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Iridovirus Infections of Captive and Free-Ranging Chelonians in the United States

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 Title Page
 Acknowledgement
 Table of Contents
 List of Tables
 List of Figures
 Abstract
 Introduction
 Ranavirus infection of free-ranging...
 Development and use of an indirect...
 Experimental transmission of a...
 The role of infected leopard frogs...
 In vitro efficacy of acyclovir...
 Conclusions and future researc...
 References
 Biographical sketch
 

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IRIDOVIRUS INFECTIONS OF CAPTIV E AND FREE-RANGING CHELONIANS IN THE UNITED STATES By APRIL JOY JOHNSON A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2006

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Copyright 2006 by April Joy Johnson

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ACKNOWLEDGMENTS Funding for this work was provided in part by the University of Florida College of Veterinary Medicine Batchelo r Foundation, grant No. D04Z0-11 from the Morris Animal Foundation, and a grant from the Disney Conser vation Fund. I also thank the University of Florida, College of Vete rinary Medicine for awarding me an Alumni Fellowship for tuition and stipend over the first three years. I would like to thank the many people who helped collect and submit turtle and tortoise samples including Dr. William Belzer Dr. Terry Norton, Jeffrey Spratt, Valorie Titus, Susan Seibert and Ben Atkinson. Also, I would like to thank all the pathologists who contributed to this work including Dr. Allan Pessier, Dr. Nanc y Stedman, Dr. Robert Wagner and Dr. Jason Brooks. I thank Drs. Jerry Stanley and Kathy Goodblood for allowing me access to chelonian and amphibi an populations at Buttermilk Hill Nature Sanctuary and to Drs. Mick Robinson and Ken Dodd for providing necropsy reports on archived cases. I would also like to thank the Mycoplasma Research Laboratory at the University of Florida including Dr. Ma ry Brown, Dr. Lori Wendland and Dina Demcovitz for providing access to their wild gopher tortoise plasma bank. I thank Yvonne Cates at the Zoological Society of San Diego for excellent histology support and Lynda Schneider from the University of Florida Electron Microscopy Core Laboratory. I also thank Dr. Harvey Ramirez, Rachelle Wright and Kevin Chadbourne with Animal Care Services for excellent ca re of the turtles and frogs. iii

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Several people went above and beyond norma l expectations in assisting with this research in various ways. I would especially like to thank Dr. Allan Pessier for all his time, patience and assistance with necropsie s and histopathology as well as making me a better scientific writer. I thank Dr. Jim We llehan for all his advice and assistance with molecular and laboratory techniqu es. I also am indebted to our laboratory technicians, Sylvia Tucker and April Childress. They both provided invaluable laboratory support and became great friends throughout my time at the University of Florida. I thank my parents for their support a nd encouragement, without which I would never have made it to this point in my career. They have always believed in me and, although they might not always understand my unique interests, have never discouraged me from pursuing my goals. For th at I will always be thankful. And last, but far from least, I am extremely grateful to my supervisory committee members: Dr. Elliott Jacobson, Dr. David Bl oom, Dr. Jack Gaskin, Dr. Jorge Hernandez and Dr. Gail Scherba. I thank them for th e advice, support and encouragement they have given me since the beginning. I thank them for helping me develop my skills and for their confidence in my abilities. I have lear ned so much from them and have thoroughly enjoyed this opportunity to work with them. iv

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TABLE OF CONTENTS page ACKNOWLEDGMENTS.................................................................................................iii LIST OF TABLES...........................................................................................................viii LIST OF FIGURES...........................................................................................................ix ABSTRACT......................................................................................................................x ii CHAPTER 1 INTRODUCTION........................................................................................................1 2 RANAVIRUS INFECTION OF FR EE-RANGING AND CAPTIVE BOX TURTLES AND TORTOISES IN THE UNITED STATES.....................................11 Introduction.................................................................................................................11 Materials and Methods...............................................................................................12 Animals................................................................................................................12 Necropsy and Histopathology.............................................................................16 Nucleotide Amplification, Seque ncing, and Sequence Analysis........................16 Virus Isolation.....................................................................................................17 Transmission Electron Microscopy.....................................................................18 Restriction Enzyme Analysis..............................................................................18 Results.........................................................................................................................19 Necropsy and Histopathology.............................................................................19 PCR and Sequence Analysis...............................................................................20 Virus Isolation.....................................................................................................21 Transmission Electron Microscopy.....................................................................21 Restriction Enzyme Analysis..............................................................................21 Discussion...................................................................................................................22 3 DEVELOPMENT AND USE OF AN INDIRECT ENZYME LINKED IMMUNOSORBENT ASSAY FOR DETECTION OF IRIDOVIRUS EXPOSURE IN GOPHER TORTOISES ( Gopherus polyphemus )...........................34 Introduction.................................................................................................................34 Materials and Methods...............................................................................................36 Virus....................................................................................................................36 v

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Antigen Preparation.............................................................................................37 Positive and Negative Reference Plasma............................................................38 ELISA Procedure.................................................................................................39 Experimentally Inoculated Turtles......................................................................40 Reproducibility....................................................................................................41 Protein Expression and Immunoblotting.............................................................42 Wild Gopher Tortoises Samples with Unknown Exposure.................................43 Results.........................................................................................................................43 Antigen Preparation.............................................................................................43 ELISA Parameters...............................................................................................44 Experimentally Inoculated Turtles......................................................................44 Reproducibility....................................................................................................45 Protein Expression and Immunoblotting.............................................................45 Wild Gopher Tortoises Samples with Unknown Exposure.................................46 Discussion...................................................................................................................46 4 EXPERIMENTAL TRANSMISSION OF A RANAVIRUS IN WESTERN ORNATE BOX TURTLES ( Terrapene ornata ornata) AND RED-EARED SLIDERS ( Trachemys scripta elegans )......................................................................65 Introduction.................................................................................................................65 Materials and Methods...............................................................................................66 Experimental Animals and Husbandry................................................................66 Pre-inoculation Sample Collection......................................................................67 DNA Preparation, Polymerase Chain R eaction and Nucleotide Sequencing......67 ELISA..................................................................................................................68 Virus Preparation.................................................................................................70 Transmission Studies...........................................................................................71 Study 1..........................................................................................................71 Study 2..........................................................................................................73 Results.........................................................................................................................74 Experimental Animals and Pre-inoculation Sampling........................................74 Transmission Studies...........................................................................................74 Study 1..........................................................................................................74 Study 2..........................................................................................................75 Discussion...................................................................................................................80 5 THE ROLE OF INFECTED LEOPARD FROGS ( Rana pipiens) IN TRANSMISSION OF A RANAVIRUS IN RED-EARED SLIDERS ( Trachemys scripta elegans) .........................................................................................................102 Introduction...............................................................................................................102 Materials and Methods.............................................................................................104 Virus Preparation...............................................................................................104 Frog Pilot Study.................................................................................................105 Full Frog Study and Pre-inoculation Sampling.................................................106 Turtles................................................................................................................107 vi

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Virus Titers........................................................................................................109 ELISA................................................................................................................110 Results.......................................................................................................................111 Frog Pilot Study.................................................................................................111 Full Frog Study..................................................................................................112 Turtles................................................................................................................112 Virus Titers........................................................................................................113 ELISA................................................................................................................113 Discussion.................................................................................................................114 6 IN VITRO EFFICACY OF ACYCL OVIR AS A POTENTIAL THERAPEUTIC AGENT FOR IRIDOVIRUS INFECTIONS IN CHELONIANS............................125 Introduction...............................................................................................................125 Materials and Methods.............................................................................................127 Cell Cultures......................................................................................................127 Virus..................................................................................................................128 Acyclovir and Concentrations...........................................................................128 Cytotoxicity Assays...........................................................................................129 Cytopathic Effects (CPE) Reduction Assays....................................................129 Virus Titer Reduction Assays............................................................................130 Results.......................................................................................................................130 Discussion.................................................................................................................130 7 CONCLUSIONS AND FUTURE RESEARCH......................................................135 LIST OF REFERENCES.................................................................................................139 BIOGRAPHICAL SKETCH...........................................................................................149 vii

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LIST OF TABLES Table page 3-1 Reproducibility of the ELISA. .................................................................................53 3-2 ELISA results of 1000 free-ranging gopher tortoise ( Gopherus polyphemus ) plasma samples by county and state. ........................................................................53 3-3 ELISA results of 658 free -ranging gopher tortoises ( Gopherus polyphemus ) from the state of Florid a are listed by region. ..........................................................55 3-4 ELISA results of 1000 free-ranging gopher tortoises ( Gopherus polyphemus ) listed by state. ...........................................................................................................55 4-1 PCR results on tissues collected at necropsy from the pi lot study box turtles (BT; Terrapene ornata ornata) and red-eared sliders (RES; Terrapene scripta elegans) in the full transmission studies. .................................................................88 4-2 Polymerase chain reaction (PCR) resu lts of oral and cloacal swabs taken on eleven different days post-inoculation (DPI) ...........................................................89 4-3 PCR results for urine collected opportunistically from turtles in the full transmission study. ...................................................................................................90 5-1 Results of the pilot frog study. ...............................................................................119 6-1 Effect of increasing doses of acy clovir on cytopathic effect reduction, cytotoxicity and TCID50 of Terrapene heart cells inoculated with BSTRV. .........134 viii

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LIST OF FIGURES Figure page 2-1 Gross lesions associated with iridoviru s infections in turtles and tortoises.............26 2-2 Esophagus, eastern box turtle ( Terrapene carolina carolina ). There is diffuse necrosis and ulceration of the mucosa and replacement by fibrin, inflammatory cell infiltrates and superfi cial bacterial colonies......................................................27 2-3 Spleen, eastern box turtle ( Terrapene carolina carolina ). There is disruption of the white and red pulp with deposits of fibrin (arrow) admixed with karyorrhectic debris, and infiltrates of small numbers of heterophils......................28 2-4 Epicardium, Burmese star tortoise ( Geochelone platynota ). Arrows depict basophilic intracytoplasmic inclusion bodies in a macrophage and endothelial cell........................................................................................................................... .29 2-5. Results of a polymerase chain reaction targeting approximately 500 bp of the major capsid protein gene........................................................................................30 2-6 Transmission electron photomicrograph of Terrapene heart cells inoculated with liver tissue from a Burmese star tortoise ( Geochelone platynota ) demonstrating cytoplasmic arrays of ir idovirus-like particles.........................................................31 2-7 Transmission electron photomicrograph of paraffin embedded spleen from a box turtle ( Terrapene carolina ) that died in 1991 in Georgia.........................................32 2-8 HindIII and XbaI restriction enzyme pattern of five iridovirus isolates...................33 3-1 Negative staining electron photomicrograph of an iridovirus particle purified by sucrose gradient ultracentrifugation.........................................................................56 3-2 Optimization of the ELISA with antige n coated at 1:100 dilution, comparing the positive to negative (P/N) ratio of two fold serial plasma dilutions of the positive control turtle (Burmese star tortoise with clinical si gns of illness) versus a negative control (Burmese star tortoi se with no history of illness)..........................57 3-3 Frequency distribution of P/N ratios from an indir ect ELISA performed on 1000 free ranging gopher tortoise ( Gopherus polyphemus ) plasma samples....................58 3-4 Individual P/N ratio values for 1000 free-ranging gopher tortoises ( Gopherus polyphemus) in increasing value..............................................................................59 3-5 P/N ratios of red-eared slider ( Trachemys scripta elegans ) plasma samples collected weekly over five months...........................................................................60 ix

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3-6 Coomassie blue staining of a SDS-P AGE gel separating proteins of iridovirusinfected and uninfected Terrapene heart cell lysates...............................................62 3-7 Western immunoblot................................................................................................63 3-8 County map of Florida. The five counties highlighted indicate where seropositive tortoise samples were identified...........................................................64 4-1 Photograph taken 12 days post-inocul ation showing development of white opaque ocular discharge in the IM inoculated box turtle ( Terrapene ornata ornata )......................................................................................................................91 4-2 Photograph taken 12 days post-inocul ation showing white caseous diphtheric plaques in the mouth of an IM inoculated red-eared slider ( Trachemys scripta elegans)....................................................................................................................92 4-3 Photograph showing exophthalmus, conjunctivitis and hyphema in an intramuscularly inoculat ed red-eared slider ( Trachemys scripta elegans )...............93 4-4 Photograph showing colonic hemorrhage in a turtle intramuscularly inoculated with Ranavirus euthanized 23 days post inoculation...............................................94 4-5 Spleen; red-eared slider ( Trachemys scripta elegans).............................................95 4-6 Spleen; red-eared slider ( Trachemys scripta elegans).............................................96 4-7 Liver; red-eared slider (Trachemys scripta elegans )................................................97 4-8 Kidney; red-eared slider ( Trachemys scripta elegans).............................................99 4-9 Colon; red-eared slider ( Trachemys scripta elegans) intramuscularly inoculated with Ranavirus .......................................................................................................100 4-10. Oral mucosa; red-eared slider ( Trachemys scripta elegans ) intramuscularly inoculated with Ranavirus ......................................................................................101 5-1 Photograph demonstrating injection of virus infected cell culture media into a ventral lymph sac in a leopard frog ( Rana pipiens)...............................................119 5-2 Photograph demonstrating the placement of a feeding tube in a red-eared slider ( Trachemys scripta elegans) for administering frog homogenates directly into the caudal esophagus..............................................................................................120 5-3 Photomicrograph demonstrating the norma l architecture of a liver in a leopard frog ( Rana berlandieri )..........................................................................................121 5-4 Photomicrograph demonstrating multifocal hepatic necrosis in a leopard frog ( Rana berlandieri ) experimentally inoculated with a Ranavirus isolated from a Burmese star tortoise (Geochelone platynota ).......................................................122 x

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5-5 Photomicrograph at higher ma gnification of the leopard frog (Rana berlandieri ) liver shown in Fig. 5-4. Arrows de note the presence of intracytoplasmic inclusion bodies consiste nt with iridovirus infections in amphibians....................123 5-6. ELISA results graphed as positive to negative (P/N) ratios. Samples were assayed weekly for the duration of the study.........................................................124 6-1. Effect of increasing concentra tions of acyclovir on the TCID50 of Terrapene cells inoculated with the Burm ese star tortoise isolate..........................................134 xi

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Abstract of Dissertation Pres ented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy IRIDOVIRUS INFECTIONS OF CAPTIV E AND FREE-RANGING CHELONIANS IN THE UNITED STATES By April J. Johnson December 2006 Chair: Elliott Jacobson Major: Veterinary Medical Sciences Iridoviruses of the genus Ranavirus are well known for causing mass mortality events of fish and amphibians with sporadic re ports of infection in reptiles. The objective of this study was to characterize Ranavirus infections of chelonians. First, histopathologic and molecular investigations of naturally occurring infections in several species of chelonian were investigated. A virus isolate (BSTRV) obtained from a captive Burmese star tortoise (Geochelone platynota ) was experimentally inoculated into western ornate box turtles ( Terrapene ornata ornata ) and red-eared sliders ( Trachemys scripta elegans). Oral transmission failed to create illnes s, however five of si x turtles inoculated intramuscularly developed clinical and hist ologic lesions consistent with naturally infected cases. Virus was re-isolated, fulfilling Koch's postulates and establishing BSTRV as a causative agent of disease and mortality in chelonians. Restriction enzyme analysis of this isolate with an isolate from a leopard frog ( Rana utricularia) obtained at the site where the tortoise died was found to have identical restriction patterns suggesting xii

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they are either the same or very closely rela ted strains. This indicates that amphibians might serve as a source of infection for ch elonians, or vice versa. BSTRV was also utilized as a coating antigen in the de velopment of an indirect enzyme linked immunosorbent assay (ELISA). Plasma from a surviving pen-mate of the Burmese star tortoise served as a positive control for optimization. A seroprevalence study of 1000 banked free-ranging gopher tortoise plasma samples found that only 1.5% of tortoises were positive for exposure to the virus. The role of amphibians in the route of transmission of virus was assessed by experimentally inocul ating leopard frogs, euthanizing them, homogenizing them, and feed ing them to turtles via feeding tubes over a six-week period. All turtles failed to de velop clinical signs or to produce antiRanavirus antibodies over three months. Last ly, the antiviral compound acyclovir was assessed at 0, 0.2, 1, 5, 10, and 25g/ml for its ability to reduce or eliminate virus replication in vitro and to create cytotoxicity in Terrapene heart cells. No cytotoxicity was observed at any concentra tion. Increasing concentration found only a slight ten fold reduction in virus titer from 10 4.8 to 10 3.8 TCID 50 xiii

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CHAPTER 1 INTRODUCTION Chelonians are long-lived animals within the class Reptilia The order Chelonia also sometimes called Testudines, consists of two suborders and thirteen families of turtles and tortoises. Chelonians worldwide are experiencing dramatic declines. The 2006 International Union for Conservation of Nature (IUCN) Red List of Threatened Animals has listed 26 species as critically endangered, 45 as endangered and 58 as vulnerable of 295 species (43.7%). An additio nal 41 (13.9%) species are listed as near threatened. Other earlier estimates indicated about 50% of all taxa to be experiencing difficulties (Jacobson et al. 1999). Reasons for declines include habitat fragmentation, increased collections for the food and pet market, change in vegetation, drought, and debilitating diseases (Jacobson et al. 1999; Dodd, 2001). Chelonians have low fecundity and low juvenile survival rates, indicati ng that a loss of adult animals can have a significant impact on population survivability (Heppell, 1998). Native species of chelonians are experiencing similar declines. Two of the three tortoises within the U.S. [desert tortoise ( Gopherus agassizii ) and gopher tortoise ( G. polyphemus )] are listed as vulnerable by the IUCN. Continuing d ecline and disappearan ce of box turtle ( Terrapene spp) populations across the genus' entire range during the last century brought the 1995 listing of all box turtle species in appendices of The Convention on In ternational Trade in Endangered Species (CITES) in an effort to slow their declines. Progress toward conserving these species is needed no w before they become endangered. 1

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2 Several important diseases ha ve been identified in populations of wild and captive chelonians. Marine turtle fibropapillomatosis is a significant health problem affecting several species of marine turtles around the world (Herbst and Jacobson, 1995). Mycoplasmosis is a chronic inf ectious disease that has been seen in wild gopher tortoises in Florida and desert tortoi ses in the southwest deserts of the United States (Brown et al. 2002). Tortoise herpesviruses have emerged as important pathogens of captive tortoises in the pet trade. Tortoise herpesvirus-1 is a causative agent of rhinitis-stomatitis complex in a variety of tort oise species (Origgi et al. 2004) and tortoise herpesvirus-2 has also been seen associated with rhinitis and st omatitis in a captive desert tortoise (Johnson et al. 2005). A study looking at all wild reptile cases submitted to the Wildlife Center of Virginia between 1991 and 2000 showed that 2% (n=694) of all cases were a result of infectious disease, although further charac terization was not described. A total of 15 cases were infectious, of whic h 14 were eastern box turtles ( Terrapene carolina carolina ) and the other case was a rat snake. Two cases had respiratory tract infections, nine had conjunctivitis while the other four had both resp iratory tract infections and conjunctivitis (Brown and Sleeman, 2002). This dissertation will demonstrate that Ranavirus es are also important emerging pathogens in wild and capti ve tortoises, and box turtles in the United States. This virus had previously been observed in a gopher tortoise (Westhouse et al. 1996) and a box turtle (Mao et al. 1997) in the United States. However, research presented here will show that it is likely res ponsible for other past a nd recent die-offs of box turtles in the eastern U.S. and may be a cause of unexplained deaths and population declines of gopher tortoises in Florida.

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3 Iridoviruses are large do uble stranded cytoplasmic DNA viruses (Williams, 1996) that were first discovered in crane fly la rvae in 1954 (Xeros, 1964) exhibiting a blue iridescence below the epidermis, which later le d to the name iridescent virus or iridovirus (Williams and Smith, 1957). Only the insect viruses are known to create iridescence, whereas vertebrate iridoviruses do not. Iridoviruses are circul arly permuted and terminally redundant (Goorha and Murti, 1982). Unlike poxviruses, which have a completely cytoplasmic site of replica tion (Schramm and Locker, 2005), iridoviruses require both the nucleus and cyt oplasm for replication (Goorha et al. 1978). The family Iridoviridae consists of four ge nera. Two genera, Chloriridovirus and Iridovirus infect insects. Viruses in the genus Iridovirus are typically smaller than those in the genus Chloriridovirus. Viruses in the genera Lymphocystivirus and Ranavirus are capable of infecting ectothermic vertebrate s. Lymphocystiviruses infect fish, while ranaviruses have been shown to infect fish, amphibians and reptiles (Mao et al. 1997). A group of unclassified erythrocytic viruse s have also been attributed to the Iridoviridae family (Johnsrude et al. 1997; Telford and Jacobs on 1993); however further characterization is needed to establish the phyl ogenetic relationship of these viruses to the classified iridoviruses. Frog virus 3 (FV3), the type species for the genus Ranavirus, was first isolated in 1966 from a renal carcinoma in a leopard frog (Granoff et al. 1965), although it was subsequently determined that there was no association of the virus to the tumor (Granoff et al. 1966). A Ranavirus also was recovered from bu llfrog tadpoles manifesting a syndrome called tadpole edema virus (TEV) (Wolf et al. 1968). In experimental studies, this virus was capable of infecting and causing significant mortality in Great Basin

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4 spadefoot toads (Scaphiopus hammondii intermontanus ), American toads (Bufo americanus ), Fowler's toads ( Bufo woodhousii fowleri ), and bullfrogs ( Rana catesbeiana) (Wolf et al. 1968). A subsequent expe rimental study demonstrat ed Fowler's toads and the newt ( Diemictylus viridiscens ) to be susceptible to TEV, as well as two other isolates from frogs (LT1 and FV1) and two ne wt isolates (T8 and T15) (Clark et al. 1969). Significant research with amphibian iri doviruses did not make much progress until the early 1990s when worldwide declines in amphibians brought new interest regarding the role of these viruses in amphibian mortality events (Bradford, 1991; Speare and Smith, 1992; Fellers and Drost, 1993; Cunningham et al. 1996; Fisher an d Shaffer, 1996; Laurance et al. 1996; Jancovich et al. 1997; Bollinger et al. 1999; Lips, 1999; Green et al. 2002; Docherty et al. 2003). In a study of sixty-f our amphibian mortality and morbidity events, iridovirus was the most common cause of mortality (Green et al. 2002). Late larval forms were more suscepti ble and epizootics were clearly associated with increased population densit ies. Affected salamanders exhibited problems with buoyancy, the inability to stay upright, swimmi ng in circles, lethargy and red spots or swollen areas on the ventrum near the gills or hind limbs (Docherty et al. 2003). While iridoviruses have been occasiona lly reported as pathogens of reptiles (Marschang et al. 2005; Hyatt et al. 2002; Drury et al. 2002; Johnsrude et al. 1997), they have not received as much attention comp ared to iridovirus infection of amphibians. In chelonians, the first report of an iri dovirus infection involved a captive Hermann's tortoise ( Testudo hermanni ) that died with necrotizing le sions in the liver, intestine and spleen (Heldstab and Bestetti, 1982). Several years later, an epidemic in a captive group of Hermann's tortoises was reported (Muller et al. 1988). While two iridoviruses were

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5 identified in an exotic tortoise ( Testudo horsfieldii ) and a box turtle ( Terrapene carolina ) in the U.S., no disease or pathology was mentioned (Mao et al. 1997). The only report in a wild tortoise involved a gopher tortoise ( Gopher polyphemus ) in Florida that had signs of respiratory disease (Westhouse et al. 1996). While viral pa rticles were seen on electron microscopy, there was no attemp t at virus isolation or molecular characterization. Recent isolates from two of seven Hermann's tortoises that died in a zoo in Switzerland were found by polymerase ch ain reaction (PCR) to have major capsid protein sequences closely related to FV3 (Marschang et al. 1999). Around this time, iridovirus infections were docum ented in soft-shelled turtles ( Trionyx sinensis ) exhibiting cervical cutaneous erythema or "red-neck di sease" at a turtle farm in China (Chen et al. 1999). In experimental infections with the iridovirus isolated from the soft-shelled turtles, the virus was shown to be a causat ive agent of the "red neck" syndrome in young inoculated turtles. Most recently, Ranavirus infections were identified in a group of seven captive eastern box turtles in North Carolina in 2002 (DeVoe et al. 2004). Six of these seven were wild caught with the most recent having been added 6 months prior to the outbreak. Clinical signs associated with Ranavirus infections in these box turtles were cutaneous abscessation, oral erosions or abscessation and respiratory distress (DeVoe et al. 2004). One of seven affected turtles also showed unilateral conjunctivitis and cellulitis of the head and neck. Ranaviruses are variably host specific and are widespread geographically (Chinchar, 2002, Daszak et al. 1999). Virus can grow in multiple types of cell lines including fish, amphibian, re ptilian, avian and mammalian, provided temperatures are conducive to growth (Chinchar, 2002). Eviden ce of viruses being capable of infecting

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6 multiple species of animals has been demons trated both naturally and experimentally. Inter-class infections with Ranavirus may occur among sympatric species in the wild, with fish or amphibians servi ng as the reservoir host (Mao et al. 1999). Moody and Owens (1994) demonstrated an anuran virus, Bohle iridovirus, to be pathogenic for a fish, Lates calcarifer. Experimental transmission of Bohle iridovirus, a virus isolated from amphibians in Australia, was experimentally in oculated into six species of reptiles of which two turtle species, Emydura krefftii and Elseya latisternum appeared to be susceptible showing increased mo rtality in inoculated hatchli ngs (Ariel, 1997). An insect iridovirus was isolated from a chameleon, two bearded dragons, and a frill-neck lizard (Just et al. 2001). Thus, some of these viruses are very unusual in their ability to infect phylogenetically distinct lineages of vertebra tes and invertebrates. However, other isolates have been shown to be very host specific and cannot be transmitted experimentally to other classes of animals (Jancovich et al. 2001). Natural transmission of iridoviruses has yet to be definitively identified, and may vary between genera or species of viruses. Experimental studies have shown that cannibalism of infected animals or ingestion of infected water may serve as a route of infection in amphibians (Jancovich et al. 2001; Pearman et al. 2004). Experimental infections of salamanders with a Ranavirus showed that both dose and host characteristics influenced the virulence of in fection (Brunner et al. 2005). The infection dose was positively correlated with mortality rate and inversely related to average survival times. Environmental temperatures have also been shown to significantly impact the percent mortality and time to death in sa lamanders experimentally infected with a Ranavirus (Rojas et al. 2005), where salamanders infect ed at 18 and 10C were more

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7 likely to die than those exposed at 26C. Two antimicrobial peptides, esculentin-2P and ranatuerin-2P, isolated from the skin of leopard frogs ( Rana pipiens) are able to inactivate Frog Virus 3 in a dose dependent manner (Chinchar et al. 2001) and may play a role in viral resistance. The immune response of reptiles to ir idoviruses has not been previously investigated; however, some research has been performed in this area for amphibians. Birds, reptiles and amphibians produce a lo w-molecular weight immunoglobulin called IgY (reviewed in Warr et al. 1995). A study of 21 wild caught cane toads ( Bufo marinus ) in Townsville, Australia, found three to ads to have anti-iridovirus antibodies cross-reactive against both epizootic haematopoetic necros is virus and bohle iridovirus on enzyme linked immunosorbent assay (ELISA). Sera from positive toads or toads that had been exposed to an iridovirus showed positive to negative (P/N) ratios of 2.81, 2.91 and 3.4 compared to sera from naive toads, which ranged from 0.55 to 1.13 (Whittington et al. 1997). Other studies in amphibians ha ve focused on the African pipid frog, Xenopus laevis. One study experimentally infected adult frogs four weeks apart. Anti-FV3 IgY was detected in plasma one week following the second injection. This appearance of antibodies correlated to the time of viral clea rance and the ameliorati on of clinical signs (anorexia and cutaneous erythema), suggesting a role of the adaptive immune system in clearing infection (Gantress et al. 2003). This study also suggests the genotype of the MHC to play a significant role in the host susceptibility. Inbred Xenopus having a decreased MHC class I expression were mo re susceptible to infection, as were Xenopus larvae, which also lack MHC cl ass I gene expression (Gantress et al. 2003). Another study looking at antibod y response to FV3 in Xenopus also showed that a second

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8 exposure was necessary for developing a dete ctable response of IgY production (Maniero et al. 2006). Antibodies were first detectable 10 days after a second exposure and levels plateaued at 14 days. IgM levels were ne ver detectable even after three exposures. Prevention of iridovirus infections has been researched primarily in fish. A vaccine against red seabream iridovirus (RSIV) has shown that genetic vaccines can be effective in protecting red seabream ( Pagrus major ) against experimental challenge with RSIV (Caipang et al. 2006). Inoculation of juvenile red seabream with DNA plasmids encoding either the major capsid protein ge ne or an open read ing frame encoding a transmembrane domain against RSIV showed upregulation of transcription of the MHC class I gene. Additionally, DNA vaccinated fish showed lower mortalities after subsequent exposure to RSIV than did non-va ccinated fish. Formalin-killed virus has also been shown to be effective in upre gulating MHC class I transcripts (Nakajima et al. 1997; Nakajima et al. 1999). A field trial testing the e fficacy of a formalin-killed virus showed a 49.3% decrease in mortality betw een vaccinated and control groups, and a statistically significant incr ease in size of fish in th e vaccinated group (Nakajima et al. 1999). A recent study evaluated an environm ental contaminant and its potential to contribute to increased mortalit ies in iridovirus infected am phibians (Forson and Storfer, 2006). Somewhat surprisingly, the authors f ound that atrazine at moderate doses may reduce the efficacy of iridoviruses and, thus, pr otect animals exposed to both atrazine and iridovirus, resulting in decrea sed mortality rates. However, high levels of exposure to atrazine could result in decreased fitness. Salamanders were metamorphosed earlier and were smaller at metamorphosis than those expos ed to no or moderate levels of atrazine (Forson and Storfer, 2006).

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9 Treatment of iridovirus infections with antiviral drugs are not reported in the literature; however previous research has sugge sted that antiviral treatment with acyclovir might be successful. Acyclovir becomes activ ated by a viral gene product, an enzyme, thymidine kinase. Sequencing of the entire genomes of several isolates has identified open reading fames encoding putative thymidin e kinase (TK) genes, and functionality was further studied in the Bohle iridovirus (BIV) (Coupar et al. 2005). The BIV TK gene was inserted into a TK gene negative mutant vaccinia virus insertion plasmid, under the control of a vaccinia virus promoter and us ed to infect human 143B (TK-) cells. The investigators were able to show that this gene expressed a functional TK gene by rescuing of the TK negative mutant. Comparison of seque nces of different iridovirus TK genes to other virus TK genes showed that unlike poxviruses, iridoviruse s, similar to the herpesviruses, appear more closely related to the mito chondrial TKs and to cellular deoxycytidine kinases, whereas the TK gene sequence in other DNA viruses including poxviruses and African swine fever virus appear more closely relate d to the cellular TK genes (Coupar et al. 2005). Thymidine kinase genes can activate nucleoside analogs such as acyclovir if the gene can recognize deoxy cytidine as an alternative substrate. The ability of different iridovirus TKs to do th is has not been evaluated. While TK genes appear well conserved among the different genera, they vary considerably between genera within the family, so each woul d need to be evaluated separately. The research presented in the following chap ters will investigate several aspects of Ranavirus infections in chelonians. Chapter 2 reports on molecular and histopathologic findings of natural cases of iridovirus infections in wild a nd captive chelonians in the United States. It describes several recent ch elonian cases and two past die-offs of box

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10 turtles dating back to 1998 and 1991 and de monstrates a new geographic range of infections from the northeast to Texas. Chapter 3 will describe the development and use of an indirect enzyme linked immunosorbent a ssay (ELISA) to evaluate the prevalence of iridovirus exposure in wild gopher tortoises. Chapter 4 wi ll describe an experimental infection study fulfilling Koch's postulates, confirming that iridovirus is a primary pathogen of chelonians. Chapter 5 reports on a study investigating a possible route of transmission in which ingestion of infected amphibians could be a source of iridovirus in a natural setting. Chapter 6 will describe the use of an antiviral drug, acyclovir, at reducing replication of iridovirus in vitro. And lastly, chapter 7 will discuss overall conclusions and directions for future research.

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CHAPTER 2 RANAVIRUS INFECTION OF FREE-RA NGING AND CAPTIVE BOX TURTLES AND TORTOISES IN THE UNITED STATES Introduction It has been suggested that chelonians (turtles and tortoi ses) face a more serious threat than that posed by the well-publici zed decline of amphibian populations (Klemens, 2000). Two thirds of all species of freshwater turtles and tortoises ar e currently listed as threatened on the IUCN Red List of Th reatened Species (Turtle Conservation Fund, 2002). Chelonians have low fecundity, low j uvenile survival ra te, and a long adult lifespan; a life history stra tegy where loss of adult animals (such as loss by disease) has a significant impact on population recovery (Heppe ll, 1998). Emerging in fectious diseases have been increasingly recogni zed as factors influencing wi ldlife health and populations (Harvell et al ., 1999; Daszak et al ., 2000). Although mycoplas mosis has been postulated to contribute to declines of some tortoise species (USFWS, 1994), the cause(s) of mass mortality events in wild chelonian populat ions often remain undetermined (Flanagan, 2000; Dodd, 2001). Among the emerging diseases of wild life, iridoviruses in the genus Ranavirus are well known for causing mass mortality events of fish and amphibians (Langdon and Humphrey, 1987; Daszak et al. 1999; Green et al ., 2002). Iridovirus infections have also been sporadically described in reptiles including snakes (Hyatt et al. 2002), lizards (Marshang et al ., 2005) and chelonians. In chelonians iridovirus infections have been reported in captive Hermann's tortoises ( Testudo hermanni ) (Heldstab and Bestetti, 1982; 11

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12 Muller et al ., 1988; Marschang et al ., 1999), farmed soft-shelled turtles (Chen et al ., 1999) and captive eastern box turtles (DeVoe et al ., 2004). Two ranaviruses identified in an exotic tortoise ( Testudo horsfieldii ) and a box turtle ( Terrapene carolina ) in the U.S., were found to be closely related to frog viru s 3 (FV3) and designated as tortoise virus 5 and turtle virus 3, respectively (Mao et al. 1997). The only report of an iridovirus infection in a free-ranging cheloni an involved a gopher tortoise ( Gopher polyphemus ) in Florida with signs of respiratory disease (Westhouse et al ., 1996). This report identifies Ranavirus infections in five recent chelonian deaths or mortality events from Georgia, Florida, Ne w York, and Pennsylvania and in archived material recovered from previously unexpl ained mass mortality events in 1991 from Georgia and 1998 from Texas. This demons trates a previously undescribed geographic extent of chelonian Ranavirus infections and suggests th at ranaviruses may be more important pathogens of free-ranging chelonians than anticipated from previous reports. We also present molecular evidence for an id entical or similar virus in frogs in the vicinity of two chelonian epizootics suggesti ng that amphibians could serve as reservoir hosts for chelonians. Materials and Methods Animals Burmese star tortoises. Three female and two male captive Burmese star tortoises ( Geochelone platynota ) were kept in an outdoor encl osure at St Catherines Island Wildlife Survival Center, Georgia (31N/ 81W) since April 2001. In early June 2003, two female and one male tortoise began showing clinical signs consisting of nasal discharge, conjunctivitis and se vere subcutaneous edema of the neck (Fig. 2-1A). The tortoises were treated with antimicrobials, a nd were soaked daily for 90 minutes in warm

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13 water. One female tortoise died three days after developing clini cal signs and yellow white caseous plaques were observed on the to ngue at necropsy. Oral antiviral therapy and intracoelomic fluids were then initiated in the surviving tortoises. Subsequently, six adult Southern leopard frogs (Rana utricularia ) were sampled from within the tortoise pens and one was sampled from a pond nearby. One of the frogs from the pen was found moribund while others appeared healthy. Tissues from the dead tortoise and the leopard frogs were submitted for histopathology, polymerase chain reaction (PCR) for determining presence of certain DNA sequences of Ranavirus and Herpesvirus and virus isolation. Gopher tortoise. A wild gopher tortoise ( Gopherus polyphemus ) was found circling on a road in north central Florida (29 86'N/82 22'W) on 25 July 2003 and was brought to the University of Florida, Co llege of Veterinary Medicine Zoological Medicine Service for evaluation and treatment On presentation it exhibited palpebral swelling and ocular and nasal discharge (Fig 2-1B). The tortoise was treated with intracoelomic fluid twice daily, was allowed to soak in shallow warm water for 20 minutes a day and was started on antimicr obial therapy. The tortoises condition continued to decline and it was euthanized with intrav enous Beuthanasia-D solution (Schering-Plough Animal Health Corp., Kenilw orth, NJ) on 29 July 2003. A complete necropsy was performed and tissues were submitted for histopathology, PCR and virus isolation. Eastern box turtles. A 200 ha gated and fenced area within a private nature sanctuary in Venango County, Pennsylvania, (41N/79W) was created as a study site for relocated box turtles ( Terrapene carolina carolina ). All box turtles within the

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14 study site were radiotelemetered and tracked regularly, and no box turtles were present at the site prior to the release of the relocated turtles. During the summer of 2003, there were 32 adult and 34 head-started juvenile turtles. Turtles were observed every five days to determine their health status and location. Fifteen of the 66 turtles (23%) died between 15 August and 20 November 2003. Many of th e turtles were considered healthy approximately four to eight days prior to being found either moribund or dead. Nine of the turtles were found dead, while six were found moribund with palpebral edema, ocular discharge, and fluid draining from the mouth. Moribund turtles were taken out of the preserve, treated with an ophthalmic ointment in the eyes, soaked daily in warm water and given a temperature gradient. Two turtle s were started on antimicrobials and one of the two also received a parasi ticide. All moribund turtles died within hours to days after being found exhibiting signs of illness. In May 2004, two freshly dead green frog tadpoles ( Rana clamitans ) with marked cutaneous erythema were collected from a pond at the nature preserve. Tissues from five turtles collected in 2003 and from the tadpoles collected in 2004 were submitted for hist opathology, PCR and virus isolation. Two wild box turtles were found moribund w ith ocular discharge and swelling, as well as aural abscesses and yellow caseous plaq ues in the oral cavity in Suffolk County, New York (40N/72W) on 2 August 2005. They were observed near the edge of a drying pond that is utilized by many pond-breed ing amphibians, including green frogs ( Rana clamitans) and bullfrogs (Rana catesbeiana). These animals were taken to a local wildlife rehabilitator, where one died overnig ht. The carcass was then put in a freezer for later evaluation. The other had the aural abscesses drained and was treated with antimicrobials, but its health continued to decline until its death on 1 September 2005.

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15 This turtle was put on ice and, along with the frozen specimen, sent to the University of Florida for histopathology, PCR and virus isolation. Florida box turtle. A wild Florida box turtle ( Terrepene carolina bauri ), found in north central Florida (29 42'N/82 23'W) in October 2004, was submitted to the Zoological Medicine Service, College of Veteri nary Medicine, University of Florida, for treatment. The box turtle exhibited palpebral edema, nasal and ocul ar discharge (Fig. 21C) and had yellow-white caseous plaques in the oral cavity (Fig. 2-1D). The turtle was administered fluid intracoelomically with B vitamins daily and was started on analgesics to alleviate pain. Due to failure to respond to therapy and a poor pr ognosis the turtle was euthanized three days after admission with intravenous Beuthanasia-D solution. A complete necropsy was performed and tissu es submitted for histopathology, PCR and virus isolation. Past mortality events. Tissues from two previous box turtle epizootics of undetermined etiology were examined. In July and August 1991 over thirty Eastern box turtles were found dead in or near wate r sources in Murray county (34'N/84'W), northwest Georgia (Dodd, 2001). Two moribund turtles were found exhibiting lethargy, ocular discharge and had caseous white plaques in the oral cavity. On e turtle also had a subcutaneous abscess caudal to the left eye. Both turtles were submitted for necropsy. In 1998, several Eastern box turtles and other unspe cified turtle species died suddenly in a private collection in Texas (Dodd, 2001). Ar chived paraffin blocks for histologic examination were obtained from two box turtles from the Georgia die off and one box turtle from Texas and re-evaluated using light and transmission electron microscopy.

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16 Necropsy and Histopathology At necropsy, tissues were collected from all major organ systems of the following tortoises and turtles: Burmese star tortoise (1), gopher tort oise (1), easte rn box turtles [Pennsylvania (5), Georgia (2), New York (1), and Texas (1)] and Florida box turtle (1). Tissues were fixed in neutral buffered 10% formalin, dehydrated in graded alcohols, embedded in paraffin, sectioned at 6 m, and stained with hematoxylin and eosin. Tongue, liver and spleen of each animal were collected and frozen at C for detecting DNA sequences of Ranavirus and Herpesvirus using PCR and virus isolation. Nucleotide Amplification, Se quencing, and Sequence Analysis DNA was extracted from chelonian and am phibian tissues and cell cultures used for virus isolation using the DNeasy kit (Qia gen, Valencia, CA, USA). Five 5m thick paraffin embedded sections from box turt les from the 1991 and 1998 mortality events were extracted using the DNeasy kit following the protocol for paraffin embedded tissue. Sense primer (5-GACTTGGCCACTTATCAC-3) and anti-sense primer (5GTCTCTGGAGAAGAAGAA-3) as pr eviously described (Mao et al ., 1997) were used to amplify approximately 500 base pairs of the Ranavirus major capsid protein gene. A 50 l reaction mixture was run which contained 4 l extracted DNA, 1 M of each primer, 200 M each of dATP, dCTP, dG TP, and dTTP, 2.5 U of Taq DNA polymerase and PCR buffer containing 50 mM KCl, 10 mM Tris-HCl, 1.5 mM MgCl 2 (Eppendorf, Westbury, New York, USA). Th e mixtures were amplified in a thermal cycler (PCR Sprint, Thermo Hybaid) with an initial denaturation at 94 C for 2.5 min, followed by 25 cycles of denaturation at 94 C for 30 sec.; annealing at 50 C for 30 sec, extension at 72 C for 30 sec., and a final extension step at 72 C for 10 min as previously described

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17 (Marschang et al. 1999). The same extracted DNA from the chelonian tissues was evaluated by PCR for the presence of cheloni an herpesvirus(es). A nested consensus PCR was performed as previously described to detect a portion of the herpesvirus DNA dependent DNA polymerase (VanDevanter et al ., 1996). Any PCR products were resolved in 1% ag arose gels and bands were excised and purified using the QIAquick ge l extraction kit (Qiagen). Pr oducts were sequenced in both directions directly using the Big-Dye Terminator Kit (Perkin-Elmer, Branchburg, New Jersey) and analyzed on ABI 377 automated DNA sequencers at the University of Floridas Sequencing Center. Virus Isolation Turtle heart cells [TH-1; American T ype Cell Culture (ATCC), Manassas, VA] were seeded into 25 cm 2 flasks (Costar, Corning, NY, USA). Cells were cultured in Dulbeccos modified Eagle medium (DMEM, Gibco, Carlsbad, CA, USA) supplemented with 5% fetal bovine serum (Gibco, Carl sbad, CA, USA), gentamicin (60 mg/liter) (Sigma, St. Louis, MO, USA), penicillin G (120,000 U/liter), streptomycin (120,000 U/liter) and amphotericin B (300 g/liter) (Sigma) and cultured to confluency. A small piece of spleen or liver from each case was hom ogenized in separate 5ml tissue grinders containing DMEM. Part of each homogenate was applied to a flask of confluent monolayer of TH cells while the other was passed through a 0.45 m filter (Costar) onto another flask of cells. Cells were incubate d at 28 C. Flasks were observed daily for cytopathic effect (CPE).

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18 Transmission Electron Microscopy Second passage TH-1 cell monolayers (75mm 2 flasks) inoculated with first passage isolates from homogenates of kidney tissue co llected at necropsy from the Burmese star tortoise and the gopher tortoise were ex amined by transmission electron microscopy (TEM). Cells were harvested 3 days after in fection (2 days after observation of CPE) and centrifuged at 4,500 x g for ten minutes. Supernatant was discarded and the remaining pellet was suspended in Trumps fixative (4 % paraformaldehyde, 1% gluteraldehyde). Cells were post-fixed in osmium tetroxide, de hydrated in graded alcohols and embedded in Spurr's resin. Tissues were viewed us ing a Hitachi H7000 transmission electron microscope at the University of Fl orida Electron Microscopy Laboratory. Paraffin embedded spleen from a box turtle from the 1991 mass mortality event in Georgia and paraffin embedded trachea from a box turtle from Texas from the 1998 mass mortality event were deparaffinized in xylene, embedded in Spurr's resin, sectioned for TEM and examined as described for cell culture. Skin from the necropsied Burmese star tortoise was placed in Trump's solution (McDowell and Trump, 1976) was submitted to the Athens Diagnostic Laboratory, University of Georgia for TEM. Tissue wa s post fixed in osmium tetroxide, dehydrated in graded alcohols and embedded in epoxy. Ultr athin sections were stained with uranyl acetate and lead citrate and examined w ith a JEOL JSM-1210 transmission electron microscope. Restriction Enzyme Analysis Frog Virus 3 was obtained from ATCC a nd served as the positive control for comparative purposes with the viruses isolat ed in this study. Fr og Virus 3 and second passage isolates of tissue homogenates from the Burmese star tortoise and Southern

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19 leopard frog that were collected at the same site were inoculated onto TH-1 cells. Once CPE was observed, viral DNA was radiolabeled with [methyl3 H] thymidine, extracted and digested with HindIII and XbaI endonucleases as previo usly described (Mao et al ., 1999). Restricted DNA fragments were separa ted on a 0.7% agarose gel, after which the gel was impregnated with Enhance (Perkin Elmer, Wellesley, MA) according to the manufacturer's directions and the radiolabel ed fragments were detected by fluorography. Results Necropsy and Histopathology Histologic findings were similar in the Bu rmese star tortoise, Gopher tortoise and box turtles. Consistent lesions in all animals were necrotizing and ul cerative stomatitis or esophagitis, fibrinous and necrotizing splenitis and multicentric fibrinoid vasculitis. Lesions in the oral cavity a nd esophagus were characterized by near diffuse mucosal erosion and ulceration with surfaces covere d by a thick coagulum comprised of fibrin, degenerate heterophils, sloughe d epithelial cells and bacter ial colonies (Fig. 2-2). Lesions in the spleen consisted of disrupti on of the white and to a lesser degree, the surrounding red pulp by deposits of fibrin ad mixed with karyorrhectic debris, and infiltrates of small numbers of heterophils (Fig. 2-3). There was frequently mild to marked red pulp congestion and/or hemorrhage. Fibrinoid vasculitis with thrombosis was observed in splenic, sheathed capillaries (ellip soids) in all animals and to varying degrees in other locations incl uding oral mucosa, esophagus, stom ach, intestine, skin, lung, heart, and liver. All animals had some degree of necr osis of hematopoietic tissue in the kidney, liver and bone marrow. Individual cases had multifocal necrotizing tracheitis (1/7), conjunctivitis (1/7) or gastri tis (2/7). Rarely, basophili c intracytoplasmic inclusion bodies suggestive of iridovirus infection were observed within epithelial cells of the oral

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20 mucosa, esophagus, stomach and trachea, and/or within endothelial cells, macrophages (Fig. 2-4) and hematopoietic progenitor cells Inclusion bodies were observed in only 3 of 7 (Burmese star tortoise, eastern box turtle, and gopher tortoi se) cases examined histologically. Necrosis of hepatic and renal hemat opoietic tissues with rare basophilic intracytoplasmic inclusion bodies consistent w ith iridovirus infecti on was observed in the moribund southern leopard frog from the Burmese star tortoise pen in Georgia and in one green frog tadpole from the site in Pennsylvania. Archived tissues from the past two morta lity events in Georgia and Texas showed similar histologic lesions as described above. These include d fibrinous splenitis in all turtles as well as a necrotizing tracheitis w ith rare intracytoplasmic basophilic inclusion bodies in sloughed respiratory epithelial cells in the box turtle from Texas. PCR and Sequence Analysis PCR for the Ranavirus major capsid protein ge ne yielded DNA fragments approximately 500 base pairs in length (Fig. 2-5). After sequencing of the fragments and excluding the primer component, the sequences of the gopher tortoise, star tortoise, all box turtles, southern leopard frog and green frog shared 100% seque nce identity. The sequences were compared to known sequen ces in GenBank (National Center for Biotechnology Information, Bethesda, Maryland), EMBL (Cambridge, United Kingdom), and Data Bank of Japan (Mishima, Shiuoka, Japan) databases using TBLASTX (Altschul, et al. 1997). TBLASTX results for the seque nces all showed the highest score with FV3 capsid protein gene (GenBa nk accession # AF157769). Comparison of sequences showed that all isolates shared 100% sequence identity with FV3 across that portion of the major capsid protein gene PCR of DNA extracted from paraffin

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21 embedded sections of tissue from box turtles from the 1991 and 1998 die offs were negative. All tortoises a nd box turtles were negative by PCR for the presence of herpesvirus. Virus Isolation All TH-1 cells infected with tissue homoge nates from the dead turtles and tortoises exhibited cytopathic effects (C PE) that consisted of cell r ounding and lysis two to three days post infection. Flasks that contained both filtrated tissue homogenate as well as unfiltered tissue homogenate showed CPE. The Burmese star tortoise isolate was also passaged onto fathead minnow cel ls (ATCC), which subsequently exhibited similar CPE. Transmission Electron Microscopy Using TEM, the cell cultures infected w ith tissue homogenates from the Burmese star and gopher tortoises showed large numbers of icosahedral shaped viral particles that were consistent in size (appr oximately 130nm) and shape with an iridovirus (Fig. 2-6). Similar viral particles were observed with in intracytoplasmic inclusion bodies in endothelial cells and macrophages of the skin in the Burmese star tortoise, within the cytoplasm of unidentified cells in the spleen of a 1991 box turtle from Georgia and within intracytoplasmic inclusion bodies of degenerate respiratory epithelial cells of the trachea in the 1998 box turtle from Texas (Fig. 2-7). Restriction Enzyme Analysis Repeated attempts failed to show discre te bands of the green frog tadpole after restriction with both enzymes, and thus, comparisons cannot be made with this isolate. Restriction with the Hind III enzyme demonstrated identical patterns between the FV3, the Burmese star tortoise and the leopard frog isolate collected at th e same location as the star tortoise (Fig. 2-8A). The eastern box turtle isolate showed a slightly different

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22 pattern. Restriction with XbaI found a similar result (Fig. 2-8B). FV3, the Burmese star tortoise and leopard frog isolates showed id entical patterns, while the box turtle isolate was different. Discussion Emerging infectious diseases are those that have newly appeared in a population or have previously existed but are rapidly increasing in in cidence or geographic range (Morse, 1995). The findings reported here sugge st that iridovirus infections in chelonians fill this description, and are emerging pathoge ns of chelonians. Infections are being discovered in new populations of turtles, and the incidence is either increasing, or our ability to detect the disease in these animal s is increasing. Infections in chelonians are more geographically widespread than has been previously documented. Previous reports from the United States had identified chel onian iridovirus infec tions in a wild gopher tortoise in Florida (Westhouse et al. 1996), captive box turtles in North Carolina (DeVoe et al. 2004) and in two chelonians where the location was not reported (Mao et al. 1997). Here we have identified more infecti ons than previously described within a twoyear period in Georgia, Florida, Pennsylvan ia, and New York, indica ting an increase in incidence of infection. Identif ication of iridovirus-like partic les in the mortality event in 1998 expands the new geographic range furthe r to include Texas. We also have described infection in a Burmese star tortoi se, a species that has previously not been documented to be susceptible to iridovirus infections. Reports of mortality events involving large numbers of box turtles and gopher tortoises have been documented in which et iologies were never definitively identified (Dodd, 2001; Rossell et al. 2002; Seigel et al ., 2003). This report demonstrates by re

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23 evaluation of archived samples that at leas t some of these mort ality events (Dodd, 2001) have been infected with a virus(es) compatib le with iridovirus and mortalities may have been caused by infection with these viruses. Polymerase chain reaction of these cases were negative, however it is unknown how long tissues had been fixed in formalin prior to being embedded into paraffin. Formalin fixation has been shown to degrade DNA and could result in false negative results (Tokuda et al. 1990). Still, TEM of the same tissues showed virus particles consistent in size and shape with iridovirus. The histologic lesions in th e chelonians in this report were relatively non-specific and the intracytoplasmic basophilic inclusion bodies suggestive of iridovirus infection (Heldstab and Bestet ti, 1982; Marschang et al. 1999; Bollinger et al. 1999; Docherty et al. 2003) were absent or were rare and could easily be missed. Clinical and pathological differential diagnoses for the animals in this report prior to the demonstration of iridoviruses included chelonian herpesvirus infection (Johnson et al. 2005) for the lesions of necrotizing stomatitis and septicemia for the fibrinous splenitis and multicentric vasculitis. While iridovirus infection should be considered in cases with lesions similar to those seen in the turtles and tort oises in this report, ancillary diagnostic tests, including viral isolation attempts a nd PCR for ranaviruses and herpesviruses, should be performed to confirm the diagnosis. All chelonian and amphibian isolates in this study shared 100 % sequence identity across a portion of the major capsid prot ein gene. The major capsid protein gene sequence is fairly conserved among iridoviru ses, although one study has shown that it contains enough diversity to be able to di stinguish closely related isolates (Tidona et al. 1998). Mao et al (1997) compared MCP sequences a nd restriction enzyme patterns of

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24 whole genomic DNA of 10 vertebrate iridoviru ses, including one virus isolated from a box turtle, Turtle Virus 3 (TV3). While th e sequence obtained from a portion of the TV3 MCP gene indicated that it was identical to that of Frog Virus 3, a restriction enzyme analysis using HindIII and XbaI showed a different restriction pattern between the two isolates. Our study found a similar result. An isolate from a box turtle from the Pennsylvania die-off shared 100% sequence id entity with FV3 acro ss a portion of the MCP gene, however the whole viral genomic re striction enzyme analysis pattern differed from FV3, and showed a similar result to that of the box turtle isolate, TV3, reported by Mao et al. (1997). This suggests that the major ca psid protein gene may be too conserved to determine if different animals are in fected with the same virus. An interesting and potentially significan t finding was identical viruses, as determined by restriction enzyme analysis, in the Burmese star tortoise and a moribund southern leopard frog found within its pen. Th is suggests that both animals were infected with the same virus. Inter-class infections have been shown previously in a natural setting where sympatric species of amphibians a nd fish were infected with the same virus species (Mao et al ., 1999) as well as through experime ntal transmission studies (Moody and Owens, 1994). There are several ways that chelonians and amphibians might be exposed to the same virus. Previous studies in salamanders have shown that transmission can occur through cannibalism of in fected individuals (Jancovich et al ., 2001; Pearman et al ., 2004). Box turtles are omnivorous, and to rtoises while normally herbivorous, may opportunistically feed on carrion. This was c onfirmed when animal caretakers at the site of the Burmese star tortoise death in Georgia observed a ra diated tortoise ( Geochelone radiata ) and a Burmese black mountain tortoise ( Manouria emys phayrei ), both normally

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25 considered herbivorous, eating dead amphibian s in nearby pens. There could also be a common environmental source of virus, such as shared bodies of water. Iridoviruses are quite resistant and thought to be capable of persisting in water sources for extended periods of time (Daszak et al ., 1999). Iridoviruses create sy stemic infections, and thus, a vector-borne route of transmission might also be a wa y that both amphibians and chelonians could become infected. In summary, this report demonstrates that Ranavirus is an emerging pathogen of chelonians and suggest that amphibians might serve as a source of infection. This describes a new geographic range for chelonian ir idovirus infections in the United States. Ranaviruses are considered a global threat to amphibian populations based on the lack of host specificity, high virulence and global distribution (Daszak et al ., 1999) so they should likewise be considered a global threat to cheloni an populations.

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26 Fig. 2-1. Gross lesions associated with iridovirus infections in tu rtles and tortoises. A) Photograph of a Burmese star tortoise ( Geochelone platynota ) with nasal discharge and palpebral and cervical edema. B) Photograph of a wild gopher tortoise ( Gopherus polyphemus ) with nasal discharge and palpebral edema. C) Photograph of a wild Florida box turtle (Terrapene carolina bauri ) with palpebral edema and ocular discharge. D) Photograph of caseous plaques in the oral cavity of a Florida box turtle.

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27 Fig. 2-2. Esophagus, eastern box turtle ( Terrapene carolina carolina ). There is diffuse necrosis and ulceration of the mucosa and replacement by fibrin, inflammatory cell infiltrates and superfic ial bacterial colonies (arrows). H&E stain.

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28 Fig. 2-3. Spleen, eastern box turtle (Terra pene carolina carolina). There is disruption of the white and red pulp with deposits of fibrin (arrow) admixed with karyorrhectic debris, and infiltrates of small numbers of heterophils. H&E stain.

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29 Fig. 2-4. Epicardium, Burmese star tortoise (Geochelone platynota ). Arrows depict basophilic intracytoplasmic inclusion bodies in a macrophage and endothelial cell. The lumen of the blood vessel contains a fibr in thrombus (F). H&E stain.

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30 Fig. 2-5. Results of a polymerase chain reaction targeting approximately 500 bp of the major capsid protein gene. Lane 1 is a 100 bp ladder. The bright band represents a 500 bp fragment. Lane 2 and 3 are positive samples from an eastern box turtle ( Terrapene carolina carolina ) and a green frog tadpole ( Rana clamitans). Lane 4 is a positive contro l sample from a Burmese star tortoise ( Geochelone platynota ) confirmed previously with nucleotide sequencing. Lane 5 is a negative control.

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31 Fig. 2-6. Transmission electron photomicrograph of Terrapene heart cells inoculated with liver tissue from a Burmese star tortoise (Geochelone platynota ) demonstrating cytoplasmic arrays of iridovirus-like particles.

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32 Fig. 2-7. Transmission electron photomicrograph of paraffin embedded spleen from a box turtle ( Terrapene carolina ) that died in 1991 in Georgia. There are icosahedral virus particles compatible with iridoviruses.

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33 A B Fig. 2-8. HindIII and XbaI restriction enzyme pattern of five iridovirus isolates. A) HindIII restriction. Lane 1 is Frog Viru s 3, the type species for the genus Ranavirus. Lane 2 is an isolate from a Burmese star tortoise ( Geochelone platynota ) and lane 3 is an isolate from a southern leopard frog (Rana utricularia ) collected from the pen adjacent to the Burmese star tortoise. Lane 4 is an isolate from an eastern box turtle ( Terrapene carolina carolina ) from Pennsylvania and Lane 5 is a green frog tadpole (Rana clamitans) isolate from the same location as the box turtle. B) XbaI restriction pattern. Lanes are the same as those in Fig. A.

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CHAPTER 3 DEVELOPMENT AND USE OF AN INDIRECT ENZYME LINKED IMMUNOSORBENT ASSAY FOR DETECTION OF IRIDOVIRUS EXPOSURE IN GOPHER TORTOISES ( Gopherus polyphemus ) Introduction Iridovirus infections of the genus Ranavirus have recently been identified in freeranging and captive native chelonians from Flor ida, Georgia, New York, North Carolina, Tennessee, and Pennsylvania (Allender et al., in press; DeVoe et al. 2004; Johnson et al., 2004). Species affected were Burmese star tortoises ( Geochelone platynota ), gopher tortoises (Gopherus polyphemus ), eastern box turtles ( Terrapene carolina carolina ), and Florida box turtles (Terrapene carolina bauri ). Evidence of iridovirus infection was also observed in archived material from previ ously unexplained mass mortality events of eastern box turtles ( Terrapene carolina carolina ) from Georgia in 1991 and Texas in 1998 (Dodd, 2001; Johnson et al., 2004). Clinical signs asso ciated with infections included those of upper respiratory tract dis ease including respiratory distress and nasal discharge, as well as oral ulceration, cu taneous abscessation, anorexia and lethargy (Westhouse et al., 1996, DeVoe et al., 2004). Consistent lesions in affected animals included necrotizing stomatitis and/or esophagiti s, fibrinous and necrotizing splenitis, and multicentric fibrinoid vasculitis Intracytoplasmic inclusion bodies were rarely observed in affected tissues. A portion of the major capsid protein (MCP) gene was sequenced from recent cases from Georgia, Florida and Pennsylvania and found to be identical across approximately 500 basepairs to each ot her and to Frog Virus 3 (FV3), the type species of the genus Ranavirus in the family Iridoviridae (Johnson et al., 2004). Koch's 34

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35 postulates were fulfilled by experi mentally inoculating a tortoise Ranavirus isolate into red-eared sliders (Johnson et al., unpublished findings). Three of four sliders developed severe clinical signs including anorexia (3/3), lethargy (3/3), oral plaques (1/3), nasal discharge (3/3), ocular disc harge (3/3) and exophthalmus conjunctivitis, and hyphema (1/3). Histologic changes were similar to those seen in naturally infected cases. Virus was isolated from tissues of each of the three turtles, fulfilling Koch's postulates and establishing iridovirus as a pr imary pathogen in chelonians. Iridoviral infections are the most common cause of mortality events in amphibians in the United States (Green et al., 2002). Iridoviruses ar e globally distributed and thus considered a threat to amphibian populations worldwide based on the lack of host specificity, high virulence and ubiquitous distribution (Daszak et al., 1999). The geographic range of Ranavirus infections in chelonians in the U.S. has recently been found to be larger than previously known. Prior to 2003 only three cases of chelonian infections had been reported in the U.S.; however only one report in cluded the location of the infected chelonian. A w ild gopher tortoise from Florida was found to have iridoviruslike particles by transmission el ectron microscopy (Westhouse et al., 1996). A box turtle and Russian tortoise ( Testudo horsfieldi ) isolate were described in another report (Mao et al., 1997), but the location was not disclosed, and as Russian tortoises are not native to the United States, it is possible they were both kept as pets. Current published and unpublished reports now show a much larger ge ographic range with ir idovirus infected chelonians identified from Texas to New York and Pennsylvania (Allender et al., in press; DeVoe et al. 2004; Johnson et al., 2004). Therefore, it is reasonable to assume

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36 that chelonians throughout the eastern United States can be exposed to iridoviruses; however, the prevalence rate of e xposure has not b een determined. Serology can be a useful tool for detec ting previous exposure to pathogens. Indirect enzyme linked immunos orbent assays (ELISA) have been used to detect exposure of various reptiles to specific pathogens (Schumacher et al., 1993; Origgi et al. 2001; Brown et al., 2001; Jacobson et al., 2005) and has been used to detect exposure of amphibians to iridovirus infections (Whittington et al. 1997; Gantress et al., 2003; Maniero et al. 2006). To determine iridovirus expos ure in wild gopher tortoises in the U.S., we developed an indirect ELISA usi ng a previously developed mouse anti-desert tortoise IgY monoclonal antibody as the secondary antibody (Schumacher et al., 1993). We also describe the results of a larger serological survey of w ild gopher tortoises from various sites in Alabama, Florida, Ge orgia, Louisiana and Mississippi. Materials and Methods Virus A Ranavirus isolated and partially characte rized from a naturally infected Burmese star tortoise in Georgia (Johnson et al. 2004), here termed BSTRV, was used as the antigen in the development of the ELISA Briefly, polymerase chain reaction (PCR) targeting a portion of ranaviral major cap sid protein genes followed by nucleotide sequencing demonstrated that the BSTRV is olate shared 100% se quence identity of approximately 500 basepairs with FV3. Tr ansmission electron microscopy of BSTRV inoculated TH-1 cells showed vi rus particles in the cytoplasm of infected cells consistent in size and shape with iridoviruses. Rest riction enzyme digests of BSTRV compared with FV3 showed identical restri ction patterns us ing the enzymes HindIII and XbaI, indicating that BSTRV is either identi cal or very closely related to FV3.

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37 Antigen Preparation Two methods of antigen preparation were used. The first method of preparation was by sucrose gradient ultracentrifugation as previously described for epizootic haematopoietic necrosis vi rus of fish (Steiner et al. 1991). Terrapene heart cells (TH-1) were acquired from the American Type Cu lture Collection (ATCC-CCL 50; Rockville, MD) and grown to confluency in 225cm 2 tissue flasks (Costar, Corning, NY). Cells were cultured in Dulbecco's modified Eagle medium (DMEM, Gibco, Carlsbad, CA) supplemented with 5% fetal bovine serum (Gibco), gentamicin (60mg/liter; Sigma, St. Louis, MO), penicillin G (120,000 U/liter), streptomycin (120,000 U/liter) and amphotericin B (300g/liter; Sigma). Cells were inoculated with a fourth passage of BSTRV and incubated at 28C in the presence of 5% CO 2 When cytopathic effects (CPE) were observed in over 70% of cells, consisting of cell rounding and detachment from the flask, the flasks were frozen and th awed 3 times with vigorous vortexing before each freeze. Supernatant was transferred to 15ml tubes and clarified by low speed centrifugation at 4,500x g for 15 minutes. Supernatant was then decanted into two sterile 1-liter bottles a nd stored at 4 C until 1.5 liters of supernatan t were obtained. Virus was then pelleted at 10,000x g for 8 hours at 4 C and the supernatant was discarded. Pellets were resuspended in the residual media, divi ded into four equal parts and overlayed on four 15-60% (w/v) sucrose gradients. Grad ients were then ultr acentrifuged at 150,000x g for 45 minutes at 4 C. Bands of purified virus we re collected by fractionation and diluted in Tris-HCl (pH 8.0) until sucrose wa s less than 20%. Bands in Tris-HCl were layered onto four 5ml 20% (w/v) sucros e cushions and ultracentrifuged at 80,000x g for one hour at 4 C. Sucrose was next decanted and pellets were resuspended in 200 l Tris

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38 HCl and stored at -80 C. Purity was assessed with negative staining electron microscopy, protein assay (Biorad, Hercules, CA), PCR and ELISA. The second way antigen was prepared wa s by creating a cell lysate from virus infected cells. Virus was inoculated onto TH-1 cell monol ayers as described above in 225 cm 2 flasks. Flasks were scraped when fl asks exhibited 100% CPE. Uninfected flasks were concurrently processed in the same manner to serve as control antigen to detect any background cross reactivity of plas ma to cellular proteins. Cells and media were transferred to 15ml tubes and centrifuged at 4,500x g for 30 minutes. Supernatant was then discarded and the cell pellets were resuspended in residual media, and then frozen and thawed three times. Tubes were vortexed before and after each freeze cycle and following the final thaw, we re centrifuged again at 4,500x g for 30 minutes. Supernatant was then transferred to a 4m l sterile cryotube. A protein assay was performed to determine the final protein concentration of the antigen. PCR was performed to confirm the presence of viral DNA. Positive and Negative Reference Plasma In July of 2003, three of five captive Burmese star tortoises became ill with clinical signs consis ting of nasal discharge, conjunc tivitis and cervical subcutaneous edema. One of the three tortoises died and histologic and molecular investigations demonstrated the presence of iridovirus in various tissues. Surv iving tortoises were treated with supportive care and all four tortoises survived. Plasma was collected at the time of infection during July and then again in September 2003. Plasma from one of these tortoises was used as the positive cont rol in development of the ELISA. Plasma

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39 was collected from Burmese star tortoise s from a zoological collection with no known history of disease to se rve as negative reference plasma for the ELISA. ELISA Procedure A checkerboard optimizati on strategy was used to determine the optimum concentrations of both antigen and plasma to be used in the ELISA. Antigen concentrations evaluated were 1:100, 1:250, 1:500, and 1:1000. Plasma concentrations evaluated were two-fold serial dilutions from 1:50 to 1:1600. The following procedure was found to be optimal utilizing the crude cell lysate antigen. Each well of a high protein binding 96 well microplate (Maxis orp F96; Nunc, Kamstrup, Denmark) was coated with 50 l of infected or uninf ected cell lysate diluted to 1:100 in 0.01M sodium phosphate buffer (pH 7.2) containing 0.15 M NaCl and 0.02% sodium azide (PBS/Az). Plates were then incubated overnight at 4 C. Antigen was then aspirated off and wells were washed four times with in ELISA wash buffer (PBS/Az with 0.05% Tween 20). This washing process was performed between each of the following steps. Wells were then blocked against non-specific binding with 300 l of Superblock blocking buffer by Pierce (Rockford, IL) for one hour at room te mperature (RT). Each remaining step was incubated for 1hr at RT. Plasma samples di luted 1:100 in blocking buffer were added at 50 l volumes to wells in triplicate. One we ll was coated with uninfected cell lysate while the other two wells were coated with in fected cell lysate. Next, a biotin-conjugated monoclonal antibody produced against the desert tortoise IgY light chain and previously used for detecting anti-mycoplasma anti bodies in desert tortoises (Schumacher et al. 1993) was diluted to a final concentration of 0.5 g/ml in PBS/Az and added to each well in 50 l volumes. Alkaline phosphatase-conjugate d streptavidin (Zymed Laboratories,

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40 Inc., San Francisco, California) was then applied to each well at 50 l of a 1:5000 dilution in PBS/Az. Next, the ELISA was developed with 100 l per well of a 1.0mg/ml P nitrophenyl phosphate prepared in 0.01M sodium bicarbonate buffer containing 2mM MgCl 2 and plates were stored in the dark. The absorbance of each well was read at A 405 using a StatFax 3200 microplate reader (Awareness Technology, Palm City, Florida) at 30 minutes. Each plasma sample was read in triplicate. Plasma was placed on one well originally coated with uninfected cell lysate and on two wells coated with infected cell lysate. The average absorbance reading of the two wells coated with infected cell lysate was calculated and the positive/negative (P/N) ratio value of each sample was determined by dividing the mean absorbance of the duplic ate average by the absorbance reading of the well from the uninfected cell lysate. Th is subtracts out any background noise, or nonspecific binding that may have o ccurred as a result of using the Terrapene heart cell line. The cut-off value for a positive test result was made by adding three times the standard deviation of the mean P/N ratio to the mean (Crowther, 2001). Experimentally Inoculated Turtles Although iridovirus infection was confirmed in the Burm ese star tortoise that died, it was not confirmed in any of its pe n-mates. So while we assumed the other clinically ill tortoises were exposed, there was no way to definitively know. To develop known positive and negative samples for use in developing the ELISA and to validate the test by detection of seroconversion, ten red-eared sliders ( Trachemys scripta elegans ) ranging in weight from 775 to 1050g were obtained from a rep tile dealer and allowed to acclimate in an animal care faci lity room at the University of Florida for three weeks.

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41 This study was performed under the approval of the Institutional Animal Care and Use Committee at the University of Florida. Oral and cloacal swabs were collected from turtles and ran by PCR as previously described (Mao et al. 1997) for the presence of Ranavirus DNA sequences to determine current infection status. Plasma was colle cted and tested by the following ELISA to determine presence of antiRanavirus antibodies at the time of arrival, which would indicate previous exposure. Turtles were randomly assigned to one of three groups. Group 1 turtles (Nos. 13, 14, 16 and 20) received 1ml of a virus infected crude cell lysate prepared as described above and dilu ted to a final concentration of 10 2 TCID 50 /ml orally (PO) by metal gavage feeding tube placed into the distal esophagus. Group 2 turtles (Nos. 12, 15, 17, 18) received the same dose of virus intramsucularly (IM), half the dose in the right and half the dose in the left p ectoral muscles. The two other turtles were assigned to a control route of inoculation. Turtle 11 was mock inoculated with the same volume of an uninfected cell ly sate by PO while turtle 19 received an uninfected cell lysate by IM. Plasma samples were collected weekly for five months from each turtle to attempt to detect the production of antibodies. Turtles were euthanized at five months, or when severe clinical signs of disease appeared including any of the following: severe lethargy, subcutaneous edema, nasal or ocul ar discharge, oral plaques, or hyphema. Tissues were collected at necropsy for PCR, histopathology and virus isolation. Reproducibility Intra-assay and inter-assay reproducib ility were determined by performing two precision runs. The positive and negative refere nce plasma samples, used to optimize the test, were used in each assay. Intra-assay reproducibility was determined by running the positive and negative sample multiple times on the same plate. Each sample is run

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42 multiple times on one well of uninfected cell lysa te and two wells of infected cell lysate. This resulted in 64 readings for each sample on the infected cell lysate and 32 on the uninfected cell lysate. Interassay reproducibility was determ ined by using the values of the reference plasma results used as cont rols in running up the wild gopher tortoise samples from multiple dates and multiple plates. The mean A 405 the standard deviation (SD) and the coefficient of va riation (CV) for the intraand inter-assay reproducibility were calculated using the optimized ELISA conditions. Protein Expression and Immunoblotting The positive control plasma was tested for its ability to detect viral proteins in a western blot using infected and uninfected TH -1 cell lysates as antig en. Four four-fold dilutions of infected and uninfected cellular lysate protei ns were separated by sodium dodecyl sulfate-poly acrylamide gel electrophores is (SDS-PAGE) under reducing conditions along with broad range molecular weight markers. The separated proteins were transferred onto 0.2m nitrocellulo se membranes (Biorad, Hercules, CA) by standard methods (Harlow and Lane, 1988). Me mbranes were then rinsed in water for five minutes in preparation for coomassie blue staining or in tris buffered saline containing 0.5% Tween20 (TTBS, pH 7.5) fo r twenty minutes prior to immunoblotting. Membranes were then stained in coomassie blue stain for 90 minutes, followed by water for 30 minutes to destain to view different ial protein profiles between infected and uninfected cell lysates. Afte r rinsing in TTBS, membranes for immunoblotting were then blocked with Superblock blocking buffer in phosphate buffered saline (Pierce) for one hour. Blocking buffer was then removed and plasma samples diluted 1:2000 in blocking buffer were added to the membranes. After one hour, the membranes were washed with TTBS for 30 minutes and the monoclonal antibody was added, diluted 1:10,000 in

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43 blocking buffer for one hour. Again the membranes were washed and AP streptavidin was added, diluted 1:5,000 in phosphate buffe red saline for one hour. Membranes were washed as previously, and the membranes were developed in substr ate buffer (0.1M TrisHCl, 1mM MgCl 2 ) containing nitroblue tetrazoliu m chloride (NBT) and 5-bromo-4chloro-3-indolylphosphate p-toluidine salt (BCIP) (Biora d). The reaction was stopped by removing the NBT-BCIP solution and adding deionized water. Membranes were allowed to air dry. Wild Gopher Tortoises Samples with Unknown Exposure Plasma samples from 1000 wild gopher tortoises ( Gopherus polyphemus) from Florida, Georgia, Alabama, Louisiana a nd Mississippi were obt ained from samples submitted to the Mycoplasma Research Laboratory at the University of Florida from 2002 to 2006. County data was recorded when available, although it was not available in 68 cases. Samples were further subdivided by region of the state including central, central east, central west, north central, north east, north west, south east, and south west. Results were also compared according to state. Results Antigen Preparation Two diffuse bands were observed on the sucrose gradients and each band was separated into two samples when placed on the sucrose cushion for pelleting. This resulted in four stocks of purified virus (two low bands 1A and 1B and two higher bands 2A and 2B). Ten microliters of each stock was submitted for negative staining electron microscopy (EM). While virus partic les were observed (Fig. 3-1), they were difficult to find by EM. A protein assay of each sample indicated protein concentrations of 1000 (1A), 375 (1B), 625 (2A) and 500 g/ml (2B). Polymerase chain reaction of

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44 each sample resulted in strong positive signals Use of the purified antigen in an ELISA indicated approximately 37.5 g/ml of the purified virus wa s necessary for coating wells to obtain the best results. Because such a small volume of purified iridovirus resulted from the sucrose gradient purification protoc ol, and such a large volume was needed for use in the ELISA, it was decided that a crude cell lysate would be used for detecting seroconversion in the transmission studies and for the seroepidemiology study. ELISA Parameters Checkerboard optimization found that d iluting the antigen 1:100 and diluting plasma samples also at 1:100 gave the larg est difference between P/N ratios of positive and negative reference samples (Fig. 3-2). Plasma cut off values were determined after running the wild gopher tortoise samples. Th e frequency distribution was determined in tenth increments from 0.5 to 4.1 and the P/N ratios of the samples were determined to have a normal distribution, skewed to the right (Fig. 3-3). Because the data was normally distributed, the mean P/N ratio of samples (1.07 8), plus three times the standard deviation (0.379) was used to determine the positive cu t-off value (2.2). P/N ratios from each sample were plotted on a graph with the samp le numbers on the x-axis in increasing order and the P/N ratio values on the y-axis (Fig. 3-4). This plot shows a gradual increase in P/N ratios followed by a sharp increase around a P/N ratio of about 2.0, confirming that our cut off value of 2.2 to be a reasonable value. Experimentally Inoculated Turtles One IM inoculated turtle (No. 15) became extremely lethargic and died 24 days post-inoculation. Cloacal swabs taken on turt le 15 were positive by PCR and sequencing for the presence of iridovirus starting 8 days before the turt le died. Oral swabs were positive the day before the turtle died. Ot her than lethargy, no clinical signs were

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45 observed in this turtle such as stomatitis, c onjunctivitis or nasal discharge. Histologic changes were consistent with those seen in naturally and experimentally inoculated turtles (DeVoe et al. Johnson et al. unpublished data). Briefl y, there was a fibrinoid vasculitis in the spleen with multifocal infiltrates of low numbers of heterophils and scattered free brown pigment granules (pre sumptively from disrupted melanomacrophage centers). There were occasional lumenal fi brin thrombi with ad mixed heterophils and karyorrhectic debris. No other turtles di ed during the five-month course of the study. Plasma samples collected weekly over the five-month period failed to detect seroconversion in all of the mock inoculat ed (Fig. 3-5A) turtle s and turtles orally inoculated with Ranavirus (Fig. 3-5B). One IM inoculated turtle (No. 12) had an increasing P/N ratio trend, but only twice did values exceed a ratio of 2, which occurred on weeks 8 and 18 (Fig. 3-5C). All other turtles remained well below the positive P/N ratio cut off value of 2.2. Reproducibility The mean A 405 SD and CV values for the intraand inter-assay pr ecision runs are shown in Table 1. Ideally, CV values shoul d be less than 15% (Crowther, 2001). Three of the CV values in the precision runs are >15% indicating that some variability still exists in this assay. Protein Expression and Immunoblotting Coomassie blue staining of proteins fr om infected cell lysates and uninfected cell lysates showed a different pa ttern of protein expression (F ig. 3-6). Immunoblotting of infected and uninfected cell lysa te showed a marked increase in binding to proteins in the infected cell lysate and very weak binding to proteins in the uninfect ed cell lysate (Fig. 37). Strong signals were seen on virus-infected cells at approximately 125 kDa, and 78

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46 kDa with weaker signals seen at 70, 65, and 28 kDa. Very faint signals were also seen at 125 and 78 kDa in the uninfected cells. Wild Gopher Tortoises Samples with Unknown Exposure Of 1000 gopher tortoise plasma samples assayed, 15 (1.5%) were positive with a P/N ratio >2.2 (Table 2). Eight seropositive tortoises came from five counties in Florida including Lake, St. Lucie, Broward, Palm Beach and Martin (Fig. 3-8). While seropositive tortoises represent three re gions including central, central east, and southeast, four of the five counties are clus tered closely together in the south including Palm Beach, Broward, Martin, and St. Lucie. The remaining seven seropositive tortoises were located in Baker, Georgia, a county in th e southwest corner of the state. Prevalence of seropositive tortoises by c ounty was quite variable (Table 3). Approximately 3% of tortoises sampled in Lake county were positiv e (n=99), 2.9% in St. Lucie (n=35), 10% in Broward (n=10), 3.4% in Palm Beach (n=58) and 6.2% in Martin county (n=16). Of tortoises tested from Baker, GA 6.5% were positive (n=113). By state, Florida had an overall prevalence of 1.2% (n=658) while Georgia had a prevalence of 3.1% (n=225) (Table 3). All tortoises tested from Alabam a, Mississippi and Loui siana were negative. Discussion Iridoviruses of the genus Ranavirus are emerging as important pathogens of chelonians. Previous report s have established their presence in Florida (Westhouse et al. 1996) and in North Carolina (DeVoe et al., 2004). Newer unpublished findings indicate a much larger range of infection including re ports from Texas, Georgia, New York and Pennsylvania (Johnson et al., 2004). The duration of illness can be rather short (Johnson et al. unpublished findings), making it difficult to observe symptomatic tortoises in wild populations. Therefore, we developed an indirect ELISA to detect anti-Ranavirus

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47 antibodies and performed a cr oss-sectional survey of gophe r tortoises utilizing banked plasma samples to attempt to determine the prevalence of exposure of free-ranging chelonians to iridoviruses. When developing a new assay, it is ideal to have another assay against which to compare results. With a reference assay, the sensitivity and specificity of the new assay can be evaluated, and thus the level of confid ence one has in its ability to detect what you want it to detect. Unfortunate ly, this was not possible with this pathogen. Detectable neutralizing antibodies are not usually found in naturally or experimentally infected animals (reviewed by Whittington et al., 1997) and thus, serum neutralization tests, which are sometimes used as gold standards for verifying ELISA results, cannot confidently be used to determine exposure to this virus. For this reason we decided to perform experimental transmission studies using red-ear sliders to at tempt to demonstrate the validity of our assay by detecting se roconversion. Our assay failed to detect antibodies in all but one slider. However, recent reports show that multiple exposures are needed in amphibians to detect IgY antibod ies against FV3 using an ELISA (Gantress et al., 2003; Maniero et al. 2006) and IgM antibodies were not detectable even after multiple exposures. Only one dose of virus was admininstered to experimentally inoculated turtles, which might explain why th e turtles in this study failed to mount an immune response. Our results did show a good corre lation between antibody production following a clinical disease in a Burmese star tortoise that was previously housed with a tortoise that died with confirmed Ranavirus infection. Plasma from a Burmese star tortoise from another facility with no known history of illness was selected as a seronegative control; antibodies to Ranavirus were not found in this tortoise.

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48 Additionally, using a western blot, we were ab le to show that plasma from the positive control turtle bound to proteins in the infected cells in the immunoblot but not in the uninfected cells indicating that the plasma was reacting with viral proteins and not cellular proteins from the lysate we were usi ng to coat the plates. For these reasons, we assume our test to be valid for use in detecting anti-iridovirus antibodies in chelonians, however further validation of this assay is warranted. Only one turtle died as a result of IM inoculati on of virus at 10 2 TCID 50 A previous study showed that 75% (n=4) of tu rtles inoculated with a higher dose of virus (10 5 TCID 50 ) died as a direct result of iridovirus inf ections (Johnson et al., unpublished data). The viral dose was extrapolated fr om experimental studies with fish and amphibians (Langdon, 1989, Moody and Owens, 1994, Bollinger, 1999, Cullen and Owens, 2002), so it was unknown what a sub-lethal dose would be in chelonians. While studies with a larger sample size might provide more accurate correlations between viral load and mortality rates, it appears that host ch aracteristics likely play a significant role in resistance or susceptibility to disease. Environmental temperatures have also been shown to significantly impact the percent mortality and time to death in salamanders experimentally inoculated with a Ranavirus (Rojas et al. 2005), where salamanders inoculated at 18 and 10C were more likel y to die than those exposed at 26C. Underlying disease conditions we re not noted at necropsy or on histologic review of tissues in the turtle that died that might have contributed to an increased susceptibility. Prevalence among free-ranging gopher tortoise s was found to be low, only 1.5% of 1000 samples being positive. This could be the true prevalence rate, although we suspect that this is an underestimate of the true rate There are several f actors that could cause

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49 the prevalence to be underestimated. Prevalen ce is a function of the incidence of disease multiplied by the average duration of the illn ess. While incidence would be difficult to determine in a natural setting, duration of illn ess can be extrapolated from experimental studies. If chelonians die quickly as a resu lt of infection, they will not have time to mount an immune response to the pathogen, a nd will not survive to be surveyed. As previously mentioned, experimental transm ission studies have shown a high rate of mortality (75%) in turtles intramuscularly inoculated with 10 5 TCID 50 of a Ranavirus infected cell lysate (Johnson et al., unpublished data). Turtles al l died within 30 days of exposure to the virus. Although, the rout e of transmission is unknown in a natural environment, if naturally exposed cases experience similar mortality rates and duration from exposure to death, a cross-sectional study evaluating the prevalence of exposure will miss many tortoises that were exposed, beca use the majority of them will die. This was demonstrated in a natural setting in Pe nnsylvania. A population of approximately 70 eastern box turtles ( Terrapene carolina carolina) was being repatriated in a nature sanctuary. Turtles were tracked every 3-5 da ys by radiotelemetry. In the summer and fall of 2003, 15 of these turtles died suddenly, with what was later idenitified to be iridovirus inf ections (Johnson et al., 2004). The following spring, the remaining 55 turtles were sampled, and plasma was run for the presence of antibodies. Only three turtles were positive on ELISA (data not shown). If we calculated this value as the true prevalence, we would estimate that ~5.5% of turtles were exposed, when we know that 21% of the population died that was not in cluded in the estimate. This severely underestimates the prevalence of disease. However, we cannot extrapolate these differences to other populations, as this was a repatriated population, and thus, subject to

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50 stressors that might not be found in wild populations, making them more susceptible to disease. Secondly, this study suggests that tu rtles must be exposed more than once to mount an immune response similar to what was found in studies with Xenopus (Gantress et al., 2003; Maniero et al. 2006) Thus, if turtles have only been exposed once, we will not detect that first exposure with our ELISA, also underestimating the true prevalence of exposure. Emerging infectious diseases have been increasingly recognized as factors influencing wildlife health and populations (Harvell et al ., 1999; Daszak et al ., 2000). Although mycoplasmosis has been postulated to contribute to declines of some tortoise species (USFWS, 1994), the cause(s) of mass mortality events in wild chelonian populations often remain undetermined (Flanagan, 2000; Dodd, 2001). Iridovirus infections in chelonians can have a high morta lity rate, but the duration of illness is short, making it difficult to observe disease outbreaks in the wild. It is po ssible that it might contribute significantly to mort ality rates in wild populations of chelonians. Utilizing a serological assay may help to determine regi ons where iridovirus infections might be more prevalent in chelonian populations. Thes e locations could then be monitored more closely for disease outbreaks in both chelonians and amphibians. Results of the serosurvey showed that counties in th e central and southeastern region of Florida were more likely to have seropositive tortoises. Interestingly, four of these counties had adjoining borders (St. Lucie, Martin, Palm Beach and Broward), suggesting that either tortoises in this area are at higher risk of exposure to iridoviruses, or alternatively, iridovirus is more endemic in this area, so lower levels of exposure allow more tortoises to seroconvert without succumbi ng to the disease. Direct comparisons of

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51 prevalence rates between countie s are difficult to make as this was a convenience sample and geographic area or tortoise densities per county data were not controlled for in sampling. Only 32 of 67 counties were sampled. The northwestern region was not well sampled, and inferences about this region can't be made. However, seven tortoises in Baker County, Georgia were seropositive and th is county is located in the southwestern portion of the state, which would be near th e northwestern portion of Florida. Earlier studies have shown that amphibians might be a source of iridovirus infections in chelonians (Johnson et al. unpublished data). A moribund leopard frog was found and euthanized at the same site in Georgia where the Burmese star tortoises became ill and one died. Restriction enzyme analysis of viral genomic DNA from an isolate obtained from the tortoise and the frog demonstrated identical restriction patterns, suggesting they are the same or very closely related viruses. Thus, conditions that might propagate amphibian iridovirus infections would likely cause an increased chance of exposure in chelonians. Green et al., (2002) found that increased precipitation and population densities were direc tly associated with increase d die-offs of amphibians. Thus, these similar settings might create higher rates of exposure in chelonians. Additionally, it has been shown that sublethally infected amphibians can cause sporadic, recurrent disease outbreaks in amphibians (Brunner et al ., 2004). Experimentally and naturally infected tiger salamander larvae and metamorphs were able to maintain sublethal, transmissible infections for over five months. Apparent ly healthy infected dispersing metamorphs were returning to wate r bodies to breed and it was speculated that these individuals were likely serving as a reservoir host for infecting newly hatched larvae, creating recurrent outbreaks of dis ease. It is unknown whether chelonians are

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52 capable of sustaining sublethal infections or capable of spreading disease to naive populations. One turtle experimentally in oculated intramuscularly with iridovirus remained clinically healthy but continued to shed virus from the cloaca detectable by PCR up until 30 days. The study ended at 30 days so it is unknown how long this turtle would have kept shedding, or if the virus bei ng shed was still infec tious. Further studies would be useful to determine the risk posed to other chelonian and amphibian populations of iridovirus infected turtles that survive the in itial infection. This assay could be useful for managing populations in wild and captive settings by identifying tortoises who might be asymptomatic carriers. In summary, this study reports the development of an indirect ELISA for detection of anti-iridovirus antibodies in chelonians. It was able to detect an tibodies in a naturally infected Burmese star tortoise whose pen-mate died with a confirmed iridovirus infection. A seroprevalence survey of banked plasma sa mples from free-ranging gopher tortoises in Florida, Georgia, Alabama, Louisiana and Mississippi found a 1.5% prevalence rate of exposure. Further studies are ne eded to characterize the true incidence of disease in wild populations of chelonians.

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53 Table 3-1. Reproducibility of the ELISA. SD = standard deviation of the mean A 405 CV = coefficient of variance expressed as a percent. ICL = values from samples run on wells coated with infected cell lysate. UCL = values from samples run on wells coated w ith uninfected cell lysate. Positive Sample Negative Sample n Mean A 405 SD CV n Mean A 405 SD CV Intra-assay ICL 64 0.363 0.022 6.06 64 0.107 0.011 10.28 UCL 32 0.094 0.009 9.57 32 0.112 0.009 8.04 Inter-assay ICL 26 0.366 0.079 21.58 26 0.130 0.023 17.69 UCL 13 0.095 0.011 11.58 13 0.111 0.017 15.31 Table 3-2. ELISA results of 1000 free-ranging gopher tortoise ( Gopherus polyphemus ) plasma samples by county and state. State County Number tested Positive Percentage AL Baldwin 2 0 0 AL Mobile 7 0 0 FL Alachua 12 0 0 FL Brevard 27 0 0 FL Broward 10 1 10.0 FL Citrus 51 0 0 FL Clay 1 0 0 FL Collier 2 0 0 FL Columbia 1 0 0 FL Hernando 24 0 0 FL Hillsborough 16 0 0 FL Indian River 2 0 0 FL Lake 99 3 3.0 FL Lee 18 0 0 FL Leon 17 0 0 FL Madison 1 0 0 FL Manatee 6 0 0 FL Marion 22 0 0 FL Martin 25 1 4.0 FL Miami-Dade 8 0 0 FL Nassau 47 0 0 FL Orange 40 0 0 FL Osceola 11 0 0 FL Palm Beach 58 2 3.4

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54 FL Pasco 26 0 0 FL Pinellas 2 0 0 FL Polk 6 0 0 FL Sarasota 1 0 0 FL Seminole 54 0 0 FL St. Johns 7 0 0 FL St. Lucie 35 1 2.9 FL Taylor 1 0 0 FL Volusia 26 0 0 FL Walton 2 0 0 GA Baker 113 7 6.2 GA Liberty 74 0 0 GA Tattnall 38 0 0 LA Washington Parish 12 0 0 MS Greene 7 0 0 MS Harrison 16 0 0 MS Perry 5 0 0 Unknown 68 0 0 TOTAL 1000 15 1.5

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55 Table 3-3. ELISA results of 658 free-ranging gopher tortoises ( Gopherus polyphemus ) from the state of Florida are listed by region. C = central, CE = centraleast, CW = centralwest, NC= northcentral, NE = northeast, NW = northwest, SE = southeast and SW = southwest. Region Total No. Tested No. Positive Percent C 232 3 1.3 CE 90 1 1.1 CW 126 0 0.0 NC 32 0 0.0 NE 55 0 0 NW 2 0 0 SE 101 4 4.0 SW 20 0 0 Table 3-4. ELISA results of 1000 free-ranging gopher tortoises ( Gopherus polyphemus ) listed by state. State Total No. Tested Total No. Positve Percent Alabama 9 0 0 Florida 658 8 1.2 Georgia 225 7 3.1 Louisiana 12 0 0 Mississippi 28 0 0

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56 Fig. 3-1. Negative staining el ectron photomicrograph of an iridovirus particle purified by sucrose gradient ultracentrifugation.

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57 ELISA Optimizatio n 0 1 2 3 4 5 6 7 8 50 100 200 400 800 1600 Reciprocal of Plasma Dilutions Neg Control Pos Control Fig. 3-2. Optimization of the ELISA with antigen coated at 1:100 dilution, comparing the positive to negative (P/N) ratio of tw o fold serial plasma dilutions of the positive control turtle (Burmese star tortoise with clinical signs of illness) versus a negative control (Burmese star tortoise with no history of illness). Plasma diluted at 1:100 showed the greatest difference between the positive and negative control.

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58 P/N Ratio Frequency Distribution0 50 100 150 200 2500 5 0 5 9 0 6 0 6 9 0 7 0 7 9 0 8 0 8 9 0 9 0 9 9 1 0 1 0 9 1 1 1 1 9 1 2 1 2 9 1 3 1 3 9 1 4 1 4 9 1 5 1 5 9 1 6 1 6 9 1 7 1 7 9 1 8 1 8 9 1 9 1 9 9 2 0 2 0 9 2 1 2 1 9 2 2 2 2 9 2 3 2 3 9 2 4 2 4 9 2 5 2 5 9 2 6 2 6 9 2 7 2 7 9 2 8 2 8 9 2 9 2 9 9 3 0 3 0 9 3 1 3 1 9 3 2 3 2 9 3 3 3 3 9 3 4 3 4 9 3 5 3 5 9 3 6 3 6 9 3 7 3 7 9 3 8 3 8 9 3 9 3 9 9 4 4 0 9 > 4 1P/N Ratios Fig. 3-3. Frequency distri bution of P/N ratios from an indirect ELISA performed on 1000 free ranging gopher tortoise ( Gopherus polyphemus ) plasma samples. Samples show an approximately normal distribution, skewed to the right.

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59 ELISA Results0 1 2 3 4 5 6 Tortoises Fig. 3-4. Individual P/N ratio valu es for 1000 free-ranging gopher tortoises ( Gopherus polyphemus) in increasing value.

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60 0.0 0.5 1.0 1.5 2.0 2.5 Pre1234567891011121314151617181920 Weeks post-inoculationP/N Ratio RES19 RES11 A 0.0 0.5 1.0 1.5 2.0 2.5Pre 1 2 3 4 5 6 7 8 9 1 0 11 1 2 1 3 14 1 5 1 6 1 7 1 8 19 2 0Weeks post-inoculationP/N Ratio RES13 RES14 RES16 RES20 B

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61 0.0 0.5 1.0 1.5 2.0 2.5Pre 1 2 3 4 5 6 7 8 9 1 0 11 1 2 1 3 14 1 5 1 6 1 7 1 8 19 2 0Weeks post-inoculationP/N Ratio RES12 RES15 RES17 RES18 C Fig. 3-5. P/N ratios of red-eared slider ( Trachemys scripta elegans) plasma samples collected weekly over five months. A) P/N ratios of mock-inoculated turtles. B) P/N ratios of turtles orally inoculated with Ranavirus C) P/N ratios of turtles intramuscularl y inoculated with Ranavirus

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62 Fig. 3-6. Coomassie blue staining of a SD S-PAGE gel separating proteins of iridovirusinfected and uninfected Terrapene heart cell lysates. La ne 1 is a broad range, pre-stained molecular weight marker with weight in kDa marked next to the lane. Lanes 2-5 are four fold serial dilu tions of iridovirus infected cell lysate. Lanes 6-8 are four fold serial dilutions of uninfec ted cell lysate.

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63 Fig. 3-7. Western immunoblot. Lanes coat ed with four fold serial dilutions of iridovirus infected cell lysate in lanes 25 and uninfected cell lysate in wells 69. Plasma from the positive control was used as the primary antibody for detection of iridovirus specific antibody binding. St rong signal was seen on virus-infected cells at approximate ly 125 kDa, and 78 kDa with weaker signals seen at 70, 65, and 28 kDa. Very faint signals were also seen at 125 and 78 kDa in the uninfected cells.

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64 Fig. 3-8. County map of Florida. Th e five counties highlighted indicate where seropositive tortoise sa mples were identified.

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CHAPTER 4 EXPERIMENTAL TRANSMISSION OF A RANAVIRUS IN WESTERN ORNATE BOX TURTLES ( Terrapene ornata ornata ) AND RED-EARED SLIDERS ( Trachemys scripta elegans) Introduction Viruses in the family Irid oviridae are large doub le stranded DNA viruses capable of infecting ectothermic vertebrates, and invertebrates (Williams et al ., 2005). Iridoviruses within the genus Ranavirus have been shown capable of infecting fish, amphibians and reptiles (Chinchar, 2002) and have emer ged as major pathogens of free-ranging amphibians worldwide (Cunningham et al ., 1996, Zupanovic et al., 1998, Daszak et al ., 1999). In a study of sixty-four amphibian mo rbidity and mortality events between 1996 and 2001, the most common cause of mortality events was infection with iridoviruses (Green et al ., 2002). Iridovirus infections in rept iles have been less well described than in amphibians and fish, with sporadic reports in chelonians, snakes, and lizards (Just et al. 2001, Hyatt et al ., 2002, Marschang et al. 2005). Infection of a tortoise was first reported in 1982 in a spur-tailed Mediterranean land tortoise ( Testudo hermanni ) that had necrotic foci in the liver, intestine and spleen (Heldstab and Best etti, 1982). Subsequently, iridoviruses were reported in other species of chelonians both in captivity and in the wild (Westhouse et al., 1996, Chen et al., 1999, Marschang et al., 1999, DeVoe et al., 2004). Clinical signs associated with infections have included si gns of upper respiratory tract disease including respiratory distress and nasal discharge, as well as oral ulceration, cutaneous abscessation, anorexia and lethargy (Westhouse et al., 1996, DeVoe et al., 2004). While 65

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66 there is circumstantial evidence that iridov irus is a primary pathogen in chelonians, Kochs postulates have never been fulfilled Ranavirus was recently identified in tissues ob tained from a variety of wild and captive chelonians that died in Georgia, Fl orida, Texas, New York and Pennsylvania (Johnson et al ., 2004). Species affected were Burmese star tortoises ( Geochelone platynota ), gopher tortoises ( Gopherus polyphemus ), eastern box turtles ( Terrapene carolina carolina ), and Florida box turtles ( Terrapene carolina bauri ). Clinical signs in these cases included lethargy, anorexia, na sal discharge, conjunctivitis, severe subcutaneous cervical edema, and necrot izing pharyngitis-stomatitis. One of the ranaviruses isolated from a Burmese star tortoise ( Geochelone platynota ) that died in an outdoor pen in a zoological collection from Georgia in 2003 wa s further characterized as either identical or closely related to Frog Virus 3 (Johnson et al ., 2004). To determine a causal relationship between the isolated Ranavirus and the clinical and histologic lesions observed in these chelonians and to fulfill Koch's postulates, we performed two experimental transmission studies using th e previously characterized Burmese star tortoise isolate that will hereafte r be termed Burmese star tortoise Ranavirus (BSTRV). The first was a pilot study to determine the suitabili ty of either western ornate box turtles ( Terrapene ornata ornata ) or red-eared sliders ( Trachemys scripta elegans ) as an experimental model for a subsequent larg er transmission study. Here we report the findings of those studies. Materials and Methods Experimental Animals and Husbandry Study 1 was a pilot study consisting of th ree western ornate box turtles (BT; Terrapene ornata ornata ) and three red-eared sliders (RES; Trachemys scripta elegans )

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67 that were purchased from a reptile supplie r in Texas. Study 2 consisted of ten RES purchased from a reptile supplier in Louisian a. Both studies were performed under the approval of the Institutional Animal Care and Use Committee at the University of Florida. Turtles were housed individually in plastic containers in a centralized animal facility room maintained at approximately 25 o C. Appropriate husbandry for each species was used that included commercially availabl e foods. Box turtles were kept on land, with overhead heat lamps provided for basking at one end of the container (average temperature of 28 o C). Red-eared sliders were kept in water (temperatures averaging between 21.3 and 25.6C), with basking platfo rms provided under an overhead heat lamp (average basking temperature of 28C). Fluorescent room lights were kept on a 12-hour light and 12-hour dark cycle. Pre-inoculation Sample Collection Upon arrival, each turtle was examined for the presence of any clinical signs of illness. Urine, oral and cloacal swabs were obtained from each turtle and tested for the presence of iridovirus utilizing a polymerase chain reaction (PCR) test (see below). Blood samples were collected from each turtle into 2ml tubes coated with lithium heparin and centrifuged at 4,500 x g for 5 min. Buffy coats were removed and also tested for iridovirus using PCR. Plasma was removed and tested for antiRanavirus antibodies using an indirect enzyme linked imm unosorbent assay (ELISA; see below). DNA Preparation, Polymerase Chain Re action and Nucleotide Sequencing Oral and cloacal swabs were combined for each turtle into one 1.5ml microcentrifuge tube and 100 l of phosphate buffered saline was added. DNA was extracted from buffy coats, oral and cl oacal swabs using the DNeasy kit (Qiagen, Valencia, CA, USA) as were tissue samples later collected at necropsy. Viral DNA from

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68 urine samples was extracted using the QIAmp Ul trasens virus kit (Qiagen). Sense primer (5-GACTTGGCCACTTATCAC-3) a nd anti-sense primer (5GTCTCTGGAGAAGAAGAA-3), were used as previously described (Mao et al ., 1997) to amplify approximately 500 basepairs of the Ranavirus major capsid protein gene. A 20 l reaction mixture was run which contained 2 l extracted DNA, 1 M of both primers, 200 M each of dATP, dCTP, dGTP, and dTTP, 1.5 U of Taq DNA polymerase and PCR buffer containing 50 mM KCl, 10 mM Tris-HCl, 1.5 mM MgCl 2 (Eppendorf, Westbury, New York). The mixtures were amplified in a thermal cycler (PCR Sprint, Thermo Hybaid) with an initial denaturation at 94 C for 2.5 min, followed by 25 cycles of denaturation at 94 C for 30 sec.; annealing at 50 C for 30 sec, extension at 72 C for 30 sec., and a final extension step at 72 C for 10 min as previously described (Marschang et al ., 1999). The PCR products were reso lved in 1% agarose gels and any bands were excised and purified using the QIAquick gel extraction kit (Qiagen). Products were sequenced in both directions directly using the Bi g-Dye Terminator Kit (Perkin-Elmer, Branchburg, New Jersey ) and analyzed on ABI 377 automated DNA sequencers at the University of Florida s Sequencing Center. The sequences were compared to known sequences in GenBank (National Center for Biotechnology Information, Bethesda, Maryland), EMBL (Cambridge, United Kingdom), and Data Bank of Japan (Mishima, Shiuoka, Japan) databases using TBLASTX (Altschul et al ., 1997). ELISA An indirect enzyme linked immunosorbent assay (ELISA) was used to determine the presence of antiRanavirus antibodies. The ELISA met hodology was similar to that developed for use in identifying the presence of anti-tortoise herpesvirus antibodies in

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69 tortoises (Origgi et al ., 2004) and anti-west Nile virus antibodies in alligators (Jacobson et al ., 2005). The BSTRV isolate was used as th e antigen in the assay. Each well of a 96 well high-protein binding micropl ate was coated overnight at 4 o C with 50 l of a 1:400 dilution of either an uninfected lysate from Terrapene heart cells (TH-1, ATCC-CCL 50, American Type Culture Collection, Rockville MD) or TH-1 cell lysate from cells infected with BSTRV. Lysates were dilu ted in 0.01 M sodium phosphate buffer (pH7.2) containing 0.15 NaCl and 0.02% NaN 3 (PBS/A). Wells were then washed four times in ELISA wash buffer (PBS/A with 0.05% Tween-20). This washing process was repeated in between all of the following st eps. Wells were blocked with 300 l of Superblock blocking buffer (Pierce) for one hour at room temperature. All remaining steps were incubated for one hour at room temperatur e. Plasma samples were added in 50 l volumes at a 1:100 dilution in blocking buffer. The secondary antibody used was a biotin-conjugated mouse anti-tortoise i mmunoglobulin (Ig) monoclonal antibody diluted to a final concentration of 0.5 g/ml in blocking buffer. Alkaline phosphataseconjugated-streptavidin (Zymed Laboratories, Inc., San Francisco, CA) was then applied to each well at 50 L of a 1:5000 dilution in PBS/A. Next, 100 l of a 1.0 mg/ml Pnitrophenyl phosphate prepared in 0.01 M sodium bicarbonate buffer containing 2 mM MgCl 2 was added to each well and the plates were then stored in the dark until being read. The optical density (OD) of each well was read at A 405 using a StatFax 3200 microplate reader (Awareness Technology, Palm City, Florida, USA) after 30 minutes. Each sample was done in triplicate: on e time on wells coated with uninfected cell lysate and in duplicate on wells initially coat ed with infected cell lysate. The replicate values of the wells coated with BSTRV ly sate were averaged and divided by the OD

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70 reading of the value of the plasma sample r un on the uninfected cell ly sate to subtract out any background binding that might be caused by cross-reactivity to th e cells. Values greater than 2 were considered positive (Jacobson et al ., 2005), suggesting previous exposure to the virus, and would preclude use of that turtle in th e study. Plasma from a surviving penmate of the above Burmese star tortoise was used as a positive control. Virus Preparation Terrapene heart cells (TH-1) were acquired from the American Type Culture Collection (ATCC-CCL 50; Rockville, MD ) and grown to confluency in 225cm 2 tissue flasks (Costar, Corning, NY). Cells were cult ured in Dulbecco's modified Eagle medium (DMEM, Gibco, Carlsbad, CA) supplemented with 5% fetal bovine serum (Gibco), gentamicin (60mg/liter; Sigma, St. Loui s, MO), penicillin G (120,000 U/liter), streptomycin (120,000 U/liter) and amphotericin B (300g/liter; Sigma). Cells were inoculated with a fourth passa ge of BSTRV and incubated at 28C in the presence of 5% CO 2 When cytopathic effect (CPE) was observed, consisting of cell rounding and detachment from the flask in over 70% of ce lls, the flasks were scraped and contents transferred to 15 ml centrifuge tubes and clarified by slow speed centrifugation at 4,500xg for 30 minutes. The supernatant wa s then discarded and the cell pellet resuspended in 10mls of cell culture media. The preparation was then vortexed, frozen and thawed three times to release virus from the cells into the supernatant. The preparation was again clarified by centrifuging for thirty minutes. The supernatant was then transferred to a new tube and the cell pellet was discarded. The live virus in the media was then quantified by a TCID 50 assay, diluted with media to create a concentration of 10 5 TCID 50 /ml, and frozen at -80C until use.

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71 Transmission Studies Study 1 Study 1 was designed as a pilot study to de termine the suitabili ty of either redeared sliders (RES) or western orna te box turtles (BT) as a model of Ranavirus infection for chelonians. Three RES and three BT we re included in the study and each turtle was allowed to acclimate for two weeks prior to infection. One of each species was assigned to one of three groups: 1) a mock inocul ated control group, with both turtles receiving 0.5ml uninfected cell lysate ora lly and 0.5ml by intramuscular in jection, 2) an orally (PO) inoculated group with both turtles receiving 1ml of infected cell lysa te containing virus at 10 5 TCID 50 /ml by metal gavage feeding tube in the caudal esophagus and 3) an intramuscularly (IM) inoculated group, with both turtles rece iving the same concentration of virus as the PO inoculated group with 0.5ml injected into both the left and right pectoral muscles. Turtles were observed da ily after inoculation for the duration of the study. Oral swabs and buffy coats were collect ed 1-week post inocul ation for evaluation by PCR for the presence of iridovirus. DNA extraction and PCR were performed as described previously for prescreening. Phys ical examinations were performed daily to assess the presence of the follo wing clinical signs: lethargy, anorexia, cervical edema, palpebral or periocular edema, ocular discharge, nasal disc harge, oral discharge, the presence of oral plaques or any other abnormalities. Turtles were euthanized if clinical signs of disease became severe or at two weeks post-inoculation. For euthanasia, ketamine was administered intramuscularly at 100 mg/kg followed by intravenous sodium pentoba rbital. Once turtles were unresponsive to painful stimuli and showed no corneal refle x, they were decapitated and a complete necropsy performed. Portions of tongue, es ophagus, stomach, small and large intestine,

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72 liver, kidney, and spleen were collected as eptically by flame sterilizing tools between each organ, and frozen at -80C for virus is olation and/or DNA extraction and PCR. The following tissues were collected and fixed in 10% neutral phosphate buffered formalin: tongue, esophagus, stomach, small and large in testine, liver, kidne y, spleen, pancreas, heart, trachea, lung, brain, urinary bladde r, thyroid gland, adrenal gland, bone, bone marrow, skin, skeletal muscle, nasal cavit y, eye and gonad. These tissues were then processed for histologic examination. Th ey were embedded into paraffin, and 6 m sections were stained with hematoxylin and eosin. Virus isolation was performed on Terrapene heart cells seeded into 25 cm 2 flasks (Costar, Corning, NY). Cells were culture d in Dulbeccos modified Eagle medium/F12 (Invitrogen, Carlsbad, CA) supplemented with 5% fetal bovine serum (Gibco, Carlsbad, CA, USA), gentamicin (60 mg/liter) (Sigma, St Louis, MO, USA), penicillin G (120,000 U/liter), streptomycin (120,000 U /liter) and amphotericin B (300 g/liter) (Sigma) and cultured to confluency. A small piece of kidney (approximately 50mg) collected aseptically at necropsy was homogenized in separate 5ml tissue grinders containing DMEM. The homogenate was passed through a 0.45 m filter (Costar) onto a flask of cells. Cells were incubated at 28C and obser ved daily. Flasks of cells were harvested when CPE was observed in over 70% of cells or at post-inoculation day 10. DNA was extracted from cells using the DNeasy kit (Qiagen) protocol for animal cells. Polymerase chain reaction and nucleotide sequencing was performed as previously described to confirm the presence of iridovirus DNA sequences.

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73 Study 2 Based on the results of the pilot stud y, a second transmission study was designed that consisted of ten RES. Turtles were as signed to one of the following three groups: 1) control; 2) IM inoculated or 3) PO inoculated. Turtles were numbered from one to ten according to increasing weight. They were th en blocked into two groups (1-5 and 6-10) and turtles within each block were randomized to 1 control, 2 PO i noculated (turtle Nos. 1, 2, 7 and 10) and 2 IM inoculated (turtle Nos. 3, 5, 6 and 8). The control turtle in the lower weight block (turtle No. 4) was PO mo ck inoculated with 1ml of uninfected cell lysate, while the control turtle in the higher weight block (turtle No. 9) received 1ml IM, half in each pectoral muscle. Turtles were monitored daily for the presence of clinical signs consistent with iridovirus infection. Or al and cloacal swabs were taken three times a week throughout the four-week study. Fr ee catch urine samples were collected opportunistically at the same time periods. Tur tles were euthanized wh en clinical signs of infection became severe or four weeks post-inoculation. Necropsies, tissue collections and virus is olation were performed as described in Study 1. In addition, spleen from one IM inoculated (No. 6) and one control (No. 9) turtle were collected in Trump's solution (4% formaldehyde, 1% glutaraldehyde in a phosphate buffer; Electron Microscopy Sciences, Hatfield, PA) for transmission electron microscopy (TEM). Tissues were embedded in Spurrs resin and ultrathin sections were obtained and stained with lead citrate and uranyl acetate for TEM at the Electron Microscopy Core Laboratory, University of Florida.

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74 Results Experimental Animals and Pre-inoculation Sampling Upon arrival, all turtles were considered healthy. Using PCR, all oral swabs, cloacal swabs, urine and buffy coats collect ed from turtles in Studies 1 and 2 were negative for iridovirus DNA sequences. All plasma samples evalua ted by ELISA were below the positive cutoff value, indicating that the turtles were seronegative for exposure to Ranavirus Transmission Studies Study 1 Control mock inoculated and PO inocul ated red-eared sliders and box turtles showed no clinical signs of disease throughout the two-week study. The IM inoculated RES and BT both showed severe clinical signs. The RES di ed 9 days post inoculation (DPI). At 8 DPI the IM inoculated RES wa s basking continuously and started showing signs of lethargy, cutaneous erythema, and kept its palpebrae closed. It was found dead on the basking platform on the morning of 9 DP I. The orally inoculated RES remained normal. At 8 DPI, the IM inoculated BT de veloped a white opaque ocular discharge (Fig. 4-1), at ten days became lethargic and anor exic, and at 12 DPI, was euthanized. The orally inoculated BT remained normal. Both IM inoculated turtles in the pilo t study were positive by PCR for iridovirus sequences using DNA extracted from oral sw abs and buffy coats collected one week post-inoculation. Oral swabs and buffy coats collected on the PO inoculated turtles and mock-inoculated control turtles were nega tive. PCR performed on DNA extracted from both IM inoculated turtles were positive for a ll eight tissues (Table 1). PCR was negative for all eight tissues of both control turt les and both PO inoculated turtles.

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75 The most consistent and si gnificant histologic lesions were observed only in the spleens (2/2) of IM inoculated BT and RES, both having similar changes. Changes were based on the normal histology of the spleen th at was previously defined for red-eared sliders (Kroese and Van Rooijen, 1982). Lesi ons were centered on th e splenic ellipsoids or sheathed capillaries and will be described in detail for turtles in Study 2 (below). Briefly, the walls of the ellipsoids were moderately to markedly expanded by homogenous to slightly fibrilla r eosinophilic material (fibrin; fibrinoid vasculitis) with multifocal infiltrates of low numbers of heterophils. There were occasional lumenal fibrin thrombi with admixed hetero phils and karyorrhectic debris. Kidney samples from both IM inoculated tu rtles (P3 and P6) that were coated onto TH-1 cells demonstrated cytopa thic effects of cell rounding and lysis within two days of incubation. Intracytoplasmic inclusion bodies were observed in infected cells and PCR and nucleotide sequencing on DNA extracted from cells from each flask were positive for iridovirus. None of the cultu res inoculated with tissues collected from PO inoculated turtles, or control turtles demonstrated any CPE up to ten days post-inoculation of cells and each were negative by PCR for the presence of Ranavirus DNA segments. Study 2 Similar to Study 1, only IM inoculated RES showed clinical signs of disease and were euthanized before the end of the four-w eek study. Three of the four IM inoculated turtles showing severe clinical signs were euthanized on days 11, 13 and 23 DPI. All three turtles became anorectic, and extremely lethargic. The turtle euthanized 13 DPI developed oral plaques on the roof of the m outh and tip of the tongue (Fig. 4-2). Turtle 3, euthanized 23 DPI, exhibited exophthalm us, conjunctivitis and hyphema (Fig. 4-3). All three turtles had clear serous ocular and nasal discharge. The fourth IM inoculated

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76 turtle (No. 8) showed ocular discharge and s ubjectively basked more than the other turtles between 16 and 25 DPI, but then recovere d and showed no clinical signs at the termination of the study. It became anor ectic 14 DPI and remained so throughout the study. Three of 4 PO inoculated turtles al so became anorectic after inoculation and remained so throughout the study. No other signs of disease were noted in those turtles. Turtles mock inoculated were negativ e by PCR on all oral and cloacal swabs collected (Table 2). Three tu rtles PO inoculated (Nos. 2, 7 and 10) were positive by PCR on oral and/or cloacal swabs 2 DPI but not in any subsequent samples. The fourth PO inoculated turtle (No. 1) was not positive on any sample date. Oral and cloacal swabs from IM inoculated turtles were positiv e by PCR for iridovirus at varying times throughout the study (Table 2). Turtle No. 8 (IM inoculated), which was euthanized at the end of the study, was never positive on any oral swabs collected, but was positive on cloacal swab on five occasions including the last two sampling dates that were 23 and 26 DPI. PCR on urine samples followed a simila r pattern. Three of four IM inoculated turtles were positive between one and five days prior to being euthanized (Table 3). One positive band from each turtle was sequenced to confirm the positive PCR results. All amplicons were of the expected size and each sequence shared 100% identity with the sequence of the original isolate. DNA extracted from eight tissues from each turtle including tongue, esophagus, stomach, small inte stine, large intestin e, kidney, spleen and liver were positive by PCR on three of four IM inoculated turtles (Nos. 3, 5 and 6) at necropsy (Table 1). Tissues from IM inocul ated turtle No. 8, and all PO inoculated turtles and control mock-inoculat ed turtles were negative.

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77 At necropsy, gross changes were observed in several turtles. Turtle No. 3 that was IM inoculated exhibited petechia in several organs including the glottis, liver, pancreas and fat. Congestion was observed in the stom ach and on the surface of the bladder. The cecum and colon demonstrated a multifocal to coalescing area of hemorrhage (Fig. 4-4). The gastrointestinal tract of turtles 3, 5 and 6 appeared thickened and edematous. Turtle 6 also exhibited petechia on th e surface of the pancreas and congestion in the stomach. No lesions were seen in PO i noculated and control group turtles. Significant histologic lesions were observed only in IM in oculated turtles. Similar to the pilot study, the most consistent lesions we re in the spleen (3/4 turtles) and centered on the splenic ellipsoids. The majority of the splenic white pulp surrounded ellipsoids (periellipsoidal lymphocyte sheath; PELS) (Fi g. 4-5A and 4-6A) with lesser white pulp surrounding arterioles (periarteriolar lymphocyt e sheath; PALS) (Fig. 4-6B). Ellipsoids (Fig. 4-6A) were characterized by plump, cuboidal, endothelial cells, a thick eosinophilic wall lacking smooth muscle (confirmed with Massons trichrome stain), and a lack of reticular fibers between lymphocytes of th e PELS using Gordon and Sweets reticulin stain. In IM inoculated turtles, the walls of the ellipsoids were moderately to markedly expanded by homogenous to slightly fibrilla r eosinophilic material (fibrin; fibrinoid vasculitis) (Fig. 4-5B and 4-6B) with multifocal infiltrates of low numbers of heterophils and scattered free brown pigment gra nules (presumptively from disrupted melanomacrophage centers). There were occasional lumenal fibrin thrombi with admixed heterophils and karyorrhectic debris (Fig. 4-6B). There was mild to moderate lymphoid depletion and dispersi on of lymphoid cells in the P ELS with relative sparing of

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78 lymphocytes in the PALS (Fig. 4-5B and 4-6B ). Replacing the PELS were combinations of karyorrhectic debris, heteroph ilic infiltrates, extravasated erythrocytes and fibrin. The liver of all turtles had mild to marked hepatocellular vacuolar change that was most pronounced in the control turtles (F ig. 4-7A) and considerably less prominent in IM inoculated turtles (depletion of hepa tocellular lipid and gl ycogen) (Fig.4-7B). Three (No. 3, 5, 6) of four tu rtles had multifocal random dilatation of hepatic sinusoids with fibrin thrombi (Fig. 4-7B) and variable single-cell necrosis of adjacent hepatocytes (Fig. 4-7C). Rarely, hepatocytes had small intracytoplasmic basophilic inclusion bodies (Fig. 4-7C). Admixed with fi brin thrombi and necrotic hepatocytes were small amounts of karyorrhectic debris, infiltrates of small numbers of heterophils, and for lesions occurring adjacent to melanomacrophage centers, small amounts of dispersed brown granular pigment. One turtle (No. 3) had moderate multifocal hemorrhage in association with liver lesions as well as moderate multifocal fibrin thrombi within small to medium portal venules and veins. In addition to vascular changes in sple nic ellipsoids, hepatic sinusoids and portal blood vessels, acute fibrin thrombi were also observed in a variety of other tissues in intramuscularly inoculated animals (Fig. 4-8 a nd 4-9). Thrombi were noted in gastric or intestinal lamina propria, submucosa and sero sa (No. 3, 5, and 6), glomerular capillaries (No. 3 and 6), esophagus (No. 3), pulmonary capillaries and veins (No. 3 and 6), meninges (No. 3 and 6), eye (No.3), nasal muco sa (No. 3), and oral mucosa (No. 3). Lesions associated with thrombi included segmental marked colonic mucosal hemorrhage (Fig. 4-9), multifocal mild to moderate meningeal hemorrhage, mild to moderate heterophilic meningitis, and mild heterophilic interstitial pneumonia.

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79 A small number of IM inoculated turtles also had multifocal mild to moderate ulcerative and heterophilic stomatitis and es ophagitis (No. 6) (Fig. 4-10), and multifocal mild to moderate heterophilic and necrotizi ng gastritis (No. 5 and No. 6). Turtles from all three groups had small granulomas in a wide variety of tissues that surrounded probable spirorchid-type trematode eggs a nd rarely, adult trematodes were observed within mesenteric blood vessels. Effete gra nulomas in the gastric submucosa and serosa of a small number of control and virus inocul ated turtles contained cross-sections of an unidentified nematode. Trematodes and nema todes were interpreted as incidental findings. Similar to study 1, cytopathic effects c onsisting of cell rounding and lysis were seen in cultures of Terrapene heart cells that were coated with kidney homogenates from three of four IM inoculat ed turtles (Nos. 3, 5 and 6). Using PCR and nucleotide sequencing, Ranavirus was identified in DNA extracted from cells from each flask. No CPE was seen in cells that received tissue hom ogenates from orally inoculated turtles, control turtles, and one of the intram uscularly inoculated turtles (No. 8). Transmission electron microscopy of splenic ellipsoids in 1 IM inoculated turtle (No. 6) in the full transmission study demonstr ated marked expansion of the vessel wall by a granular to fibrillar lightly electron dense material that was consistent with fibrin. Admixed with fibrin and within remnants of the white pulp were scattered unidentified necrotic cells with intracytopl asmic arrays of icosahedral virions consistent with an iridovirus. No virions were observed in cells associated with ellipsoids in the control turtle (No. 9).

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80 Discussion Henle-Koch's postulates provide a strict set of guidelines for proving a causal relationship between an infectious agen t and a disease (Evans, 1976). Here we performed a transmission study using a Ranavirus isolate from a captive Burmese star tortoise that became ill and died in an attemp t to determine if a causal relationship exists between infection with this Ranavirus and the clinical and histologic changes observed in the Burmese star tortoise. Si nce it is not practical to pe rform a challenge study in this critically endangered species, we decided to assess both box turtles and red-eared sliders as a model for Ranavirus infection in chelonians. Box turtles were selected since Ranavirus infection has been identified in this species (Mao et al ., 1997; Devoe et al ., 2004; Johnson et al ., 2004). We decided to also eval uate the suitability of red-eared sliders since populations of box turtles are d eclining throughout their range. Red-eared sliders, however, are being raised in the lower Mississippi Valley for the overseas pet trade, which became a factor in their ultimate selection as an experimental animal in our studies. In addition, results of the pilo t study showed that both species similarly responded when administered a Ranavirus isolate by two different routes. Both IM inoculated turtles showed se vere clinical signs and were euthanized prior to the termination of the study at tw o weeks. Oral swabs from both were positive one week post-inoculation and histologic le sions were consistent between the two species. Both the PO inoculated RES and BT did not show any cl inical or histologic le sions. Results of this study showed that both RES and BT can serve as suitable models of Ranavirus infection for chelonians. As a result, RES we re chosen as the experimental model for the larger transmission study.

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81 Histologic lesions in turtles inoculated by intramuscular injection in both studies consisted primarily of multicentric fibrinoid va sculitis and formation of fibrin thrombi in small blood vessels in numerous tissues, and in this regard, resembled recent reports of Ranavirus infection in captive and free-rangi ng box turtles and tortoises (DeVoe et al ., 2004; Johnson et al ., 2004). Lesions in blood vessels were consistent with observed Ranavirus infection of endothelial cells in a naturally-infected captive Burmese star tortoise (Johnson et al ., 2004) and with descriptions of apparent viral endotheliotropism in rainbow trout and redfin perch infected with another Ranavirus called epizootic haematopoietic necrosis virus (EHNV) (Re ddacliff and Whittington, 1996). Involvement of endothelial cells may also be part of the pathogenesis of some amphibian Ranavirus infections as suggested by multicentric hemorrhage and edema or observation of characteristic inclusion bodies within endothelial cells (Wolf et al ., 1968; Cunningham et al ., 1996; Bollinger et al ., 1999; Docherty et al ., 2003). The consistent involvement of the splenic ellipsoids (sheathed capillaries) provides a basis for the prominent necrotizing splenitis observed in some natural Ranavirus infections of chelonians (DeVoe et al ., 2004; Johnson et al ., 2004). Similar lesions of the ellipsoids were observed in redfin perch, but not rainbow trout, experimentally infected with EHNV (Reddacliff and Whittington, 1996) A filtering function for splenic ellipsoids for particulate ma terial and immune complexes has been documented in other species (Sorby et al ., 2005) and possibly, the lesi ons observed in chelonian Ranavirus infections could be a conseque nce of antigen trapping in m acrophages associated with the ellipsoid sheath during rana viral viremia.

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82 Necrotizing stomatitis, esophagi tis and/or pharyngitis are ch aracteristic lesions in many naturally occurring chelonian Ranavirus infections (Westhouse et al ., 1996; Marshang et al., 1999; DeVoe et al ., 2004; Johnson et al ., 2004). Clinical signs such as ocular and nasal discharge, conjunctivitis and palpebral edema associated with Ranavirus infection (both naturally infect ed animals and the experimental animals in this report) are often attributed to the upper respiratory trac t and overlap with thos e signs observed with mycoplasmosis caused by Mycoplasma agassizii (Brown et al ., 1999) or herpesvirus infection (Origgi, et al ., 2004; Johnson et al ., 2005). In particular, infection with tortoise herpesviruses 1 and 2 are associated with caseous or diphtheritic oral plaques that are grossly indistinguishable from oral lesions associated with Ranavirus infection. In the experimentally infected animals of this report, necrotizing stomatitis and esophagitis were observed in a single intramuscularly inocul ated red-ear slider (N o. 6). Lesions were not observed in the nasal cavity of any an imal examined. Oral lesions could be secondary to thrombus formation and infa rction in small submucosal vessels or alternatively, could be the resu lt of viral infection and necros is of oral epithelial cells. The observation of intracytoplasmic inclusion bodies consisting of Ranavirus in epithelial cells of some naturally occurring infec tions would appear to support the latter explanation. Why oral lesions were not present in more of the experimentally inoculated turtles in this report is uncertain, but it is po ssible that epithelial infection is a late event that follows viremia and hence was not observed in experimental animals. Also, the virus may have been attenuated in cell culture resulting in alte red pathogenicity. The intracytoplasmic basophilic inclusion bodies that are suggestive of iridovirus infection and prominent in many cases of Ranavirus infection in fish, amphibians and

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83 some chelonians were not prominent in the ex perimentally inoculated turtles (Reddacliff and Whittington, 1996; Westhouse et al ., 1996; Bollinger et al ., 1999; Marschang et al ., 1999; Docherty et al ., 2003). This observation is c onsistent with recently reported naturally occurring chelonian Ranavirus infections, and indicate s that inclusion bodies may be an inconsistent finding and should not be relied upon for use in formulating a histologic differential diagnosis (Devoe et al ., 2004; Johnson et al ., unpublished findings). Virions consistent with ranavi ruses were observed by transmission electron microscopy in cells within the sp leen of an IM inoculated re d ear slider and suggests that TEM may still be a useful diagnostic tool in chelonian Ranavirus infections even in the absence of visible inclusions on histologic se ction. Demonstration of intracytoplasmic virions in cells of an experimentally inoculat ed turtle is important because it shows that the virus is capable of entering and replica ting within cells, and that, lesions were not induced by the presence of inoculated non-repl icating virus. Necrotizing liver lesions have been experimentally induced in mice and rats following injection of inactivated iridovirus virions or solubilized structural proteins (Lorbacher de Ruiz, 1990). Similarly, the Ranavirus Frog Virus 3 can trigger apoptosis in tissue culture in th e absence of viral gene expression (Chinchar, 2002). Fu ture studies may better define the in-vivo mechanism of cell death associated with iridovirus infections. This study found that IM inoculated turtle s were more likely to become infected with Ranavirus show clinical signs and die compar ed to turtles that were orally inoculated. All four IM inocul ated turtles showed clinical si gns and three died as a result of infection (75%), whereas no orally inoculated turtles show ed any signs of disease or died. This could mean that turtles do not become exposed through ingestion of infected

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84 animals or water sources as has been pr eviously shown to occur in amphibians (Jancovich et al ., 2001, Pearman et al. 2004), or that abrasions naturally acquired from ingesting bones or other abrasi ve material may be necessary for virus to be introduced systemically. Another explanation for the inability to re-create disease in orally inoculated animals was that a natural expos ure was not replicated in the laboratory setting. Viral dose administered was extrapolated from studies done with fish and amphibians (Langdon, 1989, Moody and Owens, 1994, Bollinger, 1999, Cullen and Owens, 2002) but requirements for infection of turtles may be higher or repeated exposure may be necessary. Experimental infections of salamanders with a Ranavirus showed that both dose and host characteristic s influenced the virulence of infection (Brunner et al ., 2005). The infection dose was positively correlated with the mortality rate and inversely related to average survival times. Environmental temperatures have also been shown to significantly impact th e percent mortality and time to death in salamanders experimentally inoculated with a Ranavirus (Rojas et al., 2005), where salamanders inoculated at 18 and 10C were more likely to die than those exposed at 26C. While water and room temperatures averaged between approximately 21 and 25C, basking areas were kept warmer at 28 C. Eliminating heat lamps over basking areas and lowering the room temperature might have kept turtles c ooler, and altered the results in the orally inoculat ed group. Alternatively, other ro utes of transmission such as vector-borne transmission may be required for turtles to become infected in the wild. Intracytoplasmic inclusion bodies were recently identified in the circulating leukocytes of an eastern box turtle infected with iridovirus (Allender et al ., In Press). Ranaviruses are variably host specific, so virus may be able to survive in mosquitoes or other biting

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85 insects capable of transmitting virus from an infected animal to an uninfected one. This has been shown to be the case with ins ect iridoviruses, wh ere parasites were experimentally shown capable of transmitting invertebrate iridescent virus from infected to uninfected larvae (Lopez et al ., 2002). It remains unknown whether natural outbreaks of iridovirus infections in any vertebrate species o ccur as a result of in troduction of novel virus strains, recrudescen ce of latent or persistent infec tions in surviving populations as a result of stressors or other immunosuppressive causes, or viral persistence in the environment (Williams et al ., 2005). Although the mechanism of transmission of iridoviruses in natural settings is unknown, it has been shown that sublethally infected amphibians can cause sporadic, recurrent disease outbreaks in amphibians (Brunner et al ., 2004). Experimentally and naturally infected tiger salamander larvae and metamorphs were able to maintain sublethal, transmissible infections for over five months. Apparent ly healthy infected dispersing metamorphs were returning to wate r bodies to breed and it was speculated that these individuals were likely serving as a reservoir host for infecting newly hatched larvae, creating recurrent outbreaks of disease. The current study show ed that turtles may also become asymptomatic carriers, although further studies would he lp to confirm this finding. Turtle No. 8 which was inoculated intramuscularly showed transient signs of disease but then recovered. At necropsy, ti ssues collected from eight different organs were negative for iridovirus on PCR. Kidney samples inoculated onto TH-1 cells showed no cytopathic effects. However, cloacal swab s collected one, four and eight days prior to necropsy were positive using PCR. If this wa s a result of slow elimination of the virus, all orally inoculated turtles should have ha d PCR-positive cloacal samples after 2 DPI.

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86 However, all swabs taken after this time peri od were negative. Furthermore, given that IM inoculation was used in turtle No. 8, any virus shed from the GI tract would require transportation through the blood st ream and across the wall to be there. While it is possible that this was a laboratory contaminan t, all swabs were extracted and tested by PCR according to date collected as opposed to all swabs tested at the same time per turtle. If there was contamination, we would not expect to see it on five occasions from one turtle and not from any orally inoculated turtles on any date past 2 DPI. Therefore it seems more likely that virus shedding was occu rring, and the site of virus replication was missed. Some viruses show a predilection for specific cells, such as infectious bursal disease virus in chickens, where virus shows a predilection for the cells of the bursa of Fabricius, located in the cloaca (Burkhardt and Mller, 1987). Further long-term studies would help to confirm whether Ranavirus persistence occurs a nd immunohistochemical or in situ hybridization studies of tissues from infected turtles may help identify the tissue tropism for Ranavirus persistence. If turtles can serv e as asymptomatic carriers, they may also serve as a reservoir host of virus for other turtles and other susceptible species. Inter-class infections of iri dovirus have been shown naturally and experimentally in sympatric species of fish and amphibians, where both were capable of being infected with the same virus (Mao et al ., 1999, Moody and Owens, 1994). In another study (data not shown), the isolate used in this experimental study was capable of infecting leopard frogs ( Rana pipiens) that were injected intraperitoneally at the same dose. Thus, sublethally infected turtles such as one of the turtles (No. 8) in our study could serve as a reservoir host for amphibian populations in geographi c locations where the species overlap.

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87 In summary, the experime ntal inoculation of a Ranavirus in chelonians was investigated. Koch's postulates were fulfille d when intramuscular inoculation of virus into naive turtles resulted in clinical and hist ologic changes consistent with those seen in natural infections, and when the same virus was subsequently recovered. Since oral inoculation failed to result in disease or mort ality, the natural route of transmission in the wild remains unknown. The immune system of reptiles is temperature dependent (Cooper et al ., 1985) and perhaps by manipulating th e environmental temperature, and the temperature of the host, the susceptibilit y of turtles to infection can be altered. Another possibility that shoul d be investigated is the tr ansport and inoculation of Ranavirus into chelonians by arthropods. This study also suggests that sublethally infected turtles may serve as reservoir hosts of infection for other chelonians, as well as amphibians. Ranaviruses are considered a gl obal threat to amphibi an populations based on the lack of host specificity, high vi rulence and global distribution (Daszak et al ., 1999) and this study confirms that they should lik ewise be considered a threat to chelonian populations.

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Table 4-1. PCR results on tissues collected at necr opsy from the pilot st udy box turtles (BT; Terrapene ornata ornata ) and red-eared sliders (RES; Terrapene scripta elegans ) in the full transmission studies. Controls Orally Inoculated Turtles IM Inoculated Turtles Pilot Study Full Study Pilot Study Full Study Pilot Study Full Study Tissue P1 (BT) P4 (RES) 4 9 P2 (BT) P5 (RES) 1 2 7 10 P3 (BT) P6 (RES) 3 5 6 8 Liver + + + + + + Kidney + + + + + + Spleen + + + + + + Tongue + + + + + + Esophagus + + + + + + Stomach + + + + + + Small Intestine + + + + + + Large Intestine + + + + + + 88

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Table 4-2. Polymerase chain reaction (PCR) results of oral and cloacal swabs taken on eleven different days post-inoculation (DPI) with an uninfected (control) or iridovirus infected cell ly sate throughout the four-week fu ll transmission study. O = oral, C=cloacal. a "N/A", indicates the turtle was eu thanized prior to the sample date. Oral Control IM Control Red ear sliders inoculated orally Red ear sliders inoculated intramuscularly I.D. 4 9 1 2 7 10 3 5 6 8 DPI O C O C O C O C O C O C O C O C O C O C 0 2 + + + + + + 5 + 7 + + + + 9 + + + + + 12 + + N/A N/A + + 14 + + N/A N/A N/A N/A 16 + + N/A N/A N/A N/A 19 + + N/A N/A N/A N/A + 21 + + N/A N/A N/A N/A 23 + + N/A N/A N/A N/A + 26 N/A a N/A N/A N/A N/A N/A + 89

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90 Table 4-3. PCR results for urine collected opportunistically from turtles in the full transmission study. Controls Orally Inoculated Turtles IM Inoculated Turtles Days p.i. 4 9 1 2 7 10 3 5 6 8 0 N/A N/A 5 N/A 7 N/A + N/A 8 N/A N/A N/A + 11 N/A N/A N/A N/A N/ A N/A N/A + N/A N/A 12 N/A N/A N/A N/A N/A + 15 N/A N/A N/A N/A 19 N/A N/A N/A N/A 23 N/A N/A N/A N/A N/ A N/A + N/A N/A N/A 27 N/A N/A N/A

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91 Fig. 4-1. Photograph taken 12 days post-inoculat ion showing development of white opaque ocular discharge in the IM inoculated box turtle ( Terrapene ornata ornata ).

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92 Fig. 4-2. Photograph taken 12 days post-inoculation show ing white caseous diphtheric plaques in the mouth of an IM inoculated red-eared slider ( Trachemys scripta elegans).

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93 Fig. 4-3. Photograph showing exophthalmus, conjunctivitis and hyphema in an intramuscularly inoculat ed red-eared slider ( Trachemys scripta elegans ).

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94 Fig. 4-4. Photograph showing colonic hemorrh age in a turtle intramuscularly inoculated with Ranavirus euthanized 23 days post inoculation.

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95 A B Fig. 4-5. Spleen; red-eared slider ( Trachemys scripta elegans ). A) Sham inoculated turtle. The white pulp is concentrat ed in discrete cuffs around splenic ellipsoids. H&E stain. Bar = 200 um. B) Turtle intramuscularly inoculated with Ranavirus There is lymphoid depletion and expansion of the walls of splenic ellipsoids by eosinophilic material. H&E stain. Bar = 200 u m.

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96 A B Fig. 4-6. Spleen; red-eared slider ( Trachemys scripta elegans ). A) Sham inoculated turtle. A higher magnification of tw o splenic ellipsoids and associated periellipsoidal lymphocyte sheathes (PELS). H&E stain. Bar = 50 um. B) Turtle intramuscularly inoculated with Ranavirus Higher magnification of two splenic ellipsoids and one splenic arteriole (arrow). The walls of the ellipsoids are expanded by fi brin and there are lumenal fibrin thrombi. Note the sparing of the splenic arteri ole and associated lymphoid sheath (periarteriolar lymphoid sheath (PALS). H&E stain. Bar = 50 u m.

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97 A B

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98 C Fig. 4-7. Liver; red-eared slider ( Trachemys scripta elegans ). A) Sham inoculated turtle. There is marked diffuse vacuolation of hepatocytes. A melanomacrophage center is present in the upper left. H&E stain. Bar = 100 u m. B) Turtle intramuscularly inoculated with Ranavirus There is depletion of normal hepatocellular lipid and glycogen. Sinusoids are multifocally expanded by fibrin thrombi with infilt rates of low numbers of heterophils. H&E stain. Bar = 100 um. C) Turtle intramuscularly inoculated with Ranavirus. There is single cell necrosis of hepatocytes with occasional intracytoplasmic basophilic inclusion bodi es (arrow). H&E stain. Bar = 20 u m.

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99 A B Fig. 4-8. Kidney; red-eared slider ( Trachemys scripta elegans). A) Sham inoculated turtle. Normal glomerulus. H&E stain. Bar = 20 u m. B) Turtle intramuscularly inoculated with Ranavirus There are fibrin thrombi in glomerular capillary loops admixed w ith few heterophils and small amounts of karyorrhectic debris H&E stain. Bar = 20 um.

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100 A B Fig. 4-9. Colon; red-eared slider ( Trachemys scripta elegans) intramuscularly inoculated with Ranavirus A) There is mucosal and superficial submucosal hemorrhage with marked submucosal edema. The arrow indicates a submucosal blood vessel. H&E stain. Bar = 1.0 mm. B) Higher magnificat ion of submucosal blood vessel shown in 45A. There is a lumenal fibrin thrombus w ith admixed karyorrhectic debris and mild perivascular hemorrhage. H&E stain. Bar = 100 u m.

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101 Fig. 4-10. Oral mucosa; red-eared slider ( Trachemys scripta elegans ) intramuscularly inoculated with Ranavirus There is focal mucosal ulceration and replacement by a mat of heterophils and fi brin. H&E stain. Bar = 200 um.

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CHAPTER 5 THE ROLE OF INFECTED LEOPARD FROGS ( Rana pipiens) IN TRANSMISSION OF A RANAVIRUS IN RED-EARED SLIDERS ( Trachemys scripta elegans) Introduction Emerging infectious diseases have been increasingly recognized as factors influencing wildlife health and populations (Harvell et al ., 1999; Daszak et al ., 2000). Iridoviruses in the genus Ranavirus are being observed at an increasing frequency in chelonians throughout the eastern United States (DeVoe et al., 2004; Johnson et al., 2004). While the route of transmission of this virus to chelonians has yet to be identified, it is suspected that amphibians may be a reservoir host. A Burmese star tortoise ( Geochelone platynota ) with clinical signs of illness (ocular, nasal and oral discharge, palpebral and cervical edema, and stomatitis) died at a captive breeding facility in Georgia. Molecular techni ques including polymerase chain reaction and nucleotide sequencing demonstrated the presence of Ranavirus major capsid protein genes in various tissues. Transmi ssion electron microscopy of tissues and cell cultures inoculated with tissue homogenate of kidney from this tortoise demonstrated the presence of virions consistent in sh ape and size with iridoviruses (Johnson et al., 2004). Tortoises in an adjacent pen were observed ingesting dead amphibians, and so, frogs and toads from the site of the tortoi se death were collected and euthanized to determine if they were infected with Ranavirus. One frog was found to be positive by PCR, with the amplicon sharing 100% identity across approximately 500 basepairs with the Burmese star tortoise isolate (BSTRV) as well as Frog Virus 3, the type species of the 102

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103 genus Ranavirus Histologic lesions cons istent with amphibian Ranavirus infection were observed in this frog (Johnson et al., 2004). Virus was isolated and whole viral genomic restriction utilizing two endonucl eases comparing the frog and tortoise isolates showed identical restriction patterns, s uggesting that they were either infected with the same or very closely related stra ins of the same virus. Experimental transmission studies fulfilling Koch's postulates were performed confirming the BSTRV isolate is a causa tive agent of disease and mortality in experimentally inoculated western ornate box turtles ( Terrapene ornata ornata ) and redeared sliders ( Trachemys scripta elegans ). Mortality was only observed when turtles were inoculated intramuscularly and not when turtles were inoculated orally. This is different than what has been shown with e xperimentally inoculated amphibians, where ingestion of infected water and infected amphi bian conspecifics resu lted in infection and increased mortality rates (Jancovich et al ., 2001, Pearman et al ., 2004). One hypothesis for why this occurred was that the natural ro ute of exposure was not being replicated in the laboratory. Turtles were inoculated with an infected cell lysa te by gavage feeding tube into the distal esophagus, when in a natural setting they would be ingesting bones and other organic matter that might cause natura l abrasions in the gastrointestinal tract, allowing a route of entry of the virus into the blood stream. Turtles in the wild might also be subjected to multiple exposures of virus, as opposed to a single dose as was administered in the previous transmission study. To attempt to establish a route of transm ission that might be occurring in natural settings, leopard frogs were experimental ly inoculated with BSTRV, euthanized, homogenized and fed to turtles in an attemp t to determine if ingestion of infected

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104 amphibians might be responsible for infecti on of chelonians in the wild. We evaluated infection by observing for clinical signs seen in chelonians infected with Ranavirus as well as seroconversion using an indirect ELISA Here we report the results of this study. Materials and Methods Virus Preparation A previously isolated an d partially characterized Ranavirus from a Burmese star tortoise (BSTRV; Geochelone platynota ; Johnson et al. 2004) was used to inoculate frogs in this study. Briefly, polymerase chai n reaction targeting a portion of ranaviral major capsid protein genes followed by DNA se quencing demonstrated that the BSTRV isolate shared 100% sequence identity with Fr og Virus 3, the type species for the genus Ranavirus in the family Iridoviridae. Restriction enzyme digests of BSTRV compared with FV3 showed identical restri ction patterns using two enzymes, HindIII and XbaI, indicating that BSTRV is either id entical or closel y related to FV3. Terrapene heart cells (TH-1) were acquired from the American Type Culture Collection (ATCC-CCL 50; Rockville, MD ) and grown to confluency in 225cm 2 tissue flasks (Costar, Corning, NY). Cells were cult ured in Dulbecco's Modified Eagle Medium (DMEM, Gibco, Carlsbad, CA) supplemented with 5% fetal bovine serum (Gibco), gentamicin (60mg/liter; Sigma, St. Loui s, MO), penicillin G (120,000 U/liter), streptomycin (120,000 U/liter) and amphotericin B (300g/liter; Sigma). Cells were inoculated with a fourth passa ge of BSTRV and incubated at 28C in the presence of 5% CO 2 When cytopathic effect (CPE) was observed, consisting of cell rounding and detachment from the flask in over 70% of ce lls, the flasks were scraped and contents transferred to 15 ml centrifuge tubes and clarified by slow speed centrifugation at 4,500xg for 30 minutes. The supernatant wa s then discarded and the cell pellet

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105 resuspended in 10mls of cell culture media. The preparation was then vortexed, frozen and thawed three times to release virus from the cells into the supernatant. The preparation was again clarified by centrifuging for thirty minutes. The supernatant was then transferred to a new tube and the cell pellet was discarded. The live virus in the media was then quantified by a TCID 50 assay, and frozen at -80C until use. Frog Pilot Study A pilot study was undertaken to determine the time post-inoculation that would be ideal for maintaining the highest titer of virus in infected frogs to be used in a feeding study. This and the following study were performed under the approval of the Institutional Animal Care and Use Committee at the University of Florida. Thirteen wild caught adult Rio Grande leopard frogs (Rana berlandieri ) were obtained from a biological supply company. Frogs averaged 72 grams and were placed in groups of three or four in plastic containers on damp moss. A shallow water dish was available for soaking. Oral and cloacal swabs were coll ected from each individual upon arrival and assayed for the presence of iridovirus us ing a polymerase chain reaction (PCR) as previously described (Mao et al ., 1997) and further described below. Once all frogs were determined to be negative, nine frogs in three containers were inoculated with the BSTRV isolate. Each frog received 0.5mls of a 10 6.4 TCID 50 /ml infected cell lysate in its ventral lymph sac (subcutaneously; Fig. 5-1). The container housing four frogs remained uninfected as controls for normal tissue for histopathology and to see if any of the four were positive on PCR once tissues were assayed. One container of three inoculated frogs each was euthanized on days 3, 7 and 14 post-inoculation, and the four controls were euthanized 14 days post-inoculation. Frogs were euthanized by an overdose of CO 2 followed by decapitation after a loss of righting

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106 abilities was observed and there was no corneal reflex or response to painful stimuli. A small piece of liver was collected and frozen at -80C for PCR, while another portion was placed in 10% phosphate buffered formalin for histopathology. A small piece of spleen was collected and frozen for virus isolation and determining virus titer differences between individuals and between post-inoculation days. Full Frog Study and Pre-inoculation Sampling Seventy-one captive ra ised leopard frogs ( Rana pipiens) were obtained from a biological supply company and placed in te n groups of seven or eight in plastic containers in an animal care f acility room at the University of Florida. Oral and cloacal swabs were collected from each frog, and combin ed into one tube. Frogs were identified only by container and not individually at this stage. DNA was extracted from the swabs using the DNeasy kit (Qiagen, Valencia, CA) and assayed by PCR for the presence of iridovirus. Sense primer (5-GACTTGGCCA CTTATCAC-3) and anti-sense primer (5GTCTCTGGAGAAGAAGAA-3) as pr eviously described (Mao et al ., 1997) were used to amplify approximately 500 base pairs of the Ranavirus major capsid protein gene. A 20 l reaction mixture was run which contained 2 l extracted DNA, 1 M of each primer, 200 M each of dATP, dCTP, dGTP, and dTTP, 1.5 U of Taq DNA polymerase and PCR buffer containing 50 mM KCl, 10 mM Tris-HCl, 1.5 mM MgCl 2 (Eppendorf, Westbury, New York, USA). The mixtures were amplified in a thermal cycler (PCR Sprint, Thermo Hybaid) with an initial denaturation at 94 C for 2.5 min, followed by 25 cycles of denaturation at 94 C for 30 sec.; annealing at 50 C for 30 sec, extension at 72 C for 30 sec., and a final extension step at 72 C for 10 min as previously described (Marschang et al ., 1999). Two frogs were pos itive upon arrival by PCR in

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107 containers 3 and 6. All seven frogs in contai ner six were euthanized as described for the pilot study and identified by individual and co ntainer (6-1 through 6-7). A portion of the spleen was removed on each and set up for P CR. The carcasses were refrigerated until the PCR results were determined. Frog 6-4 was found to be positive for iridovirus, and it was placed in formalin and submitted for histopathology. The remaining frogs were frozen at -80 C. Thirty frogs in containers 5, 7, 8, 9, and 10 were experimentally inoculated with 500 l of a10 5.75 TCID 50 /ml virus culture injected into th eir ventral lymph sacs. Frogs in containers 1-4 remained uninoculated. Seven days post-inoculati on all the frogs were euthanized by CO 2 as described for the pilot study. A small piece of liver was collected for PCR to ensure the frogs were either inf ected or that control frogs were uninfected. Organs were removed from each frog and stored frozen at -80C in sealed plastic storage bags until PCR could be performed. Inocul ated frogs that had strong positive signals by PCR and control frogs that were not inoculated and ne gative by PCR were used for feeding to turtles. Organs were separated and homogenized in a food processor. Four skeletons were added to each group to pr ovide bits of bone and objects normally encountered when eating a carcass. The homoge nized frogs were stor ed frozen at -80C in 35ml aliquots for feeding (slightly more th an the amount deemed necessary to feed all eight turtles in the feed ing group for one day). Turtles Sixteen red-eared sliders we re obtained from an animal dealer. Seven were adults (Nos. 1-7), while nine were juveniles (Nos. 8-16). Size differences were a result of availability at the time of the study. U pon arrival turtles were randomly assigned

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108 numbers within their size groups and anesthetized with intravenous propofol. A pharyngostomy tube was inserted through the ce rvical skin, and with the inserted end passed to the esophageal-gastric junction. Th e tube was held in place using a Chinese mattress suture pattern (Fig. 5-2). Feeding tube sizes were selected such that the largest tube, as deemed appropriate for the size of the turtle, was used in order to reduce the chance of the tube clogging and to allow larger particles of food such as bone bits to pass through the tubes. The seven adults were randomly assigned to one of two feeding groups, and the juveniles were similarly randomly separated into the two groups. Eight turtles (Nos. 1, 3, 4, 5, 8, 11, 12 and 13) in group 1 received virus inoculated frog homogenate, while eight turtles in group 2 (Nos. 2, 6, 7, 9, 10, 14, 15, and 16) received uninfected homogenized frogs. Aliquots of food were thawed within three hours of the time of feeding and mixed with equal volumes of water and mixed well. This was sufficient for passing through the tubes of the adult turtles using 10ml syri nges, but was too thick for passing through feeding tubes of juveniles. The food for j uveniles were mixed 2:1 of water to food, well mixed and then filtered through sterile gauze to filter out larger partic les. Turtles were then administered food with 3ml syringes. Tu rtles were fed 1% of their total body weight in food at each feeding, twice weekly for six weeks. Wate r was administered into the feeding tubes after food was give n to ensure the full frog meal was out of the tube and in the gastrointestinal tract a nd to keep the tubes patent. Physical examinations were performed daily to assess the presence of the following clinical signs: lethargy, anorex ia, cervical edema, palpebral or periocular edema, ocular discharge, nasal discharge, oral discharge, the presence of oral plaques or any other

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109 abnormalities. Any turtle found to have severe clinical signs of dis ease was euthanized. Blood was collected from the subcarapacial vein from each turtle prio r to inoculation and once a week for the three-month study. Blood was centrifuged and plasma collected for evaluation of production of antiRanavirus antibodies by ELISA. Full necropsies were performed on all turtles at the time of death, euthanasia or at the end of the three-month study. For eu thanasia, ketamine was administered intramuscularly at 100 mg/kg followed by intr avenous sodium pentobarbital. Once turtles were unresponsive to painful stimuli and show ed no corneal reflex, they were decapitated and a complete necropsy performed. Porti ons of tongue, esophagus, stomach, small and large intestine, liver, kidney, and spleen were collected aseptically by flame sterilizing tools between each organ, and frozen at -80 C for virus isolation and/or DNA extraction and PCR. The following tissues were collected and fixed in 10% neutral phosphate buffered formalin: tongue, esophagus, stomach, small and large intestine, liver, kidney, spleen, pancreas, heart, trachea, lung, brain, urinary bladder, thyroid gland, adrenal gland, bone, bone marrow, skin, skeletal muscle, nasal cavity, eye and gonad. These tissues were then processed for histologic examina tion. They were embedded into paraffin, and 6 m sections were stained with hematoxylin and eosin. Virus Titers A TCID 50 assay as previously described was performed on the homogenized frogs at both the 1:1 and the 2:1 dilutions, before the first feeding and then again at 2 weeks and 4 weeks to determine if a ny reduction in titer was occu rring as a result of being stored in the freezer over time. For both the 1:1 and 2:1 dilutions, 100 l was taken, diluted in cell culture media and syringe filtered using a 0.45 m filter (Costar, Corning,

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110 NY) into 9.9mls of media. This 1:10 dilution was then ten-fold serially diluted. One ml of each dilution was placed into f our wells of a 24 well plate from 10 -2 to 10 -7 The plates were incubated for 5 days at 28 C and then observed for the presence of CPE. The TCID 50 was then calculated for each type of food. ELISA An indirect enzyme linked immunosorbent assay (ELISA) was used to determine the presence of antiRanavirus antibodies. The ELISA met hodology was similar to that developed for use in identifying the presence of anti-tortoise herpesvirus antibodies in tortoises (Origgi et al ., 2004) and anti-west Nile virus antibodies in alligators (Jacobson et al ., 2005). The BSTRV isolate was used as th e antigen in the assay. Each well of a 96 well high-protein binding micropl ate was coated overnight at 4 o C with 50 l of a 1:100 dilution of either an uninfected lysate from Terrapene heart cells (TH-1, ATCC-CCL 50, American Type Culture Collection, Rockville MD) or TH-1 cell lysate from cells infected with BSTRV. Lysates were dilu ted in 0.01 M sodium phosphate buffer (pH7.2) containing 0.15 NaCl and 0.02% NaN 3 (PBS/A). Wells were then washed four times in ELISA wash buffer (PBS/A with 0.05% Tween-20). This washing process was repeated in between all of the following st eps. Wells were blocked with 300 l of Superblock blocking buffer (Pierce) for one hour at room temperature. All remaining steps were incubated for one hour at room temperatur e. Plasma samples were added in 50 l volumes at a 1:100 dilution in blocking buffer. The secondary antibody used was a biotin-conjugated mouse anti-tortoise i mmunoglobulin (Ig) monoclonal antibody diluted to a final concentration of 0.5 g/ml in blocking buffer. Alkaline phosphataseconjugated-streptavidin (Zymed Laboratories, Inc., San Francisco, CA) was then applied

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111 to each well at 50 L of a 1:5000 dilution in PBS/A. Next, 100 l of a 1.0 mg/ml Pnitrophenyl phosphate prepared in 0.01 M sodium bicarbonate buffer containing 2 mM MgCl 2 was added to each well and the plates were then stored in the dark until being read. The optical density (OD) of each well was read at A 405 using a StatFax 3200 microplate reader (Awareness Technology, Palm City, Florida, USA) after 30 minutes. Each sample was done in triplicate: on e time on wells coated with uninfected cell lysate and in duplicate on wells coated with infected cell lysate. The replicate values of the wells coated with BSTRV lysate were av eraged and divided by the OD reading of the value of the plasma sample run on the uni nfected cell lysate to subtract out any background binding that might be caused by cross-reactivity to the cells. This makes up the positive to negative ratio (P/N ratio). Ratios greater than 2.2 were considered positive as previously determined by taking the mean P/N ratio of 1000 free-ranging gopher tortoise samples plus three times th e standard deviation of the mean. Results Frog Pilot Study Of the three frogs euthan ized three days post-inocul ation, only one frog liver was positive for the presence of iridovirus on PCR (Table 1). Light microscopic observation of a portion of liver of this frog demonstrated mild, multifocal sinusoidal karyorrhectic debris. The other two frogs were PCR nega tive and showed no histologic changes (Fig. 5-3). At seven days post-i noculation, one frog died naturally and two others were euthanized. The frog that died naturally had multifocal necrosis (Fig. 5-4) with basophilic cytoplasmic inclusion bodies in the hepatocytes (Fig. 5-5; arrows denote inclusion bodies), which is consistent with ir idovirus infections. The other two frogs had mild to moderate multifocal sinusoidal kary orrhectic debris with fibrin. DNA samples

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112 extracted from the livers of all three frogs were positive on PCR for the presence of iridovirus (Table 1). One frog was pos itive 14 days post-inoculation. This frog histologically showed moderate multifocal sinu soidal karyorrhectic debris with moderate marked extramedullary hematopoeisis. The other two inoculated frogs were negative on PCR and histologically were normal. PCR of liver samples from control frogs were negative and frog livers were histologically normal. Virus was isolated on TH-1 cells only from frogs with positive PCR results, while frogs with negative PCR results demonstrated no cytopathic effects (Table 1). Virus titers in positive frogs were determined between groups but did not differ significantly between days. As a result of this data, frogs in the larger study for feedi ng were euthanized 7 days post-inoculation. Full Frog Study One additional control frog was found to be positive for the presence of iridovirus when the liver samples were tested by PCR fo r a total of three of 71 (4.2%) frogs from the biological supply company being positive on arrival for iridovirus. Twenty-one control frogs were selected for blending fo r turtle food, along with four skeletons for bone matter. Twenty-two of 30 (73.3%) liver samples from experimentally inoculated frogs were positive by PCR for iridovirus at necropsy. Organs from 18 of the samples showing the strongest signal were utilized for food. Weak positives and negatives were excluded to reduce any dilutiona l effect. Four skeletons we re added for bone matter. Turtles Four juvenile turtles were euthanized or died prior to the termination of the study. Turtle 9 and 23 died 1 week post-inoculation (WPI), turtle 11 was euthanized two WPI, and turtle 8 was euthanized 4 WPI. At n ecropsy, it was observed that the pharyngostomy tubes had perforated the esophagus and that all four turtles ha d likely died as a result of

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113 sepsis from injecting the frog homogenates di rectly into the coelomic cavity. Three of the turtles were in the infected group and one was in the control group. PCR on tissues collected at necropsy failed to identify ir idovirus in two of the turtles being fed Ranavirus-infected frogs. One turtle (No. 11) wa s positive, but because food was being administered unintentionally into the coelomic cavity, it is difficult to say whether the virus was causing an active in fection or rather it was simply residual virus from the last feeding. Histopathologic observa tions suggest the latter, as lesions typically observed in other naturally and experimentally in oculated turtles were not observed. All other turtles remained clinically healthy throughout the study. PCR of tissues collected at necropsy were nega tive in each turtle. No histol ogic changes were seen that were consistent with lesions s een in naturally or experimenta lly inoculated turtles. At necropsy, the feeding tubes were in the co rrect locations and no lesions were seen observed with their presence. Virus Titers Gauze filtration of the food being fed to j uveniles actually resulted in higher titers. Food assayed for juveniles from the fi rst day of feeding had a titer of 10 3.75 TCID 50 /ml while adult food had a titer of 10 2.5 TCID 50 /ml. Titers taken at 2 weeks and 4 weeks had titers of 10 3.5 and 10 2.5 TCID 50 /ml, respectively. ELISA Plasma samples collected weekly over th e three-month study failed to detect the production of antiRanavirus antibodies. P/N ratios did not exceed 1.5 in any turtle at all time points in comparison to a positive control observed at 3.5 (Fig. 5-6).

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114 Discussion Iridoviruses of the genus Ranavirus are emerging as important pathogens of captive and free-ranging chelonians in the easte rn United States. When managing disease in populations, one of the more important que stions becomes "how is this virus being transmitted?" If a method of transmission can be identified, breaks in the chain or route of transmission become important methods of controlling and preventing the spread of disease within and between populations. L ittle is known about the transmission of Ranavirus to chelonians, although cases of infec tion are being increasingly reported (Allender et al., In Press; DeVoe et al., 2004; Johnson et al., 2004). Recent findings suggest that amphibians may serve as a reser voir host of infection for chelonians, as a frog isolate collected at the same location as a tortoise isolate shared identical restriction patterns when whole viral genomes were rest ricted with two different endonucleases (Johnson et al., unpublished data). This study was designed to explore the hypothesis that turtles might become inf ected with iridovirus by ingesting dead infected amphibians. Experimental studies with amphibians have shown that amphibians can become infected by cannibalism of in fected individuals or i ngested water (Jancovich et al ., 2001, Pearman et al, 2004). This indicates that by e ither source, infection by an oral route is possible under experimental cond itions in amphibians. We we re unable to replicate these findings in our study with red-eared sliders, either because we failed to replicate natural conditions of oral inoculation, or that chel onians are infected by a different route than amphibians. By placing feeding tubes, we we re able to control the amount of food that the turtles ingested and the fre quency with which they ate; however, this also limited the amount of food that was ingested to values established for feeding debilitated anorexic turtles in rehabilitation settings. Also, the si ze of the particulate matter that was ingested

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115 was limited in size. Wild turtles may ingest higher amounts of food than 1% of their body weight used in this study and thus inge st greater titers of virus as infected amphibians are eaten. Or, viral loads in naturally exposed amphibians may achieve higher levels than those that we induced experimentally in the inoculated leopard frogs. By placing the feeding tubes within the esophagus, we created a fixed diameter for food to pass through that is significantly smaller th an the flexible diameter of the esophagus in chelonians. So when turtles are ingesting a dead amphibian, much larger pieces can be ingested, along with bones and ot her organic matter that might be adhered to the carcass. These bones or other hard matter might cause natural abrasions that we did not replicate in our study, although placement of a feeding tube should have created a breakdown (even if temporary) in the gastrointestinal -vascular barrier. Additionally, we bypassed the oral cavity by placing the feeding tube in the caudal esophagus, which may or may not play an important role in virus-receptor interactions. Additional factors may have caused us to fail to recognize a true route of oral transmission. Studies with experimental inoculation of iri doviruses in amphibians have shown that both host and dose characteristics are important in determining the mortality rates, as well as the duration of disease (Brunner et al ., 2005). The infection dose was positively correlated with the mortality rate and inversely related to average survival times. Environmental temperatures have al so been shown to significantly impact the percent mortality and time to death in sala manders experimentally inoculated with a Ranavirus (Rojas et al ., 2005), where salamanders inoculat ed at 18 and 10C were more likely to die than those exposed at 26C. While water and room temperatures in this study averaged between approximately 21 and 25C, basking areas were kept warmer at

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116 28C. Eliminating heat lamps over basking areas and lowering the room temperature might have kept turtles cool er, and altered the results. Another possibility that we could not induce infecti on in this study is that chelonians are not infected oral ly and other routes of transm ission should be investigated. Iridoviruses are variably host specific, and we have shown that a tortoise isolate was capable of creating disease in leopard frogs. Also, we have shown in previous studies that Ranavirus infections of chelonians create a systemic disease and viral DNA sequence can be amplified from multiple tissues (Johnson et al., unpublished data). Intracytoplasmic inclusion bodies have also be en observed in circulat ing leukocytes in an infected box turtle in Tennessee (Allender et al., In Press). Thus, it is possible that arthropod-borne vectors, such as mosquitoes or other biting insects, might play a role in transmitting the virus from amphibians to chelonians or from chelonians to other chelonians. An additional finding of interest in this st udy was the ability of this virus isolated from a tortoise to create dis ease in an adult leopard frog. Other studies have shown that adult amphibians are often resistant to infection with Frog Virus 3 (Gantress et al., 2003). One frog experimentally inoculated with a tort oise isolate died naturally seven days postinoculation with histologic le sions consistent with iridovir us including hepatic necrosis and the presence of intracytoplasmic inclus ion bodies. This helps to confirm the possibility that a tortoise virus could be pat hogenic to a species of a different class such as an amphibian, and therefore, it is possible that an amphibian virus could be pathogenic to a tortoise. However we cannot rule out that this frog was infected upon arrival. While PCR of oral and cloacal swabs did not amp lify viral DNA sequences, the findings of the

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117 full study showed that only 2 of 3 infected frogs had positive PCR on swabs collected ante-mortem. Because we inoculated this fr og with virus, we cannot rule out that we super-infected this individual which may ha ve overloaded the immune system resulting in the death of the animal. Along the sa me lines, this study also provides a rough prevalence of infected amphibians being sold at biological supply companies. About 4% of the frogs ordered from the company were already infected upon arrival, although they appeared clinically healthy dur ing the two weeks they were h oused in the facility. This could have significant effects in terms of dispersal of infectio n if schools or other organizations bought them for a study and then released them into local environments. Additionally, this study has shown that PCR of oral and cloacal swabs can be a useful method for detecting active iri dovirus infections of amphibians. While we missed one of the three infected frogs, we were able to amp lify sequences in 2 frogs ante-mortem. This could provide useful for detecting the presence of dis ease in populations with low numbers of individuals where sacrific ing adult or young amphibians would be detrimental to the overall success of the population. Prevalence rates of infection however, might be underestimated. In summary, our study attempted to demons trate an oral route of transmission of a Ranavirus from infected amphibians to chelon ians. We tube-fed infected frog homogenates to turtles over six weeks and monitored them for signs of clinical disease and for production of anti-Ranavirus antibodie s which might suggest an exposure. We were unable to confirm that chelonians can become infected by ingesting dead amphibians. However, further studies a llowing turtles to eat infected amphibians ad lib might provide different results as well as altering th e conditions of the inoculation such as

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118 environmental temperatures and viral load. Additional methods of transmission should be investigated such as a vector-borne route of transmission. Determining the route of transmission of Ranavirus to chelonians will allow for better management and prevention of disease in captive a nd wild populations.

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119 Table 5-1. Results of the pilot frog study. Da y = days post inoculation that frogs were euthanized. PCR = result of the polym erase chain reaction. Histologic change is recorded as positive if lesions were observed consistent with iridovirus infections. Frog Infected Control Day PCR VI Titer (TCID 50 ) Histologic Change Inclusion Bodies 1 X 3 + + 10 4 + 2 X 3 N/A 3 X 3 N/A 4 X 7 + + 10 4.5 + + 5 X 7 + + 10 6.75 + 6 X 7 + + 10 4 + 7 X 14 + + 10 5.75 + 8 X 14 N/A 9 X 14 N/A 10 X 14 N/A 11 X 14 N/A 12 X 14 N/A 13 X 14 N/A Fig. 5-1. Photograph demonstrating injection of virus infected cell culture media into a ventral lymph sac in a leopard frog ( Rana pipiens).

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120 Fig. 5-2. Photograph demonstrating the pl acement of a feeding tube in a red-eared slider ( Trachemys scripta elegans ) for administering frog homogenates directly into the caudal esophagus.

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121 Fig. 5-3. Photomicrograph demonstrating the normal architecture of a liver in a leopard frog ( Rana berlandieri ).

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122 Fig. 5-4. Photomicrograph demonstrating multifocal hepatic necrosis in a leopard frog ( Rana berlandieri ) experimentally inoculated with a Ranavirus isolated from a Burmese star tortoise ( Geochelone platynota ).

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123 Fig. 5-5. Photomicrograph at high er magnification of the leopard frog ( Rana berlandieri ) liver shown in Fig. 5-4. A rrows denote the presence of intracytoplasmic inclusion bodies consiste nt with iridovirus infections in amphibians.

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124 Turtle Feeding Trial0 0.5 1 1.5 2 2.5 3 3.5 4 12345678910 Weeks post-inoculation 1 2 3 4 5 6 7 10 12 14 15 16 Positive Fig. 5-6. ELISA results graphed as positive to negative (P/N) ratios. Samples were assayed weekly for the duration of the study. A positive control value is included for comparison of values. All turtles remained negative throughout the study.

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CHAPTER 6 IN VITRO EFFICACY OF ACYCLOVI R AS A POTENTIAL THERAPEUTIC AGENT FOR IRIDOVIRUS INFECTIONS IN CHELONIANS Introduction Iridoviruses have been shown to be pat hogenic in chelonians both in the wild and captivity (Marschang et al. 1996; Westhouse et al. 1996; DeVoe et al. 2004). The method of transmission remains unknown, but c ould be a result of ingesting infected amphibians or exposure to infected water sources. Both have been shown to be a route of transmission in experimental inocul ations of salamanders (Jancovich et al. 2001; Pearman et al. 2004). Other routes of transmission such as vector-borne transmission could potentially play a role such as tr ansmission through mosquitoes or other biting insects, although this has not been previous ly investigated. Re gardless of route of infection, chelonians housed in out door environments at breeding facilities, in private and zoological collections may be at risk for exposure and develo pment of effective treatment methods would be valuable for captive manage ment of these infections. Previously, treatment of infected chelonians has consiste d of supportive care including intracoelomic or subcutaneous fluids, and antibiotics to reduce opportuni stic infections. Antiviral therapy has been used to treat herpesvirus infections in tortoises (Marschang et al. 1997), and while pharmacokinetics studies have not yet been performed, the recommended dose for treatment with acyclovir is 80mg/kg ora lly once daily (Funk and Diethelm, 2006). Marschang et al. (1997) evaluated the in vitro efficacy of two doses of acyclovir and 125

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126 gancyclovir (25g/ml and 50g/ml) at redu cing replication of herpesviruses in cell culture and found them to be very effective. Acyclovir is a guanine anal ogue antiviral drug. It is closely related to DNA except that it lacks the 2'and 3' carbons and 3' -hydroxyl group of the deoxyribose ring (Elion, 1993). Acyclovir needs to be phosphorylated three times to acycl ovir triphosphate in order to be an active at blocking virus repl ication. The virus thymidine kinase (TK) enzyme in some herpesviruses can perform the initial phosphorylation of the acyclovir, unlike the cellular TK, which cannot. This prevents acyclovir from stopping replication of cellular DNA, but allows for inhibition of viral DNA replication. A cellular enzyme, guanylate kinase, performs a second phosphoryl ation (Miller and Mi ller, 1980). A third phosphorylation then occurs which can be done by a number of cellular enzymes (Elion, 1993). Once phosphorylated three times, the antiviral competes with deoxyguanosine triphosphate (dGTP) for the viral DNA polymeras e. Once acyclovir is inserted into the new strand of the replicating DNA chain, replic ation stops because of the lack of a 3'hydroxyl group needed to form the phosphodiester bond to the next deoxynucleoside triphosphate (dNTP). While some species of herpesviruses have TK's that are capable of phosphorylating and thus activating acyclovir, other herpesvi ruses, such as human cytomegalovirus, are much less sensitive to acyclovir (Elion, 1993). Other large DNA viruses also have TK genes such as poxviruses (Hruby et al. 1983) and African swine fever virus (Blasco et al. 1990). Several isolates of iridoviruses ha ve been shown to have putative thymidine kinase genes including Chilo iridescent virus (Jakob et al. 2001), and grouper iridovirus (Tsai et al. 2005). Functional thymidine kinases have been found in fish lymphocystis

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127 disease virus (Scholz et al. 1988) and Bohle ir idovirus (Coupar et al. 2005). Coupar et al. (2005) sequenced and compared homologous open reading frames from epizootic haematopoietic necrosis virus (EHNV) from fish, Wamena iridovirus (WIV) from green pythons and frog virus 3 (FV3). High levels of homology at the nucleotide level were shared among isolates suggesting all to be ORF's encoding thymidine kinase. When compared to other DNA virus TK genes a nd cellular TK genes, iridoviruses and herpesviruses appeared more closely related to the mitochondrial TKs and to cellular deoxycytidine kinases, whereas poxviruses were more closely related to the cellular TKs (Coupar et al. 2005). This suggests that if iridovirus TK genes are similar to herpesvirus TK genes, then it is plausible that they too could phosphorylate a nd activate acyclovir. This study was performed to determine if acyclovir is effective at eliminating cytopathic effect in Ranavirus infected cell monolayers, and/or to determine if virus titers were reduced in cell cultures treated with varyin g concentrations of acyclovir. Results of this study will determine if the antiviral co mpound, acyclovir, should be considered for use in treating infected chelonians. Materials and Methods Cell Cultures Terrapene heart cells (TH-1) were acquired from the American Type Culture Collection (ATCC-CCL 50; Rockville, MD ) and grown to confluency in 2cm 2 wells of 24 well cell culture plates (BD Biosciences, San Jose, CA). Cells were cultured in Dulbecco's modified Eagle medium (DMEM, Gibco, Carlsbad, CA) supplemented with 5% fetal bovine serum (Gibco), gentamicin (60m g/liter; Sigma, St. Louis, MO), penicillin G (120,000 U/liter), streptomycin (120,000 U/lit er) and amphotericin B (300g/liter; Sigma). Cells were grown at 28C in the presence of 5% CO 2

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128 Virus A Ranavirus previously isolated from a Burmese star tortoise (BSTRV; Geochelone platynota ) was used to determine the efficacy of acyclovir in vitro Briefly, transmission electron microscopy of BSTRV i noculated TH-1 cells showed arrays of virus-like particles in the cytoplasm of infect ed cells consistent in size and shape with iridoviruses. Polymerase chain reaction ta rgeting a portion of the major capsid protein gene of ranaviruses followed by DNA sequenc ing demonstrated that the BSTRV isolate shared 100% sequence identity with Frog Virus 3, the type species for the genus Ranavirus in the family Iridoviridae. Restriction enzyme digests of BSTRV compared with FV3 showed identical restri ction patterns using two enzymes, HindIII and XbaI, indicating that BSTRV is either identical or closely related to FV3. Intramuscular inoculation of BSTRV into western ornate box turtles ( Terrapene ornata ornata ) and redeared sliders ( Trachemys scripta elegans ) resulted in high mortality rates. The BSTRV isolate was titered using a tissue culture infectious dose 50 (TCID 50 ) assay from infected media of a third passage isolate. One hundred microliters of the stock virus was diluted in 9.9 mls of cell culture media making a 1:100 dilution and ten fold serial dilutions were made from this dilution six times. One milliliter of each virus dilution was added to 4 wells of a 24 well plate. Cells were incubated for 5 days at 28C and then observed for cytopathic effects in cluding cell rounding and detachment from the plate. The TCID 50 value is the dilution at which 50% of the wells are infected with virus. Virus utilized in this study was determined to be 10 5 TCID 50 Acyclovir and Concentrations Acyclovir was purchased from American Pharmaceutical Partners, Inc. (Schaumburg, IL). Acyclovir was diluted into 25mls of the cell cu lture media described

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129 above at the following concentrations: 0, 0.2, 1, 5, 10, and 25g/ml. The pH was measured on an aliquot of each to ensure that the pH of the media containing acyclovir did not differ from the pH of the media. C oncentrations in media were determined from concentrations found to be eff ective against herpesviruses, another large d ouble stranded DNA virus, in vitro (Kimura et al. 1983; Buck and Loh, 1985; Marschang et al. 1997). Cytotoxicity Assays Cytotoxicity can be mistaken for cytopathic effects, so it is important to run a control for this alongside the CPE reduction assays. Four we lls of a 24 well culture plate with confluent TH-1 monolayers were incuba ted with 1ml of media containing acyclovir at one of the following six doses: 0, 0.2, 1, 5, 10 or 25g/ml. Cell cultures were incubated at 28C for four days. After f our days, cell cultures were observed for the presence of cytotoxicity. Cytopathic Effects (CPE) Reduction Assays Acyclovir was evaluated for its ability to reduce replication of virus by observing for CPE reduction in treated cells. TH-1 cells were grown to confluency in 24 well cell culture plates. Two sets of c ontrols were used. The firs t control consisted of cells receiving media without virus or acyclovir to make sure th e media was not creating any CPE. The second set of controls received virus but no acyclovir to ensure the presence and titer of virus in the media. The latter set of controls and the remainder of the cells received 1ml of virus diluted 1:10 in cell culture media and were incubated at 28C for two hours. After two hours, the media wa s removed, cells were washed once in 1x Hanks balanced salt solution (Gibco) and me dia containing acyclovir at 0, 0.2, 1, 5, 10 or 25g/ml was added. Each concentration was don e in triplicate. Cells were allowed to incubate for four days, after which, wells were observed for the presence of CPE.

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130 Virus Titer Reduction Assays The efficacy of acyclovir against iridovirus was also evaluated by virus titer reduction assays. After cell cu ltures were incubated for four days in the CPE reduction assays, the cell cultures were frozen and thawed three times and the media in the triplicate wells were placed into 15ml centrif uge tubes and centrifuged briefly to remove cell debris. Supernatant from each concentr ation was then assayed as above for the average TCID 50 resulting from each concentration including the control, which contained virus without any acyclovir. Th is indicates the quantitative reduction of virus replication resulting from the presence of acyclovir. Results After four days incubation with the various doses of acyclovir, all wells showed CPE at all doses (Table 1). Cytotoxicity or cytopathic effects resulting from the presence of the antiviral compound as opposed to those induced by virus were not observed in any of the wells (Table 1). The virus titer reduc tion assay showed a slight decrease in titer with increasing dose of acyclov ir. Cultures with no or 0.2 g/ml of acyclovir in the cell culture media had a TCID 50 of 10 4.8 /ml. As the dose increased to 1 or 5 g/ml, the titer decreased slightly to 10 4.5 TCID 50 /ml. A further decrease was seen with 10 and 25 g/ml to 10 4.2 and 10 3.8 TCID 50 respectively (Table 1, Fig. 6-1). While a trend was seen towards a decreasing titer with an increase in concentration of acyclovir, the decrease was not found to be statistically significant (p=0.287). Discussion This preliminary investigation into a pot ential treatment for chelonian iridovirus infections demonstrated that acyclovir was not capable of completely inhibiting viral

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131 replication, although it showed a non-statistically significant trend towards decreasing virus titers with increasing concentrations of acyclovir. While th is study evaluated the in vitro efficacy of acyclovir up to 25 g/ml, higher doses were not evaluated. One study evaluating the efficacy of acyclovir agai nst feline herpesvirus-1 found that 56 g/ml of acyclovir was necessary to reduce plaque numbers to 50% of untreated infections (van der Meulen et al. 2006). Therefore, evaluation of the efficacy of higher doses of acyclovir against iridovi rus replication is warranted, but should be evaluated for their harmful potential both in vitro and in vivo in the species of interest prior to being recommended as a method of treatment. U npublished reports exist of acyclovir being used successfully at the current recommende d dose of 80 mg/kg/day orally (Funk and Diethelm, 2006) to treat iridov irus infected chelonians (T. Norton, R. Ashton, personal communications), in additi on to supportive care includi ng fluid therapy and broadspectrum antibiotics. In both of these instances, th e tortoises survived the infection, two have gone on to successfully reproduce, and no apparent harmful side effects were noted. However, it remains unknown if the recovery was in fact due to the acyclovir or to the supportive care. Pharmacokinetic studies are still need ed to determine the bioavailability, elimination half-life, therap eutic dose and dosing intervals for chelonians with different routes of administration. An oral route of administration is usually more desirable than intravenous or intramuscular administration as it is less invasive, and is the currently recommended route of administration for acyclovir use in chelonians (Funk and Diethelm, 2006). Valacyclovir, however, is a pr o-drug of acyclovir th at has better oral bioavailability, and thus, requi res fewer and lower doses to achieve superior plasma

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132 acyclovir levels in humans (MacDougall and Guglielmo, 2004). Valacyclovir doses for human use are about one third of the recomm ended dose of acyclovir. Pharmacokinetic studies of both drugs would be useful for future administration of acyclovir for both iridovirus and herp esvirus infections. Another factor that might have affected th e results of this study is the timing of acyclovir administration. To mimic a realistic infection, virus was introduced prior to adding media containing acyclovir Results may have varied if cells were pretreated with acyclovir, if acyclovir was introduced at the time of virus administra tion or earlier than 2 hours post adsorption of virus to cell monolayers. A study looking at the efficacy of acyclovir in reducing the replicati on of channel catf ish herpesvirus in vitro showed that the earlier the antiviral drug is added, the more virus replic ation is suppressed (Buck and Loh, 1985). While acyclovir added at 0 and 1.5 hours after adsorption of virus to cell cultures resulted in 8199% inhibition of herpesvirus re plication, no inhibition was seen when acyclovir was added 5 hours after ad sorption. Thus, further studies could determine if addition of acycl ovir earlier than 2 hours post adsorption might result in a further increase in inhibiti on of iridovirus replication. While acyclovir may or may not be helpful in reducing in vivo virus replication of iridovirus, other drugs should similarly be eval uated for their ability to inhibit iridovirus replication. Acylcovir was chosen for this st udy based on the similarity of the iridovirus TK genes to herpesvirus TK genes, and the su ccessful use of acyclov ir to treat human and other herpesvirus infections. Other nucle oside analogue antiviral drugs should be evaluated such as gancyc lovir and vidarabine.

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133 This study has shown that there is a non-sta tistically significant trend in decreasing virus titers with increasi ng doses of acyclovir in Ranavirus infected cell culture monolayers. This represents the first investigation into an antiviral drug as a potential treatment for iridovirus in fected chelonians.

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134 Table 6-1. Effect of increasing doses of acyclovir on cytopathic effect reduction, cytotoxicity and TCID 50 of Terrapene heart cells inoculated with BSTRV. Acyclovir Concentration ( g/ml) No. of wells showing CPE (n=3) No. of wells showing cytotoxicity (n=4) Virus titer (TCID 50 ) 0 3 0 10 4.8 0.2 3 0 10 4.8 1 3 0 10 4.5 5 3 0 10 4.5 10 3 0 10 4.2 25 3 0 10 3.8 4.8 4.8 4.5 4.5 4.2 3.8 0 1 2 3 4 5 6 0 0.2 1 5 10 25 ug/ml AcyclovirTCID50 Fig. 6-1. Effect of increasing concen trations of acyclovir on the TCID 50 of Terrapene cells inoculated with the Burm ese star tortoise isolate.

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CHAPTER 7 CONCLUSIONS AND FUTURE RESEARCH It has been suggested that ch elonians face a more serious th reat of decline than that posed by the well-publicized decline of amphibian populations (Klemens, 2000). Two thirds of all species of freshwater turtles and tortoises are currently listed as threatened on the IUCN Red List of Threatened Species (Turtle Conservation Fund, 2002). Chelonians have low fecundity, low juvenile survival ra te, and a long adult lifespan; a life history strategy where loss of adult anim als (such as loss by disease) has a significant impact on population recovery (Heppell, 1998). Emerging infecti ous diseases have been increasingly recognized as f actors influencing wildlife he alth and populations (Harvell et al ., 1999; Daszak et al ., 2000) and the cause(s) of mass mortality events in wild chelonian populations often remain unde termined (Flanagan, 2000; Dodd, 2001). This study fulfills Koch's postulates, conclusively identifying Ranavirus as a causative agent of mortality in chelonians. It provides a comprehensive review of histologic changes observed in both naturally and experimentally infected chelonians, which will prove useful to pathologists, wildlife veterinarians, and biologists in the future. Review of archived materials suggest s that chelonian iridovirus infections date back to at least 1991 and recent and historic ca ses have defined a much larger geographic range of prevalence than previously known, spanning from Texas to New York. The seroprevalence rate of exposur e in free-ranging gopher tortoises was determined at 1.5%, although we suspect that this is an underestimate of the true exposure rate in natural settings : an artifact of th e severity and short duration of disease 135

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136 as determined by experimental transmission studies. The seroprev alence of free-ranging box turtles was not evaluated. Box turtles are more carnivorous than gopher tortoises, so if ingestion of amphibians is a major source of infection, we might expect to see a higher rate of prevalence in box turt les than in gopher tortoises. Similarly, fresh water turtles should be evaluated for their rates of exposure, because they are like ly in more direct contact with virus particles in infected bodies of water. Further evaluation of species susceptibility should also be performed. Wh ile three of five Burmese star tortoises became ill, of which one died, several other spec ies of tortoises were in adjacent pens and remained clinically healthy, suggesting that some susceptibility differences exist between species. Persistent infections should also be eval uated further. This is known to occur in amphibians, and result of one transmissi on study showed that one turtle remaining clinically healthy was shedding virus 30 days post-inoculation. Studies with a larger sample size and longer duration would help to confirm this finding. Also determining whether or not the virus being shed was inf ectious would be helpful in determining how to handle recovering individuals within managed populations. Another study that should be focused on in the future is confirmation that chelonians can become infected with amphibian viruses. While we were able to show that a tortoise isolate was capable of infecting other turtles, and that this virus appeared to be closely related to, if not the same strain as the frog isolate, we could not be 100% sure that this virus came from an amphibian. We did however show that the tortoise isolate was able to create disease in at least one frog, with hist ologic confirmation of lesions

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137 consistent with iridovirus infection. A furt her study taking a frog isolate and inoculating turtles would help to confir m that amphibians could serve as a source of infection. The most important question this study faile d to answer was how this virus is being transmitted. We hypothesized that amphibians ma y be a source of infection, as isolates found in amphibians at the site of two turtle deaths had 100% shared sequence identity across a portion of the major capsid protein ge ne with the turtle is olates. Restriction enzyme analysis of one tortoise and frog isol ate showed identical pa tterns suggesting that they are infected with the same strain or cl osely related strains of virus. If amphibians really are the source of infection, than chel onians worldwide are at risk of exposure in areas where they overlap with amphibian populat ions. Chelonians at the site where the frog isolate was obtained were observed on tw o occasions ingesting dead amphibians, and thus we suspected that tr ansmission might occur by way of an oral route. However, we were unable to replicat e an oral route of transm ission, both using a single concentrated dose of virus in cell culture me dia as well as by repeated dosing of turtles with infected frogs. Further studies should focus on identifying the route of transmission, as this is a critical piece of information in managing diseas ed populations or in preventing disease. Vector-borne routes of infection s hould really be considered because this virus causes systemic infections. In summary, this study has shown that iridoviruses in the genus Ranavirus can be highly pathogenic to turtles and tortoises. Th ey may be a significant cause of mortality in wild populations, and die-offs w ould likely be difficult to de tect based on the duration of disease. While we were able to provide some answers regarding the pathogenesis of this

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138 virus in chelonians, there are many questions that remain unanswered and to which future studies should be directed.

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LIST OF REFERENCES Allender, M.C., M.M. Fry, A.R. Irizarry, L. Craig, A.J. Johnson, and M. Jones. In press. Intracytoplasmic inclusions in circulating leukocytes from an eastern box turtle ( Terrapene carolina carolina ) with iridoviral infect ion. J. Wildl. Dis. Altschul, S.F., T.L. Madden, A.A. Schff er, J. Zhang, Z. Zhang, W. Miller, and D.J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucl. Acid Res. 25:3389-3402. Ariel, E. 1997. Pathology and serological aspect s of Bohle iridovirus infections in six selected water-associated reptiles in No rth Queensland. PhD thesis. James Cook University, Queensland. Blasco, R., C. Lopez-Otin, M. Munoz, E.O. Bockamp, C. SimonMateio, E. Vinuela. 1990. Sequence and evolutionary relations hips of African swine fever virus thymidine kinase. Virol. 178:301-304. Bollinger, T.K., J. Mao, D. Schock, R.M. Brigham, and V.G. Chinchar. 1999. Pathology, isolation and preliminary molecular characterization of a novel iridovirus from tiger salamanders in Saskatchewan. J. Wildl. Dis. 35:413-429. Bradford, D.F. 1991. Mass mortality and extincition in a high-elevation population of Rana muscosa. J. Herpetol. 25:174-177. Brown, M.B., G.S. McLaughlin, P.A. Klein, B.C. Crenshaw, I.M. Schumacher, D.R. Brown, and E.R. Jacobson. 1999. Upper respiratory tr act disease in the gopher tortoise is caused by Mycoplasma agassizii J. Clin. Microbiol. 37:2262-2269. Brown, D.R., I.M. Schumacher, G.S. McL aughlin, M.B. Brown, P.A. Klein, and E.R. Jacobson. 2002. Application of diagnostic tests for mycoplasmal infections of desert and gopher tortoises, with management recommendations. Chelon. Conserv. Biol. 4:497-507. Brown, D.R., I.M. Schumacher, M.F. N ogueira, L.J. Richey, L.A. Zacher, T.R. Schoeb, K.A. Vliet, R.A. Bennett, E.R. Jacobson and M.B. Brown. 2001. Detection of antibodies to a pathogeni c mycoplasma in American alligators ( Alligator mississippiensis), broad-nosed Caimans (Caiman latirostris ), and Siamese crocodiles ( Crocodylus siamensis ). J. Clin. Microbiol. 39:285-292. 139

PAGE 153

140 Brown, J.D., and J.M. Sleeman. 2002. Morbidity and mortality of reptiles admitted to the Wildlife Center of Virginia, 1991 to 2000. J. Wildl. Dis. 38:699-705. Brunner, J.L., K. Richards, and J.P. Collins. 2005. Dose and host characteristics influence virulence of ranaviru s infections. Oecologia. 144 :399-406. Brunner, J.L., D.M. Schock, E.W. Davidson, and J.P. Collins. 2004. Intraspecific reservoirs: complex life history and the pers istence of a lethal ranavirus. Ecology. 85:560-566. Buck, C.C., and P.C. Loh. 1985. In vitro effect of acyclovir and other antiherpetic compounds on the replicati on of channel catfish vi rus. Antiviral Res. 5:269-280. Burkhardt, E., H. Mller. 1987. Susceptibility of checken blood lymphoblasts and monocytes to infectious bursal disease virus (IBDV). Arch. Virol. 94:297-303. Caipang, C.M.A., T. Takano, I. Hirono and T. Aoki. 2006. Genetic vaccines protect red seabream, Pagrus major upon challenge with red seabream iridovirus (RSIV). Fish Shellfish Immunol. 2:130-138. Chen, Z., J. Zheng, and Y. Jian. 1999. A new iridovirus isolated from soft-shelled turtle. Virus Res. 63:147-151. Chinchar, V.G. 2002. Ranaviruses (family Iridoviridae): emerging cold-blooded killers. Brief review. Arch. Virol. 147 :447-470. Chinchar, V.G., J. Wang, G. Murti, C. Carey, and L. Rollins-Smith. 2001. Inactivation of frog virus 3 and channe l catfish virus by esculentin-2P and ranatuerin-2P, two antimicrobial peptides isolated from frog skin. Virol. 288:351-357. Clark, J.F., G. Gray, F. Fabian, R.F. Ziegel, and D.T. Karzon. 1969. Comparative studies of amphibian cytoplasmic virus strains isolated from the leopard frog, bullfrog, and newt, p. 310-326. In : A. Mizell (ed.), Biology of amphibian tumors. Recent results in cancer rese arch. Springer-Verlag, New York. Cooper, E.L., A.E. Klumpau, and A.G. Zabata 1985. Chapter 8: Reptilian immunity, p. 599-678. In: C. Gans (ed.), Biology of the Reptilia, Vol 14, Morphology. Academic Press, New York. Coupar, B.E.H., S.G. Goldie, A.D. Hyatt and J.A. Pallister. 2005. Identification of a Bohle iridovirus thymidine kinase gene and demonstration of activity using vaccinia virus. Arch. Virol. 150 :1797-1812. Crowther, J.R. 2001. The ELISA guidebook, p. 301-345. Humana Press, Totowa, New Jersey.

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141 Cullen, B.R., and L. Owens. 2002. Experimental challenge and clinical cases of Bohle iridovirus (BIV) in native Australi an anurans. Dis. Aquat. Org. 49:83-92. Cunningham, A.A., T.E.S. Langton, P.M. Bennett, J.F. Lewin, S.E.N. Drury, R.E. Gough, and S.K. Macgregor. 1996. Pathological and microbiological findings from incidents of unusual mortality of the common frog ( Rana temporaria ). Philos. Trans. R. Lond. B Biol. Sci. 351:1529-1557. Daszak, P., L. Berger, A.A. Cunningham, A.D. Hyatt, D.E. Green and R. Speare. 1999. Emerging infectious di seases and amphibian population declines. Emerg. Infect. Dis. 5:735-748. Daszak, P., A.A. Cunningham, and A.D. Hyatt. 2000. Emerging infectious diseases of wildlifethreats to biodiversit y and human health. Science. 287:443-449. DeVoe, R., K. Geissler, K., S. Elmore, D. Rotstein, G. Lewbart and J. Guy. 2004. Ranavirus-associated morbidity and mortality in a group of captive eastern box turtles ( Terrapene carolina carolina ). J. Zoo Wildl. Med. 35:524-543. Docherty, D., C.U. Meteyer, J. Wang, J. Mao, S.T. Case, and V.G. Chinchar. 2003. Diagnostic and molecular evaluation of three iridovirus-associated salamander mortality events. J. Wildl. Dis. 39:556-566. Dodd, K.C. 2001. North American box turtles: a natural history. University of Oklahoma Press. Norman, Oklahoma. Drury, S.E.N., R.E. Gough, and I. Calvert. 2002. Detection and isolation of an iridovirus from chameleons ( Chameleo quadricornis and Chameleo hoehnelli ) in the United Kingdom. Vet. Rec. 150:451-452. Elion, G.B. 1993. Acyclovir: discovery, mechanism of action and selectivity. J. Med. Virol. Suppl. 1:2-6. Evans, A.S. 1976. Causation and disease: the HenleKoch postulates re visited. Yale J. Biol. Med. 49:175-195. Fellers, G.M., and C.A. Drost. 1993. Disappearance of the cascades frog Rana cascadae at the southern end of its range, California, USA. Biol. Conserv. 65:177-181. Fisher, R.N. and H.B. Shaffer. 1996. The decline of amphibians in California's Great Central Valley. Conserv. Biol. 10:1387-1397. Flanagan, J. 2000. Disease and health considerations, p. 85-95. In: M.W. Klemens (ed.), Turtle conservation, Smithsonian Institution Press, Washington.

PAGE 155

142 Forson, D., and A. Storfer. 2006. Effects of atrazine and iridovirus infection on survival and life-history traits of the long-toed salamander ( Ambystoma macrodactylum ). Environ. Toxicol. Chem. 25:168-173. Funk, R.S., and G. Diethelm. 2006. Reptile formulary, p. 1119-1139. In: D.M. Mader (ed.), Reptile Medicine and Surgery, 2nd edition, WB Saunders, Philadelphia. Gantress, J., Maniero, G.D., Cohen, N., and Robert, J. 2003. Development and characterization of a model system to study amphibian immune responses to iridoviruses. Virol. 311 :254-262. Goorha, R. and K.G. Murti. 1982. The genome of frog virus 3, an animal DNA virus, is circularly permuted and terminally redundant. Proc. Natl. Acad. Sci. U.S.A. 79:248-252. Goorha, R., G. Murti, A. Granoff, and R. Tirey. 1978. Macromolecular synthesis in cells infected by frog virus 3. VIII. The nuc leus is a site of frog virus 3 DNA and RNA synthesis. Virol. 82:34-52. Granoff, A., P.E. Came, and K.A. Rafferty, Jr. 1965. The isolation and properties of viruses from Rana pipiens: Their possible relationship to the renal adenocarcinoma of the leopard frog. Ann. N.Y. Acad. Sci. 126:237-255. Granoff A., P.E. Came, and D.C. Breeze. 1966. Viruses and renal adenocarcinoma of Rana pipiens. I. The isolation and properties of virus from normal and tumor tissue. Virol. 29:133-148. Green, D.E., K.A. Converse, and A.K. Schrader. 2002. Epizootiology of sixty-four amphibian morbidity and mortality events in the USA, 1996-2001. Ann. N.Y. Acad. Sci. 969:323-339. Harlow, E., and D. Lane. 1988. Antibodies: a laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Harvell, C.D., K. Kim, J.M. Burkholder, R.R. Colwell, P.R. Epstein, D.J. Grimes, E.E. Hofmann, E.K. Lipp, A.D. Osterhaus, R.M. Overstreet, J.W. Porter, G.W. Smith, and G.R. Vasta. 1999. Emerging marine diseasesclimate links and anthropogenic factors. Science. 285:1505-1510. Heldstab, A, and G. Bestetti. 1982. Spontaneous viral hepatitis in a spur-tailed Mediterranean land tortoise ( Testudo hermanni ). J. Zoo Anim. Med. 13:113-120. Heppell, S.S. 1998. Application of life-history theo ry and population model analysis to turtle conservation. Copeia. 2:367-375.

PAGE 156

143 Herbst, L. and E.R. Jacobson. 1995. Diseases of marine turtles, p. 593-595. In : K. Bjorndal (ed.), Biology and Conservation of Sea Turtles. Smithsonian Institution Press, Washington, D.C. Hruby, D.E., R.A. Maki, D.B. Miller, and L.A. Ball. 1983. Fine structure analysis and nucleotide sequence of the vaccinia virus thymidine kinase gene. Proc. Natl. Acad. Sci. USA. 80:3411-3415. Hyatt A.D., M. Williamson, B.E. Coupar, D. Middleton, S.G. Hengstberger, A.R. Gould, P. Selleck, T.G. Wise, J. Kattenbelt, A.A. Cunningham, and J. Lee. 2002. First identification of a ranavirus from green pythons ( Chondropython viridis ). J. Wildl. Dis. 38:239-252. Jacobson E.R., J.L. Behler, and J.L. Jarchow. 1999. Health assessment of chelonians and release into the wild, p. 232-242. In: M.E. Fowler and R.E. Miller (eds.), Zoo and Wild Animal Medicine, Cu rrent Therapy 4, W.B. Saunders Co., Philadelphia. Jacobson, E.R., A.J. Johnson, J.A. Hernande z, S.J. Tucker, A.P. DuPuis II, R. Stevens, D. Carbonneau, and L. Stark. 2005. Validation and use of an indirect enzyme-linked immunosorbent assay for detection of antibodies to west Nile virus in American alligators ( Alligator mississippiensis) in Florida. J. Wildl. Dis. 41:107-114. Jakob, N.J., K. Muller, U. Bahr, and G. Darai. 2001. Analysis of the first complete DNA sequence of an invertebrate iridoviru s: coding strategy of the genome of Chilo iridescent virus. Virol. 286:182-196. Jancovich, J.K., E.W. Davidson, J.F. Morado, B.L. Jacobs and J.P. Collins. 1997. Isolation of a lethal virus from the endangered tiger salamander Ambystoma tigrinum stebbinsi Dis. Aquat.Org. 31:161-167. Jancovich, J.K., E.W. Davidson, J.F. Morado, B.L. Jacobs, and J.P. Collins. 2001. Transmission of the Ambystoma tigrinum virus to alternative hosts. Dis. Aquat. Org. 46:159-163. Johnson, A.J., J.F.X. Wellehan, A.P. Pessier, T.M. Norton, W.R. Belzer, J.W. Brooks, R. Wagner, N.L. Stedman, J. Spratt, and E.R. Jacobson. 2004. Iridovirus infections of turtles and tortoi ses. Proceedings of the Wildlife Disease Association. Johnson, A.J., A.P., Pessier, J.F.X. Wellehan, R. Brown, and E.R. Jacobson. 2005. Identification of a novel tort oise herpesvirus from a California desert tortoise ( Gopherus agassizii ). Vet. Microbiol. 111:107-116.

PAGE 157

144 Johnsrude, J.D., R.E. Raskin, A.Y. Hoge, and G.W. Erdos. 1997. Intraerythrocytic inclusions associated with iridovi ral infection in a fer de lance (Bothrops moojeni ) snake. Vet. Pathol. 34:235-238. Just, F., S. Essbauer, W. Ahne, and S. Blahak. 2001. Occurrence of an invertebrate iridescent-like virus ( Iridoviridae ) in reptiles. J. Vet. Med. B Infect. Dis. Vet. Public Health. 48:685-694. Kimura, T., S. Suzuki, and M. Yoshimizu. 1983. In vitro antiviral effect of 9-(2hydroxyethoxymethyl) guanine on the fish herpesvirus, Oncorhynchus masou virus (OMV). Antiviral Res. 3:93-101. Klemens, M.W. 2000. Turtle Conservation. Smithsonian Institution Press, Washington. Kroese, F.G., and N. Van Rooijen. 1982. The architecture of the spleen of the redeared slider, Chrysemys scripta elegans (Reptilia, Testudines). J. Morphol. 173: 279-284. Langdon, J.S. 1989. Experimental transmission and pathogenicity of epizootic haematopoietic necrosis virus (EHNV) in redfin perch, Perca fluviatilis L ., and 11 other teleosts. J. Fish Dis. 12:295-310. Langdon, J.S. and J.D. Humphrey. 1987. Epizootic haemat opoietic necrosis, a new viral disease in redfin perch, Perca fluviatilis L., in Australia. J. Fish Dis. 10:289-297. Laurance, W.F., K.R. McDonald, and R. Speare. 1996. Epidemic disease and the catastrophic decline of Australian ra in forest frogs. Conserv. Biol. 10:406-413. Lips, K.R. 1999. Mass mortality and population declin es of anurans at an upland site in Western Panama. Conserv. Biol. 13:117-125. Lopez, M., J.C. Rojas, R. Vandame, and T. Williams. 2002. Parasitoid-mediated transmission of an iridescent vi rus. J. Invertebr. Pathol. 80:160-170. Lorbacher de Ruiz, H. 1990. Hepatotoxicity of iridoviruses, p. 235-245. In: G. Darai (ed.), Molecular biology of iridoviruses, Kluwer Academic Publishers, Boston. MacDougall, C., and B.J. Guglielmo. 2004. Pharmacokinetics of valaciclovir. J. Antimicrob. Chemo. 53:899-901. Maniero, G.D., H. Morales, J. Gantress, and J. Robert. 2006. Generation of a longlasting, protective, and neutralizing anti body response to the ranavirus FV3 by the frog Xenopus. Develop. Comp. Immunol. 30:649-657.

PAGE 158

145 Mao, J., D.E. Green, G. Fellers, and V.G. Chinchar. 1999. Molecular characterization of iridovir uses isolated from sympat ric amphibians and fish. Virus Res. 63:45-52. Mao, J., R.P. Hedrick, and V.G. Chinchar. 1997. Molecular characterization, sequence analysis and taxonomic position of newly isolated fish iridoviruses. Virol. 229:212-220. Marschang, R.E., P. Becher, H. Posthaus, P. Wild, H-J. Thiel, U. Mller-Doblies, E.F. Kaleta, and L.N. Bacciarini. 1999. Isolation and characterization of an iridovirus from Herm anns tortoises (Testudo hermanni ). Arch. Virol. 144 :19091922. Marschang, R.E., S. Braun, P. Becher. 2005. Isolation of a ranavirus from a gecko ( Uroplatus fimbriatus). J. Zoo Wildl. Med. 36:295-300. Marschang, R.E., M. Gravendyck, and E.F. Kaleta. 1997. Herpesvirus in tortoises: investigations into virus isolation and the treatment of viral stomatitis in Testudo hermanni and T. graeca J. Vet. Med. 44:385-394. McDowell, E.M., and B.F. Trump. 1976. Historical fixative s suitable for diagnostic light and electron microscopy. Arch. Pathol. Lab. Med. 100:405-414. Miller, W.H. and R.L. Miller. 1980. Phosphorylation of acy clovir (acycloguanosine) monophosphate by GMP kinase. J. Biol. Chem. 255:7204-7207. Moody, N.J.G., and L. Owens. 1994. Experimental demonstration of the pathogenicity of a frog virus, Bohle iridovirus, for a fish species barramundi Lates calcarifer Dis. Aquat. Org. 18:95-102. Morse, S.S. 1995. Factors in the emergence of inf ectious diseases. Emerg. Infect. Dis. 1:7-15. Muller, M., N. Zangger, and T. Denzler. 1988. Iridovirus-epidemie bei der griechischen Landschildkrote ( Testudo hermanni hermanni ), p.271-274. Verhandl Ber 30. Int Symp Erkr Zoo-und Wildtiere, Sofia. Nakajima, K., Y. Maeno, J. Kurita and Y. Inui. 1997. Vaccination against red sea bream iridoviral disease in red sea bream. Fish Pathol. 32:205-209. Nakajima, K., Y. Maeno, A. Honda, K. Yokoyama, K. Tooriyama, and S. Manabe. 1999. Effectiveness of a vaccine against red sea bream iridoviral disease in a field trial test. Dis. Aquat. Org. 36:73-75. Origgi, F.C., P.A. Klein, K. Mathes, S. Blahak, R.E. Marschang, S.J. Tucker and E.R. Jacobson. 2001. Enzyme-linked immunosorbent assay for detecting

PAGE 159

146 herpesvirus exposure in Mediterranea n tortoises (spur-thighed tortoise [ Testudo graeca ] and Hermanns tortoise [ Testudo hermanni ]). J. Clin. Microbiol. 39:3156-3163. Orrigi, F.C., C.H. Romero, D.C. Bloom, P.A. Klein, J.M. Gaskin, S.J. Tucker, and E.R. Jacobson. 2004. Experimental transmission of a herpesvirus in Greek tortoises (Testudo graeca ). Vet. Pathol. 41:50-61. Pearman, P.B., T.W. Garner, M. Straub, and U.F. Greber. 2004. Response of the Italian agile frog ( Rana latastei ) to a Ranavirus, frog vi rus 3: a model for viral emergence in naive populations. J. Wildl. Dis. 40:660-669. Reddacliff, L.A., and R.J. Whittington. 1996. Pathology of epizzotic haematopoietic necrosis virus (EHNV) inf ection in rainbow trout ( Oncorhynchus mykiss Walbaum ) and redfin perch ( Perca fluviatilis L ). J. Comp. Pathol. 115:103-115. Rojas, S., K. Richards, J.K. Jancovich, and E.W. Davidson. 2005. Influence of termperature on Ranavirus infection in larval salamanders Ambystoma tigrinum Dis. Aquat. Organ. 63:95-100. Rossell, C. R., I.M. Rossell, M.M. Orroca, and J.W. Petranka. 2002. Epizootic disease and high mortality in a populati on of eastern box turtles. Herp Rev. 33:99101. Scholz, J., A. Rosen-Wolff, M. Touray, P. Schnitzler and G. Darai. 1988. Identification, mapping and cloning of th e thymidine kinase gene of fish lymphocystis disease virus. Virus Res. 9:63-72. Schramm, B. and J.K. Locker. 2005. Cytoplasmic organization of poxvirus DNA replication. Traffic. 6:839-846. Schumacher, I.M., M.B. Brown, E.R. Jacobson, B.R. Collins, and P.A. Klein. 1993. Detection of antibodies to a pathogeni c mycoplasma in desert tortoises (Gopherus agassizii ) with upper respiratory tract disease. J. Clin. Micro. 31:1454-1460. Seigel, R.A., R.B. Smith, and N.A. Seigel. 2003. Swine flu or 1918 pandemic? Upper respiratory tract disease and sudden mortality of gopher tortoises ( Gopherus polyphemus) on a protected habitat in Florida. J. Herpetol. 37 :137-144. Sorby, R., T.N. Wien, G. Husby, A. Espenes, and T. Landsverk. 2005. Filter function and immune complex trapping in splenic ellipsoids. J. Comp. Pathol. 132:313-321. Speare, R. and J.R. Smith. 1992. An iridovirus-like agen t isolated from the ornate burrowing frog Limnodynastes ornatus in northern Australia. Dis. Aquat. Org. 14:51-57.

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147 Steiner, K.A., R.J. Whittington, R.K. Pe tersen, C. Hornitzky,, and H. Garnett. 1991. Purification of epizootic haematopoie tic necrosis virus and its detection using ELISA. J. Virol. Meth. 33:199-209. Telford, S.R. and E.R. Jacobson. 1993. Lizard erythrocytic virus in east African chameleons. J. Wildl. Dis. 29:57-63. Tidona, C.A., P. Schnitzler, R. Kehm, and G. Darai. 1998. Is the major capsid protein of iridoviruses a suita ble target for the study of viral evolution? Virus Genes. 16:59-66. Tokuda, Y., T. Nakanure, K. Satonaka, S. Maeda, K. Doi, S. Baba, and T. Sugiyama. 1990. Fundamental study on the mechanisms of DNA degradation in tissues fixed in formaldehyde. J. Clin. Pathol. 43:748-751. Tsai, C-T., J-W. Ting, M-H. Wu, MW. Wu, I-C. Guo, and C-Y. Chang. 2005. Complete genome sequence of the grouper iridovirus and comparison of genomic organization with those of othe r iridoviruses. J. Virol. 79:2010-2023. Turtle Conservation Fund. 2002. A global action plan fo r conservation of tortoises and freshwater turtles. Strategy a nd funding prospectus 2002-2007. M.T.C. Printing, Inc., Leominster, Massachusetts. United States Fish and Wildlife Service. 1994. Desert tortoise (Mojave population) recovery plan. U.S. Fish and W ildlife Service. Portland, Oregon. Van der Meulen, K., B. Garre, S. Croubels, and H. Nauwynck. 2006. In vitro comparison of antiviral drugs against feline herpesvirus 1. BMC Vet Res. 26:13. VanDevanter, D.R. P. Warrener, L. Bennett, E.R. Schultz, S. Coulter, R.L. Garber, and T.M. Rose. 1996. Detection and analysis of diverse herpesviral species by consensus primer PCR. J. Clin. Microbiol. 34:1666-1671. Warr, G.W., Magor, K.E., and D.A. Higgins. 1995. IgY: clues to the origins of modern antibodies. Immunol. Today. 16:392-398. Westhouse, R.A., E.R. Jacobson, R.K. Harris, K.R. Winter, and B.L. Homer. 1996. Respiratory and pharyngo-es ophageal iridovirus infection in a gopher tortoise ( Gopherus polyphemus ). J. Wildl. Dis. 32:682-686. Whittington, R.J., C. Kearns, and R. Speare. 1997. Detection of antibodies against iridoviruses in the serum of the amphibian Bufo marinus J. Virol Meth. 68 :105108.

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148 Williams, R.C., and K.M. Smith. 1957. A cristallizable in sect virus. Nature. 179 :119120. Williams, T. 1996. The iridoviruses. Adv. Virus Res. 46:345-412. Williams, T., V. Barbosa-Solomieu, and V.G. Chinchar. 2005. A decade of advances in iridovirus research. Adv. Virus Res. 65:173-248. Wolf, K., G.L. Bullock, C.E. Dunbar, and M.C. Quimby. 1968. Tadpole edema virus: a viscerotrophic pa thogen for anuran amphibian s. J. Infect. Dis. 118:253262. Xeros, N. 1964. A second virus disease of the leather jacket, Tipula paludosa Nature. 174:562-563. Zupanovic, Z., G. Lopez, A.D. Hyatt, B. Green, G. Bartran, H. Parkes, R.J. Whittington, and R. Speare. 1998. Giant toads Bufo marinus in Australia and Venezuela have antibodies against ra naviruses. Dis. Aquat. Organ. 32:1-8.

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BIOGRAPHICAL SKETCH April Johnson received a Bachelor of Science in biology from Cedarville University in July 1998. She then attende d the University of Illinois, College of Veterinary Medicine, where she received a Doctor of Veteri nary Medicine in May 2002. She enrolled in the PhD program at the Univ ersity of Florida, College of Veterinary Medicine, during the fall of that same year. She concurrently enrolled in the Master of Public Health program with an emphasis in epidemiology at the University of Florida, College of Public Health and Health Professions, in the fall of 2003, which she completed in the spring of 2006. In July of 2006, she started a two-year fellowship as an Epidemic Intelligence Service officer with the Commi ssioned Corps of the U.S. Public Health Service at the Centers for Disease Control a nd Prevention in Atlanta, Georgia, where she will focus on the epidemiology and prevention of influenza. 149


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Table of Contents
    Title Page
        Page i
        Page ii
    Acknowledgement
        Page iii
        Page iv
    Table of Contents
        Page v
        Page vi
        Page vii
    List of Tables
        Page viii
    List of Figures
        Page ix
        Page x
        Page xi
    Abstract
        Page xii
        Page xiii
    Introduction
        Page 1
        Page 2
        Page 3
        Page 4
        Page 5
        Page 6
        Page 7
        Page 8
        Page 9
        Page 10
    Ranavirus infection of free-ranging and captive box turtles and tortoises in the United States
        Page 11
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    Development and use of an indirect enzyme linked immunosorbent assay for detection of iridovirus exposure in gopher tortoises
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    Experimental transmission of a ranavirus in western ornate box turtles and red-eared sliders
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    The role of infected leopard frogs in transmission of a ranavirus in red-eared sliders
        Page 102
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    In vitro efficacy of acyclovir as a potential therapeutic agent for iridovirus infections in chelonians
        Page 125
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    Conclusions and future research
        Page 135
        Page 136
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    References
        Page 139
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    Biographical sketch
        Page 149
Full Text












IRIDOVIRUS INFECTIONS OF CAPTIVE AND FREE-RANGING CHELONIANS IN
THE UNITED STATES















By

APRIL JOY JOHNSON


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


2006

































Copyright 2006

by

April Joy Johnson















ACKNOWLEDGMENTS

Funding for this work was provided in part by the University of Florida College of

Veterinary Medicine Batchelor Foundation, grant No. D04ZO-11 from the Morris Animal

Foundation, and a grant from the Disney Conservation Fund. I also thank the University

of Florida, College of Veterinary Medicine for awarding me an Alumni Fellowship for

tuition and stipend over the first three years.

I would like to thank the many people who helped collect and submit turtle and

tortoise samples including Dr. William Belzer, Dr. Terry Norton, Jeffrey Spratt, Valorie

Titus, Susan Seibert and Ben Atkinson. Also, I would like to thank all the pathologists

who contributed to this work including Dr. Allan Pessier, Dr. Nancy Stedman, Dr. Robert

Wagner and Dr. Jason Brooks. I thank Drs. Jerry Stanley and Kathy Goodblood for

allowing me access to chelonian and amphibian populations at Buttermilk Hill Nature

Sanctuary and to Drs. Mick Robinson and Ken Dodd for providing necropsy reports on

archived cases. I would also like to thank the Mycoplasma Research Laboratory at the

University of Florida including Dr. Mary Brown, Dr. Lori Wendland and Dina

Demcovitz for providing access to their wild gopher tortoise plasma bank. I thank

Yvonne Cates at the Zoological Society of San Diego for excellent histology support and

Lynda Schneider from the University of Florida Electron Microscopy Core Laboratory. I

also thank Dr. Harvey Ramirez, Rachelle Wright and Kevin Chadbourne with Animal

Care Services for excellent care of the turtles and frogs.









Several people went above and beyond normal expectations in assisting with this

research in various ways. I would especially like to thank Dr. Allan Pessier for all his

time, patience and assistance with necropsies and histopathology as well as making me a

better scientific writer. I thank Dr. Jim Wellehan for all his advice and assistance with

molecular and laboratory techniques. I also am indebted to our laboratory technicians,

Sylvia Tucker and April Childress. They both provided invaluable laboratory support

and became great friends throughout my time at the University of Florida.

I thank my parents for their support and encouragement, without which I would

never have made it to this point in my career. They have always believed in me and,

although they might not always understand my unique interests, have never discouraged

me from pursuing my goals. For that I will always be thankful.

And last, but far from least, I am extremely grateful to my supervisory committee

members: Dr. Elliott Jacobson, Dr. David Bloom, Dr. Jack Gaskin, Dr. Jorge Hernandez

and Dr. Gail Scherba. I thank them for the advice, support and encouragement they have

given me since the beginning. I thank them for helping me develop my skills and for

their confidence in my abilities. I have learned so much from them and have thoroughly

enjoyed this opportunity to work with them.
















TABLE OF CONTENTS



A C K N O W L E D G M E N T S ................................................................................................. iii

LIST OF TABLES ..................._......... .................... ......... ...... ....... ... viii

LIST OF FIGURES ......... ......................... ...... ........ ............ ix

ABSTRACT .............. ..................... .......... .............. xii

CHAPTER

1 IN T R O D U C T IO N ............................................................................. .............. ...

2 RANAVIRUS INFECTION OF FREE-RANGING AND CAPTIVE BOX
TURTLES AND TORTOISES IN THE UNITED STATES.............. .................11

Introdu action ...................................... ................................. .......... ....... 11
M materials and M methods ....................................................................... .................. 12
A nim als ................................................... .......................... 12
N ecropsy and H istopathology .......................... ............. ............... ... 16
Nucleotide Amplification, Sequencing, and Sequence Analysis .......................16
V iru s Iso latio n ................................................... ................ 17
Transmission Electron Microscopy ........................ ......... ..................18
R restriction Enzym e A analysis ........................................ ........................ 18
R results ............... ...... ......... ........ ... ....................................19
N ecropsy and H istopathology ........................................ ......... ............... 19
PCR and Sequence A analysis ........................................ .......... ............... 20
V iru s Iso latio n ............................................................................................... 2 1
Transmission Electron Microscopy ......................................... ...............21
R estriction Enzym e A analysis ........................................ ........................ 21
D iscu ssio n ...................................................................... .................... 2 2

3 DEVELOPMENT AND USE OF AN INDIRECT ENZYME LINKED
IMMUNOSORBENT ASSAY FOR DETECTION OF IRIDOVIRUS
EXPOSURE IN GOPHER TORTOISES (Gopheruspolyphemus) ..........................34

In tro du ctio n ...................................... ................................................ 3 4
M materials and M methods ....................................................................... ..................36
V iru s ............................................................................. 3 6









Antigen Preparation......................................................... 37
Positive and Negative Reference Plasm a ................................. ................ 38
ELISA Procedure ................................................ .... .. ... .. ........ .... 39
Experim mentally Inoculated Turtles ........................................... .....................40
R eproducibility .................................. .................... ........ ..... ............ 41
Protein Expression and Immunoblotting ................................. ..................42
Wild Gopher Tortoises Samples with Unknown Exposure.............................43
Results ............... ...... ............ ............. ...............43
A ntigen Preparation ......... ............................................................ ... .... ....... 43
ELISA Param eters ......... .. .............. .. .................................... ............. 44
Experim mentally Inoculated Turtles ........................................... .....................44
R eproducibility ................................... .......................... ..... ............ 45
Protein Expression and Immunoblotting ................................. ..................45
Wild Gopher Tortoises Samples with Unknown Exposure.............................46
D discussion ............... ..... .. .......... ......... .. ........ ..... ..... ........ 46

4 EXPERIMENTAL TRANSMISSION OF A RANA VIRUS IN WESTERN
ORNATE BOX TURTLES (Terrapene ornata ornata) AND RED-EARED
SLID ER S (Trachemys script elegans)........................................... .....................65

In tro d u ctio n .......................................... ... ....................................... ..................... 6 5
M materials an d M eth od s .................................................................... .....................66
Experimental Animals and Husbandry....... ................. ................... 66
Pre-inoculation Sample Collection.............................. ...............67
DNA Preparation, Polymerase Chain Reaction and Nucleotide Sequencing......67
E L IS A .........................................................6 8
V irus Preparation........... ....................................... ............ ..70
T ransm mission Studies........... ............................................... .......... ........... 7 1
S tu d y 1 ..............................................................7 1
S tu d y 2 ................................................................7 3
R results ........................................................ .. .. ............ ............... 74
Experimental Animals and Pre-inoculation Sampling .....................................74
T ran sm mission Stu dies..................................................................................... 74
S tu d y 1 ..............................................................7 4
S tu d y 2 .............................................................7 5
D isc u ssio n ............................................................................................................. 8 0

5 THE ROLE OF INFECTED LEOPARD FROGS (Ranapipiens) IN
TRANSMISSION OF A RANAVIRUS IN RED-EARED SLIDERS (Trachemys
scrip ta eleg ans) ......... ...... .................... ......................................................... 102

Introdu action ...................................................................................................102
M materials and M methods ...........................................................104
Virus Preparation ..... ........... ........ ......... .........104
Frog Pilot Study............................................. 105
Full Frog Study and Pre-inoculation Sampling .................................. 106
T u rtle s ......................................................................... 1 0 7









V iru s T ite rs ................................................................................................... 1 0 9
E L IS A ......................................................1 10
R e su lts .................................................................................................................. 1 1 1
F ro g P ilo t S tu d y ............................................................................................ 1 1 1
F ull F rog Study ..................................... ............................................................. 112
T u rtle s ....................................................1 12
Virus Titers .................................. ................................ ........ 113
E L I S A ........................................................................................................... 1 1 3
D isc u ssio n ........................................................................................1 14

6 IN VITRO EFFICACY OF ACYCLOVIR AS A POTENTIAL THERAPEUTIC
AGENT FOR IRIDOVIRUS INFECTIONS IN CHELONIANS ........................125

In tro d u ctio n .......................................................................................................... 12 5
M materials and M methods ...........................................................127
Cell Cultures ............... ......... ........ ......... 127
Virus .............. .......... .......... .................. 128
A cyclovir and Concentrations ..................................................................... 128
Cytotoxicity Assays............................................. 129
Cytopathic Effects (CPE) Reduction Assays ..................................... 129
Virus Titer Reduction Assays .................................................. ........130
R e su lts ................................ ................................................................................. 1 3 0
D isc u ssio n ................................ ........................................................................... 1 3 0

7 CONCLUSIONS AND FUTURE RESEARCH .................... ...........135

L IST O F R E FE R E N C E S ............... ............... ......................................................... 139

BIOGRAPHICAL SKETCH ............... ......... ........ ........149
















LIST OF TABLES


Table page

3-1 R eproducibility of the ELISA .......................................................................... 53

3-2 ELISA results of 1000 free-ranging gopher tortoise (Gopheruspolyphemus)
plasm a sam ples by county and state................................................ .................. 53

3-3 ELISA results of 658 free-ranging gopher tortoises (Gopheruspolyphemus)
from the state of Florida are listed by region. ......................................................55

3-4 ELISA results of 1000 free-ranging gopher tortoises (Gopheruspolyphemus)
listed by state. .......................................................................... 55

4-1 PCR results on tissues collected at necropsy from the pilot study box turtles
(BT; Terrapene ornata ornata) and red-eared sliders (RES; Terrapene script
elegans) in the full transmission studies. ..................................... ............... 88

4-2 Polymerase chain reaction (PCR) results of oral and cloacal swabs taken on
eleven different days post-inoculation (DPI) ................................. ............... 89

4-3 PCR results for urine collected opportunistically from turtles in the full
transm mission study. .................................................................. .. ..... 90

5-1 R results of the pilot frog study. ........................................................... ... ............ 119

6-1 Effect of increasing doses of acyclovir on cytopathic effect reduction,
cytotoxicity and TCID50 of Terrapene heart cells inoculated with BSTRV..........134









LIST OF FIGURES


Figure

2-1 Gross lesions associated with iridovirus infections in turtles and tortoises ............26

2-2 Esophagus, eastern box turtle (Terrapene carolina carolina). There is diffuse
necrosis and ulceration of the mucosa and replacement by fibrin, inflammatory
cell infiltrates and superficial bacterial colonies....... .................... ...............27

2-3 Spleen, eastern box turtle (Terrapene carolina carolina). There is disruption of
the white and red pulp with deposits of fibrin (arrow) admixed with
karyorrhectic debris, and infiltrates of small numbers of heterophils......................28

2-4 Epicardium, Burmese star tortoise (Geochelone platynota). Arrows depict
basophilic intracytoplasmic inclusion bodies in a macrophage and endothelial
c e ll ........................................................ .................................2 9

2-5. Results of a polymerase chain reaction targeting approximately 500 bp of the
m ajor capsid protein gene. .......................................................................... .. .... 30

2-6 Transmission electron photomicrograph of Terrapene heart cells inoculated with
liver tissue from a Burmese star tortoise (Geochelone platynota) demonstrating
cytoplasmic arrays of iridovirus-like particles. ............. ........................................31

2-7 Transmission electron photomicrograph of paraffin embedded spleen from a box
turtle (Terrapene carolina) that died in 1991 in Georgia......................................32

2-8 HindIII and XbaI restriction enzyme pattern of five iridovirus isolates...................33

3-1 Negative staining electron photomicrograph of an iridovirus particle purified by
sucrose gradient ultracentrifugation. ............................... .. ......................... 56

3-2 Optimization of the ELISA with antigen coated at 1:100 dilution, comparing the
positive to negative (P/N) ratio of two fold serial plasma dilutions of the positive
control turtle (Burmese star tortoise with clinical signs of illness) versus a
negative control (Burmese star tortoise with no history of illness).....................57

3-3 Frequency distribution of P/N ratios from an indirect ELISA performed on 1000
free ranging gopher tortoise (Gopheruspolyphemus) plasma samples ..................58

3-4 Individual P/N ratio values for 1000 free-ranging gopher tortoises (Gopherus
polyphem us) in increasing value. ........................................ ......................... 59

3-5 P/N ratios of red-eared slider (Trachemys script elegans) plasma samples
collected w eekly over five m months ........................................ ........ ............... 60









3-6 Coomassie blue staining of a SDS-PAGE gel separating proteins of iridovirus-
infected and uninfected Terrapene heart cell lysates....................... ...............62

3-7 W western im m unoblot. ....................................................................... ...................63

3-8 County map of Florida. The five counties highlighted indicate where
seropositive tortoise samples were identified .............. ........................................64

4-1 Photograph taken 12 days post-inoculation showing development of white
opaque ocular discharge in the IM inoculated box turtle (Terrapene ornata
orna ta) ................................................................................9 1

4-2 Photograph taken 12 days post-inoculation showing white caseous diphtheric
plaques in the mouth of an IM inoculated red-eared slider (Trachemys script
eleg a n s). .......................................................... ................ 92

4-3 Photograph showing exophthalmus, conjunctivitis and hyphema in an
intramuscularly inoculated red-eared slider (Trachemys script elegans) .............93

4-4 Photograph showing colonic hemorrhage in a turtle intramuscularly inoculated
with Ranavirus euthanized 23 days post inoculation....................... ...............94

4-5 Spleen; red-eared slider (Trachemys script elegans) ..........................................95

4-6 Spleen; red-eared slider (Trachemys script elegans) ..........................................96

4-7 Liver; red-eared slider (Trachemys script elegans)................... ...............97

4-8 Kidney; red-eared slider (Trachemys script elegans)................... ............... 99

4-9 Colon; red-eared slider (Trachemys script elegans) intramuscularly inoculated
w ith R anavirus. .................................................................... 100

4-10. Oral mucosa; red-eared slider (Trachemys script elegans) intramuscularly
inoculated w ith R anavirus ......... ................. ................................ ............... 101

5-1 Photograph demonstrating injection of virus infected cell culture media into a
ventral lymph sac in a leopard frog (Ranapipiens). ....................... ...........119

5-2 Photograph demonstrating the placement of a feeding tube in a red-eared slider
(Trachemys script elegans) for administering frog homogenates directly into
the caudal esophagus................................................ .. ... ....... .. ........ .... 120

5-3 Photomicrograph demonstrating the normal architecture of a liver in a leopard
frog (Rana berlandieri). ............................................... .............................. 121

5-4 Photomicrograph demonstrating multifocal hepatic necrosis in a leopard frog
(Rana berlandieri) experimentally inoculated with a Ranavirus isolated from a
Burmese star tortoise (Geochelone platynota) ...................................................... 122









5-5 Photomicrograph at higher magnification of the leopard frog (Rana berlandieri)
liver shown in Fig. 5-4. Arrows denote the presence of intracytoplasmic
inclusion bodies consistent with iridovirus infections in amphibians..................123

5-6. ELISA results graphed as positive to negative (P/N) ratios. Samples were
assayed weekly for the duration of the study. ........................................................ 124

6-1. Effect of increasing concentrations of acyclovir on the TCID50 of Terrapene
cells inoculated with the Burmese star tortoise isolate. .......................................134















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

IRIDOVIRUS INFECTIONS OF CAPTIVE AND FREE-RANGING CHELONIANS IN
THE UNITED STATES


By

April J. Johnson

December 2006

Chair: Elliott Jacobson
Major: Veterinary Medical Sciences

Iridoviruses of the genus Ranavirus are well known for causing mass mortality

events of fish and amphibians with sporadic reports of infection in reptiles. The objective

of this study was to characterize Ranavirus infections of chelonians. First,

histopathologic and molecular investigations of naturally occurring infections in several

species of chelonian were investigated. A virus isolate (BSTRV) obtained from a captive

Burmese star tortoise (Geochelone platynota) was experimentally inoculated into western

ornate box turtles (Terrapene ornata ornata) and red-eared sliders (Trachemys script

elegans). Oral transmission failed to create illness, however five of six turtles inoculated

intramuscularly developed clinical and histologic lesions consistent with naturally

infected cases. Virus was re-isolated, fulfilling Koch's postulates and establishing

BSTRV as a causative agent of disease and mortality in chelonians. Restriction enzyme

analysis of this isolate with an isolate from a leopard frog (Rana utricularia) obtained at

the site where the tortoise died was found to have identical restriction patterns suggesting









they are either the same or very closely related strains. This indicates that amphibians

might serve as a source of infection for chelonians, or vice versa. BSTRV was also

utilized as a coating antigen in the development of an indirect enzyme linked

immunosorbent assay (ELISA). Plasma from a surviving pen-mate of the Burmese star

tortoise served as a positive control for optimization. A seroprevalence study of 1000

banked free-ranging gopher tortoise plasma samples found that only 1.5% of tortoises

were positive for exposure to the virus. The role of amphibians in the route of

transmission of virus was assessed by experimentally inoculating leopard frogs,

euthanizing them, homogenizing them, and feeding them to turtles via feeding tubes over

a six-week period. All turtles failed to develop clinical signs or to produce anti-

Ranavirus antibodies over three months. Lastly, the antiviral compound acyclovir was

assessed at 0, 0.2, 1, 5, 10, and 25[tg/ml for its ability to reduce or eliminate virus

replication in vitro and to create cytotoxicity in Terrapene heart cells. No cytotoxicity

was observed at any concentration. Increasing concentration found only a slight ten fold

reduction in virus titer from 104.8 to 103 TCID50.














CHAPTER 1
INTRODUCTION



Chelonians are long-lived animals within the class Reptilia. The order Chelonia,

also sometimes called Testudines, consists of two suborders and thirteen families of

turtles and tortoises. Chelonians worldwide are experiencing dramatic declines. The

2006 International Union for Conservation of Nature (IUCN) Red List of Threatened

Animals has listed 26 species as critically endangered, 45 as endangered and 58 as

vulnerable of 295 species (43.7%). An additional 41 (13.9%) species are listed as near

threatened. Other earlier estimates indicated about 50% of all taxa to be experiencing

difficulties (Jacobson et al., 1999). Reasons for declines include habitat fragmentation,

increased collections for the food and pet market, change in vegetation, drought, and

debilitating diseases (Jacobson et al., 1999; Dodd, 2001). Chelonians have low fecundity

and low juvenile survival rates, indicating that a loss of adult animals can have a

significant impact on population survivability (Heppell, 1998). Native species of

chelonians are experiencing similar declines. Two of the three tortoises within the U.S.

[desert tortoise (Gopherus agassizii) and gopher tortoise (G. polyphemus)] are listed as

vulnerable by the IUCN. Continuing decline and disappearance of box turtle (Terrapene

spp) populations across the genus' entire range during the last century brought the 1995

listing of all box turtle species in appendices of The Convention on International Trade in

Endangered Species (CITES) in an effort to slow their declines. Progress toward

conserving these species is needed now before they become endangered.









Several important diseases have been identified in populations of wild and captive

chelonians. Marine turtle fibropapillomatosis is a significant health problem affecting

several species of marine turtles around the world (Herbst and Jacobson, 1995).

Mycoplasmosis is a chronic infectious disease that has been seen in wild gopher tortoises

in Florida and desert tortoises in the southwest deserts of the United States (Brown et al.,

2002). Tortoise herpesviruses have emerged as important pathogens of captive tortoises

in the pet trade. Tortoise herpesvirus-1 is a causative agent of rhinitis-stomatitis complex

in a variety of tortoise species (Origgi et al., 2004) and tortoise herpesvirus-2 has also

been seen associated with rhinitis and stomatitis in a captive desert tortoise (Johnson et

al., 2005). A study looking at all wild reptile cases submitted to the Wildlife Center of

Virginia between 1991 and 2000 showed that 2% (n=694) of all cases were a result of

infectious disease, although further characterization was not described. A total of 15

cases were infectious, of which 14 were eastern box turtles (Terrapene carolina carolina)

and the other case was a rat snake. Two cases had respiratory tract infections, nine had

conjunctivitis while the other four had both respiratory tract infections and conjunctivitis

(Brown and Sleeman, 2002). This dissertation will demonstrate that Ranaviruses are also

important emerging pathogens in wild and captive tortoises, and box turtles in the United

States. This virus had previously been observed in a gopher tortoise (Westhouse et al.,

1996) and a box turtle (Mao et al., 1997) in the United States. However, research

presented here will show that it is likely responsible for other past and recent die-offs of

box turtles in the eastern U.S. and may be a cause of unexplained deaths and population

declines of gopher tortoises in Florida.









Iridoviruses are large double stranded cytoplasmic DNA viruses (Williams, 1996)

that were first discovered in crane fly larvae in 1954 (Xeros, 1964) exhibiting a blue

iridescence below the epidermis, which later led to the name iridescent virus or iridovirus

(Williams and Smith, 1957). Only the insect viruses are known to create iridescence,

whereas vertebrate iridoviruses do not. Iridoviruses are circularly permuted and

terminally redundant (Goorha and Murti, 1982). Unlike poxviruses, which have a

completely cytoplasmic site of replication (Schramm and Locker, 2005), iridoviruses

require both the nucleus and cytoplasm for replication (Goorha et al., 1978).

The family Iridoviridae consists of four genera. Two genera, Chloriridovirus and

Iridovirus, infect insects. Viruses in the genus Iridovirus are typically smaller than those

in the genus Chloriridovirus. Viruses in the genera Lymphocystivirus and Ranavirus are

capable of infecting ectothermic vertebrates. Lymphocystiviruses infect fish, while

ranaviruses have been shown to infect fish, amphibians and reptiles (Mao et al., 1997). A

group of unclassified erythrocytic viruses have also been attributed to the Iridoviridae

family (Johnsrude et al., 1997; Telford and Jacobson 1993); however further

characterization is needed to establish the phylogenetic relationship of these viruses to the

classified iridoviruses.

Frog virus 3 (FV3), the type species for the genus Ranavirus, was first isolated in

1966 from a renal carcinoma in a leopard frog (Granoff et al., 1965), although it was

subsequently determined that there was no association of the virus to the tumor (Granoff

et al., 1966). A Ranavirus also was recovered from bullfrog tadpoles manifesting a

syndrome called tadpole edema virus (TEV) (Wolf et al., 1968). In experimental studies,

this virus was capable of infecting and causing significant mortality in Great Basin









spadefoot toads (Scaphiopus hammondii intermontanus), American toads (Bufo

americanus), Fowler's toads (Bufo woodhousiifowleri), and bullfrogs (Rana catesbeiana)

(Wolf et al., 1968). A subsequent experimental study demonstrated Fowler's toads and

the newt (Diemictylus viridiscens) to be susceptible to TEV, as well as two other isolates

from frogs (LT1 and FV1) and two newt isolates (T8 and T15) (Clark et al., 1969).

Significant research with amphibian iridoviruses did not make much progress until

the early 1990s when worldwide declines in amphibians brought new interest regarding

the role of these viruses in amphibian mortality events (Bradford, 1991; Speare and

Smith, 1992; Fellers and Drost, 1993; Cunningham etal., 1996; Fisher and Shaffer, 1996;

Laurance et al., 1996; Jancovich et al., 1997; Bollinger et al., 1999; Lips, 1999; Green et

al., 2002; Docherty et al., 2003). In a study of sixty-four amphibian mortality and

morbidity events, iridovirus was the most common cause of mortality (Green et al.,

2002). Late larval forms were more susceptible and epizootics were clearly associated

with increased population densities. Affected salamanders exhibited problems with

buoyancy, the inability to stay upright, swimming in circles, lethargy and red spots or

swollen areas on the ventrum near the gills or hind limbs (Docherty et al., 2003).

While iridoviruses have been occasionally reported as pathogens of reptiles

(Marschang et al., 2005; Hyatt et al., 2002; Drury et al., 2002; Johnsrude et al., 1997),

they have not received as much attention compared to iridovirus infection of amphibians.

In chelonians, the first report of an iridovirus infection involved a captive Hermann's

tortoise (Testudo hermanni) that died with necrotizing lesions in the liver, intestine and

spleen (Heldstab and Bestetti, 1982). Several years later, an epidemic in a captive group

of Hermann's tortoises was reported (Muller et al., 1988). While two iridoviruses were









identified in an exotic tortoise (Testudo horsfieldii) and a box turtle (Terrapene carolina)

in the U.S., no disease or pathology was mentioned (Mao et al., 1997). The only report in

a wild tortoise involved a gopher tortoise (Gopher polyphemus) in Florida that had signs

of respiratory disease (Westhouse et al., 1996). While viral particles were seen on

electron microscopy, there was no attempt at virus isolation or molecular

characterization. Recent isolates from two of seven Hermann's tortoises that died in a

zoo in Switzerland were found by polymerase chain reaction (PCR) to have major capsid

protein sequences closely related to FV3 (Marschang et al., 1999). Around this time,

iridovirus infections were documented in soft-shelled turtles (Trionyx sinensis) exhibiting

cervical cutaneous erythema or "red-neck disease" at a turtle farm in China (Chen et al.,

1999). In experimental infections with the iridovirus isolated from the soft-shelled

turtles, the virus was shown to be a causative agent of the "red neck" syndrome in young

inoculated turtles. Most recently, Ranavirus infections were identified in a group of

seven captive eastern box turtles in North Carolina in 2002 (DeVoe et al., 2004). Six of

these seven were wild caught with the most recent having been added 6 months prior to

the outbreak. Clinical signs associated with Ranavirus infections in these box turtles

were cutaneous abscessation, oral erosions or abscessation and respiratory distress

(DeVoe et al., 2004). One of seven affected turtles also showed unilateral conjunctivitis

and cellulitis of the head and neck.

Ranaviruses are variably host specific and are widespread geographically

(Chinchar, 2002, Daszak et al., 1999). Virus can grow in multiple types of cell lines

including fish, amphibian, reptilian, avian and mammalian, provided temperatures are

conducive to growth (Chinchar, 2002). Evidence of viruses being capable of infecting









multiple species of animals has been demonstrated both naturally and experimentally.

Inter-class infections with Ranavirus may occur among sympatric species in the wild,

with fish or amphibians serving as the reservoir host (Mao et al., 1999). Moody and

Owens (1994) demonstrated an anuran virus, Bohle iridovirus, to be pathogenic for a fish,

Lates calcarifer. Experimental transmission of Bohle iridovirus, a virus isolated from

amphibians in Australia, was experimentally inoculated into six species of reptiles of

which two turtle species, Emydura krefftii and Elseya latisternum, appeared to be

susceptible showing increased mortality in inoculated hatchlings (Ariel, 1997). An insect

iridovirus was isolated from a chameleon, two bearded dragons, and a frill-neck lizard

(Just et al., 2001). Thus, some of these viruses are very unusual in their ability to infect

phylogenetically distinct lineages of vertebrates and invertebrates. However, other

isolates have been shown to be very host specific and cannot be transmitted

experimentally to other classes of animals (Jancovich et al., 2001).

Natural transmission of iridoviruses has yet to be definitively identified, and may

vary between genera or species of viruses. Experimental studies have shown that

cannibalism of infected animals or ingestion of infected water may serve as a route of

infection in amphibians (Jancovich et al., 2001; Pearman et al., 2004). Experimental

infections of salamanders with a Ranavirus showed that both dose and host

characteristics influenced the virulence of infection (Brunner et al., 2005). The infection

dose was positively correlated with mortality rate and inversely related to average

survival times. Environmental temperatures have also been shown to significantly impact

the percent mortality and time to death in salamanders experimentally infected with a

Ranavirus (Rojas et al., 2005), where salamanders infected at 18 and 100C were more









likely to die than those exposed at 260C. Two antimicrobial peptides, esculentin-2P and

ranatuerin-2P, isolated from the skin of leopard frogs (Ranapipiens) are able to inactivate

Frog Virus 3 in a dose dependent manner (Chinchar et al., 2001) and may play a role in

viral resistance.

The immune response of reptiles to iridoviruses has not been previously

investigated; however, some research has been performed in this area for amphibians.

Birds, reptiles and amphibians produce a low-molecular weight immunoglobulin called

IgY (reviewed in Warr et al., 1995). A study of 21 wild caught cane toads (Bufo

marinus) in Townsville, Australia, found three toads to have anti-iridovirus antibodies

cross-reactive against both epizootic haematopoetic necrosis virus and bohle iridovirus on

enzyme linked immunosorbent assay (ELISA). Sera from positive toads or toads that had

been exposed to an iridovirus showed positive to negative (P/N) ratios of 2.81, 2.91 and

3.4 compared to sera from naive toads, which ranged from 0.55 to 1.13 (Whittington et

al., 1997). Other studies in amphibians have focused on the African pipid frog, Xenopus

laevis. One study experimentally infected adult frogs four weeks apart. Anti-FV3 IgY

was detected in plasma one week following the second injection. This appearance of

antibodies correlated to the time of viral clearance and the amelioration of clinical signs

(anorexia and cutaneous erythema), suggesting a role of the adaptive immune system in

clearing infection (Gantress et al., 2003). This study also suggests the genotype of the

MHC to play a significant role in the host susceptibility. Inbred Xenopus having a

decreased MHC class I expression were more susceptible to infection, as were Xenopus

larvae, which also lack MHC class I gene expression (Gantress et al., 2003). Another

study looking at antibody response to FV3 in Xenopus also showed that a second









exposure was necessary for developing a detectable response of IgY production (Maniero

et al., 2006). Antibodies were first detectable 10 days after a second exposure and levels

plateaued at 14 days. IgM levels were never detectable even after three exposures.

Prevention of iridovirus infections has been researched primarily in fish. A vaccine

against red seabream iridovirus (RSIV) has shown that genetic vaccines can be effective

in protecting red seabream (Pagrus major) against experimental challenge with RSIV

(Caipang et al., 2006). Inoculation of juvenile red seabream with DNA plasmids

encoding either the major capsid protein gene or an open reading frame encoding a

transmembrane domain against RSIV showed upregulation of transcription of the MHC

class I gene. Additionally, DNA vaccinated fish showed lower mortalities after

subsequent exposure to RSIV than did non-vaccinated fish. Formalin-killed virus has

also been shown to be effective in upregulating MHC class I transcripts (Nakajima et al.,

1997; Nakajima et al., 1999). A field trial testing the efficacy of a formalin-killed virus

showed a 49.3% decrease in mortality between vaccinated and control groups, and a

statistically significant increase in size of fish in the vaccinated group (Nakajima et al.,

1999). A recent study evaluated an environmental contaminant and its potential to

contribute to increased mortalities in iridovirus infected amphibians (Forson and Storfer,

2006). Somewhat surprisingly, the authors found that atrazine at moderate doses may

reduce the efficacy of iridoviruses and, thus, protect animals exposed to both atrazine and

iridovirus, resulting in decreased mortality rates. However, high levels of exposure to

atrazine could result in decreased fitness. Salamanders were metamorphosed earlier and

were smaller at metamorphosis than those exposed to no or moderate levels of atrazine

(Forson and Storfer, 2006).









Treatment of iridovirus infections with antiviral drugs are not reported in the

literature; however previous research has suggested that antiviral treatment with acyclovir

might be successful. Acyclovir becomes activated by a viral gene product, an enzyme,

thymidine kinase. Sequencing of the entire genomes of several isolates has identified

open reading fames encoding putative thymidine kinase (TK) genes, and functionality

was further studied in the Bohle iridovirus (BIV) (Coupar et al., 2005). The BIV TK

gene was inserted into a TK gene negative mutant vaccinia virus insertion plasmid, under

the control of a vaccinia virus promoter and used to infect human 143B (TK-) cells. The

investigators were able to show that this gene expressed a functional TK gene by rescuing

of the TK negative mutant. Comparison of sequences of different iridovirus TK genes to

other virus TK genes showed that unlike poxviruses, iridoviruses, similar to the

herpesviruses, appear more closely related to the mitochondrial TKs and to cellular

deoxycytidine kinases, whereas the TK gene sequence in other DNA viruses including

poxviruses and African swine fever virus appear more closely related to the cellular TK

genes (Coupar et al., 2005). Thymidine kinase genes can activate nucleoside analogs

such as acyclovir if the gene can recognize deoxycytidine as an alternative substrate. The

ability of different iridovirus TKs to do this has not been evaluated. While TK genes

appear well conserved among the different genera, they vary considerably between

genera within the family, so each would need to be evaluated separately.

The research presented in the following chapters will investigate several aspects of

Ranavirus infections in chelonians. Chapter 2 reports on molecular and histopathologic

findings of natural cases of iridovirus infections in wild and captive chelonians in the

United States. It describes several recent chelonian cases and two past die-offs of box









turtles dating back to 1998 and 1991 and demonstrates a new geographic range of

infections from the northeast to Texas. Chapter 3 will describe the development and use

of an indirect enzyme linked immunosorbent assay (ELISA) to evaluate the prevalence of

iridovirus exposure in wild gopher tortoises. Chapter 4 will describe an experimental

infection study fulfilling Koch's postulates, confirming that iridovirus is a primary

pathogen of chelonians. Chapter 5 reports on a study investigating a possible route of

transmission in which ingestion of infected amphibians could be a source of iridovirus in

a natural setting. Chapter 6 will describe the use of an antiviral drug, acyclovir, at

reducing replication of iridovirus in vitro. And lastly, chapter 7 will discuss overall

conclusions and directions for future research.














CHAPTER 2
RANAVIRUS INFECTION OF FREE-RANGING AND CAPTIVE BOX TURTLES
AND TORTOISES IN THE UNITED STATES

Introduction

It has been suggested that chelonians (turtles and tortoises) face a more serious

threat than that posed by the well-publicized decline of amphibian populations (Klemens,

2000). Two thirds of all species of freshwater turtles and tortoises are currently listed as

threatened on the IUCN Red List of Threatened Species (Turtle Conservation Fund,

2002). Chelonians have low fecundity, low juvenile survival rate, and a long adult

lifespan; a life history strategy where loss of adult animals (such as loss by disease) has a

significant impact on population recovery (Heppell, 1998). Emerging infectious diseases

have been increasingly recognized as factors influencing wildlife health and populations

(Harvell et al., 1999; Daszak et al., 2000). Although mycoplasmosis has been postulated

to contribute to declines of some tortoise species (USFWS, 1994), the causes) of mass

mortality events in wild chelonian populations often remain undetermined (Flanagan,

2000; Dodd, 2001).

Among the emerging diseases of wildlife, iridoviruses in the genus Ranavirus, are

well known for causing mass mortality events of fish and amphibians (Langdon and

Humphrey, 1987; Daszak et al., 1999; Green et al., 2002). Iridovirus infections have also

been sporadically described in reptiles including snakes (Hyatt et al., 2002), lizards

(Marshang et al., 2005) and chelonians. In chelonians, iridovirus infections have been

reported in captive Hermann's tortoises (Testudo hermanni) (Heldstab and Bestetti, 1982;









Muller et al., 1988; Marschang et al., 1999), farmed soft-shelled turtles (Chen et al.,

1999) and captive eastern box turtles (DeVoe et al., 2004). Two ranaviruses identified in

an exotic tortoise (Testudo horsfieldii) and a box turtle (Terrapene carolina) in the U.S.,

were found to be closely related to frog virus 3 (FV3) and designated as tortoise virus 5

and turtle virus 3, respectively (Mao et al., 1997). The only report of an iridovirus

infection in a free-ranging chelonian involved a gopher tortoise (Gopher polyphemus) in

Florida with signs of respiratory disease (Westhouse et al., 1996).

This report identifies Ranavirus infections in five recent chelonian deaths or

mortality events from Georgia, Florida, New York, and Pennsylvania and in archived

material recovered from previously unexplained mass mortality events in 1991 from

Georgia and 1998 from Texas. This demonstrates a previously undescribed geographic

extent of chelonian Ranavirus infections and suggests that ranaviruses may be more

important pathogens of free-ranging chelonians than anticipated from previous reports.

We also present molecular evidence for an identical or similar virus in frogs in the

vicinity of two chelonian epizootics suggesting that amphibians could serve as reservoir

hosts for chelonians.

Materials and Methods

Animals

Burmese star tortoises. Three female and two male captive Burmese star tortoises

(Geochelone platynota) were kept in an outdoor enclosure at St Catherine's Island

Wildlife Survival Center, Georgia (31040'N/81010'W) since April 2001. In early June

2003, two female and one male tortoise began showing clinical signs consisting of nasal

discharge, conjunctivitis and severe subcutaneous edema of the neck (Fig. 2-1A). The

tortoises were treated with antimicrobials, and were soaked daily for 90 minutes in warm









water. One female tortoise died three days after developing clinical signs and yellow

white caseous plaques were observed on the tongue at necropsy. Oral antiviral therapy

and intracoelomic fluids were then initiated in the surviving tortoises. Subsequently, six

adult Southern leopard frogs (Rana utricularia) were sampled from within the tortoise

pens and one was sampled from a pond nearby. One of the frogs from the pen was found

moribund while others appeared healthy. Tissues from the dead tortoise and the leopard

frogs were submitted for histopathology, polymerase chain reaction (PCR) for

determining presence of certain DNA sequences of Ranavirus and Herpesvirus, and virus

isolation.

Gopher tortoise. A wild gopher tortoise (Gopheruspolyphemus) was found

circling on a road in north central Florida (29086'N/82022'W) on 25 July 2003 and was

brought to the University of Florida, College of Veterinary Medicine Zoological

Medicine Service for evaluation and treatment. On presentation it exhibited palpebral

swelling and ocular and nasal discharge (Fig 2-1B). The tortoise was treated with

intracoelomic fluid twice daily, was allowed to soak in shallow warm water for 20

minutes a day and was started on antimicrobial therapy. The tortoise's condition

continued to decline and it was euthanized with intravenous Beuthanasia-D solution

(Schering-Plough Animal Health Corp., Kenilworth, NJ) on 29 July 2003. A complete

necropsy was performed and tissues were submitted for histopathology, PCR and virus

isolation.

Eastern box turtles. A 200 ha gated and fenced area within a private nature

sanctuary in Venango County, Pennsylvania, (4143'N/79093'W) was created as a study

site for relocated box turtles (Terrapene carolina carolina). All box turtles within the









study site were radiotelemetered and tracked regularly, and no box turtles were present at

the site prior to the release of the relocated turtles. During the summer of 2003, there

were 32 adult and 34 head-started juvenile turtles. Turtles were observed every five days

to determine their health status and location. Fifteen of the 66 turtles (23%) died between

15 August and 20 November 2003. Many of the turtles were considered healthy

approximately four to eight days prior to being found either moribund or dead. Nine of

the turtles were found dead, while six were found moribund with palpebral edema, ocular

discharge, and fluid draining from the mouth. Moribund turtles were taken out of the

preserve, treated with an ophthalmic ointment in the eyes, soaked daily in warm water

and given a temperature gradient. Two turtles were started on antimicrobials and one of

the two also received a parasiticide. All moribund turtles died within hours to days after

being found exhibiting signs of illness. In May 2004, two freshly dead green frog

tadpoles (Rana clamitans) with marked cutaneous erythema were collected from a pond

at the nature preserve. Tissues from five turtles collected in 2003 and from the tadpoles

collected in 2004 were submitted for histopathology, PCR and virus isolation.

Two wild box turtles were found moribund with ocular discharge and swelling, as

well as aural abscesses and yellow caseous plaques in the oral cavity in Suffolk County,

New York (4051 'N/7252'W) on 2 August 2005. They were observed near the edge of a

drying pond that is utilized by many pond-breeding amphibians, including green frogs

(Rana clamitans) and bullfrogs (Rana catesbeiana). These animals were taken to a local

wildlife rehabilitator, where one died overnight. The carcass was then put in a freezer for

later evaluation. The other had the aural abscesses drained and was treated with

antimicrobials, but its health continued to decline until its death on 1 September 2005.









This turtle was put on ice and, along with the frozen specimen, sent to the University of

Florida for histopathology, PCR and virus isolation.

Florida box turtle. A wild Florida box turtle (Terrepene carolina bauri), found in

north central Florida (29042'N/82023'W) in October 2004, was submitted to the

Zoological Medicine Service, College of Veterinary Medicine, University of Florida, for

treatment. The box turtle exhibited palpebral edema, nasal and ocular discharge (Fig. 2-

1C) and had yellow-white caseous plaques in the oral cavity (Fig. 2-1D). The turtle was

administered fluid intracoelomically with B vitamins daily and was started on analgesics

to alleviate pain. Due to failure to respond to therapy and a poor prognosis the turtle was

euthanized three days after admission with intravenous Beuthanasia-D solution. A

complete necropsy was performed and tissues submitted for histopathology, PCR and

virus isolation.

Past mortality events. Tissues from two previous box turtle epizootics of

undetermined etiology were examined. In July and August 1991 over thirty Eastern box

turtles were found dead in or near water sources in Murray county (34045'N/84047'W),

northwest Georgia (Dodd, 2001). Two moribund turtles were found exhibiting lethargy,

ocular discharge and had caseous white plaques in the oral cavity. One turtle also had a

subcutaneous abscess caudal to the left eye. Both turtles were submitted for necropsy. In

1998, several Eastern box turtles and other unspecified turtle species died suddenly in a

private collection in Texas (Dodd, 2001). Archived paraffin blocks for histologic

examination were obtained from two box turtles from the Georgia die off and one box

turtle from Texas and re-evaluated using light and transmission electron microscopy.









Necropsy and Histopathology

At necropsy, tissues were collected from all major organ systems of the following

tortoises and turtles: Burmese star tortoise (1), gopher tortoise (1), eastern box turtles

[Pennsylvania (5), Georgia (2), New York (1), and Texas (1)] and Florida box turtle (1).

Tissues were fixed in neutral buffered 10% formalin, dehydrated in graded alcohols,

embedded in paraffin, sectioned at 6[tm, and stained with hematoxylin and eosin.

Tongue, liver and spleen of each animal were collected and frozen at -800C for detecting

DNA sequences of Ranavirus and Herpesvirus using PCR and virus isolation.

Nucleotide Amplification, Sequencing, and Sequence Analysis

DNA was extracted from chelonian and amphibian tissues and cell cultures used

for virus isolation using the DNeasy kit (Qiagen, Valencia, CA, USA). Five 5tm thick

paraffin embedded sections from box turtles from the 1991 and 1998 mortality events

were extracted using the DNeasy kit following the protocol for paraffin embedded tissue.

Sense primer (5'-GACTTGGCCACTTATCAC-3') and anti-sense primer (5'-

GTCTCTGGAGAAGAAGAA-3') as previously described (Mao et al., 1997) were used

to amplify approximately 500 base pairs of the Ranavirus major capsid protein gene. A

50[tl reaction mixture was run which contained 4[l extracted DNA, 1 pM of each primer,

200 [tM each of dATP, dCTP, dGTP, and dTTP, 2.5 U of Taq DNA polymerase and PCR

buffer containing 50 mM KC1, 10 mM Tris-HC1, 1.5 mM MgC12 (Eppendorf,

Westbury, New York, USA). The mixtures were amplified in a thermal cycler (PCR

Sprint, Thermo Hybaid) with an initial denaturation at 94C for 2.5 min, followed by 25

cycles of denaturation at 94C for 30 sec.; annealing at 50C for 30 sec, extension at 72C

for 30 sec., and a final extension step at 72C for 10 min as previously described









(Marschang et al., 1999). The same extracted DNA from the chelonian tissues was

evaluated by PCR for the presence of chelonian herpesvirus(es). A nested consensus

PCR was performed as previously described to detect a portion of the herpesvirus DNA

dependent DNA polymerase (VanDevanter et al., 1996).

Any PCR products were resolved in 1% agarose gels and bands were excised and

purified using the QIAquick gel extraction kit (Qiagen). Products were sequenced in

both directions directly using the Big-Dye Terminator Kit (Perkin-Elmer, Branchburg,

New Jersey) and analyzed on ABI 377 automated DNA sequencers at the University of

Florida's Sequencing Center.

Virus Isolation

Turtle heart cells [TH-1; American Type Cell Culture (ATCC), Manassas, VA]

were seeded into 25 cm2 flasks (Costar, Coming, NY, USA). Cells were cultured in

Dulbecco's modified Eagle medium (DMEM, Gibco, Carlsbad, CA, USA) supplemented

with 5% fetal bovine serum (Gibco, Carlsbad, CA, USA), gentamicin (60 mg/liter)

(Sigma, St. Louis, MO, USA), penicillin G (120,000 U/liter), streptomycin (120,000

U/liter) and amphotericin B (300[tg/liter) (Sigma) and cultured to confluency. A small

piece of spleen or liver from each case was homogenized in separate 5ml tissue grinders

containing DMEM. Part of each homogenate was applied to a flask of confluent

monolayer of TH-1 cells while the other was passed through a 0.45[tm filter (Costar)

onto another flask of cells. Cells were incubated at 280 C. Flasks were observed daily for

cytopathic effect (CPE).









Transmission Electron Microscopy

Second passage TH-1 cell monolayers (75mm2 flasks) inoculated with first passage

isolates from homogenates of kidney tissue collected at necropsy from the Burmese star

tortoise and the gopher tortoise were examined by transmission electron microscopy

(TEM). Cells were harvested 3 days after infection (2 days after observation of CPE) and

centrifuged at 4,500 x g for ten minutes. Supernatant was discarded and the remaining

pellet was suspended in Trump's fixative (4% paraformaldehyde, 1% gluteraldehyde).

Cells were post-fixed in osmium tetroxide, dehydrated in graded alcohols and embedded

in Spurr's resin. Tissues were viewed using a Hitachi H7000 transmission electron

microscope at the University of Florida Electron Microscopy Laboratory.

Paraffin embedded spleen from a box turtle from the 1991 mass mortality event in

Georgia and paraffin embedded trachea from a box turtle from Texas from the 1998 mass

mortality event were deparaffinized in xylene, embedded in Spurr's resin, sectioned for

TEM and examined as described for cell culture.

Skin from the necropsied Burmese star tortoise was placed in Trump's solution

(McDowell and Trump, 1976) was submitted to the Athens Diagnostic Laboratory,

University of Georgia for TEM. Tissue was post fixed in osmium tetroxide, dehydrated

in graded alcohols and embedded in epoxy. Ultrathin sections were stained with uranyl

acetate and lead citrate and examined with a JEOL JSM-1210 transmission electron

microscope.

Restriction Enzyme Analysis

Frog Virus 3 was obtained from ATCC and served as the positive control for

comparative purposes with the viruses isolated in this study. Frog Virus 3 and second

passage isolates of tissue homogenates from the Burmese star tortoise and Southern









leopard frog that were collected at the same site were inoculated onto TH-1 cells. Once

CPE was observed, viral DNA was radiolabeled with [methyl-3H] thymidine, extracted

and digested with HindlII and Xbal endonucleases as previously described (Mao et al.,

1999). Restricted DNA fragments were separated on a 0.7% agarose gel, after which the

gel was impregnated with Enhance (Perkin Elmer, Wellesley, MA) according to the

manufacturer's directions and the radiolabeled fragments were detected by fluorography.

Results

Necropsy and Histopathology

Histologic findings were similar in the Burmese star tortoise, Gopher tortoise and

box turtles. Consistent lesions in all animals were necrotizing and ulcerative stomatitis or

esophagitis, fibrinous and necrotizing splenitis, and multicentric fibrinoid vasculitis.

Lesions in the oral cavity and esophagus were characterized by near diffuse mucosal

erosion and ulceration with surfaces covered by a thick coagulum comprised of fibrin,

degenerate heterophils, sloughed epithelial cells and bacterial colonies (Fig. 2-2).

Lesions in the spleen consisted of disruption of the white and to a lesser degree, the

surrounding red pulp by deposits of fibrin admixed with karyorrhectic debris, and

infiltrates of small numbers of heterophils (Fig. 2-3). There was frequently mild to

marked red pulp congestion and/or hemorrhage. Fibrinoid vasculitis with thrombosis was

observed in splenic, sheathed capillaries ellipsoidss) in all animals and to varying degrees

in other locations including oral mucosa, esophagus, stomach, intestine, skin, lung, heart,

and liver. All animals had some degree of necrosis of hematopoietic tissue in the kidney,

liver and bone marrow. Individual cases had multifocal necrotizing tracheitis (1/7),

conjunctivitis (1/7) or gastritis (2/7). Rarely, basophilic intracytoplasmic inclusion

bodies suggestive of iridovirus infection were observed within epithelial cells of the oral









mucosa, esophagus, stomach and trachea, and/or within endothelial cells, macrophages

(Fig. 2-4) and hematopoietic progenitor cells. Inclusion bodies were observed in only 3

of 7 (Burmese star tortoise, eastern box turtle, and gopher tortoise) cases examined

histologically.

Necrosis of hepatic and renal hematopoietic tissues with rare basophilic

intracytoplasmic inclusion bodies consistent with iridovirus infection was observed in the

moribund southern leopard frog from the Burmese star tortoise pen in Georgia and in one

green frog tadpole from the site in Pennsylvania.

Archived tissues from the past two mortality events in Georgia and Texas showed

similar histologic lesions as described above. These included fibrinous splenitis in all

turtles as well as a necrotizing tracheitis with rare intracytoplasmic basophilic inclusion

bodies in sloughed respiratory epithelial cells in the box turtle from Texas.

PCR and Sequence Analysis

PCR for the Ranavirus major capsid protein gene yielded DNA fragments

approximately 500 base pairs in length (Fig. 2-5). After sequencing of the fragments and

excluding the primer component, the sequences of the gopher tortoise, star tortoise, all

box turtles, southern leopard frog and green frog shared 100% sequence identity. The

sequences were compared to known sequences in GenBank (National Center for

Biotechnology Information, Bethesda, Maryland), EMBL (Cambridge, United Kingdom),

and Data Bank of Japan (Mishima, Shiuoka, Japan) databases using TBLASTX

(Altschul, et al., 1997). TBLASTX results for the sequences all showed the highest score

with FV3 capsid protein gene (GenBank accession # AF157769). Comparison of

sequences showed that all isolates shared 100% sequence identity with FV3 across that

portion of the major capsid protein gene. PCR of DNA extracted from paraffin









embedded sections of tissue from box turtles from the 1991 and 1998 die offs were

negative. All tortoises and box turtles were negative by PCR for the presence of

herpesvirus.

Virus Isolation

All TH-1 cells infected with tissue homogenates from the dead turtles and tortoises

exhibited cytopathic effects (CPE) that consisted of cell rounding and lysis two to three

days post infection. Flasks that contained both filtrated tissue homogenate as well as

unfiltered tissue homogenate showed CPE. The Burmese star tortoise isolate was also

passage onto fathead minnow cells (ATCC), which subsequently exhibited similar CPE.

Transmission Electron Microscopy

Using TEM, the cell cultures infected with tissue homogenates from the Burmese

star and gopher tortoises showed large numbers of icosahedral shaped viral particles that

were consistent in size (approximately 130nm) and shape with an iridovirus (Fig. 2-6).

Similar viral particles were observed within intracytoplasmic inclusion bodies in

endothelial cells and macrophages of the skin in the Burmese star tortoise, within the

cytoplasm of unidentified cells in the spleen of a 1991 box turtle from Georgia and within

intracytoplasmic inclusion bodies of degenerate respiratory epithelial cells of the trachea

in the 1998 box turtle from Texas (Fig. 2-7).

Restriction Enzyme Analysis

Repeated attempts failed to show discrete bands of the green frog tadpole after

restriction with both enzymes, and thus, comparisons cannot be made with this isolate.

Restriction with the HindIII enzyme demonstrated identical patterns between the FV3,

the Burmese star tortoise and the leopard frog isolate collected at the same location as the

star tortoise (Fig. 2-8A). The eastern box turtle isolate showed a slightly different









pattern. Restriction with Xbal found a similar result (Fig. 2-8B). FV3, the Burmese star

tortoise and leopard frog isolates showed identical patterns, while the box turtle isolate

was different.



Discussion

Emerging infectious diseases are those that have newly appeared in a population or

have previously existed but are rapidly increasing in incidence or geographic range

(Morse, 1995). The findings reported here suggest that iridovirus infections in chelonians

fill this description, and are emerging pathogens of chelonians. Infections are being

discovered in new populations of turtles, and the incidence is either increasing, or our

ability to detect the disease in these animals is increasing. Infections in chelonians are

more geographically widespread than has been previously documented. Previous reports

from the United States had identified chelonian iridovirus infections in a wild gopher

tortoise in Florida (Westhouse et al., 1996), captive box turtles in North Carolina (DeVoe

et al., 2004) and in two chelonians where the location was not reported (Mao et al.,

1997). Here we have identified more infections than previously described within a two-

year period in Georgia, Florida, Pennsylvania, and New York, indicating an increase in

incidence of infection. Identification of iridovirus-like particles in the mortality event in

1998 expands the new geographic range further to include Texas. We also have

described infection in a Burmese star tortoise, a species that has previously not been

documented to be susceptible to iridovirus infections.

Reports of mortality events involving large numbers of box turtles and gopher

tortoises have been documented in which etiologies were never definitively identified

(Dodd, 2001; Rossell et al., 2002; Seigel et al., 2003). This report demonstrates by re-









evaluation of archived samples that at least some of these mortality events (Dodd, 2001)

have been infected with a viruses) compatible with iridovirus and mortalities may have

been caused by infection with these viruses. Polymerase chain reaction of these cases

were negative, however it is unknown how long tissues had been fixed in formalin prior

to being embedded into paraffin. Formalin fixation has been shown to degrade DNA and

could result in false negative results (Tokuda et al., 1990). Still, TEM of the same tissues

showed virus particles consistent in size and shape with iridovirus.

The histologic lesions in the chelonians in this report were relatively non-specific

and the intracytoplasmic basophilic inclusion bodies suggestive of iridovirus infection

(Heldstab and Bestetti, 1982; Marschang et al., 1999; Bollinger et al., 1999; Docherty et

al., 2003) were absent or were rare and could easily be missed. Clinical and pathological

differential diagnoses for the animals in this report prior to the demonstration of

iridoviruses included chelonian herpesvirus infection (Johnson et al., 2005) for the

lesions of necrotizing stomatitis and septicemia for the fibrinous splenitis and

multicentric vasculitis. While iridovirus infection should be considered in cases with

lesions similar to those seen in the turtles and tortoises in this report, ancillary diagnostic

tests, including viral isolation attempts and PCR for ranaviruses and herpesviruses,

should be performed to confirm the diagnosis.

All chelonian and amphibian isolates in this study shared 100% sequence identity

across a portion of the major capsid protein gene. The major capsid protein gene

sequence is fairly conserved among iridoviruses, although one study has shown that it

contains enough diversity to be able to distinguish closely related isolates (Tidona et al.,

1998). Mao et al. (1997) compared MCP sequences and restriction enzyme patterns of









whole genomic DNA of 10 vertebrate iridoviruses, including one virus isolated from a

box turtle, Turtle Virus 3 (TV3). While the sequence obtained from a portion of the TV3

MCP gene indicated that it was identical to that of Frog Virus 3, a restriction enzyme

analysis using HindII and Xbal showed a different restriction pattern between the two

isolates. Our study found a similar result. An isolate from a box turtle from the

Pennsylvania die-off shared 100% sequence identity with FV3 across a portion of the

MCP gene, however the whole viral genomic restriction enzyme analysis pattern differed

from FV3, and showed a similar result to that of the box turtle isolate, TV3, reported by

Mao et al. (1997). This suggests that the major capsid protein gene may be too conserved

to determine if different animals are infected with the same virus.

An interesting and potentially significant finding was identical viruses, as

determined by restriction enzyme analysis, in the Burmese star tortoise and a moribund

southern leopard frog found within its pen. This suggests that both animals were infected

with the same virus. Inter-class infections have been shown previously in a natural

setting where sympatric species of amphibians and fish were infected with the same virus

species (Mao et al., 1999) as well as through experimental transmission studies (Moody

and Owens, 1994). There are several ways that chelonians and amphibians might be

exposed to the same virus. Previous studies in salamanders have shown that transmission

can occur through cannibalism of infected individuals (Jancovich et al., 2001; Pearman et

al., 2004). Box turtles are omnivorous, and tortoises while normally herbivorous, may

opportunistically feed on carrion. This was confirmed when animal caretakers at the site

of the Burmese star tortoise death in Georgia observed a radiated tortoise (Geochelone

radiata) and a Burmese black mountain tortoise (Manouria emysphayrei), both normally









considered herbivorous, eating dead amphibians in nearby pens. There could also be a

common environmental source of virus, such as shared bodies of water. Iridoviruses are

quite resistant and thought to be capable of persisting in water sources for extended

periods of time (Daszak et al., 1999). Iridoviruses create systemic infections, and thus, a

vector-borne route of transmission might also be a way that both amphibians and

chelonians could become infected.

In summary, this report demonstrates that Ranavirus is an emerging pathogen of

chelonians and suggest that amphibians might serve as a source of infection. This

describes a new geographic range for chelonian iridovirus infections in the United States.

Ranaviruses are considered a global threat to amphibian populations based on the lack of

host specificity, high virulence and global distribution (Daszak et al., 1999) so they

should likewise be considered a global threat to chelonian populations.


































Fig. 2-1. Gross lesions associated with iridovirus infections in turtles and tortoises. A)
Photograph of a Burmese star tortoise (Geochelone platynota) with nasal
discharge and palpebral and cervical edema. B) Photograph of a wild gopher
tortoise (Gopheruspolyphemus) with nasal discharge and palpebral edema.
C) Photograph of a wild Florida box turtle (Terrapene carolina bauri) with
palpebral edema and ocular discharge. D) Photograph of caseous plaques in
the oral cavity of a Florida box turtle.






































Fig. 2-2. Esophagus, eastern box turtle (Terrapene carolina carolina). There is diffuse
necrosis and ulceration of the mucosa and replacement by fibrin,
inflammatory cell infiltrates and superficial bacterial colonies (arrows). H&E
stain.






































Fig. 2-3. Spleen, eastern box turtle (Terrapene carolina carolina). There is disruption of
the white and red pulp with deposits of fibrin (arrow) admixed with
karyorrhectic debris, and infiltrates of small numbers of heterophils. H&E
stain.












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Fig. 2-4. Epicardium, Burmese star tortoise (Geochelone platynota). Arrows depict
basophilic intracytoplasmic inclusion bodies in a macrophage and endothelial
cell. The lumen of the blood vessel contains a fibrin thrombus (F). H&E
stain.


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Fig. 2-5. Results of a polymerase chain reaction targeting approximately 500 bp of the
major capsid protein gene. Lane 1 is a 100 bp ladder. The bright band
represents a 500 bp fragment. Lane 2 and 3 are positive samples from an
eastern box turtle (Terrapene carolina carolina) and a green frog tadpole
(Rana clamitans). Lane 4 is a positive control sample from a Burmese star
tortoise (Geochelone platynota) confirmed previously with nucleotide
sequencing. Lane 5 is a negative control.




































Fig. 2-6. Transmission electron photomicrograph of Terrapene heart cells inoculated
with liver tissue from a Burmese star tortoise (Geochelone platynota)
demonstrating cytoplasmic arrays of iridovirus-like particles.

















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Fig. 2-7. Transmission electron photomicrograph of paraffin embedded spleen from a
box turtle (Terrapene carolina) that died in 1991 in Georgia. There are
icosahedral virus particles compatible with iridoviruses.








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Fig. 2-8. HindIII and Xbal restriction enzyme pattern of five iridovirus isolates. A)
HindIII restriction. Lane 1 is Frog Virus 3, the type species for the genus
Ranavirus. Lane 2 is an isolate from a Burmese star tortoise (Geochelone
platynota) and lane 3 is an isolate from a southern leopard frog (Rana
utricularia) collected from the pen adjacent to the Burmese star tortoise. Lane
4 is an isolate from an eastern box turtle (Terrapene carolina carolina) from
Pennsylvania and Lane 5 is a green frog tadpole (Rana clamitans) isolate from
the same location as the box turtle. B) Xbal restriction pattern. Lanes are the
same as those in Fig. A.


5














CHAPTER 3
DEVELOPMENT AND USE OF AN INDIRECT ENZYME LINKED
IMMUNOSORBENT ASSAY FOR DETECTION OF IRIDOVIRUS EXPOSURE IN
GOPHER TORTOISES (Gopherus polyphemus)

Introduction

Iridovirus infections of the genus Ranavirus have recently been identified in free-

ranging and captive native chelonians from Florida, Georgia, New York, North Carolina,

Tennessee, and Pennsylvania (Allender et al., in press; DeVoe et al., 2004; Johnson et

al., 2004). Species affected were Burmese star tortoises (Geochelone platynota), gopher

tortoises (Gopheruspolyphemus), eastern box turtles (Terrapene carolina carolina), and

Florida box turtles (Terrapene carolina bauri). Evidence of iridovirus infection was also

observed in archived material from previously unexplained mass mortality events of

eastern box turtles (Terrapene carolina carolina) from Georgia in 1991 and Texas in

1998 (Dodd, 2001; Johnson et al., 2004). Clinical signs associated with infections

included those of upper respiratory tract disease including respiratory distress and nasal

discharge, as well as oral ulceration, cutaneous abscessation, anorexia and lethargy

(Westhouse et al., 1996, DeVoe et al., 2004). Consistent lesions in affected animals

included necrotizing stomatitis and/or esophagitis, fibrinous and necrotizing splenitis, and

multicentric fibrinoid vasculitis. Intracytoplasmic inclusion bodies were rarely observed

in affected tissues. A portion of the major capsid protein (MCP) gene was sequenced

from recent cases from Georgia, Florida and Pennsylvania and found to be identical

across approximately 500 basepairs to each other and to Frog Virus 3 (FV3), the type

species of the genus Ranavirus in the family Iridoviridae (Johnson et al., 2004). Koch's









postulates were fulfilled by experimentally inoculating a tortoise Ranavirus isolate into

red-eared sliders (Johnson et al., unpublished findings). Three of four sliders developed

severe clinical signs including anorexia (3/3), lethargy (3/3), oral plaques (1/3), nasal

discharge (3/3), ocular discharge (3/3) and exophthalmus, conjunctivitis, and hyphema

(1/3). Histologic changes were similar to those seen in naturally infected cases. Virus

was isolated from tissues of each of the three turtles, fulfilling Koch's postulates and

establishing iridovirus as a primary pathogen in chelonians.

Iridoviral infections are the most common cause of mortality events in

amphibians in the United States (Green et al., 2002). Iridoviruses are globally distributed

and thus considered a threat to amphibian populations worldwide based on the lack of

host specificity, high virulence and ubiquitous distribution (Daszak et al., 1999). The

geographic range of Ranavirus infections in chelonians in the U.S. has recently been

found to be larger than previously known. Prior to 2003 only three cases of chelonian

infections had been reported in the U.S.; however only one report included the location of

the infected chelonian. A wild gopher tortoise from Florida was found to have iridovirus-

like particles by transmission electron microscopy (Westhouse et al., 1996). A box turtle

and Russian tortoise (Testudo horsfieldi) isolate were described in another report (Mao et

al., 1997), but the location was not disclosed, and as Russian tortoises are not native to

the United States, it is possible they were both kept as pets. Current published and

unpublished reports now show a much larger geographic range with iridovirus infected

chelonians identified from Texas to New York and Pennsylvania (Allender et al., in

press; DeVoe et al., 2004; Johnson et al., 2004). Therefore, it is reasonable to assume









that chelonians throughout the eastern United States can be exposed to iridoviruses;

however, the prevalence rate of exposure has not been determined.

Serology can be a useful tool for detecting previous exposure to pathogens.

Indirect enzyme linked immunosorbent assays (ELISA) have been used to detect

exposure of various reptiles to specific pathogens (Schumacher et al., 1993; Origgi et al.,

2001; Brown et al., 2001; Jacobson et al., 2005) and has been used to detect exposure of

amphibians to iridovirus infections (Whittington et al., 1997; Gantress et al., 2003;

Maniero et al., 2006). To determine iridovirus exposure in wild gopher tortoises in the

U.S., we developed an indirect ELISA using a previously developed mouse anti-desert

tortoise IgY monoclonal antibody as the secondary antibody (Schumacher et al., 1993).

We also describe the results of a larger serological survey of wild gopher tortoises from

various sites in Alabama, Florida, Georgia, Louisiana and Mississippi.

Materials and Methods

Virus

A Ranavirus isolated and partially characterized from a naturally infected

Burmese star tortoise in Georgia (Johnson et al., 2004), here termed BSTRV, was used as

the antigen in the development of the ELISA. Briefly, polymerase chain reaction (PCR)

targeting a portion of ranaviral major capsid protein genes followed by nucleotide

sequencing demonstrated that the BSTRV isolate shared 100% sequence identity of

approximately 500 basepairs with FV3. Transmission electron microscopy of BSTRV

inoculated TH-1 cells showed virus particles in the cytoplasm of infected cells consistent

in size and shape with iridoviruses. Restriction enzyme digests of BSTRV compared

with FV3 showed identical restriction patterns using the enzymes HindII and Xbal,

indicating that BSTRV is either identical or very closely related to FV3.









Antigen Preparation

Two methods of antigen preparation were used. The first method of preparation

was by sucrose gradient ultracentrifugation as previously described for epizootic

haematopoietic necrosis virus of fish (Steiner et al., 1991). Terrapene heart cells (TH-1)

were acquired from the American Type Culture Collection (ATCC-CCL 50; Rockville,

MD) and grown to confluency in 225cm2 tissue flasks (Costar, Corning, NY). Cells were

cultured in Dulbecco's modified Eagle medium (DMEM, Gibco, Carlsbad, CA)

supplemented with 5% fetal bovine serum (Gibco), gentamicin (60mg/liter; Sigma, St.

Louis, MO), penicillin G (120,000 U/liter), streptomycin (120,000 U/liter) and

amphotericin B (300[g/liter; Sigma). Cells were inoculated with a fourth passage of

BSTRV and incubated at 280C in the presence of 5% CO2. When cytopathic effects

(CPE) were observed in over 70% of cells, consisting of cell rounding and detachment

from the flask, the flasks were frozen and thawed 3 times with vigorous vortexing before

each freeze. Supernatant was transferred to 15ml tubes and clarified by low speed

centrifugation at 4,500xg for 15 minutes. Supernatant was then decanted into two sterile

1-liter bottles and stored at 40C until 1.5 liters of supernatant were obtained. Virus was

then pelleted at 10,000xg for 8 hours at 40C and the supernatant was discarded. Pellets

were resuspended in the residual media, divided into four equal parts and overlayed on

four 15-60% (w/v) sucrose gradients. Gradients were then ultracentrifuged at 150,000xg

for 45 minutes at 40C. Bands of purified virus were collected by fractionation and

diluted in Tris-HCl (pH 8.0) until sucrose was less than 20%. Bands in Tris-HCl were

layered onto four 5ml 20% (w/v) sucrose cushions and ultracentrifuged at 80,000xg for

one hour at 40C. Sucrose was next decanted and pellets were resuspended in 200[tl Tris-









HC1 and stored at -800C. Purity was assessed with negative staining electron

microscopy, protein assay (Biorad, Hercules, CA), PCR and ELISA.

The second way antigen was prepared was by creating a cell lysate from virus

infected cells. Virus was inoculated onto TH-1 cell monolayers as described above in

225 cm2 flasks. Flasks were scraped when flasks exhibited 100% CPE. Uninfected

flasks were concurrently processed in the same manner to serve as control antigen to

detect any background cross reactivity of plasma to cellular proteins. Cells and media

were transferred to 15ml tubes and centrifuged at 4,500xg for 30 minutes. Supernatant

was then discarded and the cell pellets were resuspended in residual media, and then

frozen and thawed three times. Tubes were vortexed before and after each freeze cycle

and following the final thaw, were centrifuged again at 4,500xg for 30 minutes.

Supernatant was then transferred to a 4ml sterile cryotube. A protein assay was

performed to determine the final protein concentration of the antigen. PCR was

performed to confirm the presence of viral DNA.

Positive and Negative Reference Plasma

In July of 2003, three of five captive Burmese star tortoises became ill with

clinical signs consisting of nasal discharge, conjunctivitis and cervical subcutaneous

edema. One of the three tortoises died and histologic and molecular investigations

demonstrated the presence of iridovirus in various tissues. Surviving tortoises were

treated with supportive care and all four tortoises survived. Plasma was collected at the

time of infection during July and then again in September 2003. Plasma from one of

these tortoises was used as the positive control in development of the ELISA. Plasma









was collected from Burmese star tortoises from a zoological collection with no known

history of disease to serve as negative reference plasma for the ELISA.

ELISA Procedure

A checkerboard optimization strategy was used to determine the optimum

concentrations of both antigen and plasma to be used in the ELISA. Antigen

concentrations evaluated were 1:100, 1:250, 1:500, and 1:1000. Plasma concentrations

evaluated were two-fold serial dilutions from 1:50 to 1:1600. The following procedure

was found to be optimal utilizing the crude cell lysate antigen. Each well of a high

protein binding 96 well microplate (Maxisorp F96; Nunc, Kamstrup, Denmark) was

coated with 50[tl of infected or uninfected cell lysate diluted to 1:100 in 0.01M sodium

phosphate buffer (pH 7.2) containing 0.15 M NaCl and 0.02% sodium azide (PBS/Az).

Plates were then incubated overnight at 40C. Antigen was then aspirated off and wells

were washed four times with in ELISA wash buffer (PBS/Az with 0.05% Tween 20).

This washing process was performed between each of the following steps. Wells were

then blocked against non-specific binding with 300[tl of Superblock blocking buffer by

Pierce (Rockford, IL) for one hour at room temperature (RT). Each remaining step was

incubated for Ihr at RT. Plasma samples diluted 1:100 in blocking buffer were added at

50[tl volumes to wells in triplicate. One well was coated with uninfected cell lysate

while the other two wells were coated with infected cell lysate. Next, a biotin-conjugated

monoclonal antibody produced against the desert tortoise IgY light chain and previously

used for detecting anti-mycoplasma antibodies in desert tortoises (Schumacher et al.,

1993) was diluted to a final concentration of 0.5[tg/ml in PBS/Az and added to each well

in 50[tl volumes. Alkaline phosphatase-conjugated streptavidin (Zymed Laboratories,









Inc., San Francisco, California) was then applied to each well at 50[tl of a 1:5000 dilution

in PBS/Az. Next, the ELISA was developed with 100[tl per well of a 1.Omg/ml P-

nitrophenyl phosphate prepared in 0.01M sodium bicarbonate buffer containing 2mM

MgC12 and plates were stored in the dark. The absorbance of each well was read at A405

using a StatFax 3200 microplate reader (Awareness Technology, Palm City, Florida) at

30 minutes.

Each plasma sample was read in triplicate. Plasma was placed on one well

originally coated with uninfected cell lysate and on two wells coated with infected cell

lysate. The average absorbance reading of the two wells coated with infected cell lysate

was calculated and the positive/negative (P/N) ratio value of each sample was determined

by dividing the mean absorbance of the duplicate average by the absorbance reading of

the well from the uninfected cell lysate. This subtracts out any background noise, or non-

specific binding that may have occurred as a result of using the Terrapene heart cell line.

The cut-off value for a positive test result was made by adding three times the standard

deviation of the mean P/N ratio to the mean (Crowther, 2001).

Experimentally Inoculated Turtles

Although iridovirus infection was confirmed in the Burmese star tortoise that

died, it was not confirmed in any of its pen-mates. So while we assumed the other

clinically ill tortoises were exposed, there was no way to definitively know. To develop

known positive and negative samples for use in developing the ELISA and to validate the

test by detection of seroconversion, ten red-eared sliders (Trachemys script elegans)

ranging in weight from 775 to 1050g were obtained from a reptile dealer and allowed to

acclimate in an animal care facility room at the University of Florida for three weeks.









This study was performed under the approval of the Institutional Animal Care and Use

Committee at the University of Florida.

Oral and cloacal swabs were collected from turtles and ran by PCR as previously

described (Mao et al., 1997) for the presence ofRanavirus DNA sequences to determine

current infection status. Plasma was collected and tested by the following ELISA to

determine presence of anti-Ranavirus antibodies at the time of arrival, which would

indicate previous exposure. Turtles were randomly assigned to one of three groups.

Group 1 turtles (Nos. 13, 14, 16 and 20) received Iml of a virus infected crude cell lysate

prepared as described above and diluted to a final concentration of 102 TCID5o/ml orally

(PO) by metal gavage feeding tube placed into the distal esophagus. Group 2 turtles

(Nos. 12, 15, 17, 18) received the same dose of virus intramsucularly (IM), half the dose

in the right and half the dose in the left pectoral muscles. The two other turtles were

assigned to a control route of inoculation. Turtle 11 was mock inoculated with the same

volume of an uninfected cell lysate by PO while turtle 19 received an uninfected cell

lysate by IM. Plasma samples were collected weekly for five months from each turtle to

attempt to detect the production of antibodies. Turtles were euthanized at five months, or

when severe clinical signs of disease appeared including any of the following: severe

lethargy, subcutaneous edema, nasal or ocular discharge, oral plaques, or hyphema.

Tissues were collected at necropsy for PCR, histopathology and virus isolation.

Reproducibility

Intra-assay and inter-assay reproducibility were determined by performing two

precision runs. The positive and negative reference plasma samples, used to optimize the

test, were used in each assay. Intra-assay reproducibility was determined by running the

positive and negative sample multiple times on the same plate. Each sample is run









multiple times on one well of uninfected cell lysate and two wells of infected cell lysate.

This resulted in 64 readings for each sample on the infected cell lysate and 32 on the

uninfected cell lysate. Inter-assay reproducibility was determined by using the values of

the reference plasma results used as controls in running up the wild gopher tortoise

samples from multiple dates and multiple plates. The mean A405, the standard deviation

(SD) and the coefficient of variation (CV) for the intra- and inter-assay reproducibility

were calculated using the optimized ELISA conditions.

Protein Expression and Immunoblotting

The positive control plasma was tested for its ability to detect viral proteins in a

western blot using infected and uninfected TH-1 cell lysates as antigen. Four four-fold

dilutions of infected and uninfected cellular lysate proteins were separated by sodium

dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) under reducing

conditions along with broad range molecular weight markers. The separated proteins

were transferred onto 0.2[im nitrocellulose membranes (Biorad, Hercules, CA) by

standard methods (Harlow and Lane, 1988). Membranes were then rinsed in water for

five minutes in preparation for coomassie blue staining or in tris buffered saline

containing 0.5% Tween20 (TTBS, pH 7.5) for twenty minutes prior to immunoblotting.

Membranes were then stained in coomassie blue stain for 90 minutes, followed by water

for 30 minutes to destain to view differential protein profiles between infected and

uninfected cell lysates. After rinsing in TTBS, membranes for immunoblotting were then

blocked with Superblock blocking buffer in phosphate buffered saline (Pierce) for one

hour. Blocking buffer was then removed and plasma samples diluted 1:2000 in blocking

buffer were added to the membranes. After one hour, the membranes were washed with

TTBS for 30 minutes and the monoclonal antibody was added, diluted 1:10,000 in









blocking buffer for one hour. Again the membranes were washed and AP streptavidin

was added, diluted 1:5,000 in phosphate buffered saline for one hour. Membranes were

washed as previously, and the membranes were developed in substrate buffer (0.1M Tris-

HC1, ImM MgCl2) containing nitroblue tetrazolium chloride (NBT) and 5-bromo-4-

chloro-3-indolylphosphate p-toluidine salt (BCIP) (Biorad). The reaction was stopped by

removing the NBT-BCIP solution and adding deionized water. Membranes were allowed

to air dry.

Wild Gopher Tortoises Samples with Unknown Exposure

Plasma samples from 1000 wild gopher tortoises (Gopheruspolyphemus) from

Florida, Georgia, Alabama, Louisiana and Mississippi were obtained from samples

submitted to the Mycoplasma Research Laboratory at the University of Florida from

2002 to 2006. County data was recorded when available, although it was not available in

68 cases. Samples were further subdivided by region of the state including central,

central east, central west, north central, north east, north west, south east, and south west.

Results were also compared according to state.

Results

Antigen Preparation

Two diffuse bands were observed on the sucrose gradients and each band was

separated into two samples when placed on the sucrose cushion for pelleting. This

resulted in four stocks of purified virus (two low bands 1A and 1B and two higher

bands 2A and 2B). Ten microliters of each stock was submitted for negative staining

electron microscopy (EM). While virus particles were observed (Fig. 3-1), they were

difficult to find by EM. A protein assay of each sample indicated protein concentrations

of 1000 (1A), 375 (1B), 625 (2A) and 500 [tg/ml (2B). Polymerase chain reaction of









each sample resulted in strong positive signals. Use of the purified antigen in an ELISA

indicated approximately 37.5 tg/ml of the purified virus was necessary for coating wells

to obtain the best results. Because such a small volume of purified iridovirus resulted

from the sucrose gradient purification protocol, and such a large volume was needed for

use in the ELISA, it was decided that a crude cell lysate would be used for detecting

seroconversion in the transmission studies and for the seroepidemiology study.

ELISA Parameters

Checkerboard optimization found that diluting the antigen 1:100 and diluting

plasma samples also at 1:100 gave the largest difference between P/N ratios of positive

and negative reference samples (Fig. 3-2). Plasma cut off values were determined after

running the wild gopher tortoise samples. The frequency distribution was determined in

tenth increments from 0.5 to 4.1 and the P/N ratios of the samples were determined to

have a normal distribution, skewed to the right (Fig. 3-3). Because the data was normally

distributed, the mean P/N ratio of samples (1.078), plus three times the standard deviation

(0.379) was used to determine the positive cut-off value (2.2). P/N ratios from each

sample were plotted on a graph with the sample numbers on the x-axis in increasing order

and the P/N ratio values on the y-axis (Fig. 3-4). This plot shows a gradual increase in

P/N ratios followed by a sharp increase around a P/N ratio of about 2.0, confirming that

our cut off value of 2.2 to be a reasonable value.

Experimentally Inoculated Turtles

One IM inoculated turtle (No. 15) became extremely lethargic and died 24 days

post-inoculation. Cloacal swabs taken on turtle 15 were positive by PCR and sequencing

for the presence of iridovirus starting 8 days before the turtle died. Oral swabs were

positive the day before the turtle died. Other than lethargy, no clinical signs were









observed in this turtle such as stomatitis, conjunctivitis or nasal discharge. Histologic

changes were consistent with those seen in naturally and experimentally inoculated

turtles (DeVoe et al., Johnson et al., unpublished data). Briefly, there was a fibrinoid

vasculitis in the spleen with multifocal infiltrates of low numbers of heterophils and

scattered free brown pigment granules (presumptively from disrupted melanomacrophage

centers). There were occasional lumenal fibrin thrombi with admixed heterophils and

karyorrhectic debris. No other turtles died during the five-month course of the study.

Plasma samples collected weekly over the five-month period failed to detect

seroconversion in all of the mock inoculated (Fig. 3-5A) turtles and turtles orally

inoculated with Ranavirus (Fig. 3-5B). One IM inoculated turtle (No. 12) had an

increasing P/N ratio trend, but only twice did values exceed a ratio of 2, which occurred

on weeks 8 and 18 (Fig. 3-5C). All other turtles remained well below the positive P/N

ratio cut off value of 2.2.

Reproducibility

The mean A405, SD and CV values for the intra- and inter-assay precision runs are

shown in Table 1. Ideally, CV values should be less than 15% (Crowther, 2001). Three

of the CV values in the precision runs are >15% indicating that some variability still

exists in this assay.

Protein Expression and Immunoblotting

Coomassie blue staining of proteins from infected cell lysates and uninfected cell

lysates showed a different pattern of protein expression (Fig. 3-6). Immunoblotting of

infected and uninfected cell lysate showed a marked increase in binding to proteins in the

infected cell lysate and very weak binding to proteins in the uninfected cell lysate (Fig. 3-

7). Strong signals were seen on virus-infected cells at approximately 125 kDa, and 78









kDa with weaker signals seen at 70, 65, and 28 kDa. Very faint signals were also seen at

125 and 78 kDa in the uninfected cells.

Wild Gopher Tortoises Samples with Unknown Exposure

Of 1000 gopher tortoise plasma samples assayed, 15 (1.5%) were positive with a

P/N ratio >2.2 (Table 2). Eight seropositive tortoises came from five counties in Florida

including Lake, St. Lucie, Broward, Palm Beach and Martin (Fig. 3-8). While

seropositive tortoises represent three regions including central, central east, and

southeast, four of the five counties are clustered closely together in the south including

Palm Beach, Broward, Martin, and St. Lucie. The remaining seven seropositive tortoises

were located in Baker, Georgia, a county in the southwest corner of the state. Prevalence

of seropositive tortoises by county was quite variable (Table 3). Approximately 3% of

tortoises sampled in Lake county were positive (n=99), 2.9% in St. Lucie (n=35), 10% in

Broward (n=10), 3.4% in Palm Beach (n=58), and 6.2% in Martin county (n=16). Of

tortoises tested from Baker, GA 6.5% were positive (n=113). By state, Florida had an

overall prevalence of 1.2% (n=658) while Georgia had a prevalence of 3.1% (n=225)

(Table 3). All tortoises tested from Alabama, Mississippi and Louisiana were negative.

Discussion

Iridoviruses of the genus Ranavirus are emerging as important pathogens of

chelonians. Previous reports have established their presence in Florida (Westhouse et al.,

1996) and in North Carolina (DeVoe et al., 2004). Newer unpublished findings indicate

a much larger range of infection including reports from Texas, Georgia, New York and

Pennsylvania (Johnson et al., 2004). The duration of illness can be rather short (Johnson

et al., unpublished findings), making it difficult to observe symptomatic tortoises in wild

populations. Therefore, we developed an indirect ELISA to detect anti-Ranavirus









antibodies and performed a cross-sectional survey of gopher tortoises utilizing banked

plasma samples to attempt to determine the prevalence of exposure of free-ranging

chelonians to iridoviruses.

When developing a new assay, it is ideal to have another assay against which to

compare results. With a reference assay, the sensitivity and specificity of the new assay

can be evaluated, and thus the level of confidence one has in its ability to detect what you

want it to detect. Unfortunately, this was not possible with this pathogen. Detectable

neutralizing antibodies are not usually found in naturally or experimentally infected

animals (reviewed by Whittington et al., 1997) and thus, serum neutralization tests,

which are sometimes used as gold standards for verifying ELISA results, cannot

confidently be used to determine exposure to this virus. For this reason we decided to

perform experimental transmission studies using red-ear sliders to attempt to demonstrate

the validity of our assay by detecting seroconversion. Our assay failed to detect

antibodies in all but one slider. However, recent reports show that multiple exposures are

needed in amphibians to detect IgY antibodies against FV3 using an ELISA (Gantress et

al., 2003; Maniero et al., 2006) and IgM antibodies were not detectable even after

multiple exposures. Only one dose of virus was administered to experimentally

inoculated turtles, which might explain why the turtles in this study failed to mount an

immune response. Our results did show a good correlation between antibody production

following a clinical disease in a Burmese star tortoise that was previously housed with a

tortoise that died with confirmed Ranavirus infection. Plasma from a Burmese star

tortoise from another facility with no known history of illness was selected as a

seronegative control; antibodies to Ranavirus were not found in this tortoise.









Additionally, using a western blot, we were able to show that plasma from the positive

control turtle bound to proteins in the infected cells in the immunoblot but not in the

uninfected cells indicating that the plasma was reacting with viral proteins and not

cellular proteins from the lysate we were using to coat the plates. For these reasons, we

assume our test to be valid for use in detecting anti-iridovirus antibodies in chelonians,

however further validation of this assay is warranted.

Only one turtle died as a result of IM inoculation of virus at 102TCID50. A

previous study showed that 75% (n=4) of turtles inoculated with a higher dose of virus

(105TCID0o) died as a direct result of iridovirus infections (Johnson et al., unpublished

data). The viral dose was extrapolated from experimental studies with fish and

amphibians (Langdon, 1989, Moody and Owens, 1994, Bollinger, 1999, Cullen and

Owens, 2002), so it was unknown what a sub-lethal dose would be in chelonians. While

studies with a larger sample size might provide more accurate correlations between viral

load and mortality rates, it appears that host characteristics likely play a significant role in

resistance or susceptibility to disease. Environmental temperatures have also been shown

to significantly impact the percent mortality and time to death in salamanders

experimentally inoculated with a Ranavirus (Rojas et al., 2005), where salamanders

inoculated at 18 and 100C were more likely to die than those exposed at 260C.

Underlying disease conditions were not noted at necropsy or on histologic review of

tissues in the turtle that died that might have contributed to an increased susceptibility.

Prevalence among free-ranging gopher tortoises was found to be low, only 1.5% of

1000 samples being positive. This could be the true prevalence rate, although we suspect

that this is an underestimate of the true rate. There are several factors that could cause









the prevalence to be underestimated. Prevalence is a function of the incidence of disease

multiplied by the average duration of the illness. While incidence would be difficult to

determine in a natural setting, duration of illness can be extrapolated from experimental

studies. If chelonians die quickly as a result of infection, they will not have time to

mount an immune response to the pathogen, and will not survive to be surveyed. As

previously mentioned, experimental transmission studies have shown a high rate of

mortality (75%) in turtles intramuscularly inoculated with 105TCID50 of a Ranavirus

infected cell lysate (Johnson et al., unpublished data). Turtles all died within 30 days of

exposure to the virus. Although, the route of transmission is unknown in a natural

environment, if naturally exposed cases experience similar mortality rates and duration

from exposure to death, a cross-sectional study evaluating the prevalence of exposure

will miss many tortoises that were exposed, because the majority of them will die. This

was demonstrated in a natural setting in Pennsylvania. A population of approximately 70

eastern box turtles (Terrapene carolina carolina) was being repatriated in a nature

sanctuary. Turtles were tracked every 3-5 days by radiotelemetry. In the summer and

fall of 2003, 15 of these turtles died suddenly, with what was later identified to be

iridovirus infections (Johnson et al., 2004). The following spring, the remaining 55

turtles were sampled, and plasma was run for the presence of antibodies. Only three

turtles were positive on ELISA (data not shown). If we calculated this value as the true

prevalence, we would estimate that -5.5% of turtles were exposed, when we know that

21% of the population died that was not included in the estimate. This severely

underestimates the prevalence of disease. However, we cannot extrapolate these

differences to other populations, as this was a repatriated population, and thus, subject to









stressors that might not be found in wild populations, making them more susceptible to

disease. Secondly, this study suggests that turtles must be exposed more than once to

mount an immune response similar to what was found in studies with Xenopus (Gantress

et al., 2003; Maniero et al., 2006). Thus, if turtles have only been exposed once, we will

not detect that first exposure with our ELISA, also underestimating the true prevalence of

exposure.

Emerging infectious diseases have been increasingly recognized as factors

influencing wildlife health and populations (Harvell et al., 1999; Daszak et al., 2000).

Although mycoplasmosis has been postulated to contribute to declines of some tortoise

species (USFWS, 1994), the causes) of mass mortality events in wild chelonian

populations often remain undetermined (Flanagan, 2000; Dodd, 2001). Iridovirus

infections in chelonians can have a high mortality rate, but the duration of illness is short,

making it difficult to observe disease outbreaks in the wild. It is possible that it might

contribute significantly to mortality rates in wild populations of chelonians. Utilizing a

serological assay may help to determine regions where iridovirus infections might be

more prevalent in chelonian populations. These locations could then be monitored more

closely for disease outbreaks in both chelonians and amphibians.

Results of the serosurvey showed that counties in the central and southeastern

region of Florida were more likely to have seropositive tortoises. Interestingly, four of

these counties had adjoining borders (St. Lucie, Martin, Palm Beach and Broward),

suggesting that either tortoises in this area are at higher risk of exposure to iridoviruses,

or alternatively, iridovirus is more endemic in this area, so lower levels of exposure allow

more tortoises to seroconvert without succumbing to the disease. Direct comparisons of









prevalence rates between counties are difficult to make as this was a convenience sample

and geographic area or tortoise densities per county data were not controlled for in

sampling. Only 32 of 67 counties were sampled. The northwestern region was not well

sampled, and inferences about this region can't be made. However, seven tortoises in

Baker County, Georgia were seropositive and this county is located in the southwestern

portion of the state, which would be near the northwestern portion of Florida.

Earlier studies have shown that amphibians might be a source of iridovirus

infections in chelonians (Johnson et al., unpublished data). A moribund leopard frog was

found and euthanized at the same site in Georgia where the Burmese star tortoises

became ill and one died. Restriction enzyme analysis of viral genomic DNA from an

isolate obtained from the tortoise and the frog demonstrated identical restriction patterns,

suggesting they are the same or very closely related viruses. Thus, conditions that might

propagate amphibian iridovirus infections would likely cause an increased chance of

exposure in chelonians. Green et al., (2002) found that increased precipitation and

population densities were directly associated with increased die-offs of amphibians.

Thus, these similar settings might create higher rates of exposure in chelonians.

Additionally, it has been shown that sublethally infected amphibians can cause sporadic,

recurrent disease outbreaks in amphibians (Brunner et al., 2004). Experimentally and

naturally infected tiger salamander larvae and metamorphs were able to maintain

sublethal, transmissible infections for over five months. Apparently healthy infected

dispersing metamorphs were returning to water bodies to breed and it was speculated that

these individuals were likely serving as a reservoir host for infecting newly hatched

larvae, creating recurrent outbreaks of disease. It is unknown whether chelonians are









capable of sustaining sublethal infections or capable of spreading disease to naive

populations. One turtle experimentally inoculated intramuscularly with iridovirus

remained clinically healthy but continued to shed virus from the cloaca detectable by

PCR up until 30 days. The study ended at 30 days so it is unknown how long this turtle

would have kept shedding, or if the virus being shed was still infectious. Further studies

would be useful to determine the risk posed to other chelonian and amphibian

populations of iridovirus infected turtles that survive the initial infection. This assay

could be useful for managing populations in wild and captive settings by identifying

tortoises who might be asymptomatic carriers.

In summary, this study reports the development of an indirect ELISA for detection

of anti-iridovirus antibodies in chelonians. It was able to detect antibodies in a naturally

infected Burmese star tortoise whose pen-mate died with a confirmed iridovirus infection.

A seroprevalence survey of banked plasma samples from free-ranging gopher tortoises in

Florida, Georgia, Alabama, Louisiana and Mississippi found a 1.5% prevalence rate of

exposure. Further studies are needed to characterize the true incidence of disease in wild

populations of chelonians.









Table 3-1. Reproducibility of the ELISA. SD = standard deviation of the mean A405.
CV = coefficient of variance expressed as a percent. ICL = values from
samples run on wells coated with infected cell lysate. UCL = values from
samples run on wells coated with uninfected cell lysate.


Positive Sample Negative Sample
n Mean A405 SD CV n Mean A405 SD CV
Intra-assay ICL 64 0.363 0.022 6.06 64 0.107 0.011 10.28
UCL 32 0.094 0.009 9.57 32 0.112 0.009 8.04
Inter-assay ICL 26 0.366 0.079 21.58 26 0.130 0.023 17.69
UCL 13 0.095 0.011 11.58 13 0.111 0.017 15.31


Table 3-2. ELISA results of 1000 free-ranging gopher tortoise (Gopheruspolyphemus)
plasma samples by county and state.



State County Number tested Positive Percentage


Baldwin
Mobile
Alachua
Brevard
Broward
Citrus
Clay
Collier
Columbia
Hernando
Hillsborough
Indian River
Lake
Lee
Leon
Madison
Manatee
Marion
Martin
Miami-Dade
Nassau
Orange
Osceola
Palm Beach


0
0
0
0
10.0
0
0
0
0
0
0
0
3.0
0
0
0
0
0
4.0
0
0
0
0
3.4










FL Pasco 26 0 0
FL Pinellas 2 0 0
FL Polk 6 0 0
FL Sarasota 1 0 0
FL Seminole 54 0 0
FL St. Johns 7 0 0
FL St. Lucie 35 1 2.9
FL Taylor 1 0 0
FL Volusia 26 0 0
FL Walton 2 0 0
GA Baker 113 7 6.2
GA Liberty 74 0 0
GA Tattnall 38 0 0
LA Washington Parish 12 0 0
MS Greene 7 0 0
MS Harrison 16 0 0
MS Perry 5 0 0
Unknown 68 0 0


TOTAL 1000 15 1.5


TOTAL


1000


15 1.5












Table 3-3. ELISA results of 658 free-ranging gopher tortoises (Gopheruspolyphemus)
from the state of Florida are listed by region. C = central, CE = centraleast,
CW = centralwest, NC= northcentral, NE = northeast, NW = northwest, SE =
southeast and SW = southwest.


Region Total No. Tested No. Positive Percent
C 232 3 1.3
CE 90 1 1.1
CW 126 0 0.0
NC 32 0 0.0
NE 55 0 0
NW 2 0 0
SE 101 4 4.0
SW 20 0 0



Table 3-4. ELISA results of 1000 free-ranging gopher tortoises (Gopheruspolyphemus)
listed by state.

State Total No. Tested Total No. Positve Percent
Alabama 9 0 0
Florida 658 8 1.2
Georgia 225 7 3.1
Louisiana 12 0 0

Mississippi 28 0 0



















.;-s





















Fig. 3-1. Negative staining electron photomicrograph of an iridovirus particle purified
by sucrose gradient ultracentrifugation.







57



ELISA Optimizatior


S--*Neg Control
- Pos Control


50 100 200 400 800 1600
Reciprocal of Plasma Dilutions


Fig. 3-2. Optimization of the ELISA with antigen coated at 1:100 dilution, comparing
the positive to negative (P/N) ratio of two fold serial plasma dilutions of the
positive control turtle (Burmese star tortoise with clinical signs of illness)
versus a negative control (Burmese star tortoise with no history of illness).
Plasma diluted at 1:100 showed the greatest difference between the positive
and negative control.












P/N Ratio Frequency Distribution


o (O ^ C& 0 '-, ^ V O N ^ 0& '-, '^ ^ V ^o ON ^ C 0 '-N 'N r N rO r ; 0

P/N Ratios


Fig. 3-3. Frequency distribution of P/N ratios from an indirect ELISA performed on
1000 free ranging gopher tortoise (Gopheruspolyphemus) plasma samples.
Samples show an approximately normal distribution, skewed to the right.







59



ELISA Results


Tortoises


Fig. 3-4. Individual P/N ratio values for 1000 free-ranging gopher tortoises (Gopherus
polyphemus) in increasing value.




































Pre 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20
Weeks post-inoculation


Ne ) <0Ab5Weeks post-inoculation
Weeks post-inoculation


~/c~7A


-- RES19
-U-RES11


-*-RES13
--RES14
-RES16
--RES20



















1%V


-- RES12
-m-RES15
-RES17
--RES18


Weeks post-inoculation
Weeks post-inoculation


Fig. 3-5. P/N ratios of red-eared slider (Trachemys script elegans) plasma samples
collected weekly over five months. A) P/N ratios of mock-inoculated turtles.
B) P/N ratios of turtles orally inoculated with Ranavirus. C) P/N ratios of
turtles intramuscularly inoculated with Ranavirus.










Infected Cell Lysate
SA


Uninfected Cell Lysate
A 7 A


Fig. 3-6. Coomassie blue staining of a SDS-PAGE gel separating proteins of iridovirus-
infected and uninfected Terrapene heart cell lysates. Lane 1 is a broad range,
pre-stained molecular weight marker with weight in kDa marked next to the
lane. Lanes 2-5 are four fold serial dilutions of iridovirus infected cell lysate.
Lanes 6-8 are four fold serial dilutions of uninfected cell lysate.


MW in KDa


34.8
28.9

20.6
7.1








345


MW in
KDa


Fig. 3-7. Western immunoblot. Lanes coated with four fold serial dilutions of
iridovirus infected cell lysate in lanes 2-5 and uninfected cell lysate in wells 6-
9. Plasma from the positive control was used as the primary antibody for
detection of iridovirus specific antibody binding. Strong signal was seen on
virus-infected cells at approximately 125 kDa, and 78 kDa with weaker
signals seen at 70, 65, and 28 kDa. Very faint signals were also seen at 125
and 78 kDa in the uninfected cells.


6 7 8 9








Uninf6789ed








Uninfected Cells


209


124
80


49.1

34.8
28.9

20.6
7.1


Infected Cells




































Fig. 3-8. County map of Florida. The five counties highlighted indicate where
seropositive tortoise samples were identified.














CHAPTER 4
EXPERIMENTAL TRANSMISSION OF A RANA VIRUS IN WESTERN ORNATE
BOX TURTLES (Terrapene ornata ornata) AND RED-EARED SLIDERS (Trachemys
script elegans)

Introduction

Viruses in the family Iridoviridae are large double stranded DNA viruses capable of

infecting ectothermic vertebrates, and invertebrates (Williams et al., 2005). Iridoviruses

within the genus Ranavirus have been shown capable of infecting fish, amphibians and

reptiles (Chinchar, 2002) and have emerged as major pathogens of free-ranging

amphibians worldwide (Cunningham et al., 1996, Zupanovic et al., 1998, Daszak et al.,

1999). In a study of sixty-four amphibian morbidity and mortality events between 1996

and 2001, the most common cause of mortality events was infection with iridoviruses

(Green et al., 2002).

Iridovirus infections in reptiles have been less well described than in amphibians

and fish, with sporadic reports in chelonians, snakes, and lizards (Just et al., 2001, Hyatt

et al., 2002, Marschang et al., 2005). Infection of a tortoise was first reported in 1982 in

a spur-tailed Mediterranean land tortoise (Testudo hermanni) that had necrotic foci in the

liver, intestine and spleen (Heldstab and Bestetti, 1982). Subsequently, iridoviruses were

reported in other species of chelonians both in captivity and in the wild (Westhouse et al.,

1996, Chen et al., 1999, Marschang et al., 1999, DeVoe et al., 2004). Clinical signs

associated with infections have included signs of upper respiratory tract disease including

respiratory distress and nasal discharge, as well as oral ulceration, cutaneous

abscessation, anorexia and lethargy (Westhouse et al., 1996, DeVoe et al., 2004). While









there is circumstantial evidence that iridovirus is a primary pathogen in chelonians,

Koch's postulates have never been fulfilled.

Ranavirus was recently identified in tissues obtained from a variety of wild and

captive chelonians that died in Georgia, Florida, Texas, New York and Pennsylvania

(Johnson et al., 2004). Species affected were Burmese star tortoises (Geochelone

platynota), gopher tortoises (Gopherus polyphemus), eastern box turtles (Terrapene

carolina carolina), and Florida box turtles (Terrapene carolina bauri). Clinical signs in

these cases included lethargy, anorexia, nasal discharge, conjunctivitis, severe

subcutaneous cervical edema, and necrotizing pharyngitis-stomatitis. One of the

ranaviruses isolated from a Burmese star tortoise (Geochelone platynota) that died in an

outdoor pen in a zoological collection from Georgia in 2003 was further characterized as

either identical or closely related to Frog Virus 3 (Johnson et al., 2004). To determine a

causal relationship between the isolated Ranavirus and the clinical and histologic lesions

observed in these chelonians and to fulfill Koch's postulates, we performed two

experimental transmission studies using the previously characterized Burmese star

tortoise isolate that will hereafter be termed Burmese star tortoise Ranavirus (BSTRV).

The first was a pilot study to determine the suitability of either western ornate box turtles

(Terrapene ornata ornata) or red-eared sliders (Trachemys script elegans) as an

experimental model for a subsequent larger transmission study. Here we report the

findings of those studies.

Materials and Methods

Experimental Animals and Husbandry

Study 1 was a pilot study consisting of three western ornate box turtles (BT;

Terrapene ornata ornata) and three red-eared sliders (RES; Trachemys script elegans)









that were purchased from a reptile supplier in Texas. Study 2 consisted often RES

purchased from a reptile supplier in Louisiana. Both studies were performed under the

approval of the Institutional Animal Care and Use Committee at the University of

Florida. Turtles were housed individually in plastic containers in a centralized animal

facility room maintained at approximately 25C. Appropriate husbandry for each species

was used that included commercially available foods. Box turtles were kept on land, with

overhead heat lamps provided for basking at one end of the container (average

temperature of 28 C). Red-eared sliders were kept in water (temperatures averaging

between 21.3 and 25.60C), with basking platforms provided under an overhead heat lamp

(average basking temperature of 280C). Fluorescent room lights were kept on a 12-hour

light and 12-hour dark cycle.

Pre-inoculation Sample Collection

Upon arrival, each turtle was examined for the presence of any clinical signs of

illness. Urine, oral and cloacal swabs were obtained from each turtle and tested for the

presence of iridovirus utilizing a polymerase chain reaction (PCR) test (see below).

Blood samples were collected from each turtle into 2ml tubes coated with lithium heparin

and centrifuged at 4,500 x g for 5 min. Buffy coats were removed and also tested for

iridovirus using PCR. Plasma was removed and tested for anti-Ranavirus antibodies

using an indirect enzyme linked immunosorbent assay (ELISA; see below).

DNA Preparation, Polymerase Chain Reaction and Nucleotide Sequencing

Oral and cloacal swabs were combined for each turtle into one 1.5ml

microcentrifuge tube and 100tl of phosphate buffered saline was added. DNA was

extracted from buffy coats, oral and cloacal swabs using the DNeasy kit (Qiagen,

Valencia, CA, USA) as were tissue samples later collected at necropsy. Viral DNA from









urine samples was extracted using the QIAmp Ultrasens virus kit (Qiagen). Sense primer

(5'-GACTTGGCCACTTATCAC-3') and anti-sense primer (5'-

GTCTCTGGAGAAGAAGAA-3'), were used as previously described (Mao et al., 1997)

to amplify approximately 500 basepairs of the Ranavirus major capsid protein gene. A

20[tl reaction mixture was run which contained 2[tl extracted DNA, 1 [tM of both

primers, 200 [tM each of dATP, dCTP, dGTP, and dTTP, 1.5 U of Taq DNA polymerase

and PCR buffer containing 50 mM KC1, 10 mM Tris-HC1, 1.5 mM MgC12

(Eppendorf, Westbury, New York). The mixtures were amplified in a thermal cycler

(PCR Sprint, Thermo Hybaid) with an initial denaturation at 94C for 2.5 min, followed

by 25 cycles of denaturation at 94C for 30 sec.; annealing at 50C for 30 sec, extension

at 72C for 30 sec., and a final extension step at 72C for 10 min as previously described

(Marschang et al., 1999). The PCR products were resolved in 1% agarose gels and any

bands were excised and purified using the QIAquick gel extraction kit (Qiagen).

Products were sequenced in both directions directly using the Big-Dye Terminator Kit

(Perkin-Elmer, Branchburg, New Jersey) and analyzed on ABI 377 automated DNA

sequencers at the University of Florida's Sequencing Center. The sequences were

compared to known sequences in GenBank (National Center for Biotechnology

Information, Bethesda, Maryland), EMBL (Cambridge, United Kingdom), and Data Bank

of Japan (Mishima, Shiuoka, Japan) databases using TBLASTX (Altschul et al., 1997).

ELISA

An indirect enzyme linked immunosorbent assay (ELISA) was used to determine

the presence of anti-Ranavirus antibodies. The ELISA methodology was similar to that

developed for use in identifying the presence of anti-tortoise herpesvirus antibodies in









tortoises (Origgi et al., 2004) and anti-west Nile virus antibodies in alligators (Jacobson

et al., 2005). The BSTRV isolate was used as the antigen in the assay. Each well of a 96

well high-protein binding microplate was coated overnight at 4 oC with 50 [tl of a 1:400

dilution of either an uninfected lysate from Terrapene heart cells (TH-1, ATCC-CCL 50,

American Type Culture Collection, Rockville, MD) or TH-1 cell lysate from cells

infected with BSTRV. Lysates were diluted in 0.01 M sodium phosphate buffer (pH7.2)

containing 0.15 NaCl and 0.02% NaN3 (PBS/A). Wells were then washed four times in

ELISA wash buffer (PBS/A with 0.05% Tween-20). This washing process was repeated

in between all of the following steps. Wells were blocked with 300 [tl of Superblock

blocking buffer (Pierce) for one hour at room temperature. All remaining steps were

incubated for one hour at room temperature. Plasma samples were added in 50[tl

volumes at a 1:100 dilution in blocking buffer. The secondary antibody used was a

biotin-conjugated mouse anti-tortoise immunoglobulin (Ig) monoclonal antibody diluted

to a final concentration of 0.5[tg/ml in blocking buffer. Alkaline phosphatase-

conjugated-streptavidin (Zymed Laboratories, Inc., San Francisco, CA) was then applied

to each well at 50 iL of a 1:5000 dilution in PBS/A. Next, 100 pl of a 1.0 mg/ml P-

nitrophenyl phosphate prepared in 0.01 M sodium bicarbonate buffer containing 2 mM

MgC12 was added to each well and the plates were then stored in the dark until being

read. The optical density (OD) of each well was read at A405 using a StatFax 3200

microplate reader (Awareness Technology, Palm City, Florida, USA) after 30 minutes.

Each sample was done in triplicate: one time on wells coated with uninfected cell

lysate and in duplicate on wells initially coated with infected cell lysate. The replicate

values of the wells coated with BSTRV lysate were averaged and divided by the OD









reading of the value of the plasma sample run on the uninfected cell lysate to subtract out

any background binding that might be caused by cross-reactivity to the cells. Values

greater than 2 were considered positive (Jacobson et al., 2005), suggesting previous

exposure to the virus, and would preclude use of that turtle in the study. Plasma from a

surviving penmate of the above Burmese star tortoise was used as a positive control.

Virus Preparation

Terrapene heart cells (TH-1) were acquired from the American Type Culture

Collection (ATCC-CCL 50; Rockville, MD) and grown to confluency in 225cm2 tissue

flasks (Costar, Coming, NY). Cells were cultured in Dulbecco's modified Eagle medium

(DMEM, Gibco, Carlsbad, CA) supplemented with 5% fetal bovine serum (Gibco),

gentamicin (60mg/liter; Sigma, St. Louis, MO), penicillin G (120,000 U/liter),

streptomycin (120,000 U/liter) and amphotericin B (300tg/liter; Sigma). Cells were

inoculated with a fourth passage of BSTRV and incubated at 280C in the presence of 5%

CO2. When cytopathic effect (CPE) was observed, consisting of cell rounding and

detachment from the flask in over 70% of cells, the flasks were scraped and contents

transferred to 15 ml centrifuge tubes and clarified by slow speed centrifugation at

4,500xg for 30 minutes. The supernatant was then discarded and the cell pellet

resuspended in 10mls of cell culture media. The preparation was then vortexed, frozen

and thawed three times to release virus from the cells into the supernatant. The

preparation was again clarified by centrifuging for thirty minutes. The supernatant was

then transferred to a new tube and the cell pellet was discarded. The live virus in the

media was then quantified by a TCID5o assay, diluted with media to create a

concentration of 105TCID50/ml, and frozen at -800C until use.









Transmission Studies

Study 1

Study 1 was designed as a pilot study to determine the suitability of either red-

eared sliders (RES) or western ornate box turtles (BT) as a model ofRanavirus infection

for chelonians. Three RES and three BT were included in the study and each turtle was

allowed to acclimate for two weeks prior to infection. One of each species was assigned

to one of three groups: 1) a mock inoculated control group, with both turtles receiving

0.5ml uninfected cell lysate orally and 0.5ml by intramuscular injection, 2) an orally (PO)

inoculated group with both turtles receiving lml of infected cell lysate containing virus at

105TCID50/ml by metal gavage feeding tube in the caudal esophagus and 3) an

intramuscularly (IM) inoculated group, with both turtles receiving the same concentration

of virus as the PO inoculated group with 0.5ml injected into both the left and right

pectoral muscles. Turtles were observed daily after inoculation for the duration of the

study. Oral swabs and buffy coats were collected 1-week post inoculation for evaluation

by PCR for the presence of iridovirus. DNA extraction and PCR were performed as

described previously for prescreening. Physical examinations were performed daily to

assess the presence of the following clinical signs: lethargy, anorexia, cervical edema,

palpebral or periocular edema, ocular discharge, nasal discharge, oral discharge, the

presence of oral plaques or any other abnormalities.

Turtles were euthanized if clinical signs of disease became severe or at two weeks

post-inoculation. For euthanasia, ketamine was administered intramuscularly at 100

mg/kg followed by intravenous sodium pentobarbital. Once turtles were unresponsive to

painful stimuli and showed no corneal reflex, they were decapitated and a complete

necropsy performed. Portions of tongue, esophagus, stomach, small and large intestine,









liver, kidney, and spleen were collected aseptically by flame sterilizing tools between

each organ, and frozen at -800C for virus isolation and/or DNA extraction and PCR. The

following tissues were collected and fixed in 10% neutral phosphate buffered formalin:

tongue, esophagus, stomach, small and large intestine, liver, kidney, spleen, pancreas,

heart, trachea, lung, brain, urinary bladder, thyroid gland, adrenal gland, bone, bone

marrow, skin, skeletal muscle, nasal cavity, eye and gonad. These tissues were then

processed for histologic examination. They were embedded into paraffin, and 6 jm

sections were stained with hematoxylin and eosin.

Virus isolation was performed on Terrapene heart cells seeded into 25 cm2 flasks

(Costar, Coming, NY). Cells were cultured in Dulbecco's modified Eagle medium/F 12

(Invitrogen, Carlsbad, CA) supplemented with 5% fetal bovine serum (Gibco, Carlsbad,

CA, USA), gentamicin (60 mg/liter) (Sigma, St. Louis, MO, USA), penicillin G (120,000

U/liter), streptomycin (120,000 U/liter) and amphotericin B (300[tg/liter) (Sigma) and

cultured to confluency. A small piece of kidney (approximately 50mg) collected

aseptically at necropsy was homogenized in separate 5ml tissue grinders containing

DMEM. The homogenate was passed through a 0.45[tm filter (Costar) onto a flask of

cells. Cells were incubated at 280C and observed daily. Flasks of cells were harvested

when CPE was observed in over 70% of cells or at post-inoculation day 10. DNA was

extracted from cells using the DNeasy kit (Qiagen) protocol for animal cells. Polymerase

chain reaction and nucleotide sequencing was performed as previously described to

confirm the presence ofiridovirus DNA sequences.









Study 2

Based on the results of the pilot study, a second transmission study was designed

that consisted of ten RES. Turtles were assigned to one of the following three groups: 1)

control; 2) IM inoculated or 3) PO inoculated. Turtles were numbered from one to ten

according to increasing weight. They were then blocked into two groups (1-5 and 6-10)

and turtles within each block were randomized to 1 control, 2 PO inoculated (turtle Nos.

1, 2, 7 and 10) and 2 IM inoculated (turtle Nos. 3, 5, 6 and 8). The control turtle in the

lower weight block (turtle No. 4) was PO mock inoculated with lml of uninfected cell

lysate, while the control turtle in the higher weight block (turtle No. 9) received Iml IM,

half in each pectoral muscle. Turtles were monitored daily for the presence of clinical

signs consistent with iridovirus infection. Oral and cloacal swabs were taken three times

a week throughout the four-week study. Free catch urine samples were collected

opportunistically at the same time periods. Turtles were euthanized when clinical signs of

infection became severe or four weeks post-inoculation.

Necropsies, tissue collections and virus isolation were performed as described in

Study 1. In addition, spleen from one IM inoculated (No. 6) and one control (No. 9)

turtle were collected in Trump's solution (4% formaldehyde, 1% glutaraldehyde in a

phosphate buffer; Electron Microscopy Sciences, Hatfield, PA) for transmission electron

microscopy (TEM). Tissues were embedded in Spurr's resin and ultrathin sections were

obtained and stained with lead citrate and uranyl acetate for TEM at the Electron

Microscopy Core Laboratory, University of Florida.









Results

Experimental Animals and Pre-inoculation Sampling

Upon arrival, all turtles were considered healthy. Using PCR, all oral swabs,

cloacal swabs, urine and buffy coats collected from turtles in Studies 1 and 2 were

negative for iridovirus DNA sequences. All plasma samples evaluated by ELISA were

below the positive cutoff value, indicating that the turtles were seronegative for exposure

to Ranavirus.

Transmission Studies

Study 1

Control mock inoculated and PO inoculated red-eared sliders and box turtles

showed no clinical signs of disease throughout the two-week study. The IM inoculated

RES and BT both showed severe clinical signs. The RES died 9 days post inoculation

(DPI). At 8 DPI the IM inoculated RES was basking continuously and started showing

signs of lethargy, cutaneous erythema, and kept its palpebrae closed. It was found dead

on the basking platform on the morning of 9 DPI. The orally inoculated RES remained

normal. At 8 DPI, the IM inoculated BT developed a white opaque ocular discharge (Fig.

4-1), at ten days became lethargic and anorexic, and at 12 DPI, was euthanized. The

orally inoculated BT remained normal.

Both IM inoculated turtles in the pilot study were positive by PCR for iridovirus

sequences using DNA extracted from oral swabs and buffy coats collected one week

post-inoculation. Oral swabs and buffy coats collected on the PO inoculated turtles and

mock-inoculated control turtles were negative. PCR performed on DNA extracted from

both IM inoculated turtles were positive for all eight tissues (Table 1). PCR was negative

for all eight tissues of both control turtles and both PO inoculated turtles.









The most consistent and significant histologic lesions were observed only in the

spleens (2/2) of IM inoculated BT and RES, both having similar changes. Changes were

based on the normal histology of the spleen that was previously defined for red-eared

sliders (Kroese and Van Rooijen, 1982). Lesions were centered on the splenic ellipsoids

or "sheathed capillaries" and will be described in detail for turtles in Study 2 (below).

Briefly, the walls of the ellipsoids were moderately to markedly expanded by

homogenous to slightly fibrillar eosinophilic material (fibrin; fibrinoid vasculitis) with

multifocal infiltrates of low numbers of heterophils. There were occasional lumenal

fibrin thrombi with admixed heterophils and karyorrhectic debris.

Kidney samples from both IM inoculated turtles (P3 and P6) that were coated onto

TH-1 cells demonstrated cytopathic effects of cell rounding and lysis within two days of

incubation. Intracytoplasmic inclusion bodies were observed in infected cells and PCR

and nucleotide sequencing on DNA extracted from cells from each flask were positive for

iridovirus. None of the cultures inoculated with tissues collected from PO inoculated

turtles, or control turtles demonstrated any CPE up to ten days post-inoculation of cells

and each were negative by PCR for the presence of Ranavirus DNA segments.

Study 2

Similar to Study 1, only IM inoculated RES showed clinical signs of disease and

were euthanized before the end of the four-week study. Three of the four IM inoculated

turtles showing severe clinical signs were euthanized on days 11, 13 and 23 DPI. All

three turtles became anorectic, and extremely lethargic. The turtle euthanized 13 DPI

developed oral plaques on the roof of the mouth and tip of the tongue (Fig. 4-2). Turtle

3, euthanized 23 DPI, exhibited exophthalmus, conjunctivitis and hyphema (Fig. 4-3).

All three turtles had clear serious ocular and nasal discharge. The fourth IM inoculated









turtle (No. 8) showed ocular discharge and subjectively basked more than the other turtles

between 16 and 25 DPI, but then recovered and showed no clinical signs at the

termination of the study. It became anorectic 14 DPI and remained so throughout the

study. Three of 4 PO inoculated turtles also became anorectic after inoculation and

remained so throughout the study. No other signs of disease were noted in those turtles.

Turtles mock inoculated were negative by PCR on all oral and cloacal swabs

collected (Table 2). Three turtles PO inoculated (Nos. 2, 7 and 10) were positive by PCR

on oral and/or cloacal swabs 2 DPI but not in any subsequent samples. The fourth PO

inoculated turtle (No. 1) was not positive on any sample date. Oral and cloacal swabs

from IM inoculated turtles were positive by PCR for iridovirus at varying times

throughout the study (Table 2). Turtle No. 8 (IM inoculated), which was euthanized at

the end of the study, was never positive on any oral swabs collected, but was positive on

cloacal swab on five occasions including the last two sampling dates that were 23 and 26

DPI. PCR on urine samples followed a similar pattern. Three of four IM inoculated

turtles were positive between one and five days prior to being euthanized (Table 3). One

positive band from each turtle was sequenced to confirm the positive PCR results. All

amplicons were of the expected size and each sequence shared 100% identity with the

sequence of the original isolate. DNA extracted from eight tissues from each turtle

including tongue, esophagus, stomach, small intestine, large intestine, kidney, spleen and

liver were positive by PCR on three of four IM inoculated turtles (Nos. 3, 5 and 6) at

necropsy (Table 1). Tissues from IM inoculated turtle No. 8, and all PO inoculated

turtles and control mock-inoculated turtles were negative.









At necropsy, gross changes were observed in several turtles. Turtle No. 3 that was

IM inoculated exhibited petechia in several organs including the glottis, liver, pancreas

and fat. Congestion was observed in the stomach and on the surface of the bladder. The

cecum and colon demonstrated a multifocal to coalescing area of hemorrhage (Fig. 4-4).

The gastrointestinal tract of turtles 3, 5 and 6 appeared thickened and edematous. Turtle

6 also exhibited petechia on the surface of the pancreas and congestion in the stomach.

No lesions were seen in PO inoculated and control group turtles.

Significant histologic lesions were observed only in IM inoculated turtles. Similar

to the pilot study, the most consistent lesions were in the spleen (3/4 turtles) and centered

on the splenic ellipsoids. The majority of the splenic white pulp surrounded ellipsoids

(periellipsoidal lymphocyte sheath; PELS) (Fig. 4-5A and 4-6A) with lesser white pulp

surrounding arterioles (periarteriolar lymphocyte sheath; PALS) (Fig. 4-6B). Ellipsoids

(Fig. 4-6A) were characterized by plump, cuboidal, endothelial cells, a thick eosinophilic

wall lacking smooth muscle (confirmed with Masson's trichrome stain), and a lack of

reticular fibers between lymphocytes of the PELS using Gordon and Sweet's reticulin

stain. In IM inoculated turtles, the walls of the ellipsoids were moderately to markedly

expanded by homogenous to slightly fibrillar eosinophilic material (fibrin; fibrinoid

vasculitis) (Fig. 4-5B and 4-6B) with multifocal infiltrates of low numbers of heterophils

and scattered free brown pigment granules (presumptively from disrupted

melanomacrophage centers). There were occasional lumenal fibrin thrombi with

admixed heterophils and karyorrhectic debris (Fig. 4-6B). There was mild to moderate

lymphoid depletion and dispersion of lymphoid cells in the PELS with relative sparing of









lymphocytes in the PALS (Fig. 4-5B and 4-6B). Replacing the PELS were combinations

of karyorrhectic debris, heterophilic infiltrates, extravasated erythrocytes and fibrin.

The liver of all turtles had mild to marked hepatocellular vacuolar change that

was most pronounced in the control turtles (Fig. 4-7A) and considerably less prominent

in IM inoculated turtles (depletion of hepatocellular lipid and glycogen) (Fig.4-7B).

Three (No. 3, 5, 6) of four turtles had multifocal random dilatation of hepatic sinusoids

with fibrin thrombi (Fig. 4-7B) and variable single-cell necrosis of adjacent hepatocytes

(Fig. 4-7C). Rarely, hepatocytes had small intracytoplasmic basophilic inclusion bodies

(Fig. 4-7C). Admixed with fibrin thrombi and necrotic hepatocytes were small amounts

of karyorrhectic debris, infiltrates of small numbers of heterophils, and for lesions

occurring adjacent to melanomacrophage centers, small amounts of dispersed brown

granular pigment. One turtle (No. 3) had moderate multifocal hemorrhage in association

with liver lesions as well as moderate multifocal fibrin thrombi within small to medium

portal venules and veins.

In addition to vascular changes in splenic ellipsoids, hepatic sinusoids and portal

blood vessels, acute fibrin thrombi were also observed in a variety of other tissues in

intramuscularly inoculated animals (Fig. 4-8 and 4-9). Thrombi were noted in gastric or

intestinal lamina propria, submucosa and serosa (No. 3, 5, and 6), glomerular capillaries

(No. 3 and 6), esophagus (No. 3), pulmonary capillaries and veins (No. 3 and 6),

meninges (No. 3 and 6), eye (No.3), nasal mucosa (No. 3), and oral mucosa (No. 3).

Lesions associated with thrombi included segmental marked colonic mucosal hemorrhage

(Fig. 4-9), multifocal mild to moderate meningeal hemorrhage, mild to moderate

heterophilic meningitis, and mild heterophilic interstitial pneumonia.









A small number of IM inoculated turtles also had multifocal mild to moderate

ulcerative and heterophilic stomatitis and esophagitis (No. 6) (Fig. 4-10), and multifocal

mild to moderate heterophilic and necrotizing gastritis (No. 5 and No. 6). Turtles from

all three groups had small granulomas in a wide variety of tissues that surrounded

probable spirorchid-type trematode eggs and rarely, adult trematodes were observed

within mesenteric blood vessels. Effete granulomas in the gastric submucosa and serosa

of a small number of control and virus inoculated turtles contained cross-sections of an

unidentified nematode. Trematodes and nematodes were interpreted as incidental

findings.

Similar to study 1, cytopathic effects consisting of cell rounding and lysis were

seen in cultures of Terrapene heart cells that were coated with kidney homogenates from

three of four IM inoculated turtles (Nos. 3, 5 and 6). Using PCR and nucleotide

sequencing, Ranavirus was identified in DNA extracted from cells from each flask. No

CPE was seen in cells that received tissue homogenates from orally inoculated turtles,

control turtles, and one of the intramuscularly inoculated turtles (No. 8).

Transmission electron microscopy of splenic ellipsoids in 1 IM inoculated turtle

(No. 6) in the full transmission study demonstrated marked expansion of the vessel wall

by a granular to fibrillar lightly electron dense material that was consistent with fibrin.

Admixed with fibrin and within remnants of the white pulp were scattered unidentified

necrotic cells with intracytoplasmic arrays of icosahedral virions consistent with an

iridovirus. No virions were observed in cells associated with ellipsoids in the control

turtle (No. 9).









Discussion

Henle-Koch's postulates provide a strict set of guidelines for proving a causal

relationship between an infectious agent and a disease (Evans, 1976). Here we

performed a transmission study using a Ranavirus isolate from a captive Burmese star

tortoise that became ill and died in an attempt to determine if a causal relationship exists

between infection with this Ranavirus and the clinical and histologic changes observed in

the Burmese star tortoise. Since it is not practical to perform a challenge study in this

critically endangered species, we decided to assess both box turtles and red-eared sliders

as a model for Ranavirus infection in chelonians. Box turtles were selected since

Ranavirus infection has been identified in this species (Mao et al., 1997; Devoe et al.,

2004; Johnson et al., 2004). We decided to also evaluate the suitability of red-eared

sliders since populations of box turtles are declining throughout their range. Red-eared

sliders, however, are being raised in the lower Mississippi Valley for the overseas pet

trade, which became a factor in their ultimate selection as an experimental animal in our

studies. In addition, results of the pilot study showed that both species similarly

responded when administered a Ranavirus isolate by two different routes. Both IM

inoculated turtles showed severe clinical signs and were euthanized prior to the

termination of the study at two weeks. Oral swabs from both were positive one week

post-inoculation and histologic lesions were consistent between the two species. Both the

PO inoculated RES and BT did not show any clinical or histologic lesions. Results of

this study showed that both RES and BT can serve as suitable models of Ranavirus

infection for chelonians. As a result, RES were chosen as the experimental model for the

larger transmission study.









Histologic lesions in turtles inoculated by intramuscular injection in both studies

consisted primarily of multicentric fibrinoid vasculitis and formation of fibrin thrombi in

small blood vessels in numerous tissues, and in this regard, resembled recent reports of

Ranavirus infection in captive and free-ranging box turtles and tortoises (DeVoe et al.,

2004; Johnson et al., 2004). Lesions in blood vessels were consistent with observed

Ranavirus infection of endothelial cells in a naturally-infected captive Burmese star

tortoise (Johnson et al., 2004) and with descriptions of apparent viral endotheliotropism

in rainbow trout and redfin perch infected with another Ranavirus called epizootic

haematopoietic necrosis virus (EHNV) (Reddacliff and Whittington, 1996). Involvement

of endothelial cells may also be part of the pathogenesis of some amphibian Ranavirus

infections as suggested by multicentric hemorrhage and edema or observation of

characteristic inclusion bodies within endothelial cells (Wolf et al., 1968; Cunningham et

al., 1996; Bollinger et al., 1999; Docherty et al., 2003).

The consistent involvement of the splenic ellipsoids (sheathed capillaries) provides

a basis for the prominent necrotizing splenitis observed in some natural Ranavirus

infections of chelonians (DeVoe et al., 2004; Johnson et al., 2004). Similar lesions of the

ellipsoids were observed in redfin perch, but not rainbow trout, experimentally infected

with EHNV (Reddacliff and Whittington, 1996). A filtering function for splenic

ellipsoids for particulate material and immune complexes has been documented in other

species (Sorby et al., 2005) and possibly, the lesions observed in chelonian Ranavirus

infections could be a consequence of antigen trapping in macrophages associated with the

ellipsoid sheath during ranaviral viremia.









Necrotizing stomatitis, esophagitis and/or pharyngitis are characteristic lesions in

many naturally occurring chelonian Ranavirus infections (Westhouse et al., 1996;

Marshang et al., 1999; DeVoe et al., 2004; Johnson et al., 2004). Clinical signs such as

ocular and nasal discharge, conjunctivitis and palpebral edema associated with Ranavirus

infection (both naturally infected animals and the experimental animals in this report) are

often attributed to the upper respiratory tract and overlap with those signs observed with

mycoplasmosis caused by Mycoplasma agassizii (Brown et al., 1999) or herpesvirus

infection (Origgi, et al., 2004; Johnson et al., 2005). In particular, infection with tortoise

herpesviruses 1 and 2 are associated with caseous or diphtheritic oral plaques that are

grossly indistinguishable from oral lesions associated with Ranavirus infection. In the

experimentally infected animals of this report, necrotizing stomatitis and esophagitis

were observed in a single intramuscularly inoculated red-ear slider (No. 6). Lesions were

not observed in the nasal cavity of any animal examined. Oral lesions could be

secondary to thrombus formation and infarction in small submucosal vessels or

alternatively, could be the result of viral infection and necrosis of oral epithelial cells.

The observation of intracytoplasmic inclusion bodies consisting of Ranavirus in epithelial

cells of some naturally occurring infections would appear to support the latter

explanation. Why oral lesions were not present in more of the experimentally inoculated

turtles in this report is uncertain, but it is possible that epithelial infection is a late event

that follows viremia and hence was not observed in experimental animals. Also, the virus

may have been attenuated in cell culture, resulting in altered pathogenicity.

The intracytoplasmic basophilic inclusion bodies that are suggestive of iridovirus

infection and prominent in many cases of Ranavirus infection in fish, amphibians and









some chelonians were not prominent in the experimentally inoculated turtles (Reddacliff

and Whittington, 1996; Westhouse et al., 1996; Bollinger et al., 1999; Marschang et al.,

1999; Docherty et al., 2003). This observation is consistent with recently reported

naturally occurring chelonian Ranavirus infections, and indicates that inclusion bodies

may be an inconsistent finding and should not be relied upon for use in formulating a

histologic differential diagnosis (Devoe et al., 2004; Johnson et al., unpublished

findings). Virions consistent with ranaviruses were observed by transmission electron

microscopy in cells within the spleen of an IM inoculated red ear slider and suggests that

TEM may still be a useful diagnostic tool in chelonian Ranavirus infections even in the

absence of visible inclusions on histologic section. Demonstration of intracytoplasmic

virions in cells of an experimentally inoculated turtle is important because it shows that

the virus is capable of entering and replicating within cells, and that, lesions were not

induced by the presence of inoculated non-replicating virus. Necrotizing liver lesions

have been experimentally induced in mice and rats following injection of inactivated

iridovirus virions or solubilized structural proteins (Lorbacher de Ruiz, 1990). Similarly,

the Ranavirus Frog Virus 3 can trigger apoptosis in tissue culture in the absence of viral

gene expression (Chinchar, 2002). Future studies may better define the in-vivo

mechanism of cell death associated with iridovirus infections.

This study found that IM inoculated turtles were more likely to become infected

with Ranavirus, show clinical signs and die compared to turtles that were orally

inoculated. All four IM inoculated turtles showed clinical signs and three died as a result

of infection (75%), whereas no orally inoculated turtles showed any signs of disease or

died. This could mean that turtles do not become exposed through ingestion of infected









animals or water sources as has been previously shown to occur in amphibians

(Jancovich et al., 2001, Pearman et al., 2004), or that abrasions naturally acquired from

ingesting bones or other abrasive material may be necessary for virus to be introduced

systemically. Another explanation for the inability to re-create disease in orally

inoculated animals was that a natural exposure was not replicated in the laboratory

setting. Viral dose administered was extrapolated from studies done with fish and

amphibians (Langdon, 1989, Moody and Owens, 1994, Bollinger, 1999, Cullen and

Owens, 2002) but requirements for infection of turtles may be higher or repeated

exposure may be necessary. Experimental infections of salamanders with a Ranavirus

showed that both dose and host characteristics influenced the virulence of infection

(Brunner et al., 2005). The infection dose was positively correlated with the mortality

rate and inversely related to average survival times. Environmental temperatures have

also been shown to significantly impact the percent mortality and time to death in

salamanders experimentally inoculated with a Ranavirus (Rojas et al., 2005), where

salamanders inoculated at 18 and 100C were more likely to die than those exposed at

26C. While water and room temperatures averaged between approximately 21 and

250C, basking areas were kept warmer at 280C. Eliminating heat lamps over basking

areas and lowering the room temperature might have kept turtles cooler, and altered the

results in the orally inoculated group. Alternatively, other routes of transmission such as

vector-borne transmission may be required for turtles to become infected in the wild.

Intracytoplasmic inclusion bodies were recently identified in the circulating leukocytes of

an eastern box turtle infected with iridovirus (Allender et al., In Press). Ranaviruses are

variably host specific, so virus may be able to survive in mosquitoes or other biting









insects capable of transmitting virus from an infected animal to an uninfected one. This

has been shown to be the case with insect iridoviruses, where parasites were

experimentally shown capable of transmitting invertebrate iridescent virus from infected

to uninfected larvae (Lopez et al., 2002). It remains unknown whether natural outbreaks

of iridovirus infections in any vertebrate species occur as a result of introduction of novel

virus strains, recrudescence of latent or persistent infections in surviving populations as a

result of stressors or other immunosuppressive causes, or viral persistence in the

environment (Williams et al., 2005).

Although the mechanism of transmission of iridoviruses in natural settings is

unknown, it has been shown that sublethally infected amphibians can cause sporadic,

recurrent disease outbreaks in amphibians (Brunner et al., 2004). Experimentally and

naturally infected tiger salamander larvae and metamorphs were able to maintain

sublethal, transmissible infections for over five months. Apparently healthy infected

dispersing metamorphs were returning to water bodies to breed and it was speculated that

these individuals were likely serving as a reservoir host for infecting newly hatched

larvae, creating recurrent outbreaks of disease. The current study showed that turtles may

also become asymptomatic carriers, although further studies would help to confirm this

finding. Turtle No. 8 which was inoculated intramuscularly showed transient signs of

disease but then recovered. At necropsy, tissues collected from eight different organs

were negative for iridovirus on PCR. Kidney samples inoculated onto TH-1 cells showed

no cytopathic effects. However, cloacal swabs collected one, four and eight days prior to

necropsy were positive using PCR. If this was a result of slow elimination of the virus,

all orally inoculated turtles should have had PCR-positive cloacal samples after 2 DPI.









However, all swabs taken after this time period were negative. Furthermore, given that

IM inoculation was used in turtle No. 8, any virus shed from the GI tract would require

transportation through the blood stream and across the wall to be there. While it is

possible that this was a laboratory contaminant, all swabs were extracted and tested by

PCR according to date collected as opposed to all swabs tested at the same time per

turtle. If there was contamination, we would not expect to see it on five occasions from

one turtle and not from any orally inoculated turtles on any date past 2 DPI. Therefore it

seems more likely that virus shedding was occurring, and the site of virus replication was

missed. Some viruses show a predilection for specific cells, such as infectious bursal

disease virus in chickens, where virus shows a predilection for the cells of the bursa of

Fabricius, located in the cloaca (Burkhardt and Miller, 1987). Further long-term studies

would help to confirm whether Ranavirus persistence occurs and immunohistochemical

or in situ hybridization studies of tissues from infected turtles may help identify the tissue

tropism for Ranavirus persistence. If turtles can serve as asymptomatic carriers, they

may also serve as a reservoir host of virus for other turtles and other susceptible species.

Inter-class infections of iridovirus have been shown naturally and experimentally in

sympatric species of fish and amphibians, where both were capable of being infected with

the same virus (Mao et al., 1999, Moody and Owens, 1994). In another study (data not

shown), the isolate used in this experimental study was capable of infecting leopard frogs

(Ranapipiens) that were injected intraperitoneally at the same dose. Thus, sublethally

infected turtles such as one of the turtles (No. 8) in our study could serve as a reservoir

host for amphibian populations in geographic locations where the species overlap.









In summary, the experimental inoculation of a Ranavirus in chelonians was

investigated. Koch's postulates were fulfilled when intramuscular inoculation of virus

into naive turtles resulted in clinical and histologic changes consistent with those seen in

natural infections, and when the same virus was subsequently recovered. Since oral

inoculation failed to result in disease or mortality, the natural route of transmission in the

wild remains unknown. The immune system of reptiles is temperature dependent

(Cooper et al., 1985) and perhaps by manipulating the environmental temperature, and

the temperature of the host, the susceptibility of turtles to infection can be altered.

Another possibility that should be investigated is the transport and inoculation of

Ranavirus into chelonians by arthropods. This study also suggests that sublethally

infected turtles may serve as reservoir hosts of infection for other chelonians, as well as

amphibians. Ranaviruses are considered a global threat to amphibian populations based

on the lack of host specificity, high virulence and global distribution (Daszak et al., 1999)

and this study confirms that they should likewise be considered a threat to chelonian

populations.