Two Methods for Analyzing Aquatic Environmental DNA Meredith

Two Methods for Analyzing Aquatic Environmental DNA

Meredith Bennett (Junior, Environmental Science) Faculty Advisor: Dr. Brady Porter Duquesne University

Environmental DNA (eDNA) is a collection of DNA molecules found in the environment. Environmental DNA can be present in the environment in several forms, including tissue, feces, gametes, and dead cells. 1, 2 Environmental DNA can be found in many different substrates, including soil, permafrost, and even air. 1, 3, 4 The vast majority of eDNA studies, however, relate to aquatic ecosystems. 5-8 Environmental DNA studies have been applied to a wide range of topics. Two of the more common applications of eDNA are the study of rare and elusive species 3, 4, 9 and the monitoring of invasive species. 8, 10, 11 The Hula painted frog is a rare species recently rediscovered after it was believed to be extinct. 4 Environmental DNA methods were used to locate additional populations of the frog and determine the features that classify suitable habitats of the organism. 4 As for invasive species, invertebrates in the Laurentian Great

Lakes have been identified using two new metabarcoding eDNA assays. 10 The markers detected invasive bivalve and gastropod species while contributing to the library of assays applicable to studies of mollusks. 10 Environmental DNA can also be used to measure biodiversity 12-14 and genetic variation. 15-17 For example, metabarcoding has been used to measure the diversity of

Guianese fish species. 18 The eDNA approach was accepted as an effective measure of biodiversity, providing a detailed inventory of spatial data about the fish species. 18 Similarly, the genetic variation of perch-like fishes in Russia was analyzed using eDNA metabarcoding. 17

Researchers identified 42 official species along with many distinct cryptic species. 17

Environmental studies often concern the distribution and behavior of aquatic species. 19-21

Environmental DNA monitoring of invasive Bigheaded Carps revealed patterns of abundance over time and location. 8 Relationships exist between eDNA and Carp movement and between eDNA and hydrographic conditions. 8 Additional applications of environmental DNA will likely appear with the development of more advanced sequencing technology coupled with

2 standardized methods of eDNA collection and extraction.

Unlike studies of microbes where entire genomes are detected by sampling organisms directly 22, 23, eDNA studies target DNA molecules. 1 It is important to note that eDNA is not a scientific method, in itself. 1 Instead, environmental DNA is a useful resource that can be extracted from the environment and analyzed with scientific methods, such as polymerase chain reaction (PCR) and metabarcoding. 24 These methods are made possible by rapidly developing technology, such as next generation sequencing and other molecular techniques. 24 Once it is shed from aquatic organisms, environmental DNA becomes available for scientists to collect.

DNA is “captured” from the samples using various methods, including precipitation, centrifugation, and filtration.25 Filtration has been found to be the most successful of these capture methods.25 The eDNA is then extracted from the samples, after which it can be analyzed.

25, 26

There are two main approaches for analyzing eDNA: single-species detection and metabarcoding. The single species approach requires the development of a specific primer that only amplifies the species of interest. 4, 5, 8, 27 This highly specific primer is then used in quantitative PCR (qPCR) to make copies of a target region of DNA and signify detection through successful amplification. 4, 5, 8, 27 In metabarcoding studies, general primers are designed to amplify a broad taxonomic group of organisms. 5, 7, 10, 17 Next generation sequencing is then performed on the products of PCR, simultaneously detecting numerous species by comparison with a reference sequence database. 5, 7, 10, 17 In most eDNA studies, primers are designed to target mitochondrial DNA. 5, 28-30 Mitochondrial genes are optimal for eDNA because their high copy number per cell helps ensure survival through environmental degradation. 31 Mitochondrial

3 genes also have an extensive reference database and contain higher species-specific variation. 1,

There are important differences to consider between the two methods of eDNA analysis, single-species detection and metabarcoding. In single-species detection, the primer required for the polymerase chain reaction (PCR) must be highly specific to the single target species. 33-36

Ideally, the species-specific primer will amplify the target DNA sequence by making millions of copies during PCR. 33-36 If PCR works and the DNA sequence is amplified, the target species has been detected. 33-36 Quantitative PCR (qPCR) is often used in single-species detection studies and has several advantages over standard PCR. 37 By using fluorescent labelling probes, qPCR is more sensitive than general PCR and can provide some information about the quantity of the

DNA signal in samples. 37 For example, species-specific primers were developed to detect the

Macquarie perch (Macquaria australasica), a fish endemic to Australia. 19 The primers were designed to detect the 12S region of the mitochondrial DNA along with a specific region of nuclear DNA for the Macquarie perch. 19 Using qPCR, the primers successfully detected

Macquarie perch DNA and were used to track perch spawning activity based on abundance data.

Unlike in single-species detection, metabarcoding requires the development of a more general primer used to detect large taxonomic groups. 38-41 The primer must be general enough to capture all desired species, but not too general that it amplifies the DNA from non-target species, such as humans. 38-41 The general workflow of a metabarcoding study consists of collection, extraction, amplification, and sequencing. 24 Instead of detection through PCR amplification, metabarcoding studies rely on detection of species through sequencing data generated by next- generation sequencing. 38-41 Sequences are clustered into Operational Taxonomic Units (OTUs)

4 and identified to species by matching to a reference database. 38-41 A metabarcoding study was used to evaluate the diversity of Guianese freshwater fishes. 18 Around 8 million sequencing reads were generated and 132 Guianese fish species were identified using the metabarcoding method. 18 Because of the differences between single-species detection and metabarcoding, the two methods are useful under different circumstances.

Single-species detection and metabarcoding have advantages and disadvantages based on the circumstances of a given study. Single-species studies are convenient, because PCR amplification and detection occur simultaneously, eliminating the need for additional steps. 33-36

Single-species studies are also generally less expensive than metabarcoding, because they lack the extra expense of next-generation sequencing. 5, 27, 42 In addition, single-species analyses can be performed as samples trickle in without affecting the cost. 27 Metabarcoding becomes cost efficient only after collecting a large batch of samples 27, delaying analysis until samples are collected. As sequencing costs continue to decrease 27, 32, however, metabarcoding studies will likely become more cost efficient.

There are many instances when metabarcoding is more suitable than single-species detection. Conveniently, metabarcoding detection does not require any prior knowledge of the species that might be detected. 42 Metabarcoding also requires less development and validation of primers, because a general primer will work for a broad taxonomic group. Single-species detection, on the other hand, requires the development of a new primer set for each species, and more validation is needed to confirm the primer’s specificity. 5, 42, 43 If single-species primers are not designed correctly, they can generate false positives by cross-amplifying non-target species.

27, 42, 43 False negatives can also be produced in single-species detection by failing to amplify members of the target species with geographic or intraspecific variation. 27, 42, 43 Metabarcoding

5 also produces more reliable results than single-species detection. In metabarcoding studies, the identification of species is based on the analysis of multiple diagnostic sites compared with a reference database, while single-species detection is based only on whether PCR works or not. 5,

44 Finally, metabarcoding is generally more useful than single-species detection, because metabarcoding surveys contribute to future reference sequence databases 27, 44, increasing knowledge of regional variation and the dispersal of cryptic species. In single-species studies, your results are limited to presence/absence data for the single species you were trying to detect.

33-36 Because metabarcoding is generally more resistant to logistical errors and produces more reliable species detection and identification, we argue that this method is superior to single- species detection in most situations. There are still instances, however, when single-species detection might be advantageous over metabarcoding.

Environmental DNA survey methods are powerful tools that are generally more cost efficient and less invasive than traditional surveys. Surveys using eDNA provide valuable information about the location, movements, and genetic diversity of species based on presence/absence data. Both single-species detection and metabarcoding are useful in specific situations. Assuming good primers are validated and available, single-species methods are cost efficient and can produce results as samples trickle in. Although metabarcoding requires samples to be analyzed all at once, it is most useful for characterizing multiple species of the same taxonomic group and provides more confident identification of species through sequence analyses. Understanding the advantages and limitations of each eDNA method will help you choose the best method for your study. We believe that eDNA surveys should be used to complement and guide traditional survey methods leading to greater detection and cost efficiency.

References:

1. Lacoursière-Roussel, A.; Deiner, K., Environmental DNA is not the tool by itself. Journal of Fish Biology 2019, n/a (n/a). 2. Hoffmann, C.; Schubert, G.; Calvignac-Spencer, S., Aquatic biodiversity assessment for the lazy. Molecular Ecology 2016, 25 (4), 846-848. 3. Beans, C., Environmental DNA helps researchers track pythons and other stealthy creatures. Proceedings of the National Academy of Sciences of the United States of America 2018, 115 (36), 8843-8845. 4. Renan, S.; Gafny, S.; Perl, R. G. B.; Roll, U.; Malka, Y.; Vences, M.; Geffen, E., Living quarters of a living fossil—Uncovering the current distribution pattern of the rediscovered Hula painted frog (Latonia nigriventer) using environmental DNA. Molecular Ecology 2017, 26 (24), 6801-6812. 5. Roy, M.; Belliveau, V.; Mandrak, N. E.; Gagné, N., Development of environmental DNA (eDNA) methods for detecting high-risk freshwater fishes in live trade in Canada. Biological Invasions 2018, 20 (2), 299-314. 6. McDevitt, A. D.; Sales, N. G.; Browett, S. S.; Sparnenn, A. O.; Mariani, S.; Wangensteen, O. S.; Coscia, I.; Benvenuto, C., Environmental DNA metabarcoding as an effective and rapid tool for fish monitoring in canals. Journal of Fish Biology 2019, 95 (2), 679- 682. 7. Valentini, A.; Taberlet, P.; Miaud, C.; Civade, R.; Herder, J.; Thomsen, P. F.; Bellemain, E.; Besnard, A.; Coissac, E.; Boyer, F.; Gaboriaud, C.; Jean, P.; Poulet, N.; Roset, N.; Copp, G. H.; Geniez, P.; Pont, D.; Argillier, C.; Baudoin, J. M.; Peroux, T.; Crivelli, A. J.; Olivier, A.; Acqueberge, M.; Le Brun, M.; Møller, P. R.; Willerslev, E.; Dejean, T., Next-generation monitoring of aquatic biodiversity using environmental DNA metabarcoding. Molecular Ecology 2016, 25 (4), 929-942. 8. Erickson, R. A.; Rees, C. B.; Coulter, A. A.; Merkes, C. M.; McCalla, S. G.; Touzinsky, K. F.; Walleser, L.; Goforth, R. R.; Amberg, J. J., Detecting the movement and spawning activity of bigheaded carps with environmental DNA. Molecular ecology resources 2016, 16 (4), 957-965. 9. McKelvey, K. S.; Young, M. K.; Knotek, W. L.; Carim, K. J.; Wilcox, T. M.; Padgett- Stewart, T. M.; Schwartz, M. K., Sampling large geographic areas for rare species using environmental DNA: A study of bull trout Salvelinus confluentus occupancy in western Montana. Journal of Fish Biology 2016, 88 (3), 1215-1222. 10. Klymus, K. E.; Marshall, N. T.; Stepien, C. A., Environmental DNA (eDNA) metabarcoding assays to detect invasive invertebrate species in the Great Lakes. PLOS ONE 2017, 12 (5), e0177643. 11. Balasingham, K. D.; Walter, R. P.; Mandrak, N. E.; Heath, D., Environmental DNA detection of rare and invasive fish species in two Great Lakes tributaries. Molecular Ecology 2018, 27 (1), 112-127. 12. Bakker, J.; Wangensteen, O. S.; Baillie, C.; Buddo, D.; Chapman, D. D.; Gallagher, A. J.; Guttridge, T. L.; Hertler, H.; Mariani, S., Biodiversity assessment of tropical shelf eukaryotic communities via pelagic eDNA metabarcoding. Ecology and Evolution 2019, 9 (24), 14341- 14355.

13. Bagley, J. C.; de Aquino, P. D. P. U.; Breitman, M. F.; Langeani, F.; Colli, G. R., DNA barcode and minibarcode identification of freshwater fishes from Cerrado headwater streams in Central Brazil. Journal of Fish Biology 2019, 95 (4), 1046-1060. 14. Hou, G.; Chen, W. T.; Lu, H. S.; Cheng, F.; Xie, S. G., Developing a DNA barcode library for perciform fishes in the South China Sea: Species identification, accuracy and cryptic diversity. Molecular ecology resources 2018, 18 (1), 137-146. 15. del Carmen Gomez Cabrera, M.; Young, J. M.; Roff, G.; Staples, T.; Ortiz, J. C.; Pandolfi, J. M.; Cooper, A., Broadening the taxonomic scope of coral reef palaeoecological studies using ancient DNA. Molecular Ecology 2019, 28 (10), 2636-2652. 16. Arroyave, J.; Martinez, C. M.; Stiassny, M. L. J., DNA barcoding uncovers extensive cryptic diversity in the African long-fin tetra Bryconalestes longipinnis (Alestidae: Characiformes). Journal of Fish Biology 2019, 95 (2), 379-392. 17. Turanov, S. V.; Kartavtsev, Y. P.; Lipinsky, V. V.; Zemnukhov, V. V.; Balanov, A. A.; Lee, Y. H.; Jeong, D., DNA-barcoding of perch-like fishes (Actinopterygii: Perciformes) from far-eastern seas of Russia with taxonomic remarks for some groups. Mitochondrial DNA 2016, 27 (2), 1188-1209. 18. Cilleros, K.; Valentini, A.; Allard, L.; Dejean, T.; Etienne, R.; Grenouillet, G.; Iribar, A.; Taberlet, P.; Vigouroux, R.; Brosse, S., Unlocking biodiversity and conservation studies in high-diversity environments using environmental DNA (eDNA): A test with Guianese freshwater fishes. Molecular ecology resources 2019, 19 (1), 27-46. 19. Bylemans, J.; Furlan, E. M.; Hardy, C. M.; McGuffie, P.; Lintermans, M.; Gleeson, D. M., An environmental DNA-based method for monitoring spawning activity: a case study, using the endangered Macquarie perch (Macquaria australasica). Methods in Ecology and Evolution 2017, 8 (5), 646-655. 20. Levi, T.; Allen, J. M.; Bell, D.; Joyce, J.; Russell, J. R.; Tallmon, D. A.; Vulstek, S. C.; Yang, C.; Yu, D. W., Environmental DNA for the enumeration and management of Pacific salmon. Molecular ecology resources 2019, 19 (3), 597-608. 21. Stoeckle, M. Y.; Soboleva, L.; Charlop-Powers, Z., Aquatic environmental DNA detects seasonal fish abundance and habitat preference in an urban estuary. PLoS ONE 2017, 12 (4). 22. Coolen, M. J. L.; Muyzer, G.; Rijpstra, W. I. C.; Schouten, S.; Volkman, J. K.; Sinninghe Damsté, J. S., Combined DNA and lipid analyses of sediments reveal changes in Holocene haptophyte and diatom populations in an Antarctic lake. Earth and Planetary Science Letters 2004, 223 (1-2), 225-239. 23. Thomas, C.; Ariztegui, D., Fluid inclusions from the deep Dead Sea sediment provide new insights on Holocene extreme microbial life. Quaternary Science Reviews 2019, 212, 18-27. 24. Deiner, K.; Bik, H. M.; Mächler, E.; Seymour, M.; Lacoursière-Roussel, A.; Altermatt, F.; Creer, S.; Bista, I.; Lodge, D. M.; de Vere, N.; Pfrender, M. E.; Bernatchez, L., Environmental DNA metabarcoding: Transforming how we survey animal and plant communities. Molecular Ecology 2017, 26 (21), 5872-5895. 25. Eichmiller, J. J.; Miller, L. M.; Sorensen, P. W., Optimizing techniques to capture and extract environmental DNA for detection and quantification of fish. Molecular ecology resources 2016, 16 (1), 56-68. 26. Hermans, S. M.; Buckley, H. L.; Lear, G., Optimal extraction methods for the simultaneous analysis of DNA from diverse organisms and sample types. Molecular ecology resources 2018, 18 (3), 557-569.

27. Harper, L. R.; Lawson Handley, L.; Hahn, C.; Boonham, N.; Rees, H. C.; Gough, K. C.; Lewis, E.; Adams, I. P.; Brotherton, P.; Phillips, S.; Hänfling, B., Needle in a haystack? A comparison of eDNA metabarcoding and targeted qPCR for detection of the great crested newt (Triturus cristatus). Ecology and Evolution 2018, 8 (12), 6330-6341. 28. Balasingham, K. D.; Walter, R. P.; Heath, D. D., Residual eDNA detection sensitivity assessed by quantitative real-time PCR in a river ecosystem. Molecular ecology resources 2017, 17 (3), 523-532. 29. Andújar, C.; Arribas, P.; Yu, D. W.; Vogler, A. P.; Emerson, B. C., Why the COI barcode should be the community DNA metabarcode for the metazoa. Molecular Ecology 2018, 27 (20), 3968-3975. 30. Lacoursière-Roussel, A.; Rosabal, M.; Bernatchez, L., Estimating fish abundance and biomass from eDNA concentrations: variability among capture methods and environmental conditions. Molecular ecology resources 2016, 16 (6), 1401-1414. 31. Deiner, K.; Renshaw, M. A.; Li, Y.; Olds, B. P.; Lodge, D. M.; Pfrender, M. E., Long- range PCR allows sequencing of mitochondrial genomes from environmental DNA. Methods in Ecology and Evolution 2017, 8 (12), 1888-1898. 32. Sigsgaard, E. E.; Jensen, M. R.; Winkelmann, I. E.; Møller, P. R.; Hansen, M. M.; Thomsen, P. F., Population-level inferences from environmental DNA—Current status and future perspectives. Evolutionary Applications 2020, 13 (2), 245-262. 33. Ardura, A., Species-specific markers for early detection of marine invertebrate invaders through eDNA methods: Gaps and priorities in GenBank as database example. Journal for Nature Conservation 2019, 47, 51-57. 34. Gustavson, M. S.; Collins, P. C.; Finarelli, J. A.; Egan, D.; Conchúir, R. Ó.; Wightman, G. D.; King, J. J.; Gauthier, D. T.; Whelan, K.; Carlsson, J. E. L.; Carlsson, J., An eDNA assay for Irish Petromyzon marinus and Salmo trutta and field validation in running water. Journal of Fish Biology 2015, 87 (5), 1254-1262. 35. Horiuchi, T.; Masuda, R.; Murakami, H.; Yamamoto, S.; Minamoto, T., Biomass- dependent emission of environmental DNA in jack mackerel Trachurus japonicus juveniles. Journal of Fish Biology 2019, 95 (3), 979-981. 36. Riascos, L.; Geerts, A. N.; Oña, T.; Goethals, P.; Cevallos-Cevallos, J.; Vanden Berghe, W.; Volckaert, F. A. M.; Bonilla, J.; Muylaert, K.; Velarde, E.; Boets, P.; Van der heyden, C., DNA-based monitoring of the alien invasive North American crayfish Procambarus clarkii in Andean lakes (Ecuador). Limnologica 2018, 70, 20-25. 37. Adams, C. I. M.; Knapp, M.; Gemmell, N. J.; Jeunen, G. J.; Bunce, M.; Lamare, M. D.; Taylor, H. R., Beyond biodiversity: Can environmental DNA (eDNA) cut it as a population genetics tool? Genes 2019, 10 (3). 38. Oliveira, L. M.; Knebelsberger, T.; Landi, M.; Soares, P.; Raupach, M. J.; Costa, F. O., Assembling and auditing a comprehensive DNA barcode reference library for European marine fishes. Journal of Fish Biology 2016, 89 (6), 2741-2754. 39. Nichols, S. J.; Kefford, B. J.; Campbell, C. D.; Bylemans, J.; Chandler, E.; Bray, J. P.; Shackleton, M.; Robinson, K. L.; Carew, M. E.; Furlan, E. M., Towards routine DNA metabarcoding of macroinvertebrates using bulk samples for freshwater bioassessment: Effects of debris and storage conditions on the recovery of target taxa. Freshwater Biology 2019. 40. Jerde, C. L.; Wilson, E. A.; Dressler, T. L., Measuring global fish species richness with eDNA metabarcoding. Molecular ecology resources 2019, 19 (1), 19-22.

41. Hänfling, B.; Handley, L. L.; Read, D. S.; Hahn, C.; Li, J.; Nichols, P.; Blackman, R. C.; Oliver, A.; Winfield, I. J., Environmental DNA metabarcoding of lake fish communities reflects long-term data from established survey methods. Molecular Ecology 2016, 25 (13), 3101-3119. 42. Schneider, J.; Valentini, A.; Dejean, T.; Montarsi, F.; Taberlet, P.; Glaizot, O.; Fumagalli, L., Detection of Invasive Mosquito Vectors Using Environmental DNA (eDNA) from Water Samples. PLOS ONE 2016, 11 (9), e0162493. 43. Goldberg, C. S.; Turner, C. R.; Deiner, K.; Klymus, K. E.; Thomsen, P. F.; Murphy, M. A.; Spear, S. F.; McKee, A.; Oyler-McCance, S. J.; Cornman, R. S.; Laramie, M. B.; Mahon, A. R.; Lance, R. F.; Pilliod, D. S.; Strickler, K. M.; Waits, L. P.; Fremier, A. K.; Takahara, T.; Herder, J. E.; Taberlet, P., Critical considerations for the application of environmental DNA methods to detect aquatic species. Methods in Ecology and Evolution 2016, 7 (11), 1299-1307. 44. Lacoursière-Roussel, A.; Dubois, Y.; Normandeau, E.; Bernatchez, L., Improving herpetological surveys in eastern North America using the environmental DNA method. Genome 2016, 59 (11), 991-1007.