Diagnosis of Reptilian Viral Disease
American Association of Zoo Veterinarians Conference 1999
Elliott R. Jacobson, DVM, PhD
Department of Small Animal Clinical Sciences, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA


Starting in 1966 with the identification of the viral nature of intracytoplasmic inclusions in erythrocytes of geckos, more and more viruses have been identified as potential pathogens in reptiles. Many diseases originally thought to be bacterial in origin are now being recognized as viral diseases. Pneumonia that was first seen in a colony of vipers (Bothrops moojeni) was initially thought to be caused by Pseudomonas. Subsequent investigations identified the presence of a myxovirus and transmission studies have confirmed the pathogenic effect of these viruses. There is a very basic approach in determining the viral basis for a disease. The history of a die-off coupled with histopathologic findings may be suggestive. Ultimately, virus needs to be identified in tissues either by electron microscopy or by culturing the agent in tissue culture. Negative staining electron microscopy can be used to identify particles in feces, secretions, and tissue homogenates. Serology also has been used to determine exposure to a pathogen and PCR can be used to identify presence of specific nucleotide sequences in tissues or secretions of the host. Ultimately, virus needs to be cultured, purified and challenged back into naive animals to establish a causal relationship.


Since 1966, a wide variety of viruses have been either identified in tissue section and/or isolated from members of the orders Chelonia (turtles and tortoises), Crocodylia (alligators, crocodiles, caiman, and gharial), and Squamata (Lacerta: lizards; Ophidia: snakes).5 An endogenous retrovirus is the only virus identified in Rhynchocephalia (tuatara).14 Most viruses identified in reptiles have been only circumstantially incriminated as causes of disease, and few studies fulfill Koch’s postulates. In many cases evidence of viral infection is based upon identification in tissue section by light microscopy, with more specific identification made using electron microscopy. Of the various viruses identified in reptiles, transmission studies demonstrating a causal relationship have only been documented for the gray patch herpesvirus of green turtles (Chelonia mydas) in aquaculture10, ophidian (snake) paramyxovirus7, and a retrovirus isolated from boid snakes with inclusion body disease13. Most of the viruses identified in tissue section of reptiles have not been isolated and their role in reptile disease awaits further studies.

In this paper I will review the various diagnostic approaches used in determining the presence of viruses in reptiles.

Histopathology and Transmission Electron Microscopy

While viruses have been isolated from tissues of reptiles without concomitant histopathology, often the isolation of viruses follows light microscopic findings suggestive of viral infection. Tissue specimens at necropsy are routinely collected in neutral buffered 10% formalin and if a viral infection is suspected, additional samples can be placed in Trump’s solution (a 4% formalin/1% glutaraldehyde mixture).8 If there is the potential of performing immunohistochemical staining such as for ophidian paramyxovirus, tissues should be transferred and stored in 70% ethanol at 24 to 48 hr following initial fixation.3 Frozen tissues can be collected for demonstration of viral antigen using antibody-labeled fluorescence microscopy. This has been used to demonstrate presence of ophidian paramyxovirus in lung tissue of infected snakes.7 Based upon light microscopic findings such as the presence of intranuclear inclusions in chelonian herpesvirus infections,9 or intracytoplasmic inclusions in boid inclusion body disease,13 the next step would be evaluation of tissues by transmission electron microscopy (TEM). While in TEM it is ideal to have tissues fixed in an appropriate fixative such as Trump’s solution or 2.5% glutaraldehyde, formalin fixed tissue and even paraffin embedded tissue can be examined for presence of virus. Sections of paraffin embedded tissue can be cut from a block using a scalpel blade, post-fixed, and submitted for TEM. The cost of TEM ranges from $100.00 to $200.00 per sample for processing, sectioning, examination, and photography. Based upon the size, location, morphology and morphogenesis, a presumptive categorization into a family of virus often can be made. However, a final diagnosis is dependent upon biochemical characterization or determining specific nucleotide sequences using polymerase chain reaction.

Negative Staining Electron Microscopy

Negative staining in conjunction with TEM is a useful and rapid method of examining clinical specimens for presence of virus. The principle of negative staining is that there is no reaction between the stain and the specimen. The most commonly used negative stains are uranyl acetate (0.5–1.0%) and potassium phosphotungstate (PTA) (0.5–3.0%). Depending on the nature of the tissue, different ways of processing the sample are required to detect viral particles. Fluid from vesicles can be obtained with a sterile pipette and may be placed directly on a Formvar-coated 200 mesh copper grid, while large amounts of fluid (lung washings) require centrifugation for clarification. In these cases the supernatant after low speed centrifugation (1,500 g), or the diluted pellet after high speed centrifugation (15,000 g), is placed on the grid for staining. Fecal material requires suspension and concentration and should be placed in distilled water or phosphate buffered saline (PBS). Fecal material can be mixed with PBS to give a 20% suspension in 5 ml. After centrifuging, a drop of the supernatant is placed on a grid for examination.

Cell Culture

Viruses can be isolated from: 1) swab specimens of lesions or luminal surfaces; 2) tissue biopsies and 3) tissues collected at necropsy. When biopsies or tissues are collected at necropsy, it is important to collect samples as aseptically as possible. Tissues can be placed in a sterile petri dish and transported to the laboratory on ice, frozen in sterile plastic bags at -70°C for future viral isolation, or placed in cell culture media and frozen for future viral isolation attempts. Tissues should be sectioned into small pieces or ground in a tissue grinder, with cell culture media added. Processed samples are added to plastic cell culture flasks that contain actively dividing cells. Several reptile cell lines are commercially available from American Type Cell Culture (Bethesda, Maryland) and include: 1) viper heart cells; 2) iguana heart cells and 3) Terrapene heart cells. Cells are incubated at 30°C and are checked daily for cytopathic effects (CPE). Certain reptile viruses such as ophidian paramyxovirus have been adapted to mammalian Vero cells. This has certain advantages since it is possible to achieve faster growth of mammalian cells in culture compared to reptile cells. If changes are seen such as cell necrosis or syncytial cell formation, samples can be removed from a flask, fixed and processed for electron microscopy. Suspected viral antigen can be demonstrated in infected cells using a fluorescent antibody technique. This approach has been used for demonstrating the presence of ophidian paramyxovirus in infected viper heart cells.11


Very few serologic tests have been developed or are available for diagnosing exposure of reptiles to specific viruses. Serologic studies have been performed on free-ranging reptiles and reptiles used in experimental studies in order to determine the presence of antibodies to various togaviruses.7 However, infections with these viruses do not appear to be clinically important in reptiles. A viral neutralization test has been developed to determine exposure of tortoises (Testudo hermanii and T. graeca) to a herpesvirus suspected of causing a stomatitis/pharyngitis.5 A hemagglutination inhibition (HI) assay has been developed to determine presence of antibody against ophidian paramyxoviruses.6,12 The HI assay has been most useful because of its relative simplicity and rapid “turnaround” time.2 Briefly, serum samples collected by heart puncture or tail venipuncture are diluted 1:10 in sterile physiologic saline at 56°C for 30 min to inactivate complement, then absorbed with washed and pelleted chicken erythrocytes to remove nonspecific agglutinins (12 hr at 5°C). Using microtiter methodology, serial doubling serum dilutions are made (1:10, 1:20, 1:40, 1:80, etc.) using 0.05 ml. volumes of phosphate buffered physiologic saline containing 0.1% bovine serum albumin. The latter minimizes autoagglutination of the erythrocyte suspension used later to indicate whether active virus is present or not. To each serum dilution is added an ophidian paramyxovirus suspension diluted to contain 8 hemagglutination units/0.05 ml. This has previously been determined by titration, taking advantage of the fact that the virus causes chicken erythrocytes to bind together (hemagglutinate). After allowing the virus-serum mixtures to interact for 1 hr at room temperature, 0.1 ml of a 0.3% suspension of washed chicken erythrocytes is added to each well of the microtiter plate. The plates are placed in a refrigerator for 2–3 hr to permit settling of the erythrocytes. If antibodies are present in a particular serum dilution, they will bind to the viruses and prevent them from hemagglutinating the erythrocytes (hemagglutination-inhibition). Where antibody is present, the red cells settle into a “button” at the bottom of the microtiter well, rather than forming a “mat” due to the hemagglutinating property of the viruses. The serum antibody “titer” is read as the reciprocal of the highest serum dilution which causes hemagglutination-inhibition. An HI titer >20 is considered to be positive, indicating definite exposure to virus. Snakes which survive paramyxovirus infections may have HI antibody titers exceeding 10,240.2

A single sample is only indicative of the exposure status at the time the sample was collected. In order to demonstrate an active infection, samples should be collected at 2–4-wk intervals. A change of titer greater than one dilution would indicate an active infection.

The laboratory of the author of this paper is currently performing assays for zoological and private collections throughout the United States. Blood samples are collected and placed in lithium heparin tubes. Plasma is removed, placed in a cryotube, and submitted on dry ice. Minimally, 0.2 cc of plasma should be provided. The cost of an assay is $30.00.


Polymerase chain reaction (PCR) is a method detecting portions of DNA by amplifying short nucleotide sequences within the DNA genome. PCR can be used for detection of viruses in: 1) cell culture; 2) tissues; 3) secretions; 4) lavages and 5) other biologic samples. A primer of the sequence being amplified is needed. PCR requires oligonucleotide primers complimentary to the sequence being amplified. A PCR assay has been developed for detecting presence of the marine turtle fibropapilloma associated herpesvirus in tumors of sea turtles (Dr. Paul Klein, Department of Pathology and Laboratory Medicine, University of Florida, Gainesville, Florida).

Literature Cited

1.  Blahak S, R Biermann. 1995. Herpesvirus infection in land tortoises as a problem of chelonian conservation. Proceedings of the International Congress of Chelonian Conservation. Gonfaron, France, Pp.240–243.

2.  Gaskin JM, M Haskell, N Keller, E Jacobson. 1989. Serodiagnosis of ophidian paramyxovirus infections. In: Third International Colloquium on the Pathology of Reptiles and Amphibians. Abstracts. Orlando, Fl, Pp.21–22.

3.  Homer BL, JP Sundberg, JM Gaskin, J Schumacherand, ER Jacobson. 1995. Immunoperoxidase detection of ophidian paramyxovirus in snake lung using a polyclonal antibody. J. Vet. Diag. Investig. 7:72–77.

4.  Jacobson ER. 1986. Viruses and viral diseases of reptiles. Acta Zoologica et Pathologica Antiverpiensia. Pp. 73–89.

5.  Jacobson ER. 1993. Viral diseases of reptiles. In: Fowler ME, ed. Zoo and Wild Animal Medicine, Current Therapy 3. W.B. Saunders, Philadelphia, Pp. 153–159.

6.  Jacobson E, JM Gaskin, et al. 1981. Illness associated with paramyxo-like virus infection in a zoological collection of snakes. J. Amer. Vet. Med. Assoc. 179:1227–1230.

7.  Jacobson ER, HP Adams, TW Geisbert, SJ Tucker, B Hall, B Homer. 1997. Pulmonary lesions in experimental ophidian paramyxovirus pneumonia of Aruba Island rattlesnakes, Crotalus unicolor. Vet. Pathol. 34: 450–459.

8.  McDowell EM, BF Trump. 1976. Historical fixative suitable for diagnostic light and electron microscopy. Arch Path Lab Med. 100:405–414.

9.  Muller M, W Sachsse, N Zangger.1990. Herpesvirus-Epidemie beider griechischen (Testudo hermanni) und der maurischen Landschildkrote (Testudo graeca) in der Schweiz. Schweiz Arch Tierhelk. 32:199–203.

10.  Rebell H, A Rywlin, HA Haines. 1975. Herpesvirus-type agent associated with skin lesions of green turtles in aquaculture. Am J Vet Res. 36:1221–1224.

11.  Richter G. 1994. Determination of Optimal Culture Technique and the Study of Physicochemical and Cytopathological Aspects of Ophidian Paramyxovirus [Master of Science Thesis]. University of Florida.

12.  Richter GA, BL Homer, SA Moyer, DS Williams, G Scherba, SJ Tucker, BJ Hall, JC Pedersen, ER Jacobson. 1996. Characterization of paramyxoviruses isolated from three snakes. Virus. Res. 43: 77–83.

13.  Schumacher J, ER Jacobson, B Homer, JM Gaskin. 1994. Inclusion body disease of boid snakes. J. Zoo Wildl. Med. 25:511–524.

14.  Tristem M, T Myles, F Hill. 1995. A highly divergent retroviral sequence in a tuatara (Sphenodon). Virol. 210:206–211.


Speaker Information
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Elliott R. Jacobson, DVM, PhD
Department of Small Animal Clinical Sciences
College of Veterinary Medicine
University of Florida
Gainesville, FL, USA

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