disease (Figures 9.105–9.107), affected snakes also regurgitate food items within several days of feeding. While some snakes die within several weeks of first manifesting illness, others may survive for months. Other signs seen in affected snakes are stomatitis, pneumonia and lymphoproliferative disorders, and

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round cell tumors (Figure 9.108). In Burmese pythons the disease is more chronic, with primary signs of CNS disease; several pythons with flaccid paralysis of their entire bodies have been seen. Regurgitation is not seen in Burmese pythons. Many affected snakes may have a subclinical infection. Inclusions have been seen in tissues of boa constrictors considered to be clinically healthy. We do not know what percentage of snakes that have IBD will develop clinical signs of disease and how many will remain clinically healthy. It is possible that latent infections can persist for long periods of time. We also do not know if there are different pathogenic strains of the virus that may account for differences in the clinical appearance of infected snakes. A postmortem diagnosis of IBD is based upon the presence of eosinophilic to amphophilic intracytoplasmic inclusions in H&E-stained tissue sections. The tinctorial characteristics of the inclusions may vary with the type of hematoxylin used and differences in staining methods. In pythons, inclusions are found mostly within neurons in the CNS (Figures 9.109– 9.110) where they may stain eosinophilic or amphophilic. In boa constrictors, inclusions are also commonly seen in neurons and glial cells in the CNS (Figures 9.111–9.112), with (Figures 9.113–115) or without (Figure 9.116) encephalitis. The encephalitis is generally more severe in pythons compared to boa constrictors. In boa constrictors, inclusions also are commonly seen in: (1) mucosal epithelial cells adjacent to and overlying the esophageal tonsils (Figures 9.117–9.118), (2) lymphoid cells in esophageal tonsils (Figure 9.119), (3) epithelial cells lining the gastrointestinal tract, (4) epithelial cells in cells lining the respiratory tract (Figure 9.120), (5) hepatocytes (Figures 9.121–9.122), (6) pancreas (Figure 9.123–9.124); and (7) renal tubular epithelial cells (Figures 9.125–9.126). An antemortem diagnosis can be attempted by demonstrating inclusions in histologically processed and H&E-stained biopsy specimens. Boid snakes have well-developed esophageal tonsils (Jacobson and Collins, 1980) (Figure 9.127), and in snakes with IBD they may enlarge or abscess (Figure 9.128). Using a flexible endoscope with a biopsy device, esophageal tonsils are easily biopsied, fixed, and routinely processed for H&E staining (Figure 9.129). Inclusions are commonly seen in overlying mucosal epithelial cells (Figures 9.130). Liver and kidney biopsy specimens can also be obtained for histologic evaluation. For a more rapid diagnosis, cytologic impression smears can be made and stained with H&E (Figure 9.131) and Wright-Giemsa (Figure 9.132) stained cytological preparations of these tissues (Garner and Raymond, 2004; Jacobson, 2002). Inclusions are easier to identify in H&E-stained preparations. Inclusions are occasionally seen in lymphocytes in peripheral blood films of snakes with IBD (Figure 9.133). At a light microscopic level, the inclusions seen in IBD can overlap with inclusions having a different electron microscopic appearance (Fleming et al., 2003). Using electron

microscopy, the inclusions consist of radio-dense round particles that accumulate at the margins of the inclusion (Figures 6.82–6.87). It appears that they are nonviral in nature and consist of protein that is being deposited by surrounding polyribosomes. Recent studies have documented that the inclusions in IBD consist of a unique 68-KDa protein (Wozniak et al., 2000b). Monoclonal antibodies produced against this protein band specifically labeled inclusion bodies, demonstrating that IBD inclusions represent an intracytoplasmic accumulation of an IBDV-associated protein. In an attempt to better understand the nature of these inclusions, samples were semipurified from the liver of a boa constrictor with IBD; liver from a boa constrictor lacking inclusions was used as a control. A 68-KDa protein was identified, transferred to a membrane, and was submitted to the University of Florida’s Protein Sequencing Core Laboratory for determining its amino acid sequences. If snakes having IBD produce antibodies against this protein, and snakes free of inclusions do not, then it may be possible to develop a serologic test that is an indicator of the presence of the 68-KDa protein. While there is evidence to support a retrovirus as the causative agent of IBD (Schumacher et al., 1994b), the original isolate was subsequently lost. In a more recent study, several retroviruses were isolated from boa constrictors with IBD and were partially characterized (Jacobson et al., 2001b). Western blot analysis of viral proteins indicated that these isolates were similar. A polyclonal antibody was produced in rabbits against one of the isolates, and using electron microscopy, immunogold labeling was performed to demonstrate that the antibody recognizes the virus (Figures 6.8, 6.88). Still, we do not know if any of the isolates is the causative agent of IBD. Transmission studies are needed to determine a causal relationship, and in order to do so, serologic and molecular tests are needed to ensure that challenged snakes have not been exposed to this virus. Because of this, an enzyme-linked immunosorbent assay was developed (Lock et al., 2003) to screen boa constrictors for exposure to specific antigens. A novel endogenous retrovirus was partially sequenced from healthy blood from pythons (Python curtus), healthy appearing Burmese pythons, and Burmese pythons with signs of IBD (Huder et al., 2002). Sequences for this virus could not be demonstrated in royal (ball) pythons, reticulated pythons, carpet pythons, anacondas, or boa constrictors. Samples of kidney from boa constrictors that were positive for IBD were also negative. The pathogenic significance of this virus in Burmese pythons and blood pythons is unknown. Reoviruses and adenoviruses were isolated from boa constrictors with IBD (Marschang et al., 2001b). It is unknown whether these are incidental or possibly contribute to IBD. While the exact route of transmission has not been elucidated, direct contact is probably involved. The snake mite, Ophionyssus natricis, has been incriminated as a possible vector of the causative agent. Invariably, mites are present in snake colonies experiencing an outbreak. Venereal transmission and transmission through the placenta in viviparous

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snakes and through the egg of oviparous snakes may be possible.

9.10.6.5 Neurological Disease of Australian Pythons  An adult diamond python (Morelia spilota spilota), two carpet pythons (Morelia spilota variegata), and a Boelen’s python (Morelia boeleni) from a collection that included several species of Australian and Indonesian pythons developed signs of a neurological disease and were subsequently euthanized (Boyer et al., 2000). Using light microscopy, all snakes had intranuclear inclusions in glial cells and gliosis. Using electron microscopy, intranuclear inclusions in a carpet python were composed of filamentous material that was similar to PMV nucleocapsid strands. In a diamond python, in one area of the brain, there were 90 to 100 nm particles that were consistent with members of the family Retroviridae. The relationship of this virus with the neurological disease in these pythons remains unknown.

9.11 Reoviridae 9.11.1 General Characteristics The family Reoviridae consists of nonenveloped, spherical, linear, double-stranded RNA viruses measuring 60 to 80 nm in diameter. They have two or three shells and 10 to 12 structural proteins. There are nine genera (three of which are in plants) within the family, with each distinguished by its morphology and physicochemical properties. Differences in the outer shell exist among the nine genera. All have an inner shell with icosahedral symmetry. Replication takes place in the cytoplasm and inclusions may or may not be seen.

9.11.2 Chelonia A reovirus was isolated from the tongue, esophagus, lung, and kidney of an ill Greek tortoise (Marschang, 2000; Marschang et al., 1998). No associated pathology was reported.

9.11.3 Sauria Using electron microscopy, papillomas in a European green lizard contained particles that were morphologically similar to herpesvirus, papovavirus, and reovirus (Raynaud and Adrian, 1976). Two reoviruses isolated from a green iguana that died shortly after acquisition were compared with avian and mammalian reoviruses with respect to their serological and physicochemical properties; some similarities and differences were found (Blahak, 1994; Blahak et al., 1995). A study of iguanid lizards on Honduran islands found that while five of 14 Iguana iguana rhinolopha, 17 of 31 Ctenosaura bakeri, and one of four C. similis were seropositive for exposure to reovirus, the virus was not isolated from any of these lizards (Gravendyck et al., 1998). In a similar study, while antibodies to reovirus were found in 3 of 13 Xenosaurus grandis col-

lected in Mexico and transported to a university in the United States, no virus was isolated from these lizards (Marschang et al., 2002b). Using negative staining electron microscopy, a reovirus was identified in tissues of Uromastyx hardwickii that had been recently imported into the United Kingdom (Drury et al., 2002b). The virus was isolated and RNA was compared with that of avian reovirus. While both had 10 segments, migration patterns of the segments showed differences.

9.11.4 Ophidia A reo-like virus was isolated from 2 of 4 Chinese vipers (Azemiops feyi) that died and were found to have enteritis (Jacobson, 1986). A reovirus was isolated from a moribund ball python (Ahne et al., 1987), an emerald tree boa (Blahak and Gobel, 1991), an Aesculapian snake (Blahak et al., 1995), and from the brain of a prairie rattlesnake (Crotalus viridis) that was representative of 14 other prairie rattlesnakes showing signs of CNS disease including incoordination, loss of proprioception, and convulsions (Vieler et al., 1994). A reovirus was isolated from a juvenile Moellendorff rat snake (Elaphe moellendorffi) and a beauty snake (Elaphe taenuris) that died during an outbreak of respiratory disease in recently imported snakes. Histological examination of tissues from the Moellendorff rat snake revealed a mild proliferative pneumonia (Figure 9.134), with infiltrates of lymphocytes adjacent to and surrounding the apical faveolar muscle bundles (Figure 9.135). The isolate was found to consist of double-stranded RNA and measured 75 to 85 nm in diameter (Figure 6.77). The size and morphology was consistent with a reovirus (Lamirande et al., 1999). The virus was inoculated intratracheally into a juvenile black rat snake that subsequently died with interstitial pneumonia. The virus was re-isolated and inoculated into a second black rat snake that was killed at 40 days post-inoculation and was found to have tracheitis and pneumonia. This was the first experimental transmission of a reovirus in a reptile. Mojave rattlesnakes that died with a mild proliferative pneumonia (Figure 9.136) were infected with a reovirus that was subsequently isolated in Vero cells (Figure 9.137). Degenerate PCR primers were designed targeting conserved regions of the genome, and sequenced products indicated a novel reovirus that is probably a member of the Orthoreovirus (Wellehan et al., 2004c; Wellehan et al., 2005b).

9.12 Togaviridae 9.12.1 General Characteristics Members of the family Togaviridae are spherical, enveloped, linear, single-stranded RNA viruses that measure approximately 70 nm in diameter. The core exhibits icosahedral symmetry. Replication is within the cytoplasm, with envelopment through the cytoplasmic membrane. Inclusions are not seen. There are two genera within the family: Alphavirus and Rubivirus. Alphaviruses are transmitted by mosquitoes or other

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hematophagous arthropods. While alphaviruses are known to infect reptiles, disease has not been reported.

9.12.2 Chelonia Evidence of infections with eastern equine encephalitis (EEE) virus and western equine encephalitis (WEE) virus, either by isolation or serology, have been reported in a variety of chelonians, including the eastern box turtle, musk turtle (Sternotherus carinatus), diamondback terrapin (Malaclemys terrapin), painted turtle, gopher tortoise, Texas tortoise (Gopherus berlandieri), common snapping turtle (Chelydra serpentina), and soft-shelled turtle (Bowen, 1977; Dalrymple et al., 1972; Karstad, 1961; Shortridge and Oya, 1984).

9.12.3 Crocodylia There is evidence of an antibody to EEE virus in the blood of wild American alligators (Alligator mississippiensis) (Karstad, 1961).

9.12.4 Sauria Evidence of infections with EEE virus and WEE virus, either by isolation or serology, has been reported in a variety of lizards, including members of the families Lacertidae, Teiidae, Iguanidae, Agamidae, and Gekkonidae (Shortridge and Oya, 1984; Walder et al., 1984).

9.12.5 Ophidia Evidence of infections with EEE virus and WEE virus, either by isolation or serology, has been reported in a variety of snakes, including members of the families Colubridae, Elapidae, and Crotalidae (Gebhardt et al., 1973; Shortridge and Oya, 1984; Thomas and Eklund, 1962). Transmission studies have been performed in garter snakes to determine whether the virus can overwinter in snakes, can result in antibody production, and be infectious to mosquitoes (Milby and Reeves, 1990; Thomas and Eklund, 1960, 1962; Thomas et al., 1958; Thomas et al., 1980). Environmental (body) temperature affects viremia, with no viremia detected in experimentally infected snakes during torpor, and a lag time of several days required to detect the virus after an animal emerges and is warmed. In some experimental studies, a persistent viremia was seen. Still, the virus is infrequently isolated from the blood of wild animals.

9.13 Flaviviridae 9.13.1 General Characteristics Members of the family Flaviviridae are spherical, enveloped, linear, positive-sense, single-stranded RNA viruses that measure approximately 40 to 60 nm in diameter. The core is

spherical. Replication is within the cytoplasm; inclusions are not seen. There are three genera within the family: Flavivirus, Pestivirus, and unnamed genus for hepatitis C virus. Flaviviruses are transmitted by mosquitoes or ticks.

9.13.2 Chelonia In a survey in New York state, antibodies to St. Louis encephalitis virus (SLEV) were found in a painted turtle (Whitney et al., 1968). Antibodies to Japanese encephalitis virus (JEV) have been reported in the Chinese soft-shelled turtle (Shortridge et al., 1975). A flavivirus-like agent (based on size and morphology) was isolated from tissues of a leopard tortoise that had anemia, epistaxis, and cloacal hemorrhage and that subsequently died (Drury et al., 2001).

9.13.3 Crocodylia Antibodies to West Nile virus (WNV) were detected in Nile crocodiles in a commercial farming operation in Israel (Steinman et al., 2003) and in wild alligators in Florida (Jacobson et al., 2005a). Mortality associated with WNV was reported in farmed American alligators in Georgia (Miller et al., 2003), Florida (Jacobson et al., 2005b), and Louisiana (Nevarez et al., 2005). In Florida an epizootic of neurologic disease occurred in September and October 2002, peak months for mortality in horses from WNV infection. Three affected alligators were euthanized and necropsies were performed. The most significant microscopic lesions were a moderate heterophilic to lymphoplasmacytic meningoencephalomyelitis (Figure 9.138), necrotizing hepatitis (Figure 9.139), splenic necrosis (Figure 9.140), pancreatic necrosis (Figure 9.141), myocardial degeneration with necrosis, mild interstitial pneumonia, heterophilic necrotizing stomatitis, and glossitis. Immunohistochemistry identified the WNV antigen at highest concentrations in liver, pancreas, spleen, and brain (Figure 9.142). Virus isolation and RNA detection confirmed WNV infection in plasma and tissue samples. Of the tissues, the liver had the highest viral loads (maximum 108.9 log10 pfu/0.5cm3) while brain and spinal cord tissues had the lowest viral loads (maximum 106.6 log10 pfu/0.5cm3 each). Viremia titers in plasma ranged from 103.6 to 106.5 log10 pfu/mL, exceeding the threshold needed to infect Culex pipiens mosquitoes (105 log10 pfu/ mL). In a subsequent transmission study, 24 juvenile alligators were exposed to WNV, either parenterally or orally (Klenk et al., 2004). All became infected, and all but three sustained viremia titers > 5.0 log10 pfu/mL (a threshold considered infectious for Culex quinquefasciatus mosquitoes) for 1 to 8 days. Uninoculated tankmates also became infected. The results suggested that alligators might play an important role in WNV transmission in areas with high population densities of juvenile alligators.

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9.13.4 Sauria Reports of flavivirus infection in lizards are limited to studies with JEV virus in the grass lizard (Takydromus tachydromoides) and the Far Eastern skink (Eumeces latiscutatus) (Doi et al., 1968, 1983), and a report of WNV infection in a crocodile monitor (Varanus salavtor) (Travis et al., 2003). Ten of 44 green iguanas experimentally infected with WNV showed a low, transient viremia; one harbored detectable virus in organs (Klenk and Komar, 2003).

9.13.5 Ophidia Evidence of infection of snakes with members of the family Flaviviridae include the isolation of JEV from Chinese rat snakes (Elaphe rufodorsata) (Lee et al., 1972) and the presence of antibodies to JEV in cobras (Naja naja) in Hong Kong (Shortridge et al., 1974) and to SLEV in a rattlesnake (Crotalus viridis) in California (Milby and Reeves, 1990). Of 19 garter snakes experimentally infected with WNV, none developed a viremia, but the virus was detected in one or more organs of three snakes (Klenk and Komar, 2003).

9.14 Rhabdoviridae 9.14.1 General Characteristics Members of the family Rhabdoviridae are enveloped, bacilliform- (plant) or bullet-shaped (animal) viruses. They are single-stranded, negative-sense RNA viruses that measure from 130 to 380 nm in length. The nucleocapsid is helically coiled and cylindrical in shape. There are generally 5 viral proteins. Replication for animal rhabdoviruses generally takes place in the cytoplasm, with budding at the plasma or intracytoplasmic membranes. There are 6 genera, two of which are plant viruses. While several rhabdoviruses have been reported to infect reptiles, no disease has been associated with their presence.

9.14.2 Chelonia Antibodies to vesicular stomatitis virus, a member of the family Rhabdoviridae, were reported in spiny soft-shelled turtles (Apalone [formerly Trionyx] spinifer); no associated disease was reported (Cook et al., 1965).

9.14.3 Sauria Three antigenically distinct viruses, designated Marco, Chaco, and Tinbo viruses, were isolated from the giant ameiva (Ameiva ameiva) and Spix’s kentropyx (Kentropyx calcaratus) from Brazil (Causey et al., 1966). When first isolated they were classified as arboviruses. Based on ultrastructural morphology, Marco, Chaco, and Tinbo viruses were subsequently categorized as rhabdoviruses (Monath et al., 1979). They were serologically distinct from 34 insect and mammal rhabdovi-

ruses. It is unknown whether these viruses can cause disease in their hosts. Intracytoplasmic inclusions seen in erythrocytes of a peripheral blood film of a caiman lizard were found (using TEM) to be composed of viral particles (Figure 9.143) consistent morphologically with a rhabdovirus (Allan Pessier, personal communication).

9.14.4 Ophidia Antibodies to vesicular stomatitis virus were reported in redbellied water snakes (Nerodia erythrogaster); no associated disease was noted (Hoff and Trainer, 1973).

9.15 Caliciviridae 9.15.1 General Characteristics Members of the family Caliciviridae are small (30 to 38 nm in diameter), linear, single-stranded, positive-sense, nonenveloped RNA viruses. Replication and assembly take place in the cytoplasm. Members of this family have been categorized into four genera. Infection of reptiles with calicivirus is limited to one report of isolates obtained from snakes.

9.15.2 Ophidia Multiple caliciviruses were isolated from the following snakes in a zoological collection: Aruba Island rattlesnake, rock rattlesnake, and eyelash viper (Bothrops schlegeli) (Smith et al., 1986). The isolates were indistinguishable and were grouped into a single serotype, Crotalus calicivirus type 1 (Cro-1), within the genus Vesivirus (Matson et al., 1996). They appear to be nonpathogenic in their snake hosts. They were serologically indistinguishable from 10 calicivirus isolates obtained from three species of marine mammals (Barlough et al., 1998).

9.16 Picornaviridae 9.16.1 General Characteristics Members of the family Picornaviridae are small (28 to 30 nm in diameter) linear, single-stranded, positive-sense, nonenveloped RNA viruses. Replication and assembly take place in the cytoplasm. Members of this family have been categorized into five genera. There is one report in snakes of infection with a virus resembling picornavirus.

9.16.2 Ophidia Gastrointestinal disease in a four-lined rat snake, a boa constrictor, an Aesculapian snake, and a gaboon viper were associated with a variety of viruses including adeno-, parvo-, picorna- and herpesvirus-like viruses (Heldstab and Bestetti,

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1984). Clinical signs prior to death were a necrotizing stomatitis and suspected intestinal disturbances in the rat snake, signs of CNS disease in the boa constrictor, and loss of appetite and yellow gelatinous feces in the Aesculapian snake; the gaboon viper did not exhibit any signs of illness preceding death. Isolation attempts were not reported and a causal relationship was not established.

client-owned animals submitted to the Zoological Medicine Service, College of Veterinary Medicine, University of Florida, Gainesville. Many cases resulted in research projects designed to better understand the pathogenesis of specific diseases.

References

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Figure 9.1  Green turtle, Chelonia mydas. Cheloniidae. Gray patch disease. Coalescing areas of erosive and proliferative cutaneous lesions are scattered over the palpebrae, head, and cervical regions. A herpesvirus is the causative agent. (From Jacobson ER, 1981. Compendium on Continuing Education for the Practicing Veterinarian 3:195–200. With permission.)

Figure 9.2  Green turtle, Chelonia mydas. Cheloniidae. Gray patch disease. Photomicrograph showing a proliferation and hypertrophy of epidermal cells, with numerous amphophilic intranuclear inclusions (arrows). The surface is covered with keratin (K). H&E stain.

Figure 9.3  Green turtle, Chelonia mydas. Cheloniidae. LET (lung, eye, and tracheal) disease. Caseous material is seen adjacent to the glottal opening.

Figure 9.4  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Caseous material is seen over the globe.

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Figure 9.5  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Caseous material is seen within the trachea (arrows).

Figure 9.6  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Photomicrograph showing a diffuse area of necrosis of the mucosa and submucosa, directly anterior to the glottis. H&E stain.

Figure 9.7  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Photomicrograph showing necrosis of the tracheal mucosa with an accumulation of necrotic debris in the lumen. H&E stain.

Figure 9.8  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Photomicrograph showing amphophilic intranuclear inclusions in tracheal epithelial cells. H&E stain.

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Figure 9.9  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Photomicrograph of the lung showing a proliferative pneumonia. Inflammatory cells and necrotic debris are in air passageways. H&E stain. (From Jacobson ER et al., 1986. J Amer Vet Med Assoc 189:1020– 1023. With permission.)

Figure 9.10  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Photomicrograph of lung showing eosinophilic intranuclear inclusions (arrows) in epithelial cells lining an air passageway. H&E stain.

Figure 9.11  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Photomicrograph of a multinucleated giant cell. This is from a flask of cultured green turtle kidney cells that were inoculated with lung and trachea of green turtles with LET disease. Wright-Giemsa stain.

Figure 9.12  Green turtle, Chelonia mydas. Cheloniidae. LET disease. Transmission electron photomicrograph of the nucleus of a green turtle kidney cell infected with a herpesvirus (LETV). Unenveloped viral particles (arrows) are seen in the nucleus. Uranyl acetate and lead citrate stain.

Viruses and Viral Diseases of Reptiles  425

Figure 9.13  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Fibropapillomas are seen in the lateral canthus (arrow) of the eye.

Figure 9.14  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. A large multilobulated fibroma is growing from the conjunctiva and is covering the globe of this sea turtle.

Figure 9.15  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Multilobulated fibropapillomas are seen growing from the soft tissue at the base of a foreflipper.

Figure 9.16    Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Numerous cutaneous fibropapillomas are seen.

426  Viruses and Viral Diseases of Reptiles

Figure 9.17­  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Photomicrograph showing an arborizing fibropapilloma. H&E stain. (From Jacobson ER et al., 1989. J Comp Path 101:39–52. With permission.)

Figure 9.18  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Fibromas (arrows) are seen within the kidneys. (From Norton TM et al., 1990. J Wildl Dis 26:265–270. With permission.)

Figure 9.19  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Most of the renal tissue is displaced by fibromas (arrows).

Viruses and Viral Diseases of Reptiles  427

Figure 9.20  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Multiple fibromas are seen within the left lobe of the liver.

Figure 9.21  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Multiple fibromas are seen within the lungs.

Figure 9.22  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Photomicrograph of the liver showing discrete fibromas. H&E stain.

Figure 9.23  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Photomicrograph of the skin showing ballooning changes in epidermal cells. H&E stain. (From Jacobson ER et al., 1991. Dis Aquat Org 12:1–6. With permission.)

428  Viruses and Viral Diseases of Reptiles

Figure 9.24  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Photomicrograph of the skin showing eosinophilic intranuclear inclusions (arrows) in epidermal cells. H&E stain. (From Jacobson ER et al. 1991. Dis Aquat Org 12:1–6. With permission.)

Figure 9.25  Green turtle, Chelonia mydas. Cheloniidae. Marine turtle fibropapilloma. Transmission electron photomicrograph of the epidermis of a fibropapilloma having intranuclear inclusions. Mature enveloped herpesviral particles are seen. Uranyl acetate and lead citrate.

Figure 9.26  False map turtle, Graptemys pseudogeographica. Emydidae. Herpesvirus infection. Eosinophilic to amphophilic intranuclear inclusions (arrows) are seen filling hepatocytic nuclei. H&E stain.

Figure 9.27  False map turtle, Graptemys pseudogeographica. Emydidae. Herpesvirus infection. Eosinophilic to amphophilic inclusions (arrows) are seen within hepatocytic nuclei. Two inclusions are surrounded by a halo. H&E stain.

Viruses and Viral Diseases of Reptiles  429

Figure 9.28  Desert tortoise, Gopherus agassizii. Testudinidae. Herpesvirus infection. The oral cavity is diffusely necrotic. (From Johnson AJ et al. 2005. J Vet Micro 111:107–116. With permission.)

Figure 9.29  Desert tortoise, Gopherus agassizii. Testudinidae. Herpesvirus infection. The lingual epithelium is hyperplastic, with epithelial cells containing amphophilic intranuclear inclusions (arrows). H&E stain.

430  Viruses and Viral Diseases of Reptiles

Figure 9.30  Argentine tortoise, Geochelone chilensis. Testudinidae. Herpesvirus infection. Necrotic debris is seen on the tongue and palatine mucosa.

Figure 9.31  Argentine tortoise, Geochelone chilensis. Testudinidae. Herpesvirus infection. Photomicrograph of the palatine mucosa showing loss of the mucosal epithelial and replacement with inflammatory cells and cellular debris. H&E stain.

Figure 9.32  Argentine tortoise, Geochelone chilensis. Testudinidae. Herpesvirus infection. Photomicrograph showing intranuclear inclusions (arrows) in exfoliated epithelial cells. H&E stain. (From Jacobson ER et al. 1985b. J Amer Vet Med Assoc 187:1227–1228. With permission.)

Figure 9.33  Argentine tortoise, Geochelone chilensis. Testudinidae. Herpesvirus infection. Transmission electron photomicrograph showing mature enveloped herpesviral particles in the cytoplasm of a degenerating mucosal epithelial cell. Uranyl acetate and lead citrate.

Viruses and Viral Diseases of Reptiles  431

Figure 9.34  Hermann’s tortoise, Testudo hermanni. Testudinidae. Herpesvirus infection. Caseous material (C) is seen covering the caudal portion of the tongue (T).

Figure 9.35  Hermann’s tortoise, Testudo hermanni. Testudinidae. Herpesvirus infection. Photomicrograph of the tongue showing eosinophilic intranuclear inclusions (arrows) within epithelial cells. H&E stain.

Figure 9.36  Greek tortoise, Testudo graeca. Testudinidae. Experimental herpesvirus infection. Oral plaques (arrows) are seen on the mucosal surface at 15 days post challenge with a herpesvirus isolated from a Greek tortoise.

432  Viruses and Viral Diseases of Reptiles

Figure 9.37  European green lizard, Lacerta viridis. Lacertidae. Cutaneous papillomatosis. Multiple pigmented papillomas are seen on the body surface.

Figure 9.38  European green lizard, Lacerta viridis. Lacertidae. Cutaneous papillomatosis. Photomicrograph of a papilloma showing arborizing hyperplastic and hypertrophic epidermal cells supported by stromal connective tissue. Numerous melanophores are seen below the basement membrane. H&E stain. Provided by the National Cancer Institute’s Registry of Tumors in Lower Animals, Sterling , VA under contract N02-CB-27034.

Figure 9.39  San Esteban chuckwalla, Sauromalus varius. Iguanidae. Herpesvirus infection. Photomicrograph of the liver. Eosinophilic intranuclear inclusions (arrows) are seen within hepatocytes. H&E stain.

Figure 9.40  San Esteban chuckwalla, Sauromalus varius. Iguanidae. Herpesvirus infection. Transmission electron photomicrograph of a hepatocyte. Adjacent to the nucleus (N), an intracytoplasmic mature enveloped herpesvirus particle (arrow) with an electron-dense core is seen in the cytoplasm. Uranyl acetate and lead citrate. Courtesy of Carlos Reggiardo.

Viruses and Viral Diseases of Reptiles  433

Figure 9.41  Green tree monitor, Varanus prasinus. Varanidae. Herpesvirus infection. Caseous material is seen in the oral cavity. Courtesy of James Wellehan. (From Wellehan JFX et al. 2005a. Vet Micro 105:83– 92. With permission.)

Figure 9.42  Nile crocodile, Crocodylus niloticus. Crocodylidae. Adenovirus infection. Photomicrograph showing basophilic intranuclear inclusions (arrows) within hepatocytes. H&E stain.

Figure 9.43  Nile crocodile, Crocodylus niloticus. Crocodylidae. Adenovirus infection. Photomicrograph showing basophilic intranuclear inclusions (arrows) within enterocytes of the small intestine. H&E stain.

Figure 9.44  Nile crocodile, Crocodylus niloticus. Crocodylidae. Adenovirus infection. Transmission electron photomicrograph of an enterocyte containing intranuclear crystalline arrays of nonenveloped viral particles being released into the cytoplasm. Uranyl acetate and lead citrate. (From Jacobson ER et al. 1984. J Amer Vet Med Assoc 185:1421–1422. With permission.)

434  Viruses and Viral Diseases of Reptiles

Figure 9.45   Bearded dragon, Pogona vitticeps. Agamidae. Adenovirus infection. Photomicrograph of hepatocytes containing basophilic intranuclear inclusions (arrows). H&E stain.

Figure 9.46  Boa constrictor, Boa constrictor. Boidae. Adenovirus infection. Photomicrograph of hepatocytes containing basophilic intranuclear inclusions (arrows). H&E stain. (From Jacobson ER et al. 1985b. J Amer Vet Med Assoc 187:1226–1227. With permission.)

Figure 9.47  Boa constrictor, Boa constrictor. Boidae. Adenovirus infection. Transmission electron photomicrograph of a hepatocyte nucleus containing nonenveloped viral particles. Uranyl acetate and lead citrate. (From Jacobson ER et al. 1985b. J Amer Vet Med Assoc 187:1226– 1227. With permission.)

Viruses and Viral Diseases of Reptiles  435

Figure 9.48  Sierra mountain king snake, Lampropeltis zonata multicincta. Colubridae. Adenovirus infection. Photomicrograph showing basophilic intranuclear inclusions (arrows) within enterocytes. H&E stain. Courtesy of Edward Wozniak.

Figure 9.49  Arizona mountain king snake, Lampropeltis pyromelena. Colubridae. Adenovirus infection. Photomicrograph of the small intestine showing epithelia hyperplasia and inflammatory cell infiltrates into the lamina propria and submucosa. H&E stain.

Figure 9.50  Arizona mountain king snake, Lampropeltis pyromelena. Colubridae. Adenovirus infection. Photomicrograph of the intestine showing enterocytes with intranuclear inclusions (arrows). H&E stain.

436  Viruses and Viral Diseases of Reptiles

Figure 9.51  Spectacled caiman, Caiman crocodilus. Alligatoridae. Caiman pox. Multifocal to coalescing gray patches are scattered over the body surface.

Figure 9.52  Spectacled caiman, Caiman crocodilus. Alligatoridae. Caiman pox. Multifocal gray patches are seen on the tail.

Figure 9.53  Spectacled caiman, Caiman crocodilus. Alligatoridae. Caiman pox. A single focal gray ovoid lesion (arrow) is seen on the palpebra. (From Jacobson ER. 1981. Compendium on Continuing Education for the Practicing Veterinarian 3:195–200. With permission.)

Figure 9.54  Spectacled caiman, Caiman crocodilus. Alligatoridae. Caiman pox. Several plaques are seen on the palate and tongue.

Viruses and Viral Diseases of Reptiles  437

Figure 9.55  Spectacled caiman, Caiman crocodilus. Alligatoridae. Caiman pox. Photomicrograph of a cutaneous pox lesion. There is hypertrophy and hyperplasia of epidermal cells with hyperkeratinization. H&E stain.

Figure 9.56   Spectacled caiman, Caiman crocodilus. Alligatoridae. Caiman pox. Photomicrograph of a cutaneous pox lesion. Several cells, including nuclei, are hypertrophic and contain multiple intracytoplasmic inclusions. Large inclusions (IN) are seen accumulating in the overlying keratin. H&E stain. (From Jacobson ER et al. 1979. J Amer Vet Med Assoc 175:937–940. With permission.)

Figure 9.57  Nile crocodile, Crocodylus niloticus. Crocodylidae. Crocodile pox. Lesions are seen on the abdominal skin. Courtesy of Chris Foggin.

Figure 9.58  Nile crocodile, Crocodylus niloticus. Crocodylidae. Crocodile pox. Lesions are seen on the abdominal skin. Courtesy of Fritz W. Huchzermeyer.

438  Viruses and Viral Diseases of Reptiles

Figure 9.59  Johnston’s crocodile, Crocodylus johnstoni. Crocodylidae. Crocodile pox. Lesions are seen on the palmar surface of a forefoot. Courtesy of Gilbert N. Buenviaje and Phillip W. Ladds.

Figure 9.60  Nile crocodile, Crocodylus niloticus. Crocodylidae. Crocodile pox. Photomicrograph of a skin lesion showing hyperplasia and hypertrophy of epidermal cells, most of which contain intracytoplasmic inclusions (arrows). H&E stain. Courtesy of Chris Foggin.

Figure 9.61  Flap-necked chameleon, Chamaeleo dilepis. Chamaeleonidae. Poxvirus and chlamydia. A circulating monocyte has two circulating inclusions: one composed of chlamydia (C) and the other composed of poxvirus (P). The nucleus (N) also is seen. Wright-Giemsa stain.

Figure 9.62  Flap-necked chameleon, Chamaeleo dilepis. Chamaeleonidae. Poxvirus infection. Photomicrograph of the spleen. There are numerous macrophages containing intracytoplasmic inclusions (arrows). H&E stain.

Viruses and Viral Diseases of Reptiles  439

Figure 9.63  Gopher tortoise, Gopherus polyphemus. Testudinidae. Iridovirus infection. Photomicrograph of the trachea. There is necrosis of the mucosa with the surface covered by inflammatory cells and cellular debris. H&E stain.

Figure 9.64  Gopher tortoise, Gopherus polyphemus. Testudinidae. Iridovirus infection. Photomicrograph of a desquamated epithelial cell containing an intracytoplasmic inclusion (ICI). The nucleus (PK) is pyknotic. H&E stain.

Figure 9.65  Gopher tortoise, Gopherus polyphemus. Testudinidae. Iridovirus infection. Transmission electron photomicrograph of an epithelial cell containing numerous intracytoplasmic particles having hexagonal outlines. Uranyl acetate and lead citrate.

440  Viruses and Viral Diseases of Reptiles

Figure 9.66  Gopher tortoise, Gopherus polyphemus. Testudinidae. Ranavirus infection. Multifocal to diffuse areas of necrosis (arrows) are seen on the palatine mucosa.

Figure 9.67  Eastern box turtle, Terrapene carolina. Emydidae. Ranavirus infection. Clinical signs include rhinitis and palpebral edema.

Figure 968  Eastern box turtle, Terrapene carolina. Emydidae. Ranavirus infection. Clinical signs include rhinitis and palpebral edema.

Figure 9.69  Eastern box turtle, Terrapene carolina. Emydidae. Ranavirus infection. Photomicrograph of the kidney showing intracytoplasmic inclusions (arrows) in mononuclear cells (arrows). H&E stain. Courtesy of April Johnson and Allan Pessier.

Viruses and Viral Diseases of Reptiles  441

Figure 9.70  Eastern box turtle, Terrapene carolina. Emydidae. Ranavirus infection. Photomicrograph of the liver showing intracytoplasmic inclusions (arrows) in mononuclear cells. H&E stain. Courtesy of April Johnson and Allan Pessier.

Figure 9.71  Fischer’s chameleon, Bradypodion fischeri. Chamaeleonidae. Lizard Erythrocytic Virus infection. Photomicrograph of a peripheral blood film showing multiple intracytoplasmic inclusions. Wright-Giemsa stain.

Figure 9.72  Fischer’s chameleon, Bradypodion fischeri. Chamaeleonidae. Lizard Erythrocytic Virus infection. Photomicrograph of a peripheral blood film showing multiple intracytoplasmic inclusions. Wright-Giemsa stain.

442  Viruses and Viral Diseases of Reptiles

Figure 9.73  Plains garter snake, Thamnophis radix. Colubridae. Snake Erythrocytic Virus infection. Photomicrograph of a peripheral blood film showing red blood cells containing intracytoplasmic inclusions (arrows) containing a darker-staining punctate center. Wright-Giemsa stain.

Figure 9.74  Ribbon snake, Thamnophis sauritus. Colubridae. Snake Erythrocytic Virus infection. Photomicrograph of a peripheral blood film showing red blood cells containing intracytoplasmic inclusions (arrows) containing a darkerstaining punctate center. Wright-Giemsa stain. Courtesy of Nicole Strik.

Figure 9.75  Bolivian side-neck turtle, Platemys platycephala. Chelidae. Papillomavirus infection. There is a confluent white skin lesion posterior to the right eye. (From Jacobson ER et al. 1982b. J Amer Vet Med Assoc 181:1325–1328. With permission.)

Figure 9.76  Bolivian side-neck turtle, Platemys platycephala. Chelidae. Papillomavirus infection. Photomicrograph of a skin lesion showing epidermal hyperplasia and hyperkeratosis. No inclusions are present. H&E stain.

Viruses and Viral Diseases of Reptiles  443

Figure 9.77  Sierra mountain king snake, Lampropeltis zonata multicincta. Colubridae. Adenovirus and Dependovirus. Transmission electron photomicrograph of an enterocyte nucleus showing larger adenoviral particles and smaller dependoviral particles. Uranyl acetate and lead citrate. Courtesy of Edward Wozniak.

Figure 9.78  Painted turtle, Chrysemys picta. Emydidae. Circovirus infection. Emydidae. Photomicrograph of the spleen showing multiple intracytoplasmic inclusions within macrophages (arrows). H&E stain. Courtesy of Francisco Uzal.

Figure 9.79  Caiman lizard, Dracaena guianensis. Teiidae. Paramyxovirus infection. Photomicrograph of the lung showing a proliferative pneumonia and inflammatory cells in the airways. H&E stain.

Figure 9.80  Caiman lizard, Dracaena guianensis. Teiidae. Paramyxovirus infection. Photomicrograph of the lung showing a proliferative pneumonia and inflammatory cells in the airways. H&E stain.

444  Viruses and Viral Diseases of Reptiles

Figure 9.81  Caiman lizard, Dracaena guianensis. Teiidae. Paramyxovirus infection. Transmission electron photomicrograph of an alveolar cell showing intracytoplasmic nucleocapsid filaments (NC). Uranyl acetate and lead citrate stain. (From Jacobson ER et al. 2001a. J Vet Diag Investig 13:143–151. With permission.)

Figure 9.82  Rock rattlesnake, Crotalus lepidus. Viperidae. Paramyxovirus infection. A rock rattlesnake unable to right itself when placed in dorsal recumbency, which is indicative of central nervous system disease. (From Jacobson ER et al. 1980. J Amer Vet Med Assoc 177:796–799. With permission.)

Figure 9.83  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Paramyxovirus infection. The lumen of the lung contains blood, and caseous material is accumulating in faveolar spaces.

Figure 9.84  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxoviral infection. Experimental infection with an isolate of paramyxovirus. Blood is seen in the oral cavity.

Viruses and Viral Diseases of Reptiles  445

Figure 9.85  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Blood expelled from the respiratory tract of an experimentally infected snake is seen in the cage.

Figure 9.86  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Photomicrograph of the lung of an experimentally infected snake (day 8 following challenge) showing inflammatory cells in air passageways and a thickening of the septal walls. There is minimal hyperplasia and hypertrophy of the epithelial cells lining the airways. H&E stain.

Figure 9.87  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. A higher magnification photomicrograph of the lung of an experimentally infected snake (day 8 following challenge) showing inflammatory cells in air passageways and minimal hyperplasia and hypertrophy of epithelial cells lining the airways. H&E stain.

Figure 9.88  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Photomicrograph of the lung of an experimentally infected snake (day 15 following challenge) showing inflammatory cells in air passageways and hypertrophy and hyperplasia of epithelial cells lining the airways. H&E stain.

446  Viruses and Viral Diseases of Reptiles

Figure 9.89  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Photomicrograph of a semithin, plastic-embedded section of the lung of an experimentally infected snake (day 15 following challenge) showing inflammatory cells (IN) in airways and hypertrophy and hyperplasia of epithelial cells lining the airways. Toluidine blue stain.

Figure 9.90  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Paramyxovirus infection. Photomicrograph of the lung showing a proliferative pneumonia with infiltrates of inflammatory cells in the septal interstitium. H&E stain.

Figure 9.91  Speckled rattlesnake, Crotalus mitchelli. Viperidae. Paramyxovirus infection. Photomicrograph of the pancreas showing an area of necrosis. H&E stain.

Figure 9.92  Rock rattlesnake, Crotalus lepidus. Viperidae. Paramyxovirus infection. Photomicrograph of the brain showing perivascular cuffing. H&E stain.

Viruses and Viral Diseases of Reptiles  447

Figure 9.93  Diamond python, Morelia spilota. Pythonidae. Presumptive paramyxovirus infection. Photomicrographs showing eosinophilic intranuclear inclusions in neurons in the brain. The snake was manifesting signs of neurological disease. Transmission electron microscopy revealed that inclusions consisted of filamentous strands resembling a paramyxovirus. H&E stain.

Figure 9.94  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Paramyxovirus infection. Transmission electron micrograph of the lung of an experimentally infected snake. Virions are seen budding (arrows) from the cytoplasmic membrane of a type II cell on the surface of an airway. Uranyl acetate and lead citrate stain.

Figure 9.95  Vero cell monolayer. A syncytial cell is seen in a monolayer of Vero cells infected with paramyxovirus. Wright-Giemsa stain.

Figure 9.96  Viper heart cells. Transmission electron photomicrograph of a viper heart cell infected with paramyxovirus. Spheroidal (S) and filamentous (F) forms are seen budding from cytoplasmic extensions. Uranyl acetate and lead citrate stain. (From Jacobson ER et al. 1980. J Amer Vet Med Assoc 177:796–799. With permission.)

448  Viruses and Viral Diseases of Reptiles

Figure 9.97  Vero cells in culture. Transmission electron photomicrograph of a Vero cell infected with paramyxovirus from the lung of an experimentally infected Aruba Island rattlesnake. Filamentous and spheroidal forms are seen budding from cell membranes and accumulating in intercellular spaces. Uranyl acetate and lead citrate stain. Courtesy of Thomas Geisbert.

Figure 9.98  Viper heart cell monolayer. Fluorescent antibody staining of paramyxoviral antigen in infected viper heart cells using a labeled anti-paramyxovirus polyclonal antibody.

Figure 9.99  Aruba Island rattlesnake, Crotalus unicolor. Viperidae. Fluorescent antibody staining of a paramyxoviral antigen in a frozen section of lung of an experimentally infected snake using a labeled anti-paramyxovirus polyclonal antibody.

Figure 9.100  Berg’s adder, Bitis atropos. Viperidae. Paramyxovirus infection. Photomicrograph of the lung. Brown staining indicates the presence of paramyxovirus antigen in epithelial cells lining air passageways. Immunoperoxidase (ABC) stain and light green counterstain.

Viruses and Viral Diseases of Reptiles  449

Figure 9.101  Lance-headed viper, Bothrops moojeni. Viperidae. Photomicrograph of the kidney showing neoplastic tubules. H&E stain. Courtesy of Alma Hoge.

Figure 9.102  Burmese python, Python molurus bivittatus. Pythonidae. Undifferentiated round cell tumor. The ulcerating necrotic mass involving the left mandible was diagnosed as an undifferentiated round cell tumor.

Figure 9.103  Burmese python, Python molurus bivittatus. Pythonidae. Undifferentiated round cell tumor. Photomicrograph showing round cells in the spleen of a snake with an undifferentiated malignant round cell tumor. H&E stain.

Figure 9.104  Burmese python, Python molurus bivittatus. Pythonidae. Transmission electron photomicrograph of the spleen seen in Figure 9.103. Intracellular type C-like particles are seen within a neoplastic cell. Uranyl acetate and lead citrate stain. Courtesy of Sundeep Chandra. (From Chandra AMS et al. 2001. Vet Path 38:561–564. With permission.)

450  Viruses and Viral Diseases of Reptiles

Figure 9.105  Haitian boa, Epicrates striatus. Boidae. IBD. The abnormal posture is a sign of CNS disease.

Figure 9.106  Boa constrictor, Boa constrictor. Boidae. IBD. When placed in dorsal recumbency, this snake was unable to right itself.

Figure 9.107  Boa constrictor, Boa constrictor. Boidae. IBD. When placed in dorsal recumbency, this snake was unable to right itself.

Figure 9.108  Boa constrictor, Boa constrictor. Boidae. IBD. Biopsied skin lesions of this snake with IBD indicated a round cell tumor.

Viruses and Viral Diseases of Reptiles  451

Figure 9.109  Burmese python, Python molurus. Pythonidae. IBD. Photomicrograph of the brain showing eosinophilic intracytoplasmic inclusions (arrows). H&E stain.

Figure 9.110  Burmese python, Python molurus. Pythonidae. IBD. Photomicrograph of the brain showing numerous eosinophilic intracytoplasmic inclusions in ependymal cells. H&E stain.

Figure 9.111  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of eosinophilic intracytoplasmic inclusions in neurons of the brain. H&E stain.

Figure 9.112  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of amphophilic intracytoplasmic inclusions in neurons of the brain. H&E stain. Courtesy of Nikos Gurfield.

452  Viruses and Viral Diseases of Reptiles

Figure 9.113  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the brain showing severe inflammation. H&E stain.

Figure 9.114  Burmese python, Python molurus. Pythonidae. IBD. Photomicrograph of the brain showing perivascular cuffing. H&E stain.

Figure 9.115  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the brain showing perivascular cuffing. H&E stain. Courtesy of Brian Stacy.

Figure 9.116  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the brain. No inflammatory infiltrates are seen. H&E stain.

Viruses and Viral Diseases of Reptiles  453

Figure 9.117  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the mucosal epithelial component of an esophageal tonsil. Numerous eosinophilic intracytoplasmic inclusions are seen (arrows). H&E stain.

Figure 9.118  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of an esophageal tonsil from a necropsied snake showing a hyperplastic epithelium containing eosinophilic intracytoplasmic inclusions (arrows). H&E stain.

Figure 9.119  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of an esophageal tonsil from a necropsied snake showing numerous eosinophilic intracytoplasmic inclusions (arrows) within submucosal lymphoid cells. H&E stain.

Figure 9.120  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the lung showing eosinophilic intracytoplasmic inclusions (arrows) within epithelial cells lining an airway. H&E stain.

454  Viruses and Viral Diseases of Reptiles

Figure 9.121  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the liver showing hepatocytes containing eosinophilic intracytoplasmic inclusions. H&E stain.

Figure 9.122  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the liver showing hepatocytes containing amphophilic intracytoplasmic inclusions (arrows). H&E stain. Courtesy of Nikos Gurfield.

Figure 9.123  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the pancreas showing acinar cells containing eosinophilic intracytoplasmic inclusions. H&E stain.

Figure 9.124  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the pancreas showing acinar cells containing amphophilic intracytoplasmic inclusions (arrows). H&E stain. Courtesy of Nikos Gurfield.

Viruses and Viral Diseases of Reptiles  455

Figure 9.125  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the kidney showing renal epithelia cells containing eosinophilic intracytoplasmic inclusions. H&E stain.

Figure 9.126  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of the kidney showing renal epithelial cells containing amphophilic intracytoplasmic inclusions. H&E stain.

Figure 9.127  Reticulated python, Python reticulatus. Boidae. Multiple ellipsoidal esophageal tonsils (arrows) are seen lining the mucosal surface. (From Jacobson ER and Collins B. 1980. Develop Comp Immuno 4:703–711. With permission.)

Figure 9.128  Boa constrictor, Boa constrictor. Boidae. IBD. An abscessed esophageal tonsil (arrow) can be seen.

456  Viruses and Viral Diseases of Reptiles

Figure 9.129  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of a biopsied esophageal tonsil showing a cleft of hyperplastic epithelial cells overlying an aggregate of lymphoid cells. H&E stain.

Figure 9.130  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of a biopsied esophageal tonsil showing hyperplastic epithelial cells containing eosinophilic intracytoplasmic inclusions (arrows). H&E stain.

Figure 9.131  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of a cytologic impression of the liver. Numerous eosinophilic intracytoplasmic inclusions can be seen. H&E stain.

Viruses and Viral Diseases of Reptiles  457

Figure 9.132  Boa constrictor, Boa constrictor. Boidae. IBD. Photomicrograph of cytologic impression of the liver. Numerous basophilic intracytoplasmic inclusions (arrows) can be seen. Wright-Giemsa stain.

Figure 9.133  Boa constrictor, Boa constrictor. Boidae. IBD. Peripheral blood film with lymphocytes containing light basophilic-staining inclusions (arrows). Wright-Giemsa stain. Courtesy of Nicole Strik.

Figure 9.134  Moellendorff rat snake, Elaphe moellendorffi. Colubridae. Reovirus infection. Photomicrograph of the lung showing mild proliferative interstitial pneumonia. H&E stain.

458  Viruses and Viral Diseases of Reptiles

Figure 9.135  Moellendorff rat snake, Elaphe moellendorffi. Colubridae. Reovirus infection. Photomicrograph of the lung showing an infiltrate of small mononuclear cells into the interstitium. The overlying epithelium is hyperplastic. H&E stain.

Figure 9.136  Mojave rattlesnake, Crotalus scutulatus. Viperidae. Reovirus infection. Photomicrograph of the lung showing a mild proliferation of epithelial cells (arrows) within an airway. H&E stain. Courtesy of John Roberts.

Figure 9.137  Mojave rattlesnake, Crotalus scutulatus. Viperidae. Reovirus infection. Transmission electron photomicrograph of a Vero cell infected with a reovirus. Numerous particles are seen in the cytoplasm. Uranyl acetate and lead citrate. Bar = 200 nm. Courtesy of John Roberts.

Viruses and Viral Diseases of Reptiles  459

Figure 9.138  American alligator, Alligator mississippiensis. Alligatoridae. West Nile virus infection. Photomicrograph of the meninges of the spinal cord showing an infiltrate of heterophils. H&E stain.

Figure 9.139  American alligator, Alligator mississippiensis. Alligatoridae. West Nile virus infection. Photomicrograph of the liver showing a focal area of necrosis. H&E stain. (From Jacobson ER et al. 2005. J Wildl Dis 41:96–106. With permission.)

Figure 9.140  American alligator, Alligator mississippiensis. Alligatoridae. West Nile virus infection. Photomicrograph of the spleen showing multifocal areas of necrosis. H&E stain.

460  Viruses and Viral Diseases of Reptiles

Figure 9.141  American alligator, Alligator mississippiensis. Alligatoridae. West Nile virus infection. Photomicrograph of the pancreas showing a focal area of necrosis. H&E stain.

Figure 9.142  American alligator, Alligator mississippiensis. Alligatoridae. West Nile virus infection. Photomicrograph of the brain showing labeled antigen in neurons and leukocytes. Immunoperoxidase. (From Jacobson ER et al. 2005b. J Wildl Dis 41:96–106. With permission.)

Figure 9.143  Caiman lizard, Draecena guianensis. Teiidae. Rhabdovirus infection. Transmission electron photomicrograph of a red blood cell showing intracytoplasmic particles consistent with a rhabdovirus. Uranyl acetate and lead citrate. Courtesy of Allan Pessier.

10 Bacterial Diseases of Reptiles Elliott R. Jacobson

Contents 10.1 General Comments .............................................. 462

10.18 Chlamydia and Chlamydophila .......................... 472

10.2 Pseudomonas.......................................................462

10.18.1 Chelonia..................................................... 472

10.3 Aeromonas.......................................................... 463

10.18.2 Crocodylia.................................................. 472

10.4 Salmonella.......................................................... 463

10.18.3 Sauria......................................................... 473

10.5 Citrobacter.......................................................... 465

10.18.4 Ophidia...................................................... 473

10.6 Serratia............................................................... 465

10.19 Leptospira................................................................474

10.7 Pasteurella ......................................................... 466

10.20 Borrelia...................................................................474

10.8 Erysipelothrix...................................................... 466

10.21 Coxiella...................................................................474

10.9 Neisseria............................................................. 466

10.22 Erlichia....................................................................474

10.10 Elizabethkingia................................................... 466

10.23 Mixed and Miscellaneous Aerobic Bacterial   Infections............................................................... 475

10.11 Vibrio.................................................................. 466 10.13 Helicobacter.........................................................467

10.23.1 Spiral-Shaped Bacterium of Rhinoceros   Iguanas....................................................... 475

10.14 Streptococcus.......................................................467

10.23.2 Shell Disease of Aquatic Turtles............... 475

10.15 Anaerobes.............................................................. 467

10.23.3 Subcutaneous and Tissue Abscesses   and Masses ............................................... 475

10.12 Listeria................................................................ 466

10.16 Actinomycetales..................................................... 468 10.16.1 Mycobacterium........................................ 468 10.16.2 Nocardia................................................. 469 10.16.3 Dermatophilus........................................ 469 10.17 Mycoplasma........................................................ 470 10.17.1 Chelonia..................................................... 470 10.17.2 Crocodylia.................................................. 471 10.17.3 Sauria......................................................... 472 10.17.4 Ophidia...................................................... 472

10.23.4 Bacterial Infections of the Eye   and Orbit.................................................... 476 10.23.5 Stomatitis, Gingivitis, and Pharyngitis...... 476 10.23.6 Pneumonia................................................. 477 10.23.7 Bacteremia and Osteomyelitis.................. 478 10.23.8 Miscellaneous Bacterial Infections ......... 479 Acknowledgments............................................................ 480 References......................................................................... 480

Infectious Diseases and Pathology of Reptiles  461

462  Bacterial Diseases of Reptiles

10.1 General Comments

10.2 Pseudomonas

While a variety of bacteria have been isolated from various lesions in reptiles, in most situations they have been reported as causative agents of disease based simply upon isolation, on the body surface, within visceral structures, or in excretory products. While infections with Gram-positive bacteria do occur in reptiles, it appears that infections caused by Gram-negative bacteria are more common (Jacobson, 1987). Aeromonas, Citrobacter, Enterobacter, Escherichia, Morganella, Providencia, Pseudomonas, and Salmonella are just a few of a long list of aerobic bacteria frequently isolated from ill captive reptiles (Mayer and Frank, 1974; Jacobson, 1984a). These organisms appear to become more invasive when conditions either change the resistance of the host or select for pathogenic organisms. They may also become invasive following a primary viral disease, such as paramyxovirus pneumonia and chelonian herpesvirus stomatitis. Some groups of reptiles seem particularly prone to infection with specific types of bacteria. For instance, the American alligator (Alligator mississippiensis) is susceptible to Aeromonas hydrophila infections; Neisseria iguanae has been isolated from the oral cavity and bite wounds of the green iguana (Iguana iguana), and a chronic upper respiratory disease has been seen in the desert tortoise (Gopherus agassizii) and other tortoises, with Mycoplasma agassizii identified as a causative agent of this disease. Mycoplasma crocodyli and M. alligatoris have been identified as the cause of arthritis and pneumonia in Nile crocodiles (Crocodylus niloticus) and the American alligator, respectively. While mycobacterial diseases are not uncommon in reptiles, in most cases a diagnosis is based on identification of the organism in tissue section rather than isolation and specific identification. Infections with chlamydiae have now been identified in all major groups of reptiles. In this chapter the most important bacterial diseases of reptiles will be reviewed. Bacterial diseases will be first reviewed by pathogen. Because several different bacteria may be cultured from a lesion in a reptile, bacterial infection by certain systems will also be reviewed. For instance, aural abscesses are common in box turtles, and to a lesser degree in other turtles. While multiple bacteria have been identified in this lesion, the specific causative agent and pathogenesis have not been elucidated. The scientific names of many bacteria reviewed in this chapter have been revised since the original descriptions. It is beyond the scope of this chapter to update all the older names. For currently accepted genera of bacteria in humans, see the following online taxonomic search service (PubMed) of the National Library of Medicine and the National Institute of Health: http://www.ncbi.nlm.nih.gov/Taxonomy/taxonomyhome.html/

There are relatively few reports of pseudomonad infections in chelonians (Dieterich, 1967; Jacobson, 1978; Keymer, 1978a, b), with most reports involving lizards and snakes (Jacobson, 1984a). Pseudomonas was isolated from a Greek tortoise (Testudo graeca) with an ulcerative stomatitis and pharyngitis (Keymer, 1978a) and, without providing details, was also isolated from two ornate terrapins (Trachemys scripta callirostris [formerly Pseudemys ornata calirostris]) (Keymer, 1978b). In squamates, Pseudomonas infections may be manifested as local to diffuse integumentary lesions, oral and lingual lesions, pneumonia, and septicemias (Figures 10.1–10.6). A pseudomonad species, P. reptilovorus, was considered pathogenic for several species of lizards, with intraperitoneal inoculation in regal horned lizards (Phrynosoma solare) resulting in their death (Caldwell and Ryerson, 1940). Although Page (1961) isolated P. aeruginosa from several cases of ophidian ulcerative stomatitis, Aeromonas hydrophila was considered the most significant pathogen in this study. Different species of Pseudomonas have been commonly isolated from the oral cavity of clinically healthy snakes (Goldstein et al., 1979; Goldstein et al., 1981; Ledbetter and Kutscher, 1969; Parrish et al., 1956; Theakston et al., 1990). While healthy snakes have predominantly Gram-positive oral flora (such as Staphylococcus spp.), snakes with infectious stomatitis have predominantly Gramnegative bacteria (Draper et al., 1981). Pseudomonas aeruginosa was consistently cultured from snakes in a zoological collection that had necrotizing enteritis (Gray et al., 1966), and from the oral cavity and lungs of two boa constrictors (Boa constrictor) and an Indian python (Python molurus) having stomatitis and pneumonia (Aleksandrov and Petkov, 1985). Pseudomonas aeruginosa was considered the causative agent for the deaths of horned vipers (Cerastes cerates) and lebetine vipers (Vipera lebetina) in a vivarium (Slavtchev and Chadli, 1984). Pseudomonas aeruginosa, Aeromonas hydrophila, and Proteus spp. were cultured from a series of snakes having stomatitis, septicemia, bronchopneumonia, and abscesses (Soveri, 1984). Pseudomonas was initially thought to be the cause of a fatal infection in vipers (originally reported as fer-de-lance vipers [Bothrops atrox] but later reported as lance-headed vipers [Bothrops moojeni]) in a serpentarium in Switzerland (Foelsch and Leloup, 1976). However, further work demonstrated that a paramyxovirus was the most probable cause of this outbreak (Clark et al., 1979). Pseudomonas stomatitis may eventually result in pneumonia as necrotic debris from the oral cavity is inhaled. It is common for Pseudomonas spp. to colonize burn wounds of reptiles, such as those caused by lying on heating pads (Jacobson, 1981) (Figure 10.7). Chronic burn infections of the skin may result in Pseudomonas pneumonia. Pseudomonas was isolated from a colony of American anoles (Anolis carolinensis) in which conjunctivitis and blepharitis were seen (Millichamp et al., 1983), from periorbital abscesses of chameleons (Abou Madi and Kern, 2002; Millichamp et al., 1983; Schumacher et al.,

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1996) (Figure 10.8), and from subspectacular abscesses in snakes (Millichamp et al., 1983) (Figure 10.9).

10.3 Aeromonas Aeromonas has been identified as an important pathogen in all major groups of reptiles (Ippen and Schroder, 1977). For the most part, it is considered an opportunistic microbe. The author cultured Aeromonas from the blood of several wild common snapping turtles (Chelydra serpentina) and sliders (Trachemys spp) representative of a die-off that occurred in a population of freshwater aquatic turtles emerging following winter estivation. Aeromonas was isolated from tortoises (Keymer, 1978a) and terrapins and turtles (Keymer, 1978b) that died with evidence of septicemia. Of 144 tortoises, 11 cases (7.6%) were categorized as bacterial infections of which 9 (6.25%) were due to Aeromonas hydrophilia. In 19 of 122 necropsied terrapins and turtles, 19 (15.5%) were of bacterial origin and 7 (5.7%) were due to Aeromonas hydrophilia. One report documents a die-off associated with Aeromonas hydrophilia and A. shigelloides in turtles and American alligators (Alligator mississippiensis) in a eutrophic inland lake in Florida (Shotts et al., 1972). Aeromonas hydrophila was isolated from the anterior chamber of the eye of one of several farmed American alligators with hypopyon and iridocyclitis (Jacobson, 1984b). In a survey of 123 healthy appearing American alligators from five locations in the southeastern United States, A. hydrophila was isolated from the oral cavity and internal tissues of 85% and 70%, respectively (Gorden et al., 1979). The authors suggested that low densities of A. hydrophila in internal organs, along with high water temperatures and stress, might allow the organism to proliferate, resulting in disease. Hatchling alligators from a group kept in a holding pen died and pathologic studies revealed inflammatory cells within the lungs (Peters and Cardeilhac, 1988). Aeromonas hydrophila was the primary organism isolated from blood and tissues. In Australia, microbial examination of three farmed crocodiles with bacterial hepatitis and septicemia identified Aeromonas hydrophila as the causative agent in two cases (Buenviaje et al., 1994). Aeromonas hydrophila was also isolated from farmed Nile crocodiles in Zimbabwe (Foggin, 1987). Aeromonas is commonly associated with ulcerative stomatitis in a variety of snakes (Page, 1961, 1966). In an albino eastern diamondback rattlesnake (Crotalus adamanteus), Aeromonas was cultured from the oral mucosa, which had severe diffuse hemorrhage (Figure 10.10). As with Pseudomonas, snakes with severe (or even benign) appearing mouth lesions may aspirate necrotic debris into the respiratory system, resulting in pneumonia. The main airway of the lung (vorbronchus) and secondary air passageways may be filled with exudate. At a light microscopic level, granulomas may be seen throughout the lungs (Figure 10.11). Septicemia may follow, and may be seen as multifocal areas of ecchymotic hemorrhages and ulceration involving the integumentary system. Animals

are often anorectic, listless, and steadily decline in condition. Aspiration of necrotic debris from the oral cavity into the respiratory system may result in pneumonia. Reptiles may be presented with gaping of the mouth, labored respiration, and harsh respiratory sounds. While Aeromonas is often a secondary invader, Camin (1948) demonstrated that the snake mite Ophionyssus natricis is capable of transmitting Aeromonas hydrophila among snakes.

10.4 Salmonella Salmonella appears to be a component of the normal gastrointestinal flora of both free-ranging and captive reptiles (Hinshaw and McNeil, 1947; Jackson and Jackson, 1971; Kaura and Singh, 1968; McInnes, 1971; MacNeil and Dorward, 1986; Onderka and Finlayson, 1985). While most of the studies involve captive reptiles, a few involve free-ranging reptiles. Arizona, which is now considered a species of Salmonella, was excreted by 10.5% of 691 tortoises from southeastern and southwestern Bulgaria (Dimov, 1965). Salmonella was isolated from wild turtles in the genera Rhinoclemmys (10%) and Mauremys (16%) (Gopee et al., 2000; Greenberg and Sechter, 1992). Ten isolations of Salmonella representing six serotypes were cultured from the intestinal tract of 124 free-ranging Florida lizards (Hoff and White, 1977). In a study to determine the prevalence of Salmonella in wild reptiles submitted to a wildlife center in Virginia, of 75 reptiles representing 8 species, Salmonella was not cultured from any of the cloacal swabs collected from these animals (Richards et al., 2004). There are numerous reports of isolations of Salmonella from captive reptiles. While infection of chelonians with Salmonella is common, these animals are generally disease free. Of 127 turtles tested in a large zoological collection in the United States, 37 were positive for Salmonella (Otis and Behler, 1973). A Salmonella prevalence of 11.3% was found in 62 swabs obtained from the cloaca of several species of captive turtles or soils in their tanks from ten pet shops and four private keepers in Belgium (Pasmans et al., 2002a). Salmonella has also been isolated from the gastrointestinal tract of tortoises by several authors (Boycott et al., 1953; McNeil and Hinshaw, 1946; Milanov et al., 1966; Zwart, 1960; Zwart et al., 1970). In Germany, the feces of 125 clinically healthy tortoises were cultured and 28 different Salmonella spp. were isolated from 55 tortoises (44%) (Weber and Pietsch, 1974). Of 50 swabs of the jejunum of necropsied farm-reared Nile crocodiles in Zimbabwe, eight (16%) were positive for Salmonella; S. arizonae was the most common serotype (Obwolo and Zwart, 1993). Salmonella was also isolated from wild Nile crocodiles (Madsen et al., 1998). Salmonella was cultured from saltwater (Crocodylus porosus) and Johnston’s (freshwater [Crocodylus johnstoni]) crocodiles at two farms in the northern territories in Australia, with the prevalence varying from 5 to 30.5% and 20 to 81%, respectively (Manolis et al., 1991). In the 1990s,

464  Bacterial Diseases of Reptiles

along with the increased popularity of the green iguana as a pet, and the sale of millions of these lizards, it became clear that Salmonella was a problem that went along with the ownership of these lizards (Burnham et al., 1998). Both Salmonella Marina and S. Poona were common cloacal organisms in asymptomatic green iguanas and have been cultured from both green iguanas and infected people (Ackman et al., 1995; Boam et al., 1970a; Boam et al., 1970b). Salmonella is also common in the intestinal tract of snakes (Chiodini and Sundberg, 1981; Hinshaw and McNeil, 1944, 1946; Kennedy, 1973), with S. arizonae a common isolate (Hinshaw and McNeil, 1945; Zwart et al., 1970). Of 16 pet snakes from two breeding groups screened for Salmonella, 81% were found to harbor various serovars of S. subgroup IIIb serovars (Schröter et al., 2006). A major concern is that healthy reptiles can intermittently shed Salmonella, thereby serving as reservoirs for human infections (Barten, 1993; Fox, 1974; Tauxe et al., 1985). Some authors believe that Salmonella indigenous to reptiles are more virulent than those derived from birds and mammals (Schroder and Karasek, 1977). Salmonellosis due to a variety of Salmonella serotypes probably represents the most important zoonotic disease of captive chelonians. It was estimated that in 1970 to 1971 approximately 280,000 people in the United States acquired salmonellosis from contact with aquatic pet turtles, primarily red-eared sliders (Trachemys scripta elegans) or their maintenance water (Cohen et al., 1980; Lamm et al., 1972). Most of the cases were children under 10 years of age (Gangarosa, 1985). In 1975, the United States Food and Drug Agency placed a ban on the interstate sale of turtles with a midline carapace length of less than 4 inches. Following the ban on interstate shipment of these animals, there was a substantial decline in reptile-associated salmonellosis in humans (D’Aoust and Lior, 1978). However, with the continued popularity of reptiles as pets, reptile-associated salmonellosis persists (Morbidity and Mortality Weekly Report [MMWR], 1999). In the 1990s the green iguana surged in popularity and along with it there were numerous cases of humans becoming infected from their pet iguana. Children less than 1 year old seem to be particularly susceptible to salmonellosis and have reportedly been infected from household green iguanas (Barrett et al., 1992; Dalton et al., 1995; Mermin et al., 1997). Cases of Salmonella osteomyelitis (Andreacchio and Miller, 2001; Norwinski, 2001) and urinary tract infections (Embil and Nicolle, 1997) have been traced to pet green iguanas in households. In a zoological collection, an outbreak of salmonellosis in zoo visitors, primarily children, was traced to the contaminated wooden barrier for a Komodo monitor (Varanus komodoensis) exhibit (Friedman et al., 1997). Salmonella sepsis in two human patients was traced to a donor with a pet boa constrictor, with the isolation of the same serotype of Salmonella enterica enteriditis from the platelet products of the donor and both patients (Jafari et al., 2002). A report from Sweden showed that an import restriction

requiring certificates that reptiles are not carrying Salmonella is an effective public health measure for protecting the general population against reptile-associated Salmonella (De Jong et al., 2005). To understand the pathogenesis of Salmonella infection in turtles, S. enterica subsp. enterica serovar Muenhen was orally administered to red-eared sliders and colonization of tissues was studied (Pasmans et al., 2002b). With turtles kept at 26°C, the organism could only be isolated from the intestine for eight days following inoculation. In two of six turtles kept at 36°C, this strain was isolated from the liver and spleen, and there were increased numbers of the bacteria in the intestinal tract. No disease or signs of illness were seen. In a subsequent study, intestinal explants were used to better understand the colonization in the intestinal tract (Pasmans et al., 2003). It appears that the intestinal mucous layer provides an important site of colonization and protects the underlying tissue from invasion. Salmonella may be transmitted from animal to animal or from animal to man via urine and feces. In turtles, Salmonella has been shown to penetrate turtle eggs (Feeley and Treger, 1969) with turtles being infected at hatching (Kaufmann et al., 1972). Infection during pregnancy and transovarian passage of Salmonella has also been demonstrated for snakes (Chiodini, 1982; Schröter et al., 2006). Animals may also become populated with Salmonella via food such as slaughterhouse offal. Although in most cases reptiles infected with Salmonella appear free of overt disease and are clinically normal, still Salmonella has been cultured from lesions in reptiles (Onderka and Finlayson, 1985). Salmonella was considered to be the cause of death of two Galapagos tortoises (Geochelone nigra) at the San Diego Zoo (McNeil and Hinshaw, 1946). In a survey of acclimatized tortoises in England, only 2.8% were infected (Keymer, 1978a). In that study, Salmonella was implicated as the cause of death in two tortoises. During surveys of the Sipsey Fork in north-central Alabama, beginning in July 1985, ill flattened musk turtles (Kinosternon depressum [formerly Sternotherus depressus]) were seen (Dodd, 1988). Ill turtles basked more frequently compared to healthy turtles and had the following problems: (1) emaciation, (2) lesions on the plastron that caused the overlying scutes to peel away, and (3) discolored carapace. Several affected turtles were euthanized and submitted for pathologic evaluations. The most significant findings included multifocal areas of epidermal necrosis, areas of coagulation necrosis in the liver, edema in the submucosa of the intestinal tract, and inflammatory cells in air passageways of the lung, with edema in the interstitium. A variety of Gram-negative microorganisms were isolated from multiple organs, with Salmonella arizonae the most significant isolate. An outbreak of salmonellosis was reported in farmraised Nile crocodiles (Huchzermeyer, 1991). In Zimbabwe, over a 4-year period, 49.4% of bacterial isolates from farmreared Nile crocodiles submitted for postmortem evaluation

Bacterial Diseases of Reptiles  465

were identified as Salmonella spp. (Foggin, 1992). Group C Salmonella and Salmonella typhimurium were considered primary pathogens. Salmonella was cultured from lesions in the gastrointestinal tract, liver, spleen, and vessels of two blood pythons (Python curtus) (Cambre et al., 1980), and S. arizonae (formerly Arizona hinshawii) was cultured from a variety of tissue sites from three of four red-tailed boa constrictors (Boa constrictor) to die in another zoologic collection (Boever and Williams, 1975). Numerous yellow necrotic plaque-like lesions of fibrinonecrotic membranes were seen over the mucosa of the small and large intestine. In one animal, multiple abscesses protruded from the serosal surface of the gastrointestinal system with additional involvement of the lungs, spleen, and gallbladder. Salmonella was isolated from hepatic lesions in a hognose snake (Heterodon sp.) and twin-spotted rattlesnake (Crotalus pricei), and from epi- and myocardial granulomas in a Mexican cantil (Agkistrodon bilineatus) (Frye, 1991). Salmonella arizonae (formerly Arizona hinshawii) was cultured from the blood of an eastern diamondback rattlesnake (Crotalus adamanteus), Boelen’s python (Morelia boeleni), and two diamond pythons (Morelia spilota) having variable clinical signs (Lamberski et al., 2002). All four snakes either died or were euthanized; Salmonella arizonae was associated with necrotizing gastritis in a rosy boa (Lichanura trivirgata) and necrotizing tracheitis in a Honduran milk snake (Lampropeltis hondurensis) in a zoological collection on Gran Canaria, Spain (Oros et al., 1996). Using a rabbit antiserum to Salmonella arizonae, immunoperoxidase staining was used to label the bacteria in tissue section. In a study of osteoarthritis and osteoarthrosis in 15 snakes, Salmonella was cultured from the blood and affected bone of six snakes (Isaza et al., 2000). Osteomyelitis associated with Salmonella enterica arizonae was followed in a colony of ridgenose rattlesnakes (Crotalus willardi) over a 5-year-period (Ramsay et al., 2002). Of 19 snakes in the original study group, six with bony lesions (Figure 10.12) at the beginning of the study died or were euthanized. All extraintestinal isolates except one was S. e. arizonae serotype 56: Z4,Z24. While there were many isolates of other serotypes from the cloaca (84% of samples), this serotype was isolated from only one cloacal sample.

10.5 Citrobacter The majority of reports documenting Citrobacter infections in reptiles involve chelonians. Septicemic cutaneous ulcerative disease (SCUD) was described for a variety of pond turtles, including the genera Pseudemys, Chrysemys, and Emys (Kaplan, 1957). This disease appears to be management related because predisposing conditions are necessary to initiate an epizootic and include poor nutrition and maintenance in stagnant filthy water coupled with skin abrasions. Turtles will ultimately become lethargic with reduced muscle tone, limb paralysis, and necrosis of digits with hemorrhage and

cutaneous ulcerations (Jacobson, 1981) (Figure 10.13). There may be septicemia with necrotic foci in the liver, heart, kidney, and spleen. In a subsequent report (Jackson and Fulton, 1970), Serratia was incriminated in the pathogenesis of SCUD by its lipolytic, proteolytic action on tissues, thereby allowing Citrobacter to more effectively invade affected tissue.

10.6 Serratia Serratia has been isolated from numerous reptile species, including both healthy and ill animals. Serratia was recovered from rattlesnake venom and swabs of fangs (Ledbetter and Kutscher, 1969), from the oral cavity of captive caimans, and from the oral cavity and cloaca of free-ranging white cay rock iguanas (Cyclura rileyi). Although a nonchromogenic Serratia marcescens was isolated from pyogenic arthritis in a tegu lizard (Tupinambis teguixin), the organism was not isolated from systemic granulomatous lesions found throughout the body (Ackerman et al., 1971). Serratia marcescens biotype A4 (formerly Serratia anolium) was isolated from tumor-like lesions in the Knight anole (Anolis equestris) (Duran-Reynals and Clausen, 1937). This organism was experimentally demonstrated to cause infection when inoculated subcutaneously in a variety of poikilothermic vertebrates (Clausen and Duran-Reynolds, 1937). Serratia marcescens was also cultured from subcutaneous abscesses in a green iguana and a northeastern spiny-tailed iguana (Ctenosaura acanthura) (Boam et al., 1970a). As previously mentioned, Serratia was incriminated in the pathogenesis of SCUD in freshwater turtles (Jackson and Fulton, 1970). Serratia marcescens was isolated from an aseptically obtained blood sample of a West African dwarf crocodile (Osteolaemus tetraspis) with osteomyelitis of the right ulna and radius (Heard et al., 1988). The crocodile died following attempts at a blood transfusion, and necropsy revealed a suppurative polyarthritis involving both coxofemoral and right radiocarpal joints with associated synovitis, cellulitis, and caseating arthritis of the left femorotibial joint. Serratia marcescens was cultured from subcutaneous abscesses (Figures 10.14–10.15) in a gopher tortoise (Gopherus polyphemus) (Pye et al., 1999). Despite surgical removal and administration of an appropriate antimicrobial, abscesses recurred. Evaluation of the tortoise using magnetic resonance imaging revealed masses within the lung. The tortoise was euthanized and multiple abscesses were found in the lung, kidney, and liver. The pathogenesis of Serratia abscesses in reptiles is poorly understood. Because this organism appears to be part of the normal oral flora of several species, organisms possibly become introduced subcutaneously through a bite wound or other traumatic damage to the integument.

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10.7 Pasteurella A bacterium belonging to the genus Pasteurella was isolated from the respiratory tract of desert tortoises (Snipes et al., 1980), and eventually species status was proposed for these isolates under the name Pasteurella testudinis sp. nov. (Snipes and Biberstein, 1982; Snipes et al., 1995). Because this organism was isolated from the respiratory tract of healthy desert tortoises and those with an upper respiratory tract disease (URTD), its significance was initially unknown. In some desert tortoises with URTD, P. testudinis represented up to 100% of the aerobic  microbial isolates from the nasal cavity. Subsequently, transmission studies showed that a Mycoplasma isolated from desert tortoises with URTD (see Section 10.17 below), rather then Pasteurella, was the cause of this disease (Brown et al., 1994). Pasteurella testudinis was also found to be associated with a respiratory disease and septicemia in captive leopard (Geochelone pardalis) and other tortoises from five outbreaks at three locations in the Pretoria area of South Africa (Henton, 2003). Because there were no attempts to culture Mycoplasma or viruses, the exact causative agent was not identified. Pasteurella multocida and Staphylococcus aureus were cultured from several members of a herd of American alligators exhibiting a respiratory disease (Mainster et al., 1972). Many of these alligators had clinical signs of respiratory disease including a serous and purulent discharge from the nostrils.

10.8 Erysipelothrix Cutaneous lesions due to Erysipelothrix rhusiopathiae (formerly insidiosa) have been reported in a 100-year-old American crocodile (Crocodilus acutus) in a zoologic park in Florida (Jasmin and Baucom, 1967). Of 122 terrapins necropsied in a zoological collection, E. rhusiopathiae was isolated from the spleen of a single common snapping turtle (Keymer, 1978b).

10.9 Neisseria Bite-wound inoculation of bacteria is probably an important source of infection and abscess formation in green iguanas. In one group of green iguanas and a rhinoceros iguana, (Cyclura cornuta) a new Gram-negative diplococcus, Neisseria iguanae, was isolated from abscesses and the oral cavities of healthy and infected cagemates, indicating inoculation by intraspecific aggression (Barrett et al., 1994; Plowman et al., 1987). Several of these animals developed chronic abscesses (Figure 10.16) and septicemia that responded poorly to treatment. Gram-negative bacteria were identified in the lesions, and in some lesions the bacteria were surrounded by dense eosinophilic amorphous material radiating from the periphery (Figure 10.17). A similar change in humans responding

to certain bacteria, fungi, and parasites has been reported as a Splendore-Hoeppli reaction.

10.10 Elizabethkingia In a map turtle (Graptemys barbouri) that died with cutaneous edema, light microscopic examination of tissues stained with hematoxylin and eosin revealed numerous intracytoplasmic basophilic bodies within Kupffer cells in the liver and macrophages in multiple tissues (Jacobson et al., 1989b) (Figures 6.108, 10.18–10.19). Transmission electron microscopy (TEM) demonstrated the bodies as cytosomes that contained numerous nonflagellated bacteria. Subsequently, a bacterium was cultured from the turtle and identified as Elizabethkingia meningoseptica (formerly Flavobacterium meningosepticum).

10.11 Vibrio In juvenile Kemp’s ridley sea turtles (Lepidochelys kempi) reared for head starting at the National Marine Fisheries Service, Southeast Fisheries Centers Laboratory in Galveston, Texas, Vibrio parahemolyticus was isolated from the blood of turtles with a hemorrhagic disease (Leong et al., 1989). Affected turtles often vomited blood before death, and at necropsy the body cavity was often filled with bloody fluid. It was the opinion of the authors that the outbreak could have been induced by stresses caused by tagging with monel flipper tags. In an oceanarium in Florida, Vibrio sp. was cultured from caseous necrotic debris accumulating adjacent to the glottal opening of hatchling loggerhead sea turtles. In a survey of 102 farmed green turtles (Chelonia mydas) and two hawksbill turtles (Eretmochelys imbricata) in Australia, Vibrio alginolyticus was one of four microorganisms isolated repeatedly from cases of traumatic ulcerative dermatitis, ulcerative stomatitis, obstructive rhinitis, and bronchopneumonia (Glazebrook and Campbell, 1990a). Vibrio alginolyticus was one of four bacteria isolated from hatchling and juvenile farmed and oceanarium-reared green turtles and loggerhead sea turtles in Australia with an ulcerative stomatitis–obstructive rhinitis pneumonia disease complex (Glazebrook et al., 1993). Green turtles in Hawaii with fibropapillomas (FP) were found to be bacteremic, with four species of Vibrio spp. representing a majority of the bacteria cultured (Work et al., 2003).

10.12 Listeria Listeria monocytogenes was isolated from the brains of American alligators and swine at a farm in Florida (Frye, 1991). When swine at the farm died, they were fed to alligators, and when the alligators were butchered, the carcasses were fed to swine. An adult male bearded dragon (Pogona vit-

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ticeps) that died following a 3-day period of a nonspecific illness was found by light microscopy to have microabscesses in the brain, kidney, spleen, heart, and liver (Girling and Fraser, 2004). Swabs of multiple tissues and heart blood were cultured and resulted in the isolation of L. monocytogenes. Newborn mice had recently been added to the lizard’s diet. Cultures of cardiac blood from the mice also resulted in growth of L. monocytogenes.

10.13 Helicobacter The detection of spiral-shaped Helicobacter-like bacteria associated with gastritis in a chelonian resulted in a broader study to look for similar organisms in gastric lesions in turtles and tortoises (Busch et al., 2002). Using light microscopy, of the 28 dead chelonians (6 species), 22 had gastritis, with 5 associated with spiral-shaped Helicobacter-like organisms. The gastritis was variable with the following changes seen: acute glandular distension and necrosis with intraglandular heterophils and more chronic glandular atrophy and interstitial fibrosis as well as glandular regeneration and epithelial hyperplasia. To identify the organism, bacterial DNA was extracted, amplified, and the 16 s rRNA gene sequenced from two turtles and a tortoise. The sequences demonstrated phylogenetic homology to the genus Helicobacter. This study indicates that Helicobacter should be in the differential when attempting to determine the cause of gastritis in reptiles.

10.14 Streptococcus While infections with Gram-positive bacteria are relatively uncommon in reptiles, they do occur. Two Singapore house geckos (Gekko monarchus) died of a disseminated Grampositive bacteremia (McNamara et al., 1994). Chain-forming colonies of a Streptococcus infiltrated and effaced architecture of all examined tissues and a thick capsule surrounded groups of these chains (Figures 10.20–10.21). Electron microscopy revealed bacterial cells surrounded by two cell walls and a thick peptiglycan layer; a thin fibrous material was seen spreading out from the surface (Figure 10.22). What was unusual was a minimal to absent inflammatory response. Streptococcus agalactiae group B, serotype V, was isolated from three green tree (emerald) monitors (Varanus prasimus) that died as a result of septicemia (Hetzel et al., 2003). Using light microscopy, coccoid bacteria were identified in monocytes and heterophils in multiple tissues, and as cell free clumps and chains within cardiac atrial, ventricular, and luminae of blood vessels. Immunohistochemical staining resulted in labeling in tissue section. The bacteria were identified by routine bacteriological methods and PCR amplification of species-specific parts of the 16S rRNA gene, the 16S-23S rDNA intergenic spacer region, and the CAMP factor

gene cfb. Identical bacteria were isolated from the gastrointestinal tract of mice fed to the lizards. Pathogenic streptococcal infections have been reported in other lizards (Zwart and Cornelisse, 1972).

10.15 Anaerobes Anaerobic bacterial infections are probably more common in reptiles than reported in the literature. More than likely, this is a reflection of the relatively few cultures obtained from reptiles that are submitted for anaerobe culture. Complicating the situation is whether an isolated anaerobe is the cause of a disease or lesion or is simply a member of the normal flora that showered at death. Clostridium was reported to be an inhabitant of the oral flora of American alligators (Doering et al., 1971), and along with Bacteroides, the oral flora of snakes in a zoological collection (Draper et al., 1981). Clostridium difficile, C. innocuum, C. perfringens, and C. sordelli were frequently identified in cultures of blood samples obtained from healthy appearing captive lizards, particularly monitors (Hanel et al., 1999). Blood was obtained from the ventral coccygeal vein, and, despite cleansing the tail several times before sampling blood, spores of these bacteria could have been present in scale folds resulting in contamination of the sample. The alternative explanation would be that Clostridium or their spores were present in the blood of the healthy appearing monitors in this report. Clostridium perfringens was cultured from the blood and C. sordelii from a vertebral aspirate of a rough-necked monitor (Varanus rudicollis) with spondylosis (D’Agostino et al., 2006). Vertebrae may have been infected with Clostridium circulating in the blood. While difficult to interpret, there are a number of reports of diseases in reptiles caused by infection with anaerobes. A fatal septicemia in chelonians was attributed to Clostridium novyi (Urbain et al., 1951). Clostridium perfingens was cultured from green iguanas with gangrenous necrosis of the limbs (Fiennes, 1959). Clostridium botulinum was considered the causative agent of a neurologic disease of captive green turtles called floppy flipper disease (Jacobson, 1980). During a 2-year study period, 65 specimens from reptile clinical cases were submitted for aerobic and anaerobic culture (Stewart, 1990). Of 39 specimens that resulted in bacterial growth, 21 included the following anaerobic bacteria: Bacteroides, Fusobacterium, Clostridium, and Peptostreptococcus. The cultures were predominantly from abscesses. A Clostridium sp. was isolated from edema fluid of four gharials (Gavialis gangeticus) having swollen limbs (Misra et al., 1996). Clostridium septicum was isolated from a crocodile with hepatitis and septicemia at a farm in Australia (Buenviaje et al., 1994). Diarrhea in a red-footed tortoise was associated with an enterotoxigenic strain of Clostridium perfringens (Weese and Staempfli, 2000). Entertoxemia caused by Clostridium glycolicum resulted in circulatory collapse and death of a female ornate Nile monitor (Varanus ornatus) (Bertelsen and Weese,

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wide variety of poikilotherms. Griffith (1939) isolated Mycobacterium marinum from 71% (20 of 28 cases) of the mycobacterial cases at London Zoological Park between 1924 and 1933. Mycobacterium marinum was identified in or cultured from hepatic and splenic granulomas in two Australian snakenecked turtles (Chelodina longicollis) (Schildger et al., 1991), from the lung of a loggerhead sea turtle (Caretta caretta) with 10.16 Actinomycetales a pulmonary nodule (Leong et al., 1989), and from tissues of The order Actinomycetales consists of several families includ- one of five hawksbill sea turtles (Eretmochelys imbicata) with ing the Mycobacteriaceae, Nocardiaceae, and Dermatophi- visceral granulomas containing acid-fast organisms (Posthaus laceae. The members of this group are united by certain et al., 1997). Mycobacterium marinum was identified in cutamorphological, biochemical, and molecular attributes. All neous lesions of a group of Egyptian spiny-tailed lizards (Uroare filamentous and either form spores or reproduce by frag- mastyx aegypticus) that were formerly burned and housed mentation. Mycobacterium is the only genus within the fam- in an aquarium that was formerly used for fish (Morales and ily Mycobacteriaceae. Mycobacterium is the most common Dunker, 2001). Mycobacterium kansasii, another slow-growactinomycete infection in reptiles. While Dermatophilus has ing Runyon group I photochromagen, was identified in a been seen in all major groups of reptiles, few cases of Nocar- captive Chinese soft-shelled turtle (Pelodiscus sinensis) (Oros et al., 2003). One year after purchase, the turtle developed dia have been reported. several small white foci on its carapace, showed signs of respiratory distress, and subsequently died. At necropsy sev10.16.1 Mycobacterium eral white papular lesions were found on the neck and caraThe genus Mycobacterium is divided into those in the Myco- pace, and white nodules were found within the pulmonary bacterium tuberculosis complex and the nontuberculous parenchyma. The lung nodules consisted of granulomas, with Mycobacterium. Runyon (1970) developed a classification acid-fast bacteria in their centers. Sheets of epithelioid cells, scheme based upon the growth rate and colonial pigmen- with small numbers of acid-fast organisms, were found in the tation of the nontuberculous Mycobacterium. Growth was dermis of the skin lesions. Fungi were also seen in the skin either slow (> 7 days) or rapid (< 7 days). Runyon Group I lesions. consists of those Mycobacterium that are photochromogens. The only report of M. avium (Runyon group III) from a Members of this group form pigment in the light after being reptile is the isolation of this slow-growing nonphotochrogrown in the dark; colonies take more than 7 days to appear. mogenic mycobacterium from Pacific green turtles (Brock et Runyon group II members are the scotochromogens. Mem- al., 1976). Of 120 hatchling green turtles (from the French bers of this group form pigment in the dark or light; colonies Frigate Shoals, HI) brought to the University of Hawaii for take more than 7 days to appear. Runyon group III consists study, 22 died during the first 5 months in captivity. None of the nonphotochromogens. Members of this group do not were found to have mycobacteriosis. From the 6th through form pigment in the dark or light; colonies take more than 7 the 7th month of captivity, 9 additional turtles died. Impresdays to appear. The last group, Runyon group IV, are rapid sion smears of 4 of 5 turtles made from granulomatous growers; colonies take fewer than 7 days to appear. Most of lesions in the liver, kidney, and spleen had acid-fast stainthe mycobacteria isolated from reptiles are within Runyon ing organisms. Specimens from two turtles were sent to the groups I and IV (Brownstein, 1978). Sequencing of the Myco- National Animal Disease Center in Ames, IA, and M. avium bacterium genome is now being used to better understand was cultured. the phylogenetic relationships of the various members of Compared to M. marinum, M. thamnopheos, and M. these groups. chelonei are less frequently isolated from reptiles; they are Mycobacteriosis is a sporadic disease in well-managed rapid growers (Runyon group IV). Aronson (1929) isolated reptile collections, having an annual incidence of 0.1 to 0.5% M. thamnopheos from garter snakes (Thamnophis spp.) at (Brownstein, 1978, 1984). Because the major mycobacterial the Philadelphia Zoologic Park. M. chelonei was isolated agents infecting reptiles are ubiquitous organisms in nature from a boa constrictor with acid-fast organisms in prolifand the prevalence of infection is low, it appears that a com- erative ulcerative oral lesions (Figure 10.24), paratracheal mensal relationship probably exists under most situations, and pulmonary granulomas (Figures 10.25–10.26), and with actively infected animals being predisposed to disease. hepatic granulomas (Quesenberry et al., 1986). MycobacteThe most common isolate for reptiles is Mycobacterium mari- rium thamnopheos was isolated from another boa constricnum, followed by cases of M. chelonei and M. thamnopheos. tor with similar lesions (Kiel, 1977a) and Mycobacterium Cases of M. haemophilum and M. kansasii have also been chelonei was isolated from the elbow joint and skin nodule reported. Mycobacterium marinum is a ubiquitous, sapro- of a Kemp’s ridley sea turtle with osteoarthritis (Greer et phytic, slow-growing photochromogen (Runyon group I) in al., 2003). The turtle was subsequently euthanized and M.  warm aquatic systems and is potentially pathogenic for a 2006). The author evaluated a green iguana that developed a necrotizing cellulitis following a bite from a monitor lizard (Figure 10.23); Peptostreptococcus was cultured from the wound and eventually the limb had to be amputated.

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chelonei was isolated from the lung, liver, kidney, spleen, and pericardium. In many cases acid-fast organisms are identified in tissue section without culture and specific identification (Figures 5.59–5.66, 10.27). Acid-fast-staining organisms were identified in pulmonary tubercles in a loggerhead sea turtle (Friedmann, 1903), in plastronal and pulmonary lesions in an African soft-shelled turtle (Trionyx triunguis) (Besse, 1949), in cutaneous lesions, spleen, liver, and the lung of a Hilaire’s side-necked turtle (Phrynops hilari hillarii) (Rhodin and Anver, 1977), and in the lung of 2 of 104 necropsied sea turtles farmed in northern Australia (Glazebrook et al., 1990a). Mycobacterium was identified in the subcutis or multiple caseating granulomas in freshwater and saltwater crocodiles (Ariel et al., 1997), and in granulomas in the heart, liver, and lung (Figure 10.28) of a reticulated python (Python reticulates) (Olson and Woodard, 1974), and in necroproliferative oral lesions in an albino Texas rat snake (Elaphe obsoleta lindheimerii) (Figure 10.29). Molecular tools are being used to make specific identifications of Mycobacterium. In a retrospective study of granulomatous lesions in 90 reptile cases necropsied at the University of Zurich between 1990 and 1991, acid-fast bacilli were found in 14 cases (Soldati et al., 2004). Using PCR followed by DNA sequencing, 23 cases were positive for Mycobacterium, and the following specific identifications were made in nine of these cases: M. agri, M. chelonae (cheloni is a synonym), M. confluentis, M. haemophilum, M. hiberniae, M. neoarum, and M. nonchromogenicum. In another study, using polymerase chain reaction and DNA sequencing, M. haemophilum and M. marimum were identified in pulmonary granulomas (containing acid-fast organisms) of a royal (ball) python (Python regius) (Hernandez-Divers and Shearer, 2002). Natural cases of reptilian mycobacteriosis are generally seen as chronic diseases, although acute episodes have been described. Aside from the cases presented with cutaneous lesions, those animals systemically infected generally show nonspecific signs, including anorexia, listlessness, and chronic weight loss. Dyspnea may be seen in animals having pulmonary involvement. Involvement of the integumentary system is usually seen as either subcutaneous nodules or ulcerative skin lesions. These lesions are easy to biopsy, and acid-fast organisms can be demonstrated in impression smears or in histologic section. Cases of systemic infection following cutaneous involvement have been reported (Rhodin and Anver, 1977). In acute mycobacteriosis, there is extracellular proliferation of acid-fast organisms with associated caseous necrosis and an infiltrate of heterophils and macrophages; fibroplasia is minimal. In chronic mycobacteriosis, there is typically granulomatous inflammation with development of tubercles. Granulomas may range in development from compact nests of macrophages containing intracellular acid-fast bacilli, to more complex granulomas containing caseated centers, and finally to expansion of the caseous region to involve the entire

tubercle with large numbers of extracellular bacilli (Brownstein, 1984).

10.16.2 Nocardia A Kemp’s ridley sea turtle that stranded during a cold-stunning event developed a swelling of the right foreflipper adjacent to the carpals and metacarpals several months later (Harms et al., 2002). Radiographs indicated osteolysis of metacarpal I. Fine needle aspirates of the carpal joint revealed septate nonpigmented, nonbranching fungal hyphae; Nocardia sp. was cultured from the aspirate. The infection was controlled with a variety of antimicrobials and remodeling of the carpal bones eventually occurred. The turtle was eventually released.

10.16.3 Dermatophilus 10.16.3.1 Chelonia  Dermatophilus is a unique bacterium that forms filamentous structures with transverse segmentation resulting in the formation of packets of coccoid cells that become motile spores. An organism consistent with Dermatophilus was isolated from skin lesions of three chelonians in an Australian Zoo, found to be biochemically distinct from D. congolensis and was placed in a new species, D. chelonae (Masters et al., 1995). A D. chelonae–like bacterium was isolated from cutaneous and visceral lesions of four bowsprit tortoises (Chersina angulata) and one Egyptian tortoise (Testudo kleinmanni) in a zoologic collection in the United States (Bemis et al., 1999; Ramsey et al., 1998). In the bowsprit tortoises, the skin lesions were primarily seen on the neck and consisted of yellow-white nodules covered by dry flaking skin with tracts extending into the dermis; one tortoise had a septic arthritis. One dead Egyptian tortoise had a coelomic mass containing filamentous organisms that stained with Giemsa stain.   The author of this chapter evaluated skin biopsies of four captive desert tortoises in a private collection in Arizona, which developed hyperkeratotic skin lesions at multiple soft tissue sites (Figure 10.30). By light microscopy, a filamentous organism resembling Dermatophilus was observed. The author also observed similar appearing organisms in necrotizing skin lesions, especially involving the integument surrounding the upper and lower jaws in speckled padloppers (Homopus signatus) (Figure 10.31).

10.16.3.2 Crocodylia  A Dermatophilus-like organism was identified in ventral brown skin lesions of an American alligator (Alligator mississippiensis) in Florida, following slaughter (Jacobson, 1989). A similar skin lesion was reported in farmed American alligators in Louisiana, and was named brown spot disease (Bounds and Normand, 1991; Newton, 1992). While the lesion can commence anywhere on a scale, the lesion most commonly developed at the junction between scales. An organism resembling Dermatophilus was isolated from this lesion (Bounds and Norman, 1991). Dermatophilus

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was seen in young captive crocodiles in Papua New Guinea (Ladds and Sims, 1990) and was isolated from skin lesions in farmed saltwater crocodiles in Australia (Buenviaje et al., 1997), where it accounted for 28.1% of the pathogens identified in 203 skin lesions of 180 crocodiles with dermatitis at 10 farms (Buenviaje et al., 1998). The lesion was seen as focal brown spots, primarily on the ventral abdomen (Figure 10.32). Three microscopic stages of development were noted. In stage 1, there was a lifting of the keratin with some debris accumulation. Dermatophilus filaments were observed in this lesion (Figure 10.33). In stage 2 there was epidermal necrosis, with the stratum basale intact. In stage 3 there was ulceration of the epidermis with extension of Dermatophilus into the subcutis.

10.16.3.3 Sauria  The first report of dermatophilosis in a lizard involved a bearded dragon (Pogona barbata [formerly Amphibolurus barbatus barbatus]) submitted with several subcutaneous nodules and an abdominal nodule having a central necrotic core (Simmons et al., 1972). Using light microscopy, Gram-positive branching filamentous structures consistent with D. congolensis were seen in this lesion. Dermatophilus congolensis was isolated in pure culture and was transmitted by inoculation to four bearded dragons, one bluetongued skink (Tiliqua scincoides scincoides), and one sheep. Dermatophilosis was subsequently identified in three captive Australian bearded dragons (Montali et al., 1975) and two marble lizards (Calotes mystaceus) (Anver et al., 1976), imported into the United States. An organism microscopically compatible with Dermatophilus was identified in skin lesions of a bush anole (Polychrus marmoratus) (Figures 10.34–10.37), a collared lizard (Crotophytus collaris) (Figures 10.38– 10.39), six Senegal chameleons (Chamaeleo senegalensis) (Figures 10.40–10.41), a green iguana (Iguana iguana), a savannah monitor (Varanus exanthematicus) and a captive panther chameleon (Furcifer pardalis) (Jacobson, 1989; Jacobson, 1991; Origgi et al., 1999; Ryan, 1992). In the bush anole, chameleons, collared lizard, and iguana, the lesions were seen as raised brown multifocal encrustations scattered over the integument. The encrustations were found to consist of necrotic cellular debris, accumulating keratin, and inflammatory cells, with necrosis of the underlying epidermis. While organisms could be visualized with H&E staining, numerous branching filamentous organisms with longitudinal and transverse divisions were observed in Gram- and Giemsa-stained tissue sections. 

10.16.3.4 Ophidia  In snakes, an organism compatible with Dermatophilus was identified in subcutaneous masses removed from a boa constrictor (Jacobson, 1989) (Figures 10.42–10.43) and in several subcutaneous masses removed from a 10-year-old male king cobra (Ophiophagus hannah) in a zoologic collection (Wellehan et al., 2004a). Regarding the cobra, fourteen months after the original masses were removed, a similar appearing mass was removed

from a different site. One year later, another mass appeared at the first surgical site. An organism was cultured from one of the surgically removed masses and was identified by 16S rRNA gene sequencing as Dermatophilus chelonae.

10.17 Mycoplasma 10.17.1 Chelonia The first Mycoplasma isolated from a reptile was Mycoplasma testudinis, which was taken from the cloaca of a Greek tortoise (Hill, 1985). This organism was considered a nonpathogenic commensal. In 1988 a chronic upper respiratory tract disease (URTD) was recognized in desert tortoises in the Desert Tortoise Natural Area, Kern County, CA (Jacobson et al., 1991a). Partially because of this disease and reported population declines in the western Mojave Desert, desert tortoises north and west of the Colorado River were listed as threatened by the U.S. Department of the Interior in 1990. Since this first report, desert tortoises with signs of URTD have been seen at other locations in California, Arizona, and Utah (Homer et al., 1998). The disease is characterized clinically by serous, mucous, or purulent nasal (Figures 10.44–10.45) and ocular discharge, conjunctivitis, and palpebral edema. The major changes seen were in the nasal cavity, a complex structure located between the external nares and eyes (Figure 10.46). The nasal cavity is lined by a dorsal olfactory epithelium (Figure 10.47) and a ventral mucous epithelium with intercalated ciliated epithelial cells (Figure 10.48). With URTD, at a light microscopic level there is infiltration of the nasal cavity mucosa and submucosa with inflammatory cells accompanied by hyperplasia and degeneration of upper respiratory tract epithelium (Jacobson et al., 1991a; Brown et al., 1994; Jacobson et al., 1995) (Figures 10.49–10.51). A similar appearing disease has been seen in free-ranging gopher tortoises (Gopherus polyphemus in Florida (McLaughlin et al., 2000) (Figures 10.52–10.53), and in a variety of species of captive nonnative tortoises imported as part of the pet trade (Wendland et al., 2006). A similar disease has been seen in long-term captive Greek (spur-thighed) and Hermann’s (T. hermanni) tortoises in Europe (Lawrence and Needham, 1985). It is considered to be more severe in Greek tortoises compared to Hermann’s tortoises (Martinez-Silvestre and Mateu-de Antonio, 1997). While a wide variety of aerobic bacteria were isolated from these tortoises, none were considered to be the causative agent. Fowler (1977), in studying a respiratory disease in desert tortoises, identified bacteria from the respiratory tract of both clinically healthy tortoises and tortoises showing signs of URTD. The results of his study also failed to implicate a specific bacterial organism as a cause of respiratory disease in the studied tortoises. In follow-up studies of URTD in desert tortoises, no major differences were observed regarding microbial isolates from the respiratory tract of both clinically

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healthy tortoises and tortoises showing signs of URTD (Dickinson et al., 2001; Snipes et al., 1980). Electron microscopic examination of the upper respiratory tract mucosa of free-ranging desert and gopher tortoises with URTD demonstrated the presence of a cell membrane–associated bacteria with features compatible with Mycoplasma (Jacobson et al., 1991a; McLaughlin et al., 2000) (Figures 6.100–6.101, 10.54). A previously undescribed species of mycoplasma was cultured from nasal lavages of affected desert tortoises, and was named Mycoplasma agassizii, (Brown et al., 1995). Experimental infection studies of desert tortoises (Brown et al. 1994) and gopher tortoises (Brown et al. 1999) demonstrated Mycoplasma agassizii to be an etiologic agent of URTD. These findings led to the development of the optimized culture, polymerase chain reaction (PCR) (Brown et al., 1995), and enzyme-linked immunosorbent assay (ELISA) (Schumacher et al. 1993) tests for M. agassizii necessary to determine infection, exposure, and immune status of asymptomatic and URTD-symptomatic desert and gopher tortoises. The limitations of each test and the information they provide have been reported (Brown et al., 2002). To date, mycoplasmas have been recovered by culture or detected by PCR in the following species of chelonians: desert tortoise, gopher tortoise, Texas tortoise (Gopherus berlandieri), Chaco (Argentine) tortoise (Geochelone chilensis), leopard tortoise (G. pardalis), Indian star tortoise (G. elegans), African spurred tortoise (G. sulcata), radiated tortoise (Geochelone radiata), red-footed tortoise (Geochelone carbonaria), Travancore tortoise (Indotestudo travancorica), Forsten’s tortoise (Indotestudo forstenii), spider tortoise (Pyxis arachnoides), flat-tailed tortoise (Pyxis planicauda), spurredthighed tortoise, marginated tortoise (T. marginata), Egyptian tortoise, Russian tortoise (T. horsfieldii), Hermann’s tortoise, and eastern box turtles (Terrapene carolina carolina) in the United States (Brown et al., 2002; Wendland et al., 2006), and captive Testudo sp. in the United Kingdom (Soares et al., 2004). A mycoplasma survey of 70 free-ranging and 34 captive adult eastern box turtles conducted in New York state revealed that while free-ranging box turtles were frequently seropositive by ELISA, fewer captive box turtles were positive (Calle et al., 1998). Using PCR, a novel Mycoplasma, distinct from M. agassizii, was detected in nasal swabs from eastern box turtles with an upper respiratory disease (Feldman et al., 2004). During routine screening of desert tortoises with URTD, a second genetically distinct mycoplasma, represented by tortoise isolate H3110 (American Type Culture Collection accession 700618), was cultured from affected tortoises (Brown et al., 1995). This was distinguished from previously described mycoplasmas using serology and comparing 16S rRNA gene sequences, and was named M. testudineum (Brown et al., 2004). The ELISA developed to determine exposure of tortoises to M. agassizii may not detect exposure to M. testudineum. There is some evidence that there are strains of M. agassizii that do not cause overt clinical disease. Thus, the strain

of Mycoplasma present in a given population will influence the clinical course of disease and transmission from tortoise to tortoise. There are anecdotal reports that some tortoises may clear an infection; however, these have been very difficult to substantiate.

10.17.2 Crocodylia An unspeciated Mycoplasma was isolated from a captive gharial (Gavialis gangeticus) in India that died of a respiratory disease (Misra et al., 1996). Outbreaks of a disease characterized by polyarthritis and associated with a Mycoplasma sp. were reported in one- to five-year-old Nile crocodiles (Crocodylus niloticus) on several farms in Zimbabwe (Mohan et al., 1995). The organism was isolated from both joints and lung tissue and caused arthritis and pneumonia. The disease was reproduced following experimental inoculation of crocodiles with this organism. The causative agent was named M. crocodyli (Kirchoff et al., 1997). A study to demonstrate horizontal transmission by exposing experimentally infected crocodiles to those that were uninfected was unsuccessful (Mohan et al., 1996). This suggested that vertical transmission through the eggs of infected females was possible. A Mycoplasma sp. was also identified in the feces of disease-free Nile crocodiles using negative staining TEM (Huchzermeyer et al., 1994a) and a Mycoplasma strain similar to that from Zimbabwe, and chlamydia, were isolated from farmed crocodiles in Israel with polyarthritis (Levisohn cited in Mohan et al., 1996). In 1995 an epidemic of pneumonia, pericarditis, periportal hepatitis, fibrinous polyserositis, and multifocal arthritis was reported in a captive group of American alligators (Alligator mississippiensis) in Florida (Clippinger et al., 2000) (Figures 5.55–5.56, 10.55–10.63). While pneumonia was seen in most of the affected alligators, often secondary mixed Gram-negative bacteria were isolated from lung specimens and fungi were identified in tissue sections of some cases. One alligator had a severe meningoencephalitis (Figures 10.64–10.66). A Mycoplasma sp. was cultured from ill and dead alligators, and using electron microscopy was identified in synovial tissue (Figure 6.102). It was subsequently named M. alligatoris (Brown et al., 2001a). Transmission studies in American alligators confirmed that M. alligatoris is a pathogen in this species (Brown et al., 2001b). Alligator immunoglobulin was purified from alligator plasma and a polyclonal antibody was raised in rabbits against the purified immunoglobulin. Polyclonal anti-alligator antibodies were purified and conjugated with biotin. This was used as the secondary antibody in an indirect ELISA developed for detecting exposure of alligators to M. alligatoris (Brown et al., 2001c). In a challenge study with M. alligatoris in broad-nosed caiman (Caiman latirostris), and Siamese crocodiles (Crocodylus siamensis), while 3 of 6 caiman died of mycoplasmosis following challenge, none of the Siamese crocodiles developed mycoplasmosis (Pye et al., 2001).

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10.17.3 Sauria Recently a novel Mycoplasma was isolated from vertebrae of green iguanas with granulomatous osteomyelitis (Brown et al., 2005, 2006). The two iguanas had hindlimb paralysis and were unable to urinate and defecate. They were also found to have meningitis and demyelination of the spinal cord. Nucleotide sequences for the 16S rRNA indicated that the isolate was new and was subsequently named M iguanae sp. nov.

10.17.4 Ophidia A novel Mycoplasma sp. was isolated from a Burmese python (Python molurus bivittatus) with a proliferative tracheitis and pneumonia (Penner et al., 1997) (Figure 10.67). Using PCR, amplification of the 16S rRNA gene resulted in a product that had 90% similarity with nucleotide sequences of M. agassizii, a causative agent of URTD of desert and gopher tortoises. Using TEM, an organism compatible with mycoplasma was observed on the surface of cells lining the trachea and lung airways (Figure 6.103).

10.18 Chlamydia and Chlamydophila The order Chlamydiales consists of four families. Of these, the family Chlamydiacea contains members that are known to be pathogens in humans and other animals. A reclassification of Chlamydiacea has resulted in the recognition of nine species within the following two genera: Chlamydia (C. trachomatis, C. suis, and C. muridarum) and Chlamydophila (C. psittaci, C. pneumoniae, C. felis, C. pecorum, C. abortus, and C. caviae) (Everett et al., 1999). Chlamydia and Chlamydophila are obligate intracellular pathogens of humans and a wide range of birds and animals. They have been characterized by their unique developmental cycle, which involves the interconversion among an extracellular survival form, the elementary body (EB), and an intracellular replicating form, the reticulate body (RB).

10.18.1 Chelonia A bacterium identified as Chlamydia psittaci was isolated in chicken eggs inoculated with lung tissue from a captive Moorish (Greek) tortoise with pneumonia (Vanrompay et al., 1994). From August 1990 to June 1991, an epidemic was seen to spread through green turtles in a farming operation in Grand Cayman Island, British West Indies (Homer et al., 1994). Clinical signs included lethargy, anorexia, and inability to dive. Necropsies of eight turtles revealed multiple irregular discrete to patchy pale gray foci throughout the heart of four turtles. By light microscopy, the most consistent lesions were necrotizing myocarditis (Figure 10.68), histiocytic to fibrinous splenitis, and hepatic lipidosis and necrosis. Chlamydiae were demonstrated in macrophages of paraffin-embedded heart, liver, and spleen using a modified Macchiavello’s stain.

Chlamydial antigen was also detected in the cytoplasm of myocardial fibers and in occasional hepatocytes using a commercially available genus-specific antichlamydial monoclonal antibody and the avidin biotin peroxidase complex staining method (Figure 10.69). Electron microscopic examination of the heart of the most severely affected turtle revealed developmental stages of an organism consistent with Chlamydia (Figure 10.70). Polymerase chain reaction amplification of a 293-bp fragment of the 16s rRNA gene and a nested PCR targeting the variable domain IV of the ompA gene detected Neochlamydia, C. abortus, and C. pneumoniae in paraffinembedded heart of an affected green turtle (Bodetti et al., 2002). In a retrospective study of 90 reptile cases with granulomatous lesions, both immunoperoxidase staining with a monoclonal antibody against chlamydial LPS antigen and PCR amplification of a small 92-bp fragment of the 23S rRNA gene were used to detect chlamydiae in these tissues (Soldati et al., 2004). Of 27 chelonian cases (species not identified), one case was both immunoperoxidase and PCR positive, while a second case was positive by PCR only for the presence of C. pneumoniae. Because the granulomas in these two cases could not be classified as either heterophilic or histiocytic, they were simply classified as chronic. Using PCR, a chlamydia-like organism was identified in an additional 14 cases. Nasal lavage samples collected from 155 tortoises (mostly Testudo) that had a nasal discharge were examined for chlamydial DNA by PCR targeting the ompA, ompB, and groESL genes, in addition to the 16S rRNA signature region and the 16S–23S intergenic spacer (Hotzel et al., 2005). Of the tortoises tested, 16 were positive, with clustering of a chlamydia-like agent outside the nine species of Chlamydia and Chlamydophila. Further gene sequencing will be needed to determine the exact taxonomic identity of this clustered group. It is unknown if this organism is commensal or serves as a pathogen.

10.18.2 Crocodylia Chlamydiosis is recognized as a disease problem of farmed Nile crocodiles in Zimbabwe (Foggin, 1992) and South Africa (Huchzermeyer 2003). An acute hepatitis and chronic bilateral blepharoconjunctivitis has been described (Huchzermeyer et al., 1994b). Crocodiles with severe conjunctivitis had accumulations of fibrinous exudates under the nicitans. Chlamydia was isolated in cell culture and chicken eggs infected with liver from Nile crocodiles with hepatitis (Huchzermeyer et al., 1994b). In Zimbabwe, chlamydiosis is commonly associated with adenoviral infection of the liver (Foggin, 1992). In those cases with severe hepatitis, there were lymphoplasmacytic infiltrates, with multifocal areas of necrosis. Chlamydia was isolated along with mycoplasma from farmed Nile crocodiles in Israel that had polyarthritis (Levisohn, 1995 cited in Mohan et al., 1996).

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10.18.3 Sauria

ure 10.75), pneumonia, splenitis, and enteritis. Within the center of many granulomas, basophilic inclusion bodies of The first report of chlamydiosis in a lizard involved a flapvarious sizes were seen (Figure 10.76). At a higher magnecked chameleon, (Chamaeleo dilepis) collected in Tanzanification, the inclusions were composed of small subunits. nia (Jacobson and Telford, 1990). One of three flap-necked By electron microscopy, the inclusions were found to conchameleons that had an intracytoplasmic inclusion within sist of developmental stages of an organism consistent with a circulating monocytes was maintained in captivity and was chlamydial organism (Figure 6.104). An immunoperoxidase sequentially bled over a 55-day period. At 46 days, a secstaining technique using commercially available monoclonal ond type of inclusion was occasionally seen within monoantibodies against chlamydial lipopolysaccharide antigen was cytes. Eventually, all circulating monocytes were found to used to identify the chlamydial antigen in these lesions (Fighave either one or both types of inclusions (Figure 9.61). ure 10.77). Polymerase chain reaction amplification of a 293Blood was collected and fixed in Trump’s solution for TEM. bp fragment of the 16s rRNA gene and a nested PCR targeting Eventually the lizard became moribund and was euthanized. the variable domain IV of the ompA gene detected C. abortus Histologic examination of multiple tissues demonstrated simand C. pneumoniae in paraffin-embedded tissues of these ilar inclusions within macrophages in the spleen and liver snakes (Bodetti et al., 2002). (Figures 9.62, 10.71). Using TEM, inclusions within monoChlamydiosis was identified in a series of emerald tree cytes consisted of either developmental stages of an organboas (ETB) (Corallus caninus) that died in a private collecism compatible with chlamydia and viral particles consistent tion (Jacobson et al., 2002). Of 120 ETB, 81 (67%) developed with members of the family Poxviridae (Figure 10.72). A repetitive regurgitation (3 to 4 days after feeding) during the nested PCR targeting the variable domain IV of the ompA 23-month period after the initial acquisition, and 61 (75%) gene detected the presence of C. pneumoniae in paraffinof these died (Lock et al., 2003). Ninety-seven snakes died embedded spleen and liver of this chameleon (Bodetti et al., or were euthanized; 18 were necropsied and tissues were 2002). collected from all major organs and processed for light Two one-month-old green iguanas from a group of farmmicroscopy. The histologic changes seen in ETBs varied and bred and hatched iguanas having high mortality and showincluded enteritis (5), esophagitis (4), gastritis (4), gastric gland ing nonspecific signs of lethargy and anorexia, and imported atrophy with fibrosis (3), colitis (3), hepatitis (2), nephritis (2), into Florida from Central America in June 1996, were euthrenal tubular necrosis (2), fungal dermatitis (2), splenitis (1), anized and necropsied. While no significant gross lesions myocarditis (1), endocarditis (1), periarteritis (1), pancreatitis were noted, by light microscopy there was moderate, diffuse (1), and splenitis (1). Histiocytic granulomas were seen in the necrosis of intestinal mucosal epithelial cells. A portion of small intestine, heart, and esophageal tonsils of one ETB, the formalin-fixed small intestine of one iguana was submitted small intestine of a second ETB, and in an esophageal tonsil for TEM, revealing organisms within enterocytes that had of a third ETB (Figures 10.78–10.80). Using H&E staining, developmental stages compatible with chlamydia. Polymerase the centers of many granulomas contained small basophilic chain reaction amplification of a 293-bp fragment of the 16s punctate inclusions similar to those seen in puff adders. An rRNA gene and a nested PCR targeting the variable domain immunoperoxidase staining technique with commercially IV of the ompA gene detected C. pneumoniae and C. felis in available monoclonal antibodies against the chlamydial lipoparaffin-embedded tissues from these lizards (Bodetti et al., polysaccharide antigen were used to identify the chlamydial 2002). In another retrospective study of 15 lizard cases with antigen in granulomas in the small intestine (Figure 10.81). granulomatous lesions in one or more tissues, using PCR a Additionally, macrophages within aggregates of lymphochlamydia-like organism was identified in nine cases (Soldati plasmacytic cells in the colon, small intestine and esophaet al., 2004). geal tonsils of three additional ETBs contained the antigen (Figures 10.82–10.83). Transmission electron microscopic 10.18.4 Ophidia examination of an intestinal granuloma demonstrated develUsing TEM, chlamydia-like organisms, adenovirus, and cryp- opmental stages of organisms consistent with chlamydia (Figtosporidia were identified in an Aesculapian snake (Elaphe ures 10.84–10.85). To determine the strain of chlamydia longissima) having clinical signs of gastroenteritis (Heldstab infecting these snakes, tissue samples from five frozen snakes et al., 1989). In a subsequent report, an epizootic of chla- were tested by a quantitative TaqManPCR test and a PCRmydiosis was seen in a colony of puff adders (Bitis arietans) sequence analysis test (Jacobson et al., 2004). Of 22 samples (Jacobson et al., 1989a). All snakes occasionally regurgitated tested, 9 were categorized as either positive or weakly positive mice within two days of feeding, and one snake manifested with the TaqMan test and 6 yielded an amplicon using a serial a mild respiratory disease preceding death (Figure 10.73). PCR test, which amplified a portion of the 23S rRNA gene. A At necropsy all snakes had exudate within the pericardial PCR product suitable for sequencing was obtained from the sac and two snakes had multifocal white nodules in their heart of one of the snakes. Sequence analysis showed the livers. By light microscopy there was severe granulomatous snake had been infected with Chlamydophila pneumoniae. peri-, epi-, and myocarditis (Figure 10.74), hepatitis (Fig-

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These findings showed that C. pneumoniae, a primary human pathogen, might infect emerald tree boas. In a retrospective study of 90 reptile cases with granulomatous lesions, both immunoperoxidase staining with a monoclonal antibody against chlamydial LPS antigen and PCR amplification of a small 92-bp fragment of the 23S rRNA gene were used to detect chlamydiae in these tissues (Soldati et al., 2004). Of 48 snake cases (species not identified), 4 cases were both immunoperoxidase and PCR positive for the presence of C. pneumoniae, while an additional 3 cases were positive only by PCR. Granulomas in these cases were classified as either chronic or histiocytic. Two of these snakes had a mixed infection with C. pneumoniae and a Mycobacterium other than M. tuberculosis (MOTT) in the same granulomas, whereas a second snake had a dual infection with MOTT in lung granulomas and C. pneumoniae in pericardial granulomas. Using PCR, a chlamydia-like organism was identified in an additional 26 cases.

10.19 Leptospira Leptospira has been isolated from turtles (Glosser et al., 1974; Van der Hoeden, 1968), lizards (Plesko et al., 1962), and snakes (Ferris et al., 1961; Hyakutake et al., 1980). Garter snakes (Thamnophis sirtalis) inoculated with L. interrogans developed a leptospiremia and serum agglutinins (Abdulla and Karstad, 1962). Leptospira was demonstrable in snake kidneys 6 months after inoculation, and infections were shown to persist after a 70-day period of induced hibernation. One infected snake had an interstitial nephritis. In the same study, although the European pond (Emys orbicularis), Blanding’s (Emydoidea blandingii), and common snapping (Chelydra serpentina) turtles also were inoculated, only the Blanding’s turtle developed a leptospiremia. Antibodies against Leptospira were detected in caiman (Rossetti et al., 2003) and snakes (Stanchi et al., 1986) in Argentina.

10.20 Borrelia Lyme disease is caused by several species of spirochetes within the Borrelia burgdorferi sensu lato genogroup complex. Of the 11 genospecies found worldwide, Borrelia burgdorferi sensu stricto is the cause of Lyme disease in the United States (Clark et al., 2005). Because subadult ticks, such as Ixodes scapularis in the eastern United States and Ixodes pacificus in California, can serve as vectors of Borrelia burgdorferi and selectively feed upon lizards, several studies were conducted to determine the role of lizards in the ecology of Lyme borreliosis. Studies with western fence lizards (Sceloporus occidentalis) (Lane, 1990) and southern alligator lizards (Eglaria multicarinata) (Wright et al., 1998) in California suggested that lizards do not serve as a reservoir for B. burgdorferi.

Further, the western fence lizard’s blood was found to have borreliacidal activity (Lane and Quistad, 1998). In contrast, a laboratory study with American (green) anoles (Anolis carolinensis) and southeastern five-lined skinks (Eumeces inexpectatus) showed that they could serve as reservoirs for B. burgdorferi sensu stricto (Levin et al., 1996). In a recent study, blood was collected from wild lizards in South Carolina and Florida, and although spirochetes could not be isolated from these animals, 54% of the lizards were positive for B. burgdorferi sensu lato gene sequences using a specific PCR assay. This study suggested that Lyme disease spirochetes are well established in lizards in the southeastern United States. (Clark et al., 2005).

10.21 Coxiella Coxiella burnetii is a rickettsial organism that is the cause of Q fever, an infectious disease of humans. Coxiella burnetii agglutinins were detected in sera of 13 of 53 snakes (grass snakes [Natrix natrix], Indian rat snakes [Ptyas korros], Indian cobra [Naja naja], and Indian pythons [Python molurus]) and 2 of 16 Indian roofed turtles (Kachuga sp.) (Yadav and Sethi, 1979). Coxiella burnetii was isolated from the liver and spleen of a mangrove monitor (Varanus indicus), tortoise, and python in this study.

10.22 Ehrlichia Ehrlichiosis is a disease in humans and animals caused by members of the genus Ehrlichia. Ehrlichia ruminantium (formerly Cowdria ruminantium) is a tick-borne rickettsial pathogen that is the causative agent of heartwater in cattle. Although the United States is currently free of this disease, in 1997 the African tortoise tick, Amblyomma marmoreum, was found on several species of tortoises in a private outdoor collection in central Florida (Allan et al., 1998) (Figures 12.202–12.203). Leopard tortoises from Zambia infested with the exotic African tick (Amblyomma sparsum) were also being imported into Florida. These ticks were infected with Ehrlichia ruminantium (Burridge et al., 2000a, 2000b). Subsequent studies demonstrated that leopard tortoises are refractory to E. ruminantium infection and are unlikely to introduce heartwater into new areas (Peter et al., 2001). Viperid snakes that died peracutely and acutely following importation from Africa into the United States were necropsied and found to have pulmonary disease and pericarditis as the most consistent gross lesions (Kiel et al., 2006). A cocobacillus was isolated from liver homogenates of a dead gaboon viper inoculated into viper spleen cells. Although scanning electron microscopy was reported for infected spleen cells, no TEM was conducted so the nature of the organism was not determined. PCR was conducted on infected spleen cells

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and tissues from dead snakes using three primers for pCS20 DNA; multiple products were obtained and were considered different (but related) to those produced by authenticated E. ruminantium. Although the authors believe that their data indicate a Cowdria-related attenuated species, more work is needed to conclusively determine the nature of the isolated microorganism.

10.23 Mixed and Miscellaneous Aerobic Bacterial Infections 10.23.1 Spiral-Shaped Bacterium of Rhinoceros Iguanas In a rhinoceros iguana with a bacteremia, a spiral-shaped microorganism was seen in a blood film (Figures 3.113, 6.106), and following its death, in blood vessels and sinusoids of multiple tissues (Jacobson et al., 1980) (Figures 10.86– 10.87). A subsequent TEM study of the organism in tissues revealed that it had approximately 14 flagella at each polar extremity, was covered by an electron-lucent enveloping sheath, had a discrete electron-dense cell wall, and blebs on the cell surface (Simpson et al., 1981) (Figure 6.107). These features had some similarity to those members of the bacterial family Spirillaceae.

10.23.2 Shell Disease of Aquatic Turtles While myriad shell lesions have been described in chelonians, a specific causative agent has been identified in relatively few. Some, such as SCUD (see Citrobacter above), were originally described at a time when diagnostics for certain pathogens, such as viruses and other fastidious infectious agents, were extremely limited or nonexistent. Still, these descriptive reports have value, and with the new molecular diagnostic tools that are now available (see Chapter 7) need to be revisited. A few of the more complete studies are presented below. Wallach (1975) reported an ulcerative shell disease in sliders, musk turtles, soft-shelled turtles, side-neck turtles, and painted turtles. The disease was characterized by a blotchy dark discoloration of the shell with sloughing of scutes. The bacterium, Beneckia chitinovora, was cultured from these lesions. Transmission studies were performed in painted turtles and confirmed the infectious nature of this microorganism. River cooters (Pseudemys concinna) and yellow-bellied turtles (Trachemys scripta) have been seen with a disfiguring shell disease (Garner et al., 1997; Lovich et al., 1996) (Figures 10.88–10.89). The lesions were characterized as a segmental necrosis of the epidermis and dermis with extensive remodeling of dermal bone (Garner et al., 1997) (Figure 10.90). The following bacteria were isolated from the blood of different turtles: Morganella morganii, Aeromo-

nas hydrophila, and Bacteroides sp. However, bacteria were not identified in the acute lesions in the skin. Endoparasitism with spirorchiid trematodes and necrotizing pyogranulomatous pancreatitis also was seen. The specific cause of the shell lesion could not be determined.

10.23.3 Subcutaneous and Tissue Abscesses and Masses 10.23.3.1 Chelonia  Aural abscesses (middle and inner ear infections) have been seen in green turtles (Chelonia mydas), different species of freshwater turtles, captive tortoises, and most commonly in box turtles (Terrapene spp.) (Graham-Jones, 1961; Jackson et al., 1972; Keymer, 1978a, b). Box turtles are usually presented with unilateral or bilateral swellings below the tympanic scale that consist of caseous laminar material surrounded by a mixed inflammatory reaction (Jacobson, 1981) (Figure 10.91). A variety of bacteria including Aeromonas hydrophila, Citrobacter, Corynebacterium sp., Enterobacter, Escherichia coli, Morganella morganii, Pantoea agglomerans, Proteus vulgaris, and Pseudomonas sp. were cultured from these lesions (Willer et al., 2003). In a recent study, no single bacterium was found to be the causative agent (Joyner et al., 2006). While the route of infection probably involves retrograde migration of microorganisms up the eustachian tube, the pathogenesis of the disease is poorly understood. Organochlorine contaminants have been implicated as possible causes of these abscesses (Holladay et al., 2001; Tangredi and Evans, 1997). Of 70 Greek tortoises necropsied, 5 were found to have abscesses; 3 had abscesses on the neck and 2 on a hindlimb (Holt et al., 1979). Microbial isolation in two cases resulted in growth of Escherichia coli and Corynebacterium murium. As mentioned previously, Serratia marcescens was cultured from subcutaneous abscesses in a gopher tortoise, and following euthanasia multiple abscesses were found in the lung, kidney, and liver (Pye et al., 1999). A Burmese mountain tortoise (Manouria emys) with a necrotic plastronal lesion had a fistulous tract continuous with a large abscess in the coelomic cavity. An anorexic and depressed adult female desert tortoise was found to have a large radio-dense mass in the area of the right hepatic lobe (Berschauer and Mader, 1998). The tortoise was eventually euthanized and a solid encapsulated abscess was found in the right hepatic lobe. Corynebacterium sp. was cultured from the abscess. Abscesses have been seen on the forelimbs of green turtles in a sea turtle farming operation (Figure 10.92). This probably resulted from secondary bacterial infection of a traumatized area of the integument. 10.23.3.2 Squamata  Cutaneous, subcutaneous, and internal abscesses are more commonly reported in lizards and snakes (Jacobson, 1991; Maxwell, 2003; Russo, 1987) (Figures 10.93–10.97) than in other groups of reptiles. These lesions generally consist of caseous necrotic material (lysed

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heterophils) surrounded by granulomatous inflammation (Figure 10.98). Some lesions may have diffuse cellulitis (Figure 10.99). Various bacteria have been isolated from these lesions, including Bacterium sauromali, Corynebacterium spp., Fusobacterium necrophorum, Micrococcus spp., Morganella morganii, Neisseria iguanae, Salmonella spp., Serratia anolium, and Serratia marcescens. Often these are mixed infections, with no single bacterium identified as the causative agent. Occasionally a single bacterial species is identified. Serratia marcescens was isolated from subcutaneous abscesses in a green iguana and a spiny-tailed iguana (Ctenosaura acanthura), Salmonella Marina from a second spiny-tailed iguana, and Micrococcus from a second green iguana (Boam et al., 1970a). Corynebacterium sp. was cultured from a large rostral abscess from a green iguana, and Pseudomonas sp. was cultured from a coelomic mass of an Indonesian blue-tongued skink (Tiliqua gigas). In the latter case, a fistulous tract extended from a skin lesion, through the body wall, to the coelomic mass and apparently disseminated to the eye, causing uveitis and hypopyon (Millichamp et al., 1983). Anaerobic bacteria have also been isolated from abscesses and include Bacteroides, Fusobacterium, Clostridium, and Peptostreptococcus (Stewart, 1990). Dermatophilus has been identified in subcutaneous masses in a boa constrictor (Jacobson, 1989) and a king cobra (Wellehan et al., 2004a).   The author has seen numerous snakes with either impacted, abscessed, or fistulated anal glands. Anal glands in snakes are located immediately caudal to the cloaca, and in males, dorsal to the hemipenes. Mixed Gram-negative bacteria and fungi have been identified in infected anal glands. A client who decided to breed a large number of kingsnakes at the same time in the same cage, subsequently noted multiple pericloacal subcutaneous abscesses and impacted and abscessed anal glands in most of the snakes (Figures 10.100–10.101). This had the appearance of a sexually transmitted disease. While a variety of Gram-negative bacteria were isolated from these lesions, a specific pathogen was not identified.

10.23.4 Bacterial Infections of the Eye and Orbit Bacterial infections of the eye, adjacent tissues, and subspectacular space (in certain lizards and almost all snakes) are common in reptiles. Mycobacterium was identified in a large periocular mass in a box turtle that was in captivity for 24 years (Leonard and Shields, 1970). The mass completely covered the lateral surface of the globe and displaced it medially. Histologically it was composed of granulomatous inflammation with acid-fast bacilli in multinucleated giant cells. The author has seen Mycobacterium sp. in granulomatous palpebral infections of hatchling Atlantic (Kemp’s) ridley sea turtles in aquaculture. In those lizards with eyelids, heterophils may accumulate under or adjacent to a palpebra, resulting in a large periorbital nodular abscess. This has been seen in

green iguanas (Figure 10.102), chameleons (Figures 10.8, 10.103), and other lizards (Millichamp et al., 1983; Schumacher et al., 1996). Escherichia coli was isolated from an abscess of the lower palpebra of a desert iguana (Dipsosaurus dorsalis) that obscured the entire orbit. Bacterial conjunctivitis associated with Aeromonas liquefaciens was reported in a colony of lacertid lizards (Cooper et al., 1980). Pseudomonas was isolated from American anoles (Anolis carolinensis) with conjunctivitis and blepharitis (Millichamp et al., 1983). Tortoises with mycoplasmosis can manifest palpebral edema and conjunctivitis (Jacobson et al., 1991a). Hypopyon was reported in one of four tokay geckos that had bacterial pneumonia with Klebsiella pneumoniae (Bonney et al., 1978), in American alligators with Aeromonas hydrophila septicemia, and in an Indonesian blue-tongued skink (Tiliqua gigas) with a large coelomic mass from which Pseudomonas was cultured. In snakes and those lizards having spectacles, foreign body penetration of the spectacle and retrograde migration of bacteria from the oral cavity (especially in those animals with infections of the oral cavity) to the subspectacular space by way of the nasolacrimal duct may result in heterophils migrating into the space and distension (Figure 10.104). Bacteria isolated from this material include Aeromonas, Proteus, and Pseudomonas.

10.23.5 Stomatitis, Gingivitis, and Pharyngitis 10.23.5.1 Chelonia  Stomatitis and pharyngitis has been seen in chelonians (Figure 10.105). In Greek tortoises, gross oral hemorrhage was reported (Holt and Cooper, 1976). In a subsequent study of seventy tortoises (primarily Greek tortoises), 14 had necrotic stomatitis (Holt et al., 1979). Lesions were found on dorsal or lateral aspects of the tongue (7 cases), the tongue and caudolateral hard palate (4 cases), the tongue hard palate and pharynx (2 cases), and the anterior floor of the mouth (1 case). Acinetobacter spp., Beta-hemolytic Staphylococcus spp., Enterobacter spp., Escherichia coli, Klebsiella spp., Pasteurella spp., and Pseudomonas spp. have been isolated from tortoises with stomatitis. Both herpesvirus and iridovirus (Chapter 9) need to be ruled out as the primary pathogens in tortoises and box turtles with oral lesions.   An ulcerative stomatitis-obstructive rhinitis-pneumonia disease complex was reported in hatchling and juvenile farmed and oceanarium-reared green turtles and loggerhead sea turtles in Australia (Glazebrook et al., 1993). The obstructive rhinitis was considered secondary to stomatitis. The following three bacteria were isolated from these cases: Aeromonas hydrophila, Flavobacterium sp., and Vibrio alginolyticus. Pneumonia was also seen and was considered a consequence of aspiration of necrotic material from the oral cavity. In addition to the bacteria cultured from the oral lesions, the following fungi were isolated from the respiratory tract of turtles with pneumonia: Aspergillus sp., Fusarium sp., Paecilomyces sp., and Penicillium sp.

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10.23.5.2 Squamata  While stomatitis and gingivitis have been reported in different species of lizards (McCracken and Birch, 1994) (Figure 10.1), primary bacterial infections of the oral cavity are uncommon. In many situations, the lesions and associated Gram-negative bacteria are secondary to trauma and poor husbandry. Hypovitaminosis A is common in several species of Old World chameleons (Ferguson et al., 1996), and secondary metaplastic changes in the oral cavity may predispose chameleons to Gram-negative infections. Stomatitis in veiled chameleons (Chamaeleo calyptratus) usually involves the mucous membranes adjacent to the labial scales, the commissures of the oral cavity, or sometimes the tongue (Stahl, 1997) (Figure 10.106). Acrodont dentition may predispose them to gingival infection and osteomyelitis. Chameleons have specialized glands in the lateral commissures of the oral cavity (Coke, 1999), and abscesses at this site are seen and are typically infected with mixed Gramnegative bacteria (Figure 10.107). Recently, rough scaled (Gerrhosaurus major) and black-lined (G. nigrolineatus) plated lizards (Wellehan et al., 2004b) and green tree monitors (Wellehan et al., 2004c) with oral lesions were infected with herpesvirus (see Chapter 9). Thus, viruses need to be considered when determining the cause of stomatitis in lizards and other reptiles.   Historically, ulcerative stomatitis was a common health problem of snakes. The initial damage to the oral mucosa may progress to an osteomyelitis of the bony structures of the head (Figure 10.108). The infection is initially characterized by ulceration of the mucous membranes surrounding the maxillary, dentary, and palatine teeth, with an accumulation of caseous material within the mucous membranes. Next, there may be a bacterial pneumonia via inhalation of cellular debris into the respiratory tract (Figure 10.109) or systemic infection (Figure 10.110) via the circulatory system. While recently acquired snakes may have a preponderance of Gram-positive bacteria in their oral cavity, it appears that with time in captivity, Gram-negative microorganisms become established in the oral cavity. Under the right conditions, many of these organisms have the potential to become secondary invaders. Snake’s rubbing on objects in the cage or on screened tops, bites from rats and mice, poor nutrition, and improper environmental temperatures may all be predisposing factors. The most commonly isolated aerobic bacteria from oral lesions of snakes are Aeromonas, Escherichia coli, Morganella, Proteus, Providencia, Pseudomonas, and Salmonella.

10.23.6 Pneumonia 10.23.6.1 Chelonia  While there are few reports of primary bacterial pneumonia in chelonians, a number of box turtles have been seen with clinical signs of respiratory disease. In Illinois, chronic bacterial pneumonia was diagnosed in two wild eastern box turtles that were found half bur-

ied in a dry creek bed; they were severely emaciated and depressed (Evans, 1983). Histologically, chronic inflammation was identified in the nasal sinuses and lungs. A mixture of Gram-negative bacteria including Acinetobacter calcoaceticus, Morganella morganii, Pseudomonas sp., and Serratia marcescens.were isolated from both turtles; no other pathogens were identified.   Iridovirus and herpesvirus should be considered in the differential diagnosis when working up chelonians with signs of pharyngitis and respiratory tract disease (see Chapter 9).

10.23.6.2 Squamata  The author has seen relatively few cases of Gram-negative bacterial pneumonia in lizards as compared to snakes. However, others have reported bacterial infections (Aeromonas, Klebsiella, Pseudomonas) of the respiratory tract to be common in veiled chameleons (Stahl, 1997). As in snakes, pneumonia may follow chronic infections of the oral cavity. Caiman lizards (Draecena guianensis) with severe pneumonia were infected with a paramyxovirus (Jacobson et al., 2001a) (see Chapter 9). Thus, viruses need to be in the differential diagnosis when working up pneumonias of lizards as well as other reptiles.   Diseases of the oral cavity and respiratory system are common in snakes, and at one time were responsible for much of the mortality in snake collections. Over a 5-year period, 100% of oropharyngeal cultures from recently imported pythons and boid snakes at the Institute for Herpetological Research were culture positive for Pseudomonas (Ross and Marzec, 1984). Other isolates included Acinetobacter, Aeromonas, Citrobacter, Enterobacter, Flavobacter, Klebsiella, Proteus, Providencia, and Serratia. In a prospective study undertaken to determine the bacterial flora of the respiratory tract of eight snakes (three boa constrictors, three Indian pythons, and two reticulated pythons) initially identified as healthy revealed Providencia rettgeri to be the most common aerobe isolated from glottal swabs (Hilf et al., 1990). Coagulase-negative Staphylococcus sp. was isolated from all 8 snakes. One snake developed pneumonia before baseline bacterial cultures could be obtained; Aeromonas hydrophila, Alcaligenes odorans, Citrobacter sp., Morganella morganii, and Proteus mirabilis were cultured. The snake died 2 weeks later and a pure growth of Aeromonas hydrophila was obtained from the lung. The second snake developed pneumonia 7 months after the study began, and Salmonella was cultured from a glottal swab. Approximately 2 months later, the snake died, and at necropsy the following bacteria were isolated from the lung: Citrobacter sp., Enterococcus sp., and Salmonella sp. A pulmonary disease (Figures 10.111–10.113) was reported in two private breeding collections of adult Burmese pythons in Florida (Heard, 1997; Jacobson et al., 2001b). A similar disease has been seen in Burmese pythons elsewhere in the United States and Europe. The pneumonia was proliferative and the inflammation in the interstitium was variable (Figures 10.114–10.115). In the same collections, Burmese pythons with a thromboembolic disease (TED) were also

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seen. The index case involved a snake with pitting edema of the head and oral cavity (Figure 10.116). It subsequently was found to have a large septic thrombus in the sinus venosus (Figure 10.117), from which both Corynebacterium sp. and Salmonella arizonae were cultured (Jacobson et al., 1991b). Additional Burmese pythons with similar lesions were seen (Heard, 1997; Jacobson et al., 2001b) (Figure 10.118). At a light microscopic level, there were arteritis and thrombosis of major vessels (Figure 10.119). Other snakes developed necrosis of the distal tail (Figures 10.120–10.121). This resulted from thrombi showering to the tail where the caudal vessels were occluded, resulting in ischemic necrosis (Figure 10.122). Several of the snakes with TED also had thrombi in the lungs (Figure 10.123). Aerobic cultures of the respiratory tract of affected Burmese pythons resulted in the culture of a variety of Gramnegative bacteria. Using PCR, nucleotide sequences for Mycoplasma were identified in the trachea and lung of two affected snakes. However, Mycoplasma was not cultured from these cases. In a separate report, one of several Burmese pythons with tracheitis and pneumonia from a private collection was found by electron microscopy to have Mycoplasma sp. closely associated with cells lining air passageways (Penner et al., 1997) (Figure 6.103). While Mycoplasma was originally isolated from lung tissue of the affected snake, it was a slow grower and could not be maintained in culture, and the role of Mycoplasma in this chronic respiratory disease of Burmese pythons remains unclear. Paraffin-embedded lung tissue of two snakes with proliferative pneumonia was evaluated for the presence of chlamydial DNA using a PCR assay that amplified a 293-bp fragment of the 16s rRNA gene and a nested PCR targeting the variable domain IV of the ompA gene. Both C. abortus and C. pneumoniae were identified in the lungs of these snakes (Bodetti et al., 2002). The role of these microorganisms in this disease process remains unclear.

10.23.7 Bacteremia and Osteomyelitis 10.23.7.1 Chelonia  Of four green turtles (Chelonia mydas) with severe spirorchiidiasis, disseminated Gram-negative bacterial infections, including Citrobacter freundii, Escherichia coli, Moraxella sp., and Salmonella, were cultured from the liver, lung, or kidney of three turtles (Raidal et al., 1998). Severe, multifocal granulomatous vasculitis and microabscesses were seen in the intestines, kidney, liver, lung, and brain. Green turtles in Hawaii with fibropapillomas (FP) were found to be bacteremic, with four species of Vibrio spp. representing a majority of the bacteria cultured (Work et al., 2003). The percentage of bacteremic turtles increased with the severity of FP. Green turtles with severe FP may have been immunocompromised and thus more susceptible to bacteremia.

10.23.7.2 Crocodylia  Aeromonas hydrophila is often cultured from American alligators with septicemia. In a eutrophic lake in Florida, Aeromonas hydrophila and A. shigelloides were incriminated in the deaths of American alligators (Shotts et al., 1972). In a survey of alligators in the southeastern United States, A. hydrophila was isolated from the oral cavity and internal tissues of 85% and 70%, respectively, of 123 alligators captured from 5 locations (Gorden et al., 1979). Aeromonas was isolated from the liver and kidney of one of 6 alligators with uveitis and hypopyon (Millichamp et al., 1983). Four ill, lethargic American alligators with dermatitis were found to be septic (Novak and Seigel, 1986). One of the alligators exhibited lethargy and excessive basking behavior prior to death. Skin wounds including subcutaneous abscesses were attributed to fighting; one extended into the right shoulder joint. Citrobacter freundii, Enterobacter agglomerans, Klebsiella oxytoca, Morganella morganii, Proteus sp., and Serratia marcescens were cultured from wounds or blood of these animals. Bacterial infections were commonly diagnosed in hatchling crocodiles in Zimbabwe, and were either localized or the cause of septicemia (Foggin, 1987). Histologically there was often a purulent pneumonia and hepatic necrosis.

10.23.7.3 Squamata  Bacteremias have been seen in numerous species of lizards and snakes, with bacteria probably showering from the gastrointestinal tract to viscera, long bones, and vertebrae. As mentioned previously, Neisseria iguanae, while existing as a commensal, can also result in chronic abscesses and septic lesions in green iguanas and rhinoceros iguanas (Barrett et al., 1994; Plowman et al., 1987). The author evaluated a green iguana with a proliferative lesion of the hyoid cartilage in the dewlap from which Neisseria was isolated. Multifocal osteoarthritis (Figures 10.124­–10.125) was seen by the author in green iguanas in a farming operation in Panama in the mid-1980s. The lizards were collected from the wild as adults for egg collection and establishing a long-term breeding colony. Many of these lizards developed boney lesions several weeks after capture, from which mixed Gram-negative bacteria were cultured. This was prior to the identification of Neisseria iguanae in the green iguana (Plowman et al., 1987). Possibly this bacterium was missed and was the causative agent. Escherichia coli was cultured from the liver and heart blood of an adult black-necked spitting cobra (Naja nigricollis) that was found dead in its cage, and at necropsy had extensive subcutaneous hemorrhage and edema involving the middle third of its body (Russell and Herman, 1970). Salmonella enterica arizonae (formerly Arizona hinshawii) septicemia was reported in three boa constrictors that died within a 30-day period (Boever and Williams, 1975). Lesions seen in these snakes included an intraorbital abscess (1), stomatitis (1), pneumonia (1), enteritis (3), and multiple visceral abscesses (2). Salmonella enterica arizonae was cultured from the liver and abdominal abscesses of one snake. Fiftyone of 58 blood cultures made from 28 healthy lizards from

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8 species resulted in bacterial growth. The most common isolates were Clostridium difficile, C. innocuum, C. perfringens, and C. sordelli, (Hanel et al., 1999). Percutaneous blood samples were obtained from the ventral coccygeal vein, and although the collection site was cleansed, contamination with feces or soil from the cage of these lizards may have resulted in contaminated samples. Whereas Clostridium perfringens was cultured from the blood of a rough-necked monitor (Varanus rudicollis) with spondylosis, C. sordelii was cultured from a vertebral aspirate of the same snake (D’Agostino et al., 2006). If the vertebrae were colonized from the hematogenous route, one would expect the same organism to be cultured from both sites. Possibly, Clostridium perfringens in the blood overgrew C. sordelii. Collecting a truly uncontaminated antemortem blood sample from an animal is far more difficult than it appears to be. A variety of lytic and proliferative vertebral diseases have been seen in snakes (Figures 5.44, 10.126–10.127). Early reports described histological changes in some cases, which were interpreted as similar to those seen with osteitis deformans (Paget’s bone disease) of humans. In some reports of osteitis deformans in snakes, no infectious agent could be demonstrated (Frye and Carney, 1974), while in others there was an association with bacterial infections (Frye and Kiel, 1985; Kiel, 1983). Two southern copperheads (Agkistrodon contortrix contortrix) developed osteoarthritic vertebral lesions compatible with those seen radiographically for osteitis deformans after resolution (22 to 36 months) of a severe septicemia (Kiel, 1977b). In a report that compared 15 snakes with segmental, proliferative osteoarthritis of vertebral bodies, 3 distinct histologic groups were defined (Isaza et al., 2000). One group had evidence of active bacterial osteoarthritis (Figure 10.128), with Salmonella cultured from the bones of 2 of 5 snakes in this group. In a second group, there was noninflammatory osteoathrosis mixed with small multifocal areas of inflammation (Figure 10.129). While there was no histological evidence of bacteria, bacteria were isolated from affected bone and joint lesions suggesting a low-grade osteoarthritis. A third group had degenerative osteoarthrosis and ankylosis with minimal or no inflammation (Figure 10.130). Bacteria were not isolated from the bone or blood of these snakes. The lesions in this group resembled those seen in osteitis deformans. However, the mosaic bone pattern seen in osteitis deformans was noted by one of the authors as present in the normal bone of snakes (see Section 5.5.3, Figure 5.45) The lesions in the third group also resembled disseminated idiopathic skeletal hyperostosis (DISH) in humans and dogs. However, intra-articular bony fusion seen in these snakes is not seen in DISH. These lesions may form a continuum from early active osteoarthritis to chronic noninfectious proliferation. In a five-year prospective study in a colony of ridgenose rattlesnakes (Crotalus willardi), many had a Salmonella enterica arizonae–associated osteomyelitis (Ramsay et al., 2002) (Figure 10.12). Most of the snakes that were diagnosed with osteomyelitis at the beginning of the study

subsequently died or were euthanized. The major serotype identified was S. arizonae serotype 56:Z4,Z23. Enterococcus sp. was cultured from the bone of a rhinoceros viper (Bitis nasicornis) with severe osteomyelitis (Schrötter et al., 2005). In general, bacteria cultured from infected bones of snakes with osteomyelitis are often cultured from their blood (Isaza et al., 2000).

10.23.8 Miscellaneous Bacterial Infections 10.23.8.1 Chelonia  A variety of bacteria have been isolated from various lesions in sea turtles in aquaculture facilities. From 1977 to 1983, Kemp’s ridley and loggerhead sea turtles were reared at the U.S. National Marine Fisheries Service in Galveston, TX. Diseases, malformations, and injures in turtles in this group were classified in 27 categories, several of which were associated with certain bacteria (Leong at al., 1989). A sudden hatchling death syndrome was seen with a wide variety of bacteria, including Clostridium bifermentans and Vibrio alginolyticus, isolated from affected turtles. A papillary dermatitis affected the skin around the eyes and anus of turtles, and on the limbs and plastron. Bacteria isolated from these lesions included Aeromonas formicans, Vibrio alginolyticus, V. aglosus, and Pseudomonas sp. The bacteria cultured from lesions of turtles with focal erosive dermatitis included Aeromonas formicans, Citrobacter freundii, Vibrio alginolyticus, and V. anguillarum. Vibrio parahemolyticus was cultured from a turtle with hemorrhagic bacteriosis, a condition in which bleeding was seen from tissues and organs. In a survey of farmed green turtles (102) and hawksbill turtles (2) in Australia, four microorganisms (Aeromonas hydrophila, Cytophaga-Flavobacterium sp., Pseudomonas sp., and Vibrio alginolyticus) were isolated repeatedly from cases of traumatic ulcerative dermatitis, ulcerative stomatitis, obstructive rhinitis, and bronchopneumonia (Glazebrook and Campbell, 1990a). Pseudomonas aeruginosa and Flavobacterium sp. were isolated from ulcerative lesions of the eyelids where they invaded traumatized tissues. Pseudomonas sp. and Vibrio alginolyticus were isolated from turtles with an ulcerative shell disease. For the most part, bacterial diseases of oceanarium-reared green and hawksbill sea turtles were the same as farmed turtles (Glazebrook and Campbell, 1990b). Lesions of the gastrointestinal tract found in 84 of 136 sea turtles (128 loggerheads and 4 green turtles) in the Canary Islands included ulcerative and necropurulent stomatitis, ulcerative and fibrinous esophagitis, esophageal perforation, necropurulent gastritis, and catarrhal, fibrinous, necrotizing, and necropurulent enteritis; necrotizing enteritis was associated with intussusception caused by the ingestion of monofilament line (Oros et al., 2004). Bacteria isolated from these lesions included Bacillus sp., Escherichia coli, Pasteurella sp., Proteus sp., Staphylococcus sp., Streptococcus sp., and Vibrio alginolyticus. Twenty-nine turtles had necrotizing or granulomatous hepatitis associated with Aeromonas hydrophila,

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Citrobacter sp., E. coli, Proteus sp., Staphylococcus sp., and Vibrio alginolyticus.

10.23.8.2 Crocodylia  Escherichia coli was cultured from multiple tissues of an American alligator that was found dead in a pool that it shared with 24 other alligators (Russell and Herman, 1970). There was a purulent exudate throughout edematous lungs. Several 3- to 4-year-old mugger crocodiles (Crocodylus palustris) at a farm in India that died with an hemorrhagic enteritis were found by culture (intestines, heart blood, and pericardial fluid) to be infected with Escherichia coli (Sinha et al., 1988). Of 54 young salt water and New Guinea (C. novaeguineae) crocodiles (and two unrecorded species) that died at a crocodile farm in Papua New Guinea, 16 were infected with Aeromonas hydrophila, Chromobacterium sp., and Salmonella arizonae (Ladds and Sims, 1990). In contrast, of 38 farm-reared crocodiles in Irian Jaya, in only 2 cases was evidence of bacterial pneumonia and septicemia found (Ladds et al., 1995). Microscopic evaluation of skin lesions from crocodiles on 9 farms in Australia indicated that Dermatophilus sp. and Mycobacterium sp. accounted for 28.1% and 2.5% of the cases, respectively (Buenviaje et al., 1998). In another study evaluating diseases in crocodiles at 7 farms in Queensland and the Northern Territory of Australia, hepatitis and septicemia were found at several farms (Buenviaje et al., 1994). The following bacteria were isolated in these cases: Aeromonas hydrophila, Clostridium septicum, Edwardsiella sp., Klebsiella sp., Proteus rettgeri, Salmonella sp., and Serratia liquefaciens. Interdigital subcutaneous emphysema (a condition called bubble foot) was diagnosed in 8 hatchling saltwater crocodiles on several farms in Queensland, Australia (Turton et al., 1996). Bacteria identified in or cultured from the lesions included Citronella freundii, Enteroeba agglomerans, Mycobacterium, Providencia rettgeri, and Pseudomonas putida. Poxvirus-like inclusion bodies were seen in two cases in the skin of the affected foot, and the authors speculated that there might be a causal relationship between this virus and the presence of secondary invaders. Providencia rettgeri septicemia and meningitis was described over a 3-year period in hatchling crocodiles at one farm in Australia (Ladds et al., 1996). Of 62 saltwater crocodiles necropsied at a commercial crocodile farm in northern Australia, bacteria were isolated most commonly from the liver, heart, and lung, and included Edwardsiella sp., Enterobacter agglomerans, Morganella morganii, Pasteurella multocida, Providencia rettgeri, Pseudomonas sp., and Salmonella spp. (Hibberd et al., 1996). The author has seen oomphalitis (inflammation of yolk) in several species of crocodilians. The abdominal yolk fails to reabsorb and becomes firm and enlarges as inflammatory cells are deposited within and around the mass. They are commonly infected with Gram-negative bacteria such as E. coli and S. enterica arizonae.

10.23.8.3 Squamata  Bacterial encephalitis is uncommonly seen in lizards and snakes. As previously mentioned, an adult male bearded dragon that died following a 3-day period of a nonspecific illness was found by light microscopy to have microabscesses in multiple tissues including the brain (Girling and Fraser, 2004). Listeria monocytogenes was cultured from multiple tissues and heart blood. A rosy boa (Lichanura trivirgata roseofusca) that exhibited signs of neurologic disease was necropsied and found to have meningoencephalitis, choroiditis, and ventriculitis; bacteria were seen within heterophilic granulomas in the brain (John Roberts, personal communication) (Figures 10.131–10.132). Klebsiella oxytoca and a hemolytic Staphylococcus sp. were cultured from the brain (Eric Baitchman, personal communication). Using light microscopy, a corn snake (Elaphe guttata guttata) that had a head tilt was observed to have a focal area of necrosis and an inflammatory infiltrate in nerve VIII (Figures 10.133–10.134). This snake also had adenoviral enteritis. The author has seen oomphalitis (inflammation of yolk) in both viviparous and oviparous snakes. More than likely it also occurs in lizards. In affected animals the abdominal yolk fails to reabsorb, becomes firm, and enlarges as inflammatory cells are deposited within and around the mass (Figures 10.135 –10.136). The yolk needs to be surgically removed as soon as possible since infection with Gram-negative bacteria such as S. enterica arizonae and E. coli is common. Possibly, there is vertical infection since transovarian transmission in snakes has been reported (Chiodini, 1982).

Acknowledgments The author thanks many colleagues for providing images and papers not easily obtained in the United States. Those providing images are acknowledged in the figure legends for this chapter. Many of the cases presented in this chapter represent client-owned animals submitted to the Zoological Medicine Service, College of Veterinary Medicine, University of Florida, Gainesville. Many cases resulted in research projects designed to better understand the pathogenesis of specific diseases.

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Mohan K, Foggin CM, Muvavariwa P, Honywill J, and Pawandiwa A. 1995. Mycoplasma-associated polyarthritis in farmed crocodiles (Crocodylus niloticus) in Zimbabwe. Onderstepoort J Vet Res 62:45–49. Mohan K, Foggin CM, Muvavariwa P, and Honywill J. 1996. Is mycoplasma-associated polyarthritis in farmed crocodiles a vertically transmitted disease? in Proceedings of the 13th Work Meeting of the Crocodile Specialist Group, IUCN — The World Conservation Union, Gland, Switzerland, 299–302. Montali, R. J., Smith, E. E., Davenport, M., and Bush, M. 1975. Dermatophilosis in Australian bearded lizard. J Amer Vet Med Assoc 167:553–555. Morales P and Dunker F. 2001. Fish tuberculosis, Mycobacterium marinum, in a group of Egyptian spiny-tailed lizards, Uromastyx aegyptius. J Herp Med Surg 11:27–30. Newton J. 1992. Brown spot disease in the Louisiana alligator industry: what we know about the disease and possible control protocols, in Proceedings of the Louisiana Aquaculture Conference, Baton Rouge, LA, 46–47. Norwinski RJ. 2001. Iguana-transmitted Salmonella osteomyelitis. Orthopedics 24:694. Novak SS and Seigel RA. 1986. Gram-negative septicemia in American alligators (Alligator mississippiensis). J Wildl Dis 22:484–487. Obwolo MJ and Zwart P. 1993. Prevalence of Salmonella in the intestinal tracts of farm-reared crocodiles (Crocodylus niloticus) in Zimbabwe. J Zoo Wildl Med 24:175–176. Olson GA and Woodard JC. 1974. Miliary tuberculosis in a reticulated python. J Amer Vet Med Assoc 164:733–735. Onderka DK and Finlayson MC. 1985. Salmonellae and salmonellosis in captive reptiles. Can J Com Med 49:268–270. Origgi F, Roccabianca P, and Gelmetti D. 1999. Dermatophilosis in Furcifer (Chamaeleo) pardalis. Bull Assoc Rept Amphib Vet 9:9–12. Orós J, Rodriguez JL, Herráez P, Santana P, and Fernández A. 1996. Respiratory and digestive lesions caused by Salmonella arizonae in two snakes. J Comp Path 115:185–189. Oros J, Acosta B, Gaskin JM, Deniz S, and Jensen HE. 2003. Mycobacterium kansasii infection in a Chinese soft shell turtle (Pelodiscus sinensis). Vet Rec 152:474–476. Oros J, Calabuig P, and Deniz S. 2004. Digestive pathology of sea turtles stranded in the Canary Islands between 1993 and 2001. Vet Rec 155:169–174. Otis VS and Behler JL. 1973. The occurrence of Salmonellae and Edwardsiella in the turtles of the New York Zoological Park. J Wildlf Dis 9:4–6. Page LA. 1961. Experimental ulcerative stomatitis in king snakes. Cornell Vet 51:258–266. Page LA. 1966. Diseases and infections of snakes: A review. Bull Wildl Dis Assoc 2:111–126. Parrish HM, MacLaurin AW, and Tuttle RL. 1956. North American pit vipers: Bacterial flora of the mouths and venom glands. Virg Med Monthly 83:383–385. Pasmans F, De Herdt P, and Haesebrouck F. 2002a. Presence of Salmonella infections in freshwater turtles. Vet Rec 150:692–693. Pasmans F, De Herdt P, and Haesebrouck F. 2002b. Pathogenesis of infections with Salmonella enterica subsp. enterica serovar Muenchen in the turtle Trachemys scripta elegans. Vet Micro 87:315–325. Pasmans F, Van Immerseel F, Van den Broeck W, Bottreau E, Velge P, Ducatelle R, and Haesebrouck F. 2003. Interactions of Salmonella enterica subsp. enterica serovar Muenchen with intestinal explants of the turtle Trachemys scripta scripta. J Comp Pathol 128:119–126.

Peter TF, Mahan SM, Burridge MJ. 2001. Resistance of leopard tortoises and helmeted guineafowl to Cowdria ruminantium infection (heartwater). Vet Parasitol 98:299-307. Peters DK and Cardeilhac PT. 1988. Isolation of Aeromonas hydrophila during an outbreak of hatchling alligator syndrome (HAS), in Proceedings of the International Association of Aquatic Animal Medicine, Toronto, Ontario, 19:86–88. Penner JD, Jacobson ER, Brown DR, Adams HP, and Besch-Williford CL. 1997. A novel Mycoplasma sp. associated with proliferative tracheitis and pneumonia in a Burmese python (Python molurus bivittatus). J Comp Path 17:283–288. Plesko L, Janovicova E, and Lac J. 1962. Beitrag zur Bedeutung von Kaltblutern fur die Zirkulation der Leptospiren in der Natur. Zentralbl Bakteriol Parasitenkd Hyg Abt I Orig A 192:482–484. Plowman C, Montali R, Phillips L, Schlater L, and Lowenstine L. 1987. Septicemia and chronic abscesses in iguanas (Cyclura cornuta and Iguana iguana) associated with a Neisseria species. J Zoo Anim Med 18:86–93. Posthaus H, Bacciarini LN, Pagan O, Krampe M, and Heldstab A. 1997. Causes of mortality in hawksbill turtles (Eretmochelys imbricata) at the Basle Zoo, in Internationalen Symposiums uber die Erkrankungen der Zoo- und Wildtiere, Verhandlungsbericht des 38, Zurich, Switzerland, 39–42. Pye GW, Jacobson ER, Newell SM, Scase T, Heard DJ, and Dennis PM. 1999. Serratia marcescens infection in a gopher tortoise, Gopherus polyphemus, and use of magnetic resonance imaging in diagnosing systemic disease. Bull Assoc Rept Amphib Vet 9:8–11. Pye GW, Brown DR, Nogueira MF, Vliet KA, Schoeb TR, Jacobson ER, and Bennett RA. 2001. Experimental inoculation of Mycoplasma alligatoris in broad nosed caiman (Caiman latirostris) and Siamese crocodiles (Crocodylus siamensis). J Zoo Wildl Med 32:196–201. Quesenberry KE, Jacobson ER, Allen JL, and Cooley AJ. 1986. Ulcerative stomatitis and subcutaneous granulomas caused by Mycobacterium chelonei in a boa constrictor. J Amer Vet Med Assoc 189:1131–1132. Raidal SR, Ohara M, Hobbs RP, and Prince RI. 1998. Gram-negative bacterial infections and cardiovascular parasitism in green sea turtles (Chelonia mydas). Austral Vet J 76:415–417. Ramsay EC, Bemis DA, and Patton CS. 1998. Dermatophilus infections in a tortoise collection, in Proceedings of the American Association of Zoo Veterinarians and the American Association of Wildlife Veterinarians Joint Conference, Omaha, NE, 279–280. Ramsay EC, Daniel GB, Tryon BW, Merryman JI, Morris PJ, and Bemis DA. 2002. Osteomyelitis associated with Salmonella enterica ss arizonae in a colony of ridgenose rattlesnakes (Crotalus willardi). J Zoo Wildl Med 33:301–310. Rhodin AGJ and Anver MR. 1977. Mycobacteriosis in turtles: cutaneous and hepatosplenic involvement in a Phrynops hilari. J Wildl Dis 13:180–183. Richards JM, Brown JD, Kelly TR, Fountain AL, and Sleeman JM. 2004. Absence of detectable Salmonella cloacal shedding in free-living reptiles on admission to the Wildlife Center of Virginia. J Zoo Wildl Med 35:562–563. Ross RA and Marzec G. 1984. The Bacterial Diseases of Reptiles. Institute for Herpetological Research, Stanford, CA. Rossetti CA, Uhart M, Romero GN, and Prado W. 2003. Detection of leptospiral antibodies in caimans from the Argentinean Chaco. Vet Rec 153:632–633. Runyon EH. 1970. Identification of mycobacterial pathogens utilizing colony characteristics. Am J Clin Pathol 54:578–586.

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Russell WC and Herman KL. 1970. Colibacillosis in captive wild animals. J Zoo Anim Med 1:17–21 Russo EA. 1987. Diagnosis and treatment of lumps and bumps in snakes. Comp Cont Educ Pract Vet 9:795–802. Ryan TP. 1992. Dermatophilosis in a savannah monitor (Varanus exanthematicus). Bull Reptil Amphib Vet 2:7. Schildger BJ, Weib R, Frank H, and Wicker R. 1991. Mycobacteriosis in Australian snake-neck turtles (Chelodina longicollis), in Proceedings of the 4th International Colloquium on Pathology and Medicine of Reptiles and Amphibians, Bad Nauheim, Germany, 62–67. Schroder HD and Karasek E. 1977. Toxicity of salmonellae isolated from reptiles, in XIX Internationalen Symposiums uber die Erkrankungen der Zootiere, Akademie der Wissenschaften der DDR, Berlin, 87–91. Schröter M, Heckers KO, Rüschoff B, Laufs R, and Mack D. 2005. Severe case of spinal osteomyelitis due to Enterococcus spp. in a three-yearold rhinoceros viper, Bitis nasicornis. J Herp Med Surg 15:53–56. Schröter M, Speicher A, Hofmann J, and Roggentin P. 2006. Analysis of the transmission of Salmonella spp. through generations of pet snakes. Environ Microbiol 8:556–559. Schröter M, Speicher A, Hofmann J, and Roggentin P. 2006. Analysis of the transmission of Salmonella spp. through generations of pet snakes. Environ Microbiol 8:556–559. Schumacher IM, Brown MB, Jacobson ER, Collins BR, and Klein PA. 1993. Detection of antibodies to a pathogenic mycoplasma in desert tortoises (Gopherus agassizii) with upper respiratory tract disease. J Clin Microbiol 31:1454–1460. Schumacher J, Pellicane C, Heard DJ, and Voges A. 1996. Periorbital abscess in a three-horned chameleon (Chamaeleo jacksonii). Vet Comp Ophthamol 6:30–33. Shotts EB, Gaines JL, Martin C, and Prestwood AK. 1972. Aeromonas induced death among fish and reptiles in an eutrophic inland lake. J Amer Vet Med Assoc 161:603–607. Simmons GC, Sullivan ND, and Green PE. 1972. Dermatophilosis in a lizard (Amphibolurus barbatus). Aust Vet J 48:465–466. Simpson CF, Jacobson ER, and Harvey JW. 1981. Electron microscopy of a spiral-shaped bacterium in the blood and bone marrow of a rhinoceros iguana. Can J Comp Med 45:388–391. Sinha RP, Soman JP, Jha GJ, Prasad A, Chauman HVS, and Prasad RS. 1988. An outbreak of Escherichia coli enteritis in crocodiles. Ind J Anim Sci 58:338–340. Slavtchev RS and Chadli A. 1984. Infections and deaths of horned vipers, Cerastes cerastes (L., 1758) and lebetin vipers, Vipera lebetina (L., 1758) caused by Pseudomonas aeruginosa (Schroeter, 1885). Arch Inst Pasteur Tunis 61:415–25. Snipes KP and Biberstein EL. 1982. Pasteurella testudinis sp. nov.: a parasite of desert tortoises. Int J Syst Bact 32:201–210. Snipes KP, Biberstein EL, and Fowler ME.1980. A Pasteurella sp. associated with respiratory disease in captive desert tortoises. J Amer Vet Med Assoc 177:804–807. Snipes KP, Kasten RW, Calagoan JM, and Boothby JT. 1995. Molecular characterization of Pasteurella testudinis isolated from desert tortoises (Gopherus agassizii) with and without upper respiratory tract disease. J Wildl Dis 31:22–29. Soares JF, Chalker VJ, Erles K, Holtby S, Waters M, and McArthur S. 2004. Prevalence of Mycoplasma agassizii and chelonian herpesvirus in captive tortoises (Testudo sp.) in the United Kingdom. J Zoo Wildl Med 35:25–33. Soldati G, Lu ZH, Vaughan L, Polkinghorne A, Zimmermann DR, Huder JB, and Pospischil A. 2004. Detection of mycobacteria and chlamydiae in granulomatous inflammation of reptiles: a retrospective study. Vet Pathol 41:388–397.

Soveri T. 1984. Observations of bacterial diseases of captive snakes in Finland. Nord Vet Med 36:38–42. Stahl S. 1997. Captive management, breeding, and common medical problems of the veiled chameleon (Chamaeleo calyptratus), in Proceedings of the Association of Reptilian and Amphibian Veterinarians Fourth Annual Conference, Houston, TX, 29–40. Stanchi NO, Grisolia CS, Martino PE, and Peluso FO. 1986. Presence of antileptospira antibodies in ophidia in Argentina. Rev Argent Microbiol 18:127–130. Stewart JS, 1990. Anaerobic bacterial infections in reptiles. J Zoo Wildl Med 21:180–184. Tangredi BP and Evans RH. 1997. Organochlorine pesticides associated with ocular, nasal, or otic infection in the eastern box turtle (Terrapene carolina carolina). J Zoo Wildl Med 28:97–100. Tauxe RV, Rigau-Perez JG, Wells JG, and Blake PA. 1985. Turtleassociated salmonellosis in Puerto Rico. Hazards of the global turtle trade. J Amer Vet Med Assoc 254:237–239. Theakston RD, Phillips RE, Looareesuwan S, Echeverria P, Makin T, and Warrell DA. 1990. Bacteriological studies of the venom and mouth cavities of wild Malayan pit vipers (Calloselasma rhodostoma) in southern Thailand. Trans R Soc Trop Med Hyg 84:875–879. Turton JA, Ladds PW, and Melville LF. 1996. Interdigital subcutaneous emphysema (“bubble foot”) in Crocodylus porosus hatchlings. Aust Vet J 74:395–396. Urbain A, DeChambre E, and Piette G. 1951. Septicemie due a Clostridium oedematiens, type A, sur les tortues de la menagerie des reptiles du museum. Bull Mus Natl Hist Nat 23:247–248. Van der Hoeden J. 1968. Agglutination of Leptospirae in sera of fresh water turtles. Antonie van Leeuwenhoek 34:458–464. Vanrompay D, de Meurichy W, Ducatelle R, and Haesebrouck F. 1994, Pneumonia in Morrish tortoises (Testudo graeca) associated with avian serovar A Chlamydia psittaci. Vet Rec 135:284–285. Wallach JD. 1975. The pathogenesis and etiology of ulcerative shell diseases in turtles. J Zoo Anim Med 6:11–13. Weber A and Pietsch O. 1974. Aein Beitrag zum Vorkommen von Salmonellen bei Landschildkroten aus Zoohandlungen und Privathaushalten. Berliner und Munchener tierarztliche Wochenschrift 87:257–259. Weese JS and Staempfli HR. 2000. Diarrhea associated with enterotoxigenic Clostridium perfringens in a red footed tortoise (Geochelone carbonaria). J Zoo Wildl Med 31:265–266. Wellehan JFX, Turenne C, Heard DJ, Detrissac CJ, and O’Kelley M. 2004a. Dermatophilus chelonae in a king cobra (Ophiophagus hannah). J Zoo Wildl Med 35:553–556. Wellehan JFX, Nichols DK, Li, L-L, and Kapur V. 2004b. Three novel herpesviruses associated with stomatitis in Sudan plated lizards (Gerrhosaurus major) and a black-lined plated lizard (Gerrhosaurus nigrolineatus). J Zoo Wildl Med 35:50–55. Wellehan JFX, Johnson AJ, Latimer KS, Whiteside DP, Crawshaw GJ, Detrisac CJ, Terrell SP, Heard DJ, Childress A, and Jacobson ER. 2004c. Varanid herpesvirus-1: a novel herpesvirus associated with proliferative stomatitis in green tree monitors (Varanus prasinus). Vet Mirco 105:83–92. Wendland LD, Brown DR, Klein PA, and Brown MB. 2006. Upper respiratory tract disease (mycoplasmosis) in tortoises, in Reptile Medicine and Surgery, 2nd edition, Mader DR (Ed.,), Saunders, Elsevier, St. Louis, MO, 931–938. Willer CJ, Lewbart GA, and Lemons C. 2003. Aural abscesses in wild eastern box turtles, Terrapene carolina carolina: Aerobic bacterial isolates and distribution of lesions. J Herpetol Med Surg 13:4–9.

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Figure 10.1  Savannah monitor, Varanus exanthematicus. Varanidae. Pseudomonas infection. Chronic gingivitis and stomatitis associated with Pseudomonas infection.

Figure 10.2  Green iguana, Iguana iguana. Iguanidae. Pseudomonas infection. Necrotizing glossitis associated with Pseudomonas infection.

Figure 10.3  Blood python, Python curtus. Pythonidae. Pseudomonas infection. Chronic stomatitis associated with Pseudomonas infection.

Bacterial Diseases of Reptiles  489

Figure 10.4  Burmese python, Python molurus bivittatus. Pythonidae. Pseudomonas infection. Chronic stomatitis associated with Pseudomonas infection.

Figure 10.5  Burmese python, Python molurus bivittatus. Pythonidae. Pseudomonas infection. Trauma-induced gingivitis and stomatitis associated with Pseudomonas infection. Figure 10.6  Boa constrictor, Boa constrictor. Pseudomonas infection. Glossitis associated with Pseudomonas infection. Caseous material surrounds the tongue (arrow). The snake has been intubated and is anesthetized.

Figure 10.7  Burmese python, Python molurus bivittatus. Pythonidae. Pseudomonas infection. Necrotizing epidermal and dermal necrosis resulting from a burn. Pseudomonas was cultured from the wound. (From Jacobson ER. 1981. Compend Contin Educ Pract Vet 3:195–200. With permission.)

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Figure 10.8  Three-horned chameleon, Chamaeleo jacksonii. Chamaeleonidae. Pseudomonas infection. Pseudomonas was cultured from this periorbital abscess. Courtesy of Juergen Schumacher.

Figure 10.9  Ball python, Python regius. Pythonidae. Pseudomonas infection. Pseudomonas was cultured from caseous material (arrow) in the subspectacular space.

Figure 10.10  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Aeromonas infection. There is diffuse hemorrhage of the oral mucosa. Aeromonas hydrophila was cultured from the oral cavity.

Figure 10.11  Boa constrictor, Boa constrictor. Boidae. Aeromonas infection. A heterophilic granuloma within the lung. Aeromonas hydrophila was cultured from this lesion. H&E stain.

Bacterial Diseases of Reptiles  491

Figure 10.12  Ridgenose rattlesnake, Crotalus willardi. Viperidae. Salmonella infection. Osteomyelitis associated with Salmonella enterica arizonae infection. Adjacent to the spinal cord (SP) are multifocal areas of lysed bone surrounded by inflammation (arrows). H&E stain. Courtesy of Linden Craig.

Figure 10.13  Slider, Trachemys sp. Emydidae. Citrobacter infection. Septicemic ulcerative cutaneous disease. Citrobacter was cultured from this turtle. (From Jacobson ER. 1981. Compend Contin Educ Pract Vet 3:195–200. With permission.)

Figure 10.14  Gopher tortoise, Gopherus polyphemus. Testudinidae. Serratia infection. Multiple subcutaneous abscesses can be seen. Serratia marcescens was cultured from these lesions.

Figure 10.15  Gopher tortoise, Gopherus polyphemus. Testudinidae. Serratia infection. Photomicrograph of an abscess from a gopher tortoise infected with Serratia marcescens.

492  Bacterial Diseases of Reptiles

Figure 10.16  Green iguana, Iguana iguana. Iguanidae. Neisseria infection. Nodules are seen within the lungs of an iguana infected with Neisseria iguanae. Courtesy of Tabitha Viner and the National Zoological Park, Washington, DC.

Figure 10.17  Green iguana, Iguana iguana. Iguanidae. Neisseria infection. Photomicrograph of a dermal pyogranuloma. Within the granuloma is an eosinophilic amorphous material radiating from the periphery. H&E stain. Courtesy of Tabitha Viner and the National Zoological Park, Washington, DC. (From Plowman C et al. 1987. J Zoo Anim Med 18:86–93. With permission.)

Figure 10.18  Map turtle, Graptemys barbouri. Emydidae. Elizabethkingia infection. Elizabethkingia meningoseptica (formerly Flavobacterium meningosepticum) within cytosomes in macrophages of the liver. H&E stain.

Figure 10.19  Map turtle, Graptemys barbouri. Emydidae. Elizabethkingia infection. Elizabethkingia meningoseptica (formerly Flavobacterium meningosepticum) within cytosomes in macrophages of the liver. Giemsa stain. (From Jacobson ER et al. 1989. J Zoo Wildl Med 20: 474–477. With permission.)

Bacterial Diseases of Reptiles  493

Figure 10.20  Singapore house gecko, Gekko monarchus. Gekkonidae. Streptococcus infection. Photomicrograph of the liver containing multiple nodules of a streptococcal bacterium. An inflammatory response is lacking. H&E stain. Courtesy of Tracey S McNamara. (From McNamara TS et al. 1994. J Zoo Wildl Med 25:161–166. With permission.)

Figure 10.21  Singapore house gecko, Gekko monarchus. Gekkonidae. Streptococcus infection. Nodule within the kidney. Numerous chains of a streptococcal bacterium can be seen. H&E stain. Courtesy of Tracey S. McNamara. (From McNamara TS et al. 1994. J Zoo Wildl Med 25:161–166. With permission.)

Figure 10.22  Singapore house gecko, Gekko monarchus. Gekkonidae. Streptococcus infection. Electron photomicrograph of a streptococcal bacterium arranged in a chain. The capsule consists of a thin fibrillar material. Uranyl acetate and lead citrate. Courtesy of Tracey S. McNamara.

Figure 10.23  Green iguana, Iguana iguana. Iguanidae. Peptostreptococcus infection. Peptostreptococcus was cultured from the hindlimb of a green iguana that was bitten by a monitor, Varanus sp.

494  Bacterial Diseases of Reptiles

Figure 10.24  Boa constrictor, Boa constrictor. Boidae. Mycobacteriosis. Ulcerative proliferative mouth lesion. Mycobacterium chelonei was isolated from this lesion.

Figure 10.25  Boa constrictor, Boa constrictor. Boidae. Mycobacteriosis. Multiple granulomas consisting primarily of macrophages are seen adjacent to the trachea. H&E stain.

Figure 10.26  Boa constrictor, Boa constrictor. Boidae. Mycobacteriosis. Numerous filamentous acid-fast positive bacteria are seen within a paratracheal granuloma. Acid-fast stain.

Bacterial Diseases of Reptiles  495

Figure 10.27  Rock rattlesnake, Crotalus lepidus. Viperidae. Mycobacteriosis. Numerous filamentous acid-fast positive bacteria are seen within a pulmonary granuloma. Acid-fast stain.

Figure 10.28  Reticulated python, Python reticulatus. Pythonidae. Mycobacteriosis. White military lesions are seen within the lung. Using special staining, acid-fast bacteria were identified.

Figure 10.29  Texas rat snake, Elaphe obsoleta lindheimerii. Colubridae. Mycobacteriosis. Acidfast bacteria were identified in the necroproliferative oral lesions of this snake.

496  Bacterial Diseases of Reptiles

Figure 10.30  Desert tortoise, Gopherus agassizii. Testudinidae. Dermatophilosis. Multiple hyperkeratotic skin lesions can be seen. A filamentous organism consistent with Dermatophilus was identified in this lesion.

Figure 10.31  Padlopper tortoise, Homopus signatus. Testudinidae. Dermatophilosis. Multiple hyperkeratotic skin lesions can be seen (arrows). A filamentous organism consistent with Dermatophilus was identified in this lesion.

Figure 10.32  Saltwater crocodile, Crocodylus porosus. Crocodylidae. Dermatophilosis. Multifocal brown pitted skin lesions are seen on the abdomen. Dermatophilus was identified in these lesions. Courtesy of Gilbert N. Buenviaje and Phillip Ladds.

Figure 10.33  Saltwater crocodile, Crocodylus porosus. Crocodylidae. Dermatophilosis. A filamentous branching organism consistent with Dermatophilus is seen within multifocal brown pitted skin lesions on the abdomen. Gram stain. Courtesy of Gilbert Buenviaje and Phillip Ladds.

Bacterial Diseases of Reptiles  497

Figure 10.34  Bush anole, Polychrus marmoratus. Iguanidae. Dermatophilosis. Multifocal brown hyperkeratotic skin lesions can be seen on the body surface.

Figure 10.35  Bush anole, Polychrus marmoratus. Iguanidae. Dermatophilosis. Photomicrograph of a hyperkeratotic skin lesion. The epidermis under the thickened keratin is necrotic. H&E stain.

Figure 10.36  Bush anole, Polychrus marmoratus. Iguanidae. Dermatophilosis. Photomicrograph of the skin showing a branching filamentous bacterium compatible with Dermatophilus within necrotic cellular debris. Giemsa stain.

Figure 10.37  Bush anole, Polychrus marmoratus. Iguanidae. Dermatophilosis. Higher magnification photomicrograph of Figure 10.36 showing a branching filamentous bacterium compatible with Dermatophilus in necrotic cellular debris. Giemsa stain.

498  Bacterial Diseases of Reptiles

Figure 10.38  Collared lizard, Crotaphytus collaris. Iguanidae. Dermatophilosis. An ulcerating necrotizing epidermal lesion was found to contain organisms compatible with Dermatophilus.

Figure 10.39  Collared lizard, Crotaphytus collaris. Iguanidae. Dermatophilosis. Light microscopic photomicrograph of the skin showing a branching septate bacterium compatible within Dermatophilus in necrotic cellular debris. Giemsa stain.

Figure 10.40  Senegal chameleon, Chamaeleo senegalensis. Chamaeleonidae. Dermatophilosis. Multifocal brown hyperkeratotic skin lesions can be seen on the body surface.

Bacterial Diseases of Reptiles  499

Figure 10.41  Senegal chameleon, Chamaeleo senegalensis. Chamaeleonidae. Dermatophilosis. Photomicrograph of the skin. A filamentous bacteria with cross-striations and compatible with Dermatophilus can be seen in keratin and cellular debris. Giemsa stain.

Figure 10.42  Boa constrictor, Boa constrictor. Boidae. Dermatophilosis. Photomicrograph of a surgically removed laminated subcutaneous nodule. H&E stain.

Figure 10.43   Boa constrictor, Boa constrictor. Boidae. Dermatophilosis. Photomicrograph of a laminated subcutaneous nodule. Inset: Higher magnification image showing a Gram-positive filamentous microorganism compatible with Dermatophilus. Gram stain.

500  Bacterial Diseases of Reptiles

Figure 10.44  Desert tortoise, Gopherus agassizii. Testudinidae. Mycoplasmosis. A purulent nasal discharge can be seen.

Figure 10.45  Desert tortoise, Gopherus agassizii. Testudinidae. Mycoplasmosis. A serous nasal discharge can be seen.

Figure 10.46  Desert tortoise, Gopherus agassizii. Testudinidae. Normal nasal cavity. Subgross photomicrograph of the sagittal section of the head of a normal tortoise. A large nasal cavity (NC) is seen. H&E stain.

Figure 10.47  Desert tortoise, Gopherus agassizii. Testudinidae. Normal nasal cavity. Photomicrograph of the ventral nasal cavity showing a mucous epithelium with ciliated and mucous epithelial cells. H&E stain.

Bacterial Diseases of Reptiles  501

Figure 10.48  Desert tortoise, Gopherus agassizii. Testudinidae. Normal nasal cavity. Photomicrograph of the dorsal nasal cavity showing a multilayered olfactory epithelium. H&E stain.

Figure 10.49  Desert tortoise, Gopherus agassizii. Testudinidae. Mycoplasmosis. Photomicrograph of the nasal cavity showing diffuse infiltrates of mixed inflammatory cells into the mucosa and submucosa. H&E stain.

Figure 10.50  Desert tortoise, Gopherus agassizii. Mycoplasmosis. Photomicrograph of the nasal cavity revealing diffuse hyperplasia of the mucosal epithelium with infiltrates of heterophils and macrophages. H&E stain.

Figure 10.51  Desert tortoise, Gopherus agassizii. Mycoplasmosis. Photomicrograph of the nasal septum revealing a hyperplastic mucous epithelium and infiltrates of mixed inflammatory cells. H&E stain.

502  Bacterial Diseases of Reptiles

Figure 10.52  Gopher tortoise, Gopherus polyphemus. Mycoplasmosis. The nares have a serous discharge.

Figure 10.53  Gopher tortoise, Gopherus polyphemus. Testudinidae. Mycoplasmosis. There is depigmentation of the palpebrae.

Figure 10.54  Gopher tortoise, Gopherus polyphemus. Testudinidae. Mycoplasmosis. Transmission electron photomicrograph showing Mycoplasma (arrows) on the cell surface of an epithelial cell in the nasal cavity. Uranyl acetate and lead citrate.

Bacterial Diseases of Reptiles  503

Figure 10.55  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. There is caseous material in air passageways of the lung.

Figure 10.56  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Heart with fibrinous epicarditis.

Figure 10.57  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the heart showing a fibrinous epicarditis (arrows). H&E stain.

504  Bacterial Diseases of Reptiles

Figure 10.58  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the spleen showing lymphoid cells and macrophages within the red pulp. H&E stain.

Figure 10.59  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the liver showing periportal inflammation and fibroplasia. H&E stain.

Figure 10.60  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Higher magnification photomicrograph of the liver in Figure 10.59. Periportal inflammation and fibroplasia are seen. H&E stain.

Figure 10.61  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the lung showing interstitial inflammation. H&E stain.

Bacterial Diseases of Reptiles  505

Figure 10.62  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the lung showing granulomatous inflammation. H&E stain.

Figure 10.63  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the synovium with infiltrates of mixed inflammatory cells. H&E stain.

Figure 10.64  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the brain with perivascular cuffing and diffuse infiltrates of small mononuclear cells. H&E stain.

506  Bacterial Diseases of Reptiles

Figure 10.65  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the brain with a focal area of necrosis and infiltrates of heterophils. H&E stain.

Figure 10.66  American alligator, Alligator mississippiensis. Alligatoridae. Mycoplasmosis. Photomicrograph of the brain with infiltrates of small mononuclear cells in the choroid. H&E stain.

Figure 10.67  Burmese python, Python molurus. Pythonidae. Mycoplasmosis. Photomicrograph of the lung with a proliferation of epithelial cells lining airways and infiltrates of small mononuclear cells in the interstitium. H&E stain.

Bacterial Diseases of Reptiles  507

Figure 10.68  Green turtle, Chelonia mydas. Cheloniidae. Chlamydiosis. Photomicrograph of the heart with separation, degeneration, and necrosis of myocardial fibers, and infiltrates of mixed inflammatory cells. H&E stain.

Figure 10.69  Green turtle, Chelonia mydas. Cheloniidae. Chlamydiosis. Photomicrograph of the heart with positive (brown) staining of the chlamydial antigen. Avidin biotin peroxidase complex method. Mayer’s hematoxylin counterstain.

Figure 10.70  Green turtle, Chelonia mydas. Chlamydiosis. Transmission electron photomicrograph showing developmental stages of chlamydia in the heart. Uranyl acetate and lead citrate.

Figure 10.71  Flap-necked chameleon, Chamaeleo dilepis. Chamaeleonidae. Chlamydiosis. Photomicrograph of the spleen with basophilic chlamydial inclusions (C) within macrophages. H&E stain.

508  Bacterial Diseases of Reptiles

Figure 10.72  Flap-necked chameleon, Chamaeleo dilepis. Chamaeleonidae. Chlamydiosis. Transmission electron photomicrograph of a circulating monocyte containing poxvirus (P) and developmental stages of Chlamydia (C). Uranyl acetate and lead citrate. (From Jacobson ER and Telford SR. 1990. J Wildl Dis 26:572–577. With permission.)

Figure 10.73  Puff adder, Bitis arietans. Viperidae. Chlamydiosis. Snake with clinical signs of respiratory disease.

Figure 10.74  Puff adder, Bitis arietans. Viperidae. Chlamydiosis. Photomicrograph of the heart with multiple granulomas (G) having eosinophilic centers. H&E stain.

Figure 10.75  Puff adder, Bitis arietans. Viperidae. Chlamydiosis. Photomicrograph of the liver with multiple granulomas. H&E stain.

Bacterial Diseases of Reptiles  509

Figure 10.76  Puff adder, Bitis arietans. Viperidae. Chlamydiosis. Photomicrograph of the liver with a histiocytic granuloma containing multiple basophilic inclusions (arrows). H&E stain. (From Jacobson ER et al. 1989. J Zoo Wildlf Med 20:364–369. With permission.)

Figure 10.77  Puff adder, Bitis arietans. Viperidae. Chlamydiosis. Photomicrograph of the liver with positive (brown) staining of the chlamydial antigen within a histiocytic granuloma. Avidin biotin peroxidase complex method. Harris hematoxylin counterstain.

Figure 10.78  Emerald tree boa, Corallus caninus. Boidae. Chlamydiosis. Photomicrograph of an esophageal tonsil containing macrophages. H&E stain.

Figure 10.79  Emerald tree boa, Corallus caninus. Boidae. Chlamydiosis. Photomicrograph of histiocytic granulomas (arrows) in the submucosa of the small intestine. H&E stain.

510  Bacterial Diseases of Reptiles

Figure 10.80  Emerald tree boa, Corallus caninus. Boidae. Chlamydiosis. Photomicrograph of the colon with infiltrates of lymphocytes and macrophages in the submucosa. H&E stain. (From Jacobson ER et al. 2002. J Vet Diag Investig 14:487–494. With permission.)

Figure 10.81  Emerald tree boa, Corallus caninus. Boidae. Chlamydiosis. Photomicrograph of a small intestinal histiocytic granuloma with positive (brown) staining of the chlamydial antigen within macrophages. Avidin biotin peroxidase complex method. Harris hematoxylin counterstain.

Figure 10.82  Emerald tree boa, Corallus caninus. Boidae. Chlamydiosis. Photomicrograph of an esophageal tonsil with positive (brown) staining of the chlamydial antigen within macrophages. Avidin biotin peroxidase complex method. Harris hematoxylin counterstain.

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Figure 10.83  Emerald tree boa, Corallus caninus. Boidae. Chlamydiosis. Photomicrograph of the colon with positive (brown) staining of chlamydial antigen within macrophages in the submucosa. Avidin biotin peroxidase complex method. Harris hematoxylin counterstain. (From Jacobson ER et al. 2002. J Vet Diag Investig 14:487–494. With permission.)

Figure 10.84  Emerald tree boa, Corallus caninus. Boidae. Chlamydiosis. Transmission electron photomicrograph of a histiocyte in an intestinal granuloma. Developmental stages of chlamydia can be seen in the cytoplasm. Uranyl acetate and lead citrate. (From Jacobson ER et al. 2002. J Vet Diag Investig 14:487– 494. With permission.)

Figure 10.85  Emerald tree boa, Corallus caninus. Boidae. Chlamydiosis. Transmission electron photomicrograph of elementary bodies (arrows) and initial bodies (I) within the cytoplasm of a monocyte. Uranyl acetate and lead citrate. (From Jacobson ER et al. 2002. J Vet Diag Investig 14:487–494. With permission.)

512  Bacterial Diseases of Reptiles

Figure 10.86  Rhinoceros iguana, Cyclura cornuta. Iguanidae. Spiral-shaped bacterium. Photomicrograph of spiral-shaped microorganisms in the sinusoids of the liver. GMS stain.

Figure 10.87  Rhinoceros iguana, Cyclura cornuta. Iguanidae. Spiral-shaped bacterium. Transmission electron photomicrograph. Spiral-shaped microorganisms are seen within macrophages in bone marrow. Uranyl acetate and lead citrate stains.

Bacterial Diseases of Reptiles  513

Figure 10.88  River cooter, Pseudemys concinna. Emydidae. Disfiguring shell disease of the carapace. This represents segmental necrosis of the epidermis and dermis with extensive remodeling of dermal bone.

Figure 10.89  River cooter, Pseudemys concinna. Emydidae. Disfiguring shell disease of the plastron. This represents segmental necrosis of the epidermis and dermis with extensive remodeling of dermal bone.

Figure 10.90  River cooter, Pseudemys concinna. Emydidae. Shell disease. Photomicrograph of a shell lesion showing regenerative epidermal cells (arrows) beneath a layer of necrotic debris. H&E stain. (From Garner MM et al. 1997. J Wildl Dis 33:78–86. With permission.)

514  Bacterial Diseases of Reptiles

Figure 10.91  Eastern box turtle, Terrapene carolina. Emydidae. A large aural abscess is seen caudal to the right eye. (From Jacobson ER. 1981. Compend Contin Educ Pract Vet 3:195-200. With permission.)

Figure 10.92  Green turtle, Chelonia mydas. Cheloniidae. A large laminated caseous abscess is seen protruding from a foreflipper.

Figure 10.93  Bearded dragon, Pogona vitticeps. Agamidae. Caseous subcutaneous and submucosal abscess of the oral cavity.

Figure 10.94  Green iguana, Iguana iguana. Iguanidae. Abscess of the foot resulting in osteomyelitis.

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Figure 10.95  Green iguana, Iguana iguana. Iguanidae. Caseous abscess of the rostrum secondary to trauma.

Figure 10.96  Gaboon viper, Bitis gabonica. Viperidae. Subcutaneous abscess.

Figure 10.97  Gaboon viper, Bitis gabonica. Viperidae. Transected subcutaneous abscess revealing miliary caseous granulomas.

Figure 10.98  Burmese python, Python molurus bivittatus. Pythonidae. Photomicrograph of a heterophilic granuloma within an oral abscess. H&E stain.

516  Bacterial Diseases of Reptiles

Figure 10.99  Burmese python, Python molurus bivittatus. Pythonidae. Photomicrograph of diffuse inflammation and fibrosis in an oral abscess. H&E stain.

Figure 10.100  California king snake, Lampropeltis getula californiae. Colubridae. Anal gland infection. The anal glands are impacted and abscessed.

Figure 10.101  Sinaloan milk snake, Lampropeltis triangulum sinaloae. Colubridae. Anal gland infection. Abscessed and fistulated anal glands.

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Figure 10.102  Green iguana, Iguana iguana. Iguanidae. Subconjunctival abscess.

Figure 10.103  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Bilateral periorbital abscesses.

Figure 10.104  Green tree python, Morelia viridis. Pythonidae. Unilateral subspectacular infection.

518  Bacterial Diseases of Reptiles

Figure 10.105  Eastern box turtle, Terrapene carolina. Emydidae. Stomatitis with caseous material in the oral cavity.

Figure 10.106  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Multiple abscesses are seen on the tongue. Courtesy of Rob Coke.

Figure 10.107  Jackson’s threehorned chameleon, Chamaeleo jacksonii xantholophus. Chamaeleonidae. Glandular abscess (arrow) in the lateral commissure of the oral cavity. Courtesy of Rob Coke.

Figure 10.108  Meadow viper, Vipera ursinii. Viperidae. Stomatitis of the mucosa surrounding the maxillary bone and fang resulting in osteomyelitis.

Bacterial Diseases of Reptiles  519

Figure 10.109  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Pneumonia with caseous material filling faveolar spaces.

Figure 10.110  Cook’s tree boa, Corallus enydris. Boidae. Proteus infection. Caseous lesions within the liver and lung (far right) from which Proteus was cultured.

Figure 10.111  Burmese python, Python molurus bivittatus. Pythonidae. Chronic pulmonary disease. Mucoid material is seen within the glottis (arrow).

Figure 10.112  Burmese python, Python molurus bivittatus. Pythonidae. Chronic pulmonary disease. A tube is inserted into the respiratory tract to remove accumulating exudate.

520  Bacterial Diseases of Reptiles

Figure 10.113  Burmese python, Python molurus bivittatus. Pythonidae. Chronic pulmonary disease. Material suctioned from the lung of an affected python.

Figure 10.114  Burmese python, Python molurus bivittatus. Pythonidae. Chronic pulmonary disease. Photomicrograph of the lung showing hyperplasia of lining epithelial cells and an interstitium free of inflammatory cells.

Figure 10.115  Burmese python, Python molurus bivittatus. Pythonidae. Chronic pulmonary disease. The lining epithelium is hyperplastic and there are infiltrates of small mononuclear cells within the interstitium and the layer of hyperplastic epithelial cells.

Bacterial Diseases of Reptiles  521

Figure 10.116  Burmese python, Python molurus bivittatus. Pythonidae. Thromboembolic disease. Cephalic cervical edema secondary to a thrombus in the sinus venosus.

Figure 10.117  Burmese python, Python molurus bivittatus. Pythonidae. Thromboembolic disease. Intravenous contrast radiograph revealing a space occupying mass (arrows) in the sinus venous and right atrium. (From Jacobson ER et al. 1991. J Zoo Wildl Med 22:245–248. With permission.)

Figure 10.118  Burmese python, Python molurus bivittatus. Pythonidae. Thromboembolic disease. A large thrombus (arrow) is in the right atrium. Courtesy of John Roberts.

Figure 10.119  Burmese python, Python molurus bivittatus. Pythonidae. Thromboembolic disease. Photomicrograph of the pulmonary artery containing a thrombus consisting of fibrin with entrapped heterophils and red blood cells. H&E stain.

522  Bacterial Diseases of Reptiles

Figure 10.120  Burmese python, Python molurus bivittatus. Pythonidae. Thromboembolic disease. Necrosis of the distal end of the tail.

Figure 10.121  Burmese python, Python molurus bivittatus. Pythonidae. Thromboembolic disease. Necrosis of the distal end of the tail.

Figure 10.122  Burmese python, Python molurus bivittatus. Pythonidae. Thromboembolic disease. Photomicrograph showing thrombi in major vessels of the tail resulting in diffuse necrosis of surrounding tissues. H&E stain.

Figure 10.123  Burmese python, Python molurus bivittatus. Pythonidae. Thromboembolic disease. Thrombi in vessels of the lung are surrounded by infiltrates of inflammatory cells. H&E stain.

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Figure 10.124  Green iguana, Iguana iguana. Iguanidae. Osteoarthritis. Swollen distal right hindlimb. Caseous material was found in the joint spaces and there was osteomyelitis.

Figure 10.125  Green iguana, Iguana iguana. Iguanidae. Osteoarthritis. Swollen distal right hindlimb containing caseated material in the joint space.

Figure 10.126  Russian rat snake, Elaphe schrenckii. Colubridae. Osteomyelitis. Vertebral column and ribs with boney, pitted proliferations extending across joint spaces.

Figure 10.127  Boa constrictor, Boa constrictor. Boidae. Osteomyelitis. Vertebral column and ribs with boney, pitted proliferations extending across joint spaces. Courtesy of Ramiro Isaza.

524  Bacterial Diseases of Reptiles

Figure 10.128  Eastern hognose snake, Heterodon platyrhinos. Colubridae. Osteomyelitis. Photomicrograph showing bone lysis and chronic inflammation of a vertebra. Bacteria were identified in tissue section and Streptococcus was cultured from bone. H&E stain. (From Isaza R, Garner M, and Jacobson E. 2000. J Zoo Wildl Dis 31:20–27. With permission.)

Figure 10.129  Boa constrictor, Boa constrictor. Boidae. Osteomyelitis. Photomicrograph showing bone lysis and chronic inflammation of a vertebra. While bacteria were not identified in tissue section, Edwardsiella was cultured from bone. H&E stain. Courtesy of Ramiro Isazo and the Veterinary Pathology Service, University of Florida, Gainesville.

Figure 10.130  Boa constrictor, Boa constrictor. Boidae. Degenerative vertebral osteoarthrosis and ankylosis. There are irregular deposits of cartilage and bone resulting in thickening of the articular surfaces between adjacent vertebrae. There is an absence of inflammation. H&E stain. Courtesy of Ramiro Isazo and the Veterinary Pathology Service, University of Florida, Gainesville.

Bacterial Diseases of Reptiles  525

Figure 10.131  Rosy boa, Lichanura trivirgata roseofusca. Boidae. Meningitis. Photomicrograph of a heterophilic granuloma within the choroid. H&E stain. Courtesy of Eric Baitchman, John Roberts, and the Veterinary Pathology Service, University of Florida, Gainesville.

Figure 10.132  Rosy boa, Lichanura trivirgata roseofusca. Boidae. Meningitis. Photomicrograph showing bacteria within a heterophilic granuloma in the choroid. H&E stain. Courtesy of Eric Baitchman, John Roberts, and the Veterinary Pathology Service, University of Florida, Gainesville.

Figure 10.133  Corn snake, Elaphe guttata. Colubridae. Inner ear infection. Photomicrograph showing a focal area of necrosis (arrow) in nerve VIII adjacent to the otic labyrinth (OL). H&E stain.

Figure 10.134  Corn snake, Elaphe guttata. Colubridae. Inner ear infection. Higher magnification photomicrograph showing a focal area of necrosis and infiltrates of mixed inflammatory cells within nerve VIII. H&E stain.

526  Bacterial Diseases of Reptiles

Figure 10.135  Ball python, Python regius. Pythonidae. Oomphalitis. A mass of caseous yolk intermixed with inflammatory cells and fibrin is surgically removed from the caudal coelomic cavity.

Figure 10.136  Ball python, Python regius. Pythonidae. Oomphalitis. A mass of caseous yolk intermixed with inflammatory cells and fibrin removed from the caudal coelomic cavity.

11 Mycotic Diseases of Reptiles Jean A. Paré and Elliott R. Jacobson

Contents

11.1 General Comments

11.1 General Comments .............................................. 527 11.2 An Overview of the Fungi.................................... 528 11.3 Normal Mycobiota of Reptiles.............................. 529 11.4 Mycoses.................................................................. 529 11.4.1 Dermatomycoses....................................... 529 11.4.2 Deep or Systemic Mycoses....................... 530 11.5 Mycoses in Reptiles............................................... 530 11.5.1 Mycoses in the Chelonia........................... 531 11.5.2 Mycoses in the Crocodylia........................ 532 11.5.3 Mycoses in the Squamata......................... 532 11.6 Agents of Mycoses................................................. 533 11.6.1 Hyalohyphomycotic Agents ..................... 533 11.6.2 Dematiaceous Fungi.................................. 540 11.6.3 Yeast........................................................... 541 11.6.4 Dimorphic Fungi....................................... 542 11.6.5 Dermatophytes.......................................... 543 11.7 Conclusion............................................................. 543 References......................................................................... 544

Mycoses occur regularly in reptiles and are likely underdiagnosed. Because deep or systemic mycoses often follow a clinically silent, slow, and progressive course, they are difficult to recognize and are therefore overlooked unless imaging or laparoscopic diagnostic modalities are used, or a necropsy is performed. Dermatomycotic lesions are easier to notice but are indistinguishable grossly from those caused by bacterial infections and are often misdiagnosed as such. In the literature, fungal infections in reptiles were typically unsuspected clinically and merely diagnosed serendipitously at necropsy, often only after histopathology was performed (Austwick and Keymer, 1981; Frank, 1976; Migaki et al., 1984; Schildger et al., 1991b). Herpetological medicine has evolved to provide heightened standards of care. Reptile clinicians now have access to an arsenal of increasingly sophisticated diagnostic devices, such as ultrasound machines, computed tomography and laparoscopes, which enables them to detect and locate internal lesions. Reptile owners are both more educated and more amenable to assuming the costs of diagnostic procedures. Biopsies of cutaneous and deep lesions, essential for a diagnosis of mycosis to be established but seldom performed in the past, are now routinely part of a diagnostic workup in reptile patients so that mycoses are increasingly identified antemortem. Mycoses in reptiles were historically blamed on poor husbandry. Reptiles are ectotherms whose health depends on the surrounding temperature, especially in a captive environment where thermal gradients might not be provided, or where temperatures are maintained too low. Overcrowding of animals in an enclosure, compounded by inadequate sanitation, will lead to an increase in detritus and organic substrate in which fungi will grow. Excessive humidity and

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528  Mycotic Diseases of Reptiles

temperature also promote fungal replication and sporulation so that exposure to fungal spores may reach a magnitude that will overcome the reptile’s immune system. While deficient or inadequate husbandry undoubtedly predisposes reptiles to mycotic infection, we also now know that fungi may act as primary reptile pathogens, and that some fungal agents cause disease when housing conditions and captive care appear adequate (Paré et al., 2006a; Rose et al., 2001). Mycoses also occur in free-ranging chelonians (Rose et al., 2001), snakes (Cheatwood et al., 2003; McAllister et al., 1993), and lizards (Cork and Stockdale, 1994; Martinez-Silvestre and Galán, 1999), where husbandry is not an issue but climatic changes are. Fungal epizootics in wild reptiles have been associated with winter and colder temperatures (Cheatwood et al., 2003; Martinez-Silvestre and Galán, 1999). As more herpetological field research is conducted, it may be that we will discern a role for some fungi in the dynamics of wild reptile populations. In this chapter, a very basic description of the biology of fungi will be provided, the pathogenesis of fungal infection will be reviewed, the presentation of mycoses and dermatomycoses will be described in the various reptile taxa, and the recognized fungal pathogens and disease entities will be addressed in conducting an objective review of the literature on reptile mycoses. There are several review articles that list all documented cases of mycoses in reptiles and the interested reader is referred to them (Austwick and Keymer, 1981; Frank, 1976; Jacobson et al., 2000; Migaki et al., 1984), with the warning that in many of these cases, the causal relationship between the lesion and the putative agent is unproven and often questionable. This chapter will focus on presentation, etiopathogenesis, and pathology of mycoses in reptiles, and readers interested in the medical approach to fungal disease and antifungal therapy in reptiles are referred to Paré et al. (2006b).

11.2 An Overview of the Fungi Fungi are extremely successful organisms, and more than 100,000 species have been described forming the Kingdom Fungi (Mycota or Eumycota) or true fungi (De Hoog et al., 2000; Howard, 2003). All fungi are eukaryotic and characterized by the presence of a cell wall, and all are heterotrophic, requiring an organic substrate from which to derive nutrition (Deacon, 1997). Fungi release enzymes across the cell membrane and wall to digest substrate, and then absorb simple nutrients back across the cell wall. Most fungi are filamentous (molds) and produce elongated hyphae that grow apically, branch out, and interweave within the substrate to form a mycelium (Deacon, 1997). Fungi that grow by budding of cells and do not produce true mycelia are called yeasts. Fungi can reproduce sexually and asexually. The sexual stage is called the teleomorph and generates propagules called spores (Deacon, 1997). Taxonomy is largely based on the type of sexual

spores produced by the fungus (e.g., ascospores, basidiospores, zygospores). Most medically important fungi belong to the Ascomycetes (Phylum Ascomycota) and the Zygomycetes (Phylum Zygomycota) (Howard, 2003). In addition, all fungi reproduce asexually in a stage called the anamorph to produce asexual propagules called conidia (Deacon 1997). The teleomorph and anamorph of a fungus bear different scientific names (e.g., Arthroderma otae and Microsporum canis) and may morphologically be very different, yet they are the same organism. For some fungi there is no known sexual, or teleomorphic, stage. It may be that the right conditions for sexual reproduction of these fungi have not been met or that they simply are so successful asexually that they have lost the ability to reproduce sexually. These fungi are called imperfect, and are included in the Phylum Mitosporic Fungi (Fungi Imperfecti or Deuteromycota) (Howard, 2003). Identification of fungi can be difficult, and is based on, among other criteria, the morphology of the vegetative thallus, fruiting bodies (whether spores or conidia), and the specialized cells or structures that bear them (conidiophores) (Larone, 1995; St. Germain and Summerbell, 1996). Identification and speciation of yeasts may require biochemical tests using commercial systems (Larone, 1995). Taxonomy and nomenclature of fungi are unsettled and in constant flux, and the names of many fungi involved in documented cases of reptile mycosis have changed (e.g., Cephalosporium is now Acremonium) (De Hoog et al., 2000). In this chapter, the currently accepted scientific names will be used, and the obsolete name, under which they were previously reported, will be mentioned. We do not have a very good grasp as to which fungi carry pathogenic potential for reptiles. This is because numerous cases compiled in the literature fail to demonstrate a causative relationship between the fungal isolate and the lesions observed in the reptile. In many cases there was no culture performed, while in many others a presumptive diagnosis of mycosis was risked based on isolation of a fungus from a lesion without supportive histopathology. In addition, the recognition over the last decade of the Chrysosporium anamorph of Nannizziopsis viresii (CANV) as a major cause of hyalohyphomycoses in reptiles casts some doubt on the authenticity of previously reported cases of Trichophyton, Geotrichum, Chrysosporium, Malbranchea, and maybe even Trichosporon infections, all fungi that can easily be mistaken for it. We need to develop an appreciation for which fungi are able to opportunistically cause disease in reptiles, which ones are repeat offenders, and which ones are primary pathogens. The lines between these three categories may be blurry. Fungal virulence factors (e.g., thermotolerance, adhesions, proteases) and host predisposing factors (e.g., immune status, age, sex, species) are complex. Such factors are the subject of investigation for various human and plant pathogens, but are unexplored for reptile pathogens. It seems safe to assume that fungi that elaborate keratinases and various other tissue proteases have the potential to colonize living tissue and cause disease, but numerous other factors or combination of factors such as

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adhesins, cell wall composition, thermotolerance, magnitude of exposure, and host immune status, among others, likely determine the absence or presence of infection and modulate its outcome (Cole, 2003).

11.3 Normal Mycobiota of Reptiles The population of fungi normally found in a microenvironment, formerly called the fungal flora, is more appropriately referred to as the mycobiota because fungi are not plants. Studies investigating the cutaneous or intestinal mycobiota of reptiles are scarce, but all suggest that the mycobiota of reptiles, whether cutaneous or gastrointestinal, is rich and varied. In a study designed as a survey of normal reptile skin for the presence of the CANV, two one-by-one-inch pieces of aseptically collected, fresh exuvium from 127 different captive, healthy-looking reptiles were placed on a Mycosel agar and cultured at 28°C (Paré et al., 2003). All fungi growing from the cultures were identified to genus, and all keratinophilic fungi were keyed to species. Fungi belonging to more than 50 genera were identified from these cultures, and the mycobiota in individual animals ranged from zero to 13 different fungal genera. Penicillium and Aspergillus species were respectively isolated from 78% and 69% of these animals, underlining that isolation of these fungi from skin lesions may well be incidental, and a causative relationship needs be supported histopathologically if it is to be established (Paré et al., 2003). Fungi such as Paecilomyces lilacinus, chrysosporia, Zygomycetes, Scopulariopsis spp., Cladosporium spp., and fusaria, among others, were also recovered with consistency (Paré et al., 2003) (Figures 11.1A–B). Similarly, the reptilian gut mycobiota comprises many fungi. When the intestines of 29 African dwarf crocodiles (Osteolaemus tetraspis) were examined, 20 species of fungi were isolated (Huchzermeyer et al., 2000). The gut of 200 common agamas (Agama agama) was cultured and yielded Basidiobolus, Aspergillus, Candida, Penicillium, Fusarium and Mucor species (Enweani et al., 1997). Yeasts are present in the gut of ophidians, but even more so in herbivorous chelonians and saurians (Kostka et al., 1997; Milde et al., 2000). Yeasts were isolated from the gut, but also from the skin, eyes, liver, and less often the lungs and kidneys (Kostka, 1997). Culture of yeasts or of saprophytic opportunistic fungi from the feces of reptiles is not clinically relevant unless gastrointestinal signs are present and morphologically compatible fungal hyphae or budding yeast cells are seen on a stained fecal smear.

mycoses affecting skin or dermatomycoses, and deep or systemic mycoses affecting one or many organ systems. Mycoses may be further classified as phaeohyphomycosis or hyalohyphomycosis on the basis of the presence or absence, respectively, of pigment (usually melanin) in the cell wall of the causative fungus (De Hoog, 2000). Most phaeohyphomycoses are therefore easily diagnosed microscopically as the hyphae stain more or less dark brown on routine hematoxylin and eosin–stained (H&E-stained) sections (Figure 11.2), while special stains, such as Gomori’s silver stain (Figure 11.3) or periodic acid-Schiff (PAS) stain (Figure 11.4), are often necessary to demonstrate the presence of unpigmented or hyaline hyphae in tissues. Finally, mycoses are sometimes named after the causative agent (e.g., basidiobolomycosis, fusariomycosis, trichosporonosis) or its taxon (zygomycosis for an infection caused by a zygomycetous fungus) (De Hoog, 2000). Some disease terminology used previously in the literature is now obsolete (e.g., phycomycosis is now zygomycosis). Mycotic agents are diverse in reptiles, and only recently have several organisms been identified that are frequently implicated in disease in this group of vertebrates. Because fungi are part of the cutaneous microflora (mycobiota) and are also normally isolated from the viscera of healthy reptiles, it cannot be overly emphasized that a firm diagnosis of mycosis can never rely solely on isolation of a fungus from clinical material, and can only be established if fungal elements are demonstrated histologically within the lesions, and inflammation is present around the fungal elements, indicating that the fungus is not merely invading devitalized tissue. To implicate any isolate as the cause of a lesion, the morphology of the fungal elements in histological sections needs to be compatible with the morphology of the fungal isolate cultured from the lesion. When critically reviewing published compilations of cases of reptile mycoses, we must be wary of accounts in which these criteria are not met and causality is not clearly demonstrated. Compilations of reptile mycoses in which agents are not identified or cultured are frustrating and bring little contribution to our understanding of pathogenicity factors for reptile fungal pathogens. Only through the study of histologically confirmed cases of mycosis, in which a causative agent is clearly, or at least reasonably identified, can a pattern of fungi acting as repeat offenders be discerned. For this to happen, necropsy specimens need be available for culture, isolates need be keyed to species, and reptile pathogens, whether opportunistic or primary, should always be deposited in a microfungus depository for preservation and later study.

11.4.1 Dermatomycoses

11.4 Mycoses The terminology of mycoses may sometimes seem confusing. Mycoses are typically classified as superficial mycoses, the

Dermatomycosis is a term that refers to any fungal infection of the skin and cutaneous adnexa. By contrast, dermatophytosis is a term that is restricted to dermatomycoses caused by keratinophilic fungi in the genera Microsporum, Trichophyton, and Epidermophyton. Dermatomycoses may be superficial, with

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fungal invasion of the outer epidermal strata with or without dermal involvement, or may be deep and involve the hypodermis and subcutis. Dermatomycoses, superficial and deep, occur in all reptilian orders except the Rhynchocephalia, the tuataras (Sphenodon spp.), in which fungal disease has yet to be documented. The pathogenesis is probably very similar across reptile taxa. Superficial dermatomycoses occur when the skin comes in contact with spores or conidia of pathogenic fungi. If conditions are right, spores germinate and initially colonize the outer cornified epidermal layers. In this initial stage, true inflammation is not seen (Figure 11.5), although the odd heterophil may be seen migrating across the germinativum toward the corneum. Ecdysis may or may not rid the reptile of the fungus. This is particularly true in snakes, in which an intact exuvium is cast off. In snakes with fungal skin infection, dermatomycotic lesions can be seen on the exuvium. If the infection is superficial, all fungal elements may adhere to the exuvium. In such cases, ecdysis may act as a nonspecific barrier against deeper epidermal fungal invasion. If infection is deeper, fungi will also be seen on the outer exuvial surface. The outer surface is actually the surface in contact with the new stratum corneum because the exuvium is turned inside out when a snake sheds (Figure 11.6). Subjectively, ecdysis appears to occur more frequently in snakes with skin disease (Jacobson, 1980; Jacobson, 1981) (Figure 11.7), possibly an adaptive response. Down movement of fungal hyphae into the living stratum spinosum and stratum germinativum triggers a heterophilic response, typically but not always leading to coagulation, then liquefaction necrosis of the epidermis around fungal elements (Figure 11.8). Although hyperkeratosis is sometimes seen as infection progresses, epithelial necrosis and sloughing usually occurs and results in exposure of the underlying dermis. Hyphae penetrating the dermis are met initially with heterophils, but soon a mixed inflammatory cell infiltrate surrounds fungal elements, and in many cases, syncytial cells and fibroblasts eventually surround and wall off hyphae fragments, especially in the dermis (Figure 11.9). Granulomatous inflammation may become locally extensive and displace normal structures (Figure 11.10). In some cases, fungal hyphae appear to progress through the tissues relatively unchecked. Fungal invasion may further progress to disseminated or systemic mycosis, either transmurally across the body wall or via hematogenous or lymphatic spread (Figure 11.11). Inoculation of fungi below the skin through a cut or a puncture wound (rarely hematogenously) sometimes leads to the formation of large, more or less firm, rather well-defined subcutaneous masses called mycetomas (Figures 11.12– 11.16). Those are particularly common in snakes, especially in colubrids. They are grayish or pale yellow and may have a greasy, lipoma-like appearance on cut section.

11.4.2 Deep or Systemic Mycoses Deep mycoses may occur with or without concurrent dermatomycotic lesions. Extensive dermatomycoses may prog-

ress to deep mycoses. More often, fungal infection initiates in the lungs (Figures 11.17–11.18) and will either remain confined to the lungs or disseminate to other organ systems (Figure 11.19). Spread occurs hematogenously or by direct extension across the pleura to adjacent organs. Mycotic pneumonia is relatively common in tortoises (Figures 11.20– 11.21) and crocodilians (Figures 11.22–11.23) but occurs in all reptiles. Gastrointestinal mycoses are rarer, in spite of the omnipresence of fungal spores in the gut of reptiles. Yeast infections, mostly Candida sp., are the most common agents of gastrointestinal mycoses and often extend to the liver. Rarely, fungemia may occur. Fungal disease limited to the liver, the spleen, the urogenital system, or the central nervous system, is uncommon to rare.

11.5 Mycoses in Reptiles Agents of mycosis in reptiles are often different from those causing mycoses in other vertebrate taxa. In human medicine, there are about 100 fungi that are considered primary pathogens, and an additional 200 species that are opportunistic agents of mycosis. Pathogenic fungi include, among others, the dermatophytes (Trichophyton, Microsporum, and Epidermophyton spp.); the yeast Cryptococcus neoformans; the dimorphic fungi (Blastomyces, Coccidioides, Histoplasma), so called because they are filamentous in the environment yet adopt a yeast morphology in tissues, and others. Fungi like Aspergillus spp., Candida spp., Penicillium marneffei, and others are typically opportunistic, but with the AIDS epidemic and the advent of transplants and immunosuppressive or chemotherapeutic drugs, many of those opportunistic fungi have become extremely important clinically. Other mammals are more or less susceptible to the same fungal organisms that affect humans. In birds, aspergillosis and candidiasis are relatively common diseases, but are also thought to be correlated to some form of immune compromise, be it stress or immaturity of the immune system. Mycoses in amphibians are more common: mucormycosis and chromomycosis are welldefined clinicopathological entities, and chytridiomycosis is an important disease restricted to this class of vertebrates. In fish, Saprolegnia and Ichthyophonus, the most common fungal pathogens, have been reclassified outside the Kingdom Eumycota based on molecular data. Isolates from reptiles may be difficult for most laboratories to identify because they are not necessarily common fungi. For example, an isolate from the skin of a frilled lizard (Chlamydosaurus kingii), cultivated as part of a routine survey of squamate reptile skin for the CANV, could not be identified for the simple reason that it was unknown to science, and a new fungal genus and species, Chlamydosauromyces punctatus, were created to accommodate this isolate (Sigler et al., 2002). If an isolate cannot be identified accurately, it should be forwarded to a mycologist for proper speciation.

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Diagnosis of mycotic infections of the skin and oral cavity is relatively straightforward. Such lesions should always be biopsied (Figure 11.24) as fungal lesions may be indistinguishable from those of bacterial origin or even benign epithelial neoplasia. Diagnosis of deep or systemic mycosis is more challenging. Clinical signs in reptiles affected with deep mycosis are nonspecific. Anorexia, lethargy, and progressive weakness are merely indicative of a sick reptile. Other signs such as dyspnea, regurgitation, hematochezia or dyschezia, or neurological abnormalities reflect specific organ system involvement but not the cause. Hematological changes may include anemia, leukocytosis, and in severe cases, toxic granulocytes. But these changes are not specific. Serum biochemistry may be helpful to detect hepatic or renal involvement. Diagnosis relies on localization of lesions and subsequent biopsy. Serology for specific fungal antigens or antibodies has not been explored in reptiles. The presence of hyphae or yeast in tissue is essential to a diagnosis of mycosis. Mixed fungal and bacterial infections are common, and it may be difficult to establish which of the two is the primary offender. Close attention to the margins or leading edges of the lesion may help make that determination. Mixed fungal lesions may also occur, especially when fungi are secondary invaders, but are uncommon. A more likely scenario is that isolation of multiple fungal species or genera from a single lesion occurred due to contamination of the clinical specimen. Careful comparison of the morphology of fungal elements in tissue (e.g., hyphal width, pigmentation, septation, branching, and conidiation) to that of the isolates cultured from it will usually rule out contaminants. Repeated isolation of the same fungus from multiple biopsy sites or multiple organs is more convincing. Growth of the causative isolate is more likely to occur in, on, or around the specimen, rather than elsewhere on the agar (Figure 11.25). Fungal isolates should be forwarded to a microfungus depository for preservation. Deposited isolates are then available for later taxonomical evaluation if the speciation is disputed, can be compared to other related isolates, or can be used in experimental challenges to assess pathogenicity. While culture remains the gold standard, investigation of hyphae in tissues by means of immunohistochemistry, immunofluorescence techniques, or molecular probes is desirable if not essential, at least for publication purposes, when material suitable for culture is unavailable. Retrospective identification of fungal elements in tissue sections using such techniques would greatly enhance the scientific contribution of some publications.

11.5.1 Mycoses in the Chelonia Mycoses seem particularly prevalent in sea turtles (Duguy et al., 1998; Keymer, 1978; Turnbull et al., 2000). Mycoses were diagnosed at necropsy in 3% of freshwater turtles, but in 3 of 7 (43%) sea turtles (Keymer, 1978). Dermatomycotic lesions occur more frequently in sea turtles (Figure 11.26) and in freshwater turtles (Figure 11.27) than in terrestrial

chelonians. A retrospective study of observations and necropsies performed on 230 stranded or accidentally caught sea turtles in the Bay of Biscay included leatherback (Dermatochelys coriacea), loggerhead (Caretta caretta), Kemp’s ridley (Lepidochelys kempii), and green (Chelonia mydas) sea turtles (Duguy et al., 1998). Mycotic lesions were not reported in any of the 125 leatherback and seven green sea turtles examined. Of the 85 loggerhead sea turtles, about 50% presented plastral and carapacial dermatomycotic lesions described as white felty bordering of scutes, sometimes ulcerated to the underlying bone, and identical lesions were noted in four of 12 Kemp’s ridley turtles. Outbreaks of fungal dermatitis have been described in captive hatchling or young freshwater turtles. Mucor in Florida softshell turtles (Apalone [formerly Trionyx] ferox) (Jacobson et al., 1980) (Figures 11.28–11.30) and Paecilomyces lilacinus in Fly River turtles (Carettochelys insculpta) (Lafortune et al., 2005) (Figures 11.31–11.32) were identified as the causative agents.. Pitted lesions in the carapace of pond turtles are often thought of as being fungal in origin, but a biopsy is necessary to distinguish truly mycotic carapacial lesions from algal, oomycetous, or even bacterial lesions, or from invasion of necrotic scutes by nonpathogenic keratin decomposers. In tortoises and turtles, infection of the feet with fungi is not uncommon (Frye and Dutra, 1974; Shreve et al., 2004) and may be related to inappropriate substrate. In captive and free-ranging Texas tortoises (Gopherus berlandieri), necrotizing scute disease caused by Fusarium incarnatum (formerly F. semitectum) has been described and will be addressed later in this chapter. Sudden drops in seawater temperature often result in hypothermia in sea turtles. These cold-stunned turtles typically become weak, float, and strand. Rehabilitation is long and difficult, and hampered by sluggish recovery of the immune system and a plethora of medical issues (Turnbull et al., 2000). Severe mycoses are common sequelae of cold stunning in these animals, and antifungal drugs are often given prophylactically (Harms et al., 2002; Turnbull et al., 2000). Fungi such as Fusarium in particular are implicated, but even fungi of little-known pathogenicity, such as Colletotrichum acutatum, have caused disseminated mycoses in coldstunned turtles (Manire et al., 2002), indicating how severely immunocompromised some of these animals may become. Fungal infections have also been seen in wild stranded sea turtles in Florida (Jacobson, personal observations) unrelated to cold stunning events (Figures 11.33–11.34). Mycotic bronchopneumonia was described in 2 of 20 loggerhead sea turtles, and 1 of 6 Kemp’s ridley examined in the same study had disseminated mycotic disease (Duguy et al., 1998). Deep or systemic mycoses in chelonians chiefly affect the lungs. Dissemination to other organ systems often occurs. The giant tortoises appear overrepresented among cases of fungal pneumonia. The causative agents are heavily sporulating fungi whose conidia are ubiquitous in the air and commonly inhaled, such as Aspergillus, Paecilomyces, and Penicillium species. Low environmental temperatures were often listed in the anamnesis.

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Mycosis in chelonians is regularly documented in groups of hatchlings or very young turtles (Figure 11.35), especially in aquatic species (Jacobson et al., 1979; Jacobson et al., 2000; Lafortune et al., 2005; Lappin and Dunstan, 1992), suggesting that immaturity of the immune system, along with water quality and other husbandry issues, is probably an underlying factor.

11.5.2 Mycoses in the Crocodylia Not yet described in wild crocodilians, dermatomycoses are relatively common in captive farmed animals, especially saltwater crocodiles (Crocodylus porosus) (Buenviaje et al., 1994; Benviaje et al., 1998; Hibberd and Harrower, 1993) (Figures 11.36–11.38). Lesions are typically described as thickened, leathery, discolored plaques that may coalesce. Damage to the hide in these animals carries economic significance. The list of differential diagnoses includes bacterial dermatitis, dermatophilosis, and pox, especially in caimans. Biopsies and culture are diagnostic. Demonstration of fungal hyphae in tissue sections is necessary for a diagnosis of dermatomycosis to be made. The Chrysosporium anamorph of Nannizziopsis vriesii, Paecilomyces lilacinus, and Fusarium solani have been reliably incriminated and will be reviewed later in this chapter. The most severe outbreaks occurred in young animals, and underlying factors such as overcrowding and water quality issues are probably contributory. Fungal pneumonia is overwhelmingly the most common form of systemic mycosis in crocodilians. Lungs are thickened and have thick necrocaseous exudates (Figure 11.22). Felty fungal mats may be seen lining cavitated lesions or at the lung– air interface, the color of which may depend on the offender. Paecilomyces lilacinus and Fusarium have been repeatedly implicated in crocodilian mycotic pneumonias. Case reports of fungal disease in crocodilians usually describe simultaneous disease in more than a single individual, which suggests some predisposing factor linked with husbandry. The aquatic lifestyle of crocodilians seems to favor some opportunistic pathogens like Fusarium solani and Paecilomyces lilacinus, and underline the need for meticulous water hygiene. The ecological niche for the CANV has not been identified, but CANV infections in file snakes, tentacled snakes, and saltwater crocodiles are a testimony to its presence in the aquatic milieu.

11.5.3 Mycoses in the Squamata One of the latest developments in reptile clinical mycology is the recognition of the CANV as an agent of mycosis in squamates. This fungus has emerged as a major fungal pathogen of reptiles, particularly squamates, but may not be as much a truly emerging disease as one that has been repeatedly misdiagnosed in the past for a Trichophyton, Geotrichum, other Chrysosporium, Malbranchea, or even a Trichosporon infection. For example, a Trichophyton isolated from Malagasy

day geckoes (Phelsuma abbotti abbotti) and a Chrysosporium queenslandicum isolated from a garter snake (Thamnophis sp.) were correctly identified as the CANV after the case reports were published (Schildger et al., 1991a; Vissiennon et al., 1999). Unfortunately, none of the arthroconidiating fungi (and others) from older case reports are available for reexamination and confirmation of the initial identification, and therefore these case reports need to be interpreted cautiously.

11.5.3.1 Sauria  Dermatomycoses appear more frequently than systemic mycoses in lizards (Figures 11.39–11.40). This may simply reflect a bias, as skin lesions are more obvious to owners. Lesions can be easily mistaken for bacterial or benign proliferative epidermal papillomas. Dermatomycoses in lizards sometimes affect multiple individuals (Cork and Stockdale, 1994; Gartrell and Hare, 2005; Martinez-Silvestre and Galán, 1999; Schildger et al., 1991a). Captive geckos are well represented in the literature (Cork and Stockdale, 1994; Gartrell and Hare, 2005; Schildger et al., 1991a).   Of particular interest is the emergence of a clinicopathologically well-defined syndrome in bearded dragons (Pogona vitticeps) called yellow fungus disease, which will be addressed later in this chapter. This contagious dermatomycosis is increasingly widespread, and the CANV was demonstrated as the causative agent in all the cases that were investigated mycologically. An experimental challenge of veiled chameleons (Chamaeleo calyptratus) using a chameleon CANV isolate was recently conducted, and can serve as a useful model for dermatomycosis in lizards and possibly snakes (Paré et al., 2006a). The challenge demonstrated the ability of the CANV to act as a primary pathogen, and disclosed early hyphal proliferation in the epidermal corneum prior to downward penetration of hyphae into the dermis. Scarification of the skin simulated a breach in cutaneous integrity and resulted in an increase in the rate of infection (Paré et al., 2006a). As in snakes, prior case reports of dermatomycosis in lizards in which arthroconidiating fungi were incriminated need to be interpreted cautiously. Case reports of primary systemic infections in lizards are sparse. This may indicate a relative resistance of lizards to the common airborne spores of Aspergillus and Paecilomyces, or a lack of reporting. 11.5.3.2 Ophidia   A review of the literature suggests that dermatomycoses are more common than systemic mycoses in snakes. Dermatomycotic lesions may appear as blisters, focal hyperkeratosis and necrosis, or may be ulcerative in nature (Figures 11.41–11.47). Vesicular or bullous lesions are ephemeral and may progress to ulcers or epidermal necrosis before they are noticed by owners. Ventral scutes are particularly affected in most ophidian dermatomycoses. Fungal disease should be listed among the differential list of diagnoses whenever bullae or vesicular lesions are seen in a snake. A syndrome called necrotizing mycotic dermatitis was described in snakes (Jacobson, 1980). Various 

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species of snakes displayed necrotic skin areas, usually but not always involving ventral scutes or ventrolateral scales. Epidermal necrosis and fungal invasion were consistent histologic findings, but a variety of fungal isolates were recovered from affected snakes.   In several reports of dermatomycoses in snakes, multiple individuals were affected (Bertelsen et al., 2005; Branch et al., 1998; Miller et al., 2004; Nichols et al., 1999; Rossi and Rossi, 2000), implying either a common source of infection, deficiencies in husbandry, or both. There is scientific and anecdotal evidence to suggest that CANV infection, reviewed later in this chapter, is highly contagious, at least in chameleons, brown tree snakes, tentacled snakes, and bearded dragons. Subcutaneous mycetomas are reported with some regularity in snakes (Figures 11.14–11.16), particularly colubrids, and especially in corn snakes (Elaphe guttatta guttatta) (Figure 11.46) (Austwick and Keymer, 1981; Frye and Williams, 1995; Funk, 2002; Jacobson, 1980; Sigler and Paré, unpublished data). The CANV was confirmed as the cause in one corn snake and in an eastern milk snake (Lampropeltis triangulum triangulum) (Sigler and Paré, unpublished data). There is a possibility that the Geotrichum (Jacobson, 1980a), the Chrysosporium (Austwick and Keymer, 1981), and the Malbranchea (Funk, 2002) isolated from corn snake mycetomas were misidentified CANV isolates. Dermatomycoses in snakes may well be underdiagnosed. Lesions may be identical to those of blister disease, a poorly defined syndrome usually attributed to poor husbandry and bacterial disease. Often, only when the lesions worsen or fail to regress after a course of antibiotics, is the diagnosis of bacterial dermatitis revised and fungal infection considered. Similarly, lesions resulting from thermal injury also bear resemblance to those of mycotic dermatitis. Finally, fungal involvement should be suspected in snakes that either fail to undergo ecdysis for a protracted amount of time, or in some cases where there is increased frequency of shedding associated with skin disease. Infection may be secondary to some endocrine disturbances leading to accumulation of unshed epidermal layers and proliferation of keratinophilic fungi within those layers (Figure 11.7). Fungal infections of the spectacle and cornea occur occasionally in snakes (Colette and Curry, 1978; Zwart et al., 1973) (Figures 11.48–11.49). Aspergillus and Fusarium have been incriminated. Cases of systemic mycoses in snakes are sparser. Most initially involve the lungs and disseminate with a fatal outcome. Aspergillus species, and Paecilomyces lilacinus in particular, have been repeatedly incriminated in primarily systemic mycotic infections in ophidians.

11.6 Agents of Mycoses 11.6.1 Hyalohyphomycotic Agents 11.6.1.1 CANV and Other Chrysosporia  Chrysosporium species are readily isolated from normal reptile skin. In one

study, chrysosporia were retrieved from 26% of the sampled reptiles (Paré et al., 2003). The most common species isolated was C. zonatum, but other chrysosporia were also recovered (Paré et al., 2003). Clinically, the most important Chrysosporium species for reptiles is the CANV. This fungus is an ascomycetous teleomorphic fungus in the order Onygenales, which is known from only two isolates: the type isolate described in 1970 from the lungs of a lizard and another from soil in California (Paré et al., 1997). Over the last decade, more than 20 isolates of a fungus morphologically identical to the CANV have been deposited in the University of Alberta Microfungus and Herbarium (UAMH) collection (Figure 11.25) (Sigler and Paré, unpublished data). All but one of these isolates were cultured from lesions in various reptiles and strongly implicated as the causative agent. The phylogeny within this group of isolates is being investigated molecularly, but each of these isolates is currently cataloged as the CANV. The CANV is a fungus that is white or very slightly yellow in culture, with a wooly to powdery surface (Sigler, 2003). It has limited thermotolerance and does not grow at 37°C, except for the bearded dragon isolates (see yellow fungus disease in this chapter). Microscopically, the combined presence of pyriform or clavate, sessile or terminal aleurioconidia, and hyphae fragmenting into alternate or chained arthroconidia is characteristic of the CANV. Isolates were reliably incriminated as the cause of hyalohyphomycosis in a Parson’s chameleon (Calumma parsonii), a jewel chameleon (Furcifer lateralis), a Jackson’s chameleon (Chamaeleo jacksonii) (Paré et al., 1997), a panther chameleon (Furcifer pardalis) (Heckers, 2004), veiled chameleons (Chamaeleo calyptratus) (Paré et al., 2006b), day geckos (Phelsuma sp.) (Sigler and Paré, unpublished data), bearded dragons (Bowman and Paré, unpublished data), brown tree snakes (Boiga irregularis) (Nichols et al., 1999), corn snakes (Sigler and Paré, unpublished data), an Eastern milk snake and a Pueblan milk snake (Lampropeltis triangulum campbelli) (Sigler and Paré, unpublished data), a garter snake (Sigler and Paré, unpublished data), a ball python (Python regius) (Sigler and Paré, unpublished data), two file snakes (Acrochordus arafura) (Sigler and Paré, unpublished data) (Figure 11.50), tentacled snakes (Erpeton tentaculatum) (Bertelsen et al., 2005) (Figures 11.51–11.52), and saltwater crocodiles (Crocodylus porosus) (Thomas et al., 2002). This fungus may have a wide distribution as infections have been documented in North America, Europe, and Australia. The number of cases of mycoses caused by the CANV is substantial, especially in light of a survey that suggested this fungus was found on the skin of less than 1% of healthy captive squamate reptiles. This is in contrast with Aspergillus, Penicillium, or Paecilomyces, all extremely common on the skin of reptiles. Because the CANV is not described in mycology textbooks, isolates were always misidentified prior to being correctly recognized as the CANV (Nichols et al., 1999; Paré et al., 1997; Thomas et al., 2001; Vissiennon et al., 1999). The CANV is still repeatedly mistaken for fungi in the genus Trichophyton or for another

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Chrysosporium species based on the similarity of the aleurioconidia, or for a Geotrichum or Malbranchea because of the arthroconidia. The CANV should be suspected in any hyalohyphomycosis in squamates and in crocodiles. Infection with this fungus often leads to a fatal outcome. In most instances, lesions of CANV infection are cutaneous. They may appear as small vesicles or bullae that collapse and become encrusted and brown. They may also appear hyperkeratotic at first, with subsequent necrosis and sloughing to expose raw dermis. In crocodiles, leathery white coalescing plaques were described on the skin. If left untreated, they can progress and involve the dermis and subjacent musculature or bones. Progression may occur very rapidly, but is sometimes protracted. Infection may even extend transmurally to coelomic membranes and internal viscera (Figure 11.11). In some cases there is massive, apparently unchecked hyphal proliferation right through the dermis and musculature, and in other cases the fungus evokes a granulomatous response. Typically, hyaline septate hyphae are 2 to 4 µm wide, mostly parallel walled with occasional swellings and sparse haphazard branching (Figure 11.53). Often, dense tufts of arthroconidiating hyphae are found at the epidermal surface and are extremely suggestive of the CANV (Figure 11.54). The presence of arthroconidia dismisses other hyaline nonarthroconidiating Ascomycetes, such as fusaria, aspergilli, Paecilomyces, and Penicillium species. In published cases of CANV infection, there was always more than one animal affected, as if the infection were contagious. Experimental infection of veiled chameleons with the CANV demonstrated that skin infection can be established by exposing animals to CANV spores. The rate of infection is increased if spores are applied directly to the skin, and even higher when cutaneous integrity is breached. In this infection trial, a reliable reptile fungal infection model was created. All four of Koch’s postulates were met for CANV infection in veiled chameleons.  There is strong anecdotal evidence to suggest that CANV is highly contagious among bearded dragons (Johnson, 2004; Bowman and Paré, unpublished data). Bearded dragons develop recalcitrant cutaneous infection that slowly progresses to a systemic infection if untreated. Unshed, yellowish exuvia pieces remain adhered to the skin, and the condition has been named yellow fungus disease (YFD). Numerous commercial breeders and suppliers of bearded dragons have been affected by YFD. Lesions may start out as hyperkeratotic plaques that undergo necrosis, turn a darkish brown, and slough. The skin may be swollen, hard, and cracked with exudate seeping from the fissures. Lesions are often found around the mouth (Figure 11.55), but are usually multifocal and may occur anywhere on the body (Figures 11.56–11.58). A whole limb may be affected. Histologically, in bearded dragons, extensive granulomatous inflammation of the dermis and subjacent tissue is characteristic. The CANV elements within these granulomas often morphologically evoke yeasts rather than a mold. Arthroconidia at the surface may help in establishing a diagnosis. Culture of the CANV from squames

over the lesions or ideally from a biopsy of a lesion is diagnostic. Although other fungi might have been implicated in YFD, each of the cases investigated mycologically proved to be caused by the CANV (Bowman and Paré, unpublished data; Sigler and Paré, unpublished data). All bearded dragon isolates are morphologically identical to other CANV isolates, but differ in that they grow, albeit slowly, at 37°C. The source of infection in cases involving CANV infections has not been identified, and its ecology remains to be defined. Other Chrysosporium species, such as C. queenslandicum and C. keratinophilum, as well as unspeciated chrysosporia, have been very occasionally incriminated in reptile mycoses (Austywick and Keymer, 1981). These case reports need to be interpreted cautiously as they might have been CANV infections, prior to the CANV being recognized. For example, the Chrysosporium queenslandicum isolate from a case report of mycosis in a garter snake (Vissiennon et al., 1999) was found to be a CANV isolate upon reexamination of the fungus. Unfortunately, other Chrysosporium isolates from dated case reports are not available for study. There are reports of CANV in humans. The CANV was isolated from a brain abscess in an HIV-infected human in Nigeria (Steininger et al., 2005), and from a groin lesion in an immunosuppressed man in California (Sigler and Paré, unpublished data). The source of infection was unknown.

11.6.1.2 Paecilomycosis  Paecilomycosis refers to cutaneous or systemic hyalohyphomycoses caused by any fungus in the genus Paecilomyces. Paecilomyces species are best known for their entomopathogenicity. In humans and dogs, P. variotii may cause opportunistic infections, but Paecilomyces lilacinus is the most common cause of paecilomycosis (Summerbell, 2003). It is a ubiquitous saprophyte found in such diverse substrates as soil, decaying vegetable matter and foodstuff, insects, a variety of moist or viscous substances, and water (Summerbell, 2003). In reptiles it is a repeat offender, solidly implicated as the agent of mycoses in loggerhead sea turtles (Austwick and Keymer, 1981; Hernandez-Divers et al., 2002), green sea turtles (Chelonia mydas) (Austwick and Keymer, 1981), a hawksbill sea turtle (Eretmochelys imbricata) (Posthaus et al., 1997), Fly River turtles (Carettochelys insculpta) (Lafortune et al., 2005) (Figures 11.31–11.32), Aldabra tortoise (Dipsochelys dussumieri [formerly Geochelone gigantea]) (Heard et al., 1986) (Figures 11.59–11.60), a Greek tortoise (Testudo graeca) (Schildger et al., 1991b), a yellow-legged tortoise (Geochelone [formerly Testudo] denticulata) (Bemmel et al., 1960), an ornate slider (Trachemys ornata [formerly Pseudemys ornatus]), (Austwick and Keymer, 1981), saltwater crocodiles (Crocodylus porosus) (Davis 2001; Hibberd and Harrower, 1993; Maslen et al., 1988), a Nile crocodile (Crocodylus niloticus) (Austwick and Keymer, 1981), American alligators (Alligator mississippiensis) (Austwick and Keymer, 1981), spectacled caimans (Caiman crocodilus [formerly Caiman sclerops]) (Austwick and Keymer, 1981), New Zealand spotted sticky-toed geckos (Hoplodactylus maculatus) (Cork

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and Stockdale, 1994), a shingleback skink (Tiliqua [formerly Trachydosaurus] rugosus) (Schildger et al., 1991b), an African rock python (Python sebae) (Austwick and Keymer, 1981), and a timber rattlesnake (Crotalus horridus) (Paré, unpublished data). Paecilomyces lilacinus may also have been involved with outbreaks of mycotic pneumonia in juvenile mariculture-raised green sea turtles (Jacobson et al., 1979) and of fungal dermatitis in wild pygmy rattlesnakes (Sistrurus miliarius barbouri) in Florida (Cheatwood et al., 2003). A case of nodular necrotizing pneumonia in an Aldabra tortoise attributed to P. fumosoroseus (Georg et al., 1962) was likely due to the morphologically similar P. lilacinus (Summerbell, 2003). In addition, Paecilomyces lilacinus was formerly Penicillium lilacinum, so that some of the cases of reptile mycosis in which an unspeciated Penicillium was incriminated (Austwick and Keymer, 1981; Frye, 1991) might actually have been caused by Paecilomyces lilacinus.  Paecilomyces lilacinus is a fast-growing, strong keratin decomposer (Summerbell, 2003), and along with Fusarium solani, is a common contaminant in the humid environment of crocodile pens where meat is used as a food source (Thomas et al., 2002). Fat from the food forms a film on the water, deposits on the floor of the tank, and serves as an organic substrate in which P. lilacinus thrives (Thomas, 2001). This may explain the relatively high rate of infection in crocodilians and aquatic chelonians and why skin from three different captive, healthy yellow-lipped sea kraits (Laticauda colubrina) grew profuse pure cultures of P. lilacinus. (Paré et al., 2003). Infections with P. lilacinus are opportunistic but often fatal. Lesions may be extensive and spread rapidly. Hyphae in tissue are 2.5 to 4 µm wide, branching, septate, parallel walled, and undistinguishable from those of other hyaline ascomycetous fungi like Fusarium, Acremonium, and Penicillium, unless typical conidiophores are seen at the tissue– air interface. Fungal mats covering pulmonary tissue or air sacs may harbor a distinctive purplish to rosy hue suggestive of P. lilacinus.

11.6.1.3 Fusariomycosis  Fusariomycosis refers to hyalohyphomycoses caused by any fungus in the genus Fusarium. Histologically, hyphae are indistinguishable from those of other agents of hyalohyphomycosis unless additional features, such as macroconidia, are present at the surface. Fusaria are cosmopolitan soil saprobes and are common contaminants. Some species are opportunistic plant pathogens and many contain elaborate toxins. In humans, fusariosis caused by F. oxysporum, F. solani, F. verticilloides (includes the former species F. moniliforme), and F. proliferatum has emerged as a significant concern in immunosuppressed individuals. Fusarium solani and F. oxysporum are common causes of onychomycosis and keratitis in humans. Fusaria are commonly incriminated as pathogens in reptiles (Figures 5.70, 5.72, 11.61–11.63). Fusarium solani, F. oxysporum, F. verticilloides, and F. incarnatum (which now includes F. semitectum) have most often

been identified in cases of fusariomycosis. In one survey, fusaria were isolated from 4% of reptile skins, but were not speciated (Paré et al., 2003). Fusarium species are also cultured from the gut (Enweani et al., 1997) and lungs (Gugani and Okafor, 1980) of healthy lizards.  Fusarium solani is associated with soil and plants. It is also known from ponds, rivers, sewage facilities, water pipes, and is common in the marine environment (Cabañes et al., 1997). Not surprisingly, it seems to be the major Fusarium species involved in mycoses of aquatic reptiles. It was the fungus most frequently isolated from farmed saltwater crocodile hatchlings dying of cutaneous and disseminated mycoses in Queensland (Hibberd and Harrower, 1993). Fusarium solani was also isolated from dead-in-the-egg crocodile embryos on that same farm. It has been incriminated twice as the cause of dermatomycosis in sea turtles, both times in loggerheads (Austwick and Keymer, 1981; Cabañes et al., 1997). Fusarium solani was isolated from the sand at the bottom of the holding tank housing one affected turtle (Cabañes et al., 1997). Lesions in these turtles were described as white and scaly (Cabañes et al., 1997) or ulcerative (Austwick and Keymer, 1981). Fusarium solani is also a cause of cutaneous and systemic mycoses in farmed saltwater crocodiles in Australia (Buenviaje et al., 1994; Davis, 2001; Hibberd and Harrower, 1993). It is commonly isolated from crocodile nesting material and egg shells (Davis, 2001) and has been isolated from dead-in-the-egg crocodile embryos (Hibberd and Harrower, 1993). It was a major cause of disease in farms where winter was colder and daylight exposure was reduced (Buenviaje et al., 1994). Dense fungal mats, with a fluid-filled center, were described in the lungs of affected crocodiles (Buenviaje et al., 1994). Fusarium solani infection has also been documented in terrestrial reptiles, including a radiated tortoise (Geochelone radiata) with stomach ulcers (Austwick and Keymer, 1981) and a carpet python (Morelia spilota) with extensive tail necrosis, progressing to severe ulcerative necrotizing colitis and subsequent fatal fungemia (Holz and Slocombe, 2000). Fusarium oxysporum has a widespread distribution in soils and causes disease in a variety of plants (Summerbell, 2003). In human medicine, it is by far the most common Fusarium involved in onychomycosis and is frequently involved in keratitis (Summerbell, 2003). It may also cause serious infections in severely immunocompromised patients (Summerbell, 2003). Interestingly, F. oxysporum was isolated from the lacrimal fluid of a boa constrictor (Boa constrictor) (Migaki et al., 1984) and was the cause of infection of the spectacle in a rainbow boa (Epicrates cenchria maurus) that required enucleation (Zwart et al., 1973). Other F. oxysporum infections in reptiles include a flap-necked chameleon (Chamaeleo dilepis) with chronic skin lesions (Austwick and Keymer, 1981), a European pond turtle (Emys orbicularis) with nodular skin lesions (Schildger et al., 1991b), and a red-bellied black snake (Pseudechis porphyriacus) with a  swollen, necrotic tail that progressed to systemic infection (Holz and Slocombe, 2000).

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Necrotizing scute disease (NSD) is a fungal condition affecting captive and free-ranging Texas tortoises (Gopherus berlandieri) and possibly other chelonians caused by Fusarium incarnatum (formerly F. semitectum) (Rose et al., 2001) (Figure 11.64). It is a slowly progressing shell disease in which degradation and necrosis of the epidermal lamellae translate into whitish or discolored carapacial blemishes. The disease is disfiguring but appears to remain confined to the carapacial scutes, does not disseminate, and is not fatal. NSD affects free-ranging, presumably immunocompetent reptiles, and predisposing husbandry factors have not been identified for affected captive tortoises. The disease was reproduced experimentally in Texas tortoises, but could not be induced in ornate box turtles (Terrapene ornata). Other fusaria may sometimes express pathogenicity in reptiles. Fusarium verticilloides (formerly F. moniliforme) was solidly established as the cause of death in an alligator with severe necrotizing fungal pneumonia (Frelier et al., 1985). Unspeciated fusaria are also listed among less substantiated cases of mycoses in reptiles (Austwick and Keymer, 1981; Jacobson, 1980; Migaki et al., 1984).

11.6.1.4 Aspergillus and Penicillium  Aspergillus and Penicillium are two closely related genera of common fungal contaminants. There are over 200 fungal species within the genus Aspergillus, most of which are ubiquitous in the soil, dust, decomposing matter, and in the air. In humans, several Aspergillus species act as opportunistic pathogens, but A. flavus, A. fumigatus, and A. niger account for more than 95% of infections (Richardson and Kokki, 2003). In a skin survey of captive squamate reptiles, Aspergillus species were isolated from 69% of sampled reptiles, second only to Penicillium, and it was not uncommon to culture two or even three different species of Aspergillus from a single skin sample (Paré et al., 2003). Aspergillus species are also part of the gastrointestinal mycobiota and were readily isolated from the lungs and intestines of agamas and wall geckos (Enweani et al. 1997; Gugagni and Okafor, 1979). This underlines the need for histopathology if an Aspergillus isolate from a reptile with mycosis is to be incriminated. As an example, cutaneous lesions from three different Jackson’s chameleons all yielded the same Aspergillus flavus, yet dermatophilosis was diagnosed histologically and there was no fungal involvement (Paré, unpublished data). Culture may not be essential to confirm aspergillosis if typical conidiophores at the air–tissue interface are identified in tissue sections. Dichotomously branching, parallel-walled, septate vegetative hyphae in tissue are suggestive of Aspergillus, but numerous other fungi may appear similar and the diagnosis should ideally be supported by culture or immunohistochemistry. Aspergilli in tissues demonstrate angiotropism and tend to disseminate easily. The high prevalence of opportunistically pathogenic Aspergillus on and in reptiles would suggest it commonly causes mycosis, yet cases in which this fungus was reliably incriminated are few. Interestingly, most reported cases of

reptile aspergillosis are dated, and there are very few cases in the recent literature. Aspergillus mycoses in reptiles, as with other taxa, involve mostly the lower respiratory tract. Granulomatous or nodular pneumonia is typical, along with fungal mats over the surface of the trachea, air sacs, or coelomic membranes. Dissemination to the liver, spleen, and other viscera may occur. Three Galapagos tortoises (Geochelone nigra [formerly elephantopus]) and an Aldabra tortoise died with severe fungal pneumonia attributed respectively to Aspergillus amstelodami (now A. hollandicus) and an unspeciated Aspergillus (Austwick and Keymer, 1981). Giant tortoises appear overrepresented in cases of fungal pneumonia in the literature, suggesting some predisposition. Aspergillosis was described in another chelonian, a Hilaire’s side-necked turtle (Phrynops, [formerly Hydraspis] hilarii), with disseminated, nodular, caseating pneumonia (Austwick and Keymer, 1981). Pneumonia and dermatitis due to Aspergillus fumigatus or A. ustus occurred in captive 2- to 6-week-old alligators from a zoological collection. Infections were linked with poor water quality and inadequate holding facilities. Hygiene and sanitary corrective measures eliminated the problem. Aspergillosis in squamates is reported rather infrequently. Nodular Aspergillus pneumonia occurred in a black tegu (Tupinambis teguixin [formerly nigropunctatus]) (Austwick and Keymer, 1981), and disseminated nodular lesions due to Aspergillus were described in a puff adder (Bitis arietans) and a banded water snake (Nerodia [formerly Natrix] sipedon). Cutaneous lesions in a corn snake and two North African spiny-tailed lizards (Uromastyx acanthinurus) might have been caused by A. flavus, A. oryzae, and A. tamarii (Schildger et al., 1991b), but those reports are not well substantiated. More convincing is the account of an Aspergillus terreus infection in two captive San Esteban, or painted, chuckwallas (Sauromalus varius) (Tappe et al., 1984). The lizards had necrotic, burn-like areas over the head and thorax. Lesions on the toes of one lizard progressed to gangrene. Biopsies revealed hyperkeratosis and ulceration, along with coalescing granulomas in the dermis and around vessels. Lesions were refractory to treatment and both animals eventually died. Dissemination had occurred to the lungs and to other viscera. Immunofluorescence confirmed aspergillosis, and the aleurioconidia observed on the hyphae were most consistent with A. terreus. A case of granulomatous pneumonia with fungemia and multifocal granulomatous dermatitis due to Aspergillus niger in a Western diamondback rattlesnake (Crotalus atrox) (Boyer and Garner, 2004) further illustrates the pathogenic potential of aspergilli in reptiles. Penicillium species are cosmopolitan and are the most common environmental molds in regions of temperate climate. Penicillium and Paecilomyces share morphological similarities, including penicillate conidiophores, which may have led to misidentification in the past. Furthermore, Paecilomyces lilacinus was formerly included in the genus Penicillium and might have been the true cause of older cases of reptile

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mycoses attributed to unspeciated Penicillium. Penicillium species can be cultured as often as 78% of the time from skin samples of healthy lizards and snakes, and were the most common fungi found on the skin of reptiles in a recent survey (Paré et al., 2003). It was also readily isolated from the gut and internal organs of lizards (Enweani et al., 1997; Gugagni and Okafor, 1979). Histological substantiation of any case of suspected Penicillium infection is therefore essential. A captive Aldabra tortoise in the Canary Islands died of systemic, disseminated, nodular mycosis (Oros et al. 1996) (Figures 11.65–11.67). Lesions were most prominent around the heart. Penicillium brevicompactum was isolated from lesions and compatible hyphae were disclosed histologically within necrotic granulomas. The veracity of the only case of P. brevicompactum infection in a human is disputed based on the proven inability of P. brevicompactum to grow at 37°C (Summerbell, 2003), but reptile body temperature is obviously more variable. An unspeciated Penicillium was isolated from cutaneous lesions in wild Bocage’s wall lizards (Podarcis bocagei) in Spain (Martinez-Silvestre and Galán, 1999). Lesions were most often distributed over the ventral skin and occurred in the winter months and regressed in the spring. Hyphae consistent with Penicillium were present in biopsies of the lesions. Penicillium also was isolated (Figure 11.69) from ventral skin lesions (Figure 11.70) in a Texas rat snake (Elaphe obsolete lindheimeri) (Jacobson, unpublished findings). Fungi also were seen in a tissue section of a biopsied scale. Fatal nodular mycotic hepatitis with intralesional fungi, and from which an unspeciated Penicillium has been repeatedly isolated, has been described in a group of captive Jackson’s chameleons (Figure 11.68). There were no lesions in the intestine of affected animals and the source of infection was unclear (Potter J, personal communication). The condition awaits further investigation as histologic morphology of fungal elements is more reminiscent of yeast, and Candida has also been isolated from the nodules. There is a definite need for solid cases of reptile aspergillosis and penicilliosis to be reported, and for isolates to be duly speciated and deposited so that we better understand the pathogenic potential of such prevalent environmental fungi.

11.6.1.5 Geotrichosis  Geotrichum candidum is a cosmopolitan, ubiquitous ascomycetous mold that may be confused with yeasts because of the cream colored, yeastlike colonies on agar. This fungus is a commensal that carries a very low opportunistic pathogenic potential in humans and animals, and may have been confused with other fungi in the past (Hazen and Howell, 2003). It is a commensal that can be readily isolated from reptile intestinal contents (Gugagni and Okafor, 1979; Raiti, 1998; Ruiz et al., 1980) and cutaneous samples (Paré et al., 2003). Short arthroconidia are heavily produced, and the potential for misidentification of a CANV or a Trichosporon is substantial, and case reports of infection in reptiles with this fungus again need to be interpreted cau-

tiously. This is perhaps best illustrated by the reidentification as the CANV (UAMH 9832) of the Geotrichum and Chrysosporium queenslandicum isolates that were coincriminated in a case of mycosis in a garter snake (Vissiennon et al., 1999). There is some experimental data to suggest that Geotrichum candidum is an opportunistic agent of mycosis in reptiles, or at the very least in chelonians. It was isolated from skin lesions and the kidneys of a Galapagos tortoise with dermatomycosis and renal lesions detected histologically (Ruiz et al., 1980). Experimental infection of chelonians with Geotrichum has yielded conflicting results. Local infection was demonstrated in common snapping turtles (Chelydra serpentina) and red-eared sliders at the site of injection with Geotrichum candidum inocula (Sinclair and El-Tobshy, 1969), while in another study the authors were unable to experimentally induce geotrichosis in four Russian tortoises (Testudo horsfieldii) (Ruiz et al., 1980). Geotrichum candidum has also been incriminated in a northern water snake (Nerodia [formerly Natrix] sipedon) with subcutaneous abscesses, and has been linked with mycotic dermatitis in green iguanas (Iguana iguana) (Wissman and Parsons, 1993), a Burmese python (Python molurus) (Abou-Gabal and Zenoble, 1980), carpet pythons (Morelia spilota variegata) (McKenzie and Green, 1976), and a corn snake (Jacobson, 1980) (Figures 11.46, 11.71). In the two latter reports, the histopathology and epidemiology is quite reminiscent of CANV infection in snakes (Nichols et al., 1999; Sigler and Paré, unpublished data). Deposition of reptilepathogenic Geotrichum isolates would go a long way toward unraveling or substantiating the role of G. candidum in reptile mycoses.

11.6.1.6 Zygomycosis  Zygomycoses are infections caused by any of the fungi in the Phylum Zygomycota. The term zygomycosis has replaced the term phycomycosis, now obsolete. Phycomycetes included algae and oomycetes, which have since been transferred out of the Fungi. Zygomycetes are ubiquitous and are found in a wide variety of substrates, such as decaying vegetables, foodstuffs, fruits, soil, and excreta (Dromer and McGinnis, 2003). There are two orders within the Phylum Zygomycota: the Mucorales, to which belong the familiar genera Mucor, Rhizopus, Rhizomucor, Absidia, as well as the lesser known Syncephalastrum and Cunninghamella, and the Order Entomophthorales, which includes Basidiobolus ranarum, Conidiobolus coronatus, and C. incongruus, the causes of entomophthoromycoses in humans. The morphology in the tissues of fungi in these two orders is very different and easy to distinguish. Of the two orders within the Phylum, fungi within the Mucorales have caused most documented cases of zygomycosis in reptiles. 11.6.1.6.1 Order Mucorales, Mucormycoses  The term mucormycosis encompasses mycoses caused by any of the Mucorales. Mucorales are very common environmental fungi. In a study of the cutaneous mycobiota of healthy captive

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squamates, zygomycetous fungi were common on the skin of healthy reptiles (Paré et al., 2003). Mucor was cultured from shed skin of 19% of 127 reptiles sampled, but other zygomycetes, such as Syncephalastrum racemosum, Rhizopus spp., and Cunninghamella spp., were also frequently isolated (Paré et al., 2003). Most mucoraceous zygomycetes are fast-growing fungi, which along with aspergilli and penicillia, may quickly overshadow the growth of more clinically relevant fungi from a specimen unless cycloheximide is added to the culture medium. Mucor isolates can be implicated only as the cause of mycosis if the morphology of fungal elements in tissue is compatible with that of a zygomycetous fungus. Hyphae of mucoraceous zygomycetes are typically broad, thin walled, and pleomorphic, varying in width from 5 to 20 µm, and are aseptate or poorly septate, with irregular, rightangle branching (Chandler and Watts, 1987). The thin-walled zygomycetous hyphae collapse in tissue sections and often appear wrinkled and twisted or convoluted (Chandler and Watts, 1987). Cases of mucormycoses in reptiles are relatively rare, and even rarer are cases in which the Mucor isolate was speciated. Of 30 adult Marlborough green geckos (Naultinus manukanus) collected from Stephens Island, New Zealand, and maintained in captivity, five developed multiple black powdery cutaneous lesions and white skin nodules (Gartrell and Hare, 2005). Timely ecdysis was curative for two geckos, but the disease was fatal for the three others. In two lizards, there was necrosis of digits and fungal osteomyelitis of phalangeal bones. Wide, branching, aseptate hyphae consistent with those of zygomycetes were present in histological sections of lesions and Mucor ramosissimus, a fungus that rarely causes mucormycosis in humans (Schipper and Stalpers, 2003), was isolated from skin biopsies (Gartrell and Hare, 2005). Mucormycosis has been reliably described in Florida soft-shelled turtles (Apalone [formerly Trionyx] ferox) (Jacobson et al., 1980) and wood turtles (Glyptemys [formerly Clemmys] insculpta) (Lappin and Dunstan, 1992). In both cases, affected animals were hatchlings kept in overcrowded conditions, and necrotizing lesions were confined to the shell and skin. One case of cutaneous zygomycosis was also described in an eastern indigo snake (Drymarchon corais couperi) with a small, indurated mid-dorsal cutaneous lesion in which typical zygomycetous hyphae were disclosed histologically (Werner et al., 1978), but the actual causative agent was not determined. Mucor was isolated from skin lesions in a bearded dragon and a Timor monitor (Varanus timorensis), and from the lungs of a group of crocodilians with pneumonia, but histologic morphology of hyphae was not described and causality therefore not established (Schildger et al., 1991b). A mycotic necrotizing colitis with intussusception in a Jackson’s chameleon (Chamaeleo jacksoni) was associated with an unspeciated Mucor (Shalev et al., 1977). Mucoraceous hyphae were also disclosed histologically in the colonic and cloacal mucosa of a veiled chameleon (C. calyptratus) with a history of recurrent cloacal prolapse (John Roberts, personal com-

munication) (Figures 11.72–11.73). Members of the Zygomycota may elicit little inflammation, and it can be difficult to determine whether they are the cause of necrotic lesions, or mere invaders of devitalized tissue. A case of phycomycosis in a red milk snake (Lampropeltis triangulum syspila) with nodules along the intestinal wall and trachea (Sindler et al., 1978), and another case of phycomycosis in a massasauga rattlesnake (Sistrurus catenatus) with severe facial fungal infection (Williams et al., 1979) (Figure 11.10) were described in which fungal elements in tissue were morphologically more consistent with an ascomycetous or hyphomycetous than a zygomycetous fungus. Culture was not performed in either case. 11.6.1.6.2 Order Entomophtorales, Entomophthoromycoses  Reptiles are known intestinal carriers of Basidiobolus sp., a genus of fungi that causes disease in humans and possibly anurans (Enweani et al., 1997; Feio et al., 1999; Gugagni and Okafor, 1980; Okafor et al., 1984), yet in the literature there is only a single case of reptile basidiobolomycosis reported, in which Basidiobolus ranarum was isolated from a localized lesion in the corner of the mouth of an Aldabra giant tortoise (Migaki et al., 1984). This suggests some innate resistance of reptiles to basidiobolomycosis. A different zygomycetous fungus appears to be a serious pathogen of snakes. It is histologically compatible with an entomophthoraceous fungus, and was first demonstrated within cutaneous and disseminated granulomatous lesions in a gopher snake (Pituophis melanoleucus) and a copperhead (Agkistrodon contortrix), but was not isolated or characterized further (Jessup and Seely, 1981). A previously reported case of a corn snake with cutaneous algal infection, presumed to be Prototheca, was likely caused by this entomophthoraceous fungus (Crispens and Marion, 1975). A very similar organism, very possibly the same, was observed histologically in a corn snake with cutaneous lesions, two timber rattlesnakes with stomatitis and disseminated nodular lesions, and a gopher snake with a large mass in the oral cavity (Kaplan et al., 1983). Infections of other timber rattlesnakes with this fungus have been seen (Figures 11.74–11.75) (Jacobson, unpublished findings). This zygomycetous fungus, probably belonging to a new taxon within the Order Entomophtorales, was isolated in pure culture from one timber rattlesnake, but did not sporulate and could not therefore be speciated (Kaplan et al., 1983). Further investigations have suggested this fungus belongs to a novel genus and species, Schizangiella serpentis (Humber 1989; Humber, personal communication). The ecology of this fungus is unknown.

11.6.1.7 Beauveriosis and Metarhizium anisopliae  Beauveria bassiana and Metarhizium anisopliae are widespread, highly entomopathogenic fungi that are the cause of white and green muscardine, respectively, in a wide variety of insects. Both are very rare agents of fungal keratitis in humans, while M. anisopliae has also been incriminated in

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a single human case of disseminated infection in a leukemic child, two cases of human sinusitis, and one case of rhinitis in a cat. The pathogenic potential of B. bassiana is very limited in homeotherms as it can grow only at temperatures below 30°C, while Metarhizium can grow at temperatures up to 40°C. Neither fungi was retrieved from the intestines of largely insectivorous lizards (Enweani et al., 1997; Gugagni and Okafor, 1980), and Metarhizium, but not Beauveria, was cultured from less than 1% of skin samples from normal squamate reptiles (Paré et al., 2003). Based on the literature, spontaneous infection with these two fungi are limited to the chelonians and crocodilians, although M. anisopliae experimentally induced disseminated disease in two subcutaneously challenged Madeira wall lizards (Lacerta dugesii) and one intracoelomically challenged red-eared slider (Austwick and Keymer, 1981). Experimental oral administration of M. anisopliae var acridum spores to fringe-toed lizards (Acanthodactylus dumerili) (Peveling and Demba, 2003) caused a fungal granuloma in one of them.   Beauveria bassiana has caused fatal pneumonia in two captive American alligators (Fromtling et al., 1979). Death followed hibernation, during which mechanical failure of the heating system occurred for one-half day and the temperature dropped 8°C below desired. Air exchange was also minimal in the hibernation enclosure. Damage to the lungs was multifocal and severe. Excessive mucus, lung red hepatization, and filamentous white fluffy lesions surrounded by black consolidated parenchyma were described. Microscopically, all bronchi and alveolar spaces were obstructed with hyphae. Dissemination to the liver and spleen had occurred. Fatal B. bassiana was reported in a Galapagos and an Aldabra tortoise with pulmonary abscesses in the former and necrotic lung areas in the latter (Austwick and Keymer, 1981). A pond slider (Trachemys scripta) recently imported into Spain from Cuba died of B. bassiana pneumonia (Gonzaléz-Cabo et al., 1995). Pulmonary congestion and pleuritis, with scattered yellowish granulomatous nodules were described. Intrapulmonary experimental infection of two box turtles (Terrapene carolina) with B. bassiana spores resulted in the death of the turtle kept at 15.5°C, while the exposed turtle maintained at 22°C remained normal. This is consistent with the isolation of B. bassiana from three Kemp’s ridley turtles with disseminated mycotic disease following cold stunning (Turnbull et al., 2000). Spontaneous M. anisopliae infection has only been reported in one alligator. The lungs contained miliary nodules and bullous cavitated lesions lined with dark green fungal mats. The tongue was also affected (Austwick and Keymer, 1981).

11.6.1.8 Acremonium Infections  Cephalosporiosis refers to infections involving fungi in the genus Cephalosporium. However, the genus Cephalosporium is no longer valid, as it was transferred to the genus Acremonium (De Hoog et al., 2000; Summerbell, 2003), and cephalosporiosis is therefore

an obsolete term. Acremonium species are common environmental fungi that are generally regarded as low-grade opportunistic pathogens, often following traumatic inoculation. In humans, the relatively rare cases of Acremonium infections are usually caused by A. falciforme, A. recifei, and A. kiliense (Summerbell, 2003). In a recent survey, acremonia were isolated from the skin of 13% of the reptiles sampled (Paré et al., 2003). One isolate from a piece of exuvium of a Solomon Island skink (Corucia zebrata) represented a new species and was named A. exuviarum (Sigler et al., 2004). Acremonium mycoses are rare in reptiles. Infection with an unspeciated Acremonium was documented in spectacled caimans (Caiman crocodilus [formerly Caiman sclerops]) under the older appellation of cephalosporiosis (Trevino, 1972). A fatal diffuse granulomatous mycotic pneumonia with focal necrotizing hepatitis was diagnosed in these three 6-month-old caimans. Slender conidiophores, 2- to 4-µm-wide hyphae, cigar-shaped conidia, and chlamydospores were present histologically, which were morphologically consistent with Acremonium, and an Acremonium sp. was cultured from swabs of the pulmonary nodules.   An unspeciated Acremonium, then described as Cephalosporium, was isolated from a tumorous tracheal nodule in a ringed (grass) snake (Natrix natrix, formerly Tropidonotus natrix) (Austwick and Keymer, 1981). Hyphae were seen histologically. These are the only two reported cases of presumptive Acremonium infection in reptiles.

11.6.1.9 Others  A multitude of other hyaline fungi, including Scopulariopsis, Myriodontium, and Trichoderma species, have been implicated in disease in man (Digagni et al., 2003; Summerbell, 2003). These and others have been isolated from normal reptile skin (Paré et al., 2003), and could carry some pathogenic potential for reptiles under certain conditions. Only if lesions are cultured and isolates properly identified will we begin to determine which fungal genera have the potential to cause disease in reptiles. Some well-substantiated cases of reptile mycoses were caused by unusual fungi, not typically associated with any pathogenic potential for vertebrates. Chrysonilia sitophila (formerly Monilia sitophila), a cosmopolitan soil saprobe considered nonpathogenic (St. Germain and Summerbell, 1996), was reasonably linked with multifocal epidermal lesions in a captive black rat snake (Elaphe obsoleta) (Dillberger and Abou-Gabal, 1979). Colletotrichum acutatum, a plant pathogen, was established as the cause of disseminated mycosis in a Kemp’s ridley sea turtle (Manire et al., 2002) (Figures 11.76–11.77). This turtle was a juvenile recuperating from cold stunning. These two cases, among others, demonstrate how reptiles that otherwise survive adverse conditions are often immunocompromised, and later succumb to microbes of low pathogenicity. Chelonians were found to be experimentally susceptible to infection by Lacazia loboi (formerly Loboa loboi), the agent of lobomycosis (Sampaio et al., 1971). Lobomycosis is a chronic infection of the skin and subcutis of humans and

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river dolphins (Inia geoffroyi), which is endemic to South America, particularly the Amazon River basin. Infection probably occurs following traumatic inoculation. Grossly, the lesions in humans are plaque-like, verrucose, ulcerated, or keloid (Hay, 2003) and usually remain static. Histologically, a granulomatous reaction surrounds thick double-walled, hyaline, 5- to 12-µm budding yeast-like cells arranged in chains and connected by tube-like isthmuses (Chandler and Watts, 1987; Woods and Schnadig, 2003). The fungus has never been grown in culture, but recent molecular analysis strongly suggests it is an Ascomycete in the Order Onygenales (Howard, 2003). Three red-footed tortoises (Geochelone carbonaria), 3 yellow-legged tortoises (Geochelone denticulata), and 10 scorpion mud turtles (Kinosternon scorpioides) were injected subcutaneously with material from a human patient (Sampaio et al., 1971). All developed histologically confirmed lesions, and unusual or atypical forms of the fungus were present in the lesion of one yellow-legged tortoise. Chelonians may represent a good experimental model for lobomycosis.

11.6.2 Dematiaceous Fungi Dematiaceous fungi consist of a heterogeneous group of genera that are darkly pigmented due to the presence of melanin in the cell walls of hyphae, conidia, or both (Sanche et al., 2003). The presence of melanin, with very few exceptions, allows for easy identification of fungal elements in routine H&E-stained tissue sections. Most infections with these fungi occur following traumatic inoculation in the skin or subcutis. Infections with pigmented fungi are classified in three main categories: chromoblastomycoses, mycetomas, and phaeohyphomycoses (Sanche et al., 2003). Neither dematiaceous mycetomas nor chromoblastomycosis has been documented in reptiles.

11.6.2.1 Phaeohyphomycosis  The term phaeohyphomycosis has been broadened to include all mycotic infections characterized by the presence of pigmented septate hyphae, yeast cells, pseudohyphae-like elements, or any combination thereof, in tissue sections (Sanche et al., 2003). In humans, agents of phaeohyphomycosis include, to name a few, fungal species in the genera Exophiala, Wangiella, Hormonema, Dreschlera, Curvularia, Chaetomium, Phoma, Alternaria, and Cladosporium (Sanche et al., 2003; Schell, 2003). Many of those, especially Cladosporium, Chaetomium, and Alternaria, can be readily isolated from normal reptile skin as they are normal constituents of the cutaneous mycobiota (Paré et al., 2003). A Cladosporium species was even isolated twice from the lungs of otherwise healthy agamas (Gugagni and Okafor, 1980). Histological confirmation of phaeohyphomycosis is needed to substantiate pathogenicity attributed to such common fungi.   Ochroconis humicola (formerly Scolecobasidium humicola) was isolated from granulomatous, superficially ulcerated, papular foot lesions in an eastern box turtle (Terrapene

carolina carolina) (Weitzman et al., 1985). Lesions were also present on the tail. Dematiaceous hyphae were demonstrated in a KOH preparation of a biopsy as well as histologically. Ochroconis humicola has limited thermotolerance and does not grow at temperatures of 36°C or higher (Weitzman et al., 1985), but can be cultured from the skin of healthy reptiles (Paré et al., 2003). Exophiala jeanselmei was cultured from a cavitated phaeohyphomycotic lesion involving the limb of another eastern box turtle, but there was no evidence of dissemination at necropsy (Shreve et al., 2004). A foot lesion was probably the portal of entry (Shreve et al., 2004). Pigmented fungi were observed in shell lesions of a box turtle and a granulomatous lesion of the foot of another box turtle (Reavill et al., 2004). Cultures were not performed in either case. The strong representation of box turtles in the scant literature on reptile phaeohyphomycosis suggests a relative predisposition of this species to pigmented fungi, although a bias stemming from their popularity as pets cannot be excluded. A radiated tortoise with a mycetomatous mass in the oral cavity was diagnosed at necropsy with disseminated granulomatous mycosis (Frank, 1976). Pigmented hyphae were present in the granulomas. An Exophiala (formerly Hormodendrum) species was cultured, but was lost before it could be speciated (Frank, 1976). Focal phaeohyphomycosis was also described in a mangrove snake (Boiga dendrophila) with a subcutaneous intermandibular mass (Figures 11.78–11.79) (Jacobson, 1984). Fungal granulomatous lesions were identified histologically in part of the mass, while the remainder of the lesion was diagnosed as a fibrosarcoma. Metastasis had occurred to the liver, heart, spleen, and kidney. Cultures were negative and the fungus could not be identified. Phaeohyphomycosis was diagnosed in a Galapagos tortoise (Manharth et al., 2005). The tortoise was initially presented with uniocular lesions consisting of buphthalmia and a posterior chamber density. The eye was enucleated and phaeohyphomycosis was diagnosed. The condition of the tortoise declined, it was euthanized, and disseminated phaeohyphomycosis was seen. Culture and DNA sequencing identified the fungus as a member of the genus Exophiala. In other reported cases of phaeohyphomycosis, a causative relationship is not well demonstrated for the putative fungus that was isolated. A phaeohyphomycotic stomatitis in a green anaconda (Eunectes murinus) was documented (Marcus, 1971) where the diagnosis of Cladosporium infection was ventured based on the morphology of the pigmented fungal elements, but not confirmed. Ulocladium botrytis, Trichoderma pseudokoningii, and Cladosporium herbarum were respectively isolated from a Russian tortoise (Agrionemys [formerly Testudo] horsfieldii) with dermatitis, a mangrove monitor (Varanus indicus) with fungal pneumonia and hepatitis, and a red-eared slider with dermatitis, but there was no histopathological description of the lesions (Schildger et al., 1991b). Sporothrix schenckii, a cosmopolitan saprobe found in soil and vegetal matter, is the agent of cutaneous sporotricho-

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sis in humans and mammals. It can be found on the skin of healthy reptiles (Paré et al., 2003). It was cultured from two of many pigmy rattlesnakes with extensive cutaneous lesions (Cheatwood et al., 2003) and tentatively linked with the seasonal epizootic of dermatitis observed in these snakes. However, this filamentous fungus converts to a budding yeast at 35 to 39°C or in tissue, and hyphae in the superficial epidermis are only rarely seen histologically (Chandler and Watts, 1987; Schell, 2003; Woods and Schnadig, 2003). This yeast morphology is in contrast with the description of hyphae within granulomatous skin and visceral lesions in affected rattlesnakes.

11.6.2.2 Chromoblastomycosis  Chromoblastomycosis, also called chromomycosis, is a specific form of phaeohyphomycosis caused by a small, defined group of fungi consisting of Fonsecae pedrosoi, F. compacta, Cladophialophora (Cladosporium) carrioni, Phialophora verrucosa, Rhinocladiella aquaspersa, Exophiala jeanselmei, and E. spinifera (Sanche et al., 2003). By definition, chromoblastomycosis is characterized histologically by the presence of round, 5- to 12-µm diameter, chestnut brown, thick-walled, partitioned structures called sclerotic bodies or muriform cells, surrounded by chronic granulomatous reaction. These sclerotic bodies, once thought to be budding yeasts, are currently believed to represent a fungus arrested between the yeast and hyphal morphology because of some tissue factors (Sanche et al., 2003). Pigmented hyphae are occasionally present in the epidermis. Sclerotic bodies were not a feature of the Exophiala jeanselmi phaeohyphomycotic infection in the box turtle, nor were they described in the mangrove snake. The agents of chromoblastomycosis are soil saprobes. Lesions usually develop initially on the distal lower extremities in people, and on the ventrum in amphibians, because these body parts are in contact with soil. Reptiles live close to the ground, and the ventrum of snakes is in permanent contact with soil. Furthermore, reptiles are common in tropical and subtropical areas where chromoblastomycosis typically occurs, yet there are no documented cases of chromoblastomycosis in reptiles, suggesting some innate resistance of this class of vertebrates to fungi causing chromoblastomycosis.

11.6.3 Yeast Yeast differs from molds in that they are unicellular organisms that reproduce by budding (blastoconidiation) (Larone, 1995). Some yeasts, including Candida species, can form true hyphae but typically produce blastoconidia, which may elongate and remain attached to the parent cell, forming chains called pseudohyphae. Yeast colonies are typically smooth, glabrous, and cream colored. Most yeasts are anamorphic, although a few produce ascospores (Larone, 1995). Classification is based on morphology, but also largely on biochemical tests using commercial systems. New molecular methods are being used that may establish phylogenetic relationships

of the various yeasts to phyla within the Fungi. In reptiles, yeast infections are typically caused by Candida species, but mycoses with Trichosporon spp. and Cryptococcus neoformans have also been reported.

11.6.3.1 Candidiasis  Candidiasis refers to fungal infections caused by yeasts in the genus Candida. The classic oropharyngeal mucosal Candida infection is also commonly called thrush. Gastrointestinal candidiasis is most often seen, but Candida infection of the integumentary and respiratory systems, among others, also occur. Spread to the liver from intestinal lesions is not uncommon. Candidemia is rare in animals but rather common in immunocompromised human patients. Oval, 3- 6-µm budding cells with pseudohyphae, and occasionally hyphae, are seen in affected tissue.   Candida is one of the most commonly isolated fungi in laboratories. Candida yeasts, particularly C. albicans, are ubiquitous in the environment and are normal commensals of the cutaneous, intestinal, and respiratory mycobiota of humans and animals. In human medicine, a number of nonalbicans Candida species, including C. glabrata, C. guilliermondi, C. krusei, C. lusitaniae, C. parapsilosis, and C. tropicalis, have emerged as agents of disease in immunocompromised patients. Speciation of clinical isolates is extremely relevant clinically as some of these species show innate resistance to commonly used antifungal drugs (e.g., fluconazole). Candida species can readily be cultured from the skin and internal organs of reptiles (Kostka et al., 1997). Yeasts are highly prevalent in the gut of all reptiles, and were isolated from the intestines of 100% of the 38 chelonians sampled (Kostka et al., 1997). Yeasts were also readily isolated from the lungs, liver, and kidneys of turtles, lizards, and snakes. Other surveys of reptile mycobiota concur (Enweani et al., 1997; Gugagni and Okafor, 1979). Candida was the most common yeast genus cultured from reptiles, but Trichosporon, Torulopsis, and Rhodotorula species were also retrieved (Kostka et al., 1997). In reptiles, generic candidiasis has most often been diagnosed histologically, based on the morphology of fungal elements (budding, pseudohyphae, etc.). In many cases, a histologic diagnosis of candidiasis was not substantiated by culture, so the causative yeast might have belonged to another genus. In one survey of reptiles for yeasts, C. tropicalis was isolated twice as often as C. albicans from cultured material (Kostka et al., 1997), and it should therefore not be assumed that unspeciated Candida isolates from reptile mycotic lesions belong to C. albicans. Of 23 Candida isolated from tortoises, 13 belonged to C. tropicalis, 9 to C. albicans, and 1 to C. parapsilosis (Milde et al., 2000). According to the literature, reptile Candida infections are rare compared to such infections in mammals and birds. It may be that reptile candidiasis is underreported. A compilation of necropsy cases described lesions in a number of reptiles in which yeasts were identified, but failed to identify the agent in any of them (Reavill et al., 2004). Because some filamentous fungi can adopt a yeastlike morphology in tis-

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sue, it is difficult to draw conclusions on the incidence of candidiasis among reptiles necropsied merely on the basis of histological morphology. Typical C. albicans esophageal thrush was described in a crocodile tegu (Crocodilurus lacertinus) (Austwick and Keymer, 1981). Oronasal ulcers in a radiated tortoise (Geochelone radiata), and swelling of the oral mucosa with pustules and with miliary lesions in the liver of a smooth snake (Coronella austriaca) were attributed to C. albicans (Austwick and Keymer, 1981). Candida tropicalis was loosely linked to rhinitis, tachypnea, and steatorrhea in a Hermann’s tortoise (Testudo hermanni) (Schildger et al., 1991b). The better-documented and substantiated cases of candidiasis involved a Greek tortoise (Testudo graeca) with pneumonia (Hernandez-Divers, 2001) and a stranded loggerhead sea turtle with enteritis following ingestion of a fishing line (Orós et al., 2004). Candida albicans was isolated from both these animals. The tortoise exhibited moist audible rales, oral discharge, weakness, and decreased appetite. There was a thick, cloudy discharge at the glottis, and the right lung was diffusely radiopaque. Cytology of a transcarapacially collected pulmonary lavage revealed yeasts and inflammatory cells and culture of the fluid yielded a heavy growth of C. albicans (Hernandez-Divers, 2001). In the stranded loggerhead, enteritis was linked to intestinal mucosal damage caused by an ingested fishing line. Using an immunofluorescent antibody (IFA) technique, a fungus present in the inflamed duodenal mucosa and submucosa was identified as C. albicans. The fungal elements in tissue sections were strongly stained with a polyclonal antibody and a monoclonal antibody against C. albicans. This paucity of well-substantiated reports underlines a true need for solid reptile Candida infection case reports.

11.6.3.2 Cryptococcosis  Cryptococcosis is a disease caused by the ubiquitous basidiomycetous yeast Cryptococcus neoformans, which typically manifests as a fungal pneumonia that sometimes disseminates, usually to the central nervous system (Viviani et al., 2003). Much remains to be known about its ecology, but Cryptococcus neoformans var neoformans is found in pigeon droppings worldwide, while C. neoformans var gatti is associated with certain species of eucalyptus. In tissues, uninucleate, thin-walled, spherical, oval to elliptical, 2- to 20-µm diameter yeasts with narrowbased budding are characteristic. In immunocompetent mammals, the yeasts evoke a focal suppurative and necrotizing inflammatory reaction that often evolves to granulomatous inflammation (Viviani et al., 2003). Cryptococcosis is rare in reptiles, but has been documented in an eastern water skink (Eulamprus quoyii) (Hough, 1998) and an anaconda (Eunectes murinus) (McNamara et al., 1994) (Figures 11.80–11.82). The anaconda presented with neurological signs and had classic granulomatous cryptococcal pneumonia and meningoencephalitis, which was confirmed using a Cryptococcusspecific IFA test. In contrast, the skink presented with a small,

discrete swelling over the dorsum that was surgically excised and cryptococcosis was diagnosed histologically.

11.6.3.3 Trichosporonosis  Fungi in the basidiomycetous genus Trichosporon produce true hyphae, but are treated as yeasts on the basis that they also produce pseudohyphae. Trichosporon species are normal components of the soil biota, sometimes colonize the oropharynx and skin of humans (Maenza and Merz, 2003), and may be cultured from the intestines and feces of healthy and sick reptiles (Gugagni and Okafor, 1979; Kostka et al., 1997; Raiti, 1996). The genus has recently undergone extensive revision, and the species Trichosporon beigelii is no longer valid, having been reassigned to a novel species (De Hoog et al., 2000; Maenza and Merz, 2003). Isolates of T. beigelii involved in human clinical disease have been reexamined, and those associated with deep or disseminated infections have now been identified as T. asahii and T. mucoides, while those associated with superficial skin infections belong to T. asteroides, T. ovoides, and T. inkin. Trichosporon species are not uncommon on the skin of normal reptiles and were cultured from the skin in 9% of surveyed squamates (Paré et al., 2003). Few reptile Trichosporon isolates have been examined using the newer speciation criteria, but one cutaneous isolate from the Paré survey and another isolate from the skin of an O’Shaughnessy’s chameleon (Calumna oshaughnessyi) were T. asahii (Jean Paré, unpublished data), while in a different survey focusing on the intestinal flora, T. inkin and T. cutaneum were recovered.  In reptiles, trichosporonosis has been described mostly as an opportunistic carapacial infection in chelonians (Schildger et al., 1991b). Trichosporon may also have caused fatal infection in three banded rock rattlesnakes (Crotalus lepidus klauberi) that died with systemic, disseminated disease (Reddacliff et al., 1993). Histologically, yeasts were often observed intracellularly. Trichosporon cutaneum (formerly a synonym of T. beigelii) was isolated from conspecific bite-induced subcutaneous hematomas in American anoles (Anolis carolinensis) (Jacobson, 1991) (Figure 11.83). Trichosporon has also been incriminated in severe dermatitis in a North African spiny-tailed lizard (Uromastyx acanthinurus) (Schildger et al., 1991b), lung abscesses and intestinal ulcers in an unspeciated aquatic South American snake (Austwick and Keymer, 1981), oral lesions in an Aldabra tortoise (Austwick and Keymer, 1981), a Nile crocodile (Migaki et al., 1984), and a spectacled caiman (Migaki et al., 1984). However, Trichosporon is a common environmental contaminant, and is a heavily arthroconidiating fungus, so confusion is possible with other fungi such as Geotrichum and the CANV. Care must therefore be exercised when reviewing and interpreting reptile trichosporonosis in the literature.

11.6.4 Dimorphic Fungi Dimorphic fungi are important human pathogens. Dimorphic fungi are so called because they are filamentous in the envi-

Mycotic Diseases inof Reptiles  543

ronment, yet adopt a yeasty morphology in tissues. Blastomycosis, histoplasmosis, and coccidioidomycosis are caused by Blastomyces dermatiditis, Histoplasma capsulatum, and Coccidioides immitis, respectively. Dimorphic fungi do not appear important in reptiles. A survey for Histoplasma capsulatum in wildlife failed to recover the fungus from 18 reptiles collected in areas where mammals were repeatedly diagnosed with histoplasmosis (Menges et al., 1967). Coccidioides immitis is a mold endemic to deserts and arid regions of the southwestern United States and areas of Central and South America. When airborne conidia are inhaled and deposit in the lungs, they form large round, thick-walled 10- to 80-µm spherules that mature to contain characteristic endospores. Spherules are produced in tissues at 37°C, which may explain why coccidioidomycosis has been reported in only a single reptile, a Sonoran gopher snake (Pituophis melanoleucus affinis) (Timm et al., 1988). In this snake there were discrete, round, white, 3- to 4-mm diameter nodules scattered across the hyperemic pulmonary parenchyma. Microscopically, these were composed of macrophages, giant cells, lymphocytes, and heterophils around clusters of the abovedescribed spherules. This snake, presumably exposed to C. immitis in the wild prior to capture, was also diagnosed with a hemogregarine infection and a pancreatic adenocarcinoma at necropsy, and both may have contributed to its demise (Timm et al., 1988). Experimentally, intracoelomic inoculation of lizards with C. immitis has yielded conflicting results (Egeberg, 1985; Swatek and Plunkett, 1957), while intratracheal inoculation of rattlesnakes failed to produce disease (Egeberg, 1985). Two additional unpublished cases of coccidioidomycosis have been seen in reptiles, one in a Gila monster (Heloderma suspectum) (Figures 11.84–11.85) and one in a Texas indigo snake (Drymarchon corais errebanus) (Figures 11.86–11.87) (Jacobson, unpublished findings). Coccidioidomycosis should be listed among the differential list of diagnoses in xerophilic reptile species with pulmonary disease.

11.6.5 Dermatophytes Dermatophytosis in reptiles is very rare, if it occurs at all. Trichophyton terrestre was isolated from lesions in a group of blue-tongued skinks (Tiliqua scincoides) experiencing progressive digital necrosis (Hazell et al., 1985), but causality was unclear, as the examined tissue was necrotic and fungi may have been secondary invaders. Furthermore, T. terrestre is one of the Trichophyton for which the CANV has been mistaken. In fact, in several documented cases of infection in reptiles with the Chrysosporium anamorph of Nannizziopsis vriesii (CANV), the isolates were initially misidentified as Trichophyton. A severe dermatomycosis affecting day geckoes was attributed to a Trichophyton (Schildger et al., 1991a), but the isolate was later positively identified as the CANV (UAMH 6610). This undermines the veracity of reported Trichophyton

infections, especially when left unspeciated. A Trichophyton sp. isolated from two green anacondas with dermal lesions (Miller et al., 2004) might have been the CANV. This assumption is supported by the well-documented cutaneous and systemic CANV infection in tentacled snakes (Bertelsen et al., 2005) and the known presence of the CANV in the aquatic milieu. The Trichophyton isolate from the anacondas was not available for reexamination. When their cutaneous mycobiota was studied, no Trichophyton species were isolated from 127 healthy captive squamate reptiles, while two Microsporum boullardii and one M. gypseum, both geophilic species, were recovered (Paré et al., 2003). Microsporum cookei, another geophilic species, was cultured from the skin of a python in a survey in Australia (Rees, 1967). There is currently insufficient evidence to support the occurrence of dermatophytosis in reptiles.

11.7 Conclusion There is a need for reptile clinicians and pathologists to insist on fungal isolation and accurate identification. Proper identification carries serious clinical implications for patient management and selection of antifungal drugs. The herpetological literature, even that within the last two decades, is replete with case reports where the fungal organism causing the lesions is not cultured or identified. Such cases are of limited value to the clinician and add little to the already accepted concept that reptiles are susceptible to a variety of fungal infections. Even less desirable are cases where identification of the causative fungus is equivocal, having been risked on the basis of nonspecific morphological characteristics of the fungus in tissue sections. Rarely is the histologic morphology of a fungus sufficiently characteristic as to allow identification to genus, and exceptionally to species. Incorrect identification leads to erroneous assumptions and scientific incongruities. Even when proper techniques for collection, transport, and submission of specimens to the laboratory are followed, culture may fail to yield a fungus when it is clearly present histologically. Reasons for this are unclear, but may include antifungal treatment, bacterial overgrowth, or improper culture medium or temperature. In such cases, however, every effort should be made to identify the fungus in tissue sections using immunohistochemical techniques or molecular probes. Keying isolates to genus and species is difficult, and fungi cultured from reptile material may belong to species with which clinical laboratories are unfamiliar. When unsure, the laboratory should forward the isolate(s) to a reference mycology laboratory. Identification should be carried out to species in all cases. Well-substantiated cases of mycoses in any reptile should be published in a peer-reviewed journal, as such cases are few and far between. If an isolate is clearly demonstrated to be the agent of mycosis in a reptile, every effort should be made for the fungus to be deposited in a microfungus collection for preservation, for use in experimental infection trials,

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or simply for reexamination should its identification be challenged later or should taxonomical changes occur.

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Figure 11.1  Chuckwalla, Sauromalus obesus. Iguanidae. Culture plates with different fungi growing. (Upper right) Sabouraud agar inoculated 3 days previously with fragments of unshed skin collected from the tail of an otherwise healthy chuckwalla. Note the heavy growth of a white fungus, identified as a Fusarium sp., around all skin fragments. (Lower right) Sabouraud agar inoculated 3 days previously with fragments of unshed skin collected from the same lizard, but from the head. Note the growth of various different fungi around shed skin.

Figure 11.2  Garter snake, Thamnophis sirtalis. Colubridae. Photomicrograph showing a pigmented fungus in the epidermis. H&E stain.

Figure 11.3  African spurred tortoise, Geochelone sulcata. Testudinidae. Photomicrograph showing numerous hyphae in the keratin of a plastronal scute. GMS stain.

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Figure 11.4  Chinese alligator, Alligator sinensis. Alligatoridae. Photomicrograph showing numerous branching hyphae in a major air passageway. Courtesy of the Pathology Service, College of Veterinary Medicine, University of Florida, Gainesville. PAS stain.

Figure 11.5  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Photomicrograph showing CANV hyphae (arrows) in the epidermal stratum corneum. No inflammatory cells are present. PAS stain.

Figure 11.6  Water moccasin, Agkistrodon piscivorus. Viperidae. The exuvium contains fungal material (arrows) that has penetrated from the outer to inner surface.

Figure 11.7  King snake, Lampropeltis getula. Colubridae. This snake had diffuse fungal skin disease and would shed every one to two weeks. When it started shedding, it would immediately enter another cycle of ecdysis. (From Jacobson ER. 1991. Diseases of the integumentary system of reptiles, in Dermatology for the Small Animal Practitioner, Nesbitt, GH, Ackerman, LJ (eds) Veterinary Learning Systems, Trenton, NJ, pp. 225–239. With permission.)

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Figure 11.8  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Photomicrograph showing CANV infection associated with liquefaction necrosis of the epidermal stratum corneum and multifocal coagulation necrosis in the stratum spinosum and germinativum. PAS stain.

Figure 11.9  Eastern indigo snake, Drymarchon corais couperi. Colubridae. Photomicrograph showing hyphae (arrows) within the caseated center of a subcutaneous granuloma. GMS stain.

Figure 11.10  Massasauga rattlesnake, Sistrurus catenatus. Viperidae. Photomicrograph of a cross-section of the anterior head. Multiple coalescing granulomas (GR) are seen displacing subcutaneous tissues and the left olfactory lobe (OL). H&E stain. (From Williams LW et al. 1979. Vet Med/Small Anim Clin 74:1182–1184. With permission.)

Figure 11.11  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. Photomicrograph of CANV infection in a veiled chameleon (Chamaeleo calyptratus). Transmural invasion of the coelomic (CE) cavity across the body wall. Hyphae are seen throughout the dermis, the abdominal muscles, and through the coelomic membrane. Note the intact epidermis (EP). PAS stain.

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Figure 11.12  Giant ameiva, Ameiva ameiva, Teiidae. A subcutaneous mass (mycetoma) is seen between the forelimbs (arrows).

Figure 11.13  Giant ameiva, Ameiva ameiva. Teiidae. Surgical removal of a subcutaneous mycetoma.

Figure 11.14  Western diamondback rattlesnake, Crotalus atrox. Viperidae. A subcutaneous mass (mycetoma) is seen near the tail. Courtesy of Sally Nofs.

Figure 11.15  Western diamondback rattlesnake, Crotalus atrox. Viperidae. Excised mass, histologically confirmed as a mycetoma. Courtesy of Sally Nofs. Figure 11.16  Gopher snake, Pituophis catenifer. Colubridae. A subcutaneous mycetoma is seen (arrows). The fungus did not invade muscle (M).

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Figure 11.17  Green turtle, Chelonia mydas. Cheloniidae. Multiple nodules are seen in the left lung of a juvenile green turtle. The left lung is atelectatic and the right lung is emphysematous. Nodules consisted of fungal granulomas. (From Jacobson ER et al. 1979. J Am Vet Med Assoc 175 929–933. With permission.)

Figure 11.18  Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph showing fungal hyphae and spores in an air passageway. PAS stain. (From Jacobson ER et al. 1979. J Am Vet Med Assoc 175 929–933. With permission.)

Figure 11.19  Green turtle, Chelonia mydas, Cheloniidae. Multiple nodules are seen in the liver. A fungus was identified using light microscopy.

Mycotic Diseases inof Reptiles  553

Figure 11.20  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph showing large numbers of hyphae and inflammatory cells in an air passageway. GMS stain.

Figure 11.21  Gopher tortoise, Gopherus polyphemus. Testudinidae. Photomicrograph showing large numbers of hyphae and inflammatory cells in an air passageway. GMS stain.

Figure 11.22  American alligator, Alligator mississippiensis. Alligatoridae. Lung of an alligator with severe pneumonia. Caseous material fills air passageways.

Figure 11.23  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph showing hyphae within granulomas in the lung. GMS stain.

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Figure 11.24  Bearded dragon, Pogona vitticeps. Agamidae. Biopsy of an oral fungal granulomatous mass.

Figure 11.25  Bearded dragon, Pogona vitticeps. Agamidae. Profuse CANV growth around sections of skin biopsies on Mycobiotic agar.

Figure 11.26  Green sea turtle, Chelonia mydas. Cheloniidae. Multifocal fungal skin lesions are seen.

Figure 11.27  Bolivian side-neck turtle, Platemys platycephala. Chelidae. The carapace has diffuse areas of shell disease. Fungi were identified in biopsies of the lesion.

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Figure 11.28  Florida softshelled turtle, Apalone (formerly Trionyx) ferox. Trionychidae. Mucormycosis. Multiple ulcerative lesions are seen on the shell. (From Jacobson ER et al. 1980. J Amer Vet Med Assoc 177:835–837. With permission.)

Figure 11.29  Florida soft-shell turtle, Apalone (formerly Trionyx) ferox. Trionychidae. Mucormycosis. Photomicrograph revealing epidermal ulceration and necrosis of the shell. H&E stain, (From Jacobson ER et al, 1980. J Amer Vet Med Assoc 177:835–837. With permission.)

Figure 11.30  Florida softshelled turtle, Apalone (formerly Trionyx) ferox. Mucormycosis. Trionychidae. Photomicrograph showing numerous hyphae within an area of epidermal necrosis. GMS stain. (From Jacobson ER et al, 1980. J Amer Vet Med Assoc 177:835– 837. With permission.)

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Figure 11.31  Fly River turtle, Carettochelys insculpta. Carettochelydae. Multiple ulcerations of the carapace can be seen (arrows). Paecilomyces lilacinus was cultured from the lesions.

Figure 11.32  Fly River turtle, Carettochelys insculpta. Carettochelydae. Photomicrograph of an ulcerative shell lesion. Numerous hyphae can be seen in the lesion. Paecilomyces lilacinus was cultured. GMS stain.

Figure 11.33  Loggerhead sea turtle, Caretta caretta. Cheloniidae. A stranded dead loggerhead sea turtle was necropsied and found to have a severe diffuse fungal pneumonia. Caseous material is seen throughout the lung. Courtesy of Bruce Homer.

Figure 11.34  Loggerhead sea turtle, Caretta caretta. Cheloniidae. A stranded dead loggerhead sea turtle with severe myocarditis. Hyphae were found throughout the myocardium. Courtesy of Bruce Homer.

Mycotic Diseases inof Reptiles  557

Figure 11.35  Hawksbill sea turtle, Cheloniidae. Eretmochelys imbricata. Cheloniidae. Fungal skin lesions (arrows) were seen in these neonates at the time of emergence from their eggs.

Figure 11.36  Saltwater crocodile, Crocodylus porosus. Crocodylidae. Juvenile crocodile with multifocal areas of necrosis along the jaws and within the oral cavity. Fungi were identified in this lesion. Courtesy of Jamie Hibberd.

Figure 11.37  Saltwater crocodile, Crocodylus porosus. Crocodylidae. Juvenile crocodile with extensive fungal infection of the gular skin. Courtesy of Jamie Hibberd.

Figure 11.38  Saltwater crocodile, Crocodylus porosus. Crocodylidae. Hind foot with extensive areas of necrosis. Fungi were found in this lesion. Courtesy of Jamie Hibberd.

558  Mycotic Diseases of Reptiles

Figure 11.39  Jeweled lacerta, Lacerta lepida. Lacertidae. Cutaneous fungal disease. Areas of brown discoloration were hyperkeratotic, necrotic, and colonized by fungi.

Figure 11.40  Desert iguana, Dipsosaurus dorsalis. Iguanidae. Cutaneous fungal disease. A large area of a hindlimb was discolored, hyperkeratotic, necrotic, and colonized by fungi.

Figure 11.41  Ball python, Python regius. Pythonidae. Areas of fungal skin disease are scattered about the integument. Some are ovoid (arrows) and others affect individual scales, which develop an orange discoloration (arrowheads).

Figure 11.42  Ball python, Python regius. Pythonidae. Photomicrograph showing hyphae within the epidermis. GMS stain.

Mycotic Diseases inof Reptiles  559

Figure 11.43  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. A subcutaneous fungal granuloma extends into the pit (arrows). Courtesy of Darryl Heard.

Figure 11.44  Eastern diamondback rattlesnake, Crotalus adamanteus. Viperidae. Photomicrograph of hyphae within a subcutaneous mass. GMS stain. Courtesy of Darryl Heard.

Figure 11.45  Corn snake, Elaphe guttata guttata. Colubridae. This snake was found in the field with a diffuse severe necrotizing fungal epidermitis and granulomatous dermatitis.

Figure 11.46  Corn snake, Elaphe guttata guttata. Colubridae. Corn snake with large subcutaneous masses bulging the overlying skin, which has areas of necrosis. Two masses have been biopsied.

560  Mycotic Diseases of Reptiles

Figure 11.47  Emerald tree boa, Corallus caninus. Boidae. Multiple fungal skin lesions are seen as areas of brown discoloration and epidermal necrosis. The spectacle and eye are also affected. Courtesy of Darryl Heard.

Figure 11.48  King snake, Lampropeltis getula. Colubridae. Fungal infection of the spectacle.

Figure 11.49  Eastern indigo snake, Drymarchon corais couperi. Colubridae. Fungal infection of the spectacle.

Figure 11.50  Arafura file snake, Acrochordus arafurae. Acrochordidae. The CANV was isolated from multiple small, circular, raised white epidermal lesions. Courtesy of Annette Thomas.

Mycotic Diseases inof Reptiles  561

Figure 11.51  Tentacled snake, Erpeton tentaculatum. Colubridae. The CANV was isolated from areas of epidermal necrosis. Courtesy of Mads Frost Bertelsen.

Figure 11.52  Tentacled snake, Erpeton tentaculatum. Colubridae. Septate hyphae are seen within areas of epidermal necrosis. The CANV was isolated from this lesion. GMS stain. Courtesy of Mads Frost Bertelsen.

Figure 11.53  Veiled chameleon, Chamaeleo calyptratus, Chamaeleonidae. CANV cutaneous infection. Hyphae have occasional, irregular branching. PAS stain.

Figure 11.54  Veiled chameleon, Chamaeleo calyptratus, Chamaeleonidae. CANV cutaneous infection. A dense tuft of arthroconidiating hyphae is present at the surface of the epidermis. GMS stain.

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Figure 11.55  Bearded dragon, Pogona vitticeps, Agamidae. Coalescing smooth, raised, hyperkeratotic yellowish lesions around the mouth of a lizard with yellow fungus disease. Courtesy of Jennifer Clarke.

Figure 11.56  Bearded dragon, Pogona vitticeps, Agamidae. Circular ulcerated cutaneous lesions over the forearm and elbow of a lizard with yellow fungus disease. Courtesy of Jennifer Clarke.

Figure 11.57  Bearded dragon, Pogona vitticeps, Agamidae. Epidermal hyperkeratosis and necrosis, and extensive ulceration to the dermis over the flank of a lizard with yellow fungus disease. Courtesy of Jennifer Clarke.

Figure 11.58  Bearded dragon, Pogona vitticeps, Agamidae. Dysecdysis, with retained yellowish exuvial morcels in a lizard with yellow fungus disease. Multifocal hyperkeratotic, erosive, and ulcerative lesions are seen over the lower abdomen, legs, and feet. Courtesy of Dan Johnson.

Mycotic Diseases inof Reptiles  563

Figure 11.59  Aldabra tortoise, Dipsochelys dussumieri (formerly Geochelone gigantea). Testudinidae. Photomicrograph of the stomach with an ulcerated area containing hyphae. Paecilomyces was cultured from lesions in this tortoise. GMS stain. Courtesy of Pathology Service, College of Veterinary Medicine, University of Florida, Gainesville.

Figure 11.60  Aldabra tortoise, Dipsochelys dussumieri (formerly Geochelone gigantea). Testudinidae. Photomicrograph of the omentum with hyphae within a granuloma. Paecilomyces was cultured from lesions in this tortoise. GMS stain. Courtesy of Pathology Service, College of Veterinary Medicine, University of Florida, Gainesville.

Figure 11.61  Burmese python, Python molurus bivittatus, Pythonidae. A linear area of necrosis is seen where the lateral scales join the ventral scales. Fusarium was cultured from this lesion.

Figure 11.62  Burmese python, Python molurus bivittatus, Pythonidae. Photomicrograph of a necrotizing skin lesion containing hyphae. Fusarium was cultured from this lesion. GMS stain.

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Figure 11.63  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. There is extensive ulcerative dermatomycosis over the flank. Conidia morphologically consistent with those of Fusarium were observed histologically at the epidermal surface. Courtesy of Arno Wunschmann.

Figure 11.64  Texas tortoise, Gopherus berlandieri. Testudinidae. White areas of the shell represent areas of necrosis. Fusarium semitectum (now F. incarnatum) was cultured from this lesion. Courtesy of Francis Rose.

Figure 11.65  Aldabra tortoise, Geochelone gigantea. Testudinidae. Multifocal granulomas are seen in the kidney. A fungus was identified in this lesion and Penicillium brevicompactum was cultured. Courtesy of Jorge Oros. (From Orós J et al. 1996. Vet Rec 139:295–296. With permission.)

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Figure 11.66  Aldabra tortoise, Geochelone gigantea. Testudinidae. Multifocal granulomas are seen in the liver. A fungus was identified in this lesion and Penicillium brevicompactum was cultured. Courtesy of Jorge Oros.

Figure 11.67  Aldabra tortoise, Geochelone gigantea. Testudinidae. Photomicrograph showing numerous hyphae in the liver. Penicillium brevicompactum was cultured from this lesion. Courtesy of Jorge Oros.

Figure 11.68  Jackson’s chameleon, Chamaeleo jacksonii, Chamaeleonidae. Multiple nodules are seen within the liver. Fungal elements were present histologically within the nodules and Penicillium was repeatedly isolated from tissues. Courtesy of John Potter.

566  Mycotic Diseases of Reptiles

Figure 11.69  Texas rat snake, Elaphe obsolete lindheimerii. Colubridae. Photomicrograph of Penicillium cultured from a ventral skin lesion. A fruiting body is seen. Lactophenol cotton blue stain.

Figure 11.70  Texas rat snake, Elaphe obsolete lindheimerii. Colubridae. Necrotizing skin lesion (arrows) on a ventral scale. Penicillium was cultured from this lesion.

Figure 11.71  Corn snake, Elaphe guttata guttata. Colubridae. Photomicrograph of multiple hyphae within a subcutaneous granuloma. Geotrichum was cultured from biopsied lesions. GMS stain.

Mycotic Diseases inof Reptiles  567

Figure 11.72  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. A colonic prolapse can be seen (arrows). A fungus consistent with Mucor was identified in the prolapse. Courtesy of John Roberts.

Figure 11.73  Veiled chameleon, Chamaeleo calyptratus. Chamaeleonidae. A fungus consistent with Mucor can be seen in an area of mucosal necrosis in a prolapsed colon. GMS stain. Courtesy of John Roberts.

Figure 11.74  Timber rattlesnake, Crotalus horridus. Viperidae. Photomicrograph of a large granulomatous mass between the esophagus and lung. Schizangiella serpentis was identified in this lesion. PAS stain.

Figure 11.75  Timber rattlesnake, Crotalus horridus. Viperidae. Photomicrograph showing Schizangiella serpentis in a granulomatous mass. This fungus is arranged as pairs, triads, and tetrads. PAS stain.

568  Mycotic Diseases of Reptiles

Figure 11.76  Kemp’s ridley sea turtle, Lepidochelys kempii. Cheloniidae. The kidney (arrows) contains numerous nodules consisting of fungal granulomas. Colletotrichum acutatum was cultured from these lesions. Courtesy of Charles Manire.

Figure 11.77  Kemp’s ridley sea turtle, Lepidochelys kempii. Cheloniidae. Photomicrograph of a hyphal structure within a renal granuloma. Colletotrichum acutatum was cultured from this lesion. Courtesy of Charles Manire.

Figure 11.78  Mangrove snake, Boiga dendrophila. Colubridae. Phaeohyphomycosis was identified in the large subcutaneous mass seen near the mandible of this snake. (From Jacobson ER. 1984. J Amer Vet Med Assoc 185:1428–1430. With permission.)

Figure 11.79  Mangrove snake, Boiga dendrophila. Colubridae. Photomicrograph showing brown-staining fungal organisms (arrows) within a granuloma. H&E stain.

Mycotic Diseases inof Reptiles  569

Figure 11.80  Anaconda, Eunectes murinus. Boidae. Collections of histiocytes are seen within the interstitium of the lung. Cryptococcus was identified in this lesion. H&E stain. Courtesy of Tracey McNamara.

Figure 11.81  Anaconda, Eunectes murinus. Boidae. Multiple encapsulated yeast consistent with Cryptococcus can be seen within histiocytes in the lung. H&E stain. Courtesy of Tracey McNamara.

Figure 11.82  Anaconda, Eunectes murinus. Boidae. Multiple encapsulated yeast consistent with Cryptococcus are seen within a granuloma in the kidney. H&E stain. Courtesy of Tracey McNamara.

Figure 11.83  American anole, Anolis carolinensis. Iguanidae. The large cervical swellings represent nuchal granulomas and hematomas. Trichosporon cutaneum was identified in this lesion. Courtesy of Darryl Heard.

570  Mycotic Diseases of Reptiles

Figure 11.84  Gila monster, Heloderma suspectum, Helodermatidae. Photomicrograph of the lung showing areas of necrosis containing numerous organisms consistent with Coccidioides immitis. H&E stain.

Figure 11.85  Gila monster, Heloderma suspectum, Helodermatidae. Photomicrograph of the lung showing spherules of Coccidioides immitis. H&E stain.

Figure 11.86  Texas indigo, Drymarchon corais erebennus. Colubridae. Photomicrograph of the kidney showing areas of necrosis and numerous organisms consistent with Coccidioides immitis. H&E stain.

Figure 11.87  Texas indigo, Drymarchon corais erebennus. Colubridae. Photomicrograph of Coccidioides immitis showing spherules containing endospores. H&E stain.

12 Parasites and Parasitic Diseases of Reptiles Elliott R. Jacobson

Contents 12.1 General Comments................................................ 572 12.2 Protozoans.............................................................. 572 12.2.1 Amoebae.................................................... 572 12.2.2 Parabasalia and Euglenozoa .................... 574 12.2.3 Apicomplexa: Nonhemoparasitic.............. 575 12.2.4 Apicomplexa: Hemoparasitic.................... 579 12.3 Microsporida.......................................................... 580 12.4 Myxozoa................................................................. 581 12.5 Cestoda................................................................... 581 12.5.1 Pseudophyllidea........................................ 581 12.5.2 Proteocephalidea....................................... 582 12.5.3 Trypanorhyncha........................................ 582 12.5.4 Cyclophyllidea........................................... 582 12.6 Trematoda.............................................................. 582 12.6.1 Ochetosomatidae and Plagiorchiidae....... 583 12.6.2 Spirorchiidae.............................................. 583 12.6.3 Diplostomatidae......................................... 584 12.6.4 Rhytidodidae............................................. 584 12.6.5 Hemiuridae................................................ 584 12.7 Nematoda............................................................... 584 12.7.1 Ascaridoidea.............................................. 584 12.7.2 Cosmocercoidea........................................ 586 12.7.3 Diaphanocephaloidea............................... 586 12.7.4 Filarioidea.................................................. 586 12.7.5 Dracunculoidea......................................... 587 12.7.6 Diplotriaenoidea........................................ 587

12.7.7 Gnathostomatoidea................................... 588 12.7.8 Spiruroidea................................................ 588 12.7.9 Physalopteroidea....................................... 588 12.7.10 Dioctophymatoidea................................... 588 12.7.11 Rhabditoidea.............................................. 588 12.7.12 Oxyuroidea................................................ 589 12.7.13 Trichinelloidea........................................... 589 12.8 Acanthocephala..................................................... 590 12.8.1 Chelonia..................................................... 590 12.8.2 Squamata................................................... 590 12.9 Pentastomida.......................................................... 590 12.9.1 Chelonia..................................................... 590 12.9.2 Crocodylia.................................................. 591 12.9.3 Squamata................................................... 591 12.10 Annelida................................................................. 592 12.10.1 Hirudinea................................................... 592 12.11 Crustacea .............................................................. 592 12.11.1 Cirripedia................................................... 592 12.12 Acari....................................................................... 593 12.12.1 Parasitiformes............................................ 593 12.12.2 Acariformes................................................ 595 12.13 Diptera.................................................................... 596 12.13.1 Chelonia..................................................... 596 12.13.2 Squamata................................................... 597 Acknowledgments............................................................ 597 References......................................................................... 597

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12.1 General Comments

12.2 Protozoans

Free-ranging reptiles are infected and infested with a great diversity of endo- and ectoparasites. Considering the number of papers published, relatively few reports link parasite burdens with morbidity or mortality events in wild reptiles. Historically, the scientists interested in reptilian parasites have been taxonomists, and have not been interested in or trained to appreciate the host’s response to the parasite. In most cases the reptilian host–parasite relationship is yet to be documented. Still, lesions have been seen with certain parasitic infections. Plasmodium infections are thought to affect behavior and physiology of certain populations of lizards (Schall, 1996); the coccidian Caryospora was responsible for mortality in captive (Leibovitz et al., 1978) and wild (Gordon et al., 1993) marine turtles, and spirorchiid trematodes and their eggs caused severe lesions in multiple visceral sites in marine turtles (Gordon et al., 1998). I have also evaluated wild reptiles found ill in the field having amoebiasis, cryptosporidiosis, and various helminth infections. Individual reptiles that become ill and die in the field are seldom encountered unless there is a major mortality event. In contrast, there are many more reports on parasitic diseases of captive reptiles, especially those that were collected in the wild. Whereas some parasites, such as the protistan Entamoeba invadens, are transmitted without an intermediate host (i.e., directly from animal to animal), many other reptilian parasites require such a host for completion of their life cycles. In most captive populations, infection with parasites requiring intermediate hosts is self-limiting. The life cycles of many reptilian parasites remain to be elucidated. As is the case with higher vertebrates, reptiles may harbor parasites for considerable lengths of time before showing signs of illness; clinical disease may be seen later, when predisposing factors compromise the host immunologically and allow the development of overt infection. Some parasitic infections are insidious in development and are expressed after a prolonged prepatent period. When the disease is eventually diagnosed in such cases, the host may be beyond treatment. In this chapter, those clinically significant parasitic diseases described in the literature and those parasitic infections that I have encountered in reptiles will be reviewed. Where information is available, lesions due to infection will be described. In most instances, particularly for the helminths, specific morphological criteria for categorizing parasites will not be given. Identification of protozoans and metazoans, both intact and within tissues, can be found elsewhere (Barnard and Upton, 1994; Gardiner and Poynton, 1999; Gardiner et al., 1998). For taxonomic information on ticks infesting reptiles, see Barnard and Durden (2000). There is no single similar source of information on mites infesting reptiles. Classifications used in this chapter follow those in other texts (Anderson, 2000; Krantz, 1978; Lee et al., 2000; Marquardt et al., 2000; McDaniel, 1979).

While many protozoans have been described from peripheral blood films (Figures 12.1–12.8) of reptiles, the majority of these organisms do not seem to cause disease, and reflect a well-adapted host–parasite relationship. Some, such as Pirhemocyton, while originally thought to be a protozoan was subsequently found to be an iridovirus (see Chapter 9). Similarly, many of the commonly encountered gastrointestinal protozoans of reptiles, such as the ciliates Balantidium, and Nyctotherus, and the amoeba Hartmannella, are not known to result in lesions (Figures 12.9–12.11). For the most part they are considered commensals. While numerous species of Trypanosoma (Figure 12.7) have been reported in wild reptiles, no associated pathologic changes in the hosts have been noted. However, cases of Hepatazoon-associated hepatitis (Wozniak et al., 1998) have been seen in snakes, and, as previously mentioned, effects of Plasmodium on physiology and behavior of certain lizards has been studied (Schall, 1996). Certain coccidians, such as Caryospora in marine turtles (Gordon et al., 1993), Cryptosporidium in lizards and snakes (Brownstein et al., 1972; Uhl et al., 2001), and an intranuclear coccidian of tortoises (Jacobson et al., 1994) can result in significant clinical illness, pathologic response, and in some cases, death. The amoeba, Entamoeba invadens, is one of the first parasites reported to be a reptile pathogen (Geiman and Ratcliffe, 1936). Here I will present the major groups of protozoans known to infect and to be associated with disease processes in reptiles.

12.2.1 Amoebae The amoebae of medical importance remain uncategorized as to phylum and class (Lee et al., 2000). Amoebae are capable of forming pseudopodia (i.e., false feet). Historically, Entamoeba represents one of the most clinically significant parasites infecting captive chelonians and snakes. While several species of amoebae are known to infect reptiles (Barnard and Upton, 1994), and apparently exist in reptiles as commensals, others such as Entamoeba invadens are important pathogens. The life cycle of this parasite was one of the first described for a reptile (Geiman and Ratcliffe 1936). Reptile hosts that were studied included the water snakes (Natrix [formerly Nerodia] spp.), black racer (Coluber constrictor), and monitors (Varanus spp.). The infective stage for E. invadens, the quadrinucleate cyst, is shed in the feces. Cysts may persist in the environment for considerable periods of time. Ingested cysts pass through the gastrointestinal tract, where they develop into invasive trophozoites, which can be detected in fecal specimens using light microscopy. Studies by Barrow and Stockton (1960) demonstrated that infections in snakes were affected by the host’s body temperature. Experimentally inoculated snakes maintained at 13°C and those maintained at 33°C did not develop a clinical infection. However, snakes kept at 25°C consistently died fol-

Parasites and Parasitic Diseases of Reptiles  573

lowing inoculation. Today, while cases are still seen, they are far less common than in the 1970s and 1980s. In the past, distinguishing between Entamoeba invadens and other species of amoeba was based on host range and temperature sensitivity in culture. Today, molecular tools such as polymerase chain reaction (PCR) can be used to distinguish between the various species. Immunohistochemistry using a polyclonal antibody against E. invadens has been developed for use in snakes (Jakob and Wesemeier, 1995).

12.2.1.1 Chelonia  Hartman (1910) reported the first amoeba from a reptile with the description of E. testudinis from the Greek (Mediterranean spur-thighed) tortoise (Testudo graeca). Entamoeba testudinis was subsequently reported by Wenyon (1926) in the Argentine tortoise (Geochelone chilensis [formerly T. argentina]) and the African spurred tortoise (Geochelone sulcata [formerly T. calcarata]). Taliaferro and Homes (1924) described a second species, E. barreti, from the common snapping turtle (Chelydra serpentina). Sanders and Cleveland (1930) described the third species from a chelonian, E. terrapinae, that was recovered from the colon of the painted turtle (Chrysemys picta [formerly elegans]). Sanders and Cleveland (1930) described the third species from a chelonian, E. terrapinae, which was recovered from the colon of the painted turtle (Chrysemys elegans). For quite some time, chelonians (and crocodilians) were considered carriers because reports describing pathologic effects were lacking. This view changed when Jacobson et al. (1983) reported an epizootic of amoebiasis (compatible with E. invadens) in a captive group of recently imported red-footed tortoises (Geochelone carbonaria). Of 500 red-footed tortoises imported to southern Florida, approximately 200 died during a 2-month period. Clinical signs were nonspecific and included anorexia, listlessness, and watery diarrhea. Necropsy consistently revealed a thickened duodenum, with necrotic mucosa and multifocal to diffuse hepatic necrosis (Figures 12.12–12.13). Histologic evaluation of tissues demonstrated numerous amoebae in intestinal and hepatic lesions (Figures 12.14–12.16). Trophozoites in the wall of the intestine measured 12 to 19 µm by 10 to 13 µm, whereas those from the liver ranged from 13 to 21 µm by 11 to 18 µm; nuclei of trophozoites were 3 to 5 µm in diameter. The amoebae appeared to have spread to the liver via two pathways. One involved ascending the common bile duct to the gallbladder with spread to adjacent hepatic tissue; the second was by showering to the liver via the portal system. Thrombi within intestinal and hepatic vessels contributed to the pathologic changes. Predisposing conditions were probably involved. Tortoises were imported during the winter months and were more than likely exposed to temperatures below that for optimal growth of the amoebae. Additionally, tortoises were feeding minimally during this period. Without food, particularly carbohydrates, in the tortoise gastrointestinal tract, encystment of amoebae would not be favored (Jacobson et al.,

1983). Thus, once temperatures favored optimal growth of the parasite, the tortoises would be predisposed to tissue invasion by trophozoites. This appears to have happened during March and April when the tortoises died.  Six cases of amoebiasis were reported in flat-shelled spider tortoises (Acinixys planicauda), immediately after their importation into Japan from Madagascar (Ozaki et al., 2000). A fibrinonecrotic pseudomembrane was found along the mucosal surface of a thickened colon (due to submusosal edema); green foci were seen in the liver. Histologic examination of the colon and liver revealed numerous amoebic trophozoites (10 to 12 µm by 12 to 19 µm). While the amoebae were similar in size and morphology to E. invadens, they could not be conclusively identified. Over a one-year period, 21 juvenile tortoises and a loggerhead musk turtle (Sternotherus minor minor) in a zoological collection died; adults in the collection were not affected (Hollamby et al., 2000). Necrotizing colitis was seen in the tortoises, with the presence of intralesional protozoans consistent with Entamoeba invadens. In the musk turtles, amoebic trophozoites were present in the kidney, lung, ovary, and liver. MacNeill et al.(2002) reported an amoeba-associated mortality event in young wood turtles (Glyptemys [formerly Clemmys] marmorata). A Wright-Giemsa-stained impression smear of the liver of one turtle submitted for necropsy revealed frequent round unicellular protozoans measuring 27 to 32 µm in diameter with basophilic cytoplasm and a small eccentrically located round nucleus approximately 5.5 µm in diameter (Figure 12.17). Histologically there were areas of extensive hepatic necrosis containing 10 to 15 µm in diameter round to slightly oval protozoans that stained positive with periodic acid-Schiff (PAS) (Figure 12.18). The morphology and staining of the organisms were consistent with an Entamoeba sp. There are several reports of amoebic infections of captive green (Chelonia mydas) and loggerhead (Caretta caretta) sea turtles (Frank et al., 1976a; George et al., 1990). The major lesion seen was enterohepatitis with numerous trophozoites in the liver. I have seen several isolated cases in other turtles, including juvenile snake-neck turtles (Chelodina spp.).

12.2.1.2 Crocodylia  Amoebiasis in crocodilians is limited to two reports (Bemmel et al., 1960; Ippen, 1965). In a transmission study, American alligators (Alligator mississippiensis) inoculated with cysts of E. invadens did not become infected (Ratcliffe and Geiman, 1938).

12.2.1.3 Rhynchocephalia  There is a single report of amoebiasis in a tuatara (Sphenodon punctatus). The tuatara was in captivity for 8 years and had intestinal lesions similar to those reported in other reptiles (Frank et al., 1976b).

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12.2.1.4 Squamata   There are only a few reports of amoebiasis in lizards. One of the first recognized cases involved a lace monitor lizard (Varanus varius) that died with an E. invadens infection (Ratcliffe and Geiman 1934). In a subsequent report by Ratcliffe and Geiman (1938), infections with E. invadens were seen in water (V. salvator) and Komodo (V. komodoensis) monitors, and the blue-tongued skink (Tiliqua scincoides). Transmission studies in the American (green) anole (Anolis carolinensis) indicated they were not susceptible to infection. In another report, three months after arrival from Indonesia, a Komodo dragon at the National Zoological Park in Washington, DC, was found dead (Gray et al., 1966). Necropsy revealed a hemorrhagic colitis and enteritis. Using light microscopy, numerous amoebae were seen in ulcerative lesions in the colon and within the liver, which was diffusely necrotic. A blue-tongued skink was diagnosed with amoebiasis in another collection where large numbers of snakes were infected (Donaldson et al., 1975). Herbivorous lizards seem to be less susceptible to amoebiasis than are carnivorous lizards, and may serve as asymptomatic carriers (Frank, 1984a; Keymer, 1981). However, there are reports of systemic amoebiasis in green iguanas (Iguana iguana) (Hill, 1952; Hill and Neal, 1953; Frank, 1966a; Frank, 1974). Diffuse, severe necrotizing enterohepatitis is commonly seen in snakes suffering from amoebiasis. In spontaneous and experimental studies in snakes, the gastrointestinal tract and liver are generally the sites of infection (Ratcliffe and Geiman, 1934, Geiman and Ratcliffe, 1936). The lesions often start in the colon where focal to diffuse necrosis of the mucosa is seen (Figure 12.19). Lesions also may be seen in the small intestine and stomach. Spread of trophozoites to the liver results in hepatic necrosis (Figure 12.20). Microscopically, diffuse necrosis with inflammatory cell infiltrates is typically seen (Figure 12.21). The inflammatory response is generally due to secondary bacterial invaders such as Salmonella. While amoebae can be identified in tissue sections stained with routine hematoxylin and eosin, special staining with PAS is often used to identify trophozoites (Figure 12.22). With ulceration of the mucosa of the intestinal tract, amoebae enter portal vessels draining the intestinal tract where they can spread to the liver, and from there to almost any visceral organ. Massive enteritis, with or without hepatic necrosis, was seen in a large number of boas, pythons, and anacondas that died in an aquarium (Donaldson et al., 1975). When hepatic necrosis was seen, inflammation was minimal. Similar lesions (severe hemorrhagic colitis) were reported in ball pythons (Python regius) that died of amoebiasis in a display in Japan (Kojimoto et al., 2001). One snake in this group had necrosis in the brain with intralesional amoeba. While amoebiasis is a disease primarily of captive snakes, I have seen amoebiasis in a wild eastern king snake, Lampropeltis getula, which was found clinically ill in the field. There are no clinical signs of illness pathognomonic for amoebiasis. Affected animals may be listless and anorexic and may pass mucoid, bloody feces. Trophozoites can be

found in the feces of reptiles with signs of illness and those that are subclinical. Snakes may appear clinically normal up to 24 hours before death or may slowly decline in condition over a period lasting several weeks. A boa constrictor (Boa constrictor) and a northern Pacific coast rattlesnake (Crotalus viridis oreganus) infected with an Acanthamoeba-like protozoan, exhibited a star-gazing (opisthotonus) posture, a sign of nonspecific meningoencephalitis in snakes (Frye, 1991).

12.2.2 Parabasalia and Euglenozoa The phyla Parabasalia and Euglenozoa contain numerous species of flagellates; many have been identified in reptiles. The most significant genera include Trypanosoma and Leishmania (order Kinetoplastida), Hexamita (order Diplomonadida), and Trichomonas, Tritrichomonas, and Monocercomonas (order Trichomonadida). Although it has been suggested that reptiles may serve as reservoirs for Leishmania, which infects humans, there are no reports of naturally occurring reptilian leishmaniasis in humans (Keymer 1981). While flagellates are commonly encountered in the gastrointestinal tract of reptiles, there are few reports of lesions associated with flagellate infections in reptiles. Flagellates are often found in the intestinal lumen of most clinically healthy reptiles, and thus it is often difficult to attribute gastrointestinal disease of ill reptiles to the presence of flagellates in fecal samples. I have seen many anorectic snakes passing watery, foul-smelling feces that contained abnormally high numbers of flagellates. It is possible that a predisposing environmental condition (such as suboptimal environmental temperatures), a social stressor, or an earlier or current presence of an infectious agent allows an abnormal bloom of flagellates within the gastrointestinal tract.

12.2.2.1 Chelonia  Hexamitiasis of chelonians, caused by the parasite Hexamita parva, was reported by Zwart and Truyens (1975) for the turtles Clemmys, Cuora, Terrapene, Geoemyda, Testudo, and Geochelone. This parasite is approximately 8 µm in length and has six flagella, two of which are caudal. Clinical signs of illness are nonspecific, and include loss of weight and progressive apathy. In the above report, infected turtles had enlarged, pale kidneys at necropsy. Using light microscopy, they had acute or chronic inflammatory lesions of the kidneys and glomerular lesions consisting of proliferation of Bowman’s capsules, increased mesangial cellularity, and thickening of capillary basement membranes. Hexamita was seen within dilated renal tubules and in collecting ducts. In some cases there was infection of the bile duct system accompanied by necrosis and proliferative changes. A Home’s hingeback tortoise (Kinixys homeana), which was one of several confiscated tortoises donated to the Zoological Medicine Service at the University of Florida in Gainesville, died and had similar renal lesions (Figures 12.23–12.24).

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12.2.2.2 Ophidia  Monocercomonas (or similar-appearing parasite) is commonly seen in the gastrointestinal tract of snakes, including sea snakes (Telford, 1967). While in most cases disease is not seen, there are a few reports incriminating Monocercomonas as a pathogen of reptiles. A diamond python (Morelia spilota [formerly spilotes]) with a midbody swelling was found to have Monocercomonas-associated cholecystitis (Page and Jacobson, 1981) (Figures 12.25– 12.26). A corn snake (Elaphe guttata guttata), with a history of chronic regurgitation within several days of feeding, was found to have Monocercomonas-associated atrophic gastritis (Figures 12.27–12.28). Zwart et al.(1983) found Monocercomonas associated with a variety of illnesses in lizards and snakes. In this report, the clinical signs of illness were initially subtle; the animals became anorectic and showed reduced activity. In some cases, the reptiles developed diarrhea, and in one case a boa constrictor had a Monocercomonas infection in the caudal half of the oviducts, which were thickened and covered by seromucoid exudate. A garter snake (Thamnophis sirtalis) contracted Monocercomonas-associated pneumonia. In a study where Entamoeba invadens was initially considered the primary pathogen responsible for intestinal lesions in 51 snakes, when tissues were examined immunohistochemically using a monoclonal antibody against E. invadens, only 22 cases had intralesional protozoans identified as trophozoites of E. invadens (Jakob and Wesemeier, 1995). In 20 cases the protozoans were identified as flagellates. The flagellates were more commonly associated with changes in the small intestine, while amoebic trophozoites were more common in and around lesions in the large intestine. Tissue changes found in association with flagellates included epithelial desquamation, and inflammatory cells within a layer of fibrin on the mucosal surface. Flagellates can ascend the pancreatic duct of snakes, causing lesions resembling those caused by paramyxovirus (Michael M. Garner, personal communication). I have also seen flagellates in the oviduct of snakes having a salpingitis.

12.2.3 Apicomplexa: Nonhemoparasitic The most clinically significant nonhemoparasitic members of the phylum Apicomplexa infecting reptiles include the genera Eimeria, Isospora, Caryospora, and Cryptosporidium. Other members of the Apicomplexa known to infect reptiles are Sarcocystis and Besnoitia. However, there are few reports of associated lesions with the latter two genera. Eimeria is found in the gallbladder, bile ducts, and intestinal epithelium of snakes, lizards, and crocodilians (Figure 12.29). Sporulated oocysts of Eimeria have four sporocysts, each containing two sporozoites (Figure 12.30). Isospora is confined to the intestine (Figure 12.31) and sporulated oocysts of Isospora have two sporocysts, each containing four sporozoites (Figure 12.32). While most coccidians have intracytoplasmic development, at least 11 species of Eimeria, Isospora, and Cyclospora have been described as karyophagic, or more

accurately caryotropic, having intranuclear development (Atkinson and Ayala, 1987). Another important coccidian of wild and captive reptiles is Cryptosporidium (Upton et al., 1989). Over 57 different reptile species have been reported to be infected and include snakes, lizards, and tortoises (Donoghue, 1995). There is only a single report of Cryptosporidium in a crocodilian (Siam et al., 1994). Sequence analysis of the small subunit rRNA gene and PCR-restrictionfragment-length polymorphism was used to determine the genetic diversity of Cryptosporidium in reptiles (Xiao et al., 2004). Of 123 samples that were analyzed, 48 snake samples, 24 lizard samples, and 3 tortoise samples were positive for Cryptosporidium. Nine different types of Cryptosporidium were identified including Cryptosporidium desert monitor genotype, Cryptosporidium muris, Cryptosporidium parvum bovine and mouse genotypes, Cryptosporidium serpentis, one C. serpentis-like parasite in a lizard, two new Cryptosporidium in snakes, and one new Cryptosporidium in tortoises. Molecular and biologic characterizations indicated that the desert monitor genotype was probably Cryptosporidium saurophilum. Two host-adapted Cryptosporidium serpentis genotypes were found in snakes and lizards. Besnoitia form large cysts, which are often seen within cardiac and skeletal muscle of their host. In most situations, the cysts are seen in tissues of the host without any obvious tissue response. Sarcocystis has been reported as cysts within skeletal muscle of certain reptiles and within enterocytes in the intestinal tract. While some reptiles can serve as an intermediate host, others can serve as a final host, with stages in the intestinal tract (Matuschka, 1987). The few reptilian coccidians studied have been found to have a direct life cycle. Reptiles become infected by ingesting sporulated oocysts. Upon reaching the intestine, released sporozoites invade the mucosal epithelium. Although coccidia commonly infect free-ranging reptiles, most reports of illness associated with coccidial infections are in captive animals that are maintained under crowded, suboptimal conditions.

12.2.3.1 Chelonia 12.2.3.1.1 Eimeria, Isospora, and Unclassified Coccidians  Over 30 coccidian species are known from turtles and tortoises, but their mode of intracellular development, and even the site at which this occurs, have been poorly described. Eimeria is the most frequently reported turtle coccidian (Figure 12.33); only one species of Isospora is known from chelonians (Carini, 1942; Cerruti, 1930; Ernst et al., 1971; Upton et al., 1995). Disseminated visceral coccidiosis was reported in Indo-gangetic flap-shelled turtles (Lissemys punctata andersonii) that died in a zoo (Helke et al., 2006). Sporulated oocysts of Eimeria spp. were found in multiple tissues. An unusual unclassified intranuclear coccidian infection was reported in two radiated tortoises (Geochelone radiata) that were captive bred and reared in the

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United States (Jacobson et al., 1994). Both tortoises were in renal failure and were euthanized. Light microscopic examination of multiple tissues revealed nephritis and pancreatitis, and an intranuclear protozoan was seen in renal epithelial cells, hepatocytes, pancreatic acinar cells, and enterocytes (Figures 5.54, 12.34). Electron microscopic examination identified the organism as an intranuclear coccidian (Figure 12.35). Additional cases of intranuclear coccidiosis in tortoises were subsequently reported (Garner et al., 1998; Garner et al., 2006; Jacobson et al., 1999). In several radiated tortoises, a proliferative pneumonia was seen with an intranuclear coccidian in pulmonary epithelial cells (Figures 5.41,  12.36). Another tortoise, a leopard tortoise (Geochelone pardalis), had an inner ear infection, and coccidians were detected only in the epithelium lining the inner ear and Eustachian tube (Figures 12.37–12.38). A Bowsprit tortoise (Chersina angulata) with chronic active enteritis had numerous intranuclear coccidial organisms in enterocytes and hepatocytes. Oocysts were not recovered from any of these tortoises and identification to genus could not be made. In a recent study (Johnson et al., 2005), to determine phylogenetic relationships to other coccidians, coccidian-specific consensus primers were designed for a portion of the 18S rRNA gene. A 375-base-pair amplicon was sequenced from an intranuclear coccidian identified in six species of tortoises and found to be identical in all cases. When compared to known coccidial sequences, it was closely related to Hyaloklossia lieberkuehni, a renal coccidian of the European green frog (Rana esculenta). It is clear from the variety of cases reported that intranuclear coccidiosis is a problematic disease in imported and captive tortoises. The source of this organism(s) remains unknown. It is important to realize that diverse clinical signs and organ dysfunction can be seen, including pneumonia, enteritis, inner ear infection, pancreatitis, and nephritis. 12.2.3.1.2 Caryospora  A major epizootic in hatchling mariculture-reared green turtles in the Cayman Islands was caused by Caryospora cheloniae (Leibovitz et al., 1978; Rebell et al., 1974). Affected turtles ranged in age from 4 to 8 weeks. The major intestinal lesions involved the posterior small intestine and the colon, in which there was a loss of mucosal epithelium and the presence of necrotic cellular debris and blood, which filled the lumen. Sporulated oocysts had one sporocyst, were octozoic, and measured 33.8 to 40.1 µm by 11.0 to 14.6 µm. Gordon et al.(1993) identified Caryospora cheloniae in 24 of at least 70 subadult green turtles stranded on the beaches of Moreton Bay, southeast Queensland, Australia. The turtles were emaciated (Figure 12.39) and were found to have severe fibrinous or necrotizing enteritis (Figures 12.40–12.41) or meningoencephalitis. Various developmental stages of C. cheloniae were present in the intestinal lamina propria (Figures 12.42–12.43) as well as a variety of extraintestinal sites including the kidney, thyroid gland, and brain (Figures 12.44–12.45). Oocysts were obtained from

mucosal scrapings and were allowed to sporulate. Sporocysts were elongated (Figure 12.46) and following sporulation, the excysted sporozoites were arranged in a star formation for 2 or 3 days (Figures 12.47). 12.2.3.1.3 Cryptosporidium  There are relatively few reports of Cryptosporidium in turtles and tortoises. Heuschele et al. (1986) identified Cryptosporidium in a star tortoise (Geochelone elegans) and Funk (1988) in a red-footed tortoise. Oocysts of Cryptosporidium were identified in six of 34 (18%) fecal and intestinal samples of green turtles in Hawaii (Graczyk et al., 1997). Effects on the host were not mentioned. Graczyk et al. (1998) reported that an Egyptian tortoise (Testudo kleinmanni) that developed an enteritis and died several weeks later had heavy infection of the small intestine with Cryptosporidium sp. The lamina propria of the intestine was infiltrated with heterophils, lymphocytes, and macrophages. The author has observed Cryptosporodium lining the gastric mucosa of a chronically ill Texas tortoise (Gopherus berlandieri), and a colleague (Michael M. Garner, personal communication) identified Cryptosporidium on the surface of the gastric mucosa and small intestinal enterocytes (Figure 12.48) of a Hermann’s tortoise (Testudo hermanni) and on the gastric mucosal surface and within gastric pits (Figure 12.49) of a box turtle (Terrapene sp). 12.2.3.1.4 Sarcocystis  Lainson and Shaw (1972) reported Sarcocystis in skeletal muscles of the Brazilian scorpion mud turtle (Kinosternon scorpioides). I have seen Sarcocystis as an incidental finding in the tongue of desert tortoises (Gopherus agassizii) and skeletal muscles of other tortoises submitted to the Veterinary Pathology Service, College of Veterinary Medicine at the University of Florida in Gainesville including a Hermann’s tortoise (Testudo hermanni), an unidentified Testudo sp., and a pancake tortoise (Malacocherus tornieri). Typically no inflammatory reaction is seen (Figure 12.50).

12.2.3.2 Crocodylia 12.2.3.2.1 Eimeria, Isospora, and Goussia  Several species of coccidians have been identified in crocodilians, including Eimeria alligatori and E. hatcheri from the American alligator (Alligator mississippiensis), E. pintoi from an unidentified caiman, E. caimani and E. paraguayensis from yacare caiman (Caiman yacare), E. crocodyli from the American crocodile (Crocodylus acutus), Eimeria spp. from Nile crocodiles (Crocodylus niloticus), E. kermoganti from gavials (Gavialis gangeticus), Isospora jacarei from broad-snouted caimans (Caiman latirostris), and I. wilkei from American crocodiles (Huchzermeyer, 2003). While pathologic lesions are rarely reported in association with coccidial infections in crocodilians, with identification made primarily from their feces, coccidians have been seen in visceral lesions of crocodiles raised in farming operations in Africa, Australia, and New Guinea. Gardiner at al. (1986) described sporozoites and sporulating oocysts (20 µm in diameter) in the spleen, liver, and lungs of

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3- to 4-year-old Nile crocodiles that died at a farm in Zimbabwe (Figures 12.51–12.53). Schizonts and sporocysts have also been identified in the bile ducts and gallbladder (Foggin, 1992). Oocysts contained four sporocysts, each measuring approximately 15 µm by 6 µm. Each sporocyst contained two elongated sporozoites. While a specific identification could not be made, the coccidian had certain features of members of the genus Goussia. Of 54 crocodiles (New Guinea [Crocodylus novaeguinea] and saltwater [Crocodylus porosus] crocodiles) examined in a crocodile farm in Papua New Guinea, 17 had a Goussia-like coccidian in the intestine, liver, lung, and spleen (Ladds and Sims, 1990). Histologic changes associated with infection included loss, irregularity, and fusion of villi; epithelial hyperplasia; and inflammatory infiltrates in the lamina propria. At a crocodile farm in Irian Jaya (New Guniea), of 38 young crocodiles (New Guinea and saltwater crocodiles), 9 had coccidiosis (Ladds et al., 1995). Collection of these animals from the wild probably accounted for the parasitism. Lesions included attenuation, loss or fusion of intestinal villi, and ulceration, and one case had oocysts in the spleen. 12.2.3.2.2 Cryptosporidium  The only report of Cryptosporidium in a crocodilian was from one of nine captive Nile crocodiles in a zoo in Egypt (Siam et al., 1994). No associated pathologic change was noted.

12.2.3.3 Lizards and Snakes 12.2.3.3.1 Eimeria and Isospora  Numerous species of Eimeria and Isospora have been reported in lizards and snakes, with new species being identified annually (Asmundsson et al., 2001; Couch et al., 1996; Finkelman and Paperna, 2002; Lainson, 2003; Modry et al., 2000; Modry et al., 2001a; Modry et al., 2001b; Modry et al., 2004; Upton and Barnard, 1987; Upton and Freed, 1989; Upton and Freed, 1990; Upton et al., 1984; Upton et al., 1988; Upton et al., 1989; Upton et al., 1990) (Figures 12.54–12.57). However, many species from snakes previously reported as isosporans were found to be invalid and are probably Sarcocystis (Upton et al. 1992). Oocysts and sporocysts of a coccidian in rhinoceros vipers (Bitis nasicornis), which were originally identified as a member of the genus Isospora (Hoare, 1933), were later found to belong to two genera of coccidia: Sarcocystis and Besnoitia (Hafner and Matuschka, 1984; Matuschka and Hafner 1984). While most species of coccidians of lizards have intracytoplasmic development, several species of intranuclear Isospora are known from lizards (Atkinson and Ayala, 1987; Paperna and Finkelman, 1998a). Of the various Eimeria and Isospora described from lizards and snakes, relatively few have been associated with health problems. Zwart (1973) associated intestinal coccidiosis in chameleons with intussusception. A problematic parasite of captive-bred inland bearded dragons (Pogona vitticeps) in the United States is Isospora amphiboluri (Figure 12.31). This coccidian was originally described from

the bearded dragon (Pagona barbata [formerly Amphibolurus barbatus]) by Cannon (1967) and was redescribed from the inland bearded dragon by McAllister et al. (1995a). This parasite is firmly established in breeding groups of bearded dragons in the United States and its presence has been associated with mortality in neonate and juvenile lizards. Oocysts measure 23 to 26 µm by 23 to 26 µm. Combined infections of Isospora with adenovirus and dependovirus have been reported in inland bearded dragons (Kim et al., 2002). Eimeria bitis was reported to be pathogenic for several species of snakes including the puff adder (Bitis arietans), garter snake, ribbon snake (T. sauritus), and smooth green snake (Opheodrys vernalis), where necrotizing cholecystitis was reported (Fantham and Porter, 1950; Fantham and Porter, 1953–1954). Because it is unusual for a species of Eimeria to be shared by hosts in three different genera, this needs to be substantiated. Oocysts for E. bitis measure 28 to 36 µm by 18 to 24 µm. Developmental stages of Eimeria cascabeli in the prairie rattlesnakes (Crotalus viridis viridis) and South Pacific rattlesnake (C. v. helleri) were found in epithelial cells of the gallbladder and extrahepatic ducts where there was a proliferation of connective tissue and erosions (Vetterling and Widmer, 1968). Lehman (1972) reported that infection with Eimeria resulted in intense catarrhal and diphtheroid inflammation of the small and large intestine of reticulated pythons (Python reticulates) and golden tree snakes (Chrysopelea ornate). Fantham and Porter (1953–1954) reported that Isospora naviae was pathogenic in the Cape cobra (Naja nivea) and the timber rattlesnake (Crotalus horridus). Subsequently, Upton et al. (1992) considered the classification of Isospora naviae in these snakes invalid and felt it probably represented a species of Sarcocystis. 12.2.3.3.2 Caryospora  Several different species of Caryospora have been identified in lizards and snakes, including C. simplex by Modry et al. (1997) from the common viper (Vipera berus) and sand viper (Vipera ammodytes) and by Wilber et al. (1995) in Kaznakov’s viper (Vipera kaznakovi), C. varaniornati by Modry et al. (2001c) from the ornate Nile monitor (Varanus ornatus), C regentensis and C. legeri by Daszak and Ball (2001) from the eastern green mamba (Dendroaspis angusticeps) and hissing sand snake (Psammophis sibilans sibilans), respectively, C. matatu by Modry et al. (2002) from the horned bush viper (Atheris ceratophorus), C. duszynskii by McAllister et al. (1995b) from several species of colubrid snakes and C. bigenetica from several species of viperid snakes in the southwestern United States, C. ernsti by Upton et al. (1984) from the American (green) anole (Figure 12.58), C. gracilis by Upton et al. (1992) from the flat-headed snake (Tantilla gracilis) (Figure 12.59), C. serpentis by Upton et al. (1990) from the Malagasy giant hognose snake (Leioheterodon madasgascariensis) (Figure 12.60), and C. madagascariensis by Upton et al. (1990) from the Madagascar common snake (Madagascarophis colubrinus) (Figure 12.61). No lesions were reported with any of these descriptions.

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12.2.3.3.3 Cryptosporidium  Infections with Cryptosporidium have been identified in at least 15 species of lizards and 40 species of snakes (O’Donoghue, 1995; Upton, 1990). In the first authenticated report of Cryptosporidium in reptiles, this parasite was associated with hypertrophic gastritis in numerous species of harmless and poisonous snakes (Brownstein et al., 1972). Other reports in snakes followed, with many cases showing postprandial regurgitation (generally within 3 days of feeding), and progressive wasting (Godshalk et al., 1986; Klingenberg, 1996). I have seen Cryptosporidium in the stomach of several species of snakes including the Argentine boa constrictor (Boa constrictor occidentalis), bull snake (Pituophis catenifer [formerly melanoleucus] affinis), San Diego mountain king snake (Lampropeltis zonata pulchra), corn snake (Elaphe guttata guttata), eastern indigo snake (Drymarchon corais couperi), emerald tree boa (Corallus canina), Louisiana pine snake (Pituophis melanoleucus lodingi), Texas rat snake (Elaphe obsoleta lindheimeri), tiger rattlesnake (Crotalus tigris), timber rattlesnake (Crotalus horridus), and Trans-Pecos rat snake (Bogertophis [formerly Elaphe] subocularis). The clinical course in snakes is quite different than in mammals because in the former there is generally chronic gastric disease. Although affected snakes are often adults, a juvenile wild-caught corn snake was submitted to the author with typical signs and lesions. Snakes with severe cryptosporidiosis have a palpably firm stomach that may cause the surrounding body wall to bulge (Figure 12.62). Cryptosporidium can be diagnosed by demonstrating oocysts in mucus-coated, regurgitated food, gastric lavage specimens, impression smears of a gastric biopsy specimen (Figure 12.63), and in acid-faststained fecal smears (Figure 12.64). Gross lesions include thickened longitudinal rugae and an abnormal amount of mucus adhered to the mucosa. Histologically, gastric tissue has widely dilated tubules and increased interstitial connective tissue (Figures 5.37,  12.65). Cryptosporidium attach to the glycocalyx, with oocysts measuring 2.6 µm to 6.0 µm. The Giemsa staining method is often used to demonstrate these organisms (Figure 12.66). While primarily a parasite of the gastric mucosa of snakes, snakes with intestinal cryptosporidiosis have also been seen (Figure 12.67). A juvenile scaleless Texas rat snake with severe distension of the body wall (Figure 12.62) had Cryptosporidium in both a hyperplastic stomach and the small intestine where there was a proliferation of enterocytes. Cryptosporidium was also reported on the microvillous borders of the gallbladder and intra- and extrahepatic bile ducts in two of six corn snakes with gastric cryptosporidiosis (Cimon et al., 1996). Transmission studies (Graczyk and Cranfield, 1998) confirmed the pathogenicity of reptilian Cryptosporidium in rat snakes (Elaphe obsoleta). In this study both Cryptosporidium serpentis and Cryptosporidium sp. oocysts from turtles, tortoises, and lizards were infectious to rat snakes. Gastric hyperplasia developed in nine of ten snakes infected with nonsnake reptile oocysts.

Identification of pathogenic effects of Cryptosporidium in lizards lagged behind reports in snakes. As more cases were reported, it was clear that Cryptosporidium in lizards tends to have a wider range of tissue tropism than in snakes. Gastric cryptosporidiosis was reported by Dillehay et al. (1986) in a subadult Senegal chameleon (Chamaeleo senegalensis) and by Frost et al. (1994) in two ocellated (jewelled) lacertas (Lacerta lepida). Klingenberg (1996) reported enteric cryptosporidiosis in a panther chameleon (Furcifer pardalis). In the latter report, a savannah monitor (Varanus exanthematicus) had Cryptosporidium in both the stomach and small intestine. I have identified Cryptosporidium attached to hyperplastic enterocytes in the small intestine of a black-andwhite tegu (Tupinambis teguixin). A two-month-old freckled monitor (Varanus tristis orientalis) had cryptosporidiosis of the small intestine and colon (Nathan, 1996). A Madagascan giant day gecko (Phelsuma madagascariensis grandis) had lesions within the cloaca (Upton et al., 1989). There are several reports of Cryptosporidium in leopard geckos (Eublepharis macularius) having a history of weight loss, lethargy, and diarrhea (Coke and Tristan, 1998; Graczyk et al., 1999; Taylor at al., 1999). In another study, geckos were emaciated (Figure 12.68) and had Cryptosporidium sp. on the gastric mucosa and at the apical aspect of enterocytes lining the intestinal villi (Terrell et al., 2003) (Figures 12.69–12.70). There was associated mucosal epithelial hyperplasia and mononuclear cell inflammation of the small intestine (Figure 12.71). Renal cryptosporidiosis was reported in a green iguana and a Parson’s chameleon (Calumma parsonii cristifer), and Cryptosporidium was identified in an enlarged hyperplastic salivary gland of a green iguana (Frye et al., 1999). Aural (protrusion of the tympanic membrane) and pharyngeal polyps were seen associated with cryptosporidiosis in several green iguanas (Fitzgerald et al., 1998; Uhl et al., 2001). Large pedunculated cystic masses protruded into the oral cavity and from the ear canal (Figures 12.72–12.73). Cryptosporidium was easily seen along the apical surface of epithelial cells in these masses (Figures 12.74–12.76). Cryptosporidium saurophilum, a new species, was described from the small intestine of Schneider’s skinks (Eumeces schneideri); no pathologic change was noted (Koudela and Modry, 1998).

12.2.3.4 Sarcocystis and Besnoitia  Sarcocystis is known to infect lizards and snakes, and three genera of lizards were reported to be infected with Besnoitia (Barnard and Upton, 1994). As stated previously, many coccidia originally identified as Isospora were found to be misidentified and were determined to be members of the genus Sarcocystis (Matuschka, 1987; Upton et al., 1992). Transmission studies have demonstrated the susceptibility of rodents to certain snake Sarcocystis (Lindsay et al., 1991; Paperna and Finkelman, 1998b), and the giant ameiva (Ameiva ameiva) in Brazil can serve as an intermediate host for the Rio tropical racer (Mastigodryas bifossatus) (Lainson and Paperna, 2000).

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In a search of the literature, there are very few reports of Sarcocystis-associated pathologic changes in tissues of lizards and snakes, either due to oocytes in the enteron or asexual stages in skeletal muscle. Similarly, while Besnoitia has been reported in several species of lizards, no pathologic changes have been seen. In a Fischer’s chameleon (Chameleo fischeri), limb, masseter, and occipital muscles containing cysts of Sarcocystis were swollen (Frank, 1966b). A black-headed monitor (Varanus tristis tristis) that died without manifesting signs of illness, had large numbers of Sarcocystis-like parasites within epaxial musculature; there was necrosis of the musculature in association with this parasite (Hollamby and Creeper, 1997). There is only a single report of pathologic change associated with Sarcocystis infection in a snake. Daszak and Cunningham (1995) identified severe intestinal sarcocystosis in a naturally infected bull snake (Pituophis melanoleucus sayi). The lamina propria was packed with sporulated oocysts with two sporozoites; oocysts also were present in enterocytes. Typically, the pathologic change associated with Sarcocystis is caused by the schizonts in the intermediate host.

12.2.3.5 Klossiella  Vegetative stages of Klossiella boae were reported in epithelial cells of renal tubules of the boa constrictor (Zwart, 1964) (Figure 12.77). In a survey of recently imported boa constrictors, 0.5% of the snakes were infected. 

12.2.4 Apicomplexa: Hemoparasitic This group of blood-borne intracellular sporozoans includes the hemogregarine Coccidia (Haemogregarina, Hepatozoon, Karyolysus, and Haemolivia), Hemococcidia (Schellackia and Lainsonia), and Plasmodium, Haemocystidium, Fallisia, Saurocytozoon, and Haemoproteus. While circulating hemogregarines primarily parasitize red blood cells, they have also been seen less commonly in white blood cells. Plasmodium, Fallisia, and Saurocytozoon all can parasitize white cells, and the latter two do, without using red blood cells, except occasionally for Saurocytozoon. These hemoparasites have been identified in all groups of reptiles except marine turtles. Snakes are host to the hemogregarines Plasmodium and Haemoproteus, turtles to hemogregarines and Haemoproteus, crocodiles and the tuatara to hemogregarines, and lizards to all members of this group, and are the only hosts for hemococcidia. Arthropods and annelids are involved in their transmission. Schellackia and Lainsonia use vectors for mechanical transfer only, with sporogony occurring in the vertebrate host. Knowledge of the parasite’s development in both the vector and the reptile is necessary for classification at the generic level. Haemogregarina, Hepatozoon, Haemolivia, and Karyolysus have sporogenic development in invertebrate vectors (ticks, mites, true bugs, mosquitoes, leeches) and belong to the Adeleiina. Schellackia and Lainsonia undergo both sporogony and gametogony in the reptile host and are in the

Eimeriina (Telford, 1984). Developmental stages in the invertebrate host are necessary for distinguishing among Haemogregarina, Hepatozoon, Haemolivia, and Karyolysus. While Plasmodium, Haemocystidium, and Haemoproteus have somewhat similar morphology, haemoproteiids do not have asexual reproduction in circulating blood cells; schizogony is within visceral organs. Although wild reptiles are commonly infected with these parasites, lesions are seldom reported. In the European viviparous lizard (Lacerta vivipara), hemogregarine infection was associated with negative effects on hemoglobin concentration and oxygen consumption at rest (Oppliger et al., 1996).

12.2.4.1 Haemogregarina and Hepatozoon 12.2.4.1.1 Chelonia  Hemogregarines have been reported in all families of freshwater and terrestrial turtles (Telford, 1984). They are uncommon in tortoises and have not been reported in sea turtles. 12.2.4.1.2 Crocodylia  Hemogregarines have been reported from several species of crocodilians. Hemogregarina crocodilinorum is commonly seen in peripheral blood films of wild American alligators (Figure 12.78). Haemogregarina was commonly seen in red blood cells of New Guinea and saltwater crocodiles at a farm in Irian Jaya, New Guinea (Ladds et al., 1995). Hepatozoon schizonts were seen throughout the small intestine of wild-caught African dwarf crocodiles (Osteolaemus tetraspis) brought to market in the Congo Republic (Huchzermeyer and Agnagna, 1994). A recent classification places all crocodilian Haemogregarina in the genus Hepatozoon (Smith, 1996), despite the clear statement by Khan et al. (1980) that nonsporocystic oocysts are formed during sporogony of H. crocodilinorum in the leech vector. 12.2.4.1.3 Rhynchocephalia   Haemogregerina tuatarae was reported from the tuatara (Laird, 1950). 12.2.4.1.4 Squamata  Hemogregarines have been described from numerous species of lizards and are the most common intraerythrocyte parasites seen in snakes (Figure 12.79), with most identified as members of the genus Hepatozoon. Schizonts may be found in the liver, lungs, pancreas, and spleen (Figures 12.80). One or two bananashaped organisms may be seen within a single red blood cell, which is often distorted in size and shape. Some species in snakes have a very narrow host range (Telford et al., 2001), while others are polytopic, capable of parasitizing multiple species (Telford et al., 2004). Although severe infections have been associated with anemia and dehemoglobinization, in most cases severe infections are seen without concomitant clinical signs of illness or significant lesions in tissues. Experimental infections can result in inflammatory changes in infected tissues (Wozniak and Telford, 1991). A group of Mojave Desert sidewinders (Crotalus cerastes cerastes) that were parasitemic with a

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hemogregarine had marked erythrocytic hypertrophy and dehemoglobinization (Wozniak et al., 1994). One snake had severe anisocoria and was euthanized. Using light microscopy, there were prerythrocytic meronts throughout the liver and within pulmonary endothelial cells. No inflammation was seen around any of the prerythrocytic meronts. Necrotizing lesions were found in association with hemogregarine meronts in a California king snake (Lampropeltis getula californiae) (Keymer, 1976). A crotalid snake in a zoo collection had meronts associated with inflammatory lesions in the liver (Griner, 1983). This snake was showing signs of neurologic disease before it died, and light microscopy revealed meronts in the brain. A wild-caught southern water snake (Nerodia fasciata pictiventris) that had numerous hemogregarines (Hepatozoon) in a peripheral blood film was euthanized; using light microscopic examination of the liver revealed granulomatous inflammation around Hepatozoon meronts (Wozniak et al., 1998). I have seen inflammation, necrosis, and fibrosis in association with prerythrocytic meronts of a hemogregarine in the liver of an emerald tree boa (Corallus caninus) (Figures 12.81–12.82). Because hemogregarines are transmitted through an invertebrate vector, in most cases transmission in captivity is self-limiting. However, intrauterine transmission has been documented in the Brazilian lancehead viper (Bothrops moojeni) and the Neotropical rattlesnake (Crotalus durissus) (De Biasi et al., 1971) and in the boa constrictor (Telford, 1984).

12.2.4.2 Plasmodium  There are 96 described species and subspecies of Plasmodium in reptiles; most have been reported for lizards, with only six species found in snakes and none in crocodilians and turtles (Telford, 1984; Sam R Telford Jr, personal communication). Two additional plasmodiid genera occur in lizards as well, Fallisia and Saurocytozoon, both genera parasitizing nonerythroid cells. Fallisia, with 10 species, has both schizonts and gametocytes in leukocytes and thrombocytes, while Saurocytozoon (2 species), a common parasite of tegu lizards and Brazilian and Southeast Asian skinks, may have a transitory asexual cycle in circulating cells, with gametocytes occupying lymphocytes (Telford, 1978). All undergo sporogony in an insect vector and both schizogony and gametogony in the erythrocytic series of the reptile host (Figures 12.1–12.3). During an active infection, secondary exoerythrocytic schizonts may be seen in lymphocytes, monocytes, and thrombocytes. Schizonts of P. mexicanum have been identified in cerebral capillaries (Ayala, 1970). Few reports associate pathologic lesions with Plasmodium infections in reptiles. Pathogenicity of the various species of Plasmodium has been related to each parasite’s size and ability to infect immature blood cells. Telford (1984) reported an anemia in the Japanese grass lizard (Takydromus tachydromoides) and the green grass lizard (T. smaragdinus) infected with P. sasai. Severe anemia has also been reported by Lehman (1972) in the Amazon lava lizard (Tropi-

durus torquatus) experimentally infected with P. tropiduri. A natural infection of the grass anole (Norops auratus) with P. colombiense resulted in an initial severe anemia with erythroblastosis, which was followed by a chronic infection with a normal blood picture (Ayala and Spain, 1976). Behavioral and physiological changes were seen in lizards infected with Plasmodium (Schall, 1990a, 1990b, 1996). Infection resulted in increased numbers of immature red blood cells in peripheral blood, decline in hemoglobin concentrations, reduced oxygen supply to tissues, and ultimately reduction in running stamina. For infected males, the testes size was reduced by 37%, and for females, the clutch size was reduced by 20%. In a wild population, lizards classified as dominant were not infected, while those classified as submissive were infected. Thus infection with Plasmodium can affect behavior and the ability of a male lizard to defend its territory. The common agamid lizard (Agama agama) infected with Plasmodium agamae and P. giganteum suffered similar hematological and physiological effects (Schall, 1996). Infection did not reduce clutch size, but may lengthen the time between clutches.

12.2.4.3 Haemoproteus  Approximately 27 species of Haemoproteus have been reported from turtles, lizards, and snakes. Telford (1996) assigned the Haemoproteus species described from lizards by Paperna and Landau (1991) to the genus Haemocystidium. Only the cobra parasite Haemoproteus mesnili, and the 10 haemoproteid species reported from turtles, should be considered Haemoproteus (Sam R. Telford, Jr. personal communication). Although it is believed that an insect vector is necessary for transmission, a natural vector (a tabanid fly Chrysops) has been reported only for H. metchnikovi. There are limited reports of the pathological conditions associated with Haemoproteus infections in reptiles. A dehemoglobinization of erythrocytes has been reported in the common wonder gecko (Teratoscincus scincus) infected with H. kopki (Telford, 1984). The continuing destruction of red blood cells that occurs with Plasmodium infections is not seen in infections with Haemoproteus. As with other hemoparasites, transmission should be self-limiting in captivity if there is good vector control.

12.3 Microsporida The phylum Microsporida consists of obligate intracellular unicellular protozoans that are collectively termed microsporidians. Although formerly classified with the Protozoa, recent DNA sequencing studies indicate that the phylum Microspora should be classified under the Fungi kingdom. Over 100 genera and almost 1000 species have been reported in a wide variety of invertebrates and all classes of vertebrates (Weber et al., 1994). Members of this phylum have only asexual reproduction. Infection begins with injection of sporoplasm into the host cell followed by a proliferating

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merogonic phase. Eventually a sporogonic phase begins in which meronts of simple structure transform into sporonts of relatively complex structure. In the past, the morphology, internal and external, of both stages was used to distinguish microsporidia. Today, DNA sequencing has replaced morphological features for determining phylogenetic relationships of various microsporidia. In reptiles, Pleistophora was reported in the tuatara in a zoological collection (Liu and King, 1971); Encephalitozoon lacertae in the European wall lizard (Podarcis muralis) in France (Canning, 1981); Pleistophora (formerly Glugea) danilewskyi in the European grass snake (Natrix [formerly Tropidonotus] natrix), in Italy (Guyenot and Naville, 1922; Guyenot et al., 1925); and Pleistophora atretii in the split keelback snake (Atretium schistosum) in India (Narasimhamurti et al., 1982). Microsporidian spores were identified in fecal smears of 19 (21%) of 90 snakes from a zoological collection (Graczyk and Cranfield, 2000). Inland bearded dragons (Pogona vitticeps) have become popular in the reptile pet trade in the United States and Europe. As a consequence of large-scale breeding, several pathogens have been identified as causes of morbidity and mortality in colonies of these lizards, one of which is Microsporidium (Jacobson et al., 1998). Light microscopic examination of hematoxylin and eosin-stained tissue sections obtained from a bearded dragon that died showing nonspecific signs of illness revealed clusters of light basophilic microorganisms (2 to 3 µm in diameter) packing renal tubules (Figure 12.83). The organisms were acid-fast (Figure 12.84). With a Gram stain, they stained positive (Figure 12.85). There was also severe hepatic necrosis with clusters of similar appearing intracytoplasmic microorganisms packing and distending hepatocytes and free in areas of necrosis (Figure 12.86). Similar microorganisms were within (1) alveolar epithelial cells, (2) gastric mucosal epithelial cells, (3) enterocytes, and (4) capillary endothelial cells and ventricular ependymal cells in the brain. In two other lizards, similar appearing microorganisms were in macrophages in granulomatous inflammation in the colon, adrenal glands, and ovaries. In addition to being Gram-positive and acid-fast, the organism had a small polar granule that stained positive with the periodic acid-Schiff reaction. Electron microscopic examination of deparaffinized liver revealed merogonic and sporogonic stages of a protozoan compatible with members of the phylum Microsporida (Figure 12.87). Because one of the lizards was laboratory hatched and reared, and was fed only crickets, transmission may have been through infected crickets or vertically through the egg.

12.4 Myxozoa The phylum Myxozoa was once grouped with the Microsporida in a single taxon that was called the Cnidospora because members of both have coiled filaments in the spore stage.

Based on DNA sequencing, the Myxozoa are now classified within the kingdom Metazoa. While the Microsporida have only asexual reproduction, the Myxozoa have an elementary type of sexual reproduction and the spores are multicellular in origin (Marquardt et al., 2000). While most are parasites of teleost fish, there are several reports of myxozoan infection of amphibians including wild frogs and toads in the United States (McAllister and Trauth, 1995; Eiras, 2005). Renal infection in Asian horned frogs (Megophrys nasuta) in zoological collections with Chloromyxum sp. (Duncan et al., 2004) was reported. In reptiles, myxozoans have been reported from a variety of aquatic turtles (Eiras, 2005). Kudo (1919) described Myxidium americanum from the urinary tract of the soft-shelled turtle Apalone (formerly Trionyx) ferox spinifer. Myxidium chelonarum was reported from the gallbladder and bile ducts of the yellow-bellied slider (Pseudemys scripta scripta) and 13 other species of aquatic turtles from the eastern and southeastern United States (Johnson, 1969). An unidentified myxozoan was found in dilated renal tubules of a red-eared slider (Trachemys scripta elegans) from a farming operation in Louisiana (John Roberts, personal communication) (Figures 12.88–12.89). Myxobolus sp. was identified in dilated renal tubules of a yellow-spotted Amazon River turtle (Podocnemis unifilis) that had chronic interstitial nephritis (Jose Catão-Dias, personal communication) (Figures 12.90–12.91). Myxidium mackiei was identified in the renal tubules of two Indo-Gangetic flap-shelled turtles (Lissemys punctata andersonii) that died in a zoo collection (Helke and Poynton, 2005). Spores occluded 10% of the renal proximal convoluted tubule, and the microvilli of cells within these tubules were sheared, compressed, or missing. Renal myxozoanosis was associated with renal disease in the crowned river turtle (Hardella thurjii) (Garner et al., 2006). Using DNA sequencing, this was named Myxidium hardella n. sp.

12.5 Cestoda 12.5.1 Pseudophyllidea Members of this order include the genera Duthiersia, Bothridium, Bothriocephalus, Scyphocephalus, and Spirometra. There are few reports describing the life cycles of these tapeworms. Eggs are released through a uterine pore into the intestinal system of the definitive host and pass in the feces. Bothridium and Bothriocephalus are mainly parasites of boid snakes. In most situations they are found incidentally at necropsy (Figure 12.92). Although heavy infections are not rare, there are nevertheless few reports of associated pathological effects. A reticulated python infected with Bothridium pithonis had severe edema and hemorrhage of the intestinal mucosa at the site of attachment (Wiesenhütter, 1964). Toft and Schmidt (1975) described mild chronic 

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enteritis associated with Bothridium in two green tree pythons (Morelia [formerly Chondropython] viridis). Spirometra is a pseudophyllidean cestode that is widely distributed in different species of snakes, which serves in most cases as an intermediate or paratenic host. A paratenic host is one used to transport infectious stages of a parasite to the definitive host. There is no additional development of the parasite in the paratenic host. When released from the definitive host, each egg releases a larva (coracidium), which when ingested by a copepod, develops into a procercoid. Upon ingestion of the latter stage by a second intermediate host (amphibian, reptile, or mammal), procercoids develop into plerocercoids that are known as spargana, and can be found throughout the body. When present subcutaneously, spargana may result in soft swellings of the body. Edema and hemorrhage of soft tissues may be associated with this stage. The definitive host is generally a mammalian carnivore (most often a felid). Diagnosis of pseudophyllidean infections is based upon the identification of the Spirometra larvae or the identification of eggs in the feces of animals infected with Bothridium and Bothriocephalus (Figure 12.93). The latter two parasites have operculated eggs that appear similar to trematode eggs. In order to differentiate these genera, eggs can be placed in shallow water until the oncosphere or coracidium develops (Frank, 1981). Differing from most other cestodes, eggs of pseudophyllideans float in salt solutions, thus aiding in their diagnosis.

12.5.2 Proteocephalidea This order of cestodes contains a majority of the tapeworm species found in reptiles, and includes the genera Proteocephalus, Acanthotaenia, Crepidobothrium, and Ophiotaenia. The major genus, Ophiotaenia, consists of approximately 50 species. The life cycle includes an intermediate invertebrate (copepod) host, and reptiles serve as the definitive host. Tadpoles, frogs, and fish may serve as paratenic hosts. Adult parasites are found within the small intestine. Most infections in reptiles are subclinical. High numbers of tapeworms may result in mechanical obstruction of the gastrointestinal tract or in competition with the host for essential nutrients. A green anaconda (Eunectes murinus) with 1547 Crepidobothrium parasites in the intestine was emaciated at the time of death, even though it had been eating regularly (Kutzer and Grunberg, 1965). Diagnosis is based upon demonstration of adult parasites in feces or identification of eggs containing characteristic oncospheres in fecal smears or fecal sediment samples. Eggs of Ophiotaenia measure approximately 50 µm by 40 µm (Figures 12.94–12.95).

12.5.3 Trypanorhyncha Larvae of certain members of this tapeworm order are known to infect sea turtles. Plerocercoids were found in two of 33 loggerhead sea turtles from a fish market in Alexandria, Egypt (Sey, 1977). Large numbers of cysts were attached to the serosa of the stomach and the outer surface of the lungs. Their morphology suggested affinities to Lacistorhynchus or Eutetrarhynchus. I have seen larval trypanorhynchan cysts in the wall of the stomach, small intestine, surface of the liver, peritoneum, and limb musculature of stranded loggerhead sea turtles and green turtles in Florida (Figures 12.96– 12.97). They are surrounded by a bladderlike structure (Figure 12.98). In tissue section the bladder is often surrounded by a thin connective tissue capsule (Figure 12.99). Members of this group use elasmobranchs (sharks) as the definitive host.

12.5.4 Cyclophyllidea The most important and unique cyclophyllideans in lizards and snakes are in the family Mescocestoididae, genus Mesocestoides. Members of this genus require three hosts to complete their life cycle, with mammals and birds serving as the final host. Arthropods (mites) serve as the first intermediate host, in which oncospheres develop into a cysticercoid. The larva that infects reptiles is called the tetrathyridium and development in reptiles occurs after they ingest an infected arthropod containing the cysticercoid. In reptiles, tetrathyridia may be found within the intestines or at a variety of extraintestinal cells. Goldberg (1985) found tetrathyridia of Mesocestoides within white nodules in the liver, on the intestine, within the coelom, and within the mesenteries of the island night lizard (Xantusia riversiana). There was minimal inflammatory response, with the parasites surrounded by a thin fibrous capsule. Other cases of Mesocestoides tetrathyridia in the island night lizard have been seen (Figure 12.100). Mesocestoides also has been seen in the liver of a captive inland bearded dragon from California (Figures 12.101–12.102). I received tissues of an Arizona mountain king snake (Lampropeltis pyromelana) that had approximately 80% of the pancreas replaced by tetrathyridia (Figure 12.103). While the inflammatory response to these parasites is often minimal, mechanical damage to organs and the replacement (loss) of normal cells probably results in organ dysfunction.

12.6 Trematoda Numerous trematodes have been reported from all groups of reptiles. Marine turtles may have more genera and species than any other reptile. At least 51 species of trematodes have been reported from the green turtle alone (Lauckner, 1985). However, relatively few have been associated with any dis-

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ease process. Those of clinical and pathological significance are discussed below.

12.6.1 Ochetosomatidae and Plagiorchiidae Several genera of digenetic trematodes within the families Ochetosomatidae (Dasymetra, Lechriorchis, Zeugorchis, Ochestosoma) and Plagiorchiidae (Stomatrema) are commonly encountered in the oral cavity (Figure 12.104), pulmonary system (Figure 12.105), and upper esophagus (Figure 12.106) of snakes. These parasites have been collectively called the renifers. Amphibians are intermediate hosts, but even when this prey is removed from the diet, infections appear to persist for prolonged periods of time in captive snakes. Possibly, larval stages within the gastrointestinal tract mature at relatively slow rates or adults are long lived. Adult members of the above genera migrate from the oral cavity, by way of the glottis, into the lungs and air sacs. Although in most infections no signs of illness are seen, renifers must still be considered potential pathogens. In the lungs, adult parasites attach to the epithelial lining, where they produce focal lesions. In several instances, secondary bacterial pneumonia with Gram-negative microorganisms (Providencia, Pseudomonas) has been found associated with pulmonary trematode infections. Diagnosis is based upon identification of adult parasites in the gastrointestinal system, oral cavity, and respiratory system. Parasite eggs are yellow-orange, measure approximately 40 µm by 25 µm, and have an operculum (Figure 12.107). Eggs shed within the lungs are coughed up, and with those in the oral cavity, are swallowed and eliminated in the feces. Eggs are easily identified in lung washings of infected shedding snakes or (with more difficulty) in a fecal sediment sample. Other trematode species inhabiting the gastrointestinal and urogenital systems produce eggs that are easily confused with eggs of these parasites. Another member of the family Plagiorchiidae known to infect snakes is Styphlodora. This digenetic trematode inhabits the urinary system of snakes where infections of collecting tubules and ureters are seen. Frank (1981) found several hundred in the ureter of a young reticulated python, and 37 in a boa constrictor. Lesions associated with this parasite include renal tubular dilatation, buildup of intraluminal tubular debris, and chronic interstitial nephritis. A capsule of fibrous connective tissue often surrounds Styphylodora. In addition to boa constrictors, I have seen this trematode in a tropical rat snake (Spilotes pullatus) (Figure 12.108) and an eastern king snake (Lampropeltis getula). The vast majority of cases are probably subclinical. Antemortem diagnosis depends upon identification of eggs in a direct smear of urine or within urate sediment. Eggs are yellow in color and measure approximately 40 µm by 18 µm. Diagnosis is generally made upon histological examination of sections from the kidney and ureter. In most

cases, infection is of little clinical importance. Although the life cycle of this parasite is unknown, an intermediate host is most likely involved because an intermediate host is needed for all digenetic flukes; thus, infection in captivity should be self-limiting.

12.6.2 Spirorchiidae Adult members of the family Spirorchiidae inhabit the circulatory system of susceptible reptiles, with turtles appearing to be the most commonly infected reptilian host. Adult parasites are generally found within the great vessels leaving the heart or within the heart chambers, where focal endothelial hyperplasia has been seen. The most significant lesions resulting from these parasites are attributed to the release of eggs within the vascular compartment. Eggs ultimately become trapped within terminal vessels almost anywhere in the animal’s body; at these places a severe inflammatory response (often granulomatous) may be seen.

12.6.2.1 Chelonia  Digenetic trematodes of the family Spirorchiidae utilize freshwater turtles and sea turtles as their definitive hosts. At least 8 genera and 20 species of spirorchiids infect loggerhead, green, and hawksbill (Eretmochelys imbricata) sea turtles, and are considered the most pathogenic of sea turtle parasites (George, 1997; Lauckner, 1985). Spirorchiids are vascular system generalists, with a preference for the heart and arterial system of their turtle hosts (Platt and Brooks, 1997). Lesions have been seen in Spirorchis scripta infected wild emydine turtles in the United States (Holliman and Fisher, 1968), spirorchiid infected stranded loggerhead sea turtles in the eastern United States (Wolke et al., 1982), Learedius learedi and Neospirorchis schistosomatoides infected stranded green turtles in Bermuda (Rand and Wiles, 1984), L. learedi, Hapalotrema dorsopora, and Carettacola hawaiiensis infected green turtles in Hawaii (with and without fibropapillomatosis) (Aguirre et al., 1998), Hapalotrema mehrai, H. postorchis, and N. schistosomatoides infected wild green turtles in Australia (Gordon et al., 1998), L. learedi infected farmed green turtles in the Cayman Islands (Greiner et al., 1980), and Haplotrema infected farmed, oceanarium-reared and wild sea turtles in Australia (Glazebrook and Campbell, 1990a, 1990b). Turtles with heavy infections may be in a severely debilitated state. I have seen shell lesions in emydine turtles infected with Spirorchis (Figures 12.109–12.110). Microscopically, eggs (with and without associated granulomas) may be found in almost any tissue including the brain, heart, lung, liver, spleen, urinary bladder, salt glands, and shell. In stranded sea turtles, adult parasites caused endocarditis, arteritis, and thrombosis of the blood vessels (Gordon et al., 1998). In stranded green and loggerhead sea turtles in Florida, large collections of eggs in the intestinal tract may be seen as black, raised, linear to confluent serpentine lesions (Fig-

584  Parasites and Parasitic Diseases of Reptiles

ures 12.111–12.112). Nodules may be seen protruding from the serosal surface (Figure 12.113). Microscopic examination of these lesions revealed large numbers of eggs surrounded by cellular debris and fibrous connective tissue (Figures 12.114–12.115). Spirorchiid eggs also may be associated with a moderate to severe inflammatory response (Figures 5.77, 12.116). Large numbers of eggs are commonly seen in the spleen (Figures 12.117–12.119). In addition to direct pathological effects in the vicinity of the adult parasite, eggs released within the vascular system may be transported to remote areas almost anywhere in the animal’s body where they lodge in small vessels, often initiating a mild to severe granulomatous inflammatory response. The eggs can also migrate through blood vessel walls, causing tissue damage and inflammation in adjacent tissues (Gordon et al., 1998). Turtles may develop edema of the limbs, which is the result of vascular occlusion. Severe lesions have been seen in the gastrointestinal mucosa and submucosa, and in the spleen, liver, heart, kidneys, and lungs. The author of this chapter has seen turtles with severe granulomatous lung lesions exhibiting flotation abnormalities. Meningitis and encephalitis have been reported in green turtles and loggerhead sea turtles in response to parasitism by intravascular spirorchiid trematode eggs (Gordon et al., 1998). Adults and eggs (Figure 12.120) of Neospirorchis have been seen in meningeal vessels of the brain and spine of loggerhead sea turtles diagnosed with a neurological disease (Jacobson et al., 2006). In painted turtles (Chrysemys picta) experimentally infected with Spirorchis parvus, one developed hemiplegia (Holliman et al., 1971). The left side of the fourth ventricle of this turtle had adult flukes and was necrotic. Spirorchiid parasitism may promote secondary Gram-negative bacterial infections (Raidal et al., 1998). Diagnosis of spirorchid infections is generally made at necropsy or upon identification of eggs within tissue sections. For making a specific identification, the parasite or eggs need to be removed and mounted on glass slides for microscopic identification. Eggs vary in size and morphology among the various genera. Eggs of Spirorchis elegans (infecting freshwater pond turtles) measure approximately 100 µm by 75 µm, Learedius learedi, which infects the green turtle, produces eggs measuring approximately 300 µm by 70 µm, Haplotrema eggs measure approximately 325 µm by 25 µm, and Neospirorchis measure approximately 50 µm by 40 µm (Figures 12.121–12.126).

12.6.3 Diplostomatidae The family Diplostomatidae has genera (Alaria and Fibricola) with larvae that have been identified in subcutaneous tissues in snakes. An adult male Texas indigo snake (Drymarchon corais erebennus) developed large fluctuant subcutaneous masses in both the intermandibular region and 10 cm distal of the tail (Wright et al., 1989). Needle aspirates of the

masses revealed numerous trematode larvae. The cervical mass was surgically removed and the affected portion of the tail was resected. Numerous immature trematodes were seen and identified as mesocercariae of Alaria marcianae (Figures 12.127–12.128). In the same report, mesocercariae of Alaria mustelae and metacercariae of Fibricola crater were found in subcutaneous lesions and coelomic fat in red-sided garter snakes (T. s. parietalis) collected in central Manitoba, Canada.

12.6.4 Rhytidodidae Rhytidodoides similis, a member of the family Rhytidodidae, has been identified as a parasite of the gallbladder of green turtles, where papillomatous hyperplasia of the mucosa near the entrance to the cystic duct was seen (Smith et al., 1941). Sometimes the entire mucosa was affected.

12.6.5 Hemiuridae Members of this family are parasites of sea snakes (family Hydrophidae) and marine teleost fish. Hydrophitrema gigantica is a large trematode (17 mm to 26 mm in length) infecting the respiratory tract of the annulated sea snake (Hydrophis cyanocinctus) from the waters near Hong Kong (Ko et al., 1975). Mucoid exudate was seen in the lung along with focal hyperplasia, hemorrhage, necrosis, and inflammatory cell infiltrates at sites of attachment. Pulmovermis cyanovitellosus infects the lung of the Chinese sea snake (Laticauda semifasciata) from waters off the southern coast of Taiwan (Coil and Kuntz, 1960), and was reported as Laticaudatrema amamiensis from the trachea, lung, and air sac of the Chinese sea snake from Amami Island, Japan (Telford, 1967).

12.7 Nematoda While numerous nematodes have been described from reptiles (primarily wild reptiles), relatively few have been incriminated as significant pathogens. Some of the more important nematode parasites of reptiles are presented below.

12.7.1 Ascaridoidea The more important members of this superfamily infecting reptiles include the genera Angusticaecum (in tortoises), Anisakis (sea turtles), Sulcascaris (sea turtles), Dujardinascaris, Brevimulticaecum, Gedoelstascaris, and Multicaecum (in crocodilians), and Ophidascaris, Polydelphis, and Hexametra (snakes and lizards). Ascarids may cause lesions either as larvae migrating through visceral structures or as adults embedded within the gastrointestinal mucosa. Clinical signs (when present) associated with ascarid infection are nonspecific. Infected reptiles may be anorectic and may slowly lose weight. Reptiles, especially snakes, that do eat may regurgitate partially digested food within several days of feeding.

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Diagnosis of ascarid infection depends on identification of adult parasites in regurgitated food, or on postmortem evaluation, within the gastrointestinal tract. Migrating larvae may be identified on histological examination of tissues. Ascarid eggs can be identified in stomach washings and in a fecal flotation specimen (Figures 12.129–12.131); they are typically thick-shelled, unembryonated, and measure approximately 80 to 100 µm by 60 to 80 µm.

12.7.1.1 Chelonia  The ascarid Angusticaecum is common in the Greek tortoise (Holt et al., 1979). Migrating larvae of Angusticaecum may have been responsible for pulmonary lesions in a Greek tortoise, and in a few cases there was obstruction of the intestinal tract (Keymer, 1978). Angusticaecum was removed from the ear canal of a Greek tortoise, which was filled with debris that distended the right tympanic membrane (Cutler, 2004).  Members of the genus Anisakis are common parasites of the stomach of sea turtles in Australia (Burke and Rodgers, 1982). Most of the larvae were found coiled below the serosal surface of the stomach and intestine. In one turtle, 20 larvae were encysted below the capsule of the liver. There were also hemorrhagic ulcers in the pyloroduodenal region of the gastrointestinal tract associated with the presence of larvae. The source of infection was sardines (Harengula ovalis) that were being fed to the turtles. In a survey of diseases of farmed marine turtles in northern Australia, larval nematodes identified as Anisakis type I were found in the gastrointestinal tract and coelomic cavity of 15 of 104 sea turtles (mostly green turtles) (Glazebrook and Campbell, 1990a). Yearlings were more heavily infected compared to juveniles. Larvae were seen penetrating the wall of the gastrointestinal tract (where ulceration was seen) and migrating to the liver, spleen, and lung. Migrations resulted in adhesions between the gastrointestinal tract and peritoneal wall. Encysted parasites were seen in the peritoneal wall and capsule of the liver. In a report from the Canary Islands, Spain, Anasakis was seen embedded in the stomach of loggerhead sea turtles (Oros et al., 2004) (Figure 12.132). Larvae caused mucosal ulceration and granulomatous inflammation as they migrated from the luminal surface of the serosa (Figure 12.133). Larvae also migrated into the coelomic cavity where they caused perihepatitis. Anisakis has not been seen in sea turtles in Florida. Sulcascaris sulcata is another important ascarid that parasitizes sea turtles in warm and temperate waters around the world (Berry and Canon, 1981). Fourth stage larvae are found both in molluscan (scallops; Amusium balloti) intermediate hosts and in the stomach (near the esophageal juncture) of sea turtles, where large masses may be embedded in the stomach wall resulting in focal necrosis and inflammatory infiltrates. Adults are found in the stomach of sea turtles, where they attain considerable size (up to 111 mm in length) (Figure 12.134). Eggs measure approximately 61 to 75 µm by 86 to 100 µm (Figure 12.129). While originally

described from green turtles, Sulcascaris sulcata more commonly infects loggerhead sea turtles. This is consistent with their molluscan diet.

12.7.1.2 Crocodylia  Numerous genera of ascaridoids have been identified in crocodilians including Brevimulticaecum, Dujardinascaris, Gedoelstascaris, Goezia, Hartwichia, Multicaecum, Ortleppascaris, Tarranova, and Trispiculascaris. As adults they inhabit the gastrointestinal tract. Of 54 crocodiles (New Guinea and saltwater crocodiles) examined in a crocodile farm in Papua New Guinea, 22 had Gedoelstascaris mawsoni (synonym Dujardinascaris mawsoni) in the stomach (Ladds and Sims, 1990). In three they caused ulcers and granulomas in the stomach. I have necropsied several American alligators in Florida that were infected with Dujardinascaris in the stomach (Figure 12.135). In most cases, overt clinical signs of disease were not seen. 12.7.1.3 Squamata  Several species of Hexametra are recognized from lizards and snakes in Africa and Southeast Asia (Sprent, 1978). Hexametra angusticaecoides was reported from chameleons from Madagascar (Chabaud and Brygoo, 1960; Caballero Rodriguez, 1968), with infection occurring after ingesting an infected intermediate host (Chabaud et al., 1962). Despite the common occurrence of these parasites in lizards and snakes, there are few reports of associated mortality. A captive panther chameleon that died acutely without signs of illness was found to have H. angusticaecoides in the stomach and intestine (Coke, 1997). In the same report, a one-year-old pet veiled chameleon (Chamaeleo calyptratus), which was euthanized after manifesting respiratory distress, had 20 to 30 immature Hexametra in the coelomic cavity. Ascarid infections are common in wild snakes, where pathologic lesions have been reported. Third-stage larvae of Polydelphis quadrangularis will cause gastric ulceration in the definitive host, the Neotropical rattlesnake (Crotalus durissus terrificus), if they are less than 173 days of age (Araujo, 1971). Older larvae infecting a susceptible snake will complete the life cycle in the stomach. In experimental studies, P. quadrangularis reaches mature size at approximately 1 year after infection. Pythons are infected with several species of ascarids, where adults embed in the gastric mucosa and larvae migrate through visceral tissues creating mechanical damage. The inflammatory response with wandering larvae ranges from minimal to severe. A Papuan python (Apodora papuana [formerly Liasis papuanus]) that was in captivity for 4 years and died after 3 weeks of anorexia had adhesions between the aorta and esophagus (Hamir, 1986). The aortic wall contained numerous fibrous nodules, the endothelial surface was roughened, and many aneurysms were seen throughout the length of the aorta. Similar nodules were seen in the peritoneum of the coelomic cavity. One or more large nematodes were removed from each nodule and were identified as Ophidascaris pap-

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uanus; adults were found in the intestine. In addition to these lesions, using light microscopy, a bronchopneumonia and a large area of necrosis of one lung was seen. Other ascarids seen by Sprent (1963; 1970a, 1970b) from pythons include Ophidascaris moreliae from the carpet python (Morelia spilota variegata), Polydelphis anoura from carpet pythons, scrub pythons (Morelia amethistina), and the woma (Aspidites ramsayi), and Amplicaecum robertsi in the carpet python, where encapsulated larvae were seen in the aorta (Sprent 1963). Third-stage larvae of Ophidascaris moreliae were either found free or attached to the surface of the lungs (Sprent 1970a). Adults of Ophidascaris moreliae are found in the stomach and intestine. For several species of ascarids in snakes, adult parasites can cause lesions in the cranial gastrointestinal tract. Adult Ophidascaris are generally encountered within the caudal esophagus and stomach, where they often embed deep in the submucosa (Figure 12.136). Numerous ascarids may be seen projecting from a single focal ulcerating lesion, which gives it a medusoid appearance. A tremendous sclerotic inflammatory response is associated with these embedded parasites. Adults may be found free within the intestinal lumen and the bile and pancreatic ducts, where they may cause obstruction or perforation. Immature parasites may be found on the serosal surface of the stomach and adjacent to coelomic vessels and viscera (Figures 12.137–1.138). Secondary bacterial infection, particularly with Gram-negative microorganisms, is a common sequela to the gastrointestinal lesions.

12.7.2 Cosmocercoidea The superfamily Cosmocercoidea includes the family Atractidae, members of which are viviparous and have a direct life cycle. The eggs hatch and third-stage larvae develop in utero; they are capable of autoinfecting the host, resulting in very large infections. Pathologic lesions have been seen with the genus Proatractis, an inhabitant of the lower digestive tract of tortoises. Verminous colitis diagnosed in eight red-footed tortoises and three leopard tortoises was attributed to Proatractis (Rideout et al., 1987). Six of the tortoises also had Proatractis in the small intestine. Clinical signs in infected tortoises were either nonspecific or absent. Grossly, there was roughening and thickening of the caecum and proximal colon. Myriads of tiny nematodes (up to 1.0 cm) were seen on the mucosal surface and within the lumen of the colon; in six tortoises, nematodes were also seen in the small intestine. Microscopically there was extensive mucosal necrosis (Figure 12.139), with myriads of proatractids (up to 1.0 cm) embedded in the mucosa (Figure 12.140) and inflammatory infiltrates extending into the tunica muscularis. Proatractis was also seen within the colonic contents (Figure 12.141). One tortoise also had colitis due to Entamoeba invadens, and several tortoises were also infected with Kalicephalus.

12.7.3 Diaphanocephaloidea The superfamily Diaphanocephaloidea is one of several superfamilies within the order Strongylida. The most clinically significant member of this family in reptiles is Kalicephalus, the intestinal hookworm of snakes. They have a large complex oral cavity (Figure 12.142). Approximately two dozen species, each having a low degree of host specificity, are known. This is a relatively small parasite (1 to 1.5 cm long), which is often overlooked on postmortem evaluation. This nematode has a direct life cycle, and transmission occurs either orally through contaminated food or water or by the percutaneous route. Depending upon the species, the prepatent period may vary from two to four months. Most infections of Kalicephalus appear to be subclinical. Snakes with heavy infections may exhibit nonspecific signs of lethargy, debility, and anorexia. Severely infected snakes may pass bloody feces. Lesions include ulcerative enteritis and hemorrhage (Cooper, 1971). Gastric impaction, gastritis, and ulceration in a milk snake (Lampropeltis triangulum) were associated with Kalicephalus infection (Klaphake et al., 2005). Secondary Gram-negative microorganisms may invade the ulcerative lesions and further complicate the problem. Antemortem diagnosis depends upon demonstration of adult parasites or eggs within the feces. Eggs are thinly walled, measure approximately 70 to 100 µm by 40 to 50 µm (Figure 12.143), and may be embryonated by the time the feces are eliminated. Kalicephalus eggs must be differentiated from Rhabdias and Strongyloides eggs, which tend to be smaller and always contain larvae.

12.7.4 Filarioidea The superfamily Filarioidea (order Spirurida) includes several genera of filarial nematodes known to infect reptiles. Some of the more important genera include Foleyella, Macdonaldius, and Oswaldofilaria. Limited studies have shown that there seems to be a low degree of host specificity for these genera. As adults, all members of this group are found in extraintestinal sites (lungs, circulatory system, subcutaneous areas); they are either ovoviviparous or viviparous. Microfilariae are released into the circulatory system and transmission is achieved through bloodsucking arthropods, usually ticks and mosquitoes. Most reptilian filarial infections are diagnosed at necropsy, with relatively few reports documenting clinical signs and gross lesions associated with infection. Telford (1965) reported occlusion of the post vena cava by Macdonaldius in certain Mexican snakes.

12.7.4.1 Crocodylia  Several genera of filaroids have been reported in crocodilians. Of 54 crocodiles (New Guinea and saltwater crocodiles) examined in a crocodile farm in Papua New Guinea, Micropleura were found free in the abdominal cavity in two New Guinea crocodiles (Ladds and Sims, 1990). In three crocodiles at a farm in Irian Jaya, Micropleura were also found free in the coelomic cavity (Ladds et al., 1995). No

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pathologic lesions were reported associated with these infections. Other filaroids in crocodilians include the descriptions of Oswaldofilaria kanbaya by Manzanell (1986) from the coelomic cavity of saltwater crocodiles, O. medemi by Marinkelle (1981) from the smooth-fronted caiman (Paleosuchus trigonatus), and O. versterae by Bain et al.(1982) in Nile crocodiles.

12.7.4.2 Squamata  Three species of filaroids have been identified in the Gila monster (Heloderma suspectum). Smith (1910) was the first to report a filaroid from the Gila monster, which he named Filaria mitchelli. Chabaud and Frank (1961a) renamed the genus as Piratuba mitchelli, and Goldberg and Bursey (1990) provided a redescription of its microfilaria. Hannum (1941) recovered adults and microfilaria of a filaroid from a Gila monster, which he named Chandlerella corophila. Yamaguti (1961) reassigned this to the genus Splendidofilaria. Chabaud and Frank (1961a, b) identified Macdonaldius andersoni in the arterial system of Gila monsters. The author of this chapter received tissues from a Gila monster from Tucson, Arizona that had severe nephritis, with numerous microfilariae in interstitial vessels (Figures 12.144–12.145) and adult filaroids in renal arteries (Figure 12.146). While the arrangement of somatic nuclei and measurements of the microfilariae were compatible with those of P. mitchelli, a specific identification could not be made (Charles Bursey, personal communication). Foleyella is a common filaroid of certain agamid lizards and chameleons. Folyella philistinae infects the starred lizard (Agama stellio) in Lebanon (Schacher and Khalil, 1967), and F. agamae infects the common agama (Agama agama) in Nigeria (Obiamiwe et al., 1995). Foleyella commonly infects the subcutaneous tissues and coelomic cavity of chameleons (Figure 12.147). Bolette (1998a) collected F. candezei from the lateral abdominal and thoracic subcutaneous tissues of a Fischer’s chameleon (Bradypodion fischeri;). Frank (1981) identified F. furcata in the body cavity of an Oustalet’s chameleon (Chamaeleo oustaleti). Five live 3- to 6-cm long Foyella were surgically removed from a swollen right fluid-filled superior eyelid of another Oustalet’s chameleon (Thomas et al., 1996). An adult imported wild-caught Senegal chameleon was noted to have a nematode under the skin in the lateral region of the abdomen (Szell et al., 2001). Blood was obtained from the heart and smears revealed large numbers of microfilariae. The chameleon was treated with ivermectin and subsequently died. Both male and female nematodes were recovered from the chameleon and were identified as Foleyella furcata. Immunohistochemical detection of microfilariae of Foleyella was reported in an Oustalet’s chameleon using a polyclonal antibody against excretory and secretory products of an adult Dirofilaria immitis (Oros et al., 2002). Filaroids in the genus Madathamugadia parasitize gerrhosaurid and iguanid lizards of Madagascar, Turkmenistan, and Africa, and those in the genus Thamugadia parasitize geckonid lizards in North Africa and India, agamids in Leba-

non, and lacertids and agamids in Turkmenistan. Both genera are members of the filaroid subfamily Splendidofilarinae (Anderson, 2000). A filaroid nematode, which was removed from a subcutaneous mass in a female Madagascan giant day gecko (Phelsuma madagascariensis grandis), was identified as a member of the subfamily Splendidofilarinae (Haag et al., 2005). Several months later, because of dystocia, a celiotomy was performed; several additional filaroids were found encapsulated in the coelomic cavity and were removed. Because no male filaroids were found, they could not be assigned to a genus. Several species of Macdonaldius are found in snakes. Hull and Camin (1959) reported the death of a bull snake due to obstruction of the portal vein with Macdonaldius seetae. No premonitory signs of illness were seen. Macdonaldius oschei was identified as a cause of necrotizing dermatitis in reticulated pythons (Python reticulatus) in a zoological collection (Frank, 1964). Numerous parasites were found in the mesenteric arteries, with obstruction of peripheral capillaries by microfilariae, resulting in ischemic necrosis of the skin. Transmission was presumably from boa constrictors kept in the same exhibit, with an argasid tick (Ornithodoros talaje) possibly serving as the vector. Telford (1965) identified Macdonaldius oschei and M. colimensis from several species of snakes from Colima, Mexico, including the lyre snake (Trimorphodon biscutatus) and boa constrictor (Figures 12.148–12.149). Microfilaria found in a peripheral blood film of a wild-collected Trans-Pecos rat snake were identified as Macdonaldius seetae (Smith, 1997).

12.7.5 Dracunculoidea Members of the superfamily Dracunculoidea (order Spirurida) have been identified in several species of snakes. Three boa constrictors, one eastern king snake, and a monocled cobra (Naja naja kaouthia) with raised pustular lesions scattered over the body surface (Figure 12.150) were found to have numerous nematode larvae within impressions smears, wet mounts, and biopsy specimens of the lesions (Jacobson et al., 1986). Long slender nematodes identified as members of the Dracunculoidea were associated with each pustule (Figure 12.151). In the cobra, a subcutaneous unpustulated dracunculid was found in a lateral area of the midbody (Figure 12.152). Because liquefied pus rarely develops in response to pathogens in reptiles, perhaps substances released by these parasites resulted in the liquefaction of the necrotic material.

12.7.6 Diplotriaenoidea Diplotriaenoidea is a superfamily within the order Spirurida. Hastospiculum is a member of this superfamily that infects lizards and snakes. It is considered a primitive filarial nematode, having microfilariae in thick-shelled eggs. In tissue section, eggs are often in a cluster, with each surrounded by a

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clear area. The adults usually live in the lung where the eggs are deposited and are coughed up, swallowed, and passed in feces.

12.7.6.1 Squamata  Several Hastospiculum spp. have been identified in monitor lizards (Varanus spp.) (Figures 12.153– 12.154). Eggs released in the surrounding tissues can induce an inflammatory and fibrous connective tissue response (Figure 12.155). Adults of Hastospiculum bipinnatum, a parasite of the desert monitor (Varanus griseus), were found in the coelomic cavity (Ashour, 1994). Hastospiculum spiralis infects the Indonesian mangrove monitor (V. indicus) (Bolette, 1998b). Snakes are also infected with Hastospiculum. Araujo (1970) described Hastospiculum onchocercum major in the Neotropical rattlesnake (Crotalus durissus terrificus). The author of this chapter received tissues of a cloud-forest boa constrictor (B. c. nebulosis) that had eggs of Hastospiculum in a granulomatous mass extending from the submucosa of the esophagus to the pleura surface of the lung (Figure 12.156).

serving as the definitive host and beetles serving as the intermediate host.

12.7.9 Physalopteroidea Physalopteroidea is a superfamily within the order Spirurida. The single family, Physalopteridae, is subdivided into three subfamilies. Adult members of this family generally embed in the gastric mucosa. Several species of insects from different orders can serve as intermediate hosts. Lee (1957) described the life cycle of Skrjabinoptera phrynosoma, a common parasite of the Texas horned lizard (Phrynosoma cornutum). The intermediate host is the Texas agricultural ant (Pogonomyrmex barbatus). Physaloptera retusa has been identified in the gastrointestinal tract of lizards (Bursey and Goldberg, 1991) and snakes (Pfaffenberger et al., 1989) in North America. Pathologic change has not been reported in reptiles infected with these parasites.

12.7.10 Dioctophymatoidea

The Gnathostomatoidea is a superfamily within the order Spirurida. The spirurid genus Tanqua has been described from white-throated monitors (Varanus albigularis) and Nile monitors (Varanus niloticus) in South Africa (Hering-Hagenbeck and Boomker, 2000). Tanqua was identified in several monitors evaluated by the Zoological Medicine Service at the University of Florida in Gainesville (Figure 12.157). While no histological lesions were seen, parasitized lizards were anorexic, and when parasites were removed using a flexible endoscope inserted into the stomach, their appetites improved.

The superfamily Dioctophymatoidea (order Dioctophymatida) has members whose larvae are known to infect reptiles. Lichtenfels and Lavies (1976) reported that larvae of the Eustrongylides caused dermal lesions in red-sided garter snakes. These parasites are most often encountered at subcutaneous sites, in the lungs, and free within the coelomic cavity. Eustrongylides was embedded in the gastric serosa of one New Guinea crocodile in a farm in Irian Jaya, New Guinea (Ladds et al., 1995). Intermediate hosts of these parasites include fish and frogs. Adult parasites are located in the mucosa of the esophagus, proventriculus, and intestine of fish-eating birds. Reptiles probably function as a paratenic host.

12.7.8 Spiruroidea

12.7.11 Rhabditoidea

The superfamily Spiruroidea (order Spirurida) consists of four families, one of which is Spirocercidae. Members of this family require two hosts: insects in the orders Coleoptera and Odonata serve as the intermediate host, and certain mammals and birds as the final host; reptiles serve as a paratenic host. Third-stage larvae of the spirocercid Physocephalus sexalatus were identified in submucosal and mesenteric granulomas in a western diamondback rattlesnake (C. atrox), a mottled rock rattlesnake (C. lepidus lepidus), and a Mojave rattlesnake (C. scutulatus) (McAllister et al., 2004). Goldberg and Bursey (1988) reported finding larval nematodes of another spirocercid, Ascarops sp., in hepatic granulomas in the wild populations of western fence lizard (Sceloporus occidentalis). Ascarops sp. was also identified in gastric granulomas in the sagebrush lizard (S. graciosus). No signs of illness were seen in these lizards. Lizards were considered a paratenic (reservoir) host (Goldberg and Bursey, 1989), with wild and domestic pigs (and other mammals)

The superfamily Rhabditoidea is of clinical significance to snakes and includes the genera Rhabdias and Strongyloides. These parasites may exist in a free-living or a parasitic, parthenogenetic phase. Transmission is direct, and infection can occur without parasites going through the free-living stage. The parasitic phase is associated with adult nematodes in the lungs (Rhabdias) or intestinal tract (Strongyloides). There are several reports of clinical disease in snakes infected with these parasites. Infective larvae may directly penetrate the skin or may be ingested through contaminated food or water. Ingested larvae penetrate the oral mucosa, gain access to the circulatory system, and ultimately become distributed to the lungs. In the genus Rhabdias, the larvae will mature in the lung. Strongyloides larvae will ultimately pass up the trachea to the oral cavity and develop into adults in the intestines. Both genera produce similar appearing embryonated eggs. Clinical signs of illness may include respiratory disease, when adult Rhabdias are in the lungs or when larvae of

12.7.7 Gnathostomatoidea

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Strongyloides migrate through this organ. Snakes may exhibit open-mouth breathing, an extended glottis, and severe pneumonia, including a secondary bacterial pneumonia. A mucus-laden exudate may accumulate around the nares and glottis. At necropsy, exudate and necrotic cellular debris are often found in the major air passageways (Figure 12.158). A proliferative pneumonia may be seen in association with Rhabdias pulmonary infection (Figures 12.159–12.160). A ball python with stomatitis had a Rhabdias-like nematode in a tooth sulcus and sections of larval nematodes were seen in granulomas in the submucosa of the oral cavity (David Taylor, personal communication). Snakes with adult Strongyloides infection in the intestinal tract show nonspecific signs of illness including anorexia, weight loss, and lethargy. Infected snakes may develop diarrhea. Wiesman and Greve (1982) found Strongyloides mirzai associated with proliferative enteritis in a green tree python. It was believed that Strongyloides allowed secondary bacterial invasion by Gram-negative microorganisms. After examining a Strongyloides infection in a king snake, Holt et al. (1978) believed that loss of fluids and electrolytes in the intestine contributed to the death of the snake. Antemortem diagnosis of Rhabdias and Strongyloides infection depends upon demonstration of embryonated eggs, measuring approximately 60 µm by 35 µm, in a lung washing (Figures 12.161–12.162). It is virtually impossible to distinguish between embryonated eggs of Strongyloides and Rhabdias. First-stage Rhabdias larvae may also be found in a lung washing and can be differentiated from third-stage larvae of Strongyloides by the presence of a small, rhabditoid esophagus in the former and a very long esophagus in the latter,. Embryonated eggs of Rhabdias are often found within fecal flotation specimen and should be distinguished from the larger embryonated eggs of Kalicephalus.

12.7.12 Oxyuroidea Members of the superfamily Oxyuroidea are commonly encountered as colonic parasites of turtles and lizards. For the most part, oxyuroids have developed a commensal relationship with their host. In a study of oxyuroids in the gopher tortoise (Gopherus polyphemus), Bolson tortoise (Gopherus flavomarginatus), and desert tortoise, the following 15 species and subspecies were identified: Alaeuris mazzottii, A. paramazzottii, A. gopheri gopheri, A. gopheri pudica, A. gopheri macrolabiata, A. caballeroi, A. kinsellai kinsellai, A. kinsellai sonorae, A. longicollis, Gopheruris aspicula, Oxyuris, Tachygonetria macrolaimus tetrapapillata, T. dentata nearctica, Thaparia macrocephala, and Th. microcephala (Petter and Douglass, 1976). As many as nine oxyuroid species have been described in the green iguana, including those of the genera Alaeuris, Ozolaimus, Pseudalaeuris, and Tachygonetria (Yamaguti, 1961). In iguanids, each species may inhabit a different region of the large

intestine and may serve to mechanically break up ingesta and prevent cellulose impactions (Iverson, 1982). Only under unusual circumstances are oxyuroids pathogenic. Kane et al.(1976) reported a fatal impaction with oxyuroids (Alaeuris brachylophi) in a captive Fiji island iguana (Brachylophus fasciatus). An oxyuroid nematode in the genus Tachygonetria was probably pathogenic in a Hermann’s tortoise that died in a zoological collection (Keymer 1978). Oxyurid eggs are commonly seen in feces of herbivorous chelonians and lizards. Adults rarely cause health problems. The eggs are large, measuring approximately 130 µm by 40 µm (Figures 12.163–12.164).

12.7.13 Trichinelloidea The superfamily Trichinelloidea has members known to infect reptiles. For the most part, eggs of certain members such as Capillaria are identified in the feces of reptiles that are otherwise clinically healthy. Capillaria eggs have a thick shell and bipolar plugs (Figures 12.165–12.166). Some members of this group are identified only in tissues of infected reptiles. Crocodilians are the most important group of reptiles infected with these nematodes.

12.7.13.1 Crocodylia­  The genera of trichinelloids infecting crocodilians are Capillaria, Paratrichosoma, and Trichinella. Large numbers of Capillaria were identified in the stomach of a New Guinea crocodile in a crocodile farm in Irian Jaya, New Guinea (Ladds et al., 1995). There was a mixed inflammatory cell infiltrate in response to these parasites. The genus Paratrichosoma causes serpentine lesions in the skin of a number of species of crocodilians (Asford and Muller, 1978). They are typically seen on the ventral abdominal scales (Figure 12.167). Migration of the parasite through the keratin and cellular layers of the epidermis results in cystic spaces; eggs (27 µm by 73 µm) are deposited in these spaces (Buenviaje et al., 1998). Paratrichosoma crocodylus has been seen in crocodiles in New Guinea (Asford and Muller, 1978; Buenviaje et al., 1998) and Paratrichosoma recurvum from Morelet’s crocodiles (Crocodylus moreletii) in Mexico (Moravec and Vargas-Vazquez, 1998). Trichinella zimbabwensis was identified in skeletal muscles of 40% of farm-raised Nile crocodiles in Zimbabwe (Pozio et al., 2002), and T. papuae was identified in skeletal muscles of 22% of wild-born saltwater crocodiles in Papua New Guinea (Pozio et al., 2004). Both species of Trichinella are capable of experimentally infecting reptiles and mammals. Infection of crocodiles in Papua New Guinea probably occurred after collection by local people who fed them wild pig meat from a local market before sending them on to a crocodile farm in another province (Pozio et al., 2005).

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12.8 Acanthocephala The phylum Acanthocephala represents a small group of endoparasites that are commonly called spiny-headed worms. They are distinguished by their spine-covered proboscis and total lack of a digestive tract. In reptiles, adult parasites are generally found in the intestinal tract, whereas immature forms may migrate extensively; they can be found throughout serosal surfaces of the gastrointestinal tract and attached to the capsule of visceral organs. An arthropod serves as an essential intermediate host, although in many cases a reptile may act as a paratenic host. Adult acanthocephalan infections are generally diagnosed by identifying characteristic multienveloped ellipsoidal eggs in a fecal sample or within tissues (Figure 12.168).

12.8.1 Chelonia In the United States, several species of Neoechinorhynchus were identified by Fisher (1960), Cable and Fisher (1961), and Acholonu (1969) in the red-eared slider, Missouri slider (Pseudemys floridana hoyi), midland painted turtle (C. picta marginata), common snapping turtle, smooth softshell turtle (Apalone mutica [formerly Trionyx muticus]), spiny softshell turtle (Apalone [formerly Trionyx] spinifer), Blanding’s turtle (Emys blandingii), common map turtle (Graptemys geographica), false map turtle (G. pseudogeographica), Mississippi map turtle (G. kohni), and eastern box turtle (T. carolina carolina). While heavy infections may result in occlusion of the intestine, they are often found incidentally at necropsy and in tissue section (Figures 12.169–12.170).

12.8.2 Squamata An acanthocephalan, Porrorchis sp., was identified in one of 30 Tokay geckoes (Gekko gecko) evaluated for the presence of endoparasites (Reese et al., 2004). An adult king cobra (Ophiophagus hannah) was infected with mature Sphaerechinorhynchus serpenticola (identified by Brent Nickol) (Figure 12.171). They were located in the intestinal tract and attached to the capsule of the liver (Figure 12.172). Mature parasites in king cobras may be a result of the ophiophagous nature of these snakes. Cystacanths of the acanthocephalan Macracanthorhynchus ingens were identified in several species of snakes in Louisiana (Elkins and Nickol, 1983). Cystacanths of Centrorhynchus spinosus and Macracanthorhynchus ingens were identified in eastern indigo snakes in Florida (Foster et al. 2000). Cystacanths were encysted in the mesenteries and serosal surface of the small intestine, with fewer in the muscularis and even fewer in the lamina propria. An indigo snake that was submitted to the Pathology Service at the University of Florida in Gainesville was found to have severe colitis, and was infected with an acanthocephalan within the muscularis of the colon (Figure 12.173).

Acanthocephalan eggs within the wall of the colon were surrounded by a mild inflammatory response (Figure 12.174).

12.9 Pentastomida As adults, these wormlike parasites are superficially segmented; they range in size from 0.5 to 12 cm. Two compound pairs of hooks surround the mouth (Figures 12.175–12.176). Histologic features used in identifying these parasites in tissue section are acidophilic glands adjacent to the intestine (Figure 12.177) and sclerotized openings in the body wall (Figure 12.178). All require intermediate hosts for completion of their life cycle, and although approximately 90% of adult pentastomids are parasites of reptiles, Linguatula infects domestic mammals and Reighardia sternae infects gulls. The most important genera infecting reptiles include Sebekia (in crocodilians), Raillietiella (in lizards and snakes), and Kiricephalus, Porocephalus, and Armillifer (in snakes). Ingested larvae migrate through the gastrointestinal tract and may undergo extensive visceral migrations before they reach maturity either in the lung and air sac or in subcutaneous tissues. Pentastomids can cause localized lesions where direct damage to tissues can occur. Pentastomids may molt multiple times in their reptile host, with the molted cuticle capable of inducing antigenic stimulation (Riley, 1986). Pathologic changes may not be seen once molting has stopped. Mammals and fish serve as intermediate hosts, and several species within the genus Armillifer may infect humans. Numerous paratenic hosts have been identified for certain genera. Antemortem diagnosis depends upon identification of eggs in a lung washing or feces (Figures 12.179–12.181). For several genera, eggs have a distended, thinly walled capsule and measure up to 130 µm in diameter. Larvae, which contain hooklets, may be seen within the eggs. For some species, autoinfection may be possible. Often the inflammatory response in the lung is mild in reptiles with adult pentastomids. There is evidence suggesting that lung-inhabiting pentastomids, and also nymphs encysted in the tissues of mammalian intermediate hosts, evade immune surveillance and reduce inflammation by coating their chitinous cuticle with their own stage-specific surfactant (Riley and Henderson, 1999).

12.9.1 Chelonia Diesingia (family Sebekidae) is the major genus of pentastomids known to infect turtles (Riley, 1994). Diesingia megastoma was first recovered from the lungs of the Geoffrey’s side-neck turtle (Phrynops geoffroyanus), and later in the South American snake-neck turtle (Hydromedusa tectifera) from Brazil (Junker et al., 2003). While originally named Pentastoma megastomum, it was renamed Diesingia megastoma (synonym Sebekia megastoma). Diesingia kachugensis was described from the liver of the red-crowned roof turtle

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(Kachuga kachuga) from Southeast Asia. Lesions have not been described for Diesingia infection in turtles.

12.9.2 Crocodylia Five genera of pentastomids, four from the family Sebekidae and one from the family Subtriquetridae, infect the respiratory tract of crocodilians (Junker et al., 1999). Pentastomids in the genus Alofia have been reported in saltwater crocodiles in the Philippines (Riley, 1994). Multiple species in the genus Sebekia have been described from several species of crocodilians, including the American crocodile, an unidentified crocodile in Africa, the Nile crocodile, caiman (Caiman crocodylis [formerly sclerops]), and American alligator (Riley et al., 1990). In Africa, the Nile crocodile is infected with the genera Alofia, Leiperia, Sebekia, and Selfia (Riley, 1994; Riley and Huchzermeyer, 1995a; Riley et al., 1990). In a study in the Republic of the Congo, lungs and viscera of African dwarf crocodiles collected at a market had three species of pentastomids: Sebekia okavangoensis, Alofia parva, and an unidentified Sebekia sp. (Riley and Huchzermeyer, 1995b). Two Nile crocodiles (one clinically healthy and one debilitated) from Kruger National Park, South Africa, were necropsied and were found to be infected with A. nilotici, A. simpsoni, Leiperia cincinnalis, S. cesarisi, S. okavangoensis, and S. wedli (Junker et al., 1999). In the clinically healthy crocodile, areas of the respiratory tract where the parasites attached had an area of mild coagulative necrosis with infiltrates of heterophils, with hemorrhage into surrounding tissue. A pentastome was in the aortic lumen, attached to the endothelial lining. At the point of attachment there was ulceration of the lining and a mixed inflammatory response in the adjacent tissue. In a debilitated crocodile, females and nymphs of L. cincinnalis obstructed part of the bronchi and pulmonary aorta. Pentastomid migration tracts covered the outer surface of the trachea. There was chronic multifocal granulomatous pneumonia associated with intralesional pentastomids. Fungal hyphae were seen in these lesions. In the United States, Sebekia oxycephala is considered a common parasite of the pulmonary system of wild American alligators (Cherry and Ager, 1982; Deakins, 1971; Hazen et al., 1978). Pulmonary edema, hemorrhage, and necrosis have been seen associated with migration and the presence of Sebekia in lungs of American alligators (Figure 12.182). Several 4-week-old captive hatchling American alligators that developed signs of respiratory disease were euthanized, and were necropsied along with several that died; nymphal S. oxycephala were found in the pulmonary parenchyma, liver, and coelomic cavity (Boyce et al., 1984). Histologically there was pulmonary hemorrhage; no inflammatory response was noted (Figure 12.183). Mosquito fish (Gambusia affinis) that were fed to the alligators had a high prevalence of these pentastomes. Larvae dissected from fish and fed to the alligators resulted in similar clinical signs and pulmonary hemorrhage. In a subsequent report, Sebekia of American alligators was

considered distinct from S. oxycephala (originally described from African crocodiles) and was named S. mississippiensis (Overstreet et al., 1985). Clinically healthy American alligators in northern Florida had S. mississippiensis infection rates of 90 to 100% (Boyce, 1985). Although fish serve as the major intermediate host for S. mississippiensis, certain turtles, snakes, and mammals may also serve as sources of infection (Overstreet et al., 1985). Significant lesions were reported in paratenic hosts (mice, hamsters, the Florida red belly turtle [Pseudemys nelsoni], and a pig frog [Rana grylio]) experimentally infected with nymphs of Sebekia mississippiensis (Boyce and Kazacos, 1991). Mixed inflammatory cells were seen at sites where nymphs were located, and dead nymphs were surrounded by a granulomatous inflammatory response. Fatal pentastomiasis with S. oxycephala was reported in captive African dwarf crocodiles after they were fed wild-caught mosquito fish from the Silver Springs area of Florida (Adams et al., 2001). Using light microscopy, sections of these parasites were found in the lungs, between the liver and heart, between the liver and stomach, and embedded within the muscular tunic of the stomach. Of 54 New Guinea and saltwater crocodiles examined in a crocodile farm in Papua New Guinea, two had pneumonia associated with the presence of Sebekia sp. (Ladds and Sims, 1990). In these cases there was consolidation of the lungs or presence of Sebekia-containing dark foci under the pleura. Histologically, there was interstitial pneumonia, bronchiectasis, and hyperplasia of bronchiolar epithelium. In a crocodile farm in Irian Jaya, New Guinea, of 38 young New Guinea and saltwater crocodiles that either died or were euthanized, four (all New Guinea crocodiles) had pentastomiasis (Ladds et al., 1995). Pentastomes were seen as multiple subpleural black foci up to 15 mm in diameter. Although a specific identification was not made, they were more than likely Sebekia. Focal or diffuse exudative pneumonia was associated with these parasites or their eggs. Sebekia also has been reported in crocodiles in farming operations in Australia (Buenviaje et al., 1994). Dark foci seen in the lungs were composed of eggs and adult pentastomes, hemorrhage, and granulomatous inflammation.

12.9.3 Squamata Pentastomids infect a variety of lizards (Figure 12.184); the major genera are Elenia, Raillietiella, and Sambonia (Riley, 1986). Raillietiella has approximately 31 species, some of which also parasitize amphisbaenians and snakes (Riley, 1986). Raillietiella affinis, R. frenatus, and R. gehyrae have been reported from Tokay geckos (Reese et al., 2004), with R. frenatus and R. gehyrae using cockroaches as intermediate hosts (Ali and Riley, 1983). Gretillat and Brygoo (1959) reported R. chamaelonis from the Malagasy chameleon (Chamaeleo verrucosus) and Oustalet’s chameleon (C. oustaleti). Brygoo (1963) reported R. hemidactyli from the Malagasy chameleon, and McAllister et al. (1992) reported

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a new species of Raillietiella from Malagasy chameleons in Madagascar. A wild-caught Bosc’s (savannah) monitor, which died after exhibiting signs of chronic pneumonia, was found to have adult pentastomids of an undescribed Sambonia species in the lungs (Flach et al., 2000). Snakes are infected by the genera Armillifer, Cubirea, Gigliolella, Kiricephalus, Parasambonia, Porocephalus, Raillietiella, and Waddycephalus (Riley, 1986). Porocephalus crotali infect the lungs and air sacs of water moccasins (Agkistrodon piscivorus) and the eastern diamondback rattlesnake (Crotalus adamanteus) in the southeastern United States (Forrester et al. 1970) (Figure 12.185). Kiricephalus pattoni parasitizes different species of Asian (Taiwan) snakes, with adults located in both the lung and subcutaneous tissues (Self and Kuntz, 1967). Kiricephalus coarctatus infects eastern indigo snakes in the southeastern Unites States. Of 12 indigo snakes in Florida (Foster et al., 2000), eight were infected with K. coarctatus. Parasites distributed in subcutaneous sites may cause bulging of the overlying skin. Sometimes when an infected snake is handled, the adults will emerge through the skin (Self and Kuntz, 1967). Little to no inflammation will be seen at these locations. Adult pentastomids in snakes may be found in the coelomic cavity and attached to the liver. Armillifer armillatus and A. grandis infect pythons and vipers (Fain, 1961a). Armillifer moniliformis parasitizes primarily Southeast Asian pythons. There are several reports associating lesions with pentastomid infections. Kiricephalus coarctatus was associated with lung lesions in a garter snake (T. sirtalis); the embedded parasite penetrated into the hypaxial musculature (Deakins, 1972). I have seen pneumonia associated with Armillifer armillatus infections in gaboon vipers (Figures 12.186– 12.187). Deaths of snakes have been attributed to pentastomids occluding the trachea. Damage to the lung may result in secondary bacterial infections. Four of nine boa constrictors with pulmonary damage due to infection with Porocephalus dominicana, died of bacteremia and pericarditis (Riley and Walters, 1980).

12.10 Annelida 12.10.1 Hirudinea This subclass contains hundreds of species of marine, freshwater and terrestrial leeches. Leeches are common ectoparasites of aquatic reptiles, particularly turtles and crocodilians. The most common leeches in freshwater turtles in the southeastern United States are Placobdella papillifera, P. ornata, and P. parasitica. They are commonly found on the soft tissue along the neck (Figure 12.188) and also in the inguinal area. Placobdella nuchalis parasitizes freshwater turtles and alligators (Klemm, 1995). Other leeches reported on crocodilians include the following: Helobdella, Hirudinaria manillensis, Philobdella gracilis, Placobdella multilineata, Placobdella

papillifera, and Placobdella multistriatus (Huchzermeyer, 2003). Leeches may be found in the oral cavity of alligators (Figure 12.189). The most common leeches parasitizing sea turtles are Ozobranchus branchiatus and O. margoi. They have both posterior and anterior suckers. Ozobranchus branchiatus has seven pairs of branchiae (gills) and O. margoi has five pairs (Figure 12.190). Adult leeches attach to soft and hard tissues anywhere on the animal’s body, including the conjunctiva around the globes (Figure 12.191) and soft tissues around the limbs. Eggs are also deposited in mats almost anywhere on the body (Figure 12.192). In most cases there is little clinical significance associated with leech infestations. In severe cases, anemia and traumatic damage to tissues have been seen. Fibropapillomas seen in adult free-ranging green turtles are commonly infested with Ozobranchus. In Florida, green turtles with fibropapillomas had up to 50 leeches per 1.3 cm2 of tumor surface (Nigrelli and Smith, 1943). The ability of leeches to complete their entire life cycle on turtles contributes to the heavy burdens seen in debilitated turtles. In addition to anemia, leeches may serve as vectors for certain pathogens. Marine leeches (Ozobranchus spp.) were found to carry very high fibropapilloma-associated turtle herpesvirus DNA loads, with some approaching 10 million copies per leech (Greenblatt et al., 2004). This finding implicated the marine leech as a mechanical vector for the fibropapillomaassociated turtle herpesvirus.

12.11 Crustacea The subphylum Crustacea includes several classes of primarily estuarine and marine invertebrates. Of these, the order Cirripedia (barnacles) is known to parasitize sea turtles and sea snakes.

12.11.1 Cirripedia Barnacles are common external parasites of estuarine and marine turtles (Figure 12.193). Heavy barnacle burdens are commonly seen in chronically ill or dead, stranded sea turtles (Figures 12.194–12.197). Members of the families Balanidae and Lepadidae are facultative ectoparasites of marine turtles, while members of the family Cornulidae are obligate ectoparasites. Barnacles identified on the ornate diamondback terrapin (Malaclemys terrapin macrospilota) in Florida include Balanus eburneus, B. improvisus, Chelonibia manati lobatibasis, and C. testudinaria (Jackson et al., 1973; Seigel, 1983). In marine turtles the following barnacles have been reported: Conchoderma auritum, C. virgatum, Lepas anatifera, L. anserifera, and L. hillii on loggerhead sea turtles in Australia (Monroe and Limpus, 1979); Co. virgatum and Cylindrolepas darwini on Pacific (olive) ridley sea turtles (Lepidochelys olivacea) caught off the coast of California (Hubbs, 1977); B. amphitrite, B. enurneus, Chelonibia caretta, C. tes-

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tudinaria, Chthamalus fragilis, Chthamalus stellatus, L. anatifera and L. pectinata on loggerhead sea turtles in Georgia (Frick et al., 1998); Chelonibia testudinaria on green turtles, hawksbill sea turtles, and Kemp’s ridley sea turtles (Lepidochelys kempii) (Lauckner, 1985); Platylepas hexastylos on green turtles, loggerhead sea turtles, and hawksbill sea turtles (Monroe and Limpus, 1979; Schwartz, 1960); Platylepas coriacea on the leatherback sea turtle (Dermochelys coriacea) (Monroe and Limpus, 1979); Stephanolepas muricata in green turtles, hawksbill, and loggerhead sea turtles (Monroe and Limpus, 1979); and Stomatolepas spp. from green turtles, loggerhead sea turtles, Pacific (olive) ridley, and leatherback sea turtles (Monroe and Limpus, 1979). Barnacles can be either encrusting or embedding. Embedding barnacles, such as Stephanolepas and Stomatolepas, can cause local tissue damage (Figure 12.198). In addition to marine turtles, certain barnacles parasitize sea snakes. Jeffries and Voris (1979) reported Octolasmis grayi, Zann (1975) reported Platylepas ophiophilus, and Zann et al.(1975) reported Lepas anserifera, L. anatifera, and Conchoderma spp. from sea snakes.

Even under captive conditions, ticks rarely reach the burdens achieved by mites. Still, they are significant potential pathogens, causing anemia or toxicosis, and producing focal ulcerating skin lesions. While they can serve as vectors for the transmission of a variety of pathogens in mammals, there are very few reports of tick-transmitted pathogens of reptiles. As mentioned previously, Macdonaldius oschei was identified in captive reticulated pythons that developed a necrotizing dermatitis (Frank, 1964). Transmission was thought to have originated from boa constrictors kept in the same exhibit, with an argasid tick serving as the vector.

12.12.1.1.1 Chelonia  A variety of ixodid and argasid ticks have been reported to infest tortoises and turtles. Whereas ticks most commonly embed in the soft tissues around the limbs and cervical region, some may embed in the suture lines between adjacent scutes. Amblyomma dissimile, a generalist reptile parasite that more commonly infests the green iguana, was reported on the mud turtle (Kinosternon), side-neck river turtle (Podocnemis), and painted wood turtle (Rhinoclemmys) (Burridge and Simmons, 2003). Other neotropical Amblyomma ticks that are more of a chelonian specialist and frequently arrive in the United States on imported animals include A. argentinae, 12.12 Acari A. humerale (on tortoises), and A. sabanerae (on turtles) (James Mertins, personal communication). A native tick, Within the class Arachnida, the arthropodan subclass Acari Amblyomma tuberculatum, commonly infests wild gopher consists of mites and ticks. Aside from being a distinct lintortoises in the southeastern United States (Cooney and eage, ticks are simply a group of large mites. While detailed Hays, 1972) (Figure 12.199). Amblyomma clypeolatum was taxonomic information on ticks infesting reptiles can be found reported on Indian star tortoises (Geochelone elegans) (Senelsewhere (Barnard and Durden, 2000), there is no single eviratna, 1965), Hyalomma aegyptium on Hermann’s torsimilar source of information for mites that infest reptiles. toises (Sixl, 1971) and Greek (Mediterranean spur-thighed) The two major orders of mites having families with members tortoises (James W. Mertins, personal communication), and that are relevant to reptiles are the Acariformes and ParasitiAmblyomma chabaudi on Malagasy spider tortoises (Pyxis formes. The important families within these two orders with arachnoides) (Michel Burridge, personal communication) members that are ectoparasites of reptiles are listed below. (Figure 12.200). Commonly encountered ectoparasites of For the most part, there is little information on pathologic desert tortoises are the argasid ticks, Ornithodoros parkeri effects of mites on reptiles. and O. turicata (Ryckman and Kohls, 1962; Woodbury and Hardy, 1948). Ornithodoros turicata is also found in bur12.12.1 Parasitiformes rows of gopher tortoises in Florida (Figure 12.201). The only two argasid ticks that strictly parasitize tortoises are 12.12.1.1 Ixodidae and Argasidae (Suborder Ixodida)  A Ornithodoros transverus, which parasitizes the Galapagos wide variety of ticks infest reptiles including several marine giant tortoise (Geochelone nigra [formerly elephantopus]) host species. Taxonomically, ticks that infest reptiles are (Hoogstraal et al., 1973) and O. compactus, which parasitplaced in the family Ixodidae (hard ticks) or in Argasidae izes tortoises in Africa (Hoogstraal and Aeschlimann, 1982). (soft ticks). The more important hard-bodied genera infesting Ornithodoros transverus is the only tick in the world that reptiles include Amblyomma, Bothriocroton, and Hyalomma. spends its entire life cycle, including the egg stage, on its The most significant soft-bodied ticks on reptiles are Argas, host only. In 1997 in the United States, the African tortoise tick, Carios, and Ornithodoros species. Imported reptiles are commonly infested with tick ectoparasites. From 1962 to 2001, at Amblyomma marmoreum, was found on several species least 29 species of exotic ticks were imported and intercepted of tortoises in a private outdoor collection in central Florin the United States on imported reptiles (Burridge and Sim- ida (Allan et al., 1998) (Figures 12.202–12.203). Leopard mons, 2003). In Florida alone, at least eight exotic tick species tortoises from Zambia infested with the exotic African ticks (Amblyomma sparsum) were also being imported into Florwere identified on imported reptiles (Burridge, 2005). ida. These ticks were infected with Ehrlichia ruminantium,

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the causative agent of heartwater, a fatal disease of domestic and wild ruminants (Burridge et al., 2000a, b). Subsequent studies demonstrated that leopard tortoises are refractory to E. ruminantium infection, and are unlikely to introduce heartwater into new areas (Peter et al., 2001). 12.12.1.1.2 Crocodylia  Ixodid ticks have rarely been reported from crocodilians, and most records are doubtful. In what seems to be a valid occurrence, Amblyomma dissimile was reported from the Morelet’s crocodile (Crocodylus moreletii) in Belize (Rainwater et al., 2001). 12.12.1.1.3 Squamata  Numerous species of ixodid ticks (mostly Amblyomma) have been identified on lizards and snakes (Burridge and Simmons, 2003). Although some, such as Amblyomma (formerly Aponomma) komodoense and A. robinsoni, are host specific (e.g., infesting only Komodo monitors [Varanus komodoensis]), many are not so specialized (Burridge et al., 2004). For instance, as previously mentioned, the iguana tick, Amblyomma dissimile, though commonly infesting the green iguana, also has a very wide range of hosts including the boa constrictor and many other New World snakes, lizards, and chelonians from multiple families. Although not encountered as commonly as A. dissimile, A. scutatum has been found on the green iguana, other species of iguanid lizards, teiid lizards, and several species of snakes. By far the most common reptile tick in the international pet trade, and in public and private reptile collections, is the African snake tick, Amblyomma latum (formerly Aponomma). It is commonly seen in newly imported shipments of ball pythons from West Africa (Keirans and Durden, 2001). Three other Amblyomma reptile ticks, all former Aponomma spp., would probably rank as the next most common species on imported reptiles, especially monitors (James Mertins, personal communication). They are A. exornatum, A. flavomaculatum, and A. varanense. The first is from Asia and the East Indies, and the latter two are African. Less commonly seen is Amblyomma nuttalli, which has been identified on ball pythons and other African snakes and lizards imported into the United States. The Australian tick, Bothriocroton (formerly Aponomma) hydrosauri, has been reported from a wide variety of lizards and snakes in Australia. However, because of extremely limited exportation of reptiles from Australia, it is rarely seen outside that country. Lizards and snakes are also parasitized by argasid ticks. Carios (formerly Ornithodoros) darwini and C. (formerly Ornithodoros) galapagensis parasitizes the Galapagos marine iguana (Amblyrhynchus cistatus) and the Galapagos land iguanas (Conolophus pallidus and Co. subcristatus); and C. galapagensis additionally parasitizes the lava lizard (Microlophus [formerly Tropidurus] albimarlensis) in the Galapagos Islands (Keirans et al., 1980). Ornithodoros talaje has been identified as a parasite of rainbow boas (Epicrates cenchria) in Panama (Dunn, 1933).

Tick infestations are common on wild imported reptiles, but overt disease or health problems are rarely seen; however focal lesions may be seen at sites of attachment. Amblyomma exornatum, a tick commonly found in axillary areas and adjacent to the cloaca (Figure 12.204) of African monitor lizards, especially Nile monitors (Varanus niloticus), is also found in the nasal passages, where it can result in suffocation and death when infections are heavy (Norval, 1985; Young, 1965). This tick can also infest tortoises and snakes. Amblyomma flavomaculatum also similarly parasitizes African monitors, especially the savannah monitor. Ulceration at the site of attachment of the green iguana tick (Amblyomma dissimile) on rainbow boas (Epicrates cenchria) was reported (Dunn, 1918). Ticks are also known to serve as vectors of hemogregarines in several species of lizards and snakes (Burridge, 2005). Ornithodoros turicata may serve as a potential vector of Hepatozoon spp. in snakes in Florida (Wozniak and Telford, 1991). Transmission of the filarid Macdonaldius oschei from a boa constrictor to reticulated pythons in a zoological collection was attributed to an argasid tick (O. talaje) (Frank, 1964). Because certain wall-less bacteria (Spiroplasma) have been found in ticks in the United States, Ornithodoros parkeri were collected from desert tortoises and were cultured for Mycoplasma, a causative agent of upper respiratory disease of desert and other tortoises (Tully et al., 1998). Mycoplasma was not cultured from tick homogenates.

12.12.1.2 Ascidae (Suborder Mesostigmata)  Members of this family are free living and generally found in litter or soil humus. Nearly all known species are normally predators of small invertebrates, although a few species may also feed on pollen or fungi, and one Proctolaelaps species is possibly parasitic on a cockroach (Krantz 1978). Whereas infections in reptiles have not been previously reported, a Florida banded water snake (Nerodia fasciata pictiventris) was found with a heavy infestation of Proctolaelaps pygmaeus (Figure 12.205). Some opacity in the spectacle was seen along with thickening of the tissues at the fornix between the spectacle and the periocular scales. 12.12.1.3 Diplogyniidae (Suborder Mesostigmata)   Although members of the Diplogyniidae typically parasitize beetles, the unique species Ophiocelaeno sellnicki was identified from snakes in the Solomon Islands (Johnston and Fain, 1964).

12.12.1.4 Entonyssidae (Suborder Mesostigmata)  This poorly studied family consists of six or seven named genera worldwide (Fain, 1961b). The snake lung mites Entonyssus spp. and Hamertonia spp. are small and easily missed at necropsy. No significant pathologic change has been noted with their presence. Mites in the genus Mabuyonyssus parasitize the respiratory passages of certain skinks in Africa.

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12.12.1.5 Heterozerconidae  (Suborder Mesostigmata)   (1948) demonstrated that O. natricis is capable of transmitHeterozerconids are mostly parasites of the Diplopoda (millipedes). Infestations of reptiles by heterozerconids are limited to Heterozercon oudemansi from an imported rainbow boa (Epicrates cenchris) in a zoological collection (Finnegan, 1931), H. elegans from several species of snakes in Brazil (Lizaso, 1978/1979), and Zeterohercon amphisbaenae from a worm lizard (Amphisbaena) in Brazil (Flechtmann and Johnston, 1990).

ting the Gram-negative bacterium Aeromonas hydrophila. Some bacterial strains are extremely pathogenic and may result in fatal septicemia. Under unusual conditions, mites are capable of infesting humans, who may develop focal skin lesions. Mites have been circumstantially implicated in the transmission of the infectious agent of inclusion body disease of boid snakes (Chapter 9), but this has not been scientifically confirmed.

12.12.1.6 Ixodorhynchidae    (Suborder Mesostigmata)   12.12.1.8 Paramegistidae (Suborder Mesostigmata)  AlMembers of this group are exclusively associated with snakes. Within this family are the following genera: Ixodorhynchus (from snakes in North America, Cuba, and Mexico), Ixobioides (from snakes in Brazil), Hemilaelaps (from snakes in North America, Africa, Europe, and Asia), Asiatolaelaps (from snakes in Asia), and Strandtibbettsia (one species from Korea and one from Brazil) (Fain, 1962). Voss (1967) described H. philippinensis from the snakes Cyclocorus lineatus and Oligodon modestus in the Philippines. He considered Asiatolaelaps Fain a synonym of Hemilaelaps Ewing. Lizaso (1983) described two new genera (and two new species for each genus), Ophiogongylus and Chironobius, on colubrid snakes in Brazil.

though most members of this family are parasites of arthropods, the genus Ophiomegistus occurs on snakes (Fain, 1962; Voss, 1966) and skinks (Voss, 1966). Ophiomegistus alainae sp. nov.,O. brennani sp. nov., O. luzonensis, O. mabuyae, O. nihi sp. nov., O. novaguinea sp. nov., and O. sarawakensis sp. nov. were identified from skinks (Emoia, Mabuya, Sphenomorphus) in New Guinea, Java, and Sarawak (Goff, 1980). Domrow (1984) described O. blumi, O. iriani sp. nov., and O. joppae from skinks (Sphenomorphus and Emoia) in New Guinea, O. australicus from an Australian Major skink (Egernia frerei), and O. luzonensis from the Pacific coral snake (Micropochis ikaheka) in New Guinea.

12.12.1.7 Macronyssidae (Suborder Mesostigmata)  The

though members of the Schizogyniidae typically parasitize beetles, there is a unique species (Indogynium lindbergi) in this family that is limited to burrowing snakes in India (Sellnick, 1954).

family Macronyssidae includes Ophionyssus, the most common genus of mites that infest captive lizards and snakes. Of the 15 species within this genus, 13 are confined to lizard hosts (Bannert et al., 2000). The lacertid Gallot’s lizard (Gallotia galloti eisentrauti), which is confined to the Canary Island of the Tenerife Islands is infested with Ophionyssus galloticolus (Bannert et al., 2000). Ophionyssus natricis is the mite most commonly seen on captive snakes (Figures 12.206–12.209) and has been reported on wild snakes and lizards, where integumental lesions have been seen (Goldberg and Bursey, 1991b). Severe burdens are commonly encountered in recently imported snakes maintained under crowded, filthy conditions. The entire life cycle requires approximately 10 to 32 days, and a single female can lay up to 80 eggs. Eggs are deposited off the host in the immediate environment of the snake. Eggs produce larvae that transform into protonymphs, then into deutonymphs, and finally into adults. Only protonymphs and adults feed on reptiles. The larval and deuteronymphal stage are brief, free living, and mostly quiescent (Wozniak and DeNardo, 2000). While fertilized eggs develop into females, males are produced parthenogenetically.   Severe infestations of Ophionyssus may result in a debilitated, anemic snake. Mites often occur in the axes between scales and in the sulcus formed between the spectacle and the periocular scales. The conjunctiva may become swollen and edematous. Inflammatory cells may infiltrate feeding sites. Ophionyssus saurarum can serve as a vector for the hemogregarine Karyolysus (Svahn, 1975). Additionally, Camin

12.12.1.9 Schizogyniidae (Suborder Mesostigmata)  Al-

12.12.1.10 Uropodidae (Suborder Mesostigmata)  Lizards may serve as phoretic (transport) hosts for uropodid (Fuscuropoda marginata) deutonymphs (Mertins and Hartdegen, 2003). Fuscuropoda marginata may attach to skinks when its preferred hosts (beetles) are absent. Other uropodids in the Australian region may attach to lizards in sufficient numbers to cause morbidity.

12.12.2 Acariformes 12.12.2.1 Cloacaridae (Suborder Prostigmata)  The family Cloacaridae represents a highly evolved family of exclusive parasites of chelonians. Camin et al. (1967) reported on the presence of members of this family of mites within the cloaca of snapping turtles and painted turtles, and Pence and Wright (1998) reported Chelonacarus elongatus n. gen., s. sp. in green turtles. Adult mites are in a submucosal location and nymphs are on the mucosal surface. They are microscopic mites, approximately 300 µm long.

12.12.2.2 Ophioptidae (Suborder Prostigmata)  Members of the family Ophioptidae are ectoparasites of certain snakes from widely scattered geographic areas of the world. The type genus and species Ophioptes parkeri was described

596  Parasites and Parasitic Diseases of Reptiles

by Sambon (1928) from the colubrid snake Aesculapian false coral snake (Erythrolamprus aesculapii) from Bolivia. Mites were found embedded in the keratin of infested scales. Ophioptes samboni was described by Southcott (1956) from the narrow-banded snake (Simoselaps [formerly Rhynchoelaps] fasciolatus) in Australia. Fain (1962) listed nine additional species of Ophioptes and described Afrophioptes, a new genus of mites on colubrid snakes in central Africa. Lizaso (1980/1981) described Ophioptes brevipilis sp. nov. and O. longipilis sp. nov. from the false coral snake (Oxyrhopus trigeminus) and O. brevipilis sp. nov. from Boettger’s sipo (Chironius flavolineatus).

12.12.2.4 Trombiculidae and Leeuwenhoekiidae (Suborder Prostigmata)  Larval members of the mite families

Trombiculidae and Leeuwenhoekiidae are the chiggers or chigger mites. They have a worldwide distribution. The larval stages of all species feed parasitically on other animals, primarily vertebrates, but the postlarval stages are free-living predators on small soil arthropods. Most chiggers are not very host specific, but many species sometimes infest a variety of terrestrial reptiles, including chelonians. Of 178 Texas tortoises, one was heavily infested with the trombiculid mite Eutrombicula cinnabaris (formerly alfreddugesi) (Goff and Judd, 1981). This mite is also commonly seen on the gopher tortoise in the southeastern United States (Wharton and 12.12.2.3    Pterygosomatidae (Suborder Prostigmata)   Fuller, 1952). Numerous species of lizards are also parasitized The family Pterygosomatidae parasitizes both Old and New with trombiculid mites (Wharton and Fuller, 1952). Seven speWorld lizards, including members of the families Gekkoni- cies of trombiculid mites were described from Yarrow’s spiny dae, Agamidae, Cordylidae, Eublepharidae, Teiidae, Xantusi- lizard (Bennett, 1977). Neotrombicula californica was identiidae, and Iguanidae (Frank, 1984b). The nine genera known fied on the sagebrush lizard and the side-blotched lizard (Uta to infest lizards include Cyclurobia, Geckobia, Geckobiella, stansburiana) (Goldberg and Bursey, 1991a). Chiggers and Hirstiella, Ixoderma, Pterygosoma, Scaphotrix, Tequisist- some other mites may inhabit mite pockets, skin invaginations lana, and Zonurobia. Certain Hirstiella spp. are common in the cervical region, axillary areas, and postfemoral region on chuckwallas (Sauromalus spp.), lizards native to the in lizards (Arnold, 1986). Chiggers are also common at other southwestern United States and Mexico, and green igua- sites, such as around tympanic membranes (Figure 12.212). nas imported into the United States (Figure 12.210). They Eutrombicula lipovskyana were identified in mite pockets may serve as vectors for hemogregarines (Lewis and Wag- of Yarrow’s spiny lizard from Arizona, where they remained ner, 1964). Newell and Ryckman (1964) reported Hirstiella attached up to 52 days (Goldberg and Bursey, 1993). The pyriformis from captive San Esteban chuckwallas (Sauroma- trombiculid mites Vatacarus ipoides and V. kuntzi occur in lus varius) originating from San Esteban Island in the Gulf the air sacs and trachea of sea snakes in the genus Laticauda of California, and Werman (1983) studied the population (Nadchatram and Audy, 1965; Nadchatram and Radovsky, dynamics of Hirstiella pyriformis on wild chuckwallas (S. 1971; Southcott, 1957), and V. intermedius in the nasal fossae obesus) in the vicinity of Baker, California. Larvae accounted of the Galapagos marine iguana (Amblyrhynchus cristatus) for approximately 92% of the mites collected from these (Vercammen-Grandjean and Watkins, 1965). lizards. The most frequently infested sites were the dorsal and lateral surfaces of the tail. In another report (Mader et al., 1986), a colony of chuckwallas was infested with 12.13 Diptera Hirstiella trombidiiformis. Hirstiella stamii was described from specimens collected from captive green iguanas in the Numerous species of adult Diptera of several families (CeratoAmsterdam Zoological Gardens, Netherlands (Jack, 1961). pogonidae, Culicidae, Psychodidae, Tabanidae) are known to Hirstiella diolii was described from an infested colony of bite, feed upon, or obtain blood meals from reptiles (Barnard captive rhinoceros iguanas (Cyclura cornuta) at the London and Durden, 2000), and many may transmit infectious agents Zoo (Baker, 1998), and it was later found on imported Fijian to reptiles. Several species of flies are known to parasitize crested and banded iguanas (Brachylophus vitiensis and B. reptiles directly in their larval (maggot) stages, with many fasciatus, respectively), green iguanas, and rhinoceros igua- such cases of myiasis involving terrestrial chelonians. Myiasis nas at the Taronga Zoo, Australia (Walter and Shaw, 2002). of lizards or their eggs also has been seen. Several examples Pterygosomatids are generally found at periocular sites, will be presented. tympanic scale, axillae, and at the bases of cutaneous spines found on the dorsal midline of certain iguanid lizards (Fig12.13.1 Chelonia ure 12.211). Some Geckobia and Pterygosoma spp. reside in pocket-like skin folds at the base of the host’s neck on North American box turtles seem particularly prone to infesgekkonids and agamids, respectively (Arnold, 1986; Ber- tation with sarcophagid (flesh) fly larvae. In severe infestatrand and Modry, 2004). An acute dermatitis was reported tions, maggots may be diffusely distributed throughout many in a captive Yarrow’s spiny lizard (Sceloporus jarrovii) that subcutaneous sites. Larval stages of the flesh fly Cistudinowas diffusely infested with Hirstiella sp. (Goldberg and Hol- myia cistudinis frequently parasitize native box turtles and shuh, 1993). gopher tortoises in the southeastern United States. This parasite was first described in 1916 as Sarcophaga cistudinis, and

Parasites and Parasitic Diseases of Reptiles  597

was subsequently renamed Cistudinomyia (Townsend, 1917). Work on the life cycle demonstrated that adult flies deposit live larvae directly on the host (Knipling, 1937). However, larvae are unable to penetrate intact skin, requiring breaks in the integument to gain access to subcutaneous sites. The larvae can cause significant tissue damage and death. In Florida, five exotic Aldabra tortoises (Dipsochelys dussumieri [formerly Geochelone gigantea]) in outdoor enclosures were found parasitized by larvae of this dipteran, with lesions located primarily on the perineum and caudal aspects of the hind legs (Stover et al., 1989). Following removal and cleaning of the cavity with dilute povidone-iodine solution, the lesions healed in approximately five weeks. Infestations with Cistudinomyia cistudinis also were seen in exotic Hermann’s and Greek tortoises kept in outdoor pens in Florida. Other dipteran larvae are known to parasitize and kill turtle eggs (Acuna-Mesen and Hanson, 1990; Iverson and Perry, 1994; Lopes, 1982).

12.13.2 Squamata Myiasis of several species of lizards has been reported. Sarcophagid fly larvae (Anolisimyia blakeae) were removed from a subcutaneous lesion in the American (green) anole (Dodge, 1955) and Khan and Khan (1984) reported sarcophagid fly larvae (Sarcophaga crassipalpis) from Uromastyx in Pakistan. An internal infection was seen with the sarcophagid fly Balesoxipha plintopyga in the Texas spotted whiptail lizard (Cnemidophorus gularis) (Whitworth and Wangberg, 1985). Pharyngeal myiasis with larvae of sarcophagid flesh flies was reported in two common ameivas (Ameiva chrysolaema) from the Dominican Republic (Smith et al., 1994). Sarcophagid fly larvae infest clutches of eggs of the fence lizard (Sceloporus undulatus) (Trauth and Mullen, 1990) and corn snake eggs incubated under laboratory conditions were parasitized by dipteran larvae of the family Phoridae (Wilson, 2000).

Acknowledgments The author thanks many colleagues for providing images and papers not easily obtained in the United States. Those providing images are acknowledged in the figure legends for this chapter. Many of the cases presented in this chapter represent client-owned animals submitted to the Zoological Medicine Service, College of Veterinary Medicine, University of Florida, Gainesville. Many cases resulted in research projects designed to better understand the pathogenesis of specific diseases. Thanks go to Chris Gardiner, Michael M. Garner, Stephen Goldberg, Ellis Greiner, and James W. Mertins for reviewing this chapter and offering comments and suggestions for improvement.

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Wozniak EJ, McLaughlin GL, and Telford SR Jr. 1994. Description of the vertebrate stages of a hemogregarine species naturally infecting Mojave desert sidewinders (Crotalus cerastes cerastes). J Zoo Wildl Med 25:103–110. Wozniak EJ, Telford SR, DeNardo DF, McLaughlin GL, and Butler JF. 1998. Granulomatous hepatitis associated with Hepatozoon sp. meronts in a southern water snake (Nerodia fasciata pictiventris). J Zoo Wildl Med 29:68–71. Wright K, Tousignant A, Overstreet R, Shoop W, Jacobson E, and Greiner E. 1989. Mesocercaiae infections in a Texas indigo snake and red-sided garter snakes, in Third International Colloquium on the Pathology of Reptiles and Amphibians, Abstracts, Orlando, FL, 54. X iao L, Ryan UM, Graczyk TK, Limor J, Li L, Kombert M, Junge R, Sulaiman IM, Zhou L, Arrowood MJ, Koudela B, Modry D, and Lal AA. 2004. Genetic diversity of Cryptosporidium spp. in captive reptiles. Appl Environ Microbiol 70:891–899. Yamaguti S.1961. Systema Helminthum, Volume III. The Nematodes of Vertebrates. Part II. Interscience Publishers, Inc., New York. Young E. 1965. Aponnoma exornatum (Koch) as a cause for mortality among monitors. J S Afr Vet Med Assoc 36:579.

Zann LP. 1975. Biology of a barnacle (Platylepas ophiophilus Lanchester) symbiotic with sea snakes, in The Biology of Sea Snakes, Dunson WA (Ed.), University Park Press, Baltimore, MD, 267–286. Zann LP, Cuffey RJ, and Kropach C 1975. Fouling organisms and parasites associated with the skin of sea snakes, in The Biology of Sea Snakes, Dunson WA (Ed.), University Park Press, Baltimore, MD, 251–265. Zwart P. 1964. Intraepithelial protozoan, Klossiella boae n. sp., in the kidneys of a boa constrictor. J Protozool 11:261–263. Zwart P. 1973. Ziekten van reptielen V: infectiezikten. Lacerta 31:116–120. Zwart P and Truyens EHA. 1975. Hexamitiasis in tortoises. Vet Parasitol 1:175–183. Zwart P, Tennis SFM, and Cornelissen JMM. 1983. Monocercomoniasis in reptiles, in Proceedings of the First International Colloquium on Pathology of Reptiles and Amphibians, Vago C, Matz G (Eds.), Les Presses de l’Université d’Angers, Angers, France, pp. 73-76.

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Figure 12.1  Common agama, Agama agama. Agamidae. Photomicrograph of schizont of Plasmodium giganteum in a red blood cell. Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 12.2  Japanese grass lizard, Takydromus tachydromoides. Lacertidae. Photomicrograph of gametocytes of Plasmodium sasai in multiple red blood cells. Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 12.3  Puerto Rican anole, Anolis pulchellus. Iguanidae. Photomicrograph of schizont of Plasmodium floridense in a red blood cell. Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 12.4  Dwarf gecko, Lygodactylus capensis grotei. Gekkonidae. Photomicrograph of Haemocystidium lygodactyli in multiple red blood cells. Giemsa stain. Courtesy of Sam R. Telford Jr.

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Figure 12.5  Japanese grass lizard, Takydromus tachydromoides. Lacertidae. Photomicrograph of Aegyptianella in a red blood cell. Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 12.6  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of Hepatozoon fusifex in a red blood cell. Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 12.7  Turnip-tailed gecko, Thecadactylus rapicauda. Gekkonidae. Photomicrograph of Plasmodium aurulentum (arrow) and Trypanosoma thecadactyli in a peripheral blood film. Giemsa stain. Courtesy of Sam R. Telford Jr.

Figure 12.8  Flathead leaf-toed gecko. Hemidactylus platycephalus. Gekkonidae, Photomicrograph of Sauroleishmania in a red blood cell. Giemsa stain. Courtesy of Sam R. Telford Jr.

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Figure 12.9  Island night lizard, Xantusia riversiana. Xantusiidae. Photomicrograph of Balantidium in a fecal specimen. Trichrome stain. Courtesy of Sam R. Telford Jr. Bar = 20 µm.

Figure 12.10  Island night lizard, Xantusia riversiana. Xantusiidae. Photomicrograph of Nyctotherus in a fecal specimen. Note the size difference with Balantidium in Figure 12.9. Trichrome stain. Bar = 20 µm. Courtesy of Sam R. Telford Jr.

Figure 12.11  Desert tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of Nyctotherus in the colon lumen. There is no inflammatory response. H&E stain. Bar = 100 µm.

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Figure 12.12  Red-footed tortoise, Geochelone carbonaria. Testudinidae. Amoebiasis. A diffusely necrotic liver (L) is seen directly caudal to the heart (H).

Figure 12.13  Red-footed tortoise, Geochelone carbonaria. Testudinidae. Amoebiasis. The liver has large areas of necrosis (N) and the mucosal surface of the small intestine (arrowheads) is diffusely necrotic. (From Jacobson ER et al. 1983. J Amer Vet Med Assoc 183:1192–1194. With permission.)

Figure 12.14  Red-footed tortoise, Geochelone carbonaria. Testudinidae. Amoebiasis. Photomicrograph of the small intestine with a diffusely necrotic mucosal surface. H&E stain.

Figure 12.15  Red-footed tortoise, Geochelone carbonaria. Testudinidae. Amoebiasis. Photomicrograph of trophozoites (arrows) of Entamoeba invadens in the submucosa of the small intestine. H&E stain.

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Figure 12.16  Red-footed tortoise, Geochelone carbonaria. Testudinidae. Amoebiasis. Photomicrograph of trophozoites of Entamoeba invadens in the liver. H&E stain.

Figure 12.17  Wood turtle, Glyptemys (Clemmys) insculpta. Emydidae. Amoebiasis. Photomicrograph of trophozoites of Entamoeba in an impression smear of the liver. Wright-Giemsa stain. Courtesy of Amy MacNeill.

Figure 12.18  Wood turtle, Glyptemys (Clemmys insculpta). Emydidae. Amoebiasis . Photomicrograph of trophozoites of Entamoeba in the liver. PAS stain. Courtesy of Amy MacNeill.

Figure 12.19  Rat snake, Elaphe sp. Colubridae. Amoebiasis. Photomicrograph showing diffuse necrotizing colitis. Entamoeba was identified in the mucosa and submucosa.

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Figure 12.20  Boa constrictor, Boa constrictor. Boidae. Amoebiasis. Hepatic necrosis is seen. Trophozoites reached the liver via the portal tract from the intestine.

Figure 12.21  Russell’s viper, Vipera russellii. Viperidae. Amoebiasis. Photomicrograph of the intestine showing areas of necrosis (arrows) of the mucosa and infiltrates of mixed inflammatory cells. H&E stain. Courtesy of John Roberts.

Figure 12.22  Russell’s viper, Vipera russellii. Viperidae. Amoebiasis. Photomicrograph of trophozoites (arrows) in the submucosa of the intestine. Mucous epithelial cells (arrowheads) stain more intensely positive than the trophozoites. PAS stain. Courtesy of John Roberts.

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Figure 12.23  Home’s hingeback tortoise, Kinixys homeana. Testudinidae. Hexamitiasis. Photomicrograph of renal tubules that are dilated and filled with Hexamita. H&E stain. Bar = 50 µm. Courtesy of the Veterinary Pathology Service, University of Florida, Gainesville.

Figure 12.24  Home’s hingeback tortoise, Kinixys homeana. Testudinidae. Hexamitiasis. Higher magnification photomicrograph of Figure 12.23. Hexamita is seen within a dilated renal tubule. H&E stain. Bar = 20 µm. Courtesy of the Veterinary Pathology Service, University of Florida, Gainesville.

Figure 12.25  Diamond python, Morelia spilota. Pythonidae. Monocercomoniasis. Photomicrograph of Monocercomonas (arrows) within the gallbladder lumen, adjacent to the mucosa (M). Processing tissues for light microscopy causes the organism to become round. The flagella are also difficult to visualize. Giemsa stain. Bar = 50 µm.

Figure 12.26  Diamond python, Morelia spilota. Pythonidae. Monocercomoniasis. Photomicrograph of Monocercomonas in an impression smear from the gallbladder. A more typical morphology is seen compared to the same organism (Figure 12.25) in tissue processed for histology. Wright-Giemsa stain.

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Figure 12.27  Corn snake, Elaphe guttata guttata. Colubridae. Monocercomoniasis. Photomicrograph of Monocercomonas (arrows) within dilated gastric glands in a biopsy specimen. PAS stain.

Figure 12.28  Corn snake, Elaphe guttata guttata. Colubridae. Monocercomoniasis. Higher magnification photomicrograph of a gastric biopsy specimen from a corn snake. Monocercomonas (arrows) is seen within a dilated gland. PAS stain.

Figure 12.29  Helmeted iguana, Corytophanes cristatus. Iguanidae. Coccidiosis. Photomicrograph of Eimeria sp. (arrows) on the mucosal epithelial surface in the gallbladder. Inset: Higher magnification of Eimeria. H&E stain.

Figure 12.30  Mediterranean gecko, Hemidactylus turcicus. Gekkonidae. Nomarski interference-contrast photomicrograph of a sporulated oocyst of Eimeria turcicus recovered from the gallbladder. (From Upton SJ et al. 1988. J Protozool 35:24–25. With permission.)

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Figure 12.31  Inland bearded dragon, Pogona vitticeps. Agamidae. Coccidiosis. Photomicrograph of Isospora amphiboluri (arrows) in enterocytes. H&E stain.

Figure 12.32  Yellow-headed gecko, Gonatodes albogularis. Gekkonidae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Isospora albogularis recovered from the feces. SB = Stieda body; RB = refractile body. (From Upton SJ and Freed PS. 1989. Can J Zool 68:1266–1267. With permission.)

Figure 12.33  Wood turtle, Glyptemys (Clemmys) insculpta. Emydidae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Eimeria lecontei recovered from the feces. (From Upton SJ et al. Acta Protozool 34:57–60. With permission.)

Figure 12.34  Radiated tortoise, Geochelone radiata. Testudinidae. Intranuclear coccidiosis. Photomicrograph of intranuclear coccidial organisms (arrows) in the colon of a radiated tortoise. Toluidine blue stain. (From Jacobson ER et al. 1994. J Zoo Wildl Med 25:95–102. With permission.)

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Figure 12.35  Radiated tortoise, Geochelone radiata. Testudinidae. Intranuclear coccidiosis. Transmission electron photomicrograph of the colon. Stages of a coccidial organism are seen within nuclei. Uranyl acetate and lead citrate stain. (From Jacobson ER et al. 1994. J Zoo Wildl Med 25:95–102. With permission.)

Figure 12.36  Radiated tortoise, Geochelone radiata. Testudinidae. Intranuclear coccidiosis. Photomicrograph of intranuclear coccidial organisms in a pulmonary epithelial cell (arrow). H&E stain.

Figure 12.37  Leopard tortoise, Geochelone pardalis. Testudinidae. Intranuclear coccidiosis. Photomicrograph of intranuclear coccidial organisms (arrows) within cells lining the inner ear. Courtesy of Michael M. Garner. H&E stain.

Figure 12.38  Leopard tortoise, Geochelone pardalis. Testudinidae. Intranuclear coccidiosis. Transmission electron photomicrograph of an intranuclear coccidian within cells lining the inner ear. Uranyl acetate and lead citrate stain. Courtesy of Michael M. Garner.

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Figure 12.39  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. An emaciated turtle with disseminated Caryospora infection. Courtesy of Anita Gordon.

Figure 12.40  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. The intestinal mucosa is hyperemic and small amounts of fibrinous material can be seen on the surface. Courtesy of Anita Gordon.

Figure 12.41  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. Diffuse necrosis of the intestine is seen. Courtesy of Anita Gordon.

Figure 12.42  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. Photomicrograph of a duodenal mucosa with a hyperplastic epithelium. Type I meronts (arrows) of Caryospora are within enterocytes. H&E stain. Bar = 20 µm. (From Gordon AN et al. 1993. J Wildl Dis 29:490-494. With permission.)

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Figure 12.43  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. Photomicrograph of Caryospora gametogony in small intestinal epithelium. Macrogametocytes (short arrows), microgametocytes (long arrows), and oocysts (O) are seen. H&E stain; Bar = 50 µm. Courtesy of Anita Gordon.

Figure 12.44  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. Photomicrograph of forebrain with Type II meronts around a blood vessel. H&E stain. Bar = 50 µm. (From Gordon AN et al. 1993. J Wildl Dis 29:490–494. With permission.)

Figure 12.45  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. Photomicrograph of Type III meront (M) of Caryospora in the thyroid. H&E stain. Bar = 25 µm. Courtesy of Anita Gordon.

Figure 12.46  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. Photomicrograph of an unstained wet mount of sporocysts and sporozoites of Caryospora cheloniae. Bar = 20 µm. (From Gordon AN et al. 1993. J Wildl Dis 29:490-494. With permission.)

Figure 12.47  Green turtle, Chelonia mydas. Cheloniidae. Coccidiosis. Caryospora infection. Photomicrograph of a star formation following excystment of C. cheloniae sporozoites. Two sporocysts also can be seen. Bar = 20 µm. (From Gordon AN et al. 1993. J Wildl Dis 29:490–494. With permission.)

620  Parasites and Parasitic Diseases of Reptiles

Figure 12.48  Hermann’s tortoise, Testudo hermanni. Testudinidae. Cryptosporidiosis. Photomicrograph of Cryptosporidium on the surface of small intestinal enterocytes. H&E stain. Courtesy of Michael M. Garner.

Figure 12.49  Box turtle, Terrapene sp. Emydidae. Cryptosporidiosis. Photomicrograph of Cryptosporidium on the gastric mucosa. H&E stain. Courtesy of Michael M. Garner.

Figure 12.50  Hermann’s tortoise, Testudo hermanni. Testudinidae. Sarcocystis infection. Photomicrograph of Sarcocystis in skeletal muscle. H&E stain; Bar = 100 µm. Courtesy of John Roberts.

Parasites and Parasitic Diseases of Reptiles  621

Figure 12.51  Nile crocodile, Crocodylus niloticus. Crocodylidae. Coccidiosis. Photomicrograph of Goussia-like coccidial organisms (arrows) in enterocytes. H&E stain. Bar = 20 µm.

Figure 12.52  Nile crocodile, Crocodylus niloticus. Crocodylidae. Coccidiosis. Photomicrograph of sporozoites (arrows) of a Goussia-like coccidial organism in the spleen. H&E stain; Bar = 20 µm.

Figure 12.53  Nile crocodile, Crocodylus niloticus. Crocodylidae. Coccidiosis. Higher magnification photomicrograph of sporozoites of a Goussia-like coccidial organism within sporocysts in the spleen. H&E stain. Bar = 20 µm. (From Gardiner CH et al. 1986. J Wildl Dis 25:575–577. With permission.)

622  Parasites and Parasitic Diseases of Reptiles

Figure 12.54  Common house gecko, Hemidactylus frenatus. Gekkonidae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Isospora frenatus recovered from the feces. SB=Steida body. Courtesy of Steve Upton.

Figure 12.55  Madagascan giant day gecko, Phelsuma madagascariensis grandis. Gekkonidae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Eimeria brygooi recovered from the feces. (From Upton SJ and Barnard SM. 1987. J Protozool 4:452–454. With permission.)

Figure 12.56  Tree monitor, Varanus prasinus becarri. Varanidae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Eimeria becarri recovered from the feces. (From Upton SJ and Freed PS. 1990. Syst Parasitol 16:181–184. With permission.)

Figure 12.57  Flat-headed snake, Tantilla gracilis. Colubridae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Isospora wilsoni recovered from the feces. SR = sporocyst residuum. (From Upton SJ et al. 1992. Trans Amer Microsc Soc 111:50–60. With permission.)

Parasites and Parasitic Diseases of Reptiles  623

Figure 12.58  Green anole, Anolis carolinensis. Iguanidae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Caryospora ernsti recovered from the feces. OW = oocyst wall; SW = sporocyst wall; SB = Steida body. Courtesy of Steve Upton.

Figure 12.59  Flat-headed snake, Tantilla gracilis. Colubridae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Caryospora gracilis recovered from the feces. SR = sporocyst residuum; SB = Stieda body. Courtesy of Steve Upton.

Figure 12.60  Madagascar hognose snake, Leioheterodon madagascariensis. Colubridae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Caryospora serpentis recovered from the feces. (From Upton SJ et al. 1990. Can J Zool 68:2368–2375. With permission).

Figure 12.61  Madagascar common snake, Madagascarophis colubrinus. Colubridae. Nomarski interference-contrast photomicrograph. Sporulated oocyst of Caryospora madagascariensis recovered from the feces. PG = polar granule; SB = Steida body. (From Upton SJ et al. 1990. Can J Zool 68:2368–2375. With permission.)

624  Parasites and Parasitic Diseases of Reptiles

Figure 12.62  Texas rat snake, Elaphe obsoleta lindheimeri. Colubridae. Cryptosporidiosis. An enlarged stomach is distending the body wall.

Figure 12.63  Bull snake, Pituophis melanoleucus. Colubridae. Cryptosporidiosis. Photomicrograph of a gastric biopsy impression smear of a gastric epithelial cell with Cryptosporidium on the surface (arrows). Wright-Giemsa stain.

Figure 12.64  Unidentified snake. Cryptosporidiosis. Cryptosporidiosis. Photomicrograph of acid-fast stained Cryptosporidium within a fecal specimen, These organisms are not acidfast in histological preparations. Courtesy of Ellis Greiner.

Figure 12.65  Timber rattlesnake, Crotalus horridus. Viperidae. Cryptosporidiosis. Photomicrograph of Cryptosporidium on the gastric mucosal surface. The glandular epithelium has been replaced with fibrous connective tissue. H&E stain. Bar = 50µm.

Parasites and Parasitic Diseases of Reptiles  625

Figure 12.66  Bull snake, Pituophis melanoleucus. Colubridae. Cryptosporidiosis. Photomicrograph of Cryptosporidium within dilated gastric glands. Giemsa stain. Bar = 50 µm.

Figure 12.67  Trans-Pecos rat snake, Bogertophis subocularis. Colubridae. Cryptosporidiosis. Photomicrograph of Cryptosporidium on the mucosal surface of the small intestine. The mucosa is mildly hyperplastic and there is a mild inflammatory cell infiltrate in the lamina propria. H&E stain. Bar = 20 µm.

Figure 12.68  Leopard gecko, Eublepharis macularius. Eublepharidae. Cryptosporidiosis. Emaciated geckoes infected with Cryptosporidium. The tail base is thin (arrow heads) and the intestine (arrows) is thickened. (From Terrell SP et al. 2003. J Zoo Wildl Med 34:69–75. With permission.)

Figure 12.69  Leopard gecko, Eublepharis macularius. Eublepharidae. Cryptosporidiosis. Photomicrograph of Cryptosporidium on mucosal surface of dilated gastric glands. H&E stain. Bar = 50 µm. Courtesy of Scott Terrell.

626  Parasites and Parasitic Diseases of Reptiles

Figure 12.70  Leopard gecko, Eublepharis macularius. Eublepharidae. Cryptosporidiosis. Mucosal epithelial cell proliferation and inflammatory cell infiltrates in the lamina propria of the intestine can be seen. H&E stain. Courtesy of Scott Terrell.

Figure 12.71  Leopard gecko, Eublepharis macularius. Eublepharidae. Cryptosporidiosis. Photomicrograph of Cryptosporidium on enterocytes in the small intestine. Mucosal epithelial cell proliferation can be seen. H&E stain. Bar = 50 µm. Courtesy of Scott Terrell.

Figure 12.72  Green iguana, Iguana iguana. Iguanidae. Cryptosporidiosis. Cystic pedunculated aural-pharyngeal masses are seen. Courtesy of Elizabeth W. Uhl.

Figure 12.73  Green iguana, Iguana iguana. Iguanidae. Cryptosporidiosis. Bilateral tympanic protrusions are seen. Courtesy of Elizabeth W. Uhl.

Parasites and Parasitic Diseases of Reptiles  627

Figures 12.74–12.76  Green iguana, Iguana iguana. Iguanidae. Cryptosporidiosis. Photomicrographs of cryptosporidial organisms along the apical surface of epithelial cells in aural-pharyngeal polyps. There is moderate mucosal proliferation and exocytosis of inflammatory cells. H&E stain.

Figure 12.77  Boa constrictor, Boa constrictor. Boidae. Klossiella infection. Photomicrograph of Klosiella boae in an impression smear of the liver. H&E stain. Courtesy of Sam R. Telford Jr.

628  Parasites and Parasitic Diseases of Reptiles

Figure 12.78  American alligator, Alligator mississippiensis. Alligatoridae. Hemogregarina infection. Photomicrograph of Hemogregarina crocodilinorum (arrows) in red blood cells of a peripheral blood film. Wright-Giemsa stain. Bar = 10 µm.

Figure 12.79  Northern water snake, Nerodia sipedon sipedon. Colubridae. Hemogregarina infection. Photomicrograph of Hemogregarina sp. (arrows) in red blood cells of a peripheral blood film. (From Jacobson ER. 1986. Zoo and Wild Animal Medicine, 2nd edition, Fowler ME (Ed.), Saunders, Philadelphia, PA, 162–181. With permission.)

Figure 12.80  Grass snake, Natrix natrix. Colubridae. Hepatozoon infection. Photomicrograph of schizonts (arrows) of Hepatozoon sp. in the lung. The inflammatory response is minimal. H&E stain.

Parasites and Parasitic Diseases of Reptiles  629

Figure 12.81  Emerald tree boa, Corallus caninus. Boidae. Hepatozoon infection. Photomicrograph of schizonts of Hepatozoon in the liver. There is no inflammatory response. H&E stain.

Figure 12.82  Emerald tree boa, Corallus caninus. Boidae. Hepatozoon infection. Photomicrograph of another area of the liver seen in Figure 12.81. Schizonts of Hepatozoon are within an area of necrosis. An infiltrate of inflammatory cells is also seen. H&E stain.

Figure 12.83  Inland bearded dragon, Pogona vitticeps. Agamidae. Microsporidium infection. Photomicrograph of Microsporidium (arrows) within renal tubules. H&E stain. Bar = 20 µm.

Figure 12.84  Inland bearded dragon, Pogona vitticeps. Agamidae. Microsporidium infection. Photomicrograph of Microsporidium (arrows) within renal tubules. Acid-fast stain. Bar = 20 µm.

630  Parasites and Parasitic Diseases of Reptiles

Figure 12.85  Inland bearded dragon, Pogona vitticeps. Agamidae. Microsporidium infection. Photomicrograph of Microsporidium (arrows) within renal tubules of an inland bearded dragon. Gram stain. Bar = 20 µm.

Figure 12.86  Inland bearded dragon, Pogona vitticeps. Agamidae. Microsporidium infection. Photomicrograph of a semithin section of the liver of an inland bearded dragon with numerous microsporidial organisms (arrows) within an area of necrosis. Toluidine blue stain. Bar = 20 µm.

Figure 12.87  Inland bearded dragon, Pogona vitticeps. Agamidae. Microsporidium infection. Transmission electron photomicrograph of the liver. Developmental stages of Microsporidium are seen: SB = sporoblast; M = meront; S = spore. Uranyl acetate and lead citrate stain. Bar = 5 µm. (From Jacobson ER et al. 1998. J Zoo Wildl Med 29:315–323. With permission.)

Parasites and Parasitic Diseases of Reptiles  631

Figure 12.88  Red-eared slider, Trachemys scripta elegans. Emydidae. Myxozoan infection. Photomicrograph of organisms consistent with a myxozoan within a renal tubule. H&E stain. Bar = 50 µm. Courtesy of John Roberts.

Figure 12.89  Red-eared slider, Trachemys scripta elegans. Emydidae. Myxozoan infection. Photomicrograph of organisms consistent with a myxozoan within a renal tubule. Gram stain. Bar = 50 µm. Courtesy of John Roberts.

Figure 12.90  Yellow-spotted Amazon river turtle, Podocnemis unifilis. Pelomedusidae. Myxozoan infection. Photomicrograph of a renal tubule is dilated and myxozoan spores (arrows) are seen in the lumen. H&E stain. Bar = 50 µm. Courtesy of José Catão-Dias.

Figure 12.91  Yellow-spotted Amazon river turtle, Podocnemis unifilis. Pelomedusidae. Myxozoan infection. Photomicrograph of the kidney at a higher magnification. Organisms identified as Myxidium are seen in the lumen of a tubule and within epithelial cells. H&E stain. Bar = 50 µm. Courtesy of José Catão-Dias.

632  Parasites and Parasitic Diseases of Reptiles

Figure 12.92  Timor python, Python timoriensis. Pythonidae. Photomicrograph of proglottids of Bothridium sp. found in the intestine at necropsy.

Figure 12.93  Unidentified python, Python sp. Pythonidae. Photomicrograph of eggs of Bothridium sp. Bar = 50 µm.

Figure 12.94  Unidentified python, Python sp. Pythonidae. Photomicrograph of egg of Ophiotaenia sp. Bar = 50 µm.

Figure 12.95  False water cobra, Hydronastes gigas. Colubridae. Photomicrograph of a tapeworm egg. Bar = 45 µm.

Parasites and Parasitic Diseases of Reptiles  633

Figure 12.96  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Cysts of a trypanorhynch tapeworm in the gastric wall.

Figure 12.97  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Cysts of a trypanorhynch tapeworm in the peritoneum.

Figure 12.98  Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph of a larval trypanorhynch removed from a cystic lesion in a wild green turtle. Harris hematoxylin and fast green stain. Bar = 500µm. Courtesy of Ellis Greiner.

Figure 12.99  Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph of a larval trypanorhynch in a green turtle surrounded by eosinophilic necrotic debris and a capsule of fibrous connective tissue. H&E stain. Courtesy of Chris Gardiner.

634  Parasites and Parasitic Diseases of Reptiles

Figure 12.100  Island night lizard, Xantusia riversiana. Xantusiidae. Mesocestoides infection. Photomicrograph of Mesocestoides in the liver. H&E stain. Courtesy of Sam R. Telford Jr.

Figure 12.101  Inland bearded dragon, Pogona vitticeps. Agamidae. Mesocestoides infection. Photomicrograph of multiple cross-sections of Mesocestoides in the liver. H&E stain. Bar = 500 µm. Courtesy of Deryck Read.

Figure 12.102  Inland bearded dragon, Pogona vitticeps. Agamidae. Mesocestoides infection. Photomicrograph of Mesocestoides in the liver surrounded by a connective tissue capsule. H&E stain. Bar = 200 µm. Courtesy of Deryck Read.

Figure 12.103  Arizona mountain king snake, Lampropeltis pyromelena. Colubridae. Mesocestoides infection. Photomicrograph of Mesocestoides on the serosal surface of the duodenum and within the pancreas (P). H&E stain. Bar = 500 µm. (From Jacobson ER. 1986. Zoo and Wild Animal Medicine, 2nd edition, Fowler ME (ed), Saunders, Philadelphia, PA, 162–181. With permission.)

Parasites and Parasitic Diseases of Reptiles  635

Figure 12.104  Eastern indigo snake, Drymarchon corais couperi. Colubridae. Renifers are seen in the oral cavity. (From Jacobson ER. 1986. Zoo and Wild Animal Medicine, 2nd edition, Fowler ME (ed), Saunders, Philadelphia, PA, 162–181. With permission.)

Figure 12.105  Mexican garter snake, Thamnophis eques. Colubridae. Renifers are seen in the lung along with purulent material.

Figure 12.106  Eastern indigo snake, Drymarchon corais couperi. Colubridae. Photomicrograph of a renifer attached to the esophageal mucosa. H&E stain.

Figure 12.107  Florida banded water snake, Nerodia fasciata. Colubridae. Photomicrograph of multiple renifer eggs from a lung washing. Bar = 40 µm. (From Jacobson ER. 1986. Zoo and Wild Animal Medicine, 2nd edition, Fowler ME (Ed.), Saunders, Philadelphia, PA, 162–181. With permission.)

636  Parasites and Parasitic Diseases of Reptiles

Figure 12.108  Tropical rat snake, Spilotes pullatus. Colubridae. Styphlodora infection. Photomicrograph of Styphlodora within a collecting duct. H&E stain. Bar = 200 µm.

Figure 12.109  Chicken turtle, Deirochelys reticularia. Emydidae. Spirorchidiasis. Multifocal ulcerations are seen in the plastron.

Figure 12.110  Chicken turtle, Deirochelys reticularia. Emydidae. Spirorchidiasis. Photomicrograph of a spirorchiid egg in a vessel in the dermal bone of the plastron. H&E stain.

Figure 12.111  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Spirorchidiasis. Myriads of spirorchiid trematode eggs in the small intestine are seen as black serpentine tracts.

Parasites and Parasitic Diseases of Reptiles  637

Figure 12.112  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Spirorchidiasis. Black serpentine tracts of spirorchiid eggs in the intestinal mucosa of a loggerhead sea turtle.

Figure 12.113  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Spirorchidiasis. Nodules protruding from the serosal surface of the small intestine were found to contain numerous spirorchiid eggs.

Figure 12.114  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Spirorchidiasis. Photomicrograph of a section of a nodule from the small intestine seen in Figure 12.113. Spirorchiid trematode eggs with a golden brown shell are surrounded by eosinophilic necrotic debris (arrows). H&E stain.

Figure 12.115  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Spirorchidiasis. Higher magnification photomicrograph of Figure 12.114 revealing spirorchiid trematode eggs with a golden brown shell surrounded by eosinophilic necrotic debris. H&E stain.

638  Parasites and Parasitic Diseases of Reptiles

Figure 12.116  Chicken turtle, Deirochelys reticularia. Emydidae. Spirorchidiasis. Photomicrograph of spirorchiid eggs within vessels in the lamina propria of the small intestine. There is an associated inflammatory response that includes macrophages. H&E stain. (From Jacobson ER. 1986. Zoo and Wild Animal Medicine, 2nd edition, Fowler ME (Ed.), Saunders, Philadelphia, PA, 162–181. With permission.)

Figure 12.117  Painted turtle, Chrysemys picta. Emydidae. Spirorchidiasis. Photomicrograph of numerous Spirorchis eggs within the spleen. H&E stain. Courtesy of Francisco Uzal.

Figure 12.118  Painted turtle, Chrysemys picta. Emydidae. Spirorchidiasis. Photomicrograph of microgranulomas containing Spirorchis eggs within the spleen. H&E stain. Courtesy of Francisco Uzal.

Parasites and Parasitic Diseases of Reptiles  639

Figure 12.119  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Spirorchidiasis. Photomicrograph of numerous Neospirorchis eggs in the spleen. Courtesy of Brian Stacy. H&E stain.

Figure 12.120  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Spirorchidiasis. Photomicrograph of Neospirorchis eggs in the meninges of the brain. H&E stain.

Figure 12.121  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of a Haplotrema egg. Bar = 230 µm. Courtesy of Ellis Greiner.

Figure 12.122  Hawksbill sea turtle, Eretmochelys imbricata. Cheloniidae. Photomicrograph of a Haplotrema egg. Bar = 170 µm. Courtesy of Ellis Greiner.

640  Parasites and Parasitic Diseases of Reptiles

Figure 12.123  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of Neospirorchis egg. Bar = 50 µm. Courtesy of Brian Stacy.

Figure 12.124  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of a Carettacola egg. Bar = 90 µm. Courtesy of Ellis Greiner.

Figure 12.125  Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph of a Learedius learedi egg. Bar = 150 µm. Courtesy of Ellis Greiner.

Figure 12.126  Green turtle, Chelonia mydas. Cheloniidae. Photomicrograph of a Learedius learedi egg. Bar = 150 µm. Courtesy of Ellis Greiner.

Parasites and Parasitic Diseases of Reptiles  641

Figure 12.127  Texas indigo snake, Drymarchon corais erebennus. Colubridae. Alaria infection. Photomicrograph of mesocercariae of Alaria (arrows) in the skeletal muscle of the tail. H&E stain.

Figure 12.128  Texas indigo snake, Drymarchon corais erebennus. Alaria infection. Colubridae. Photomicrograph of mesocercariae of Alaria in the skeletal muscle of the tail. H&E stain.

642  Parasites and Parasitic Diseases of Reptiles

Figure 12.129  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Photomicrograph of eggs of Sulcascaris. Bar = 50 µm. Courtesy of Ellis Greiner.

Figure 12.130  American alligator, Alligator mississippiensis. Alligatoridae. Photomicrograph of a Dujardinascaris egg. Bar = 75 µm. Courtesy of Ellis Greiner.

Figure 12.131  Blood python, Python curtus. Pythonidae. Photomicrograph of an ascarid egg. Bar = 50 µm.

Parasites and Parasitic Diseases of Reptiles  643

Figure 12.132  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Anasakiasis. Anasakis embedded in the gastric mucosa. Courtesy of Jorge Oros.

Figure 12.133  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Anasakiasis. Photomicrograph of a cross-section of Anasakis in the gastric wall. Courtesy of Jorge Oros.

Figure 12.134  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Ascariasis. Numerous Sulcascaris are seen in the stomach near the junction with the esophagus.

Figure 12.135  American alligator, Alligator mississippiensis. Alligatoridae. Ascariasis. Numerous Dujardinascaris are embedded in gastric ulcers.

644  Parasites and Parasitic Diseases of Reptiles

Figure 12.136  Burmese python, Python molurus bivittatus. Pythonidae. Ascariasis. Ascarids are embedded in the gastric mucosa.

Figure 12.137  Eastern hognose snake, Heterodon platyrhinos. Colubridae. Ascariasis. Ophidascaris are seen on the gastric serosal surface.

Figure 12.138  Water snake, Nerodia sipedon. Colubridae. Ascariasis. Photomicrograph of Ophidascaris on the serosal surface of the duodenum. H&E stain.

Parasites and Parasitic Diseases of Reptiles  645

Figure 12.139  Red-footed tortoise, Geochelone carbonaria. Testudinidae. Proatractis infection. Photomicrograph of the colon revealing severe necrosis and inflammatory infiltrates with numerous Proatractis (arrows) in the necrotic debris on the surface. H&E stain. Courtesy of Bruce Rideout.

Figure 12.140  Red-footed tortoise, Geochelone carbonaria. Testudinidae. Proatractis infection. Photomicrograph of a cross-section of a female Proatractis within the colon. Larvae (LA) are within the uterus (UT). The intestine (IN) of the nematode is also seen. H&E stain. (From Rideout BA et al. 1987. J Wildl Dis 23:103–108. With permission.)

Figure 12.141  Red-footed tortoise, Geochelone carbonaria. Testudinidae. Proatractis infection. Proatractis is seen within colonic contents. Courtesy of Bruce Rideout.

646  Parasites and Parasitic Diseases of Reptiles

Figure 12.142  Eastern king snake, Lampropeltis getula. Colubridae. Photomicrograph of the anterior end of Kalicephalus.

Figure 12.143  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of a Kalicephalus egg from a fecal sample. Bar = 40 µm .

Figure 12.144  Gila monster, Heloderma suspectum. Helodermatidae. Filariasis. Photomicrograph of the kidney. There is a mild fibrosis and tubular dilatation with numerous microfilariae (arrows) in vessels within the interstitium (arrow).

Figure 12.145  Gila monster, Heloderma suspectum. Helodermatidae. Filariasis. Photomicrograph of microfilariae in an interstitial vessel within the kidney. H&E stain.

Parasites and Parasitic Diseases of Reptiles  647

Figure 12.146  Gila monster, Heloderma suspectum. Helodermatidae. Filariasis. Photomicrograph of cross-sections of filariae in a renal artery. H&E stain.

Figure 12.147  Panther chameleon, Chamaeleo pardalis. Chamaeleonidae. Filariasis. Foyella is seen in the subcutaneous tissues adjacent to the ribs. Courtesy of Brian Stacy.

Figure 12.148  Boa constrictor, Boa constrictor. Boidae. Filariasis. Photomicrograph of a microfilaria of MacDonaldius oschei is seen in a peripheral blood film. Courtesy of Sam R. Telford Jr.

Figure 12.149  Boa constrictor, Boa constrictor. Boidae. Filariasis. Photomicrograph of MacDonaldius oschei is seen within an artery. Silver stain. Courtesy of Sam R. Telford Jr.

648  Parasites and Parasitic Diseases of Reptiles

Figure 12.150  Boa constrictor, Boa constrictor. Boidae. Dracunculiasis. An elevated pustule is seen in the skin.

Figure 12.151  Boa constrictor, Boa constrictor. Boidae. Dracunculiasis. Photomicrograph of cross-sections of an adult and larval (arrow) dracunculid within subcutaneous tissue. H&E stain. (From Jacobson ER et al. 1986. J Amer Vet Med Assoc 189:1133–1134. With permission.)

Figure 112.152  Monacled cobra, Naj naja kaouthia. Boidae. Dracunculiasis. Dracunculid removed from subcutaneous tissues.

Parasites and Parasitic Diseases of Reptiles  649

Figure 12.153  Monitor, Varanus sp. Varanidae. Hastospiculum infection. Photomicrograph of an adult female Hastospiculum varani in the lung. H&E stain. Courtesy of the Armed Forces Institute of Pathology.

Figure 12.154  Monitor, Varanus sp. Varanidae. Hastospiculum infection. Photomicrograph of male Hastospiculum varani in the lung. Sperm (arrow) are seen within a spicule (S). H&E stain. Courtesy of the Armed Forces Institute of Pathology.

Figure 12.155  Monitor, Varanus sp. Varanidae. Hastospiculum infection. Photomicrograph of eggs of Hastospiculum varani surrounded by fibrous connective tissue. H&E stain. Courtesy of the Armed Forces Institute of Pathology.

Figure 12.156  Boa constrictor, Boa constrictor. Boidae. Hastospiculum infection. Photomicrograph of Hastospiculum eggs within granulomatous inflammation extending from the esophagus to the lung. H&E stain.

650  Parasites and Parasitic Diseases of Reptiles

Figure 12.157  Monitor, Varanus sp. Varanidae. Tanqua sp. identified using a flexible endoscope inserted into the stomach. Courtesy of Darryl Heard.

Figure 12.158  Corn snake, Elaphe guttata. Colubridae. Rhabdiasis. Photomicrograph of exudate within air passageways associated with Rhabdias infection. (From Jacobson ER. 1986. Zoo and Wild Animal Medicine, 2nd edition, Fowler ME (Ed.), Saunders, Philadelphia, PA, 162–181. With permission.)

Figure 12.159  Bushmaster, Lachesis muta. Viperidae. Rhabdiasis. Photomicrograph of proliferative pneumonia associated with Rhabdias infection. H&E stain.

Figure 12.160  Bushmaster, Lachesis muta. Viperidae. Rhabdiasis. Photomicrograph of larvae of Rhabdias in the lung surrounded by hypertrophied pneumocytes.

Parasites and Parasitic Diseases of Reptiles  651

Figure 12.161  Corn snake, Elaphe guttata. Colubridae. Photomicrograph of an embryonated egg of Rhabdias in a fecal sample. Bar = 35 µm.

Figure 12.162  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of a Strongyloides egg from a fecal sample. Bar = 35 µm. Courtesy of Ellis Greiner.

Figure 12.163  Gopher tortoise, Gopherus agassizii. Testudinidae. Photomicrograph of an oxyurid egg in a fecal specimen. Bar = 60 µm. Courtesy of Ellis Greiner.

Figure 12.164  Pancake tortoise, Malacocherus tornieri. Testudinidae. Photomicrograph of an oxyurid egg in a fecal sample. Bar = 55 µm. Courtesy of Ellis Greiner.

652  Parasites and Parasitic Diseases of Reptiles

Figure 12.165  Boa constrictor, Boa constrictor. Boidae. Photomicrograph of a Capillaria egg in a fecal sample. Bar = 30 µm. Courtesy of Ellis Greiner.

Figure 12.166  Ball python, Python regius. Pythonidae. Photomicrograph of an egg of Capillaria in a fecal sample. Bar = 30 µm. Courtesy of Ellis Greiner.

Figure 12.167  Mugger crocodile, Crocodylus palustris. Crocodylidae. Paratrichosoma infection. Serpentine tracks of Paratrichosoma are seen in the ventral skin.

Parasites and Parasitic Diseases of Reptiles  653

Figure 12.168  King cobra, Ophiophagus hannah. Elapidae. Photomicrograph of an egg of the acanthocephalan Sphaerechinorhynchus serpenticola. Bar = 40 µm. Courtesy of Brent Nickol.

Figure 12.169  Red-eared slider, Trachemys scripta elegans. Emydidae. Acanthocephalan infection. Photomicrograph of Neoechinorhynchus in the esophagus. Note the thin cuticle (C) and thickened hypodermis (H). H&E stain. Bar = 500 µm. Courtesy of John Roberts.

Figure 12.170  Red-eared slider, Trachemys scripta elegans Emydidae. Acanthocephalan infection. Photomicrograph of Neoechinorhynchus. Note the thin cuticle (C) and thickened hypodermis (H). H&E stain. Courtesy of Chris Gardiner.

Figure 12.171  King cobra, Ophiophagus hannah. Elapidae. Acanthocephalan infection. Photomicrograph of the anterior end of Sphaerechinorhynchus serpenticola adjacent to the capsule of the liver (L). H&E stain

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Figure 12.172  King cobra, Ophiophagus Hannah. Elapidae. Acanthocephalan infection. Photomicrograph of the anterior end of Sphaerechinorhynchus serpenticola recovered from the coelomic cavity. Note the spines on the proboscis. Courtesy of Brent Nickol.

Figure 12.173  Eastern indigo snake, Drymarchon corais couperi. Colubridae. Acanthocephalan infection. Photomicrograph of an acanthocephalan in the wall of the colon. GMS stain.

Figure 12.174  Eastern indigo snake, Drymarchon corais couperi. Colubridae. Acanthocephalan infection. Photomicrograph of an acanthocephalan egg surrounded by an inflammatory response in the wall of the colon. H&E stain.

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Figure 12.175  American alligator, Alligator mississippiensis. Alligatoridae. Pentastomidiasis. Scanning electron photomicrograph of the anterior end of Sebekia mississippiensis. The mouth is surrounded by two compound pairs of hooks. Courtesy of Walter Boyce.

Figure 12.176  Gaboon viper, Bitis gabonica. Viperidae. Pentastomidiasis. Photomicrograph of a sectioned head of Armillifer armillatus. Multiple hooks (arrows) can be seen. H&E stain.

Figure 12.177  Gaboon viper, Bitis gabonica. Viperidae. Pentastomidiasis. Photomicrograph of a cross-section through the mid-body of a female Armillifer armillatus that was removed from the lung. Eggs (arrows) are seen within the uterus, and acidophilic glands (A) are adjacent to the intestine. H&E stain.

Figure 12.178  Gaboon viper, Bitis gabonica. Viperidae. Pentastomidiasis. Photomicrograph of a sclerotized opening (arrow) in the body wall of Armillifer armillatus. This is a unique structure of pentastomes. H&E stain.

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Figure 12.179  Gaboon viper, Bitis gabonica. Viperidae. Pentastomidiasis. Photomicrograph of an egg of Armillifer armillatus obtained from a lung washing. Bar = 60 µm. (From Jacobson ER. 1986. Zoo and Wild Animal Medicine, 2nd edition, Fowler ME (Ed.), Saunders, Philadelphia, PA, 162–181. With permission.)

Figure 12.180  Gaboon viper, Bitis gabonica. Viperidae. Pentastomidiasis. Photomicrograph of an embryonated egg of Armillifer armillatus. Bar = 60 µm. Courtesy of Ellis Greiner.

Figure 12.181  Nile monitor, Varanus niloticus. Viperidae. Pentastomidiasis. Photomicrograph of a larva hatching from a pentastomid egg. Courtesy of Ellis Greiner.

Figure 12.182  American alligator, Alligator mississippiensis. Pentastomidiasis. Alligatoridae. Several Sebekia mississippiensis are seen (arrows) in the lungs. There is mild pulmonary disease.

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Figure 12.183  American alligator, Alligator mississippiensis. Alligatoridae. Pentastomiasis. Sebekia mississippiensis is seen in the lung. While there is compression of adjacent air passageways, no inflammatory response is present. H&E stain.

Figure 12.184  Monitor, Varanus sp. Varanidae. Pentastomiasis. Photomicrograph of a cross section of an unidentified pentastomid in the lung. There is no inflammatory response. H&E stain. Courtesy of the Armed Forces Institute of Pathology.

Figure 12.185  Water moccasin, Agkistrodon contortrix. Viperidae. Pentastomiasis. Porocephalus is seen in the air sac which is located on the dorsal surface of the liver.

Figure 12.186  Gaboon viper, Bitis gabonica. Viperidae. Pentastomiasis. Armillifer armillatus removed from the lung using a flexible endoscope.

Figure 12.187  Gaboon viper, Bitis gabonica. Viperidae. Pentastomiasis. Armillifer armillatus from the lung of a snake that died of pulmonary disease. Caseous exudate can be seen within faveolar spaces. (From Jacobson ER. 1986. Zoo and Wild Animal Medicine, 2nd edition, Fowler ME (Ed.), Saunders, Philadelphia, PA, 162–181. With permission.)

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Figure 12.188  Yellow-belly slider, Trachemys scripta scripta. Emydidae. Leeches are seen on the soft tissue around the head.

Figure 12.189  American alligator, Alligator mississippiensis. Alligatoridae. Numerous leeches are seen in the oral cavity. Courtesy of Steve Stiegler.

Figure 12.190  Loggerhead sea turtle, Caretta caretta. Cheloniidae. The marine leech Ozobranchus margoi is identified by five pairs of branchiae (gills).

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Figure 12.191  Green turtle, Chelonia mydas. Cheloniidae. Leeches are present on the conjunctiva.

Figure 12.192  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Leeches and mats of leech eggs (E) are seen on the soft tissue adjacent to a limb. Courtesy of Brian Stacy.

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Figure 12.193  Loggerhead sea turtle, Caretta caretta. Cheloniidae. A clinically healthy wild turtle with several barnacles on the head.

Figure 12.194  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Stranded cachectic turtle with a heavy barnacle load on the carapace. Courtesy of Brian Stacy.

Figure 12.195  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Stranded cachectic turtle with numerous barnacles on the head.

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Figure 12.196  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Stranded cachectic turtle with numerous barnacles on the head. Courtesy of Brian Stacy.

Figure 12.197  Loggerhead sea turtle, Caretta caretta. Cheloniidae. Stranded cachectic turtle with numerous small barnacles on the neck and limbs.

Figure 12.198  Green turtle, Chelonia mydas. Cheloniidae. Stephanolepas embedded (arrows) in the subcutaneous tissues of a foreflipper.

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Figure 12.199  Gopher tortoise, Gopherus polyphemus. Testudinidae. Acariasis. Amblyomma tuberculatum is seen embedded in the soft epidermal tissue at the base of a forelimb.

Figure 12.200  Spider tortoise, Pixys arachnoides. Testudinidae. Acariasis. Amblyomma chabaudi on soft tissue adjacent to a hindlimb. Courtesy of Michael Burridge.

Figure 12.201  Ornithodoros collected in the vicinity of a gopher tortoise burrow. Courtesy of Ellis Greiner.

Figure 12.202  Leopard tortoise, Geochelone pardalis. Testudinidae. Acariasis. Multiple Amblyomma marmoreum embedded in the cervical skin.

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Figure 12.203  Dipsochelys dussumieri Aldabra tortoise. Testudinidae. Acariasis. Multiple Amblyomma marmoreum embedded in the soft tissue near a hindlimb. Courtesy of Michael Burridge.

Figure 12.204  Nile monitor, Varanus niloticus. Varanidae. Acariasis. Amblyomma embedded in the skin at the base of the tail, adjacent to the cloaca.

Figure 12.205  Florida banded water snake, Nerodia sipedon fasciata. Colubridae. Acariasis. Numerous Proctolaelaps pygmaeus cover the head. Some opacity in the spectacle is seen along with thickening of the tissues at the fornix between the spectacle and the periocular scales.

Figure 12.206  Snake mite, Ophionyssus natricis. Photomicrograph of Ophionyssus natricis removed from a rat snake. A single egg can be seen within the mite.

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Figure 12.207  Carpet python, Morelia spilota. Pythonidae. Acariasis. Mites (arrows) can be seen in the fornix where the spectacle and conjunctiva join. Also, note that the conjunctiva is swollen. Courtesy of Stephen Barten.

Figure 12.208  Carpet python, Morelia spilota. Pythonidae. Acariasis. Mites can be seen under gular scales and within the midline gular fold (arrows). Courtesy of Stephen Barten.

Figure 12.209  Russell’s viper, Vipera russellii. Viperidae. Acariasis. Photomicrograph of Ophionyssus natricis attached to the skin. H&E stain. Courtesy of John Roberts.

Figure 12.210  Green iguana, Iguana iguana. Iguanidae. Acariasis. Hyperpigmented, patchy skin necrosis caused by pterygosmid mites in the genus Hirstiella. (From Jacobson ER. 2003. Biology, Husbandry, and Medicine of the Green Iguana. Krieger Publishing Company, Malabar, FL. With permission.)

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Figure 12.211  Cayman Island ground iguana, Cyclura nubila. Iguanidae. Acariasis. Cyclurobia is present at the base of the dorsal spines. Courtesy of Brian Stacy.

Figure 12.212  Common agama, Agama agama. Agamidae. Acariasis. Trombiculid mites are seen on the tympanic membrane.

13 Isolation of Pathogens Francesco C. Origgi and Jean A. Paré

Contents

13.1 General Comments

13.1 General Comments................................................ 667 13.2 Sample Handling................................................... 667 13.2.1 Sample Collection...................................... 667 13.2.2 Sample Conservation and Shipping......... 668 13.2.3 Sample Reception...................................... 669 13.3 The Approach to Microorganism Isolation ......... 669 13.3.1 Viral Isolation............................................ 669 13.3.2 Bacterial Isolation.......................................670 13.3.3 Fungal Isolation..........................................671 Acknowledgments............................................................ 672 References......................................................................... 672

The isolation of pathogens is not molecular based but rather relies on the recovery of the whole live and replication-competent microorganism(s) associated with a specific disease. Conventionally, pathogens include bacteria, viruses, fungi, and parasites. While molecular tools (see Chapter 7) have revolutionized the way a diagnostician can identify a specific pathogen, there is still a need to isolate pathogens. Pathogens (and their antigens) are still a very important component of many serodiagnostic assays (see Chapter 8). Determining the most important antimicrobial drug to use in bacterial infections is best done when the organism has been isolated. For the most part, fungi are best identified with an isolate rather than making an identification in tissue section. Even when isolated, there are relatively few mycologists who have experience with identification of fungi from reptiles. In this chapter, techniques used in isolating various pathogens in reptiles will be reviewed.

13.2 Sample Handling 13.2.1 Sample Collection The quality of the collected sample and appropriateness of the procedure adopted to obtain it influence the entire isolation process. Consequently, the development of a set of standard operative procedures (SOP) could prove helpful for consistency and would be of use when training new personnel. Fundamental considerations must be kept in mind when collecting specimens. Koneman et al. (1997a) pointed out the critical steps for appropriate sampling leading to a successful isolation. First, the sample must be collected from the specific site of infection, and maximum care needs to be taken Infectious Diseases and Pathology of Reptiles  667

668  Isolation of Pathogens

to avoid any possible contamination from tissue, organs, or secretions close to the site of interest. For example, when sampling highly contaminated tissues, such as those collected during necropsy, it is ideal to first cauterize the organ surface with a flamed or heated metal prior to inserting a sterile swab through an incision made in the tissue with sterile scissors or a scalpel blade. Instruments are commonly dipped in alcohol and flamed using an alcohol lamp or propane burner. Sometimes it is difficult to determine if the alcohol is burning on the instrument and the diagnostician may dip the instrument back in alcohol for re-flaming. This may set the entire alcohol container on fire. Second, the time of collection is critical for successful isolation. Viral isolation is strongly influenced by the time of sampling. In human herpesvirus infection, for example, viral shedding is not constant in infected individuals (Sacks et al., 2004). There is evidence suggesting that a similar phenomenon occurs in tortoises infected with tortoise herpesvirus (Origgi et al., 2004). Very few reptile infectious diseases have been investigated experimentally and only limited information is available on pathogenesis so that routes of entry, replication sites, shedding intervals, and other disease parameters remain largely unknown. Third, the quantity of specimen collected will be proportional to the number of diagnostic tests (or replicates) that it will be possible to set up for the investigation and eventually to the probability of a successful isolation. Collection of a very limited amount of material could result in a sample with insufficient bacterial load or viral titer for culture. It is therefore important to try to collect as much sample as reasonably possible. Fourth, it is critical to use the appropriate instrumentation, storage devices and media to collect and preserve specimens. Specific containers for bacterial, viral and fungal samples are commercially available. As a general rule, different types of samples require different types of sampling devices. For biopsies, surgical instruments are required; for blood culture for bacteria, special sterile bottles containing growth medium broth are required (Figures 13.1–13.2). Swabs are used for collection of different types of specimens such as exudates, tissues, and fluids (Figure 13.3). Different types of swabs might be needed for different types of pathogens. For example, calcium alginate swabs may be inappropriate for human herpesvirus 1 sampling (Crane et al., 1980), whereas they have been suggested for sampling tortoises with suspected Mycoplasma spp. infection. For sampling tortoises for herpesvirus collection, dacron polyester-tipped swabs are recommended (Origgi and Jacobson, 2000). If infection with anaerobic bacteria is suspected, aspiration of the sample using a syringe and a needle is often the method of choice, with the sample being immediately added to a liquid media anaerobic collection system. Care needs to be taken to avoid contact of the specimen with air (Koneman et al., 1997a). Finally, when drug therapy is needed, samples should be obtained from the animal prior to the beginning of the

treatment. If the animal is already on antimicrobial drug therapy, trypticase soy broth with resins should be used. The resins will absorb antimicrobials in blood, thus increasing the likelihood of recovering bacteria (Figure 13.2). After collection, the sample needs to be labeled appropriately with permanent markers so that the risk for confusion is minimized. This cannot be overstated: clear permanent labeling is needed. The author has received samples of tissue for pathogen identification in which the writing on the container or tube is illegible. In many instances, samples have been received in which the labeling has started to come off. The investigator should never have to guess the correct spelling or numbering of a label on a sample.

13.2.2 Sample Conservation and Shipping Collected samples are often shipped to a diagnostic laboratory. Precautions need to be taken to ensure that the sample will be preserved as well as possible until delivery. Overnight reliable delivery companies are needed. Specific storage and transport media for viruses, bacteria, and fungi are available. Bacteria and fungi are commonly collected with swabs maintained in an appropriate storage medium. For viral isolation, several media are available. Sterile phosphate buffer saline (PBS) or saline solution (0.9% NaCl) can be used. For best results, samples for virus isolation should be kept refrigerated until delivery to the diagnostic laboratory, but if delayed it is better to freeze the samples at −80°C for long-term storage. The viral titer tends to decrease over time when the sample is refrigerated at +4°C. In general, maintaining the appropriate temperature is one of the most difficult obstacles to preservation. Exposure of the sample to extreme temperatures can compromise its viability. For refrigerated shipping, more precautions need to be taken. The shipping destination, the transit time that the package is likely to be in the mail or in the hands of the shippers, and the size of the samples are all factors to be considered. Ice packs or dry ice are commonly used for shipping material. When sending frozen samples, dry ice is preferred over ice packs to ensure that frozen samples remain frozen. Make sure to use a well-insulated box such as a Styrofoam box shipped within a cardboard box. Specialized Styrofoam boxes for diagnostic samples are commercially available (Figure 13.4). When the latter is used, a special warning needs to be placed on the shipping container. When shipping hazardous material, such as infected samples, extra safety precautions are required. Never use glass containers. Always use resistant plastic containers that can be sealed. The containers with the samples should then be placed in another container to prevent leaking. Never ship blood or other potentially infectious material in simple paper envelopes. Potentially zoonotic material needs to be shipped in specific containers and handled by specialized personnel. Finally, it is always best to call the diagnostic laboratory to

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alert them before a sample is shipped. Always use an overnight delivery service.

13.2.3 Sample Reception Upon reception, package integrity needs to be checked immediately. Many samples arrive in damaged containers or the tissues have not been properly refrigerated (Figures 13.5–13.6). Appropriate temperature and container conditions need to be assessed. The shipping facility needs to be notified immediately about any damage to the container or samples. Samples may have questionable value if sent using an overnight delivery company but not delivered until several days later.

13.3 The Approach to Microorganism Isolation In this age of genomics and proteomics, demonstration of the presence of a microorganism in a specimen can be performed with one or more molecular-based techniques (see Chapter 7). Such modern techniques have shown the limits of traditional culture-based isolation approaches (Kroes et al., 1999; Ward et al., 1990), and have revealed the existence of a great number of microorganisms that were totally unknown or unsuspected before (Clarridge, 2004). However, a traditional culture-based pathogen isolation approach is still the only method that allows recovery of whole microorganisms for further study. Here we review the traditional culturebased isolation techniques that are pertinent to the isolation of reptile pathogens. For molecular-based approaches, see Chapter 7.

13.3.1 Viral Isolation Viral isolation requires the use of cell cultures or embryonated eggs because viruses require live cells to replicate. Cells are typically grown in plastic flasks and are checked each day using an inverted microscope (Figure 13.7). Cell cultures may be primary cell cultures, cell lines, or established cell lines. Primary cell cultures are derived from fresh tissues that are treated with a proteolytic enzyme such as trypsin. The cells are then collected and grown in a sterile flask with a cell culture medium (enriched with fetal bovine serum [FBS]) (Figure 13.8). Antibiotics and antimycotics are added to the medium to prevent the growth of contaminating bacteria and fungi. A primary cell culture can be maintained in vitro for a limited time. A subcultured primary cell culture is defined as a cell line. A cell line that has been passed more than 70 times is considered an established cell line (Koneman et al., 1997b). Established cell lines are able to grow indefinitely in conventional cell culture medium. A large number of established cell lines are available through the American Type Cell Collection (ATCC, Rockville, MD, U.S.A.) and other national or international repositories. Among the established cell lines, Terrapene

heart cells (TH-1) (ATCC-CCL 50 Sub-line B1), viper heart cells (VH-2) (ATCC-CCL 140), and African green monkey kidney cells (VERO) (ATTC-CLL 81) have been extensively used for culturing several reptile viruses such as tortoise herpesvirus (Origgi et al., 2001), iridovirus (Marschang et al., 1999), boid retroviruses (Jacobson et al., 2001), snake parvovirus (Farkas et al., 2004), ophidian reovirus (Lamirande et al., 1999), and ophidian paramyxovirus (Mayr et al., 2000) (Figures 13.9–13.15). In general, different cell lines may require different growth and maintenance conditions, such as cell culture media, FBS concentration, temperature, and buffering systems, which might need to be optimized for each cell system. ATCC makes most of the cell culture conditions for each of the cell lines available through their website: http://www.atcc.org. When cells are not required for virus isolation or growth, they can be frozen. A reservoir of several aliquots of each of the cell lines used in the laboratory should be kept stored in liquid nitrogen for future needs. A freezing medium composed of 90% filtered FBS and 10% DMSO is used by the author. Cells are first kept for 2 days at −80°C and then they are stored in liquid nitrogen. Samples collected for viral isolation generally require further processing before inoculation onto cell culture. Sample storage before inoculation onto the cell cultures may influence the outcome of the isolation attempt. Samples may be freshly collected or stored either at +4°C (short term, for a few days) or at −80°C (long term). Viral titers tend to decrease with storage, especially if stored inappropriately, such as at −20°C in a no-frost freezer. The systematic defrosting that occurs in these appliances could compromise the viability of the viruses contained in the sample. Antibiotic- and antimycotic-enriched cell culture media can be used for storage to preserve the samples and to minimize the growth of contaminants. When these media are not available, phosphate buffered saline (PBS) or saline (0.9%) can be used. Aliquots of cell medium for use as storage medium can be kept in the clinic refrigerator or freezer (for long-term storage) ready to be used for samples that are routinely collected. Swabs should be premoistened with the selected storage medium before sample collection, and then placed back in a tube with the tip immersed in the same medium. In general, it is advisable to avoid too much medium because viral titer of the sample will be diluted, thereby compromising the ability to isolate the virus. For non-cell-associated viruses, swabs can either be treated with a few rapid cycles of freezing and thawing or briefly sonified in order to disrupt the cells or the cell material that is attached to the swab tip and free intracellular virions (Schat and Purchase, 1989). These are general and standard procedures, but we remind the reader that the protocol can be adapted to different viruses on an ad hoc basis. For example, R. Marschang (personal communication) reported problems in recovering herpesviruses when the freezing and thawing procedure was adopted. The swab is then centrifuged at low speed and the supernatant is collected. The same procedure is adopted when the

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sample consists of a nasal, tracheal, or oral wash. The supernatant may be either directly inoculated onto the cells or filtered with a 0.22 or 0.45 µm syringe filter. Filtering can result in a moderate to great loss of titer, and it is not recommended when a low titer is suspected. Collection of the samples using a sterile procedure becomes extremely critical in these cases because it could help eliminate the need for filtering. It is crucial to equilibrate the filter with tissue culture medium before filtering the sample. The FBS contained in the medium will serve to equilibrate the filter and help prevent the proteinmediated trapping of viral particles. Additionally, it is important to consider that the filter could block both large viruses and viruses that are bound to cell debris. If a cell-associated virus is suspected, both freezing and thawing and sonification are inappropriate, and there is no centrifugation step. When tissues are collected for viral isolation (biopsies, tissue from necropsies) they need to be first homogenized before being processed as described for swabs. The cellular pellet is generally separated out by low-speed centrifugation and the supernatant can be either directly inoculated or filtered and then inoculated onto the cell cultures. For cell-associated virus isolation, the viral-containing infected cells are allowed to settle on the cell monolayer. The volume of the inoculum may vary. There needs be sufficient volume to cover the monolayer, but not too much to significantly dilute the viral titer. A 1/10 dilution in culture medium is common, but it can be modified. The inoculum is left in contact with the cell monolayer for a limited amount of time, which can vary from 1 to 2 hours (for cell-free virus only). This time is generally long enough for the viral particles to attach to the cell membrane of the target cells. Precaution is to be taken to avoid drying of the cell monolayer. The early removal of the inoculum helps to minimize the damage to the cell culture by toxic components in the inoculum and the risk of bacterial or fungal growth. Fresh medium is added to the culture and the flask is then placed in the incubator at an appropriate temperature. If the inoculum is highly contaminated, the monolayer is washed two or three times with culture medium first, and up to 5 times the normal amount of antibiotics and antimycotics are added to the cell medium (Schat and Purchase, 1989). Optimization, and customization of these protocols are very common. Besides the temperature requirements for each specific cell line as mentioned above, it is critical to also consider the temperature requirements for all the suspected or cultivated viruses, which can differ. Experience is very important and consulting with personnel with extensive training and experience in cell culture and pathogen isolation is strongly advised. Cultures need to be checked daily to assess the cell monolayer conditions. Virus infection of cells (both in vivo and in vitro) is complex and may result in morphological and biochemical changes or death in infected cells. These changes are collectively known as cytopathic effect (CPE). While the variety of CPE is somewhat limited, it is not pos-

sible to unequivocally associate a CPE pattern to a specific virus. Additionally, not all viruses cause CPE. Common CPE observed in cell cultures infected with reptile viruses include cell rounding and cell lysis (herpesvirus and iridovirus) (Marschang et al., 1997, 1999) (Figures 13.10–13.12), syncytia (boid retrovirus, ophidian reovirus, and ophidian paramyxovirus) (Schumacher et al., 1994; Lamirande et al., 1999; Mayr et al., 2000) (Figure 13.13); giant cells (Mayr et al., 2000) (green turtle LET herpesvirus and ophidian paramyxovirus) (Figures 9.11, 9.95); vacuolization (iridoviruses and boid retrovirus) (Marschang et al., 1998; Schumacher et al., 1994); formation of intranuclear with herpesvirus (Origgi et al., 2004) (Figure 13.12) and adenovirus (Jacobson et al., 1985) (Figure 13.14); and intracytoplasmic inclusions with IBD (inclusion body disease) (Schumacher et al., 1994) (Figure 13.15). Signs of cell toxicity may be confused with CPE. The appearance of CPE may be delayed, and blind passages may be needed to recover the virus. Reptile caliciviruses may require six or more passages before CPE are seen (Matson et al., 1996). Furthermore, changing the cell culture medium too often can influence the appearance and the development of CPE. Fresh medium would enhance the growth of the healthy cells and reduce the viral titer of the supernatant (non-cellassociated virus), thereby affecting the amount of CPE in the cultured cells. The daily collection of cell culture medium from an infected culture from day 0 (time of inoculation) can be tested using the qualitative or quantitative molecular detection approach described in Chapter 7 (Southern/northern blot, qualitative PCR/RT-PCR or real-time PCR/RT-PCR), specific for the suspected virus. This may determine if active replication of the virus is occurring long before the appearance of CPE. Positive and negative staining transmission electron microscopy also can be used to determine the presence of a virus in cell culture (see Chapter 6). Finally, when isolation is successful, the virus can then be expanded and partially or fully purified for further study and identification (see Chapter 7). The isolated virus may be used in transmission studies to determine its pathogenicity, to produce antigen for serological assay development, for polyclonal or monoclonal antibody production, and for genome sequencing studies.

13.3.2 Bacterial Isolation A series of considerations need to be kept in mind when a sample is processed for bacterial isolation. Factors such as the selection of the appropriate primary culture medium and the culture microenvironment (temperature, humidity, CO2/O2%) are critical. A decision then needs to be made as to which of the isolates recovered on the primary culture medium are clinically relevant and need to be further characterized. Several hundred types of primary culture media are available on the market and many others can be prepared in-house, offering a wide variety of media for the isolation of many different organisms. Agar plates are commonly used.

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Enrichment broths may be used for body fluids (Koneman et al., 1997a) such as blood (Figure 13.1) or certain samples prior to inoculation on solid substrates. For instance, as previously mentioned, if there is an attempt to isolate bacteria from an animal receiving antimicrobial therapy, an enrichment broth with resins (Figure 13.2) would be best to use. The media can be either selective or nonselective, and the choice depends on the type of specimen and on the isolation approach that has been adopted. Nonselective media are used when a full representation of the bacterial population present in the collected sample is needed. Selective media are used when it is necessary to give a selective advantage to one or more bacterial species that would be difficult to recover on conventional nonselective media because of fastidious growth requirements or because of overgrowth of contaminating organisms. For example, for Salmonella spp. isolation from stool samples, a series of selective media are used to allow the enrichment of Salmonella spp, virtually preventing the growth of all the other bacteria present in the sample. Growth of other microorganisms is suppressed when media selective for a specific organism are used. The choice of the culture microenvironment (temperature, humidity, CO2/O2%) is also very critical because temperature, humidity, and the presence of oxygen are additional factors that will affect the isolation of bacteria. For the most part, the ideal culture requirements have not been established for the array of bacterial pathogens present in reptiles. Most bacteria grow at humidity of 70% or more, and culture media tend to deteriorate if the humidity is low (Koneman et al., 1997a). The presence or absence of oxygen is required for aerobic and anaerobic bacteria, respectively. Sealed oxygen-free systems are available for anaerobic bacteria. Temperature strongly influences bacterial growth, and multiple incubators set at different temperatures can maximize the chance of recovery of poorly characterized microorganisms whose optimal temperature requirements are not well established. This is particularly important when samples are collected from nonmammal subjects such as reptiles, whose microbial ecology is yet to be defined. Many bacteria from reptiles do not key out using the schemes developed for mammals, and identification of new microorganisms will continue for many years to come. Among the media used for bacterial isolation, blood agar (BA) (Figure 13.16), brain heart infusion agar (BHI), and MacConkey agar (MA) (Figures 13.17–13.18) are the most popular. While BA and BHI are two nonselective media, MA is a selective medium that inhibits the growth of Gram-positive bacteria. For isolation of Salmonella, several different selective and differential agars are commercially available. One is Hextoen enteric agar (Figure 13.19). Organisms such as Salmonella, which reduce sulfur to hydrogen sulfide, will produce black colonies or blue-green colonies with a black center.  Conventionally, the sample is inoculated onto agar plates and then incubated at a specific temperature under aerobic or anaerobic conditions. The agar plates should be checked

every 24 hours and maintained in culture for no less than 72 hours for aerobes before being discarded. For anaerobes it is recommended that the plates first be examined 48 hours after inoculation, but they must be kept in culture for no less than 7 days (Donahue, 1990). Fastidious organisms such as Brucella spp. require longer incubation times measured in weeks (Nicoletti, 1990) or with Mycobacterium spp., months (Thoen, 1990). However, there are rapid-growing Mycobacteria in reptiles (see Chapter 10) that grow rather quickly. Slow growth is also a feature of Mycoplasma agassizii, a well-known pathogen of tortoises (Brown et al., 1994; Brown et al., 1999; Brown MB et al., 2001), whose primary isolation may require up to 6 weeks. In contrast, Mycoplasma alligatoris, a pathogen of the American alligator (Alligator mississippiensis), is a rapid grower (Brown DR et al., 2001). Once the bacteria are grown, a preliminary identification of the bacterial population is performed. Representative colonies are lifted, fixed on a glass slide, stained with Gram stain (Carter, 1990a) and examined using light microscopy. This procedure allows one to distinguish Gram positive (G+) from negative (G−) bacteria and to determine if they are bacilli or cocci. Each of the identified colonies is then transferred to a separate plate where they are cultured as a pure isolate (Figure 13.16). This is required to proceed to the final genus and species identification. Several biochemical test-based kits for bacterial identification are commonly used for the determination of the bacterial genus. For speciation, additional tests, such as serum agglutination for Salmonella spp., may be required (Carter and Chengappa, 1990). Sometimes conventional biochemical methods cannot provide an unequivocal and unambiguous identification of the bacterium recovered; this is more frequent for bacteria that are not commonly considered human pathogens (Clarridge, 2004). In the near future, molecular tests (see Chapter 7) will more than likely become the primary method for the identification of novel microorganisms. The final and unequivocal identification of one or more bacteria could be considered the end of the isolation procedure, but this information always needs to be interpreted in the context of other diagnostic data and procedures. The identification of a pathogen could in fact be misleading in itself if not supported by complementary diagnostic tests. Histological observation of organisms within lesions that are consistent with the organism that was cultured from the same sample supports a causative relationship: if histological lesions are absent or organisms are not seen within affected tissue sections, then isolation of the organism may represent an incidental finding.

13.3.3 Fungal Isolation The fungal microbiota of the reptile integument is rich and varied (Paré et al., 2003). Considerations previously discussed in the bacterial isolation section are also appropriate for fungal agents. Fungal isolation can be achieved on different solid

672  Isolation of Pathogens

media. Fast-growing, heavily sporulating saprophytes, such as Mucor and Aspergillus species, can rapidly overgrow the medium and mask slower growing fungi, such as the Chrysosporium anamorph of Nannizziopsis vriesii (CANV) (Paré et al., 2003). Therefore it is prudent to inoculate a nonselective medium and a selective one. Sabouraud’s dextrose (2%) agar, a nonselective medium (Figure 13.20), is commonly used (Carter, 1990b) even if it has been described as more suitable for subculturing fungi recovered on enriched medium (Koneman et al., 1997c). A blood agar and brain heart infusion agar with 1% blood can also be used (Carter, 1990b). Inhibitory mold agar, or Sabhi agar could be used when the recovery of some fastidious or slow-growing fungi is needed (Koneman et al., 1997c). Mycosel or Mycobiotic agars are particularly useful to isolate keratinophilic fungi (Figure 13.20). These agars contain cycloheximide, a compound that inhibits or severely restricts the growth of most saprophytic contaminants and also contain an antibiotic to prevent bacterial growth. Other selective media enriched with a combination of antibiotics may also be used. If bacterial contamination occurs in spite of the presence of antibiotics in the agar, the sample may be removed, dipped into a wide-spectrum antibiotic mix, and reinoculated onto a fresh agar. Sabouraud and Mycosel agars inoculated with reptile material are best incubated at 25 to 28°C. Incubation at 37°C may be helpful in speciating fungi with limited thermotolerance. Plates should be checked daily, but fungi can be very slow growing. The CANV may take 5 to 10 days to grow, and other fungi may be even slower: the agent of epizootic lymphangitis, Histoplasma farciminosum, can take up to 8 weeks to grow (Carter, 1990b). Commonly, more than one fungal agent will grow from reptile material, often differing in the morphology of the colony and in the coloration of the reverse side of the colony (Figure 11.1). Each fungal isolate can be subcultured by cutting out a small cube of cultured agar and placing it upside down on a Sabouraud’s dextrose (2%) agar plate for propagation. A portion of the mold can be collected and mounted in a drop of lactophenol blue stain on a microscope slide. Observation of the mold under light microscopy allows visualization of the vegetative thallus and the conidia, which are useful in attempting preliminary identification of the organism. Speciation of fungi is challenging and can prove very difficult. We strongly suggest consulting a mycology laboratory with personnel whose expertise extends beyond that of fungi traditionally considered important in human medicine. Sequence-based identification is rapidly gaining ground, and molecular-based identification should be considered whenever possible. As discussed for bacterial isolation, mere isolation of a fungus from a lesion is not clinically relevant unless morphologically consistent fungal elements are demonstrated in tissue sections. Clinicians need to exercise sound judgment when interpreting fungal culture results. Isolation of a fungus in pure culture from different lesions, or repeated isolation of a fungus from identical lesions, provides some evidence of causality, but causality cannot be estab-

lished without histopathology. Finally, fungal isolates with proven pathogenicity for reptiles should be preserved and forwarded to recognized depositories so they can be studied further. The main U.S. collection is the American Type Culture Collection ATCC (http://www.atcc.org/). The University of Alberta Microfungus Collection and Herbarium (UAMH) (http://www.devonian.ualberta.ca/uamh/) holds many reptile fungal isolates. The Centraalbureau voor Schimmelcultures (CBS), an institute of the Royal Netherlands Academy of Arts and Sciences (KNAW) is situated in Utrecht (http://www. cbs.knaw.nl/databases/). The web site http://wdcm.nig.ac.jp/ lists world culture collections that are members of the World Federation of Culture Collections.

Acknowledgments The authors thank Rachel Marschang for reviewing this chapter and Mark Hoffenberg for photographic support.

References Brown MB, Schumacher IM, Klein PA, Harris K, Correll T, and Jacobson ER. 1994. Mycoplasma agassizii causes upper respiratory tract disease in the desert tortoise. Infect Immun 62:4580–4586. Brown MB, McLaughlin GS, Klein PA, Crenshaw BC, Schumacher IM, Brown DR, and Jacobson ER. 1999. Upper respiratory tract disease in the gopher tortoise is caused by Mycoplasma agassizii. J Clin Microbiol 37:2262–2269. Brown DR, Farley JM, Zacher LA, Carlton JM, Clippinger TL, Tully JG, and Brown MB. 2001. Mycoplasma alligatoris sp. nov., from American alligators. Int J Syst Evol Microbiol 51:419–24. Brown MB, Brown DR, Klein PA, McLaughlin GS, Schumacher IM, Jacobson ER, Adams HP, Tully JG. 2001. Mycoplasma agassizii sp. nov., isolated from the upper respiratory tract of the desert tortoise (Gopherus agassizii) and the gopher tortoise (Gopherus polyphemus). Int J Syst Evol Microbiol 51:413–418. Carter GR. 1990a. Appendix A. Staining procedures, in Diagnostic Procedure in Veterinary Bacteriology and Mycology, Carter GR and Cole JR Jr (Eds.), Academic Press, San Diego, CA, 519–528. Carter GR. 1990b. Mycology: introduction, in Diagnostic Procedure in Veterinary Bacteriology and Mycology, Carter GR and Cole JR Jr (Eds.), Academic Press, San Diego, CA, 371–379. Carter GR and Chengappa MM. 1990. Enterobacteria, in Diagnostic Procedure in Veterinary Bacteriology and Mycology, Carter GR and Cole JR Jr (Eds.), Academic Press, San Diego, CA, 371–379.  Clarridge JE 3rd. 2004. Impact of 16S rRNA gene sequence analysis for identification of bacteria on clinical microbiology and infectious diseases. Clin Microbiol Rev 17:840–862. Crane LR, Gutterman PA, Chapel T, and Lerner AM. 1980. Incubation of swab materials with herpes simplex virus. J Infect Dis 141:531.

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Donahue J. M. 1990. Nonsporeforming anaerobic bacteria, in Diagnostic Procedure in Veterinary Bacteriology and Mycology, Carter GR and Cole JR Jr (Eds.), Academic Press, San Diego, CA, 177–200. Farkas SL, Zadori Z, Benko M, Essbauer S, Harrach B, and Tijssen P. 2004. A parvovirus isolated from royal python (Python regius) is a member of the genus Dependovirus. J Gen Virol 85:555–61. Jacobson ER, Gaskin JM, and Gardiner CH. 1985. Adenoviruslike infection in a boa constrictor. J Amer Vet Med Assoc 187:1226–1227. Jacobson, ER, Oros J, Tucker S, Pollock D, Kelley KL, Munn RJ, Lock BA, Mergia A, and Yamamoto JK. 2001. Partial characterization of retroviruses from boid snakes with inclusion body disease. Amer J Vet Res 62:217–224. Koneman EW, Allen SD, Janda WM, Schreckemberger PC, and Win WC Jr. 1997a. Introduction to microbiology, Part I: The role of the microbiology laboratory in the diagnosis of infectious diseases: Guidelines to practice and management, in Color Atlas and Textbook of Diagnostic Microbiology, Lippincott, Philadelphia, PA, 69–80. Koneman E. W., S. D. Allen, W. M. Janda, P.C. Schreckemberger, and W.C. Win Jr. 1997b. Diagnosis of infections caused by viruses, chlamydia, rickettsia and related organisms, in Color Atlas and Textbook of Diagnostic Microbiology, Lippincott, Philadelphia, PA, 1177–1293. Koneman E. W., S. D. Allen, W. M. Janda, P.C. Schreckemberger, and W.C. Win Jr. 1997c. Mycology, in Color Atlas and Textbook of Diagnostic Microbiology, Lippincott, Philadelphia, PA.983– 1069. K roes I, Lepp PW, and Relman DA. 1999. Bacterial diversity within the human subgingival crevice. Proc Natl Acad Sci USA 96:14547–14552. Lamirande EW, Nichols DK, Owens JW, Gaskin JM, and Jacobson ER. 1999. Isolation and experimental transmission of a reovirus pathogenic in rat snakes (Elaphe species). Virus Res 63:135–141. Marschang, RE, Gravendyck M, and Kaleta EF. 1997. Investigation into virus isolation and the treatment of viral stomatitis in T. hermanni and T. graeca. J Vet Med Series B 44:385– 394. Marschang RE, Posthaus H, Gravendyck M, Kaleta EF, and Bacciarini LN. 1998. Isolation of viruses from land tortoises in Switzerland, in Proceedings of the American Association of Zoo Veterinarians and American Association of Wildlife Veterinarians Joint Conference, Omaha, NE, 281– 284. Marschang RE, Becher P, Posthaus H, Wild P, Thiel HJ, MullerDoblies U, Kalet EF, and Bacciarini LN. 1999. Isolation and characterization of an iridovirus from Hermann’s tortoises (Testudo hermanni). Arch Virol 144:1909–1922.

Matson DO, Berke T, Dinulos MB, Poet E, Zhong WM, Dai XM, Jiang X, Golding B, and Smith AW. 1996. Partial characterization of the genome of nine animal caliciviruses. Arch Virol 141:2443–2456. Mayr A, Franke J, and Ahne W. 2000. Adaptation of reptilian paramyxovirus to mammalian cells (Vero cells). J Vet Med B Infect Dis Vet Public Health 47:95–98. Nicoletti P. 1990. Brucella. Introduction, in Diagnostic Procedure in Veterinary Bacteriology and Mycology, Carter GR and Cole JR Jr (Eds.), Academic Press, San Diego, CA, 95–105. Origgi F and Jacobson ER. 2000. Diseases of the respiratory tract of chelonians. Vet Clin N Amer: Exot Anim Pract 3:537–549. Origgi FC, Klein PA, Mathes K, Blahak S, Marschang RE, Tucker SJ, and Jacobson ER. 2001. Enzyme-linked immunosorbent assay for detecting herpesvirus exposure in Mediterranean tortoises (spur-thighed tortoise [Testudo graeca] and Hermann’s tortoise [Testudo hermanni]). J Clin Microbiol 39:3156–63. Origgi FC, Romero CH, Bloom DC, Klein PA, Gaskin JM, Tucker SJ, and Jacobson ER. 2004. Experimental transmission of a herpesvirus in Greek tortoises (Testudo graeca). Vet Pathol 41:50–61. Paré, JA, Sigler L, Rypien KL, and Gibas CFC. 2003. Cutaneous mycobiota of captive squamate reptiles with notes on the scarcity of Chrysosporium anamorph of Nannizziopsis vriesii. J Herpetol Med Surg 13:10–15. Ramis A, Fernandez-Bellon H, Majo N, Martinez-Silvestre A, Latimer K, and Campagnoli, R. 2000. Adenovirus hepatitis in a boa constrictor (Boa constrictor). J Vet Diagn Invest 12:573–576. Sachse K. 2003. PCR Detection of Microbial Pathogens. Humana Press, Totowa, NJ. Sacks SL, Griffiths PD, Corey L, Cohen C, Cunningham A, Dusheiko GM, Self S, Spruance S, Stanberry LR, Wald A, and Whitley RJ. 2004. HSV shedding. Antiviral Res. 63 Suppl 1:S19–26. Schat K A and Purchase HG. 1989. Cell culture methods, in A Laboratory Manual for the Isolation and Identification of Avian Pathogens, 3rd edition, Purchase HG, Arp LH, Domermuth CH, and Pearson JE (Eds.), Kendall/Hunt Publishing Co., Dubuque, IA, 167–175. Schumacher J, Jacobson ER, Homer BL, and Gaskin GM. 1994. Inclusion body disease in boid snakes. J Zoo Wildl Med 25:511–524. Thoen C. O. 1990. Mycobacterium, in Diagnostic Procedure in Veterinary Bacteriology and Mycology, Carter GR and Cole JR Jr (Eds.), Academic Press, San Diego, CA, 287–298. Ward DM, Weller R, and Bateson MM. 1990. 16S rRNA sequences reveal numerous uncultured microorganisms in a natural community. Nature 345:63–65. 

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Figure 13.1  Broth culture media in bottles for whole-blood bacterial culture or enrichment.

Figure 13.2  Trypticase soy broth with resins is used if the animal has been on antibiotic therapy. The resins will bind the antibiotics and will increase the likelihood of culturing bacteria. Courtesy of Elliott Jacobson.

Figure 13.3  Mini sample collection set composed of a sterile swab, sterile broth culture medium, and a sterile syringe. Courtesy of Elliott Jacobson.

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Figure 13.4  An appropriate shipping container for biological specimens. A thick Styrofoam container should also be shipped within a well-marked cardboard box. Courtesy of Elliott Jacobson.

Figure 13.5  A damaged shipping container. Styrofoam containers should always be shipped in a thick cardboard box. Courtesy of Elliott Jacobson.

Figure 13.6  Plasma samples for serology often arrive in a thawed condition. Unless the icepacks and samples are together within a thick Styrofoam box, the samples often thaw and warm. Courtesy of Elliott Jacobson.

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Figure 13.7  A cell culture growing in a plastic flask is observed under an inverted-phase light microscope.

Figure 13.8  Tissue culture medium and enrichments used in viral isolation.

Isolation of Pathogens  677

Figure 13.9  A stained monolayer of normal viper heart cells that was grown in a specialized chambered slide. The cells are spindle-shaped and may take two weeks to grow over the surface of a plastic flask. WrightGiemsa stain. Courtesy of Elliott Jacobson.

Figure 13.10  Photomicrograph of a herpesvirus-infected Terrapene heart (TH-1) cell monolayer. An area of lysis (CPE) is seen in this monolayer. The dark brown edges of the area consist of the viral-infected (tortoise herpesvirus) cells, which have been stained with an antitortoise herpesvirus antiserum using an immunohistochemical staining technique.

Figure 13.11  Photomicrograph of uninfected TH-1 cells. The DNA in the nuclei stains purple, while the RNA in the cytoplasm stains light blue. Schiff-methylene blue technique.

Figure 13.12  Photomicrograph of TH-1 cells at six days following infection with a tortoise herpesvirus. Compared to Figure 13.11, note the fainter staining, which likely reflects the lower amount of RNA in the infected cells. Also note the darker edges of the nuclei compared to the center, reflecting margination of chromatin material and presence of intranuclear inclusions. Schiff-methylene blue technique.

678  Isolation of Pathogens

Figure 13.13  Photomicrograph of paramyxovirus-infected viper heart cells. Note the syncytial cells that formed (arrows). WrightGiemsa stain. Courtesy of Elliott Jacobson.

Figure 13.14  Photomicrograph of viper heart cells infected with an adenovirus isolated from a boa constrictor. Basophilic intranuclear inclusions (arrows) are seen. Wright-Giemsa stain. Courtesy of Elliott Jacobson.

Figure 13.15  Primary kidney cell cultures from a boa constrictor with inclusion body disease. Intracytoplasmic inclusions (arrows) are seen. Courtesy of Elliott Jacobson.

Isolation of Pathogens  679

Figure 13.16  Blood-agar plate with a pure bacterial culture (isolated bacterial colonies).

Figure 13.17  Pale colonies of a nonlactose fermenting bacterium are seen growing on this plate of MacConkey’s agar. Courtesy of Elliott Jacobson.

Figure 13.18  Pink colonies of a lactose fermenting bacterium are seen growing on this plate of MacConkey’s agar. Courtesy of Elliott Jacobson.

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Figure 13.19  Colonies of Salmonella having black centers are seen growing on this plate of Hextoen enteric agar. Courtesy of Elliott Jacobson.

Figure 13.20  Sabouraud’s dextrose and Mycosel or Mycobiotic agars are commonly used in isolating fungi from reptiles. Sabouraud’s dextrose is a nonselective agar. Mycobiotic agar is particularly useful in isolating keratinophilic fungi. Courtesy of Elliott Jacobson.

Index 70% alcohol, use in necropsy, 222

A Abnormal heterophil morphology, 179 Abscess in Burmese python oral cavity, 515, 516 in Eastern rock turtle, 514 in green iguana, 514, 515, 517 in green turtle, 514 in Jackson’s chameleon, 518 in veiled chameleon, 517, 518 Acanthocephala, 590 in Chelonia, 590 in Eastern indigo snake, 654 in King cobra, 653, 654 in Squamata, 590 Acari, 593 acariformes, 595–596 parasitiformes, 593–595 Acariformes Cloacaridae, 595 Ophioptidae, 595–596 Pterygosomatidae, 596 Trombiculidae, 596 Acidophils as heterophils, 258 in histologic section, 258 resemblance to inflammatory heterophilic infiltrates, 260 Acremonium infections, 539 Acrodont dentition, in lizards, 6 Actinomycetales infections, 468 Dermatophilus, 469–470 Mycobacterium, 468–469 Nocardia, 469 Acute lymphocytic leukemia, in snakes, 182 Adaptive immunity, 131. See also Cell-mediated immune responses; Humoral response macrophage contributions to, 134 role of complement in, 133 Adenoviridae, 401 in Argentine boa constrictor, 331 in Arizona mountain king snake, 435 in bearded dragon, 330, 331, 434

in boa constrictors, 402, 434 in chameleons, 401 in Chelonia, 401 in colubrid snakes, 402 in Crocodylia, 401 EM micrography of, 308 identification by EM, 308 infected viper heart cells, 678 isolation from moribund snakes, 402–403 in Jackson’s chameleon, 330 in lizards, 401–402 in Nile crocodile, 433 in Ophidia, 402–403 in Sauria, 401–402 in Sierra mountain king snake, 435, 443 slow mutation rates of, 363 in viperid snakes, 402 Adrenal cortical cells in gaboon viper, 111 in timber rattlesnake, 111 Adrenal gland in American alligator, 110 in Burmese python, 110, 246 in corn snake, 89, 111 in gaboon viper, 111 in gopher tortoise, 82 in green iguana, 88, 96, 110, 111 in green turtle, 77 in loggerhead sea turtle, 110, 252 in monocled cobra, 111 necropsy examination of, 225 in reptiles, 20 in snakes, 227 in turtles/tortoises, 228 Aegyptianella, in Japanese grass lizard, 609 Aeromonas infections, 463 in boa constrictor, 490 in Eastern diamondback rattlesnake, 490 Affinity maturation, in immunoglobulins, 142 African beauty racer mixed lymphocyte reaction in, 147 NK cell activity in, 135–136 seasonal and hormonal effects in, 148

Index  681

682  Index

African spurred tortoise fungal infection in, 548 reactive monocytes in, 206 Agama lizard, Plasmodium in, 210 Agamidae spp., bearded dragon, 47, 124 Agglutinating antibodies, 145 Air sac, in neotropical rattlesnake, 80 Alaria infection, in Texas indigo snake, 641 Aldabra tortoises, 534 ascariasis in, 653 fungal lesions in liver, 565 granulated thrombocytes in, 208 hematologic values, 171 melanomacrophages in, 216 mycosis in, 537 Penicillium infection in kidney of, 564 Penicillium infection in liver of, 565 reactive monocytes in, 206 Alignment algorithms, 365 Alimentary tract, in reptiles, 9 Alkaline phosphatase (AP) use in direct ELISA, 383 use in immunoperoxidase tests, 384 Alligatoridae spp. American alligator, 31, 46, 50, 52, 65, 83, 92, 94, 95, 96, 97, 101, 110, 112, 126, 165 thymus anatomy in, 138 Allografts, cell-mediated immune responses to, 146–147 Alpha epidermal layer, 5 Alternative pathway, of complement activation, 133–134 Alveolar tissue, in Aruba Island rattlesnake, 80 American alligator ascariasis in, 643 atretic follicles in, 101 basophilic punctuate inclusions in, 196 brain and anterior spinal cord in, 112 coccygeal venipuncture in, 169, 191 coelomic cavity in, 65 complement mediated experiments in, 134 cornea in, 31 degranulated basophils in, 202 degranulated heterophils in, 199, 200 Dujardinascaris egg in, 642 ELISA tests in, 146 eye in, 126 fungal pneumonia, 539 GI tract in, 32 hemogregarine infection in, 628 heterophil activity in, 135, 197 kidney in, 83 leeches in, 658 mycoplasmic infection in, 344, 504 mycoplasmosis in, 503, 505, 506 oral cavity in, 31, 52 ovary in, 94, 95, 96 pedal abscess exudate in, 275 penis in, 92 Pentastomida in, 655, 657 pericarditis in, 289 production of tetanus antibodies in, 145 Sebekia mississippiensis in, 656 serological testing for arboviruses in, 389 skull in, 46 splenic anatomy in, 139, 164, 165 supravertebral vessel blood sampling, 190 T-cell-like activity in, 137

teeth in, 50 TEM micrograph of blood cells, 346 tongue in, 52 vitellogenic follicles in, 95, 97 West Nile virus in, 338, 459, 460 American anole, Trichosporon infection in, 569 American crocodile, 32 eosinophils in, 201 toxic heterophils in, 201 American Type Cell Collection (ATCC), 669, 672 Ammonium heparin, 170 Ammonium molybdate, immunolabeling for electron microscopy, 303 Amniotic eggs, 3 in reptiles, 1 Amoebae, 572–573 in boa constrictor, 613 in Chelonia, 573 in Crocodylia, 573 in rat snake, 612 in red-footed tortoise, 611, 612 in Rhynchocephalia, 573 in Russell’s viper, 613 in Squamata, 574 in wood turtle, 612 Amphicoelous vertebrae, 7 Amphisbaenia, 3 Amyloid-like material in green anaconda, 287 in inflammation, 264 Anaconda, Cryptococcus infection in, 569 Anaerobe infections, 467–468 Anal gland infection in California king snake, 516 in Sinaloan milk snake, 516 Anamorphs, 528 Anasakis, in loggerhead sea turtle, 643 Anatomy. See also Reptile anatomy thymus variations in reptiles, 138 Anemia binucleated thrombocytes in, 183 use of MCV and MCHC to characterize, 174 Annealing step, in PCR, 357 Annelida, hirudinea, 592 Anterior telencephalon in gopher tortoise, 115 in green iguana, 116 Antibodies, 140–146. See also Immunoglobulins limited shelf life in protein arrays, 376 normalization in reptiles, 145 Antibody assays, 381 Antibody-dependent cell-mediated cytotoxicity (ADCC), 135 Antibody detection, in western blotting, 355 Antibody microarray, 375 Antibody response, 145 in chelonians, 145 in Crocodylia, 145 in Ophidia, 146 recent serologic applications, 146 in Rhynchocephalia, 145 in Sauria, 145–146 Anticoagulants, 170 Antigen binding, for electron microscopy, 303 Antigens, 131 differential responses to, 150 Aorta in corn snake, 105

Index  683

in reticulated python, 105 Apicomplexa hemoparasitic, 579–580 nonhemoparasitic, 575–580 Apophyses, 7 Arachidonic acid metabolites, 136 Arafura file snake, CANV lesions in, 560 Arboviruses serodiagnostics in reptiles, 389 West Nile virus infection of crododilians, 389 Argasidae, 593 in Chelonia, 593–594 in Crocodylia, 594 in Squamata, 594 Argentine boa constrictor, adenovirus infection in, 330 Argentine tortoise, herpesvirus infection in, 329, 398, 430 Arizona mountain king snake adenovirus infection in, 435 mesocestoides in, 634 Arrays, 374 gene-expression arrays, 374–375 protein arrays, 375–376 Aruba Island rattlesnake alveolar tissue in, 80 paramyxovirus antigen in, 408, 448 paramyxovirus in, 284, 336, 337, 444, 445, 446, 447 rhabdovirus infection in, 414 splenopancreas in, 285 Vero cells in paramyxovirus infection, 448 Ascaridoidea, 584–585 in Aldabra tortoise, 653 in American alligator, 643 in blood python, 642 in Burmese python, 644 in Caiman Island ground iguana, 665 in carpet python, 664 in Chelonia, 585 in common agama, 665 in Crocodylia, 585 in Eastern hognose snake, 644 in Florida banded water snake, 653 in gopher tortoise, 652 in green iguana, 664 in leopard tortoise, 652 in loggerhead sea turtle, 643 in Nile monitor, 653 in rat snake, 653 in Russell’s viper, 664 in spider tortoise, 652 in Squamata, 585–586 in water snake, 644 Ascidae, 594 Asexual reproduction, 17. See also Parthenogenesis Asia minor spur-thigh tortoise, herpesvirus in, 398 Aspergillus, 536–537 cell culture precautions, 672 Aspirates, sample collection for NSEM, 306 Atretic follicles in American alligator, 101 in loggerhead sea turtle, 101 in Mexican spiny-tailed iguana, 101 Australian pythons, neurological disease of, 411–412 Australian snakeback turtle, 30 Autopsy. See Necropsy Axial skeleton in red-eared slider, 53

in reptiles, 7 Azurophilic monocytes, 136, 182 in green iguana, 201 Azurophils in histologic section, 259–260 in indigo snake, 207 in mangrove monitor lizard, 207 in rainbow boa, 207

B B-cell immunity, reptilian analogues, 137 Bacteremia in Chelonia, 478 in Crocodylia, 478 in Squamata, 478–479 Bacteria in boa constrictor, 217 Chlamydophila, 265–266 Chlamydophila identification by EM, 311–312 detection in tissue section, 265–266 in Fischer’s chameleon, 218 identification by EM, 311 lactose and nonlactose fermenting, 679 miscellaneous identification by EM, 312 Mycobacterium, 265 mycoplasma identification by EM, 311 in peripheral blood, 185–186 in rhinoceros iguana, 218 tissue handling in necropsy for, 232 Bacterial bronchopneumonia, in mugger crocodile, 272 Bacterial culture, laboratory fees for, 221 Bacterial diseases, 461–462 Actinomycetales infections, 468–470 Aeromonas infections, 463 anaerobe infections, 467–468 bacteremia, 478–479 bacterial infections of eye and orbit, 476 Borrelia infections, 474 Chlamydia and Chlamydophila infections, 472–474 Citrobacter infections, 465 Coxiella infections, 474–475 Dermatophilus infections, 469–470 Elizabethkingia infections, 466 Erysipelothrix infections, 466 gingivitis, 476–477 Helicobacter infections, 467 Leptospira infections, 474 Listeria infections, 466–467 miscellaneous, 475, 479–480 mycoplasma infections, 470–472 Neisseria infections, 466 Nocardia infections, 469 osteomyelitis, 478–479 Pasteurella infections, 466 pharyngitis, 476–477 pneumonia, 477–478 Pseudomonas infections, 462–463 Salmonella infections, 463–465 Serratia infections, 465 shell disease of aquatic turtles, 475 stomatitis, 476–477 Streptococcus infections, 467 subcutaneous and tissue abscesses and masses, 475–476 Vibrio infections, 466 Bacterial exposure Coxiella, 391

684  Index

Leptospira, 391 mycoplasmosis, 390 serodiagnostics of, 390 Bacterial identification, with 16S ribosomal RNA gene, 361 Bacterial isolation, 670–671 blood-agar plate for, 679 culture media selection for, 670 tripticase soy broth culture media for, 674 Bacterial pneumonia, 477 in Chelonia, 477 in Eastern diamondback rattlesnake, 519 in Squamata, 477 Bacterial taxonomy, 362 Balantidium, in island night lizard, 610 Ball python basophilic punctuate inclusions in, 195 capillaria in, 652 fungal dermatitis in, 558 heterophils in, 197 immature lymphocytes in, 205 liver in, 66 lung in, 245 oomphalitis in, 526 pelvic girdle remnants in, 54 Pseudomonas infection in, 490 serial blood sampling in, 170 Barbour’s map turtle, Elizabethkingia meningoseptica infection in, 346 Barnacles, in loggerhead sea turtle, 660, 661 Basic Local Alignment Search Tool (BLAST), 259 Basophilia with blood parasites, 181 in liver of boa constrictor, 402, 457 in Nile crocodile, 401 Basophilic intranuclear inclusions, in carpet python, 319 Basophilic punctuate inclusions, 177, 195, 196 Basophils, 136, 180–181 in American alligator, 202 in blood python, 202 in green iguana, 201, 202 in histologic section, 259 as predominant circulating leukocyte in chelonians, 259 in spiny turtle, 271 Bayesian methods Bayesian posterior probabilities, 370 defined, 376 Beaded lizard, 3 Bearded dragon adenovirus infection in, 330, 331, 402, 434 CANV infection in, 534, 554 Dependovirus infection in, 336 dermatomycoses in, 532 eye in, 124 hepatic necrosis in, 401 histiocytic microsporidial hepatitis in, 295, 297 microsporidia infection in, 349 oral fungal infection in, 554 plasmacytoid lymphocytes in, 204 reactive lymphocytes in, 204 skull in, 47 subcutaneous abscess in, 515 yellow fungus disease in, 562 Beauveriosis, 538–539 Behavioral fever, 261 Behavioral thermoregulation, in reptiles, 4 Berg’s adder, paramyxovirus infection in, 448 Besnoitia, in lizards and snakes, 578–579

Beta epidermal layer, 5 Binucleated thrombocytes, 183, 210 Biological measurements, in necropsy, 223 Biology. See Reptile biology Biopsies, sample collection for electron microscopy, 304–305 Bipedidiae spp., five-toed worm lizard, 33, 46 Black and white tegu coelomic cavity in, 77, 239 internal examination in necropsy, 239 lung in, 77 splenopancreas in, 242 Black-tailed rattlesnake necropsy examination of, 244 trachea and esophagus in, 244 visceral examination of, 245 BLAST software. See Basic Local Alignment Search Tool (BLAST) BLASTN search tool, 359 Blood agar, 671 for bacterial isolation, 679 Blood cells sample collection for electron microscopy, 304 TEM micrograph in American alligator, 346 Blood-forming organs, bone marrow, 137 Blood parasites, 183. See also Hemoparasites basophilia with, 181 Blood python ascarid egg in, 642 basophils in, 202 binucleated thrombocytes in, 210 chronic granuloma in, 282 granuloma formation in, 276 heterophils in, 202, 210 palatine vein blood collection in, 169, 191 Pseudomonas infection in, 488 reactive lymphocytes in, 24 Blood samples cardiac sampling, 168, 190 in Chelonia, 168–169 collection and handling, 167–168, 386 in Crocodylia, 169 in Ophidia, 169–170 paired for acute and convalescent phase, 381 in Sauria, 169 serial, 170 Blotting techniques, 351 northern blotting, 352, 354 Southern blotting, 352, 353 western blotting, 354–356 Blue panther chameleon, internal examination in necropsy, 240 Boa constrictor, 36, 37 abscessed esophageal tonsil in, 455 adenovirus-associated hepatic necrosis in, 402 adenovirus infection in, 434 Aeromonas infection in, 490 amoebiasis in, 613 bacterial inclusions in, 217 brain inflammation in, 452 capillaria egg in, 652 cardiac puncture in, 192 Dermatophilus infection in, 499 dracunculiasis in, 648 ELISA tests in, 146 eosinophilic cytoplasmic inclusions in, 451 esophageal tonsils in, 453, 456 filariasis in, 647 Hastospiculum varani in, 649

Index  685

heart and lung in, 104 hepatocytes with intracytoplasmic inclusions in, 454 Hepatozoon in, 210, 609 herpesvirus infection of, 400 histiocytic granuloma formation in, 277 IBD inclusions in, 212, 213 inclusion body disease in, 339, 340, 450 indirect ELISA for IBD in, 389–390 intranuclear inclusion bodies in, 288 Kalicephalus in, 646 kidney cells in IBD, 678 kidney with eosinophilic cytoplasmic inclusions in, 455 Klossiella infection in, 627 liver with eosinophilic intracytoplasmic inclusions in, 456, 457 lung with intracytoplasmic inclusions in, 453 microfilaria in, 648 mixed viral-associated GI disease in, 400 monocytes in, 217 mycoplasmosis with ulcerative mouth lesion in, 494 normal brain cells in, 452 normal vertebrae in, 286 osteoarthritis in, 524, 525 osteomyelitis in, 523, 524 pancreas with intracytoplasmic inclusions in, 454 peripheral blood in, 457 Pseudomonas infection in, 489 retrovirus identification by EM in, 319, 340, 341, 342 reversal lines in bone remodeling, 263, 286 skin shedding in, 37 skull and mandible in, 48, 49 strongyloides egg in, 651 thyroid gland in, 107 tissue-specific azurophil function in, 259 Body cavity fluids, preparation for electron microscopy, 306 Body condition index, 223 Boidiae spp. Boa constrictor, 36, 37, 48, 67, 104, 107 Dumeril’s ground boa, 38, 39, 40, 44, 56, 60, 76, 84, 118, 119, 120, 123 Emerald tree boa, 126, 130 green anaconda, 126 Madagascan tree boa, 71 serodiagnostics of IBD in, 389–390 Bolivian side-neck turtle fungal shell disease in, 554 papillomaviruses in, 336, 442 Bone marrow, 19 in carpet python, 271 hematopoeisis in, 260 immunological role, 137–138 Bone remodeling, in boa constrictor, 286 Bony shell, in Chelonia spp., 2 Bootstrapping, 409 as confidence measure, 369–370 defined, 376 Borrelia infections, 474 Bothridium, in Timor python, 632 Bothrops, liver in, 67 Bouin’s fixative, 229 Box turtle cryptosporidiosis in, 620 Ranavirus infection identification by EM, 320 Brain in American alligator, 112 in American alligator with mycoplasmosis, 505, 506 in American alligator with West Nile virus, 460 in boa constrictor, 452

in Burmese python, 113 in Chinese alligator, 243 in corn snake, 116 in death adder, 113 with eosinophilic inclusions in Burmese python, 451 in gopher tortoise, 112 in green iguana, 112, 117, 121 with inflammation in boa constrictor, 452 with intracytoplasmic inclusions in boa constrictor, 451 necropsy examination of, 226, 243 removal in Burmese python, 247 removal in loggerhead sea turtle, 253, 254 removal in sea turtles, 228 removal in snakes, 227 in Schneider’s dwarf caiman, 243 Branch lengths, 368 Bronchi, in chelonians, 12–13 Bronchoscopy, with electron microscopy, 306 Broth culture media, 668 for pathogen isolation, 674 Brother Island tuatara, 3 Brown anole, epidermis in, 43 Bull snake, cryptosporidiosis in, 624, 625 Burmese mountain tortoise hematologic values, 171 serological testing for iridovirus in, 388 Burmese python adrenal gland in, 110, 246 ascariasis in, 644 brain and anterior spinal cord in, 113 brain removal in, 247 brain with eosinophilic cytoplasmic inclusions in, 451 chronic pulmonary disease in, 519, 520 epithelial cells and thymocytes in, 160 fusariomycosis in, 563 gallbladder in, 430 GI tract in, 12 heart in, 245 inclusion body disease in, 452 liver in, 69 lung in, 78 mycoplasmic infection in, 344 mycoplasmosis in, 506 necrotizing skin lesion in, 563 oral abscess in, 515 parathyroid gland in, 108 Pseudomonas infection in, 489 respiratory tract in, 245 retrovirus infection in, 342 round cell tumor in, 410, 449 spinal cord in, 120 spinal cord removal in, 248, 249 spleen with red and white pulp in, 166 thromboembolic disease in, 521, 522 thymic lobules in, 160, 161 uterus in, 103 Burn-in period, in Bayesian methods, 368 Bursa of Fabricius, 136 Bush anole, Dermatophilus infection in, 497 Bush viper kidney in, 85 lung in, 79 Bushmaster snake fungal dermatitis in, 294 granulomatous dermatitis in, 293 rhabdiasis in, 650

686  Index

C C-opsonins, 135, 145 Caecum, in reptiles, 10 Caiman Island ground iguana, ascariasis in, 665 Caiman lizard exudative pneumonia in, 274 heterophilic granuloma formation in, 276, 291 paramyxovirus infection in, 443, 444 rhabdovirus infection in, 414, 460 serological testing for paramyxovirus in, 388 Caiman poxvirus, 403 in spectacled caiman, 436, 437 Caliciviridae, 414 in Ophidia, 414 California king snakes anal gland infection in, 516 mesos layer in, 5 California red-sided garter snake granuloma formation in, 276 macrophage-rich heterophilic granuloma in, 280 Candidiasis, 541–542 in crocodile tegu, 542 CANV, 534 in Arafura file snake, 560 in bearded dragon, 554 in humans, 534 as hyalohyphomycotic agents, 533–534 in squamates, 532 in tentacled snake, 561 in veiled chameleon, 549, 550, 561 Capillaria in ball python, 652 in boa constrictor, 652 Capillary agglutination test for coxiella in snakes, 391 Capture ELISA, 384 Carcass preservation, 221–222 Cardiac sampling, 168 in boa constrictor, 192 in gopher tortoise, 190 Cardiovascular system, 18–19 Carettacola, in loggerhead sea turtle, 640 Carotid artery venipuncture, in chelonians, 169 Carpet python ascariasis in, 664 basophilic intranuclear inclusions in, 319 bone marrow in, 271 CNS disease in, 411–412 intranuclear inclusions in, 318 proliferative osteoarthritis in, 286 Caryospora, 576 in Chelonia, 576 in flat-headed snake, 623 in green anole, 623 in lizards and snakes, 577 in Madagascar common snake, 623 in Madagascar hognose snake, 623 Caudal telencephalon, in gopher tortoise, 116 Caudal vertebrae, in reptiles, 7 Cell-associated virus isolation, 670 Cell cultures for pathogen isolation, 676 recommended culture time, 670 sample collection for electron microscopy, 304 temperature requirements, 670 Cell degeneration, 232

Cell lines, 669 Cell-mediated immune responses, 132, 146 to allografts and xenografts, 146–147 Graft versus Host Reaction (GVHR), 147 mixed lymphocyte reaction (MLR), 147 Centraalbureau voor Schimmelcultures (CBS), 672 Centrifugation, for serodiagnostics, 386 Cerebellar cortex, in green iguana, 118 Cerebellum in Dumeril’s ground boa, 119 in reptiles, 21 Cerebrum, in reptiles, 20 Cervical sinus venipuncture, in loggerhead turtles, 190 Cervical vertebrae, in reptiles, 7 Cestoda Cyclophyllidea, 582 Proteocephalidea, 582 Pseudophyllidea, 581–582 Trypanorhyncha, 582 Chaco tortoise, thrombocyte clumps in, 193 Chamaeleonidae adenovirus infection of, 401 Jackson’s chameleon, 48, 51 Senegal chameleon, 88 veiled chameleon, 43, 44, 78, 95, 109, 161 Chelonia, 1, 2 Acanthocephala in, 590 adenoviruses in, 401 amoebae in, 573 antibody response in, 145 Argasidae in, 593–594 Ascaridoidea in, 585 Australian snakeneck turtle, 30 bacteremia and osteomyelitis in, 478 bacterial pneumonia in, 477 blood sampling techniques, 168–169 bronchi in, 12–13 Caryospora in, 576 Chlamydia and Chlamydophila infections in, 472 circoviruses in, 407 Dermatophilus infections in, 469 Diptera in, 596–597 ear anatomy, 23 Eimeria, Isospora, and unclassified coccidians in, 575–576 embryonic shields in, 4 eye anatomy, 22 flaviviruses in, 413 green turtle, 52, 54, 55, 65, 73, 77, 81, 106, 125, 158, 159 Haemogregarina in, 579 Hawksbill sea turtle, 45, 50 Hepatozoon in, 579 herpesviruses in, 396–399, 396–400 immunoglobulins in, 143–144 iridoviruses in, 404–405 Ixodidae in, 593–594 Lacazia loboi infections in, 539–540 loggerhead sea turtle, 86, 95, 100, 101, 102, 110, 125, 159, 161, 162 miscellaneous bacterial infections in, 479–480 mycoplasma infection in, 470–471 mycoplasmosis serodiagnostics, 390 mycoses in, 531–532 nonhemoparasitic apicomplexa in, 575–576 papillomaviruses in, 406 Parabasalia in, 574 paramyxoviruses in, 407 pectoral girdle in, 7

Index  687

Pentastomida in, 590–591 Poxviridae in, 403 reoviruses in, 412 retroviruses in, 409 rhabdoviruses in, 414 rhamphotheca in, 6 secondary palate in, 6 serology of iridoviruses in, 388 shell measurements in necropsy, 223 skulls in, 6 slow mutation rates in, 363 sperm conveyance in, 15 Spirorchiidae in, 583–584 spleen investigations in, 139 stomatitis, gingivitis, and pharyngitis in, 476 subcutaneous abscesses/masses in, 475 thymus anatomy in, 138 thymus lobulation in, 138 togaviruses in, 413 trimmed toenail blood sampling in, 168 vestibulum nasi in, 12 Chelydridae spp., common snapping turtle, 45 Chemotactic factors, 133 Chicken turtle, Spirorchiidae in, 636, 638 Chinese alligator brain necropsy examination in, 243 fungal hyphae in, 549 retrovirus infection in, 409 Chinese dragon, toxic heterophils in, 198 Chlamydia inclusion, 182, 207 in Emerald tree boa, 218 in flap-necked chameleon, 333, 438 in green turtle, 345 identification by EM, 311–312 in puff adder, 345 Chlamydia infection, 472 in Chelonia, 472 in Crocodylia, 472 in Emerald tree boa, 509, 510, 511 in flap-necked chameleon, 507, 508 in green turtle, 507 in Ophidia, 473–474 in puff adder, 508, 509 in Sauria, 473 Chlamydophila, detection in tissue section, 265–266 Chromoblastomycosis, 541 Chromosome mapping, by in situ hybridization, 371 Chronic granulomas, 277 in blood python, 282 in Emerald tree boa, 282 as end-stage lesions, 261 mineralization in, 262, 282 in Western diamondback rattlesnake, 295 Chronic interstitial nephritis, in Dumeril’s boa, 279 Chronic pulmonary disease, in Burmese python, 519, 520 Chrysosporium anamorph of Nannizziopsis viresii (CANV), 528. See also CANV Chuckwalla, fungi growth in, 548 Circoviridae, 406 in Chelonia, 407 identification by EM, 309 in painted turtle, 336 spleen with inclusions, 443 Circulating inflammatory cells, 167 erythrocytes and erythrocyte responses, 176–179 hematological concepts and, 167

infectious agents in peripheral blood, 183–186 leukocytes and leukocyte responses, 179–182 thrombocytes and thrombocyte responses, 182–183 Cirripedia, 592–593 Citrobacter infections, 465 in slider, 491 Class Reptilia, 1 Class-switching recombination, in immunoglobulins, 142 Classical pathway, of complement activation, 133–134 Cloacaridae, 595 Clostridium infection, in Dumeril’s ground boa, 290 Clotting system, 136 CNS disease in diamond python and carpet python, 411–412 in Haitian boa, 408, 410, 450 Cobra, liver in, 71 Coccidioides infection, 543 in Chelonia, 575–576 in gila monster, 570 in green turtle, 618, 619 in helmeted iguana, 615 in inland bearded dragon, 616 in kidney of Texas indigo, 570 in leopard tortoise, 617 in Nile crocodile, 621 in radiated tortoise, 616, 617 Coccygeal venipuncture, in American alligator, 169, 191 Coding system, 233 for necropsy diagnoses, 234 Coelomic cavity in American alligator, 65, 110 in black and white tegu, 77, 225, 239, 240 in corn snake, 78 in diamond python, 78 exposing in necropsy internal examination, 239 in forest turtle, 250 in gopher tortoise, 104 in green iguana, 66, 82 in green turtle, 65, 77, 81, 250 necropsy exam techniques in turtles/tortoises, 227 in Nile crocodile, 240 in red-eared slider, 65 in snakes, 244 in water snake, 81 Cold stunning, and development of mycoses, 531 Collared lizard, Dermatophilus infection in, 498 Colon in corn snake, 64, 81 in desert tortoise, 64 in Emerald tree boa with Chlamydia infection, 511 in reptiles, 10–11 in water snake, 81 Colorimetric techniques, 385 for in situ hybridization, 370 Colubridae spp. adenovirus infection in, 402 corn snake, 33, 34, 67, 68, 74, 81, 84, 85, 89, 92, 94, 105, 107, 111, 114, 115, 116, 119, 126, 128, 129 European grass snake, 38 king snake, 79 tentacled snake, 81 Tolucan ground snake, 103 Trans-Pecos rat snake, 127 water snake, 81 Columbian slider, 30 Common agama

688  Index

ascariasis in, 665 Plasmodium infection in, 608 Common boa, ulcerative enteritis in, 294 Common chameleon, hematologic values, 173 Common house gecko, Isospora in, 622 Common snapping turtle, skull and mandible in, 45 Compacted course-cancellous bone, 8 Competitive ELISA, 383–384 Complement, 131, 136 as nonspecific humoral factor, 133–134 Complement activation, immunoglobulins and, 145 Complement fixation, by antibodies, 145 Complement inflammatory cascade, 133 Complementary determining regions (CDRs), 141 Complete blood counts (CBCs), 170 Computational simplicity, of distance-based trees, 367 Confidence bootstrapping method, 369–370 methods of measuring, 368–369 Conidia, 528 Consensus PCR, 357 Consensus sequences, 365 Contamination avoiding in sample handling for pathogen isolation, 668 in PCR analysis, 360 Cook’s tree boa, Proteus infection in, 519 Copperhead snake, pit organ in, 130 Corn snake, 33, 34 adrenal gland in, 89, 111 brain transverse section, 116 coelomic cavity in, 78 colon in, 64, 81 dorsal aorta in, 105 eye in, 126 fungal dermatitis in, 559 Geotrichum infection in, 566 head transverse section, 114 hemipenes in, 92 inner ear infection in, 525 kidney in, 81, 84 lung in, 78 mid-coelomic cavity in, 67 monocercomoniasis in, 615 olfactory nerves in, 114 olfactory tract in, 115 osseous and otic labyrinth in, 128 pancreas in, 74 pars pylorica region of stomach, 62 posterior medulla in, 119 rhabdiasis in, 650, 651 sexual segment in, 85 spatial relationships of pancreas, spleen, gallbladder, 68 testes in, 89 thyroid gland in, 107 vitellogenic follicles in, 94 vomero-nasal organ in, 129 Coronaviridae, 415 in Crocodylia, 415 Corpora lutea in green iguana, 99 in Karasberg tree skink, 100 in loggerhead sea turtle, 100 in reptiles, 16 Corpus albicans, in green iguana, 100 Cosmocercoidea, 586 Cost issues, necropsy-related, 220–221

Cottonmouth granulomatous periadenitis in salivary gland, 294 polychromatophils in, 193, 194, 209 reactive thrombocytes in, 209 rubricytes in, 194 Coverslip blood sampling method, 170, 192 Coxiella, 474–475 serodiagnostics of, 391 Crevice spiny lizard, trypanosomes in, 211 Critical thermal maximum (CTMax), 4 Critical thermal minimum (CTMin), 4 Crocodile pox in Johnson’s crocodile, 438 in Nile crocodile, 437 Crocodile tegu, candidiasis in, 542 Crocodylia, 1, 2–3 adenoviruses in, 401 American crocodile, 32 amoebae in, 573 antibody response in, 145 Argasidae in, 594 Ascaridoidea in, 585 bacteremia and osteomyelitis in, 478 blood sampling techniques in, 169 Chlamydia and Chlamydophila infections in, 472 coronaviruses in, 415 Cryptosporidium in, 577 Dermatophilus infections in, 469–470 dissection and internal examination in necropsy, 224–226 ear anatomy, 23 Eimeria, Isospora, and Goussia in, 576–577 extrapulmonary bronchi in, 225, 242 eye anatomy, 22 family Alligatoridae, 2 family Crocodylidae, 2 family Gavialidae, 2 Filarioidea in, 586–587 flaviviruses in, 413 Haemogregarina in, 579 hematologic values, 172 Hepatozoon in, 579 herpesviruses in, 399 immunoglobulins in, 144 Ixodidae in, 594 lack of urinary bladder in, 13 miscellaneous bacterial infections in, 479–480 multichambered lungs in, 13 mycoplasmosis in, 471 mycoses in, 532 necropsy examination in, 240 paramyxoviruses in, 407 Pentastomida in, 591 Poxviridae in, 403 retroviruses in, 409–410 saltwater crocodile, 31 serodiagnostics of mycoplasmosis in, 391 serodiagnostics of West Nile virus in, 389 splenic anatomy in, 139 temporal skull openings in, 6 thymus anatomy in, 138 togaviruses in, 413 Trichinelloidea in, 589 Crocodylidae spp., Siamese crocodile, 96 Cross-linking fixatives, problems for in situ hybridization, 372 Crustacea, 592 cirripedia, 592–593

Index  689

Cryptococcosis, 542 in anaconda, 569 Cryptodira spp., 2 Cryptosporidiosis, 262 in box turtle, 620 in bull snake, 624, 625 in Chelonia, 576 in Crocodylia, 577 in esophageal tonsils of Nile crocodile, 297 in gopher snake, 283 in green iguana, 347, 626, 627 in Hermann’s tortoise, 620 identification by EM, 312–313 in leopard gecko, 347, 625, 626 in lizards and snakes, 578 serodiagnostics of, 391 in sidewinder, 297 in Texas rat snake, 624 in timber rattlesnake, 624 in Trans-Pecos rat snake, 625 Cuban crocodile internal organs in necropsy exam, 242 necropsy examination, 240 Culture-based pathogen isolation, 669. See also Pathogen isolation Cutaneous papillomatosis, in European green lizard, 432 Cyclophyllidea, 582 Cytology techniques, in necropsy, 228–229 Cytopathic effect (CPE), 670 Cytoplasmic granules, in desert tortoise, 327, 328 Cytotoxic T-lymphocytes, 132, 136

D 2D-PAGE, 373 advantages and disadvantages, 374 interpretation of results, pitfalls, limitations, 374 procedure, 373–374 schematic representation, 373 Data gathering, for necropsy, 220, 223 Davidson’s solution, 229 Death adder brain and anterior spinal cord in, 113 venom gland in, 58 Deckert’s rat snake intranuclear inclusion in, 321 nonviral inclusions in, 321 Decomposition, issues in necropsy, 221 Decontamination techniques, after necropsy, 233 Degenerate primers, use in PCR, 357 Degenerate probes, 371 Degranulated heterophils, 179, 198 in American alligator, 200 Dehydration, anemia and, 178 Dematiaceous fungi, 540 chromoblastomycosis, 541 phaeohyphomycosis, 540–541 Denaturation step in PCR, 357 schematic representation, 358 Dendovirus, in Sierra mountain king snake, 443 Dendritic cells, 134–135 Dependovirus infection, in bearded dragon, 336 Dermal bone, in desert tortoise, 36 Dermal pyogranuloma, in green iguana, 492 Dermatitis, in bushmaster snake, 293 Dermatomycoses, 529–530 Dermatophilus infections, 469

in boa constrictor, 499 in bush anole, 497 in Chelonia, 469 in collared lizard, 498 in Crocodylia, 469–470 in desert tortoise, 496 in Ophidia, 470 in padlopper tortoise, 496 in saltwater crocodile, 496 in Sauria, 470 in Senegal chameleon, 498, 499 Dermatophytes, 543 Dermochelydae spp., leatherback sea turtle, 59 Description of findings, in necropsy, 220 Desert iguana cutaneous fungal disease in, 558 responses to Salmonella, 146 temperature effects on inflammatory responses, 261 transferrin experiments in, 133 Desert tortoise, 30, 34, 35 bone marrow samples, 138 calculating body condition index for, 223 colon in, 64 cytoplasmic granules in, 327, 328 dermal bone in, 36 Dermatophilus infection in, 496 ELISA tests in, 146 enlarged melanomacrophages in, 273 eosinophils in, 200 esophagus in, 59 fungal hyphae in, 553 gallbladder in, 68 head cross-section in, 76 hematological values, 171 hemogregarines in, 200 hemoparasites in, 200 herpesvirus in, 318, 398, 429 jugular venipuncture in, 190 liver in, 68, 70, 72 mandible in, 40 melanin granules in, 328 melanomacrophages in liver of, 259, 273, 327 mental gland, 41 monocytes in, 205 mycoplasma infection identified by EM, 321, 344 mycoplasmosis in, 500, 501 nonviral inclusions in, 185, 323 normal nasal cavity in, 500, 501 Nyctotherus in, 610 parathyroid gland in, 108 seminiferous tubules in, 90 small intestine villi in, 63 spermatozoa in, 90 spleen in, 162 testes in, 90, 91 thymic lobule in, 160 tongue in, 57 urinary bladder in, 86, 87 Dewlap, in green iguana, 52, 53 Diagnostics, 258 in necropsy, 219 Diamond python CNS disease of, 411 coelomic cavity in, 78 lung in, 78 monocercomoniasis in, 614

690  Index

paramyxovirus infection in, 447 Diamondback terrapin EM micrograph of liver in, 324 TEM micrograph of liver, 324 Diaphanocephaloidea, 586 Diencephalon in green iguana, 122 in reptiles, 20 Dif-Quick stain, 229 Differential leukocyte counts, 170, 174 Digestive system, in reptiles, 8–12 Digging, adaptations in Amphisbaenia spp., 3 Digital cameras, 224 use in electron microscopy, 300 use in necropsy, 222 Dimorphic fungi, 543 Dioctophymatoidea, 588 Diplogyniidae, 594 Diplostomatidae, 584 Diplotriaenoidea, 587 in Squamata, 588 Diptera, 596 in Chelonia, 596–597 in Squamata, 597 Direct ELISA, 383 Direct leukocyte count, 174 Discrete, defined, 376 Discrete methods, in tree building, 365 Disposal, of necropsied carcasses, 221–222 Dissection of lizards and crocodilians, 224–226 in necropsy, 224 of snakes, 226–227 of turtles and tortoises, 227–228 Distance-based trees, 365–367, 366 Distance matrix, 366 Distance methods, in tree building, 365 DNA amplification, 359 and hydrogen bonding between bases, 357 with PCR techniques, 356 DNA Data Bank of Japan, 364 DNA Data Bank of Japan (DDBJ), 359 DNA databases, 359, 364 confirming PCR results with, 361 DNA degradation, 361 DNA integrity, for ISH, 373 DNA sequence, chromatogram of, 358 DNA sequencing, in PCR, 359 DNA translation, 359 DNA viruses, 396 Documentation, of necropsy findings, 223–224 Dorsal aorta in corn snake, 105 in reticulated python, 105 Dot plot, for sequence alignment, 364 Dracunculiasis in boa constrictor, 648 in monacled cobra, 648 Dracunculoidea, 587 Drying artifacts in Fischer’s chameleon, 217 in RBCs, 170, 192 Dujardinascaris, in American alligator, 642 Dumeril’s ground boa, 38, 61, 62 biopsied scale photomicrograph, 44 cerebellum in, 119

chronic interstitial nephritis in, 279 esophagus in, 60 fundic region of stomach, 61, 62 head cross-section in, 76 head musculature, 56 kidney in, 84 mast cells in, 272 oberhautchen in, 39, 40 optic tectum in, 118 pineal gland in, 123 scales in, 39 small intestine in, 63, 64 spinal cord in, 120 tonsils in, 60 yolk sac with Clostridium infection, 290 Duodenum in Burmese python, 246 in green turtle, 73 in New Guinea snakeneck turtle, 73 Dwarf gecko, Haemocystidium lygodactyli in, 608

E Ear

abscess in Eastern box turtle, 514 anatomy in reptiles, 23 in Chelonia, 23 in Crocodylia, 23 inner ear infection in corn snake, 525 in Ophidia, 23 in Sauria, 23 squamous metaplasia in wood turtle, 287 in tuatara, 23 Eastern box turtle aural abscess in, 514 iridovirus in, 215, 216, 288 Ranavirus infection in, 440, 441 stomatitis in, 518 Eastern diamondback rattlesnake Aeromonas infections in, 490 bacterial pneumonia in, 519 fungal granuloma in, 559 fungal hyphae in, 559 hemipenes in, 92 junctional complexes in, 326 lysosomes in, 326 pancreas in, 74 paramyxovirus in, 444, 446 pinocytic vesicles in, 326 rough endoplasmic reticulum in, 325 skull and mandible in, 49 smooth endoplasmic reticulum in, 325 TEM of normal kidney, 324 TEM of normal liver, 324, 325 uterus in, 103 vertebrae in, 158 Eastern equine encephalitis (EEE), 413 Eastern hognose snake ascariasis in, 644 osteoarthritis in, 524 osteomyelitis in, 524 Eastern indigo snake acanthocephalan infection in, 654 fungal hyphae in, 550 fungal infection of spectacle in, 560 Hepatozoon in, 210 melanomacrophages in, 216

Index  691

renifers in, 635 Eastern king snake fungal dermatitis in, 549 Kalicephalus in, 646 Ecdysis effect on lymphocyte counts, 181 in snakes with fungal disease, 530 stages in reptiles, 4–5 Ectotherms behavioral fever in, 261 reasons for mycoses in, 527–528 vs. endotherms, 4 Eimeria in Chelonia, 575–576 in Crocodylia, 576 in lizards and snakes, 577 in Madagascan giant day gecko, 622 in Mediterranean gecko, 615 in tree monitor, 622 in wood turtle, 616 Elapidae spp. corn snake, 62, 64 death adder, 58, 113 monocled cobra, 70, 71, 74 Naja spp., 83, 85 Electric cutting tools, for necropsy use, 237 Electron microscopy historical perspectives, 30 in necropsy examination, 232 negative staining transmission electron microscopy (NSEM), 303 pathogen identification with, 299–300 positive staining transmission electron microscopy (PSEM), 300 sample preparation in paraffin block, 318 scanning electron microscopy (SEM), 304 sectioning procedures, 302, 318, 319 tissue preparation and sectioning for, 303, 319, 320 ultrathin sections in, 300 Electrophoresis, in Southern blotting, 352 Elizabethkingia infections, 466 in map turtle, 492 Elizabethkingia meningoseptica, in Barbour’s map turtle, 346 Embedding, for PSEM investigations, 302 EMBL data bank, 364 Embryonic shields, in Chelonia spp., 4 Emerald tree boa Chlamydia inclusions in, 218 Chlamydia infection in, 509, 510, 511 chronic granuloma in, 282 fungal skin lesions in, 560 Hepatozoon in, 629 labial pits in, 130 shed spectacle in, 126 Emydidae spp. Columbian slider, 30, 69, 70, 102 painted wood turtle, 125 red-eared slider, 30, 53, 65, 68, 70, 73, 88, 93, 163, 164 Endocrine system adrenal gland, 20 control of reproduction, 17 pancreatic islets of Langerhans, 20 parathyroid gland, 20 pituitary gland, 19 thyroid gland, 19–20 ultimobranchial body, 20 Endosteal bone, 8 Enrichment broths

for bacterial isolation, 671 for viral isolation, 676 Enteritis, in Russell’s viper, 296 Entomophthoromycoses, 538 Entonyssidae, 594 Environmental contaminant studies, necropsy for, 220 Enzymatic digestion, 361 in Southern blotting, 352, 353 Enzyme-linked immunosorbent assay (ELISA), 146, 382, 383 capture ELISA, 384 competitive ELISA, 383–384 direct ELISA, 383 indirect ELISA, 383 plasma consumption as disadvantage of, 390 statistical methods for cutoff determination, 386 Eosinophils, 136, 180 in boa constrictor, 451, 455 in Burmese python, 451 in desert tortoise, 200 in diamond python, 408 in histologic section, 258–259 in liver of boa constrictor, 456 Epidermal growth, in reptiles, 4 Epidermal layers, 5 Epiphysis in green iguana, 55 in green turtle, 54, 55 Epoxy resin 812, 302 Erysipelothrix infections, 466 Erythrocyte evaluation, 174 anemia, 178–179 in disease, 176–179 polycythemia, 178–179 Erythrocyte mitosis, 176, 194 Erythrocyte Unopette system, 174 Erythrocytes abnormalities in, 177–178 immature, 176, 193–194 life span of, 177 mature, 176, 193 normal morphology and function, 176–177 Erythroid regenerative response, 177 Erythroparasites, 178 Erythroplastids, 176, 195 Esophageal tonsils, 140. See also Tonsils abscessed in boa constrictor, 455 in boa constrictor, 411, 453, 456 cryptosporidiosis in Nile crocodile, 297 in IBD, 411 in reticulated python, 455 Esophagus in black-tailed rattlesnake, 244 in desert tortoise, 59 in Dumeril’s ground boa, 60 in green iguana, 59 in horned viper, 59 in leatherback sea turtle, 59 in reptiles, 9 Established cell cultures, 669 Estrogen, immune effects in female Indian leaf-toed gecko, 150 Eublepharidae spp., leopard gecko, 43, 124 European Bioinformatics Institute (EBI), 359 European grass snake, 38 skin shedding in, 38 European green lizard cutaneous papillomatosis in, 432

692  Index

herpesviruses in, 262, 432 lysozymes in, 133 natural killer cell activity in, 135 papillomas in, 399, 406 European tortoises, iridovirus infection of, 404 Exocrine acinar cells, in red-eared slider, 73 Extension step, in PCR, 357–358 External ear, 23 External examination, in necropsy, 224 Extrapulmonary bronchi, in crocodiles, 225, 242 Exudates in inflammatory response, 260 otitis media in wood turtle, 274 in pedal abscess of American alligator, 275 in pneumonia of Caiman lizard, 274 solid nature in reptiles, 260 Eye in American alligator, 126 bacterial infections of, 476 in Chelonia, 22 in corn snake, 126 in Crocodylia, 22 in green iguana, 121 in green tree python, 127 in leopard gecko, 124 in loggerhead sea turtle, 125 necropsy examination in snakes, 249 in New Caledonian bumpy gecko, 124 in Ophidia, 22–23 in painted wood turtle, 125 in reptiles, 21 in Sauria, 22 in Trans-Pecos rat snake, 127 in tuatara, 33, 122, 123

F False confidence intervals, 369 False map turtle, herpesvirus infection in, 329, 428 False negatives, in PCR, 361 False positives in PCR analysis, 360, 361 in in situ hybridization, 371 False tegu, paramyxovirus infection in, 407 False water cobra, tapeworm egg in, 632 Fat body in American alligator, 32 in Crocodylia spp., 2–3, 225, 240 Fat-tail gecko, intranuclear inclusion bodies in, 401 Feces samples, sample collection for NSEM, 306 Female reproductive system anatomy and histology, 15–16 in green iguana, 41 in leopard gecko, 43 Femoral pores in green iguana, 41 in male lizards, 5–6 Fetal bovine serum (FBS), in cell culture medium, 669 Fibrinogen, evaluation, 175–176 Fibropapilloma, 397 in green turtle, 391, 428 Field biologists, 223 Field necropsy, 220 Field observations, in necropsy, 223 Filarioidea, 586 in boa constrictor, 647 in Crocodylia, 586–587

in gila monster, 646, 647 in panther chameleon, 647 in Squamata, 587 Fine-needle aspiration, 229 Fischer’s chameleon bacterial inclusions in, 218 drying artifacts in, 217 erythrocyte virus infection in, 334 iridovirus infection in, 405 LEV infection in, 441 melanomacrophages in, 217 toxic heterophils in, 198 Fixatives, for PSEM, 301–302 Flap-necked chameleon bacterial inclusions in, 186 Chlamydia infection in, 507, 508 chlamydial infection in, 333, 438 Fusarium infection in, 535 iridovirus infection in, 405 lizard erythrocyte virus infection in, 334 monocyte with Chlamydia inclusion in, 207 poxvirus in, 207, 333, 403–404, 438 Flat-headed snake Caryospora in, 623 Isospora in, 622 Flathead leaf-toed gecko, Sauroleishmania in, 609 Flaviviridae, 413 in Chelonia, 413 in Crocodylia, 413 identification by EM, 310 in Ophidia, 414 in Sauria, 413–414 Florida banded water snake ascariasis in, 653 renifers in, 635 Florida soft-shelled turtle, mucormycoses in, 555 Florida worm lizard, 33 Flowerback box turtle, abnormal eosinophil morphology in, 180 Flukes. See Spirorchiidae Fluorescent techniques, for in situ hybridization, 370 Fluorochromes, 384 Fly river turtle, 534 fungal dermatitis in, 556 Follicle stimulating hormone (FSH), 17 Follicular development, in reptiles, 15 Forebrain, in reptiles, 20 Forest turtle, coelomic cavity examination in, 250 Fork-nosed chameleon, melanomacrophages in liver, 273 Formaldehyde, use in northern blotting, 354 Formalin long-term storage problems, 233, 235 necropsy issues, 221, 222 neutral buffered 10%, 302 tissue collection for fixation in, 255 Four-toed tortoise, herpesvirus outbreak in, 399 Fragment crystallizable (Fc), 142 Fragments antigen binding (Fab), 142 Free radicals, neutralization by melanin, 260 Freshwater turtles, herpesvirus infection of, 397–398 Fundic gland region in Dumeril’s ground boa, 61 in green iguana, 61 Funding limitations, for 2D-PAGE studies, 374 Fungal dermatitis in ball python, 558 in bushmaster snake, 294

Index  693

in corn snake, 559 in desert iguana, 558 in Eastern king snake, 549 in Emerald tree boa, 560 in Fly river turtle, 556 in garter snake, 548 in green sea turtle, 554 in Hawksbill sea turtle, 557 in jeweled lacerta, 558 in saltwater crocodile, 557 Fungal granuloma, in Eastern diamondback rattlesnake, 559 Fungal hyphae in African spurred tortoise, 548 in ball python, 558 in Chinese alligator, 549 in desert tortoise, 553 in Eastern diamondback rattlesnake, 559 in Eastern indigo snake, 550 in gopher tortoise, 553 in green turtle, 552 in lung of American alligator, 553 in stomach of Aldabra tortoise, 563 Fungal infections in bearded dragon, 554 detection in tissue section, 266 and immunosuppression, 530 origination in lower respiratory tissues, 530 of spectacle in King snake, 560 in timber rattlesnake, 567 Fungal isolation, 671–672 Sabouraud’s dextrose and Mycosel agar for, 680 Fungal myocarditis, in loggerhead sea turtle, 556 Fungal pneumonia, 531 in alligators, 536 in American alligator, 539, 553 in Crocodylia, 532 in loggerhead sea turtle, 556 Fungal stains, 229 Fungal tracheitis, in loggerhead sea turtle, 289, 293 Fungi cultures from chuckwalla, 548 identification by isolation, 667 overview of, 528–529 tissue handling in necropsy for, 232 Fusariomycosis, 535–536, 548 in Burmese python, 563 in Texas tortoise, 564 in veiled chameleon, 564

G Gaboon viper adrenal cortical cells in, 111 mixed viral-associated GI disease in, 400 Pentastomida in, 655, 656, 657 spleen and pancreas in, 75 subcutaneous abscess in, 515 Gallbladder in American alligator, 65 in boa constrictor, 67 in Burmese python, 246 in corn snake, 67, 68 in desert tortoise, 68 in green iguana, 66 in green turtle, 65 necropsy examination in snakes, 227 in red-eared slider, 65, 68

in reptiles, 11 Garter snakes fungal infection in, 548 time course of initial inflammatory response in, 261 Gastrointestinal candidiasis, 541 Gastrointestinal mycoses, yeasts as causative agents of, 530 Gastrointestinal tract in American alligator, 32 effect of body temperature on, 12 viral-associated gastrointestinal diseases of snakes, 406 Gekkonidae, New Caledonian bumpy gecko, 124 GenBank, 359, 364 Gender differences, in lymphocyte counts, 181 Gene expression arrays, 374 advantages and disadvantages, 375 interpretation of results, pitfalls, limitations, 374–375 procedure, 374 schematic representation, 375 Gene splicing, 363 GeneChip arrays, 374, 375 Geotrichosis, 537 in corn snake, 566 Germ cell lines, mutations in, 363 Giant amelvia, mycetoma in, 551 Gila monster, 3 adenovirus infection in, 401 coccidioides infection in, 543, 570 filariasis in, 646, 647 microfilaria in, 212 skin shedding in, 36 venom glands in, 9 Gingivitis in Chelonia, 476 in Squamata, 476–477 Glomerulonephritis, 264 in Wagler’s viper, 287 Glutaraldehyde, as PSEM fixative, 301 Gnathostomatoidea, 588 Gomon methamine silver (GMS) stain, 229 Gonads, in reptiles, 14–15 Gopher snake cryptosporidiosis in, 283 granulomatous hepatitis in, 294 mycetoma in, 551 thyroid in, 244 Gopher tortoise anterior telencephalon in, 115 ascariasis in, 652 basophilic punctuate inclusions in, 195 brain and anterior spinal cord in, 112 cardiac puncture in, 190 caudal telencephalon in, 116 coelomic cavity in, 104 drying artifacts in RBCs, 170, 192 ELISA tests in, 146 fungal hyphae in, 553 heterophils in, 197 hindbrain in, 119 immature heterophils in, 199 iridovirus in, 334, 335, 404, 439 kidney in, 83 kidneys and adrenals in, 82 lymphocytes in, 175 mesencephalon in, 116 mycoplasma infection identified by EM, 321, 344 mycoplasmosis in, 502

694  Index

olfactory bulb in, 115 optic tectum in, 117 ovary in, 93 oxyurid egg in, 651 parathyroid gland in, 108 plasma cells in, 205 plasmacytoid lymphocytes in, 205 Ranavirus infection in, 440 Serratia infection in, 491 spinal cord in, 120 testes in, 87 thrombocyte clumps in, 193 thymus in, 108 Goussia, in Crocodylia, 576–577 Graft-versus-host-reaction (GVHR), 147 Gram stain, 229 Granular lymphocytes, in monitor lizard, 203 Granuloma formation in blood python, 276 in Caiman lizard, 276 in California red-sided garter snake, 276 chronic granulomas, 262 heterophilic granulomas, 261 histiocytic granulomas, 262, 277 in inflammatory response, 261 in lung of green turtle, 552 in Massasauga rattlesnake, 550 in mugger crocodile, 272 in mycobacteriosis of boa constrictor, 494 Grass snake, Hepatozoon in, 628 Gray patch disease, in green turtle, 422 Greek spur-thigh tortoise experimental herpesvirus infection in, 431 herpesvirus in, 288, 398 indirect ELISA for herpesvirus in, 387 reovirus infection in, 412 Green anaconda amyloid-like collagen deposits in, 287 shed spectacle behind eye in, 126 Green anole, Caryospora in, 623 Green eosinophils, 180, 201 Green iguana, 33 adrenal glands in, 88, 96, 110, 111 anterior telencephalon in, 116 azurophilic monocytes in, 201 basophils in, 201, 202 brain and anterior spinal cord in, 112 brain in, 121 caseous abscess in, 515 cerebellar cortex in, 118 coelomic cavity in, 66, 82 corpora albicans in, 100 corpora lutea in, 99 cryptosporidiosis in, 347, 626, 627 dermal pyogranuloma in, 492 dewlap in, 52, 53 diencephalon in, 122 epiphysis in, 55 erythrocyte inclusions in, 196 erythroplastids in, 195 esophagus in, 59 femoral pores in, 41, 42 gastric glands, 61 green eosinophils in, 201 heart in, 105, 241 hematologic values, 173

hemococcidia in, 184 heterophils in, 197 hindbrain in, 117 interparietal scales in, 121 kidneys in, 82, 83, 84, 243 liver in, 66, 69, 71 long bone in, 56 lung in, 77, 79 middle ear cavity in, 128 monocytes in, 205 necropsy exam of thyroid gland, 241 Neisseria infection in, 492 nonviral inclusions in, 185 olfactory nerves and bulb in, 114 optic tectum in, 117, 118 osteoarthritis in, 523 ovaries in, 94 ovary with multiple follicles in, 98, 99, 110 parathyroid glands in, 241 parietal eye in, 121 peroxidase activity in heterophils of, 258 pituitary gland in, 106 plasmacytoid lymphocytes in, 204 polychromatophils in, 195 previtellogenic follicle in, 98 proliferative osteoarthritis in, 263 Pseudomonas infection in, 488 reactive lymphocytes in, 204 right renal vein in, 88 sagittal section, nasal cavity, 114 seminiferous tubules in, 89 serological testing for paramyxovirus in, 388 skull bones in, 158 skull in, 47 small intestine in, 63 spermatozoa in, 90, 91 spleen in, 165 Splendore-Hoeppli reaction in, 264, 492 stomach, 61 Streptococcus infection in, 493 subconjunctival abscess in, 517 subgross sagittal section of head, 113 testes in, 88, 89, 90, 91 thrombocytes in, 208 thyroid gland in, 107 tongue in, 57 ultimobranchial body in, 109 urinary bladder in, 87 uterus in, 103 vitellogenic follicles in, 96, 98, 99 Green sea turtle, fungal dermatitis in, 554 Green tree monitor herpesvirus by ISH in, 380 herpesvirus in, 262–263, 433 herpesvirus infection in, 433 proliferative stomatitis in, 400 Green tree python pupil in, 127 Ranavirus infection in, 335, 405–406 subspectacular infection in, 517 Green turtle abscess in, 514 Chlamydia infection in, 345, 507 coccidiosis in, 618, 619 coelomic cavity in, 65, 77, 81 duodenum in, 73

Index  695

ELISA for spirorchidiasis in, 391 epiphysis in, 54, 55 fibropapilloma in, 428 fungal granulomas in lung of, 552 fungal hyphae in, 552 gray patch disease in, 422 heart in, 106 hematologic values, 172 herpesvirus-associated fibropapillomas in, 286, 426 herpesvirus infection in, 329 Learedius egg in, 640 leeches in, 659 LET disease in, 397, 422, 423, 424 liver with fungal nodules in, 552 lung in, 77 marine turtle fibropapilloma in, 425, 426, 427, 428 necropsy exam of, 250 necropsy internal exam of, 252 pectoral girdle in, 54 salt glands in, 126 spleen in, 73 stephanolepas in, 651 tapeworm in, 633 teeth in, 52 thymus in, 158, 159 thyroid gland in, 106 western blotting for herpesvirus in, 387 Guide trees, 365 Gular valve, in crocodilians, 9 Gut-associated lymphoid tissue (GALT), 9, 140

H Haemocystidium lygodactyli, in dwarf gecko, 608 Haemogregarina in Chelonia, 579 in Crocodylia, 579 in Rhynchocephalia, 579 in Squamata, 579–580 Haemoproteus, 580 Haitian boa, CNS disease in, 450 Half-life, of specific mRNA, 354 Haplotrema in Hawksbill sea turtle, 639 in loggerhead sea turtle, 639 Hard palate, in green turtle, 52 Hastospiculum varani in boa constrictor, 649 in monitor lizard, 649 Haversian bone, 7 Hawksbill sea turtle fibropapillomatosis in, 397 fungal dermatitis in, 557 Haplotrema in, 639 mandible in, 50 skull and mandible in, 45 Hazardous material, shipping precautions, 668 Head cross-section in corn snake, 114 in desert tortoise, 76 in Dumeril’s ground boa, 76 in green iguana, 113 Heart in American alligator with mycoplasmosis, 503 in boa constrictor, 104 in Burmese python, 245 cells in viper with paramyxovirus, 447

four-chambered in Crocodylia, 2 in green iguana, 66, 105 in green turtle, 106 location in reptiles, 18 in loggerhead sea turtle, 251 necropsy exam in lizards, 224, 241 in rhinoceros viper, 104 in snakes, 245 in timber rattlesnake, 105 in tortoises/turtles, 228, 251 Heart cells adenovirus-infected, 678 herpesvirus-infected, 677 from normal viper, 677 Hedge cells, 13 Helicobacter infections, 467 Helmeted iguana, coccidiosis in, 615 Helodermatidiae spp., gila monster, 36 Hemagglutination inhibition (HI) tests, 146, 383 Hemagglutination tests, 383 Hemagglutinin-neuramidase (HN) genes, in snakes, 409 Hematocrit, reference values for, 175 Hematologic values for crocodilians, 172 for lizards, 173 for tortoises, 171 for turtles, 172 Hematology blood sample collection and handling, 167–179 general concepts, 167, 176 procedures, 170–176 Hematology procedures, 170–174 erythrocyte evaluation, 174 leukocyte evaluation, 174–175 reference intervals for, 176 thrombocyte evaluation, 175 total protein and fibrogen evaluation, 175–176 Hematopoietic tissue, distribution of, 260 Hemipenes in corn snake, 92 in Eastern diamondback rattlesnake, 92 Hemiuridae, 584 Hemococcidia, 184 Hemoglobin concentration (Hb), 170, 174 Hemogregarines, 183, 608–609, 678 in American alligator, 628 in desert tortoise, 200 in Northern water snake, 628 Hemoparasites, 183 in desert tortoise, 200 hemococcidia, 184 hemogregarines, 183 microfilaria, 184 piroplasmida, 184 Plasmodium, 184 Sauroleishmania, 184 Spirorchiidae, 184 trypanosomes, 184 Hemoparasitic apicomplexa, 579 Haemogregarina, 579–580 Haemoproteus, 580 Hepatozoon, 579–580 Plasmodium, 580 Hemopoietic system, 19 Heparin, as anticoagulant, 179 Hepatic necrosis

696  Index

adenovirus-associated in lizards, 401–402 in bearded dragon, 401 in boa constrictors, 402 in San Esteban chuckwlla, 400 Hepatitis in bearded dragon, 295, 297 in gopher snake, 294 in Russell’s viper, 296 Hepatozoon in boa constrictor, 210, 609 in Chelonia, 579 in Crocodylia, 579 in Eastern indigo snake, 210 in Emerald tree boa, 629 in grass snake, 628 in Rhynchocephalia, 579 in Squamata, 579–580 in terrestrial and aquatic snakes, 183 Hermann’s tortoise cryptosporidiosis in, 620 differential response to antigens, 150 hematologic values, 171 herpesvirus in intestinal contents of, 398 herpesvirus infection, 431 herpesvirus infection identification by EM, 320 immunoglobulin studies, 143 immunological memory in, 147 indirect ELISA for herpesvirus in, 387 iridovirus infection in, 404 poxvirus infection in, 403 sarcocystis in, 620 seasonality and hormonal effects in, 148 temperature effects on immunity, 148 testes in, 87 virus X in, 415 Herpesviridae, 306 amplifying with degenerate primers, 357 in Argentine tortoise, 329, 430 in Chelonia, 396–399 in Crocodylia, 399 in desert tortoise, 318, 429 in European green lizard, 262, 432 in false map turtle, 329, 428 false map turtle in, 329 in freshwater turtles, 397–398 in Greek spur-thigh tortoise, 288, 431 in green tree monitor, 262–263, 433 in green turtle, 329, 426, 428 in Hermann’s tortoise, 431 in Hermann’s tortoise by EM identification, 320 host generation time, 364 identification by EM, 307–308 infected heart cell monolayer, 677 lacertid lizard papillomas, 399–400 in Mojave rattlesnake, 329 in Ophidia, 400 proliferative host responses to, 262–263 in San Esteban chuckwalla, 432 in Sauria, 399–400 in sea turtles, 396–397 serodiagnostics of, 382, 387 serology in marine turtles, 387 serology in tortoises, 387 by in situ hybridization in green tree monitor, 380 in situ hybridization techniques, 371 slow mutation rates of, 363

timing of viral shedding, 668 in tortoises, 398–399 venom gland herpesvirus, 400 Heteropenia, 180 Heterophilic gastritis, in viper species, 290 Heterophilic granulomas, 261, 277 in Caiman lizard, 276, 291 in Dumeril’s ground boa, 279 formation of, 261, 278 macrophages within, 261, 280 in mugger crocodile, 280 necrosis in, 261, 278 Heterophilic pneumonia, in Malaysian giant turtle, 271 Heterophilic steatitis, in rattlesnake, 291 Heterophils, 179–180 in American alligator, 197 in ball python, 197 in blood python, 202 in bone marrow of carpet python, 271 as functional equivalents to mammalian neutrophils, 179, 258 in gopher tortoise, 197 in green iguana, 197 in histologic section, 258 immunological functions in reptiles, 135 in initial inflammatory response, 260 in loggerhead sea turtle, 258, 271 resemblance in degenerate form to macrophages, 258, 271 in spiny turtle, 259, 271 toxic, 197 in water monitor acute bacterial infection, 275 Heterozerconidae, 595 Heuristic algorithms defined, 376 in maximum parsimony, 367 Monte Carlo simulation, 368 for sequence alignment, 365 Hexametiasis, in Home’s hingeback tortoise, 614 Hextoen enteric agar, 671, 680 Hibernation anemia during, 178 effect on lymphocyte counts, 181 Hindbrain in gopher tortoise, 119 in green iguana, 117 Hirudinea, 592 Histamine, 136 Histiocytic granulomas, 262 in boa constrictor, 277 in Kenya horned viper, 282 in liver of Emerald tree boa, 293 in McGregor’s tree viper, 281 in mugger crocodile, 291 progression of, 281 Histiocytic pneumonia, in loggerhead sea turtle, 292 Histiocytic steatitis, in rattlesnake, 291 Histology female reproductive system, 15–16 male reproductive system, 15 Histopathologic examination processing fees for, 221 tissue samples in necropsy, 230 Home’s hingeback tortoise, hexametiasis in, 614 Homogenization, of tissues for viral isolation, 670 Homology defined, 376 vs. homoplasy, 363

Index  697

Homoplasy, 362, 367 defined, 376 vs. homology, 363 Hormonal influences, on immune response, 148–150 Horned viper, esophagus in, 59 Horseradish peroxidase (HRP) use in direct ELISA, 383 use in immunoperoxidase testing, 384 Host response to infectious agents, 257–258 proliferative, 262–263 Humidity, and development of mycoses, 527–528 Humoral response, 132. See also Antibody response B-cells in, 136 Hyalohyphomycotic agents, 533 acremonium infections, 539 Aspergillus, 536–537 Beauveriosis, 538–539 CANV and other Chrysossporia, 533–534 chromoblastomycosis, 541 entomophthoromycoses, 538 fusariomycosis, 535–536 geotrichosis, 537 Metarhizium anisopliae, 538–539 miscellaneous agents, 539–540 mucormycoses, 537–538 paecilomycosis, 534–535 Penicillium, 536–537 zygomycosis, 537 Hydration status, indicators in necropsy, 224 Hypersensitivity response, in reptiles, 145 Hypovitaminosis A, differentiating from solid exudates, 260

I Iguanidae spp. brown anole, 43 green iguana, 33, 41, 42, 47, 52, 53, 55, 56, 57, 59, 61, 63, 66, 69, 71, 77, 79, 82, 83, 84, 87, 88, 89, 90, 91, 94, 96, 98, 99, 100, 103, 105, 106, 107, 109, 110, 111, 112, 113, 114, 116, 117, 118, 121, 122, 128, 158, 165 marine iguana, 51 Mexican spiny-tailed iguana, 96, 97, 98, 101, 102, 104 Immature erythrocytes, 176, 193–194 Immature heterophils, in green tortoise, 199 Immature lymphocytes, 181 in ball python, 205 Immature thrombocytes, 183 Immune complex-associated glomerulonephritis, 264 Immune response. See also Immunology differential response to type and amount of antigen, 150 effect of route of administration on, 150 factors affecting, 148–151, 392 hormonal influences on, 148–150 intrinsic factors, 150–151 maturation in reptiles, 144–145 seasonal influences on, 148–150 temperature effects on, 148 and vaccination, 151 Immunofluorescence test, 384 Immunoglobulins, 140–141 in avian species, 141 in Chelonia, 143–144 complement activation and, 145 in Crocodylia, 144 functions in reptiles, 145 in mammals, 141–143

and maturation of immune response, 144–145 in nonmammalian, nonreptilian vertebrates, 143 in Ophidia, 144 papain degradation, 142 pepsin degradation, 142 precipitating and agglutinating antibodies, 145 in reptiles, 143–144 and reptilian antibody neutralization, 145 in Rhynchocephalia, 144 in Sauria, 144 serological assays of, 382 valence and affinity in reptiles, 144 Immunohistochemistry, and in situ hybridization, 266–267 Immunolabeling, for electron microscopy, 303 Immunological memory, 147–148 Immunology, 131 general concepts, 131–132 innate defense mechanisms, 132–136 lymphocytes in, 136–137 specific defense mechanisms, 136–151 Immunoperoxidase test, 384 Immunostaining, fixation for PSEM, 302 In situ hybridization, 231, 370–371, 396 hybridization step, 372 limitations of, 372–373 probes in, 371–372 schematic representation, 370, 371 signal detection in, 372 tissue preparation for, 372 Inclusion body disease (IBD), 185 in boa constrictor, 212, 213, 339, 340, 450 of boid snakes, 410–411 in Burmese python, 452 infected boa constrictor kidney cells, 678 in Jamaican boa, 213 in liver of boa constrictor, 457 peripheral blood in boa constrictor with, 457 postmortem diagnosis criteria, 411 in rainbow boa, 212 serodiagnostics in boid snakes, 389–390 Incubation temperature, and sex ratios, 18 Incubation times for bacterial isolation, 671 for Histoplasma farciminosum, 672 Indel mutations, 362 defined, 376 Indian cobra, herpesvirus in, 400 Indian leaf-toed gecko macrophages studies in, 134 testosterone and estrogen-induced immune effects in, 150 thymic involution in, 149 Indian star tortoise, 36 penis everted in, 91 Indigo snake, azurophils in, 207 Indirect ELISA, 383 for cryptosporidiosis in snakes, 391 for herpesviruses in tortoises, 387 for iridovirus infection of chelonians, 388 for mycoplasmosis in chelonians, 390 for paramyxovirus in snakes, 388 for spirorchidiasis, 391 for West Nile virus in crocodilians, 389 Infectious agents bacteria, 185–186 detection in tissue section, 264–266 hemoparasites, 183–184

698  Index

host response to, 257–258 in peripheral blood, 182 viral inclusions in blood cells, 185 Inflammation abnormal heterophil morphology in, 179 amyloid-like material in, 264 chemical mediators of, 136 immune complex-associated glomerulonephritis, 264 lesions and tissue deposits in, 263–265 polymorphic thrombocytes in, 183 Splendore-Hoeppli reaction, 264 tissue responses secondary to, 263–264 Inflammatory cells. See Circulating inflammatory cells Inflammatory response, 260 granuloma formation in, 261–262 gross appearance of exudates in, 260 temperature effects on, 261 time course of, 260–261 Infrared detection organs, in reptiles, 24 Infundibulum, in Mexican spiny-tailed iguana, 102 Inhibitory mold agar, 672 Inland bearded dragon coccidiosis in, 616 hematologic values, 173 mesocestoides in, 634 Microsporidium infection in, 629, 630 Innate defense mechanisms, 131, 132 azurophilic monocytes, 136 basophils, 136 chemical mediators of inflammation, 136 eosinophils, 136 macrophage contributions to, 134 monocytes, 136 nonspecific cellular factors, 134–136 nonspecific humoral factors, 133–134 skin and mucosal surfaces, 132 surface barriers, 132–134 Inner ear, 23 Instrumentation, for sample handling, 668 Integumentary system biopsies for electron microscopy, 305 in reptiles, 4–6 Interferons, in reptiles, 133 Internal examination black and white tegu, 239 of blue panther chameleon, 240 in Cuban crocodile, 242 of lizards and crocodilians, 224–226 in necropsy, 224 of snakes, 226–227 of turtles and tortoises, 227–228 Intestinal cells, 10 Intranuclear coccidiosis, in radiated tortoise, 348, 617 Intranuclear inclusion bodies, 232 in boa constrictor, 288 in carpet python, 318 in Deckert’s rat snake, 321 in gray patch disease of green turtles, 422 in Greek spur-thigh tortoise, 264, 288 identifying with transmission electron microscopy, 306 Iridoviridae, 185, 404 in Chelonia, 404–405 in Eastern box turtle, 215, 216, 288 in European tortoises, 404 in gopher tortoise, 334, 335, 439 identification by EM, 308–309

in Japanese grass lizard, 214 in Lacertilia, 405 in Ophidia, 405–406 Ranavirus infection of green tree pythons, 405–406 serology in chelonians, 358 snake erythrocyte virus, 405–406 in soft-shelled Chinese turtles, 405 in U.S. tortoises and turtles, 404–405 Island night lizard Balantidium in, 610 mesocestoides in, 634 Nyctotherus in, 610 pituitary gland in, 106 Islets of Langerhans, 20 in corn snake, 74 in Eastern diamondback rattlesnake, 74 in monocled cobra, 74 in white-throated monitor, 75 Isospora in Chelonia, 575–576 in common house gecko, 622 in Crocodylia, 576–577 in flat-headed snake, 622 in lizards and snakes, 577 in yellow-headed gecko, 616 Ixodidae, 593 in Chelonia, 593–594 in Crocodylia, 594 in Squamata, 594 Ixodorhynchidae, 595

J Jackson’s chameleon abscesses in, 518 adenovirus infection in, 330, 331, 401 mycosis in, 537 Penicillium infection in liver of, 565 skull and mandible in, 48 teeth in, 51 Jacobson’s organ, in snakes, 3 Jamaican boa, IBD inclusions in, 213 Japanese encephalitis virus (JEV), 413 Japanese grass lizard Aegyptianella in, 609 LEV inclusions in, 214 piroplasmida in, 211 Plasmodium infection in, 608 Jeweled lacerta, cutaneous fungal disease in, 558 Johnson’s crocodile, crocodile pox in, 438 Joining fragment (J), 144 Jugular venipuncture, in chelonians, 169 Junctional complexes, in Eastern diamondback rattlesnake, 326 Juxtasplenic body, 12

K Kalicephalus in boa constrictor, 646 in Eastern king snake, 646 Karasberg tree skink, corpora lutea in, 100 Kemp’s ridley sea turtle, 539 kidney with fungal granuloma in, 568 Kenya horned viper bacterial splenitis in, 292 histiocytic granuloma formation in, 282 Keratin layer, absence in reptilian mucosa, 132

Index  699

Keyhole limpet homocyanin (KLH), 140 Kidneys in American alligator, 83 in bush viper, 85 cells from boa constrictor with IBD, 678 in corn snake, 81, 84 in Dumeril’s ground boa, 84 with eosinophilic intracytoplasmic inclusions in boa constrictor, 455 with fungal granuloma in Kemp’s ridley sea turtle, 568 in gopher tortoise, 82, 83 in green iguana, 82, 83, 84, 243 in green turtle, 77, 81 grinding for electron microscopy, 303 in loggerhead sea turtle, 253 necropsy examination of, 225, 242 with neoplastic tubules in lance-headed viper, 449 Penicillium infection in Aldabra tortoise, 564 in snakes, 246 in spitting cobra, 83 TEM of Eastern diamondback rattlesnake, 324 in tentacled snake, 81 variation in reptiles, 14 in water snake, 81 in Western diamondback rattlesnake, 82, 246 King cobra acanthocephalus in, 653, 654 Sphaerechinorhynchus serpenticola in, 653 King snake fungal infection of spectacle, 560 lung in, 79 nonviral inclusions in, 322 Kinin system, 136 Klossiella in boa constrictor, 627 in lizards and snakes, 579 Komodo dragon parthenogenesis in, 17 septic coelomitis in, 280

L Labeling, of samples for pathogen isolation, 668 Labial pits, in Emerald tree boa, 130 Laboratory fees, for necropsy, 220–221 Lacazia loboi, in chelonians, 539 Lacertid lizard, papillomas in, 399–400 Lacertilia, iridoviruses in, 405 Lacrimal gland, absence in snakes, 22 Lamellar bone, 7 Lamina bone, 8 Lance-headed viper kidneys with neoplastic tubules in, 449 paramyxovirus infection in, 407 renal cell carcinoma in, 410 retrovirus infection in, 343 Latent viruses, molecular evolution of, 364 Learedius egg, in green turtle, 640 Leatherback sea turtle, esophagus in, 59 Lectin pathway, of complement activation, 133–134 Leeches in American alligator, 658

in yellow-bellied slider, 658

in green turtle, 659 in loggerhead sea turtle, 658, 659 Left shifting, in heterophils, 179, 199 Leopard gecko

cryptosporidiosis in, 347, 625, 626 eye in, 124 precloacal pores in, 43 Leopard tortoise, 35 adenovirus reports in, 401 ascariasis in, 652 coccidiosis in, 617 virus X in, 415 Leptospira, 474 serodiagnostics of, 391 Leukocyte evaluation, 174–175 azurophils, 182 basophils, 180–181 in disease, 179 eosinophils, 180 heterophils, 179–180 lymphocytes, 181 monocytes, 181–182 plasma cells, 181 Leukocyte morphology, 174 Leukocytes acidophils, 258 azurophils, 259–260 basophils and mast cells, 259 effects of temperature on, 261 eosinophils, 258–259 heterophils, 258 host responses to, 258–260 macrophages, 259–260 monocytes, 259–260 unknown functions in reptiles, 257 Leukotrienes, 136 Light microscopy, in necropsy, 229 Likelihood, defined for molecular diagnostics, 376 Lipofuscin granules in Eastern diamondback rattlesnake, 326 electron micrography of, 307 Listeria infections, 466–467 Liver, TEM micrograph in Eastern diamondback rattlesnake324 in American alligator, 32, 65 in ball python, 66 with basophilic intracytoplasmic inclusions in IBD-infected boa constrictor, 457 in bothrops, 67 in Burmese python, 69, 246 candidiasis in, 541 close association with heart, 18 in cobra, 71 in desert tortoise, 68, 70, 72 in diamondback terrapin, 324 fungal infection in veiled chameleon, 293 fungal nodules in green turtle, 552 in green iguana, 66, 69, 71 in green turtle, 65 grinding for electron microscopy, 303 histiocytic granuloma in Emerald tree boa, 293 histiocytic hepatitis in bearded dragon, 297 with intracytoplasmic inclusions in boa constrictor, 454, 456 in Jackson’s chameleon with Penicillium infection, 565 in Kenya horned viper, 282 in Madagascan tree boa, 71 melanomacrophages in desert tortoise, 273, 327 in monocled cobra, 70 necropsy examination in snakes, 227 with necrosis in West Nile virus, 459 in neotropical rattlesnake, 70

700  Index

in prehensile-tailed skink, 346 in puff adder with Chlamydia infection, 508 in red-eared slider, 65, 69, 70 in red-footed tortoise with amoebiasis, 611 in reptiles, 11 in rhinoceros viper, 104 TEM micrograph in diamondback terrapin, 324 TEM micrograph in Eastern diamondback rattlesnake, 325 Lizard erythrocytic virus (LEV), 185, 405 in Fischer’s chameleon, 334, 441 in flap-neck chameleon, 334 Lizards adenovirus-associated hepatic necrosis of, 401 azurophils in, 182, 207 Caryospora in, 577 Cryptosporidium in, 578 dissection and internal examination in necropsy, 224–226 Eimeria and Isospora in, 577 femoral pores in, 5–6 Helodermatidae, 3 hematologic values, 173 hemococcidia in, 184 herpesvirus-associated oral lesions in, 400 immunoglobulin isotypes in, 144 Klossiella in, 579 nonhemoparasitic apicomplexa in, 577–579 precloacal glands in, 5 red cells in, 174 sarcocystis and Besnoitia in, 578–579 splenic anatomy in, 139 tail loss in, 7 thymus anatomy in, 138 viviparity in, 16 Lobomycosis, chelonians as experimental model for, 539–540 Loggerhead sea turtle adrenal gland in, 110, 252 Anasakis in, 643 ascariasis in, 643 bacterial pneumonia in, 291 barnacles in, 660, 661 brain removal in, 253, 254 Carettacola egg in, 640 cervical sinus venipuncture, 190 corpora lutea in, 100 epibiota accumulation, 224, 238 fibropapilloma in, 397 fungal myocarditis in, 556 fungal pneumonia in, 556 fungal tracheitis in, 289, 293 haplotrema egg in, 639 heterophils in, 258, 271 histiocytic pneumonia in, 292 kidneys in, 252 leeches in, 658, 659 neospirorchis egg in, 640 ovaries in, 95, 252 pericardium in, 251 salt glands behind eye, 125 Spirorchiidae in, 636, 637, 639 spleen and pancreas in, 251 spleen with red and white pulp in, 162 Sulcascaris egg in, 642 tapeworm in, 633 thymic granuloma formation in, 295 thymic involution in, 161 thymus in, 159, 251

thyroid in, 251 urinary bladder in, 86 uterine tube in, 102 vitellogenic follicles in, 100, 101 western blotting for herpesvirus in, 387 Long bones, in green iguana, 56 Long-branch attraction, 367 Lower respiratory tract, 12 Lung in American alligator with mycoplasmosis, 504, 553 bacterial embolus in water monitor, 295 in ball python, 245 in black and white tegu, 77 in boa constrictor, 104 in Burmese python, 78 in Burmese python with mycoplasmosis, 506 in Burmese python with thromboembolic disease, 522 in bush viper, 79 in corn snake, 78 in diamond python, 78 eye, and trachea (LET) disease, in green turtle, 397, 422, 423, 424 in gila monster with coccidioides infection, 570 in green iguana, 77, 79 in green turtle, 77 in green turtle with fungal granulomas, 552 with intracytoplasmic inclusions in boa constrictor, 453 in king snake, 79 origination of fungal infection in, 530 with proliferative pneumonia in caiman lizard, 443 as site of systemic mycoses, 531 in snakes, 227 surface area in reptiles, 13 in Western diamondback rattlesnake, 79, 284 Lung washes, for electron microscopy, 306 Luteinizing hormone (LH), 17 Lymph nodes, 140 absence in reptiles, 260 Lymphoblasts in lymphoid leukemia, 182 in lymphoma, 208 Lymphocytes, 136–137, 175, 181 functional B- and T-like activity in reptiles, 137 in gopher tortoise, 193 in pancake tortoise, 196 in plateau spiny lizard, 202 Lymphoid aggregations/accessory structures, 140 seasonal effects on, 148–149 Lymphoid cells, organ locations in reptiles, 260 Lymphoid follicular involution, in reptiles, 392 Lymphoid leukemia, in Aruba Island rattlesnake, 182 Lymphoid organs, 137 bone marrow, 137–138 spleen, 138–140 thymus, 138 Lymphoid tissue, distribution of, 260 Lymphoid tissue involution, seasonal, 149 Lymphoma, in yellow-footed tortoise, 208 Lymphopenia, 181 Lysosomes in Eastern diamondback rattlesnake, 326 electron micrography of, 307

M MacConkey’s agar, 671 bacterial isolation in, 679 Macronyssidae, 595

Index  701

Macrophages, 134–135 within heterophilic granulomas, 261, 280 in histologic section, 259–260 host responses to, 258–260 in mugger crocodile, 272 in progression of histiocytic granuloma, 281 respiratory bursts of, 135 Madagascan giant day gecko, Eimeria infection in, 622 Madagascan tree boa, liver in, 71 Madagascar common snake, Caryospora in, 623 Madagascar hognose snake, Caryospora in, 623 Major histocompatibility (MHC) molecules, 132 Malagasy spider tortoise, pancreatitis in, 289 Malarial parasites, 184 Malaysian giant turtle, heterophilic pneumonia in, 271 Male reproductive system, anatomy and histology, 15 Mandible in boa constrictor, 49 in common snapping turtle, 45 in desert tortoise, 40 in Eastern diamondback rattlesnake, 49, 50 in hawksbill sea turtle, 45, 50 in Jackson’s chameleon, 48 in lizards, 6 in Nile monitor, 48 in reticulated python, 49 Mangrove monitor lizard, azurophils in, 207 Mangrove snake, phaeohyphomycosis in, 540, 568 Map turtles die-off of captive, 397 Elizabethkingia infection in, 492 Marine iguana, teeth in, 51 Marine turtle fibropapilloma in, 425 herpesvirus serology in, 387 Marine turtle fibropapilloma, 425 in green turtle, 424, 425, 426, 427, 428 Massasauga rattlesnake facial fungal infection in, 538 fungal granulomas in, 550 Mast cells in Dumeril’s ground boa, 272 in histologic section, 259 Mathematical algorithms, for sequence alignment, 364 Mature erythrocytes, 176, 193 Maximum likelihood, 369 as tree-building method, 367–368 Maximum parsimony, as tree-building method, 367 McGregor’s tree viper chronic splenic granuloma in, 292 histiocytic granuloma formation in, 281 Meadow viper, stomatitis in, 518 Mean corpuscular hemoglobin concentration (MCHC), 174 Mean corpuscular hemoglobin (MCH), 174 Mean corpuscular volume (MCV), 174 during regenerative response, 177 Medical history, for necropsy, 223 Mediterranean gecko, Eimeria infection in, 615 Melanin granules, in desert tortoise, 328 Melanomacrophages, 134, 185 in Aldabra tortoise, 216 in boa constrictor, 217 in desert tortoise, 327 in Eastern indigo snake, 216 enlargement in pathology, 260, 273 in histologic section, 259–260

in liver of desert tortoise, 259, 327 in liver of fork-nosed chameleon, 273 Memory. See Immunological memory Meninges, in reptiles, 21 Meningitis, in rosy boa, 525 Mental gland, in desert tortoise, 41 Mesencephalon, in gopher tortoise, 116 Mesocestoides in Arizona mountain king snake, 634 in inland bearded dragon, 634 in island night lizard, 634 Mesos epidermal layer, 5 Messenger RNA, detection by northern blotting, 354 Metabolic rates, variations in reptiles, 12 Metarhizium anisopliae, 538–539 Metazoan parasites, tissue handling in necropsy for, 232–233 Methemoglobin, elevations in healthy reptiles, 177 Metropolis-coupled Monte Carlo Markov chain, 368 Mexican garter snake, renifers in, 635 Mexican spiny-tailed iguana atretic follicle in, 101 immunological memory in, 147 infundibulum in, 102 ovary in, 96 previtellogenic follicles in, 98 transplantation experiments, 146 vagina in, 104 vitellogenic follicles in, 97 Microbiology techniques, in necropsy, 231 Microfilaria, 184 in boa constrictor, 648 in gila monster, 212 in monitor lizard, 212 Microsporidia, 580–581 in bearded dragon, 349 identification by EM, 313 in inland bearded dragon, 629, 630 Mid-coelomic cavity in boa constrictor, 67 in corn snake, 67 in Western diamondback rattlesnake, 67 Midbrain, in reptiles, 20 Middle ear, 23 in green iguana, 128 Minimum evolution, defined, 376 Minimum evolution tree, 367 Mixed lymphocyte reaction (MLR), 146 evidence in reptiles, 147 Moellendorf rat snake, reovirus infection in, 412, 457, 458 Mojave rattlesnake adenovirus infection in, 402 herpesvirus infection in, 329, 400 reovirus infection in, 458 Molecular clock, 363 Molecular diagnostic tests, 351 array testing, 374–376 blotting techniques, 351–356 2D-PAGE, 373–374 gene-expression arrays, 374–375 glossary of terms, 376 molecular phylogeny, 361–370 in necropsy, 231 northern blot analysis, 352, 354 polymerase chain reaction, 356–361 protein arrays, 375–376 in situ hybridization, 370–373

702  Index

Southern blot analysis, 352, 353 vs. pathogen isolation, 667, 669 western blot analysis, 354–356 Molecular evolution, 362–364 Molecular phylogeny, 361–362 confidence interval measurement, 368–370 molecular evolution and, 362–364 sequence alignment in, 364–365 sequence selection for, 364 tree-building methods, 365–368 Molecular sequence data, 362 Monacled cobra, dracunculiasis in, 648 Monitor lizard granular lymphocytes in, 203 Hastospiculum varani in, 649 microfilaria in, 212 monocytoid leukocytes in, 206 Pentastomida in, 657 tanqua infection in, 650 thyroid glands in necropsy examination, 241 Monitors, in situ hybridization techniques, 371 Monocercomoniasis in corn snake, 615 in diamond python, 614 Monocled cobra adrenal gland in, 111 islets of Langerhans in, 74 liver in, 70 pancreas in, 74 Monoclonal antibodies defined, 376 for detection of mycoplasmosis, 390 in indirect ELISA, 383 in western blotting, 354, 356 Monocytes, 136, 181–182 in boa constrictor, 217 with Chlamydia inclusion, 182, 207 in desert tortoise, 205 in green iguana, 205 Monocytoid leukocytes, 182 in monitor lizard, 206 in red tegu lizard, 206 Mott cells, 181 Mucormycoses, 537–538 cell culture precautions, 672 in Florida soft-shelled turtle, 555 rareness in reptiles, 538 in veiled chameleon, 567 Mucosal surfaces, as immunological barriers, 132–133 Mugger crocodile bacterial bronchopneumonia with macrophages in, 259, 272 extrapulmonary bronchi in, 225, 242 granuloma formation in, 272 heterophilic granuloma in, 280 histiocytic granuloma in, 291 paratrichosoma infection in, 652 Multiplex PCR, 359 Musculoskeletal system, in reptiles, 6–8 Mutations, 362. See also Indel mutations; Molecular evolution; Recombination mutations; Substitution mutations advantageous vs. deleterious, 363 Mycetoma, 530 in giant amelva, 551 in gopher snake, 551 subcutaneous, 533 in Western diamondback rattlesnake, 551

Mycobacterium in American alligator, 289, 344 in Burmese python, 344 in desert tortoise, 344 detection in tissue section, 265 in gopher tortoise, 344 in heart of twin spotted rattlesnake, 292 identification by EM, 311 in mugger crocodile, 291 in rat snake, 495 in rattlesnake, 291 in reticulated python, 495 in rock rattlesnake, 495 with ulcerative mouth lesion in boa constrictor, 494 Mycobacterium infections, 468–469 Mycobiota, 529 in normal reptiles, 529 Mycobiotic agars, for fungal isolation, 680 Mycoplasmosis in American alligator, 503, 504, 505, 506 in Burmese python, 506 in Chelonia, 470–471 chelonian, 390 crocodilian serodiagnostics, 391 in Crocodylia, 471 in desert tortoise, 500, 501 in gopher tortoise, 502 in lung of American alligator, 553 in Ophidia, 472 serodiagnostics of, 390 Mycosel agar, 672 for fungal isolation, 680 Mycoses, 529 agents of, 533–543 in Chelonia, 531–532 and cold stunning phenomenon, 531 in Crocodylia, 532 dematiaceous fungi, 540–541 dermatomycoses, 529–530 dermatophyte agents of, 543 dimorphic fungi agents of, 543 hyalohyphomycotic agents of, 533–534 misdiagnosis as bacterial infections, 527 nomenclature of, 529 in Ophidia, 532–533 in reptiles, 530–533 in Sauria, 532 in Squamata, 532–533 underdiagnoses in reptiles, 527 yeast agents of, 541–542 Mycotic diseases, 527–528 dermatomycoses, 529–530 fungi overview, 528–529 mycoses, 529–530 in reptiles, 530–531 vs. normal mycobiota of reptiles, 529 Myxozoa, 581 in red-eared slider, 631 in yellow-spotted Amazon river turtle, 631

N Nasal cavity in desert tortoise, 76 in Dumeril’s ground boa, 76 in normal desert tortoise, 500, 501 sagittal section in green iguana, 114

Index  703

Natt-Herrick’s solution, 174, 192 for leukocyte counts, 174, 175 Natural cytotoxic cells (NCC), 135–136 Natural killer cells (NK), 132, 135–136 Nearly neutral mutations, 363 Necropsy biological measurements in, 223 equipment requirements, 222, 237 in fungal disease, 529 objectives, 220 rationale for, 219–220 standard form, 236 Necropsy reports, 222, 223–224, 233 electronic storage, archiving, and retrieval of, 233 standard form for, 236 Necropsy techniques, 219 archiving and retrieval of reports, 233 associated costs, 220–221 basics, 220 carcass preservation, shipping, disposal, 221–222 cleanup, 233 coding system for, 234 cytology, 228–229 data gathering, 223 dissection and internal examination, 224–228 documentation and description, 223–224 electronic storage, 233 external examination, 224 light microscopy, 229 microbiology, 231 molecular diagnostic tests, 231 necropsy components, 223–228 postnecropsy considerations, 233–235 precautions and zoonotic disease concerns, 233 routine tissue samples, 230 tissue archives, 233, 235 tissue handling for specific pathogen diagnosis, 231–232 toxicology, 231 transmission electron microscopy, 231 Necrotizing mycotic dermatitis, in snakes, 532 Necrotizing scute disease (NSD), 536 Negative predictive value (NPV), 386 Negative selection, 363 Negative staining transmission electron microscopy (NSEM), 299, 303 feces, aspirates, and washings for, 306 processing for, 303 Neighbor joining (NJ), 365, 366 defined, 376 Neisseria infections, 466 in green iguana, 492 Nematoda, 584–585. See also Microfilaria Ascaridoidea, 584–586 Diaphanocephaloidea, 586 Dioctophymatoidea, 588 Diplotriaenoidea, 587–588 Dracunculoidea, 587 Filarioidea, 586–587 Gnathostomatoidea, 588 Oxyuroidea, 589 Physalopteroidea, 588 Rhabditoidea, 588–589 Spiruoidea, 588 Trichinelloidea, 589 Neoechinorhynchus, in red-eared slider, 653 Neoplasms, retroviruses associated with, 410 Neospirorchis, in loggerhead sea turtle, 640

Neotropical rattlesnake liver and air sac in, 80 liver in, 70 paramyxovirus infection in, 337 pituitary gland in, 106 venom gland in, 58 vertebrae in, 158 Nephrons, in reptiles, 14 Nervous system, in reptiles, 20–21 Nested PCR, 359 additional amplification step in, 360 Neutral buffered 10% formalin, as PSEM fixative, 302 New Caledonian bumpy gecko, eye in, 124 New Guinea snakeneck turtle, spleen and pancreas in, 73 New methylene blue stain, 229 Newcastle disease virus, in Crocodylia, 407 Nick translation, 371 Nile crocodile adenovirus infection in, 401, 433 coccidiosis in, 621 coelemic cavity exposure in necropsy, 240 crocodile pox in, 437 erythrophagocytosis in, 323 hepatic necrosis in, 401 intralesional coccidia in small intestine of, 296 mycoplasmosis in, 391 necrotic heterophilic granuloma in, 278 nonviral inclusions in, 322 poxvirus in, 403 reproductive tract in, 242 retrovirus infection in, 410 TEM micrograph of spleen, 323 Nile monitor ascariasis in, 653 mandible in, 48 Pentastomida in, 656 skull in, 48 teeth in, 51 NJ algorithm, 366. See also Neighbor joining (NJ) Nocardia infections, 469 Nomarski interference-contrast photomicrographs, 622–623 non-cell-associated viruses, 669 Nonhemoparasitic apicomplexa, 575 Caryospora, 576, 577 in Chelonia, 575–576 in Crocodylia, 576–577 Cryptosporidium, 576, 577, 578 Eimeria and Isospora, 575–576, 576–577, 577 Goussia, 576–577 Klossiella, 579 in lizards and snakes, 577–578 sarcocystis and Besnoitia, 578–579 Nonlamellar bone, 7–8 Nonradioactive labels, in in situ hybridization, 371 Nonregenerative anemia, 178 Nonselective media, 672 for bacterial isolation, 671 Nonspecific cellular factors, 134 heterophils, 135 macrophages and dendritic cells, 134–135 natural cytotoxic cells (NCC), 135–136 natural killer cells (NK), 135–136 phagocytes, 134 phagocytic activation, 135 Nonspecific humoral factors, 133 complement, 133–134

704  Index

interferons, 133 lysozyme, 133 transferrin, 133 Nonvascular bone, 7 Nonviral inclusions in Deckert’s rat snake, 321 in desert tortoise, 323 in King snake, 322 in Nile crocodile, 322 Nonviral pneumonia, in viperid species, 285 Normal cells, EM ultrastructure of, 307 Northern blotting, 351, 352, 354 advantages and disadvantages, 354 interpretation of results, pitfalls, limitations, 354 procedure, 354 Northern water snake, hemogregarine infection in, 628 Nucleic acid detection, 351 Nucleic acid hybridization, 231 Nucleotide sequence alignment. See Sequence alignment Nutrition condition, indicators in necropsy, 224 Nyctotherus in desert tortoise, 610 in island night lizard, 610

O Oberhautchen, 6 in Dumeril’s ground boa, 39, 40 Ochetosomatidae, 583 Olfactory tracts in corn snake, 114, 115 in gopher tortoise, 115 in green iguana, 114 in reptiles, 20–21 Oligonucleotide probes, 371, 374 One-dimensional protein gel electrophoresis (1D-PAGE), 373 Oomphalitis, in ball python, 526 Ophidia, 3 adenoviruses in, 402–403 antibody response in, 146 blood sampling techniques in, 169 Boidae and Pythonidae families, 3 caliciviruses in, 414 Chlamydia infections in, 473–474 Dermatophilus infections in, 470 ear anatomy, 23 eye anatomy, 22–23 flaviviruses in, 414 immunoglobulins in, 144 iridoviruses in, 405–406 mycoplasmosis in, 472 mycoses in, 532–533 Parabasalia in, 575 paramyxoviruses in, 407–409 parvoviruses in, 406 picornaviruses in, 414 reoviruses in, 412 retroviruses in, 410–412 rhabdoviruses in, 414 togaviruses in, 413 Ophioptidae, 595–596 Ophiotaenia, in python, 632 Opisthocoelous vertebrae, 7 Opsonization, by antibodies, 145 Optic tectum in Dumeril’s ground boa, 118 in gopher tortoise, 117

in green iguana, 117, 118 Oral cavity, in Western diamondback rattlesnake, 57 Orbital sinus sampling, in turtles and tortoises, 168 Osmium tetroxide, 306 fixation of blood cells for EM in, 304 as PSEM fixative, 301 Osseous, in corn snake, 128 Osteoarthritis. See also Proliferative osteoarthritis in boa constrictor, 524 in Eastern hognose snake, 524 in green iguana, 523 noninflammatory, 263 in Russian rat snake, 523 Osteoarthrosis, in squamates, 263 Osteomyelitis in boa constrictor, 523, 524 in Chelonia, 478 in Crocodylia, 478 in Eastern hognose snake, 524 in Squamata, 478–479 Otic labyrinth, 23 in corn snake, 128 Otitis media, in wood turtle, 274 Ovaries in American alligator, 94, 96 in gopher tortoise, 93 in green iguana, 94, 98, 99 in loggerhead sea turtles, 95, 252 in Mexican spiny-tailed iguana, 96 in necropsy exam of crocodile, 242 in red-eared slider, 93 in reptiles, 15 in reticulated python, 94 in Siamese crocodile, 96 in veiled chameleon, 95 Overcrowding, and development of mycoses, 527 Overnight delivery services, for pathogen isolation samples, 668 Oviduct, in reptiles, 16 Oviparious reproduction, 16 in lizards, 3 Ovulatory surge, 17 Oxyuroidea, 589 in gopher tortoise, 651 in pancake tortoise, 651

P Packed cell volume (PCV), 174 Padlopper tortoise, Dermatophilus infection in, 496 Paecilomyces lilacinus, 535 Paecilomycosis, 534–535 Painted wood turtle circovirus infection in, 336, 407 eye in, 125 Spirorchiidae in, 638 Palatine vein, blood collection in Ophidia, 169, 191 Palestine viper, venom glands in, 58 Palm viper, adenovirus infection in, 402 Pancake tortoise erythrocyte mitosis in, 194 herpesvirus outbreak in, 399 lymphocytes in, 196 oxyurid egg in, 651 Pancreas in boa constrictor, 67 in Burmese python, 430 in corn snake, 67, 68, 74

Index  705

in Eastern diamondback rattlesnake, 74 in gaboon viper, 75 in genus Varamus, 12 with intracytoplasmic inclusions in boa constrictor, 454 islets of Langerhans, 20 in loggerhead sea turtle, 251 in monocled cobra, 74 necropsy examination of, 225, 242 necrosis in American alligator with West Nile virus, 413, 460 in New Guinea snakeneck turtle, 73 in Nile crocodile, 322 in red-eared slider, 73 in reptiles, 11 in rhinoceros viper, 74 in Savannah monitor, 75 in turtles/tortoises, 228 in white-throated monitor, 75 Pancreatitis in Aruba Island rattlesnake, 285 in Malagasy spider tortoise, 289 in speckled rattlesnake, 446 Panther chameleon, filariasis in, 647 Papain, immunoglobulin degradation, 142 Papillomaviridae, 406 in Bolivian side-neck turtle, 336, 442 in Chelonia, 406 identification by EM, 309 in Sauria, 406 Parabasalia, 574 in Chelonia, 574 in Ophidia, 575 Paraffin blocks, for long-term tissue sample storage, 235 Paraffin-embedded tissues, sample collection for electron microscopy, 305–306 Paramegistidae, 595 Paramyxoviridae, 407 in Aruba Island rattlesnake, 284, 336, 337, 444, 445, 446, 447 in Berg’s adder, 448 in Caiman lizard, 443, 444 in Caiman lizard by EM identification, 320 in Chelonia, 407 in Crocodylia, 407 cross-reactivity between isolates, 388 in diamond python, 447 in Eastern diamondback rattlesnake, 444, 446 in false tegu, 407 identification by EM, 309–310 infected viper heart cells, 678 in lance-headed viper, 407 in neotropical rattlesnake, 327 in Ophidia, 407–409 proliferative host responses to, 262 in rock rattlesnake, 407, 444, 446 in Sauria, 407 in serodiagnostics, 388–389 serological assays for, 382 in speckled rattlesnake, 446 in Vero cells, 447 Parasite exposure cryptosporidiosis, 391 incubation time, 572 serodiagnostics of, 391 spirorchidiasis, 391 Parasite identification cryptosporidium, 312–313 by electron microscopy, 312–313 laboratory fees for, 221

microsporidia, 313 in tissue section, 266 Parasites, tissue handling in necropsy for, 232–233 Parasitic diseases, 571–572 Acanthocephala, 590 Acari, 593–596 Annelida, 592 Cestoda, 581–582 Crustacea, 592–593 Diptera, 596–597 Microsporidia, 580–581 Myxozoa, 581 Nematoda, 584–589 Pentastomida, 590–592 protozoans, 572–580 Trematoda, 582–584 Parasitiformes Ascidae, 594 Diplogyniidae, 594 Entonyssidae, 594 Heterozerconidae, 595 Ixodidae and Argasidae, 593–594 Ixodorhynchidae, 595 Macronyssidae, 595 Paramegistidae, 595 Schizogyniidae, 595 Uropodidae, 595 Parasitology, laboratory fees for, 221 Parathyroid gland, 20 in Burmese python, 108 in desert tortoise, 108 in gopher tortoise, 108 necropsy exam in green iguana, 241 in veiled chameleon, 109 Paratrichosoma, in mugger crocodile, 652 Parietal eye in green iguana, 121 in tuatara, 122, 123 Parietal-pineal complex, in reptiles, 21 Pars pylorica, tubular glands in, 10 Parthenogenesis, in reptiles, 17–18 Parvoviridae, 406 identification by EM, 309 in Ophidia, 406 in Sauria, 406 viral-associated gastrointestinal diseases of snakes, 406 Pasteurella infections, 466 Pathogen identification of bacteria by EM, 311–313 with electron microscopy, 299–300, 306–313 by molecular diagnostics, 351 by in situ hybridization, 371 in tissue section, 257–258 and ultrastructure of normal cells, 307 of viruses by EM, 307–311 Pathogen isolation, 667 approach to, 669 bacterial isolation, 670–671 broth culture media for, 674 fungal isolation, 671–672 importance of time of collection, 668 sample collection for, 667–668 sample handling for, 667–669 sample reception for, 559 viral isolation, 669–670 PCR bands, 380

706  Index

PCR labeling, 371 Pectoral girdle, 30 development in reptiles, 7 in green turtle, 54 Pedal abscess, in American alligator, 275 Pelvic girdle in green turtle, 81 in red-eared slider, 53 remnants in ball python, 54 Penicillium, 536–537 in Aldabra tortoise, 564, 565 in Texas rat snake, 566 Penis in American alligator, 92 in Indian star tortoise, 91 Pentastomida, 590 in American alligator, 655, 657 in Chelonia, 590–591 in Crocodylia, 591 in gaboon viper, 655, 656, 657 in monitor lizard, 657 in Nile monitor, 656 in Squamata, 591–592 in water moccasin, 657 Pepsin, immunoglobulin degradation, 142 Peptide arrays, 375, 376 Periarteriolar lymphoid sheaths (PALS), 139 in red-eared slider, 164 Pericarditis, in American alligator, 289 Pericardium, in loggerhead sea turtle, 251 Periellipsoidal lymphoid sheaths (PELS), 139 in red-eared slider, 163, 164 Perilymphoid fibrous zone (PLFZ), 140 in Burmese python, 166 Periosteal bone, 8 Peripheral blood cells, in IBD-infected boa constrictor, 457 Phaehyphomycosis, 540–541 in mangrove snake, 540, 568 Phagocytes, 131, 134 activation of, 135 Pharyngitis in Chelonia, 476 in Squamata, 476–477 Phloxine B solution, 174, 175 Phosphate buffered saline (PBS), 669 Phylogenetic trees, confidence problems, 369 Physalopteroidea, 588 Pineal gland, in Dumeril’s ground boa, 123 Pinocytic vesicles, in Eastern diamondback rattlesnake, 326 Piroplasmida, 184 in Japanese grass lizard, 211 Pirornaviridae, 414 in Ophidia, 414 Pit organ in copperhead snake, 130 in crotalids, 24 in Western diamondback rattlesnake, 130 Pituitary gland in green iguana, 106 in island night lizard, 106 in neotropical rattlesnake, 106 in reptiles, 19 and squamate shedding, 5 Plagiorchiidae, 583 Plains garter snake, SEV infection in, 405, 442 Plasma cells, 181, 205

Plasma proteases, 136 Plasma samples, guidelines for handling, 675 Plasmacytoid lymphocytes, 181 in bearded dragon, 204 in gopher tortoise, 205 in green iguana, 204 Plasmodium, 183, 184, 580, 608–609 in Agama lizard, 210 in common agama, 608 in Japanese grass lizard, 608 in Puerto Rican anole, 608 in tegu lizard, 211 in turnip-tailed gecko, 608, 609 Plastron, removal in necropsy exam of turtles/tortoises, 227, 250 Plateau spiny lizard, lymphocytes and erythrocytes in, 202 Platelet-activating factor (PAF), 136 Pleurodont dentition, in lizards, 6 Plexiform bone, 8 Pneumonia. See also Bacterial pneumonia in loggerhead sea turtle, 291 nonviral, 285 in radiated tortoise, 285 Poikilocytosis, 177 Poly-T primers, 360 Polychromasia, in anemic reptiles, 177, 194 Polychromatophils, 176 in cottonmouth snake, 193, 194, 209 in green iguana, 195 Polyclonal antibodies defined, 376 for detection of mycoplasmosis in crocodilians, 391 in indirect ELISA, 383 in paramyxovirus-infected viper heart, 448 in western blotting, 354, 356 Polycythemia, 178–179 Polymerase chain reaction (PCR), 231, 356, 396 denaturation, annealing, and extension steps, 347–358 interpretation of results, 360–361 method, 357, 359 PCR bands, 380 reagents in, 356–357 results, 359 reverse-transcription-PCR (RT-PCR), 359–360 schematic representation, 357 use of poly-T primers in, 360 use of random primers in, 360 variations of, 359 Positive predictive value (PPV), 386 Positive selection, 363 Positive staining transmission electron microscopy (PSEM), 300 fixation for immunostaining, 302 fixatives and fixation, 301–302 glutaraldehyde as fixative, 301 infiltration and embedding, 302 neutral buffered 10% formalin as fixative for, 302 osmium tetroxide as fixative, 301 sectioning, staining and labeling for, 302–303 Trump’s solution as fixative, 301–302 Posterior medulla, in corn snake, 119 Posterior probability, 368, 370 defined, 376 Postmortem shears, 222 Postmortem specimens, sample collection for electron microscopy, 305 Power instruments, for necropsy use, 237 Poxviridae, 185, 403 in caimans, 403

Index  707

in Chelonia, 403 in Crocodylia, 403 in flap-necked chameleon, 207, 333, 403–404, 438 identification by EM, 308 in Johnson’s crocodile, 438 in Nile crocodile, 437 in Sauria, 403–404 in spectacled caiman, 332, 333 in tegus, 403 Prairie rattlesnake spleen in, 166 splenopancreas in, 162 Precipitating antibodies, 145 Precipitative fixatives, for in situ hybridization, 372 Precloacal glands in leopard gecko, 43 in lizards, 5 Preferred optimum temperature zone (POTZ), 4 Prehensile-tailed skink, liver TEM micrograph in, 346 Prehybridization step, in Southern blotting, 352 Previtellogenic follicles in green iguana, 98 in Mexican spiny-tailed iguana, 98 Primary antibody, use in electron microscopy, 303 Primary cell cultures, 669, 670 Primary vascular bone, 7 Prior probability, 368 Proatractis, in red-footed tortoise, 645 Probes, for in situ hybridization, 371–372 Procoelous vertebrae, 7 Proliferative gastritis in gopher snake, 283 in green tree monitor, 400 Proliferative host responses, 262 cryptosporidiosis, 262 in viral diseases and proliferative lesions, 262–263 Proliferative lesions, 262 Proliferative osteoarthritis, 263 in carpet python, 286 in green iguana, 263 Prosector, in necropsy, 220 Prostaglandins, 136 Protein arrays, 375 advantages and disadvantages, 376 interpretation of results, pitfalls, limitations, 376 procedure, 375–376 Protein databases, 359 Protein detection, with western blotting, 351, 354–356 Protein migration, in western blotting, 356 Proteomics, 373, 374 Proteus infection, in Cook’s tree boa, 519 Protocephalidea, 582 Protozoan parasites, 572 amoebae, 572–574 Euglenozoa, 594–595 nonhemoparasitic apicomplexa, 575–580 Parabasalia, 594–595 tissue handling in necropsy for, 232 Pseudohyphae, in yeast, 541 Pseudomonas infection in blood python, 488 in boa constrictor, 489 in Burmese python, 489 in green iguana, 488 in three-horned chameleon, 490 Pseudomonas infection in, in ball python, 490

Pseudomonas infections, 462–463 in Savannah monitor, 488 Pseudophyllidea, 581–582 Pterygosomatide, 596 Puerto Rican anole, Plasmodium in, 608 Puff adder, Chlamydia infection in, 345, 508, 509 Pyknotic erythrocytes, 176, 195 Pythonidae spp. ball python, 54, 66 Burmese python, 69, 78, 103, 108, 110, 113, 120, 160, 161, 166 diamond python, 78 green tree python, 127 reticulated python, 49, 60, 94, 105

Q Quantitations, in serodiagnostics, 385 Quick stains, 170

R Radiated tortoise, 34 coccidiosis in, 616, 617 intranuclear coccidiosis in, 348, 617 proliferative pneumonia in, 285 Radioactive labels, 372 in in situ hybridization, 371 Rainbow boa azurophils in, 207 Fusarium infection in, 535 IBD inclusions in, 212 Ranavirus infection, 404 in Eastern box turtle, 440, 441 in gopher tortoise, 440 in green tree python, 335 identification by EM in box turtle, 320 serological testing for, 388 Random primers, 360, 371 Rat snake amoebiasis in, 612 ascariasis in, 653 mycobacteriosis in, 495 Reactive lymphocytes in African spur-thigh tortoise, 203 in bearded dragon, 204 in blood python, 204 in green iguana, 204 in spur-thigh tortoise, 204 Reactive monocytes, 182 in African spurred tortoise, 206 in Aldabra tortoise, 206 Reactive thrombocytes, 183, 209 Reagents challenges for protein arrays, 376 in PCR, 356–357 Recognition of nonself, 131 Recombination mutations, 362, 363 Red-bellied water snakes, rhabdovirus infection in, 414 Red blood cell (RBC) count, 170, 176 Red blood cells (RBCs), interactions with viral antigens, 383 Red cell indices, 174 Red-eared slider, 30 axial skeleton in, 53 coelomic cavity in, 65 effects of temperature on inflammatory responses in, 261 erythroid regenerative response in, 177 evidence of dendritic cells in, 135

708  Index

exocine acinar cells in, 73 gallbladder in, 68 immune response to Salmonella, 132 immunoglobulin studies in, 144 liver in, 69, 70 myxozoan infection in, 631 Neoechinorhynchus in, 653 ovary in, 93 pancreas in, 73 pelvic girdle in, 53 periarteriolar lymphoid sheath in, 164 periellipsoidal lymphoid sheath in, 163, 164 spleen with red and white pulp in, 163 testes in, 87 uterine tube during vitellogenesis, 102 vitellogenic follicles in, 93 Red-footed tortoise amoeba in, 611, 612 Proatractis in, 645 Red neck disease, 405 Red pulp in Burmese python, 166 in green iguana, 165 in loggerhead sea turtle, 162 in prairie rattlesnake, 166 in red-eared slider, 163 in spleen, 139 Red tegu lizard monocytoid leukocytes in, 182 ruptured thrombocytes in, 208 Reference intervals, for hematology procedures, 176 Renal cell carcinoma, in lancehead vipers, 410 Renal vein, in green iguana, 88 Renifers in Eastern indigo snake, 635 in Florida banded water snake, 635 in Mexican garter snake, 635 Reoviridae, 412 in Chelonia, 412 in Moellendorf rat snake, 412, 457, 458 in Mojave rattlesnake, 458 in Ophidia, 412 proliferative host responses to, 262 in Sauria, 412 in serodiagnostics, 389 in viper heart 2 cell, 338 Reproductive cycle, 16–17 endocrine control and environmental influences, 17 parthenogenesis, 17–18 Reproductive system female anatomy and histology, 15–16 incubation temperature and sex ratios, 18 male anatomy and histology, 15 reproductive cycles, 16–17 in reptiles, 14–15 Reproductive tract, necropsy exam of, 242 Reptile anatomy cardiovascular system, 18–19 digestive system, 8–12 ear, 23 endocrine organs, 19–29 eye, 21–23 hemopoietic system, 19 infrared detection organs, 24 integumentary system, 4–6 musculoskeletal system, 6–8

nervous system, 20–21 reproductive system, 14–18 respiratory system, 12–13 review by system and organ, 4 salt glands, 24 urinary system, 13–14 vomeronasal organ, 24 Reptile biology, 1–2 extant taxonomic orders, 2–3 thermal, 4 Reptile gene sequences, lack of availability, 375, 376 Reptiles, as transitional group between amphibians and birds, 137 Residual infectivity, 382 Respiratory system in Burmese python, 245 in reptiles, 12–13 Restriction endonucleases, 361 Restriction fragment length polymorphism (RFLP), 359 Reticulated python dorsal aorta in, 105 esophageal tonsils in, 60, 455 evidence of dendritic cells in, 135 mycobacteriosis in, 495 ovary in, 94 skull and mandible in, 49 splenic anatomy in, 140 vitellogenic follicles in, 94 Retroviridae, 409 in boa constrictor, 319, 340, 341, 342 in Burmese python, 342 in Chelonia, 409 in clinically healthy snakes, 410 in Crocodylia, 409–410 host generation time vs. molecular evolution, 364 identification by EM, 310–311 inclusion body disease of boid snakes, 410–411 in lance-headed viper, 343 neoplasm-associated, 410 neurological disease of Australian pythons, 411–412 in Ophidia, 410–412 in Rhynchocephalia, 410 in Sauria, 410 Reversal lines, in bone remodeling, 263, 286 Reverse array, 375, 376 Reverse transcription-PCR (RT-PCR), 259–260 limitations of, 360 method, 360 Rhabditoidea, 588–589 in bushmaster snake, 650 in corn snake, 650, 651 Rhabdoviridae, 414 in caiman lizard, 414, 460 in Chelonia, 414 in Ophidia, 414 in Sauria, 414 Rhineuridae spp., Florida worm lizard, 33 Rhinoceros iguana bacterial inclusions in, 218 spiral-shaped bacterium in, 345, 512 Rhinoceros viper heart and liver in, 104 pancreas in, 74 tail vein venipuncture in, 191 Rhynchocephalia, 1, 3 amoebae in, 573 antibody response in, 145

Index  709

Haemogregarina in, 579 Hepatozoon in, 579 immunoglobulins in, 144 retroviruses in, 410 subclass Lepidosauria, 3 Rhytidodidae, 584 Ribbon snake, SEV infection in, 442 Ridgenose rattlesnake, Salmonella infection in, 491 River cooter, shell disease in, 513 RNA detection with northern blotting, 351, 352, 354 RNA stability issues, 354 stability limitations, 360 RNA integrity, for in situ hybridization, 373 RNA replication, 409 RNA viruses, 396, 415 Rock rattlesnake mycobacteriosis in, 495 paramyxovirus infection in, 407, 444, 446 Romanowsky-type stains, 170 eosinophils with, 180 for heterophils, 179 Rosy boa, meningitis in, 525 Rough endoplasmic reticulum, in Eastern diamondback rattlesnake, 325 Round cell tumor, in Burmese python, 410, 449 Route of administration, effects on immune response, 150 Royal Netherlands Academy of Arts and Sciences (KNAW), 672 Rubricytes, in cottonmouth snake, 194 Ruptured thrombocytes, in red tegu lizard, 183, 208 Russell’s viper amoebiasis in, 613 ascariasis in, 664 histiocytic enteritis in, 296 necrotizing hepatitis in, 296 Russian rat snake, osteoarthritis in, 523

S 16S ribosomal RNA gene, 361 Sabhi agar, 672 Sabouraud’s dextrose, 672 fungal isolation using, 680 Sacral vertebrae, in reptiles, 7 Salivary gland, granulomatous periadenitis in cottonmouth, 294 Salmonella infections, 463–465 cell culture for bacterial isolation, 680 pathogen isolation, 671 in ridgenose rattlesnake, 491 Salt glands in green turtle, 125 in loggerhead sea turtle, 125 in reptiles, 24 Saltwater crocodile, 31 Dermatophilus infection in, 496 fungal skin lesions in, 557 Sample collection from animals undergoing drug therapy, 668 from biopsies, 304–305 blood cells and cell cultures, 304 for electron microscopy, 304, 305 feces, aspirates, and washings for NSEM, 306 for formalin fixation, 255 importance of quantity for pathogen isolation, 668 mini sample collection set, 674 in necropsy, 220 in paraffin block for EM, 318 paraffin-embedded tissues, 305–306

for pathogen isolation, 667–668 postmortem specimens, 305 Sample conservation and shipping, for pathogen isolation, 668–669 Sample containers, for necropsy use, 237 Sample handling for pathogen isolation, 667 sample collection, 667–668 sample conservation and shipping, 668–669 sample reception, 669 Sample reception, for pathogen isolation, 669 San Esteban chuckwalla hepatic necrosis in, 400 herpesvirus infection in, 432 Sandwich antibody microarray, 375–376 Sarcocystis in Hermann’s tortoise, 620 in lizards and snakes, 578–579 Sauria, 3 adenoviruses in, 401–402 antibody response in, 145–146 blood sampling techniques in, 169 Chlamydia infections in, 473 Dermatophilus infections in, 470 ear anatomy, 23 eye anatomy, 22 flaviviruses in, 413–414 herpesviruses in, 399–400 immunoglobulins in, 144 mycoses in, 532 papillomaviruses in, 406 paramyxoviruses in, 407 parvoviruses in, 406 Poxviridae in, 403–404 reoviruses in, 412 retroviruses in, 410 rhabdoviruses in, 414 togaviruses in, 413 Sauroleishmania, 184 in flathead leaf-toed gecko, 609 in tree gecko, 211 Savannah monitor Pseudomonas infection in, 488 spleen and pancreas in, 75 Scales in Dumeril’s ground boa, 39 in green iguana, 121 Scanning electron microscopy (SEM), 304 coating, 394 fixation and processing, 304 in gopher tortoise and desert tortoise, 321 history of, 300 mounting, 304 Scanning transmission microscopy, 300 Schizangiella infection, in timber rattlesnake, 567 Schizogyniidae, 595 Schneider’s dwarf caiman necropsy exam of brain, 243 snout-vent length in, 238 Scincidae spp., Karasberg tree skink, 100 Scutes, 4, 34, 35 Sea turtles herpesvirus infection of, 396–397 prevalence of mycoses in, 531 Sear and stab technique, 231, 256 Seasonal influences, 137 on development of mycoses, 528

710  Index

on hematology values, 175 on heterophil counts, 180 on immune response, 148–150 on lymphocyte counts, 181 on spleen function, 140 on thymic involution, 138 Sebekia mississippiensis, in American alligator, 656 Secondary antibody, use in electron microscopy, 303 Selective media, for bacterial isolation, 671 Semidirect leukocyte count, 174, 175 Seminiferous tubules, 15 in desert tortoise, 90 in green iguana, 89 Senegal chameleon Dermatophilus infection in, 498, 499 testis in, 88 Sensitivity for indirect ELISA of herpesviruses, 387 of molecular diagnostics, 351 of PCR techniques, 356 in serodiagnostics, 386 of serodiagnostics, 384 of Southern blotting, 352 Septic coelomitis, in Komodo dragon, 280 Septicemia bacterial inclusions in, 185 in Dumeril’s ground boa, 279 in rhinoceros iguana, 186, 218 Sequence alignment bootstrapping in, 369 in molecular phylogeny, 364–365 Sequence selection, in molecular phylogeny, 364 Serial blood sampling, 170 Seroconversion, 381 factors influencing, 392 Serodiagnostics, 381–382, 667 for arboviruses, 389 for bacterial exposure, 390–391 for Coxiella, 391 for cryptosporidiosis, 391 enzyme-linked immunosorbent assay (ELISA), 383–384 establishing test cutoffs in, 384–385 factors affecting immune response in, 392 hemagglutination and hemagglutination inhibition, 383 for herpesviruses, 387 immunofluorescence test, 384 immunoperoxidase test, 384 for inclusion body disease of boid snakes, 389–390 for iridoviruses, 388 for Leptospira, 391 for mycoplasmosis, 390 negative predictive values in, 386 for paramyxovirus, 388–389 for parasite exposure, 391 pitfalls, 385 positive predictive values in, 386 quantitations in, 385 for reovirus, 389 sample collection and handling, 386 sensitivity in, 386 serum neutralization test (SNT), 382 specificity in, 386 for spirorchidiasis, 391 titers in, 385 titrations in, 385 types of serological assays, 382–385

for viral exposure, 387–390 western blot assay, 384–385 Serologic tests, immunology-based, 146 Serological assays. See Serodiagnostics Serotonin, 136 Serratia infections, 465 in gopher tortoise, 491 Serum neutralization test (SNT), 382 as gold standard for virus-specific antibody testing, 384 Sex ratios, and incubation temperature, 18 Sex segments in necropsy examination, 226, 240 in snakes, 227 in Western rattlesnake, 246 Sexual segment in bush viper, 85 in corn snake, 85 in snakes, 14 in spitting cobra, 85 Shed spectacle in Emerald tree boa, 126 in green anaconda, 126 Shell, anatomy in reptiles, 4 Shell disease of aquatic turtles, 475 in Bolivian side-necked turtle, 554 in river cooter, 513 Shell scrapings, in necropsy exam, 229 Shelled eggs, in reptiles, 1 Shipping of carcasses for necropsy, 221–222 guidelines for pathogen isolation samples, 668–669 Shipping container, for biological specimens, 675 Siamese crocodile, ovary in, 96 Sidewinder, cryptosporidial stomach infection in, 297 Sierra mountain king snake adenovirus and dendovirus in, 402, 443 adenovirus infection in, 435 parvovirus infection in, 406 Signal detection, for in situ hybridization, 372 Silent mutations, 363 Sinaloan milk snake, anal gland infection in, 516 Singapore house gecko, Streptococcus infection in, 493 Site of infection, collecting samples from, 667 Skin, as immunological surface barrier, 132 Skin shedding in boa constrictors, 37 in European grass snake, 38 Skull in American alligator, 46 in bearded dragon, 47 in boa constrictor, 49 in common snapping turtle, 45 in Eastern diamondback rattlesnake, 49, 50 in five-toed worm lizard, 46 in green iguana, 47, 158 in hawksbill sea turtle, 45 in Jackson’s chameleon, 48 in Nile monitor, 48 in reptiles, 6 in reticulated python, 49 Slider, Citrobacter infection in, 491 Small intestine with chronic granuloma in Emerald tree boa, 282 in desert tortoise, 63 in Dumeril’s ground boa, 63, 64

Index  711

in green iguana, 63 intralesional coccidia in Nile crocodile, 296 in red-footed tortoise with amoebiasis, 611 in reptiles, 10 up- and down-regulation of, 12 villi in desert tortoise, 63 Smooth endoplasmic reticulum, in Eastern diamondback rattlesnake, 325 Snake erythrocyte virus (SEV), 185, 405–406 in plains garter snake, 405, 442 in ribbon snake, 442 in terciopelo, 215 Snakes absence of urinary bladder in, 13 acute lymphocytic leukemia in, 182 adrenal glands in, 227 bone marrow of, 138 brain removal in, 227 Dermatophilus infections in, 470 dissection and internal examination in necropsy, 226–227 evidence of secondary immune response in, 147–148 eye anatomy in, 21 eye removal in necropsy examination, 249 faveolar parenchyma in, 13 heart in, 245 Hepatozoon in, 183 herpesvirus infection of boa constrictors, 400 immunoglobulin isotypes in, 144 isolation of adenoviruses from tissues of moribund, 402–403 kidneys in male, 14 liver and gallbladder in, 227 mixed viral-associated gastrointestinal diseases of, 400, 402 necropsy examination of, 226–227, 244 necrotizing mycotic dermatitis in, 532–533 nonhemoparasitic apicomplexa in, 577–579 palatine vein blood sampling in, 169, 191 retroviruses in clinically healthy, 410 serodiagnostics of IBD in, 389–390 in situ hybridization techniques, 371 splenic anatomy in, 139–140 thyroid gland necropsy examination in, 244 underdiagnosis of dermatomycoses in, 533 venom gland herpesvirus in, 400 viral-associated gastrointestinal diseases of, 406 viviparity in, 16 Snout-vent length, 238 necropsy measurements of, 223 Sodium heparin, 170 Soft-shelled Chinese turtles, iridovirus infection of, 405 Somatic hypermutation, in immunoglobulins, 142 Southern blotting, 351, 352 advantages and disadvantages, 352 capillary DNA transfer in, 353 comparison with PCR analysis, 352 interpretation of results, pitfalls, limitations, 352 procedure, 352 schematic representation, 353 Spatial resolution, of transmission electron microscopy, 299 Specialized oral glands, 9 Specific immune mechanisms, 136 in bacterial diseases, 151 cell-mediated immune responses, 146–147 factors affecting immune response, 148–151 immunoglobulins (antibodies), 140–146 lymphocytes, 136–137 lymphoid organs, 137–140 memory, 147–148

in parasitic diseases, 151 vaccination, 151 in viral diseases, 151 Specificity of indirect ELISA for herpesviruses in tortoises, 387 of molecular diagnostics, 351 in serodiagnostics, 386 of serodiagnostics, 384 of Southern blotting, 352 Speckled rattlesnake, paramyxovirus in, 446 Spectacle fungal infection in Eastern indigo snake, 560 fungal infection in King snake, 560 Spectacled caiman caiman pox in, 436, 437 poxvirus infection in, 332, 333 toxic heterophils in, 198 Spermatozoa in desert tortoise, 90 in green iguana, 90, 91 Sphaerechinorhynchus serpenticola, in King cobra, 653 Sphenodontidae spp., tuatara, 45, 46, 61, 122, 123 Spider tortoise, ascariasis in, 652 Spinal cord in American alligator, 112 in Burmese python, 113, 120 in death adder, 113 in Dumeril’s ground boa, 120 in gopher tortoise, 112, 120 in green iguana, 112 necropsy examination of, 226, 248 removal in Burmese python, 248, 249 Spiny-tailed iguanas serological testing for paramyxovirus in, 388 serological testing for reovirus in, 389 Spiny turtle, basophils and heterophils in, 271 Spiral-shaped bacterium, in rhinoceros iguana, 345, 512 Spirorchiidae, 184, 583 in Chelonia, 583–584 in chicken turtle, 636, 638 in loggerhead sea turtle, 636, 637, 639 in painted turtle, 638 serodiagnostics of, 391 Spiruroidea, 588 Spitting cobra kidney in, 83 sexual segment in, 85 Spleen in American alligator, 164, 165 in Burmese python, 166, 430 of Burmese python with round cell tumor, 449 with circovirus inclusions, 443 in corn snake, 67, 68 in desert tortoise, 162 in flap-necked chameleon with poxvirus, 404, 438 function and seasonal variation, 140 in gaboon viper, 75 granulopoiesis in Western diamondback rattlesnake, 274 in green anaconda, 287 in green iguana, 165 in green turtle, 73 grinding for electron microscopy, 303 gut-associated lymphoid tissue (GALT) and, 140 immunological role, 138–140 in loggerhead sea turtle, 162, 251 lymph nodes and other lymphoid structures and, 140

712  Index

lymphoid aggregations and accessory structures, 140 in McGregor’s tree viper, 281 necropsy examination of, 225, 242 necrosis in American alligator with West Nile virus, 459 in New Guinea snakeneck turtle, 73 in prairie rattlesnake, 166 as principle reptilian lymphoid organ, 260 in red-eared slider, 163 role in hemopoiesis, 19 in Savannah monitor, 75 TEM micrograph in Nile crocodile, 323 in white-throated monitor, 75 Splendore-Hoeppli reaction, 264 Splenic involution, 149 Splenopancreas in Aruba Island rattlesnake, 285 in black and white tegu, 242 in Kenya horned viper, 292 in McGregor’s tree viper, 292 in prairie rattlesnake, 162 Spur-thigh tortoise immature toxic heterophils in, 200 reactive lymphocytes in, 203, 204 toxic heterophils in, 198, 199 Spurr’s resin, 302 Squamata, 1, 3 Acanthocephala in, 590 amoebae in, 574 Amphisbaenia spp., 3 Argasidae in, 594 Ascaridoidea in, 585–586 bacteremia and osteomyelitis in, 478–479 bacterial pneumonia in, 477 Diplotriaenoidea in, 588 Diptera in, 597 Filarioidea in, 587 Haemogregarina in, 579–580 Hepatozoon in, 579–580 Ixodidae in, 594 miscellaneous bacterial infections in, 480 mycoses in, 532–533 Ophidia spp., 3 Pentastomida in, 591–592 proliferative osteoarthritis and osteoarthrosis in, 263 Sauria spp., 3 stomatitis, gingivitis, and pharyngitis in, 476–477 subcutaneous abscesses/masses in, 475–476 Squamate shedding, 5 Squamous metaplasia, as secondary inflammatory response, 263–264, 287 Staining techniques, for necropsy exams, 229 Standard deviation (SD), 386 Standard operating procedures (SOP), 667 Starvation, anemia due to, 178 Stephanolepas, in green turtle, 661 Stomach in Aldabra tortoise with fungal hyphae, 563 in American alligator, 32 in Burmese python, 246 in Dumeril’s ground boa, 61, 62 in green iguana, 61 pars pylorica in corn snake, 62 in reptiles, 10 in Western diamondback rattlesnake, 283 Stomatitis in Chelonia, 476 in Eastern box turtle, 518

in meadow viper, 518 in Squamata, 476–477 Streptococcus infections, 467 in green iguana, 493 in Singapore house gecko, 493 Stress, effects on immunity in Indian leaf-toed gecko, 134 Stringency, for in situ hybridization, 372 Strongyloides, in boa constrictor, 651 Stryker saw, 222, 237 Styphlodora, in tropical rat snake, 636 Styrofoam containers, 668 for pathogen samples, 675 Subcarapacial venipuncture, in Chelonia, 168 Subcutaneous abscesses/masses in bearded dragon, 514 in Chelonia, 475 in gaboon viper, 515 in Squamata, 475–476 Substitution mutations, 362 Substitution parameters, 368 Sulcascaris, in loggerhead sea turtle, 642 Supravertebral vessel, blood sampling from, 169, 190 Surface barriers, 132 skin and mucosal surfaces, 132 Surgical biopsy, laboratory fees for, 221 Swabs, for sample collection, 674 Syncytia formation, 232 Systematized nomenclature of medicine (SNOMED), 233 Systematized nomenclature of veterinary medicine (SNOVET), 233 Systemic mycoses, 529 in lung tissue, 531

T T-cell immunity, reptilian analogues, 137 T-cell receptor (TCR), 132 Tail loss, in lizards, 7 Tail vein venipuncture, 388 in rhinoceros viper, 191 Tanqua, in monitor lizard, 650 Tapeworm in false water cobra, 632 in green turtle, 633 in loggerhead sea turtle, 633 Target sequence, in Southern blotting, 352 Taxonomic orders Chelonia, 2 Crocodylia, 2–3 extant, 1, 2 Rhynchocephalia, 3 Squamata, 3 TBLASTX search tool, 359 Teeth in American alligator, 50 differences among reptiles, 6 in Jackson’s chameleon, 51 in marine iguana, 51 in Nile monitor, 51 in Tuatara, 51 Tegu lizard mature erythrocytes in, 193 Plasmodium in, 211 poxvirus infection in, 403 Teleomorphs, 528 Telidae spp., black and white tegu, 77 Temperate climates, penicillium species in, 536 Temperature

Index  713

effects on immune response, 132–133, 148 effects on inflammatory responses, 261 effects on phagocytic activity, 134 guidelines for shipping of pathogen samples, 675 importance for sample conservation, 66 Tentacled snake CANV in, 561 kidney in, 81 Terciopelo SEV inclusions in, 214, 215 viral inclusions in, 215 Test cutoffs, establishing in serodiagnostics, 384–385 Testes in corn snake, 89 in desert tortoise, 90, 91 in gopher tortoise, 87 in green iguana, 88, 89, 91 in Hermann’s tortoise, 87 in red-eared slider, 88 in Senegal chameleon, 88 Testosterone, immune effects in male Indian leaf-toed gecko, 149–150 Testudinae spp. desert tortoise, 30, 34, 35, 36, 40, 41, 57, 59, 63, 64, 68, 70, 72, 76, 86, 87, 90, 91, 108, 160, 162 gopher tortoise, 82, 83, 93, 104, 108, 112, 115, 116, 117, 119, 120 Hermann’s tortoise, 87 Indian star tortoise, 36, 91 leopard tortoise, 35 radiated tortoise, 34 Texas indigo snake Alaria infection in, 641 coccidioides infection in kidney of, 570 Texas rat snake cryptosporidiosis in, 624 Penicillium infection in, 566 TH-1 cells, infected and normal, 677 Thermal biology, in reptiles, 4 Three-horned chameleon, Pseudomonas infection in, 490 Thrombocyte clumps, 175, 193 in Chaco tortoise, 193 in gopher tortoise, 193 Thrombocytes, 182–183 in Aldabra tortoise, 208 in American alligator, 202 bacteria and, 185–186 in cottonmouth, 209 evaluation procedures, 175 in green iguana, 208 viral inclusions in blood cells, 185 Thromboembolic disease, in Burmese python, 521, 522 Thymic involution, 138, 149 in loggerhead sea turtle, 161 Thymic lobules in Burmese python, 160, 161 in desert tortoise, 160 Thymocytes, in Burmese python, 160 Thymus anatomical variations in reptiles, 138 in gopher tortoise, 108 granuloma formation in loggerhead sea turtle, 295 in green turtle, 158, 159 immunological role, 138 in loggerhead sea turtle, 159, 251 as principle reptilian lymphoid organ, 260 seasonal involution, 138 in turtles/tortoises, 227

in veiled chameleon, 161 Thyroid gland in boa constrictor, 107 in corn snake, 107 in gopher snake, 244 in green iguana, 107, 241 in green turtle, 106 in loggerhead sea turtle, 251 in monitor lizard, 241 necropsy exam in turtles/tortoises, 227 in necropsy examination, 225 in reptiles, 19–20 Timber rattlesnake adrenal cortical cells in, 111 cryptosporidiosis in, 624 entomophthoromycosis in, 538 fungal infection in, 567 heart base in, 105 Timor python, Bothridium in, 632 Tissue archives, post-necropsy handling techniques, 233, 235 Tissue cassettes, 229, 255 Tissue cutting board, 222 Tissue handling for bacteria and fungi, 232 for protozoan parasites, 232 for specific pathogen diagnosis in necropsy, 231 for viruses, 231–232 Tissue preparation, for in situ hybridization, 372 Tissue samples for histopathologic examination, 230 typical thickness, 229 Titer, in serodiagnostics, 385 Titrations, in serodiagnostics, 385 Togaviridae, 412 in Chelonia, 413 in Crocodylia, 413 identification by EM, 310 in Ophidia, 413 in Sauria, 413 serological testing for arboviruses in, 389 Tolucan lined ground snake, uterus in, 103 Tongue in American alligator, 52 in desert tortoise, 57 in green iguana, 57 variations in reptiles, 8–9 Tonic muscle fibers, in reptiles, 8 Tonsils. See also Esophageal tonsils in Dumeril’s ground boa, 60 esophageal, 9 in reticulated python, 60 in Western diamondback rattlesnake, 274 Tortoises, 22 adrenal glands in, 228 dissection and internal examination in necropsy, 227–228 hematologic values, 171 herpesvirus serology in, 387 herpesviruses in, 398–399 iridovirus infection in U.S., 404–405 orbital sinus sampling in, 168 postoccipital venous plexus blood sampling in, 168 reproductive cycles, 17 stomatitis-rhinitis and herpesvirus in, 399 Total protein, hematological evaluation, 175–176 Total red blood cell count (TRBC), 174 Touch impressions, 255

714  Index

in necropsy exam, 229 Toxic cell debris, elimination of, 133 Toxic heterophils, 179, 197 in American crocodile, 201 Toxicology laboratory fees for, 221 in necropsy, 231 Toxin exposure, anemia due to, 178 Trachea in black-tailed rattlesnake, 244 in gopher tortoise, 335 in loggerhead sea turtle with fungal tracheitis, 289 in snakes, 227 Trans-Pecos rat snake cryptosporidiosis in, 625 eye structures in, 127 Transferrin, as nonspecific humoral factor, 133 Transition, 362 defined, 376 Transmission electron microscopy (TEM), 299, 397 adenovirus in boa constrictor, 434 adenovirus in Nile crocodile, 433 Chlamydia infection in green turtle, 507 of Emerald tree boa with Chlamydia infection, 511 history of, 300 of Mojave rattlesnake with reovirus infection, 458 of mycoplasmosis in gopher tortoise, 502 in necropsy, 231 use in biopsies, 304 Transmission studies, establishing test cutoffs using, 384 Transversion, 362 defined, 376 Tree-building methods Bayesian posterior probabilities, 370 bias in, 369 distance-based trees, 365–367 maximum likelihood, 367–368 maximum parsimony, 367 in molecular phylogeny, 365 Tree gecko, Sauroleishmania in, 211 Tree monitor, Eimeria infection in, 622 Tree topology, 368 Trematoda, 582 Diplostomatidae, 584 Hemiuridae, 584 Ochetosomatidae, 583 Plagiorchiidae, 583 Rhytidodidae, 584 Spirorchiidae, 583–584 Trichinelloidea, 589 in Crocodylia, 589 Trichosporonosis, 542 in American anole, 569 Triplet codons, 363 Trombiculidae, 596 Tropical rat snake, Styphlodora in, 636 True positives, masking of, 371 Trump’s solution, as PSEM fixative, 301–302 Trunk vertebrae, in reptiles, 7 Trypanorhyncha, 582 Trypanosomes, 183, 184, 608–609 in crevice spiny lizard, 211 in turnip-tailed gecko, 608 Tuatara, 3, 46 absence of fungal disease in, 530 ear anatomy, 23

evidence of T-cell activity in, 137 immunoglobulin isotypes in, 144 parietal eye in, 122, 123 single-chambered lungs in, 13 skin shedding in, 5 skulls in, 6 teeth in, 51 tuatara, 33, 45 white and red splenic pulp in, 139 Tumors, of hematopoietic tissue, 182, 208 Turnip-tailed gecko, Plasmodium and trypanosomes in, 608, 609 Turtles, 22 adrenal gland in, 228 brain removal in, 228, 253 dissection and internal examination in necropsy, 227–228 family Dermochelyidae, 2 green sea turtle, 2 hawksbill, 2, 45 hematologic values, 172 iridovirus infections in U.S., 404–405 Kemp’s ridley, 2 loggerhead, 2 olive ridley, 2 orbital sinus sampling in, 168 shell disease of aquatic, 475 spleen and pancreas in, 228 uric acid excretion in, 14 Twin spotted rattlesnake, mycobacterial cardiac infection in, 292 Two-dimensional polyacrylamide gel electrophoresis (2D-PAGE). See 2D-PAGE

U Ulcerative enteritis, in common boa, 294 Ultimobranchial body in green iguana, 109 in reptiles, 20 Ultrafreezers, for long-term tissue storage, 235 Ultrametric, defined, 376 Ultrathin sections, in electron microscopy, 300, 302 University of Alberta Microfungus Collection and Herbarium, 672 Unweighted pair group matching using averages (UPGMA), 365, 366, 376 Upper respiratory tract, 12 mycoplasmosis in, 390 Uranyl acetate, 319, 322 immunolabeling for electron microscopy, 303 Urinary bladder absence in crododilians, 13, 14 absence in snakes, 3, 14 in desert tortoise, 86, 87 in green iguana, 87 in green turtle, 81 in loggerhead sea turtle, 86 Urinary system, in reptiles, 13–14 Uropodidae, 595 Uterus. See also Oviduct in Burmese python, 103 in Eastern diamondback rattlesnake, 103 in green iguana, 103 in loggerhead sea turtle, 102 in red-eared slider, 102 in Tolucan lined ground snake, 103

V Vaccination, and immune response, 151 Vacuolated thrombocytes, 183, 209

Index  715

Vagina in Mexican spiny-tailed iguana, 104 in reptiles, 16 Varanidae spp. Nile monitor, 48, 51 Savannah monitor, 75 white-throated monitor, 75 Vascular catheterization, for serial blood sampling, 170 Veiled chameleon, 43 abscess in, 518 CANV in, 532, 549, 550, 561 fusariomycosis in, 564 hepatic fungal infection in, 293 left lateral view, 78 mucormycosis in, 567 ovaries in, 95 parathyroid gland in, 109 periorbital abscess in, 517 skin micrograph, 44 thymus in, 161 Venom glands in beaded lizard and gila monster, 3, 9 in death adder, 58 herpesvirus infection of, 400 in neotropical rattlesnake, 58 in Palestine viper, 58 Venoms, chemical constituents, 9 Ventral tail vein venipuncture, in Sauria, 169, 191 Vero cell monolayer, 412, 447 in Aruba Island rattlesnake, 448 Vertebrae, 7, 8 in boa constrictor, 286 in Eastern diamondback rattlesnake, 158 in neotropical rattlesnake, 158 Vestibulum nasi, in chelonians, 12 Veterinary diagnostic laboratories, 220, 221 Veterinary pathologists, 224 necropsy diagnostics by, 220 Vibrio infections, 466 Viperidae spp. adenovirus infection in, 402 Aruba Island rattlesnake, 80 boa constrictor, 67 bush viper, 79, 85 copperhead snake, 130 Eastern diamondback rattlesnake, 49, 50, 74, 82, 92, 103, 158 fer-de-lance, 67 gaboon viper, 75, 111 heart cells infected with paramyxovirus, 447, 449 heterophilic gastritis in, 290 horned viper, 59 Naja spp., 111 neotropical rattlesnake, 58, 70, 80, 106, 158 Palestine viper, 58 prairie rattlesnake, 162, 166 rhinoceros viper, 74, 104 timber rattlesnake, 105, 111 Western diamondback rattlesnake, 57, 67, 79, 130 Viral diseases, 395–396. See also Viruses detection in tissue section, 264–265 herpesviruses, 262–263 paramyxoviruses and reoviruses, 262 proliferative host responses in, 262 Viral inclusions in blood cells, 185 in erythrocytes, 178

inclusion body disease (IBD), 185 iridovirus, 185 poxvirus, 185 Viral infection anemia pursuant to, 178 basophilia with, 181 Viral isolation, 669–670 adenovirus in viper heart cells, 678 paramyxovirus-infected viper heart cells, 678 tissue culture medium and enrichments for, 676 tortoise herpesvirus, 677 Viral latency, in tortoises, 399 Viral taxonomy, revision due to molecular speciation, 362 Viral titers decrease with continued storage, 669 loss through filtering, 670 Virus assembly pools/factory areas, 405 Virus X, 415 Viruses Adenoviridae, 401–403 Adenoviridae identification by EM, 308 Calciviridae, 414 Circoviridae, 406–407 Circoviridae identification by EM, 309 Coronaviridae, 415 Flaviviridae, 413–414 Flaviviridae identification by EM, 310 Herpesviridae, 396–400 Herpesviridae identification by EM, 307–308 identification by EM, 307–311 Iridoviridae, 404–406 Iridoviridae identification by EM, 308–309 miscellaneous, 415 non-cell-associated, 669 Papillomaviridae, 406 Papillomaviridae identification by EM, 309 Paramyxovidiae, 407–409 Paramyxovidiae identification by EM, 309–310 Parvoviridae, 406 Parvoviridae identification by EM, 309 Picornaviridae, 414 Poxviridae, 403–404 Poxviridae identification by EM, 308 Reoviridae, 412 Retroviridae, 409–412 retrovirus identification by EM, 310–311 Rhabdoviridae, 414 tissue handling in necropsy for, 231–232 Togaviridae, 412–413 Togaviridae identification by EM, 310 Visceral necrosis, in Sauria with herpesvirus, 400 Vitellogenic follicles in American alligator, 95, 97 in corn snake, 94 in green iguana, 96, 98, 99 in loggerhead sea turtle, 100, 101 in Mexican spiny-tailed iguana, 97 in red-eared slider, 93 in reticulated python, 94 Viviparous reproduction, in lizards and snakes, 16 Vomeronasal organ (VNO) in corn snake, 129 in reptiles, 24

W Wagler’s viper, glomerulonephritis in, 287

716  Index

Washings, sample collection for NSEM, 306 Water moccasin fungal infection in, 549 pentastomiasis in, 657 Water monitor bacterial embolus of lung in, 295 heterophils in acute bacterial infection, 275 Water snake ascariasis in, 644 colon in, 81 kidneys in, 81 mid-coelomic cavity in, 81 retrovirus-like particles associated with RBC inclusions in, 410 West Nile virus in American alligator, 338, 459, 460, 3898 in Crocodylia, 413 serodiagnostics in crocodilians, 382, 389 Western blotting, 351, 354 advantages and disadvantages, 356 antibody detection in, 355 electrophoretic protein transfer in, 355 interpretation of results, pitfalls, limitations, 356 for marine turtle herpesviruses, 387 procedure, 354, 356 schematic representation, 355 for serodiagnostics, 384–385 Western diamondback rattlesnake chronic granuloma in mesentery of, 295 granulopoiesis in spleen of, 274 kidney in, 82, 246 liver in, 67 lung in, 79 lymphoid tonsils in, 274 mycetoma in, 551 normal lung in, 284 normal stomach in, 283 pit organ in, 130 tongue in, 57 Western equine encephalitis (WEE) virus, 396, 413 White blood cell (WBC) count, 170, 174 White pulp in Burmese python, 166 in Caspian turtle, 148–149

in green iguana, 165 in loggerhead sea turtle, 162 in prairie rattlesnake, 166 in red-eared slider, 163 in spleen, 139 White-throated monitor, pancreas in, 75 Wood turtle amoebiasis in, 612 Eimeria infection in, 616 exudative otitis media in, 274, 275 squamous metaplasia in middle ear, 546 Worm lizards, 3 five-toed, 33, 46 Florida worm lizard, 33 forelimb-only, 7 Wound healing, thrombocytes in, 183

X Xantusiidae spp., island night lizard, 106 Xenografts, cell-mediated immune responses to, 146–147

Y Yeast, 528, 541 candidiasis, 541–542 cryptococcosis, 542 difference from molds, 541 and gastrointestinal mycoses, 530 trichosporonosis, 542 Yellow-bellied slider, leeches in, 658 Yellow-footed tortoise, lymphoma in, 208 Yellow fungus disease, in bearded dragon, 562 Yellow-headed gecko, Isospora infection in, 616 Yellow-spotted Amazon river turtle, myxozoan infection in, 631 Yellow tortoise, pyknotic erythrocytes in, 195 Yolk sac, in Dumeril’s ground boa, 290

Z Zip fastener model, of epidermis, 5 Zoonotic disease, necropsy concerns, 233 Zygomycosis, 537 Zygopophyses, 7