Abstract
In the past decade, quantification of environmental DNA (eDNA) has become firmly established as an effective method for detecting the presence of organisms of research and conservation interest. Salamanders of the family Ambystomatidae are large, fossorial species; adults are rarely encountered above ground outside of their brief reproductive season. Larvae develop rapidly, in ephemeral pools or streams, often with multiple species coexisting in a single habitat. We developed species-specific PCR assays for Ambystoma barbouri, Ambystoma jeffersonianum, Ambystoma opacum, Ambystoma maculatum, and Ambystoma tigrinum based on locally sequenced specimens and tested each in silico and in vitro as well as conducted field surveys, using both end-point and quantitative PCR to test all in situ except A. tigrinum. In silico and in vitro tests confirm specificity of each primer. In situ larvae surveys were conducted and water samples collected from known Ambystoma breeding sites in central and eastern Kentucky, USA. Larvae of one or more species were observed at 12/13 sites. A total of 14 separate species sightings were observed, and in each case, eDNA from the observed species was detected via qPCR. Additionally, nine qPCR detections were observed at sites where species were not field observed. End-point PCR results were less effective in detecting field observed species, and primers appeared to bind to DNA from non-salamander targets. These primers, when used with a probe in a qPCR assay, provide an effective means of determining species presence rapidly and definitively and therefore offer to increase the ease of range delineation and spawning habitat studies.
Similar content being viewed by others
References
Altig R, McDiarmid RW, Bauer AM (2017) Handbook of larval amphibians of the United States and Canada. Cornell University Press, Ithaca
Anderson MA, Campbell JR, Carey AN et al (2014) Population survey of the streamside salamander in the Nashville Basin of Tennessee. Southeast Nat 13:101–107. https://doi.org/10.1656/058.013.0108
Anderson T, Ousterhout BH, Peterman WE et al (2015) Life history differences influence the impacts of drought on two pond-breeding salamanders. Ecol Appl 25:1896–1910. https://doi.org/10.1890/14-2096.1
Baldigo BP, Sporn LA, George SD, Ball JA (2017) Efficacy of environmental DNA to detect and quantify brook trout populations in headwater streams of the Adirondack Mountains, New York. Trans Am Fish Soc 146:99–111. https://doi.org/10.1080/00028487.2016.1243578
Bell FF, Flores AF, Sena KL et al (2022) Development and validation of qPCR assays for use in eDNA detection of southern two-lined (Eurycea cirrigera) and northern dusky (Desmognathus fuscus) salamanders. Herpetol Conserv Biol 17:398–412
Bi K, Bogart JP (2010) Time and time again: unisexual salamanders (genus Ambystoma) are the oldest unisexual vertebrates. BMC Evol Biol 10:1–14. https://doi.org/10.1186/1471-2148-10-238
Bogart JP, Lowcock LA, Zeyl CW, Mable BK (1987) Genome constitution and reproductive biology of hybrid salamanders, genus Ambystoma, on Kelleys Island in Lake Erie. Can J Zool 65:2188–2201. https://doi.org/10.1139/z87-333
Brandon RA (1964) An annotated and illustrated key to multistage larvae of Ohio salamanders. Ohio J Sci 64:252
Brandon R, Bremer D (1966) Neotenic newts, Notophthalmus viridescens louisianensis, in southern Illinois. Herpetologica 22:213–217
Breton BA, Beaty L, Bennett AM et al (2022) Testing the effectiveness of environmental DNA (eDNA) to quantify larval Amphibian abundance. Environ DNA 4:1229–1240. https://doi.org/10.1002/edn3.332
Bylemans J, Gleeson DM, Duncan RP et al (2019) A performance evaluation of targeted eDNA and eDNA metabarcoding analyses for freshwater fishes. Environ DNA 1:402–414. https://doi.org/10.1002/edn3.41
Canada E (2016) Recovery Strategy for the Jefferson Salamander (Ambystoma jeffersonianum) in Canada. Species at risk act recovery strategy series. Environment Canada, Ottawa
Clarke GS, Phillips BL, Shine R (2019) Clipping the tail fin enables cohort identification of small Anuran Tadpoles. Copeia 107:71–77. https://doi.org/10.1643/CE-18-128
Crawford JA, Peterman WE, Kuhns AR, Eggert LS (2016) Altered functional connectivity and genetic diversity of a threatened salamander in an agroecosystem. Landsc Ecol 31:2231–2244. https://doi.org/10.1007/s10980-016-0394-6
Czechowski P, de Lange M, Heldsinger M et al (2021) Environmental DNA analysis needs local reference data to inform taxonomy-based conservation policy—a case study from Aotearoa/New Zealand. bioRxiv. https://doi.org/10.1101/2021.10.22.465527
DiLeo K (2016) Species status review of Amphibian and reptiles. Trenton, New Jersey
Doyle JM, Whiteman HH (2008) Paedomorphosis in Ambystoma talpoideum: effects of initial body size variation and density. Oecologia 156:87–94. https://doi.org/10.1007/s00442-008-0977-2
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 9:1–8. https://doi.org/10.1371/journal.pone.0112611
Everts T, Halfmaerten D, Neyrinck S et al (2021) Accurate detection and quantification of seasonal abundance of American bullfrog (Lithobates catesbeianus) using ddPCR eDNA assays. Sci Rep 11:11282. https://doi.org/10.1038/s41598-021-90771-w
Everts T, Van Driessche C, Neyrinck S et al (2022) Using quantitative eDNA analyses to accurately estimate American bullfrog abundance and to evaluate management efficacy. Environ DNA 4:1052–1064. https://doi.org/10.1002/edn3.301
Ficetola GF, Miaud C, Pompanon F, Taberlet P (2008) Species detection using environmental DNA from water samples. Biol Lett 4:423–425. https://doi.org/10.1098/rsbl.2008.0118
Garcia TS, Sih A (2003) Color change and color-dependent behavior in response to predation risk in the salamander sister species Ambystoma barbouri and Ambystoma texanum. Oecologia 137:131–139. https://doi.org/10.1007/s00442-003-1314-4
Goldberg CS, Pilliod DS, Arkle RS, Waits LP (2011) Molecular detection of vertebrates in stream water: a demonstration using rocky mountain tailed frogs and Idaho giant salamanders. PLoS One 6:e22746. https://doi.org/10.1371/journal.pone.0022746
Goldberg CS, Turner CR, Deiner K et al (2016) Critical considerations for the application of environmental DNA methods to detect aquatic species. Methods Ecol Evol 7:1299–1307. https://doi.org/10.1111/2041-210X.12595
Goldberg CS, Strickler KM, Fremier AK (2018) Degradation and dispersion limit environmental DNA detection of rare amphibians in wetlands: increasing efficacy of sampling designs. Sci Total Environ 633:695–703. https://doi.org/10.1016/j.scitotenv.2018.02.295
Gregory T, Mable R (2005) BK polyploidy in animals. The evolution of the genome. Academic Press, London
Guivas RA, Brammell BF (2020) Use of environmental DNA to determine fantail darter (Etheostoma flabellare) density in a laboratory setting: effects of biomass and filtration method. Int J Zool. https://doi.org/10.1155/2020/4731686
Harper LR, Lawson Handley L, Hahn C et al (2018) Needle in a haystack? a comparison of eDNA metabarcoding and targeted qPCR for detection of the great crested newt (Triturus cristatus). Ecol Evol 8:6630–6641. https://doi.org/10.1002/ece3.4013
Harper LR, Buxton AS, Rees HC et al (2019) Prospects and challenges of environmental DNA (eDNA) monitoring in freshwater ponds. Hydrobiologia 826:25–41
Hinlo R, Gleeson D, Lintermans M, Furlan E (2017) Methods to maximize recovery of environmental DNA from water samples. PLoS One 126:e0179251. https://doi.org/10.1371/journal.pone.0179251
Hobbs J, Round JM, Allison MJ, Helbing CC (2019) Expansion of the known distribution of the coastal tailed frog, Ascaphus truei, in British Columbia, Canada, using robust eDNA detection methods. PLoS One 14:e0213849. https://doi.org/10.1371/journal.pone.0213849
Hossack BR, Lemos-Espinal JA, Sigafus BH et al (2021) Distribution of tiger salamanders in Northern Sonora, Mexico: comparison of sampling methods and possible implications for an endangered subspecies. Amphibia Reptilia 43:13–23. https://doi.org/10.1163/15685381-bja10072
Hubbs NW, Hurt CR, Niedzwiecki J et al (2022) Conservation genomics of urban populations of streamside salamander (Ambystoma barbouri). PLoS One. https://doi.org/10.1371/journal.pone.0260178
Huver JR, Koprivnikar J, Johnson PTJ, Whyard S (2015) Development and application of an eDNA method to detect and quantify a pathogenic parasite in aquatic ecosystems. Ecol Appl 25:991–1002. https://doi.org/10.1890/14-1530.1
Irschick DJ, Shaffer HB (1997) The polytypic species revisited: morphological differentiation among tiger salamanders (Ambystoma tigrinum) (Amphibia: Caudata). Herpetologica 53:30–49
Jerde CL, Mahon AR, Chadderton WL, Lodge DM (2011) Sight-unseen detection of rare aquatic species using environmental DNA. Conserv Lett. https://doi.org/10.1111/j.1755-263X.2010.00158.x
Kaganer AW, Stapleton GS, Bunting EM, Hare MP (2022) Aquatic eDNA can advance monitoring of a small-bodied terrestrial salamander and Amphibian pathogen. Environ DNA. https://doi.org/10.1002/edn3.316
Kieran SR, Hull JM, Finger AJ (2020) Using environmental DNA to monitor the spatial distribution of the California tiger salamander. J Fish Wildl Manag 11:609–617
King AC, Krieg R, Weston A, Zenker AK (2022) Using eDNA to simultaneously detect the distribution of native and invasive crayfish within an entire country. J Environ Manage 302:113929. https://doi.org/10.1016/j.jenvman.2021.113929
Klymus KE, Marshall NT, Stepien CA (2017) Environmental DNA (eDNA) metabarcoding assays to detect invasive invertebrate species in the Great Lakes. PLoS One. https://doi.org/10.1371/journal.pone.0177643
Klymus KE, Merkes CM, Allison MJ et al (2020) Reporting the limits of detection and quantification for environmental DNA assays. Environ DNA 2:271–282. https://doi.org/10.1002/edn3.29
Kraus F, Petranka JW (1989) A new sibling species of Ambystoma from the Ohio River drainage. Copeia 1989:94–110. https://doi.org/10.2307/1445610
Kraus F, Shaffer HB (1991) When molecules and morphology clash: a phylogenetic analysis of the North American Ambystomatid salamanders (caudata: Ambystomatidae). Syst Biol 40:284–303. https://doi.org/10.1093/sysbio/40.3.284
Langlois VS, Allison MJ, Bergman LC et al (2020) The need for robust qPCR-based eDNA detection assays in environmental monitoring and species inventories. Environ DNA 3:519–527. https://doi.org/10.1002/edn3.164
Lodge DM (2022) Policy action needed to unlock eDNA potential. Front Ecol Environ 20:448–449. https://doi.org/10.1002/fee.2563
Mauvisseau Q, Tönges S, Andriantsoa R et al (2019) Early detection of an emerging invasive species: EDNA monitoring of a parthenogenetic crayfish in freshwater systems. Manage Biol Invasions 10:461–472. https://doi.org/10.3391/mbi.2019.10.3.04
Moritz C, Schneider CJ, Wake DB (1992) Evolutionary relationships within the Ensatina eschscholtzii complex confirm the ring species interpretation. Syst Biol 41:273–291. https://doi.org/10.1093/sysbio/41.3.273
Moss WE, Harper LR, Davis MA et al (2022) Navigating the trade-offs between environmental DNA and conventional field surveys for improved Amphibian monitoring. Ecosphere 13:e3941. https://doi.org/10.1002/ecs2.3941
Mott CL, Sparling DW (2016) Seasonal patterns of intraguild predation and size variation among larval salamanders in ephemeral ponds. J Herpetol 50:416–422. https://doi.org/10.1670/15-029
Mott CL, Steffen MA, Uzzardo JM, Wiley BK (2007) Ambystoma opacum habitat. Herpetol Rev 38:316–317
Mullin SJ, Klueh S (2009) Demographics of a geographically isolated population of a threatened salamander (Caudata: Ambystomatidae) in central Illinois. Herpetol Conserv Biol 4:161
Nathan LM, Simmons M, Wegleitner BJ et al (2014) Quantifying environmental DNA signals for aquatic invasive species across multiple detection platforms. Environ Sci Technol 48:76–83. https://doi.org/10.1021/es5034052
Olson ZH, Briggler JT, Williams RN (2012) An eDNA approach to detect eastern hellbenders (Cryptobranchus a. alleganiensis) using samples of water. Wildl Res 39:629–636. https://doi.org/10.1071/WR12114
Othman SN, Chuang MF, Kang H et al (2020) Methodological guidelines for minimally invasive tail-clipping: a case study on Rana huanrenensis tadpoles. Asian J Conserv Biol 9:188–195
Petranka JW (1998) Salamanders of the United States. Smithsonian Institution Press, Washington
Phillips CA (1992) Variation in metamorphosis in spotted salamanders Ambystoma maculatum from Eastern Missouri. Am Midl Nat 128:276–280. https://doi.org/10.2307/2426461
Phillips CA, Dreslik JM, Johnson JR, Petzing JE (2001) Application of population estimation to pond breeding salamanders. Trans Ill State Acad Sci 94:111–118
Plante F, Bourgault P, Dubois Y, Bernatchez L (2021) Environmental DNA as a detection and quantitative tool for stream-dwelling salamanders: a comparison with the traditional active search method. Environ DNA 3:1128–1141. https://doi.org/10.1002/edn3.233
Porej D, Micacchion M, Hetherington TE (2004) Core terrestrial habitat for conservation of local populations of salamanders and wood frogs in agricultural landscapes. Biol Conserv 120:399–409. https://doi.org/10.1016/j.biocon.2004.03.015
Roussel JM, Paillisson JM, Tréguier A, Petit E (2015) The downside of eDNA as a survey tool in water bodies. J Appl Ecol 52:823–826
Sever DM, Kloepeer NM (1993) Spermathecal cytology of Ambystoma opacum (Amphibia: Ambystomatidae) and the phylogeny of sperm storage organs in female salamanders. J Morphol 217:115–127. https://doi.org/10.1002/jmor.1052170110
Shollenberger KR, Janosik AM, Johnston C (2023) Detection of the threatened snail darter Percina tanasi in the Tennessee River system using environmental DNA. J Fish Biol 102:373–379. https://doi.org/10.1111/jfb.15269
Shu L, Ludwig A, Peng Z (2020) Standards for methods utilizing environmental DNA for detection of fish species. Genes (Basel) 11:296
Simmons M, Tucker A, Chadderton WL et al (2015) Active and passive environmental DNA surveillance of aquatic invasive species. Can J Fish Aquat Sci 73:76–83. https://doi.org/10.1139/cjfas-2015-0262
Stangel PW (1988) Premetamorphic survival of the salamander Ambystoma maculatum, in Eastern Massachusetts. J Herpetol 22:345–347. https://doi.org/10.2307/1564160
Takahara T, Minamoto T, Yamanaka H et al (2012) Estimation of fish biomass using environmental DNA. PLoS One. https://doi.org/10.1371/journal.pone.0035868
Tennessee Wildlife Resources Agency (2018) Rules and regulations for in need of management, threatened, and endangered species. Chap. 1660-01-32, https://publications.tnsosfiles.com/rules/1660/1660-01/1660-01-32.20180830.pdf
Thalinger B, Deiner K, Harper LR et al (2021) A validation scale to determine the readiness of environmental DNA assays for routine species monitoring. Environ DNA 3:823–836. https://doi.org/10.1002/edn3.189
Tompkins R (1978) Genie control of axolotl metamorphosis. Integr Comp Biol 18:313–319. https://doi.org/10.1093/icb/18.2.313
Turner CR, Miller DJ, Coyne KJ, Corush J (2014) Improved methods for capture, extraction, and quantitative assay of environmental DNA from asian bigheaded carp (hypophthalmichthys spp.). PLoS One 9:1–20. https://doi.org/10.1371/journal.pone.0114329
Vitt LJ, Caldwell JP (2013) Herpetology: an introductory biology of Amphibians and reptiles, 4th edn. Academic Press, London
Voss SR (1995) Genetic basis of paedomorphosis in the axolotl, Ambystoma mexicanum: a test of the single-gene hypothesis. J Hered 86:441–447. https://doi.org/10.1093/oxfordjournals.jhered.a111618
Whiteman HH (1994) Evolution of facultative paedomorphosis in Salamanders. Q Rev Biol 69:205–221. https://doi.org/10.1086/418540
Wilcox TM, McKelvey KS, Young MK et al (2013) Robust detection of rare species using environmental DNA: the importance of primer specificity. PLoS One 8:e59520. https://doi.org/10.1371/journal.pone.0059520
Williams SJ (2012) When molecules and morphology clash: revisiting species tree reconstruction of Ambystomatid Salamanders using multiple nuclear loci. University of Kentucky
Witzel NA, Taheri A, Miller BT et al (2020) Validation of an environmental DNA protocol to detect a stream-breeding amphibian, the Streamside Salamander (Ambystoma barbouri). Environ DNA 2:554–564. https://doi.org/10.1002/edn3.83
Wood SA, Pochon X, Laroche O et al (2019) A comparison of droplet digital polymerase chain reaction (PCR), quantitative PCR and metabarcoding for species-specific detection in environmental DNA. Mol Ecol Resour 19:1407–1419. https://doi.org/10.1111/1755-0998.13055
Xia Z, Zhan A, Johansson ML et al (2021) Screening marker sensitivity: optimizing eDNA-based rare species detection. Divers Distrib. https://doi.org/10.1111/ddi.13262
Acknowledgements
We thank Jarrett Johnson (A. tigrinum) and Bret Kuss (A. talpoideum and A. texanum) for the generous donation of tissue. We thank John McGregor (Kentucky Department of Fish and Wildlife Resources) for generously sharing his knowledge and providing advice. Aside from donated samples, salamander tissue was collected under Kentucky Department of Fish and Wildlife Service Permit #SC1811153 (Ben Brammell).
Funding
Research was funded by an internal faculty development grant from Asbury University (Ben Brammell, Fall 2021). The Asbury University Department of Science and Health, Shaw School of Science, provided additional support.
Author information
Authors and Affiliations
Contributions
BB served as the PI of the lab from which this research originated, he obtained funding, participated and supervised all lab work, and wrote the manuscript text, preparing all figures and tables. ES developed and tested the A. tigrinum assays and conducted additional supporting lab work. SB, RP, and CM conducted a great deal of the lab and field work for this study. CM collected the majority of the water samples and conducted all field surveys including field identifying all salamander larvae. MS and CS also contributed significant lab work for this project. All authors reviewed the manuscript.
Corresponding author
Ethics declarations
Competing interests
The authors declare no competing interests. The authors have no relevant financial or non-financial interests to disclose.
Additional information
Publisher’s Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Supplementary Information
Below is the link to the electronic supplementary material.
Rights and permissions
Springer Nature or its licensor (e.g. a society or other partner) holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law.
About this article
Cite this article
Brammell, B.F., Strasko, E.K., Brewer, S.A. et al. Detecting fossorial salamanders using eDNA: Development and validation of quantitative and end-point PCR assays for the detection of five species of Ambystoma. Conservation Genet Resour 15, 187–198 (2023). https://doi.org/10.1007/s12686-023-01322-6
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1007/s12686-023-01322-6