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.

344  Identifying Reptile Pathogens Using Electron Microscopy

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.)

Identifying Reptile Pathogens Using Electron Microscopy  345

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.

346  Identifying Reptile Pathogens Using Electron Microscopy

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.

Identifying Reptile Pathogens Using Electron Microscopy  347

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.

348  Identifying Reptile Pathogens Using Electron Microscopy

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.)

Identifying Reptile Pathogens Using Electron Microscopy  349

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 Cons