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 cornied 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.
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 - firstname.lastname@example.org.
Northeastern Naturalist Vol. 20, No. 2
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).
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
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
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.
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.
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
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.
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 scientic 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-
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