Conservation Genetics Resources

, Volume 10, Issue 4, pp 631–633 | Cite as

Development, validation, and evaluation of an assay for the detection of wood frogs (Rana sylvatica) in environmental DNA

  • Mark A. SpanglerEmail author
  • Falk Huettmann
  • Ian C. Herriott
  • J. Andrés López
Open Access
Technical Note


We developed and describe a qPCR assay for the detection of wood frogs (Rana sylvatica) using environmental DNA (eDNA) sampling. A single primer set was designed to amplify a 115-bp region of the wood frog cytochrome B gene and assessed for target specificity. There was no evidence of amplification in 11 non-target species. We evaluated the utility of the primer set in qPCR assay by conducting geo-referenced eDNA field surveys in Interior Alaska. Results indicate that the assay consistently detects wood frog DNA in the environment to 1.83 × 10−3 pg/μL. The assay provides a complement to traditional survey methods and can be readily applied in a wider conservation and management context.


eDNA Wood frog Rana sylvatica Cytochrome B Alaska 

Wood frogs (Rana sylvatica, ITIS TSN: 7751171) are widely distributed across North America. Northern wood frogs are a sentinel species for amphibian response to climate change and land development (Benard 2015; Davenport et al. 2016; Winter et al. 2016). A species of greatest conservation need in Alaska, increased monitoring and research efforts are needed (Fields and Gotthardt 2009). Wood frog distribution in Alaska is not well-defined (Online Resource 1), nor are state population trends well-known (Anderson 2004; Gotthardt et al. 2014). Wood frog monitoring efforts in the north are complicated due to challenges associated with surveying large expanses of uninhabited wilderness. Further, an abbreviated aquatic breeding period limits acoustic survey opportunities.

Environmental DNA (eDNA) monitoring refers to the detection of trace macro-organismal DNA in the environment, most often water, soil, or feces (Bohmann et al. 2014). It is increasingly being used in quantitative surveys of aquatic ecosystems (Thomsen and Willerslev 2015). eDNA assays often provide improved detectability over traditional survey methods, but they also pose unique challenges including non-standardized protocols, PCR inhibition, and environmental influences on DNA degradation rates (Olson et al. 2012; Bohmann et al. 2014; Biggs et al. 2015). Further, the use of eDNA techniques for the detection of semi-aquatic species in ephemeral wetlands (e.g. wood frogs) has not been extensively assessed (McKee et al. 2015a). Conditions in ephemeral wetlands may be unfavorable for preservation and detection of DNA due to elevated temperatures, high sediment load, and high acidity contrasted with lakes and streams (Dejean et al. 2011; Barnes et al. 2014; Eichmiller et al. 2014). Here, we report the design and validation of a qPCR assay for detection of wood frogs in eDNA at the northern extent of the species’ range.

We designed a species-specific primer set to target the cytochrome B gene of the wood frog mitochondrial genome. Sequences from the western clade of wood frogs were obtained from GenBank (PopSet 166030264, Lee-Yaw et al. 2008). The Rasy_00 primer set (Rasy_00_F: TCCTTCATCAAACAGGATCATCTA, Rasy_00_R: CCTAGTATAATGGTGAAGCCGAAT) was developed using Primer3Plus and tested for specificity in silico using NCBI Primer-BLAST (Online Resources 2 and 3). We tested Rasy_00 in vitro to ensure positive amplification of six high-quality wood frog genomic DNA isolates, as well as 500 mL of eDNA filtrate obtained from the aquarium of a live individual (Alaska Department of Fish and Game fish resource permit #SF2016-029; Online Resource 4). No other amphibians co-occur at the northern range of the wood frog, though to assess specificity in vitro we tested Rasy_00 against genomic isolates from closely related and/or co-occurring aquatic species using the qPCR assay described below. Rasy_00 consistently amplified wood frog DNA with 100% specificity (Table 1).

Table 1

Primer Rasy_00 species specificity, as tested in vitro using genomic DNA extracts

Species + ITIS TSN


Wood frog (Rana sylvatica, 775117)


Columbia spotted frog (Rana luteiventris, 550546)

American bullfrog (Rana catesbeiana, 775084)

Northern leopard frog (Rana pipiens, 775108)

Rough-skinned newt (Taricha granulosa, 173620)

Arctic Grayling (Thymallus arcticus, 162016)

Least cisco (Coregonus sardinella, 161938)

Sockeye salmon (Oncorhynchus nerka, 161979)

Northern pike (Esox lucius, 162139)

Alaska blackfish (Dallia pectoralis, 162159)

Arctic lamprey (Lethenteron camtschaticum, 622287)

Slimy sculpin (Cottus cognatus, 167232)

We collected 1 L water samples from 60 wetlands near Fairbanks, AK throughout the breeding season to assess field performance of the eDNA assay (Online Resource 5). Opportunistic visual and acoustic observations were recorded at each site. Water samples (n = 155) were kept cool and dark and filtered within 24 h of collection. We vacuum filtered water samples through 0.45 μm cellulose-nitrate membranes until they became clogged (0.1–1 L). Each batch of sample filtrations included a filter blank of distilled water (n = 18). Filter membranes were preserved at − 80 °C for less than 6 months until DNA isolation.

We isolated total genomic eDNA from filter membranes using a modified phenol–chloroform protocol (Renshaw et al. 2015; Each batch of DNA isolations included a negative control with no filter membrane (n = 10). All pre-PCR work was conducted in a PCR-free building. DNA isolates were used as templates in a qPCR assay with the Rasy_00 primers. All qPCRs were conducted in replicate (4×) on an Applied Biosystems 7900HT Sequence Detection System. PCR conditions were as follows for 20 μL reactions: 10 μL 2× KAPA SYBR Universal MasterMix, 0.4 μL 10 μM each primer, 0.4 μL 50× ROX dye, 1.25 μL 100% DMSO, and 5 μL template DNA (diluted 1:200, as determined by serial dilution). Thermal cycling conditions were 1× (94 °C/4 min), 40× (94 °C/30 s, 55 °C/45 s, 72 °C/45 s) and a melt-curve analysis of 1× [94 °C/15 s, 55 °C/15 s, 94 °C/15 s (2% ramp rate)]. Results were scored as the number of positive replicates. Stochastic variation among replicates was observed due to low eDNA concentrations. A relaxed interpretation (qPCR score = 1) risks false positive detection resulting from sample contamination, necessitating a cutoff score ≥ 2 to confidently infer species presence (Table 2). All sites with visual/acoustic detection (n = 13) had scores ≥ 2. Non-target amplifications resulting from primer-dimer artefacts were produced in the absence of template molecules in both negative control and unknown samples. They were excluded from the results through melt-curve analysis (Gudnason et al. 2007). A subset (n = 8) of positive samples were confirmed as wood frog DNA via Sanger sequencing (GenBank accession #MG002391–MG002398). The limit of detection for the qPCR assay was assessed on a Qubit 2.0 Fluorometer using a dilution series of DNA extracted from wood frog liver tissue (UAM:Herp:122) at 1.83 × 10−3 pg/μL.

Table 2

qPCR results of wood frog eDNA field surveys


Successful qPCR replicates (n of 4)






Sites (n = 60)






Negatives (n = 28)

 Filter blanks






 Isolate blanks






Raw data available via Dryad,

Our findings suggest eDNA detection is a viable survey method for semi-aquatic species in ephemeral wetlands. The assay described here may be improved by substituting DNA template dilution with a pre-PCR column-based purification step to reduce assay variance (McKee et al. 2015b). The widespread use of this assay can provide baseline northern wood frog occurrence data for use in spatial analyses (Spangler et al. unpublished).


  1. 1.

    ITIS considers Rana sylvatica invalid, but see Yuan et al. 2016.



Partial funding was kindly provided by the Alaska Herpetological Society, the University of Alaska Fairbanks Department of Biology and Wildlife (Calvin J. Lensink Graduate Fellowship in Wildlife Biology), and the University of Alaska Fairbanks Institute of Arctic Biology (IAB Summer Graduate Research Fellowship). We thank J. Ream and A. Wenninger for providing assistance in collecting eDNA samples from captive amphibians, G. Zedda for laboratory assistance, and A. Matter for guidance with eDNA techniques. The University of Alaska Museum of the North provided tissue and DNA samples. B. Barnes and K. Kielland assisted with field site selection and access.

Supplementary material

12686_2017_881_MOESM1_ESM.pdf (324 kb)
Supplementary material 1 (PDF 324 KB)


  1. Anderson BC (2004) An opportunistic amphibian inventory in Alaska’s national parks 2001–2003. Anchorage. National Park Service, Inventory and Monitoring Program, AlaskaGoogle Scholar
  2. Barnes MA, Turner CR, Jerde CL, Renshaw MA, Chadderton WL, Lodge DM (2014) Environmental conditions influence eDNA persistence in aquatic systems. Environ Sci Technol 48(3):1819–1827CrossRefGoogle Scholar
  3. Benard MF (2015) Warmer winters reduce frog fecundity and shift breeding phenology, which consequently alters larval development and metamorphic timing. Glob Chang Biol 21.3:1058–1065CrossRefGoogle Scholar
  4. Biggs J, Ewald N, Valentini A et al (2015) Using eDNA to develop a national citizen science-based monitoring programme for the great crested newt (Triturus cristatus). Biol Conserv 183:19–28CrossRefGoogle Scholar
  5. Bohmann K, Evans A, Gilbert MTP, Carvalho GR, Creer S, Knapp M, Yu DW, De Bruyn M (2014) Environmental DNA for wildlife biology and biodiversity monitoring. Trends Ecol Evol 29(6):358–367CrossRefGoogle Scholar
  6. Davenport JM, Hossack BR, Fishback L (2016) Additive impacts of experimental climate change increase risk to an ectotherm at the Arctic’s edge. Glob Chang Biol. doi: 10.1111/gcb.13543 CrossRefPubMedPubMedCentralGoogle Scholar
  7. Dejean T, Valentini A, Duparc A, Pellier-Cuit S, Pompanon F, Taberlet P, Miaud C (2011) Persistence of environmental DNA in freshwater ecosystems. PLoS ONE 6(8):e23398. doi: 10.1371/journal.pone.0023398 CrossRefPubMedPubMedCentralGoogle Scholar
  8. Eichmiller JJ, Bajer PG, Sorensen PW (2014) The relationship between the distribution of common carp and their environmental DNA in a small lake. PLoS ONE 9(11):e112611. doi: 10.1371/journal.pone.0112611 CrossRefPubMedPubMedCentralGoogle Scholar
  9. Fields T, Gotthardt T (2009) The Alaska species ranking system: setting priorities for wildlife conservation. Final report. Prepared for the Alaska Department of Fish and Game, Nongame Program. Alaska Natural Heritage Program, Environment and Natural Resources Institute, University of Alaska Anchorage, Anchorage, AK. Accessed 7 July 2016
  10. Gotthardt T, Pyare S, Huettmann F, Walton K, Spathelf M, Nesvacil K, Baltensperger A, Humphries G, Fields T (2014) Predicting the range and distribution of terrestrial vertebrate species in Alaska. The Alaska Gap Analysis Project, University of Alaska, Alaska, pp 1–40Google Scholar
  11. Gudnason H, Dufva M, Bang DD, Wolff A (2007) Comparison of multiple DNA dyes for real-time PCR: effects of dye concentration and sequence composition on DNA amplification and melting temperature. Nucleic Acids Res 35(19):e127CrossRefGoogle Scholar
  12. Lee-Yaw JA, Irwin JT, Green DM (2008) Postglacial range expansion from northern refugia by the wood frog, Rana sylvatica. Mol Ecol 17(3):867–884CrossRefGoogle Scholar
  13. McKee AM, Calhoun DL, Barichivich WJ, Spear SF, Goldberg CS, Glenn TC (2015a) Assessment of environmental DNA for detecting presence of imperiled aquatic amphibian species in isolated wetlands. J Fish Wildl Manag 6(2):498–510CrossRefGoogle Scholar
  14. McKee AM, Spear SF, Pierson TW (2015b) The effect of dilution and the use of a post-extraction nucleic acid purification column on the accuracy, precision, and inhibition of environmental DNA samples. Biol Conserv 183:70–76CrossRefGoogle Scholar
  15. Olson ZH, Briggler JT, Williams RN (2012) An eDNA approach to detect eastern hellbenders (Cryptobranchus alleganiensis) using samples of water. Wild Res 39(7):629–636CrossRefGoogle Scholar
  16. Renshaw MA, Olds BP, Jerde CL, McVeigh MM, Lodge DM (2015) The room temperature preservation of filtered environmental DNA samples and assimilation into a phenol–chloroform–isoamyl alcohol DNA extraction. Mol Ecol Res 15(1):168–176CrossRefGoogle Scholar
  17. Thomsen PF, Willerslev E (2015) Environmental DNA: an emerging tool in conservation for monitoring past and present biodiversity. Biol Conserv 183:4–18CrossRefGoogle Scholar
  18. Winter M, Fiedler W, Hochachka WM, Koehncke A, Meiri S, De la Riva I (2016) Patterns and biases in climate change research on amphibians and reptiles: a systematic review. R Soc Open Sci 3(9):160158CrossRefGoogle Scholar
  19. Yuan ZY, Zhou WW, Chen X et al (2016) Spatiotemporal diversification of the true frogs (Genus Rana): a historical framework for a widely studied group of model organisms. Syst Biol 65(5):824–842CrossRefGoogle Scholar

Copyright information

© The Author(s) 2017

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.

Authors and Affiliations

  • Mark A. Spangler
    • 1
    • 2
    Email author
  • Falk Huettmann
    • 1
  • Ian C. Herriott
    • 3
  • J. Andrés López
    • 2
    • 4
  1. 1.EWHALE Lab - Institute of Arctic Biology, Biology and Wildlife DepartmentUniversity of Alaska Fairbanks (UAF)FairbanksUSA
  2. 2.University of Alaska Museum of the North (UAM)FairbanksUSA
  3. 3.DNA Core Facility, Institute of Arctic BiologyUniversity of Alaska Fairbanks (UAF)FairbanksUSA
  4. 4.Fisheries Division, College of Fisheries and Ocean SciencesUniversity of Alaska Fairbanks (UAF)FairbanksUSA

Personalised recommendations