The fungal pathogen Ophidiomyces ophiodiicola, the causative agent of snake fungal disease, has been implicated in declines of North American snake populations since 2006 and the geographic range of this pathogen is still not fully known. In Tennessee, US, O. ophiodiicola has been detected since 2012, but large portions of the state have not been surveyed for this pathogen. Our primary objective was to monitor the prevalence of O. ophiodiicola in the Interior Plateau ecoregion of Tennessee by swabbing all snakes that were encountered during road cruising survey efforts in 2017 and 2018. Eleven snakes of four species, copperhead (Agkistrodon contortrix), common water snake (Nerodia sipedon), black kingsnake (Lampropeltis nigra), and smooth earthsnake (Virginia valeriae), tested positive for the presence of O. ophiodiicola. Overall, 9.2% (11/120) of snakes sampled tested positive for the presence of O. ophiodiiola, and we further observed a seasonal trend in detections with summer months having the greatest frequency of detections. Our results extend the known geographic range of O. ophiodiicola in Tennessee by adding four previously unconfirmed O. ophiodiicola-positive counties. Further sampling will need to be conducted across west Tennessee because this is the most data-deficient region of the state. Our results offer additional evidence of the presence of this pathogen in Tennessee and will help researchers further understand the geographic distribution and host range.

Emerging pathogen outbreaks have occurred at alarming rates in various wildlife populations around the world and have been implicated as one of the primary causes of global biodiversity loss (Daszak et al. 2000; Hof et al. 2011). The fungal pathogen Ophidiomyces ophiodiicola, since its discovery in 2000 (Allender et al. 2016), represents the primary causative agent of snake fungal disease (Lorch et al. 2016). Severe infections of O. ophiodiicola result regularly in mortality of infected snakes (Allender et al. 2011; Dolinski et al. 2014); however, sublethal effects often manifest as behavioral changes that include increased molting and basking frequency (Clark et al. 2011; Lorch et al. 2015; Tetzlaff et al. 2017) that can make snakes more susceptible to predation or other sources of mortality. The role this pathogen plays in snake population declines is still relatively unknown (Allender et al. 2015) and additional prevalence data for this pathogen will benefit wildlife managers to better understand potential threats to snake conservation.

Ophidiomyces ophiodiicola has been detected in at least 23 states within the US. Detections have been confirmed in 33 snake species that comprise six snake families (Lorch et al. 2016; Thompson et al. 2018). Ophidiomyces ophiodiicola was first confirmed in Tennessee in 2012 on a timber rattlesnake (Crotalus horridus) and has since been detected on 15 snake species across 16 counties (Grisnik et al. 2018). However, prevalence sampling for this pathogen in Tennessee has not spanned the entire state or across all native snake species. To date, there has been little to no sampling in the western portions of Tennessee, specifically in the Interior Plateau ecoregion.

Our objective was to assess the prevalence of O. ophiodiicola in free-ranging snake populations in the Interior Plateau ecoregion of Tennessee. Results from our study can be used to help determine the prevalence of O. ophiodiicola in Tennessee and confirm which snake species might be at greatest risk from this pathogen. Ultimately, our results can be used to identify potential areas of conservation concern and implement action plans to reduce snake population losses caused by this pathogen. This research was approved under Institutional Animal Care and Use Committee #1803SS at Tennessee State University (Nashville, Tennessee, USA).

During the 2017 and 2018 field seasons (May–October), we conducted 43 road transect surveys, 30 of which were repeatedly surveyed three times (early summer [10 May–30 June], midsummer [25 June–10 August], and late summer [11 August–30 October]). We conducted road transect surveys shortly after dusk. We surveyed each transect (about 16 km) via a double pass approach (i.e., an out-and-back approach; Sutherland 2009). After each snake was identified to species, we collected the following descriptive data: sex (via probing), mass (g), individual mark via a passive integrated transponder tag (Bio-mark, version 601, Boise, Idaho, USA), and snout-vent length (mm). We released each sampled snake at the site of capture.

We swabbed all snakes encountered (i.e., living and recently deceased) during road cruising surveys and from incidental encounters. We used 15.2 cm Puritan Sterile Cotton Tipped Applicators (Puritan Medical Products Company LLC, Guilford, Maine, USA) to swab captured snakes, using 10 strokes (Hileman et al. 2018). We wore nitrile gloves during each snake capture and changed gloves after each snake was swabbed. We stored swab samples in sterile 1.5 mL microcentrifuge tubes filled with 1 mL of 95% ethanol and stored vials at room temperature after collection and then at –20 C until subsequent DNA extraction. We restrained venomous snakes in sterilized acrylic snake tubes and only swabbed the portion of the body not inside the acrylic tube. We washed tubes with 10% bleach solution after each snake capture to reduce contamination between samples. All snakes were visually assessed for clinical signs of skin lesions and recorded if present.

We followed the protocols used in Grisnik et al. (2018) for DNA extraction. Specifically, we used the Qiagen DNeasy Powersoil 96 HTP kit (Qiagen, Hilden, Germany) following the manufacturer's recommended protocol to extract genomic DNA from swab tips. We detected the molecular presence of O. ophiodiicola via the quantitative PCR assay described in Bohuski et al. (2015) by amplifying the internal transcribed spacer region. We performed 10 µL reactions that consisted of 5 µL of 2× Quantabio PerfecCTa quantitative PCR ToughMix (Quantabio, Beverly, Massachusetts, USA), 0.4 µL of IDT forward primer (10 µM), 0.4 µL of IDT reverse primer (10 µM), 0.1 µL IDT probe (20 µM), 2.1 µL PCR grade water, and 2 µL of DNA template. Thermal cycling conditions consisted of 3 min at 95 C, followed by 40 cycles of 95 C for 10 s, and 60 C for 30 s. We included a DNA extraction negative control blank on each plate. In addition, we included a no-template negative control blank on each 96-well plate that was run in triplicate, and three positive control reactions of 0.5 ng/µL of O. ophiodiicola DNA. We considered a snake to be positive for O. ophiodiicola if one or two of the three triplicate samples were positive in two consecutive runs or if all three triplicates were positive in the first run. We considered threshold cycle values of <39 cycles as confirmation of positive detections to reduce the occurrence of false positives.

We swabbed 120 individual snakes during the 2017 (n=78) and 2018 (n=42) field seasons (Table 1). We detected O. ophiodiicola on 11 snakes that represented four species (Table 2). Six copperheads (Agkistrodon contortrix), two common water snakes (Nerodia sipedon), two black kingsnakes (Lampropeltis nigra), and one smooth earthsnake (Virginia valeriae) accounted for the detections (Table 2). Detections had threshold cycle values that ranged 31.47–38.87, with the lowest value recorded from a common water snake. We detected O. ophiodiicola in four counties (Benton, Decatur, Montgomery, Stewart; Fig. 1). We detected O. ophiodiicola on five snakes (three copperheads; one black kingsnake; one smooth earthsnake) in Stewart County, four snakes (two copperheads; one common water snake; one black kingsnake) in Benton County, one snake (a copperhead) in Decatur County, and one snake (a common water snake) in Montgomery County (Tables 2, 3). We detected a seasonal trend after correcting for survey effort per month, with the majority of positive detections (64%; 7/11) detected during summer (June–August) compared to spring (9%; 1/11) and fall (27%; 3/11; Fig. 2). Further, none of the snakes swabbed had clinical signs of skin lesions during this study.

Table 1

Number of swabs obtained from each snake species during snake road-transect surveys to determine prevalence of Ophidiomyces ophiodiicola in the Interior Plateau ecoregion of Tennessee, USA.

Number of swabs obtained from each snake species during snake road-transect surveys to determine prevalence of Ophidiomyces ophiodiicola in the Interior Plateau ecoregion of Tennessee, USA.
Number of swabs obtained from each snake species during snake road-transect surveys to determine prevalence of Ophidiomyces ophiodiicola in the Interior Plateau ecoregion of Tennessee, USA.
Table 2

Locality and individual data for snakes with positive cases of Ophidiomyces ophiodiicola sampled during snake road-transect surveys in the Interior Plateau ecoregion of Tennessee, USA.

Locality and individual data for snakes with positive cases of Ophidiomyces ophiodiicola sampled during snake road-transect surveys in the Interior Plateau ecoregion of Tennessee, USA.
Locality and individual data for snakes with positive cases of Ophidiomyces ophiodiicola sampled during snake road-transect surveys in the Interior Plateau ecoregion of Tennessee, USA.
Figure 1

Counties in Tennessee, USA, previously confirmed (gray shaded counties; Grisnik et al. 2018), confirmed in this study (black dots), and unconfirmed in this study (white and gray diagonal lines) to have positive Ophidiomyces ophiodiicola detections.

Figure 1

Counties in Tennessee, USA, previously confirmed (gray shaded counties; Grisnik et al. 2018), confirmed in this study (black dots), and unconfirmed in this study (white and gray diagonal lines) to have positive Ophidiomyces ophiodiicola detections.

Close modal
Table 3

Number of swabs collected from individual snakes in each county surveyed, the number of individual species sampled in each county, and the number of positive cases of Ophidiomyces ophiodiicola in each county during snake road-transect surveys in the Interior Plateau ecoregion of Tennessee, USA.

Number of swabs collected from individual snakes in each county surveyed, the number of individual species sampled in each county, and the number of positive cases of Ophidiomyces ophiodiicola in each county during snake road-transect surveys in the Interior Plateau ecoregion of Tennessee, USA.
Number of swabs collected from individual snakes in each county surveyed, the number of individual species sampled in each county, and the number of positive cases of Ophidiomyces ophiodiicola in each county during snake road-transect surveys in the Interior Plateau ecoregion of Tennessee, USA.
Figure 2

Percent positive cases of Ophidiomyces ophiodiicola per month (total detections per month divided by the total snakes sampled per month) for 2017 and 2018 field seasons. The number of snakes sampled per month was: April, n=2; May, n=7; June, n=23; July, n=9; August, n=21; September, n=38; October, n=20.

Figure 2

Percent positive cases of Ophidiomyces ophiodiicola per month (total detections per month divided by the total snakes sampled per month) for 2017 and 2018 field seasons. The number of snakes sampled per month was: April, n=2; May, n=7; June, n=23; July, n=9; August, n=21; September, n=38; October, n=20.

Close modal

Species in the family Viperidae have been reported with relatively high prevalence of infection (Clark et al. 2011; McBride et al. 2015; Tetzlaff et al. 2015) and we detected a similar trend with copperhead snakes having the greatest O. ophiodiicola prevalence. High prevalence of O. ophiodiicola has been detected in water snakes (Nerodia spp.), with various cases across Tennessee (Grisnik et al. 2018), and our results further confirm this pattern. Recent research has suggested that the greatest frequency of infections in wild populations occurs in summer months (i.e., June, July, August, and September; Lorch et al. 2016; McKenzie et al. 2019). We observed a similar trend, with relatively greater prevalence of O. ophiodiicola during July (22%; 2/9 of sampled snakes). Further, we observed a general increase in the percent of positive cases from May to July (i.e., May, 14% [1/7]; June, 17% [4/23]; July, 22% [2/9]), compared to the period of August through October, despite having a fewer number of sampled snakes (n=39; Fig. 2).

Grisnik et al. (2018) found that 30% of snakes sampled in Tennessee tested positive for the presence of O. ophiodiicola, which is substantially greater than the prevalence (9.2%) that we reported. This disparity could be due to bias toward snakes that were detected during road cruising surveys, and differences between ecoregions in east Tennessee (Grisnik et al. 2018) and the Interior Plateau ecoregion evaluated during the current study. Further sampling will need to be conducted across West Tennessee in the Southeastern Plains and the Mississippi Valley Loess Plains ecoregions because these are the most data-deficient ecoregions in Tennessee, and they also have high snake diversity compared to other ecoregions in the state (Niemiller et al. 2013). Our results provide further evidence of O. ophiodiicola prevalence in Tennessee and will help researchers develop action plans to mitigate snake population losses due to this pathogen. As impacts of climate change increase, O. ophiodiicola is predicted to become a greater threat for snake conservation (Allender et al. 2015), which makes consistent monitoring for O. ophiodiicola prevalence a conservation priority.

We thank the Tennessee Wildlife Resources Agency and the Tennessee Valley Authority for providing funding, field site access, and the necessary permits to complete this research. We also thank the Wildlife Ecology Laboratory and the Department of Agricultural and Environmental Sciences at Tennessee State University for field assistance and logistical support, respectively. We thank two anonymous reviewers for comments on earlier versions of the manuscript.

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