Resource Manager's eDNA Toolbox

• Why Use Environmental DNA?
• Common Questions When Beginning an eDNA Surveillance Program
• Advantages and Limitations
• The Process of eDNA
• Molecular Methods
• References

Why use environmental DNA as a surveillance tool? 

Environmental DNA sampling is a powerful surveillance tool. Instead of relying on the capture of live organisms in the environment, utilizing eDNA methods allows you to indirectly monitor species without the need to physically observe them. eDNA also enhances your ability to detect the genetic traces of rare species that would be difficult to monitor using traditional surveying methods. This allows scientists and managers to gain a better understanding of what species live in a specific environment. Water is the most common medium sampled for eDNA but air and soil can also be sampled. There are many advantages to using eDNA. Environmental DNA techniques can be used for:

• the early detection of invasive species.
• detection of cryptic or endangered species.
• revealing important information about biodiversity. 
• monitoring the health of the ecosystem.

What is environmental DNA?

DNA (deoxyribonucleic acid) is a large, helical shaped biological molecule present in organisms that contains genetic information. The complete set of genetic information is referred to as the genome. The differences in the sequence or order of the DNA building blocks (base pairs) makes it possible to identify different species. An identifiable sequence at a recognized location in the genome is called a marker.

Environmental DNA (eDNA) is DNA present in various aquatic or terrestrial environments (water, soil, sediment). It's defined as “genetic material obtained directly from environmental samples without any obvious signs of biological source material."^[20] eDNA originates from cellular material shed by organisms (such as skin, hair, excrement, etc.), and it can be extracted from complex environmental samples and analyzed using molecular techniques. 

While eDNA is a powerful tool that resource managers can use for conservation efforts, biodiversity monitoring, and ecosystem management, it should not be used to replace traditional surveying methods. eDNA is used to contribute to the weight of evidence used for decision making and should be used to complement traditional methods.

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Common questions when beginning an eDNA surveillance program

Where do I begin when starting an eDNA surveillance program?

When beginning an eDNA surveillance program, you must clearly define your objectives. What is it that you hope to achieve with this program? Field and lab protocols must be developed before any sort of sampling or lab work can start. Pilot studies may be needed so that the techniques can be refined. Initial data collection may lead you to some preliminary conclusions that will cause you to alter your techniques or realize additional data collection is needed. Beginning an eDNA surveillance program takes time, commitment, and financial support for long-term planning. An effective communication plan must be established for all relevant parties, see eDNA Communication Strategies for details.

You must also decide on what happens after a positive detection. Do you begin physical sampling? Is your work done? Does your partner take over? There must be a clear plan in place once a positive detection occurs that defines what your next steps are.

What are the costs associated with beginning an eDNA surveillance program?

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There is no hard number for the cost of eDNA surveillance programs as each program and study will be different. The costs involved in eDNA surveillance programs can vary. Some costs you must consider include the costs for beginning multiple pilot studies, funding for marker development and validation, for qPCR machines, and funding for research and development. The laboratory analysis costs are typically greater than field sampling costs.

When deciding between eDNA and traditional sampling methods, the time requirement is a key factor. Traditional sampling could take days of sampling whereas eDNA could be hours. You must also consider the lab processing costs. For eDNA certain costs will include sample collection, lab analysis, lab personnel, supplies, and reagents. The choice between using traditional sampling methods and eDNA will rely on the study and target species.

What should labs conducting eDNA studies consider?

Labs that are used for eDNA surveillance programs should have certain criteria needed to be properly used for eDNA purposes. There must be strict decontamination protocols in place and a dedicated space for analysis. See eDNA Best Practices for recommendations on laboratory analysis.

Advantages and limitations

Environmental DNA is a promising biosurveillance tool that you can utilize to assess biodiversity and ecosystem health. There are certain advantages and limitations to using eDNA as a monitoring tool. 

Advantages

• It’s cost effective, while the costs of eDNA studies will depend on your target species, many eDNA studies were found to be less expensive than traditional surveying methods which can be labor intensive.^[20] However, eDNA analysis is not cheap as there are many costs to keep in mind. 
• It’s non-invasive, there’s no need to disturb targeted species by trapping them to determine their presence in the environment. Capturing and handling fish or a species of interest is stressful on them and can actually harm the individuals caught up in the surveys. This is especially helpful if your target species is rare. With eDNA, scientists can see if a targeted species’ population is rising or declining without having to physically count them.
• It’s sensitive, due to the sensitivity of eDNA methods, there is a higher chance of detecting species even at a lower density.
• You have the ability to access protected or unreachable environments.
• eDNA samples may be archived and saved for future studies.
• Sampling teams can be made up of individuals from all backgrounds, not necessarily just experts in ecology.

Limitations

• It can be challenging to find where the DNA came from, whether it came from the habitat or if it was transported by other means such as sewage or feces. A detection of eDNA doesn't necessarily mean that a live organism is present in the environment.
• eDNA cannot determine the life stage of the target species.
• False positives and false negatives can negatively impact eDNA surveys. It’s important to establish proper protocols to help reduce the chance of detecting false negatives or false positives.
• eDNA degrades at different rates due to factors such as microbial activity, heat, acidity, and ultraviolet light. It's important to preserve your samples as soon as possible to prevent degradation.
• Primer bias can occur, where one species' DNA is preferentially amplified over that of another during PCR.^[22] This can happen when using metabarcoding and general primers.

The process of eDNA

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Molecular methods

Quantitative polymerase chain reaction (qPCR)

The technique most often used to detect eDNA from complex environmental samples is polymerase chain reaction (PCR). Targeted methods (single species or genera) eDNA assays exclude all related species or co-occurring species in the sample. PCR works by quickly replicating millions or billions of copies of a short fragment of specific DNA sequence (a marker). Quantitative PCR is often used because of its high specificity due to carefully designed forward and reverse primers and probe. Combined with its high sensitivity, this makes qPCR an ideal method for single species detection.

Droplet digital polymerase chain reaction (ddPCR)

Droplet digital PCR is a technology that can be used for eDNA analysis. ddPCR works using water-in-oil emulsions, separating the sample into thousands of droplets, each as small as 1 nanoliter. These droplets are run through a PCR cycle like any other probe-based method. The fluorescence is measured in each individual droplet after amplification and the positive number of droplets is calculated. ddPCR is highly sensitive and precise, making it an acceptable method for eDNA. However, it tends to be more expensive than qPCR.  

Metabarcoding

To detect multiple species at once, metabarcoding is a method that is commonly used. By using non-specific primers, followed by high-throughput sequencing, it's possible to find hundreds of species in a single sample. Barcoding is sequencing a section of DNA to identify a species. Metabarcoding is simultaneously sequencing millions of DNA fragments to identify species represented in a pool of DNA. This method is useful for surveying environments to learn what species may be present. The number of reads for each species can often be correlated to some degree with species abundance or biomass. However, you must consider eDNA movement (eDNA entering a habitat by other means such as sewage, feces from other animals, etc.) and degradation as well. 

Point of use assays

Another method is the point of use assays, typically using either LAMP or CRISPR technologies. These assays allow for rapid on-site detection of single species or genera. They do not rely on the amplification of DNA through a PCR. Point of use assays need simple reagents and instruments for applications in the field or at port of entry.

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References

¹ Abbott C., Coulson M., Gagné N., Lacoursière‐Roussel A., Parent G.J., Bajno R., Dietrich C., May-McNally S. 2021. Guidance on the Use of Targeted Environmental DNA (eDNA) Analysis for the Management of Aquatic Invasive Species and Species at Risk. Fisheries and Oceans Canada, Ottawa, ON, Canada, Canadian Science Advisory Secretariat, 2021/019: iv+42 pp.

² Bohmann, K.; Evans, A.; Gilbert, M.T.; Carvalho, G.R.; Creer, S.; Knapp, M.; Yu, D.W.; de Bruyn, M. 2014. Environmental DNA for wildlife biology and biodiversity monitoring. Trends Ecol. Evol. 29, 358–367.

³ Bustin S.A., Benes V., Garson J.A., Hellemans J., Huggett J., Kubista M., Mueller R., Nolan T, Pfaffl MW, Shipley GL, Vandesompele J, Wittwer CT. 2009. The MIQE Guidelines: Minimum Information for Publication of Quantitative Real-Time PCR Experiments. Clinical Chemistry 55: 611–622.

⁴ 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.

⁵ Erickson R.A., Merkes C.M., Mize E.L. 2019. Sampling designs for landscape-level eDNA monitoring programs. Integrated Environmental Assessment and Management. 15(5): 760-771.

⁶ Ferrante J.A., Daniel W.M., Freedman J.A., Klymus K.E., Neilson M.E., Passamaneck Y., Rees C.B., Sepulveda A., Hunter M.E. 2022. Gaining decision-maker confidence through community consensus: developing environmental DNA standards for data display on the USGS Nonindigenous Aquatic Species database. Management of Biological Invasions 13(4): 809–832.

⁷ 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.; et al. 2016. Critical considerations for the application of environmental DNA methods to detect aquatic species. Methods Ecol. Evol. 7, 1299–1307.

⁸ Hobbs, J., Round, J. M., Allison, M. J., & Helbing, C. C. 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(3).

⁹ Hoorfar J., Malorny B., Abdulmawjood A., Cook N., Wagerner M., Fach P. 2004. Practical considerations in design of internal amplification controls for diagnostic PCR assays. J. Clin. Microbiol. 42(5), 1863-1868.

¹⁰ Hunter M.E., Ferrante J.A., Meigs-Friend G., Ulmer A. 2019. Improving eDNA yield and inhibitor reduction through increased water volumes and multi-filter isolation techniques. Sci Rep. 9(1):5259. 

¹¹ Klymus K.E., Merkes C.M., Allison M.J., Goldberg C.S., Helbing C.C., Hunter M.E., Jackson C.A., Lance R.F., Mangan A.M., Monroe E.M., Piaggio A.J., Stokdyk J.P., Wilson C.C., Richter C.A. 2020. Reporting the limits of detection and quantification for environmental DNA assays. Environmental DNA 2: 271–282.

¹² Langlois V.S., Allison M.J., Bergman L.C., To T.A., Helbing C.C. 2021. The need for robust qPCR‐based eDNA detection assays in environmental monitoring and species inventories. Environmental DNA 3: 519–527.

¹³ Lesperance, M. L., Allison, M. J., Bergman, L. C., Hocking, M. D., & Helbing, C. C. (2021). A statistical model for calibration and computation of detection and quantification limits for low copy number environmental DNA samples. Environmental DNA, 3, 970–981.

¹⁴ Morisette J., Burgiel S., Brantley K., Daniel W., Darling J., Davis J., Franklin T., Gaddis K., Hunter M., Lance R., Leskey T., Passamaneck Y., Piaggio A., Rector B., Sepulveda A., Smith M., Stepien C., Wilcox T., Wm D., Stepien S. 2021. Strategic considerations for invasive species managers in the utilization of environmental DNA (eDNA): steps for incorporating this powerful surveillance tool. Management of Biological Invasions 12: 747–775.

¹⁵ Sepulveda A.J., Nelson N.M., Jerde C.L., Luikart G. 2020b. Are Environmental DNA Methods Ready for Aquatic Invasive Species Management? Trends in Ecology & Evolution 35: 668– 678.

¹⁶ Shu, L., Ludwig, A., Peng, Z. 2020. Standards for Methods Utilizing Environmental DNA for Detection of Fish Species. Genes. 11, 296.

¹⁷ Stein, E. D., Jerde, C.L., Allan, E.A., Sepulveda, A., Abbott, C.L., Baerwald, M.R., Darling, J., Goodwin, K.D., Meyer, R.S., Timmers, M.A., & Thielen, P.M. 2023. Critical considerations for communicating environmental DNA science. Environmental DNA, 00, 1-12. 

¹⁸ Taberlet, P., Coissac, E., Hajibabaei, M., Rieseberg, L.H. 2012. Environmental DNA. Mol. Ecol. 21, 1789–1793.

¹⁹ Thalinger, B., Deiner, K., Harper, L. R., Rees, H. C., Blackman, R. C., Sint, D., Traugott, M., Goldberg, C. S., & Bruce, K. 2021. A validation scale to determine the readiness of environmental DNA assays for routine species monitoring. Environmental DNA, 3(4), 823– 836.

²⁰ Thomsen, P.F., Willerslev, E. 2015. Environmental DNA—An emerging tool in conservation for monitoring past and present biodiversity. Biol. Conserv. 183, 4–18.

²¹ Ulibarri R.M., Bonar S.A., Rees C.B., Amberg J.J., Ladell B., Jackson C. 2017.Comparing Efficiency of American Fisheries Society Standard Snorkeling Techniques to Environmental DNA Sampling Techniques. North American Journal of Fisheries Management. 37, 644-651.

²² U.S. Fish and Wildlife Service. 2023. Environmental DNA (eDNA) Best Management Practices for Project Planning, Deployment, and Application. U.S. Fish and Wildlife Service. Anchorage, AK: Conservation Genetics Laboratory.