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Prevalence of Ranavirus in Virginia Turtles as Detected by Tail-Clip Sampling Versus Oral-Cloacal Swabbing

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Ranaviruses are emerging infectious diseases that infect amphibians, fish, and reptiles. Several cases of morbidity and mortality in captive and natural populations of reptiles have been attributed to ranaviruses, but research in this taxon has been limited. We used oral-cloacal swabs and tail clips to survey two species, Chrysemys picta picta (Eastern Painted Turtles) and Sternotherus odoratus (Common Musk Turtles), in three water bodies in central Virginia to determine if ranaviruses were present. Prevalence of ranavirus in C. p. picta ranged from 4.8–31.6% at the three sites. Ranavirus was not detected in S. odoratus, but only oral-cloacal swabs were used in this species because of the cornified tail tip. While tail-tip tissues from all three study sites indicated presence of ranavirus in C. p. picta, no oral-cloacal swabs from these same turtles tested positive. We therefore suggest that oral-cloacal swabbing may yield false negatives when ranavirus is present in turtles, and that tissue sampling may be more appropriate for monitoring ranavirus in turtles.
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NORTHEASTERN NATURALIST
2013 20(2):325–332
Prevalence of Ranavirus in Virginia Turtles as Detected by
Tail-Clip Sampling versus Oral-Cloacal Swabbing
Rachel M. Goodman1,*, Debra L. Miller2,3, and Yonathan T. Ararso1
Abstract - Ranaviruses are emerging infectious diseases that infect amphibians, sh, and
reptiles. Several cases of morbidity and mortality in captive and natural populations of
reptiles have been attributed to ranaviruses, but research in this taxon has been limited.
We used oral-cloacal swabs and tail clips to survey two species, Chrysemys picta picta
(Eastern Painted Turtles) and Sternotherus odoratus (Common Musk Turtles), in three
water bodies in central Virginia to determine if ranaviruses were present. Prevalence
of ranavirus in C. p. picta ranged from 4.8–31.6% at the three sites. Ranavirus was not
detected in S. odoratus, but only oral-cloacal swabs were used in this species because of
the cornied tail tip. While tail-tip tissues from all three study sites indicated presence of
ranavirus in C. p. picta, no oral-cloacal swabs from these same turtles tested positive. We
therefore suggest that oral-cloacal swabbing may yield false negatives when ranavirus
is present in turtles, and that tissue sampling may be more appropriate for monitoring
ranavirus in turtles.
Introduction
Biodiversity is declining worldwide, and many biologists believe we are
witnessing the sixth mass extinction in the history of life (Barnosky et al.
2011, Wake and Vredenburg 2008). Nearly half of all amphibian populations
are in decline (IUCN et al. 2008), and reptiles may face similar levels of en-
dangerment (Gibbons et al. 2000, IUCN 2010, Reading et al. 2010). Many
factors have contributed to population declines and extirpations: habitat de-
struction and degradation, pollution, global climate change, introduction of
non-native species, and emerging infectious diseases (Wells 2007, Wilcove
et al. 1998). Globally, two thirds of freshwater turtle and tortoise species are
considered threatened or endangered (IUCN 2010), and infectious diseases
may be a contributing factor (Ernst and Lovich 2009). Emerging infectious
diseases contribute to population declines, and ranaviruses (family Irido-
viridae; genus Ranavirus) are of concern because they infect multiple taxa,
including fish, reptiles, and amphibians (Chinchar 2002). Currently we have
limited research on the susceptibility of this wide range of potential hosts and
the potential for transfer among species.
Ranaviruses are double-stranded DNA viruses that infect reptiles, am-
phibians, and fish and have caused mortality events in each taxon (reviewed
in Chinchar 2002). The importance of ranaviruses in amphibian population
1Biology Department, Box 74, Hampden-Sydney College, Hampden-Sydney, VA 23943.
2Veterinary Diagnostic and Investigational Laboratory, University of Georgia, Tifton, GA
31793. 3Current address - University of Tennessee, Center for Wildlife Health, Knoxville,
TN 37996. *Corresponding author - rgoodman@hsc.edu.
Northeastern Naturalist Vol. 20, No. 2
326
declines has only recently been recognized, although they have caused more
die-offs in North America than the more-studied fungal pathogen Batracho-
chytrium dendrobatidis (Daszak et al. 1999, Duffus 2009, Gray et al. 2009).
Among fish, iridovirus infections have been reported on several continents
and can cause economic damage in commercial freshwater fisheries (Ahne et
al. 1997, Whittington et al. 2010). The importance of ranaviruses for reptilian
population dynamics is unknown, but several cases of morbidity and mortal-
ity in captive and natural populations have been attributed to the pathogen
(De Voe et al. 2004, Hyatt et al. 2002, Marschang et al. 2011). Research thus
far has been limited to description and isolation of viruses from infections in
captive and wild species (Chen et al. 1999, De Voe et al. 2004, Johnson et al.
2008, Marschang et al. 1999, Westhouse et al. 1996), and clinical challenges
of two North American species, Terrapene ornata ornata Agassiz (Ornate Box
Turtle) and Trachemys scripta elegans Weid-Neuwied (Red-eared Slider),
and two Australian species, Emydura krefftii Gray (Krefft's River Turtle)
and Eiseya latisternum Gray (Saw-shelled Turtle) (Ariel 1997, Johnson et al.
2007). Signs of ranavirus infection in turtles reported in these studies include
lethargy, respiratory distress, anorexia, cutaneous erythema, ocular and nasal
discharge, and oral ulceration and plaques. Surveillance of ranavirus in reptile
populations is important to determine whether associated disease threatens
persistence, and whether sub-lethally infected reptiles may serve as reservoirs
for the pathogen that threatens co-occurring species. Also, this work in reptiles
is necessary to gain an understanding of the complete epidemiology, including
interspecific transmission, of ranaviruses. In the current study, we used and
compared oral-cloacal swabbing and tissue sampling for ranavirus surveillance
in two species of turtles, Chrysemys picta picta Schneider (Eastern Painted
Turtles) and Sternotherus odoratus Latreille (Common Musk Turtles), in three
water bodies in Virginia.
Field Site Description
The study was conducted at three sites in Prince Edward County, VA: Briery
Creek Lake in Briery Creek Wildlife Management Area (north end; 37°12.0'N,
78°27.0'W), and two ponds on the campus of Hampden-Sydney College (HSC),
Chalgrove (37°14.5'N, 78°27.8'W) and Tadpole Hole (37°14.7'N, 78°27.2'W).
Chalgrove and Tadpole Hole are both approximately 1 ha and located 0.8 km apart.
Briery Creek Lake is a 342-ha lake managed by the Virginia Department of Game
and Inland Fisheries and is located 4.5 km south of the HSC ponds (Fig. 1).
Methods
Turtles were collected during 24 May–1 July 2010. We changed trapping
sites every week, and trapped at each site twice, with 6–10 visits per site.
Traps were set 1–2 m from shore and included four Promar collapsible crab/
fish traps with dual-ring entrance, a Sundeck turtle trap with a bait tower (Item
#840876, Heinsohn’s Country Store, http://www.texastastes.com/outdoors.
R.M. Goodman, D.L. Miller, and Y.T. Ararso2013 327
htm), and a floating turtle tunnel (Item#840460, Heinsohn’s Country Store).
Because all turtle traps could capture more than one turtle at a time, there was
a small risk that pathogen transmission could occur among individuals within
the traps.
Upon removal from traps, turtles were weighed, measured for mass and
length, and individually marked using combinations of notches filed into
scutes. We used and compared two methods of sampling for ranavirus, oral-
cloacal swabbing and tail clips, for use in the polymerase chain reaction (PCR).
We swabbed turtles with plastic, sterile, cotton-tipped applicators (Puritan
model 25-806 2PC), first rolling it inside the mouth and then inside the cloaca
for 3–5 seconds each. The distal-most 0.5 cm of the tip of the tail was collected
only from species not possessing cornified tail tips (i.e., C. p. picta) using a
new, sterile scalpel blade for each animal. Both tissue samples and swabs were
stored in 1-ml vials containing 70% ethanol. Turtles were released at the site of
capture immediately after sampling.
A total of 106 turtles, including C. p. picta (n = 63) and S. odoratus (n = 43),
were captured, and all turtles appeared clinically normal. Chrysemys picta picta
Figure 1. Map of three water bodies in central Virginia where turtles were sampled for
ranavirus: Chalgrove, Tadpole Hole, and Briery Creek Lake. The star indicates the area at
Briery Creek Lake where turtle trapping was conducted (across most of shoreline at other
sites). GPS coordinates are given in the Methods section.
Northeastern Naturalist Vol. 20, No. 2
328
were collected at all sites, whereas S. odoratus were only collected from Briery
and Chalgrove (Table 1). Among the samples collected, only those from species
and sites with sample sizes of approximately 20 were tested. All traps, rubber
boots and waders, and other gear were scrubbed, soaked in a 1% chlorhexidine
diacetate (Fort Dodge Nolvasan Solution) for at least one minute, and rinsed in
water between use at different water bodies.
Genomic DNA was extracted from the tissues or swabs using a commer-
cially available kit (DNeasy Blood and Tissue Kit, Qiagen, Inc., Valencia, CA).
Negative and positive extraction controls were included. Conventional PCR
was performed using the protocol and primer sets (MCP4 and MCP5) found in
Mao et al. (1996, 1997) and targeting an approximately 500-base pair sequence
of the major capsid protein (MCP) gene. The PCR products were resolved via
electrophoresis on a 1.0% agarose gel. Controls for each PCR run included two
negative controls (water and gDNA extracted from a ranavirus-negative tad-
pole) and two positive controls (cultured ranavirus and gDNA extracted from
a ranavirus-positive tadpole). The PCR protocol was repeated once more on all
samples to verify results.
Results
Only oral-cloacal swabs were tested for S. odoratus, and none were positive
for ranavirus (Table 1). While tail tips from all three study sites indicated pres-
ence of ranavirus among C. p. picta, none of the oral-cloacal swabs from these
same turtles tested positive (Table 1). However, two of the eight ranavirus-
positive individuals that tested positive for ranavirus via tissue samples did not
have accompanying oral-cloacal swabs because they were too small for effec-
tive use of technique (i.e., juveniles). Based on tail-tissue sampling, prevalence
of ranavirus in C. p. picta was 4.8% in Briery, 31.6% in Chalgrove, and 17.4%
in Tadpole Hole.
Discussion
We found evidence of ranavirus infection in C. p. picta in our three study
sites using tail-tissue sampling; however, oral-cloacal swab sampling failed to
detect the pathogen. These ndings suggest that oral-cloacal swabbing may yield
false negatives when ranavirus is present in turtles, and that tissue sampling may
be more appropriate. Gray et al. (2012) conducted a controlled infection study
with Lithobates catesbeianus Shaw (American Bullfrog) tadpoles and found
Table 1. Ranavirus infections in turtles from three water bodies in central Virginia.
Chrysemys picta picta Sternotherus odoratus
Tissues Swabs Swabs
Water body n Ranavirus + n Ranavirus + n Ranavirus +
Briery 21 1 (4.8%) 21 0 (0.0%) 21 0 (0.0%)
Chalgrove 19 6 (31.6%) 8 0 (0.0%) 22 0 (0.0%)
Tadpole Hole 23 4 (17.4%) 21 0 (0.0%) - -
R.M. Goodman, D.L. Miller, and Y.T. Ararso2013 329
false-negative and false-positive rates of 20% and 6% for tail samples, and 22%
and 12% for swabs, respectively, using liver samples as the standard for virus
infection. Those results suggest a similar rate of false negatives for tail and swab
samples in an amphibian, whereas our eld surveillance study suggests a differ-
ence between the methods in a reptile. Further comparisons in additional species
may be warranted.
Necropsy and histology provide the most certain evidence for ranaviral dis-
ease (Miller and Gray 2010); however, lethal sampling is not desirable in the
absence of morbidity or mortality events. Oral-cloacal swabbing is the least
invasive method of sampling, but the current study indicates the sensitivity
of this testing method may be low. While not compared to testing internal or-
gans, tail-tip sampling appears to be more sensitive than oral-cloacal swabbing
and was able to detect ranavirus in C. p. picta using moderate sample sizes of
around twenty individuals. Future research may investigate other potential
areas for superficial tissue sampling on catch-and-release specimens, particu-
larly for species with a cornified or boney tail tip that is used in courtship and
copulation (Ernst and Lovich 2009). In such species, we recommend an ap-
proximately 5-mm diameter skin (epidermal and dermal) biopsy taken from
the mid-dorsal tail.
Compared to common rates for amphibians, prevalence of ranavirus in-
fection in turtles was low in the three water bodies sampled. Research in
amphibians indicates that prevalence can vary widely, depending on the spe-
cies and date of sampling. Using tissue collected from all major organs, Gray
et al. (2007) found ranavirus prevalence of 15–57% in tadpoles in undisturbed
and cattle accessed ponds, depending on the species and sampling period.
Using tail tips and liver samples, Brunner et al. (2004) found prevalence of
46–100% in dispersing metamorph salamanders following an epidemic, but
only 7% prevalence in adults returning to ponds in the following spring. A
recent survey of injured/rehab and free-ranging Terrapene carolina carolina
L. (Eastern Box Turtle) found prevalence of ca. 3% from blood samples col-
lected from injured/sick turtles submitted to rehab centers/medical facilities in
the southeastern US (Allender et al. 2011). This same study was able to detect
ranavirus in swab samples collected from injured/sick T. c. carolina, a species
that spends large amounts of time on dry land, submitted to the medical facility
in Tennessee. Our study differs from Allender et al. (2011) in that we surveyed
a heavily aquatic species and, if it holds true that water is an excellent me-
dium for ranavirus (Chinchar 2002), one would expect greater prevalence in
turtles spending more time in water. However, a recent survey of free-ranging
C. picta and Emydoidea blandingii Holbrook (Blanding’s Turtle) in Illinois
found 0% prevalence for ranavirus using blood samples and oral swabs (Al-
lender et al. 2009). Explanations for the low prevalence and lack of ranavirus
in the two species we studied include possible resistance to infection or ability
to clear infection in these species. Infection rates and ability to clear ranavirus
vary among amphibian species exposed to standardized virus treatments, and
also according to ranavirus isolate (Hoverman et al. 2010, 2011). Thus far, this
Northeastern Naturalist Vol. 20, No. 2
330
comparative analysis of infection rates has not been investigated in turtles or
any reptile. Given our findings, the marked declines of turtle populations, and
the fact that many turtle species are syntopic with amphibians and fish (poten-
tial hosts of ranaviruses), further investigation, including controlled laboratory
studies, is needed to determine the impact of ranaviruses on turtles.
Acknowledgments
We thank Hampden-Sydney College, the Biology Department, and the Honors Pro-
gram for providing funding and support for this research. All work in this study was
approved by the Hampden-Sydney College Animal Care and Use Committee and per-
formed under scientic collection permit 38354 from Virginia Department of Game and
Inland Fisheries. We thank Briery Creek Wildlife Management for permitting us to work
on the site, Lisa Whittington for assistance conducting laboratory tests at the University
of Georgia, and Zach Harrelson, Allen Luck, Sam Smith, and Erica Rutherford for as-
sistance in the eld.
Literature Cited
Ahne, W., M. Bremont, R.P. Hedrick, A.D. Hyatt, and R.J. Whittington. 1997. Iridovi-
ruses associated with epizootic haematopoietic necrosis (EHN) in aquaculture. World
Journal of Microbiology and Biotechnology 13:367–373.
Allender, M.C., M. Abd-Eldaim, A. Kuhns, and M. Kennedy. 2009. Absence of ranavirus
and herpesvirus in a survey of two aquatic turtle species in Illinois. Journal of Herpe-
tological Medicine and Surgery 19:16–20.
Allender, M.C., M. Abd-Eldaim, J. Schumacher, D. McRuer, L.S. Christian, and M.
Kennedy. 2011. PCR prevalence of ranavirus in free-ranging Eastern Box Turtles
(Terrapene carolina carolina) at rehabilitation centers in three southeastern US states.
Journal of Wildlife Diseases 47:759–764.
Ariel, E. 1997. Pathology and serological aspects of Bohle Iridovirus infections in six
selected water-related reptiles in north Queensland. Ph.D. Dissertation. James Cook
University of North Queensland, Australia. 176 pp.
Barnosky, A.D., N. Matzke, S. Tomiya, G.O.U. Wogan, O.U. Wogan, B. Swartz, T.B.
Quental, C. Marshall, J.L. McGuire, E.L. Lindsey, K.C. Maguire, B. Mersey, and
E.A. Ferrer. 2011. Has the Earth’s sixth mass extinction already arrived? Nature
471:51–57.
Brunner, J.L., D.M. Schock, E.W. Davidson, and J.P. Collins. 2004. Intraspecic res-
ervoirs: Complex life history and the persistence of a lethal ranavirus. Ecology
85:560–566.
Chen, Z.X., J.C. Zheng, and Y.L. Jiang. 1999. A new iridovirus isolated from soft-shelled
turtle. Virus Research 63:147–151.
Chinchar, V.G. 2002. Ranaviruses (family Iridoviridae): Emerging cold-blooded killers.
Archives of Virology 147:447–470.
Daszak, P.L., A.A. Berger, A.D. Cunningham, A.D. Hyatt, D.E. Green, and R. Speare.
1999. Emerging infectious diseases and amphibian population declines. Emerging
Infectious Diseases 5:735–748.
De Voe, R., K. Geissler, S. Elmore, D. Rotstein, G. Lewbart, and J. Guy. 2004. Ranavirus-
associated morbidity and mortality in a group of captive Eastern Box Turtles (Terra-
pene carolina carolina). Journal of Zoo and Wildlife Medicine 35:534–543.
R.M. Goodman, D.L. Miller, and Y.T. Ararso2013 331
Duffus, A. 2009. Chytrid blinders: What other disease risks to amphibians are we miss-
ing? EcoHealth 6:335–339.
Ernst, C.H., and J. Lovich. 2009. Turtles of the United States and Canada. 2nd Edition.
Johns Hopkins University Press, Baltimore, MD. 840 pp.
Gibbons, J.W., D.E. Scott, T.J. Ryan, K.A. Buhlmann, T.D. Tuberville, B.S. Metts, J.L.
Greene, T. Mills, Y. Leiden, S. Poppy, and C.T. Winne. 2000. The global decline of
reptiles, déjà vu amphibians. BioScience 50:653–666.
Gray, M.J., D.L. Miller, A.C. Schmutzer, and C.A. Baldwin. 2007. Frog virus 3 preva-
lence in tadpole populations inhabiting cattle-access and non-access wetlands in Ten-
nessee, USA. Diseases of Aquatic Organisms 77:97–103.
Gray, M.J., D.L. Miller, and J.T. Hoverman. 2009. Ecology and pathology of amphibian
ranaviruses. Diseases of Aquatic Organisms 87:245–266.
Gray, M.J., D.L. Miller, and J.T. Hoverman. 2012. Reliability of non-lethal surveillance
methods for detecting ranavirus infection. Diseases of Aquatic Organisms 99:1–6.
Hoverman J.T., M.J. Gray, and D. L. Miller. 2010. Anuran susceptibilities to ranaviruses:
Role of species identity, exposure route, and a novel virus isolate. Diseases of Aquatic
Organisms 89:97–107.
Hoverman, J.T., M.J. Gray, N.A. Haislip, and D.L. Miller. 2011. Phylogeny, life history,
and ecology contribute to differences in amphibian susceptibility to ranaviruses. Eco-
Health 8:301–319.
Hyatt, A., 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 iden-
tication of a ranavirus from Green Pythons (Chondropython viridis). Journal of
Wildlife Diseases 38:239–252.
International Union for Conservation of Nature (IUCN), Conservation International, and
NatureServe. 2008. Global Amphibian Assessment. Available online at http://www.
globalamphibians.org. Accessed 1 October 2011.
IUCN. 2010. IUCN Red list of threatened species, version 2010. Available online at
http://www.iucnredlist.org. Accessed 1 October 2011.
Johnson, A.J , A.P. Pessier, and E.R. Jacobson. 2007. Experimental transmission and
induction of ranaviral disease in Western Ornate Box Turtles (Terrapene ornata
ornata) and Red-Eared Sliders (Trachemys scripta elegans). Veterinary Pathology
44:285–297.
Johnson, A.J., A.P. Pessier, and J.F. Wellehan. 2008. Ranavirus infection of free-ranging
and captive box turtles and tortoises in the United States. Journal of Wildlife Diseases
44:851–863.
Mao J., T.N. Tham, G.A. Gentry, A. Aubertin, and V.G. Chinchar. 1996. Cloning, se-
quence analysis, and expression of the major capsid protein of the iridovirus frog
virus 3. Virology 216:431–436.
Mao, J., R.P. Hedrick, and V.G. Chinchar. 1997. Molecular characterization, sequence
analysis, and taxonomic position of newly isolated sh iridoviruses. Virology
229:212–220.
Marschang, R.E., P. Becher, H. Posthaus, P. Wild, H.J. Thiel, U. Müller-Doblies, E.F.
Kalet, and L.N. Bacciarini. 1999. Isolation and characterization of an iridovirus from
Hermann’s Tortoises (Testudo hermanni). Archives of Virology 144:1909–1922.
Marschang, R.E., S. Braun, and P. Becher. 2011. Isolation of a ranavirus from a gecko
(Uroplatus mbriatus). Journal of Zoo and Wildlife Medicine 36:295–300.
Miller, D.L., and M.J. Gray. 2010. Amphibian decline and mass mortality: The value of
visualizing ranavirus in tissue sections. The Veterinary Journal 186:133–134.
Northeastern Naturalist Vol. 20, No. 2
332
Reading, C.J., L.M. Luiselli, G.C. Akani, X. Bonnet, G. Amori, J.M. Ballouard, E.
Filippi, G. Naulleau, D. Pearson, and L. Rugiero. 2010. Are snake populations in
widespread decline? Biology Letters 6:777–780.
Wake, D.B., and V.T. Vredenburg. 2008. Are we in the midst of the sixth mass extinc-
tion? A view from the world of amphibians. Proceedings of the National Academy of
Sciences of the United States of America 105:11467–11473.
Wells, K.D. 2007. The Ecology and Behavior of Amphibians. University of Chicago
Press, Chicago, IL. 1400 pp.
Westhouse, R., E.R. Jacobson, R.K. Harris, K.R. Winter, and B.L. Homer. 1996. Respi-
ratory and pharyngo-esophageal iridovirus infection in a Gopher Tortoise (Gopherus
polyphemus). Journal of Wildlife Diseases 32:682–686.
Whittington, R.J., J.A. Becker, and M.M. Dennis. 2010. Iridovirus infections in nsh:
Critical review with emphasis on ranaviruses. Journal of Fish Diseases 33:95–122.
Wilcove, D.S., D. Rothstein, J. Dubow, A. Phillips, and E. Losos. 1998. Quantifying
threats to imperiled species in the United States. Bioscience 48:607–615.
... Ranaviruses have been identified in reptiles and amphibians across the United States, often leading to mass mortality events (Gray et al. 2009;Duffus et al. 2015), but in other cases causing persistent infections in the absence of die-offs (e.g., Johnson et al. 2008;Greer et al. 2009). While ranavirus infections have been documented in the Southeast, Midwest, and numerous areas across the United States (Allender et al. 2011;Gray et al. 2012;Goodman et al. 2013;Duffus et al. 2015), infections have not been detected in Arkansas. ...
... Ranaviruses have been identified in reptiles and amphibians across the United States, often leading to mass mortality events (Gray et al. 2009;Duffus et al. 2015), but in other cases causing persistent infections in the absence of die-offs (e.g., Johnson et al. 2008;Greer et al. 2009). While ranavirus infections have been documented in the Southeast, Midwest, and numerous areas across the United States (Allender et al. 2011;Gray et al. 2012;Goodman et al. 2013;Duffus et al. 2015), infections have not been detected in Arkansas. ...
... We deployed five baited hoop nets (diameter 1 m, mesh 2.5 cm) on 10 October 2015 at a single site in WNWR (Fig. 1) and checked them the following day. Upon capture, we determined the species, sex, mass, and length of each individual, and checked for any clinical signs of ranavirus infection such as ocular discharge, dermatitis, or necrotic oral tissue (Allender et al. 2013). In order to assess ranavirus prevalence and species specificity, we collected a tail clip (~1.0 cm) from each turtle with a sterile razor blade, placed it in a snaptop tube (Fisherbrand®, Cat. ...
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... A second positive case was also found in 2018, in a wood turtle (Glyptemys insculpta) (Canadian Wildlife Health Cooperative blog 2018). In the USA, the majority of reported cases of ranavirus have involved the eastern box turtle (Terrapene carolina carolina) (e.g., De Voe et al., 2004;Allender et al., 2011;Winzeler et al., 2018); a study on its prevalence in Eastern painted turtles was carried out in Virginia, USA (Goodman, Miller & Ararso, 2013). ...
... Signs can appear similar to those of other infectious agents such as mycoplasma and herpesvirus infections, bacterial infection secondary to trauma, as well as non-infectious issues such as Vitamin A deficiency. Evidence also suggests that reptiles can be asymptomatic carriers of ranviruses (e.g., Goodman, Miller & Ararso, 2013;Goodman, Hargadon & Carter, 2018). ...
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Background Ontario, Canada is home to eight native species of turtles; all eight are federally listed as Species At Risk, due to anthropogenic threats. However, until recently, reports of infectious disease have been lacking. Ranavirus is seen as an emerging threat for ectotherms globally, with mass die-offs most often reported in amphibians. Ranavirus has been detected in Ontario’s amphibian populations, can be transmitted via water, and can be transmitted from amphibians to turtles. However, no studies on the prevalence of this virus in Ontario’s turtles have previously been carried out. With recent reports of two confirmed positive case of ranavirus in turtles in Ontario, a knowledge of the ecology of ranavirus in Ontario’s turtles has become even more important. This study estimates the prevalence of ranavirus in Ontario’s turtles, and investigates the hypothesis that this is a newly emergent disease. Methods Sixty-three samples were tested for ranavirus via PCR. These included a variety of turtle species, across their home range in Southern Ontario. Fifty-two of the samples originated from the liver and kidney of turtles who had succumbed to traumatic injuries after being admitted to the Ontario Turtle Conservation Centre; ten of the samples were taken from cloacal swabs, lesion swabs, or tail clips collected from live turtles showing signs of clinical disease. One of the live turtles was later euthanized for humane reasons and PCR was also carried out on the liver/kidney. Results None of the 63 samples were found to be positive for ranavirus via PCR. The zero prevalence found in this study translates into a population prevalence estimate of less than 5%, with no change in prevalence from 2014–2018. Discussion This is the first report on the prevalence of ranavirus in Ontario’s turtles, and will help build an understanding of the ecology of this virus in Ontario. Ranavirus has historically been underreported in reptiles, but there has been an increase in global reports recently, most likely due to increased awareness. A carrier state is thought to exist in reptiles which makes surveillance in the population via random sampling a viable method of detection of prevalence. The first report of ranavirus in Ontario turtles occurred in 2018. This study suggests a continued low population prevalence for the years 2014–2018, however. Ongoing surveillance is necessary, as well as investigation of the eDNA presence in waterways as compared to the PCR of resident turtles, to further understand the sensitivity of these species to ranavirus infection. The utilization of qPCR would be helpful, to better quantify any positives encountered.
... During 24 May -1 July of 2010, turtles were trapped for a ranavirus surveillance study by Goodman et al. (2013). We trapped at each site twice for one week during this period, using four Promar collapsible crab/fish traps with dual-ring entrance, a Sundeck turtle trap with a bait tower (Item #840876, Heinsohn's Country Store), and a floating turtle tunnel (Item#840460, Heinsohn's Country Store). ...
... 4.5 km from H-SC water bodies; Goodman, pers. obs.,Goodman et al. 2013). Trapping efforts during 2010 yielded several captures of Chrysemys picta picta and Sternotherus odoratus at both Chalgrove and Tadpole, indicating resident populations of both species on campus. ...
... Studies on ranavirus in turtles have included isolation of ranaviruses from natural populations and reports of deaths and declines in captive and wild species [12,18,19]; experiments have examined susceptibility and transmission of ranavirus between species and the importance of exposure dose and rearing temperature to host susceptibility and mortality [18,[20][21][22][23][24]. While ranavirus outbreaks with high mortality may result in disconcerting headlines, limited surveys have also found ranavirus infection occurring at low prevalence in populations without apparent die-offs [25][26][27]. Ranavirus is now well documented on six continents [12], but significant questions remain regarding distribution and factors influencing morbidity and mortality in wild reptile populations. Experimental studies examining the effect of environmental stressors on ranavirus infection are especially lacking in reptiles. ...
... All samples were tested in duplicate using Applied Biosystems™ StepOne Real-time PCR machine with two negative and two positive controls in each run (pure water and DNA extracted from cultured FV3 ranavirus). Samples with Ct values <30 for both runs were considered positive for ranavirus, according to standards established for this machine using known negative and positive controls from water, cultured ranavirus, and ranavirus-infected reptiles [25,26]. If Ct values from two samples for an individual were not both <30 or if one approximated 30, we ran two additional PCR reactions. ...
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Ranaviruses are an important wildlife pathogen of fish, amphibians, and reptiles. Previous studies have shown that susceptibility and severity of infection can vary with age, host species, virus strain, temperature, population density, and presence of environmental stressors. Experiments are limited with respect to interactions between this pathogen and environmental stressors in reptiles. In this study, we exposed hatchling red-eared slider turtles (Trachemys scripta elegans) to herbicide and ranavirus treatments to examine direct effects and interactions on growth, morbidity, and mortality. Turtles were assigned to one of three herbicide treatments or a control group. Turtles were exposed to atrazine, Roundup ProMax®, or Rodeo® via water bath during the first 3 weeks of the experiment. After 1 week, turtles were exposed to either a control (cell culture medium) or ranavirus-infected cell lysate via injection into the pectoral muscles. Necropsies were performed upon death or upon euthanasia after 5 weeks. Tissues were collected for histopathology and detection of ranavirus DNA via quantitative PCR. Only 57.5% of turtles exposed to ranavirus tested positive for ranaviral DNA at the time of death. Turtles exposed to ranavirus died sooner and lost more mass and carapace length, but not plastron length, than did controls. Exposure to environmentally relevant concentrations of herbicides did not impact infection rate, morbidity, or mortality of hatchling turtles due to ranavirus exposure. We also found no direct effects of herbicide or interactions with ranavirus exposure on growth or survival time. Results of this study should be interpreted in the context of the modest ranavirus infection rate achieved, the general lack of growth, and the unplanned presence of an additional pathogen in our study
... Similar to amphibians, susceptibility to ranavirus infections and manifestation of clinical signs in turtles vary depending on developmental stages (Duffus et al. 2015), species, and temperature . A study of Eastern box turtles (Terrapene carolina carolina) in the US reported a prevalence less than 5% in a population without abnormal mortality events (Allender et al. 2013), while in asymptomatic wild Eastern painted turtles (Chrysemys picta picta) the reported prevalence was between 4.8-31.6% in ponds without mortality events (Goodman et al. 2013). While ranavirus cases have been confirmed in North America, the few reports in Canada are potentially from a combination of a lack of surveillance in turtles, as ranavirus surveys are focused on amphibians, and subtle phenotypic manifestations of disease in turtles in general. ...
... Unfortunately, since the virus may be transmitted via the water between ectotherm species (Bandín and Dopazo, 2011;Brenes et al. 2014), the ongoing carrier state does presents opportunities to perpetuate the virus in the water bodies and therefore act as a reservoir for other species. While amphibians have been suggested as potential reservoir hosts for chelonians (Johnson et al. 2008), other studies suggested that reptiles can act as asymptomatic carriers of ranaviruses (Goodman et al. 2013(Goodman et al. , 2018Brenes et al. 2014). Reptiles can act as host and transmit ranavirus to amphibians, and while reptiles do not experience mortality, infected amphibian larvae can experience up to 100% mortality (Brenes et al. 2014). ...
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Ranaviruses have been associated with chelonian mortality. In Canada, the first two cases of ranavirus were detected in turtles in 2018 in Ontario, although a subsequent survey of its prevalence failed to detect additional positive cases. To confirm the prevalence of ranavirus in turtles in Ontario, we used a more sensitive method to investigate if lower level persistent infection was present in the population. Here we report results via a combination of qPCR, PCR, Sanger sequencing and genome sequencing from turtles from across Ontario, with no clinical signs of illness. We found 2 positives with high viral load and 5 positives with low viral load. Histopathology found subtle histological changes. DNA sequences identified two types of frog virus 3 (FV3), and genome sequencing identified a ranavirus similar to wild-type FV3. Our results show that the virus has been present in Ontario’s turtles as subclinical infections.
... This could be due either to the types of samples used or the detection method used. Previous studies have shown that more invasive samples, for example, tail clips, may be better for ranavirus detection (Goodman et al., 2013). ...
... 53,54 Virus has also been detected in cloacal swabs and blood of infected animals, 53,55 and there is some indication that more invasive sampling of tissues may lead to higher rates of identification of infected animals. 56 An ELISA has been described for the detection of antibodies against ranaviruses in chelonians, but is not commercially available. 57 ...
Methods for the detection of pathogens associated with respiratory disease in reptiles, including viruses, bacteria, fungi, and parasites, are constantly evolving as is the understanding of the specific roles played by various pathogens in disease processes. Some are known to be primary pathogens with high prevalence in captive reptiles, for example, serpentoviruses in pythons or mycoplasma in tortoises. Others are very commonly found in reptiles with respiratory disease but are most often considered secondary, for example, gram-negative bacteria. Detection methods as well as specific pathogens associated with upper- and lower-respiratory disease are discussed.
Chapter
Although reptiles have often been overlooked in research, information on viruses of reptiles has been growing steadily in recent decades as has our understanding of the importance of these animals in the ecosystem. As ectotherms, their immune systems are dependent on temperature, among other factors, and interactions between infection and disease are complex and dependent on host, pathogen, and environmental factors. This chapter provides an overview of the viruses described in reptiles so far, as well as insight into some of the diseases caused by viruses in this group of animals. It also discusses the reptile immune system and the host reaction to infection. Influences of the environment on development of disease are in many cases not well understood, and this chapter includes a discussion of some important progress in this field. Studies of the effects of viruses on wild, pet, and farmed reptiles are limited, but indicate that viral disease can strongly affect individual populations in the wild, and that human action and the animal trade likely play a role in disease epidemiology.
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