U.S. Department of the Interior

U.S. Geological Survey

Fact Sheet 2012–3146

January 2013 Printed on recycled paper

Application of Environmental DNA for Inventory and 

Monitoring of Aquatic Species

This fact sheet was created to help biologists and resource managers 

understand emerging methods for detecting environmental DNA and 

their potential application for inventorying and monitoring aquatic 

species. It is a synthesis of published information.

What is Environmental DNA?

DNA, short for deoxyribonucleic acid, is the hereditary 

material in organisms that contains the biological instructions for 

building and maintaining them. The chemical structure of DNA 

is the same for all organisms, but differences exist in the order of 

the DNA building blocks, known as base pairs. Unique sequences 

of base pairs, particularly repeating patterns, provide a means to 

identify species, populations, and even individuals.

Environmental DNA (eDNA) is nuclear or mitochondrial 

DNA that is released from an organism into the environment. 

Sources of eDNA include secreted feces, mucous, and gametes; 

shed skin and hair; and carcasses. eDNA can be detected in cellular 

or extracellular (dissolved DNA) form. 

In aquatic environments, eDNA is diluted and distributed by 

currents and other hydrological processes (fig. 1), but it only lasts 

about 7–21 days, depending on environmental conditions (Dejean 

and others, 2011). Exposure to UVB radiation, acidity, heat, and 

endo- and exonucleases can degrade eDNA.

Use of eDNA for Inventory and Monitoring

Improved Detection of Native Species

Protocols using eDNA may allow for rapid, cost-effective, 

and standardized collection of data about species distribution and 

relative abundance. For small, rare, secretive, and other species that 

are difficult to detect, eDNA provides an attractive alternative for 

aquatic inventory and monitoring programs. Increasing evidence 

demonstrates improved species detection and catch-per-unit effort 

compared with electrofishing, snorkeling, and other current field 

methods. Thus, detection of species using eDNA may improve 

biodiversity assessments and provide information about status, 

distribution, and habitat requirements for lesser-known species.

Early Detection of Invasive Species 

eDNA may also be an effective tool for early detection of 

aquatic invasive species. Recent studies have focused on Asian 

carp (Jerde and others, 2011) and American bullfrogs (Dejean and 

others, 2012), but protocols are being developed for New Zealand 

mudsnails, zebra mussels, quagga mussels, and others. Application 

of eDNA methods for invasive species monitoring may include 

Figure 1. This tadpole of the Rocky Mountain tailed frog secretes enough 

DNA into the surrounding water that it can be detected in a water sample 

collected downstream (see Goldberg and others, 2011). Photograph taken 

by Jason Jones, Nevada Department of Wildlife. 

periodically collecting water samples and screening them for several 

invasive species at once. Boat-ballast water, a source of introduction 

for many invasive species including mollusks, also could be sampled. 

Some intensive eradication programs for invasive species fail when a 

few surviving individuals recolonize the ecosystem. eDNA methods 

may provide a means of confirming eradication of all invaders.

Developing eDNA Protocols for Species Monitoring

Primer and Probe Design 

Conventional Polymerase Chain Reaction (PCR) methods 

can be used for analyzing eDNA, but they may cross-amplify and 

provide false-positive results. Quantitative PCR (qPCR) methods 

are preferable to conventional PCR because they are likely more 

sensitive. In qPCR, primers are used to amplify a region of DNA 

that is specific to a target organism, and a probe is used to provide 

additional specificity and quantitative information. Several eDNA

primer and probe sequences for individual species have been 

published. Other primer/probe sets must be designed for new 

species or specific needs.
Figure 2. A water sample can be collected easily in the field 

in a sterile bottle and, if needed, stored temporarily in a cooler 

or refrigerator before eDNA is concentrated through filtration 

or centrifugation. Photograph taken by Matthew Laramie, U.S. 

Geological Survey.

Laboratory Optimization 

The primer and probe design must be tested to ensure that the 

qPCR reaction always results in a positive detection in the presence 

of target-organism DNA and that amplification of non-target DNA 

does not occur. DNA from the target species should be screened 

prior to analyzing environmental samples to ensure test sensitivity. 

Preferably, this will involve 10 or more samples collected across 

the range of the species where the test will be applied. Closely 

related, co-occurring species also should be screened to ensure 

specificity prior to analyzing environmental samples, preferably 

involving five or more samples of each. 

DNA can be extracted from preserved tissue samples using 

readily available kits. It is critical that no cross contamination 

occurs between species during the tissue-storage or 

extraction processes. 

How to Design Species-Specific Primers and 

Probe for qPCR

1. Create an inclusive consensus sequence that incorporates all within-species variability for a species 

in a well-known region of DNA. Mitochondrial 

DNA is preferred because it is more abundant 

than nuclear DNA, and more sequence data are 

available. Use sequences published in GenBank 

(National Center for Biotechnology Information, 

2012), or sequence tissue samples of target species. 

It is important that the data incorporated include 

adequate sampling in the geographic area where the 

test will be applied.

2. For the selected probe chemistry, set appropriate qPCR primer software to design short, unique 

sequences for use as forward- and reverse-primers 

and probe. Optimal probe length will differ by 

chemistry. These primers and probe will allow for 

amplification and detection of the target sequence 

(90–120 base-pair length is recommended).

3. Compare the resulting design to sequences in 

GenBank to determine if the sequences are likely to 

cross-amplify with other species. Try to incorporate 

as many differences as possible (at least 2 on each 

primer and 2 on the probe, including 1 toward the 

3’ end) between the primer/probe design for your 

target species and any other species in the database.

water filtered (volumes of 1–10 L are common). Filter methods 

also require either freezing of the filter paper (Jerde and others, 

2011; Takahara and others, 2012) or dehydration of the filter paper 

in vials with molecular-grade ethanol (Goldberg and others, 2011). 

Although all these methods have been successful, ongoing testing, 

standardization, and optimization of field and laboratory protocols 

will continue to improve applications for inventory and monitoring 

programs.

DNA Extraction and Amplification

DNA extracted from the preserved samples is stable once 

it has been purified and preserved, and only a portion is used in 

each PCR reaction. This preserved DNA can be later tested for 

additional species if desired. 

Following DNA extraction, qPCR analysis provides detection 

information about the target species’ DNA. Although the amount of 

target DNA present in field samples may be quantified (Thomsen 

and others, 2012; Takahara and others, 2012), this fact sheet is 

limited to presence/absence information.

Sources of Error

Identifying sources of error or uncertainty is a critical process 

in any study, especially for monitoring programs where results 

could influence future management decisions. Darling and Mahon 

(2011) provide an excellent overview of potential sources of 

uncertainty associated with DNA-based methods for monitoring 

aquatic species. The following points are important when using 

eDNA methods.

Design of Molecular Assay 

Assay design must account for the variation within a species 

and the variation among species. Failure to incorporate the full range 

of genetic variation of a target species can lead to false negatives, 

whereas failure to incorporate the full range of genetic variation 

in closely related, co-occurring species can lead to false positives. 

Therefore, it is important to select a genetic region that maximizes 

the amount of genetic information available for target and related 

non-target species. For some species, this may require sequencing 

additional samples to ensure the assay is both sensitive and specific.

Field Sampling

Four methods for field sampling have been developed to 

date: (1) collect 15 mL of water, preserve using ethanol and 

sodium acetate, and freeze immediately (Ficetola and others, 2008; 

Thomsen and others, 2012), (2) filter water through a cellulose 

nitrate filter (Goldberg and others, 2011), (3) filter water through 

a glass fiber filter (Jerde and others, 2011), and (4) filter water 

through carbonate filter (Takahara and others, 2012) (table 1; 

figs. 2–4). The latter three methods require pumps (either in-line, 

such as a peristaltic pump, or vacuum-line) and measurement of 
Figure 3. An example of streamside water filtration using a peristaltic pump 

and sterile filter funnel. Photograph taken by Matthew Laramie, 

U.S. Geological Survey.

Figure 4. Filtration allows a specified quantity of water to be 

“tested” for eDNA. During filtration, DNA in the water is trapped on 

the filter paper, which is frozen or dehydrated. Photograph taken by 

Matthew Laramie, U.S. Geological Survey.

Table 1. Methods used to detect eDNA from aquatic organisms in freshwater environments.

[Abbreviations: L, liter; ha, hectare; mL, milliliter; M, Molar; °C, degrees Celsius; μm, micrometer; cm, centimeter; m3, cubic meter; km3, cubic kilometer]

Source Environment Species Volume 

sampled Filter type Preservationmethod

Ficetola and others 

(2008)

Aquariums (3-L) 

Ponds (0.1–1 ha

surface area)

American bullfrog (Lithobates 

catesbeianus)

15 mL No filter used—entire sample 

centrifuged

Samples combined with 1.5 mL 

sodium acetate (3 M) and 

33 mL absolute ethanol, then 

stored at -20°C

Jerde and others 

(2011)

Large river/canal 

complex

Big headed carp (Hypophthichthys 

nobilis) 

Silver carp (H. molitrix)

2 L 1.5-μm pore size glass fiber 

filters

-20°C

Goldberg and others 

(2011)

Streams (1–2nd 

order)

Idaho giant salamander 

(Dicamptodon aterrimus) 

Rocky Mountain tailed frog 

(Ascaphus montanus)

5 L and 

10 L

0.45-μm pore size cellulose 

nitrate filter

95 percent ethanol

Thomsen and others 

(2012)

Ponds

Lakes

Streams

Common spadefoot (Pelobates 

fuscus)

Great crested newt (Triturus 

cristatus)

European weather loach (Misgurnus 

fossilis)

Eurasian otter (Lutra lutra)

White-faced darter (Leucorrhinia 

pectoralis) 

Tadpole shrimp (Lepidurus apus)

15 mL No filter used—entire sample 

centrifuged

Samples combined with 1.5 mL 

sodium acetate (3 M) and 

33 mL absolute ethanol, then 

stored at -20°C

Dejean and others 

(2011); 

Dejean and others 

(2012)

Glass beakers 

(900-mL)

Ponds

American bullfrog (Lithobates 

catesbeianus) 

Sturgeon (Acipenser baerii)

15 mL No filter used—entire sample 

centrifuged

Samples combined with 1.5 mL 

sodium acetate (3 M) and 

33 mL absolute ethanol, then 

stored at -20°C

Takahara and others 

(2012)

Aquariums 

(30×45×25 cm)

Ponds (41.1–54.5 m3

volume)

Freshwater lagoon 

(0.49 km3 volume)

Common carp 

(Cyprinus carpio)

20 mL 

and 

2 L

3.0-μm pore size polycarbonate 

filter/12-μm pore size 

polycarbonate pre-filter 

with 0.8-μm pore size 

polycarbonate filter/3.0‐μm 

pore size cellulose acetate 

filter

-18°C and -25°C
Front page banner photograph: Environmental DNA holds promise for tracking 

non-native species like this brook trout in the Boise River, Idaho. Photograph 

taken by Jason Jones, Nevada Department of Wildlife.

Front page bullfrog photograph: Environmental DNA has been used to detect 

non-native species, like this American bullfrog, in lakes and ponds in Europe 

(Dejean and others, 2012). Photograph taken by John Cossel, Department of 

Biology, Northwest Nazarene University.

Publishing support provided by the U.S. Geological Survey

Tacoma Publishing Service Center

Authors: David S. Pilliod, U.S. Geological Survey, Forest and 

Rangeland Ecosystem Science Center, Boise, Idaho 83706, USA; 

dpilliod@usgs.gov.

Caren S. Goldberg, Fish and Wildlife Sciences, University of Idaho, 

Moscow, Idaho 83844-1136, USA; cgoldberg@uidaho.edu

Matthew B. Laramie, U.S. Geological Survey, Forest and Rangeland

Ecosystem Science Center, Boise, Idaho 83706, USA; 

mlaramie@usgs.gov.

Lisette P. Waits, Fish and Wildlife Sciences, University of Idaho, 

Moscow, Idaho 83844-1136, USA; lwaits@uidaho.edu.

For information contact:

Director, Forest and Rangeland Ecosystem Science Center

U.S. Geological Survey

777 NW 9th Street, Suite 400, Corvallis, Oregon 97330

http://fresc.usgs.gov/

Quality Control 

Positive and negative controls are necessary to ensure 

quality and reliability of results at each stage of the study. All 

DNA extractions should include a negative control, so that cross 

contamination between extracts can be detected. Each well of the 

PCR plate should include an internal positive control to ensure 

that the reaction is not inhibited. All eDNA extractions and qPCR 

setups should be conducted in a PCR-free laboratory space where 

concentrated (such as from tissue) DNA samples have not been 

handled. Thermocyclers and real-time PCR machines should be 

located outside of this space.

Detection Probability 

Like other field-based sampling, results of eDNA detection 

may have some inaccuracy, and replicate samples are required 

to estimate occupancy while accounting for uncertainty. In other 

words, not detecting DNA of a species does not mean it is absent. 

The lower limits of detection for species are currently unknown and 

likely vary depending on the species and its density, size, behavior, 

and habitat.

Field Negative Controls

Two types of negative controls are often employed in the 

field to increase accuracy. First, samples from a few sites outside 

the range of a target species are used to confirm non-detection 

in locations where the species is not present. Second, samples 

are collected from distilled water using the field protocol at each 

site to ensure that cross contamination is not occurring between 

replicate samples within a site and between sites. Sterile gloves, 

filters, water collection bottles, and sample containers reduce risk 

of contamination. High-quality sample tubes placed individually 

inside plastic, sealable bags can reduce cross-contamination should 

leakage occur when samples are stored or shipped to laboratories. 

Timing of Sampling

The timing of sampling may need to coincide with the 

life history or behavior of a target species. For example, during 

reproduction when young-of-year are present, eDNA may be 

abundant. The arrival of migratory species can be detected 

assuming no other life stages of the species remain in the system.

PCR Replication

Degraded, low-quantity DNA samples are often analyzed 

in triplicate to ensure detection of DNA (Waits and Paetkau, 

2005) and to assess potential false-positives. Using this approach, 

additional analysis is required if results are not uniform. Standard 

curves should be developed based on DNA obtained from tissue 

samples of target species and span the range of sample results.

Future Directions

Although eDNA methods show great potential for inventory 

and monitoring aquatic species, there are still details to resolve. 

Thomsen and others (2012) point out that it is necessary to gain a 

better understanding of how field methods, laboratory protocols, 

and environmental conditions influence the detection of eDNA. 

Further, there is little information about factors that influence lower 

limits of detection, production of DNA, and persistence of DNA 

in different types of aquatic systems. These factors will likely 

vary among species and life stages. Before adoption of standard 

procedures for eDNA sampling and analysis, further development 

and comparative testing of protocols are necessary.

References Cited

Darling, J.A., and Mahon, A.R., 2011, From molecules to management—

Adopting DNA-based methods for monitoring biological invasions in 

aquatic environments. Environmental Research, v. 111, iss. 7, p. 978–988, 

doi:10.1016/j.envres.2011.02.001.

Dejean, T., Valentini, A., Duparc, A., Pellier-Cuit, S., Pompanon, F., Taberlet, 

P., and Miaud, C., 2011, Persistence of environmental DNA in freshwater 

ecosystems: PLoS ONE, v. 6, no. 8, e23398, doi:10.1371/journal.

pone,0023398.

Dejean, T., Valentini, A., Miquel, C., Taberlet, P., Bellemain, E., and Miaud, C., 

2012, Improved detection of an alien invasive species through environmental 

DNA barcoding—The example of the American bullfrog Lithobates 

catesbeianus: Journal of Applied Ecology, v. 49, iss. 4, p. 953–959, doi: 

10.1111/j.1365-2664.2012.02171.x. 

Ficetola, G.F., Miaud, Claude, Pompanon, François, and Taberlet, Pierre, 2008, 

Species detection using environmental DNA from water samples: Biology 

Letters, v. 4, p. 423–425, doi:10.1098/rsbl.2008.0118.

Goldberg, C.S., Pilliod, D.S., Arkle, R.S., and Waits, L.P., 2011, Molecular 

detection of vertebrates in stream water—A demonstration using Rocky 

Mountain tailed frogs and Idaho giant salamanders: PLoS ONE v. 6, no. 7, 

e22746, doi:10.1371/journal.pone.0022746.

Jerde, C.L., Mahon, A.R., Chadderton, W.L., and Lodge, D.M., 2011, 

“Sight-unseen” detection of rare aquatic species using environmental 

DNA: Conservation Letters, v. 4, iss. 2, p. 150–157, doi:10.1111/j.1755-

263X.2010.00158.x.

National Center for Biotechnology Information, 2012, NCBI: U.S. National 

Laboratory of Medicine, National Center for Biotechnology Information 

database, accessed October 4, 2012, at http://www.ncbi.nlm.nih.gov/genbank/.

Takahara, T., Minamoto, T., Yamanaka, H., Doi, H., and Kawabata, Z., 2012, 

Estimation of fish biomass using environmental DNA: PLoS One, v. 7, iss. 4, 

e35868, doi:10.1371/journal.pone.0035868.

Thomsen, P.F., Kielgast, J., Iversen, L.L., Wiuf, C., Rasmussen, M., Gilbert, 

M.T.P., Orlando, L., and Willerslev, E., 2012, Monitoring endangered 

freshwater biodiversity using environmental DNA: Molecular Ecology, v. 21, 

iss. 11, p. 2565–2573, doi:10.1111/j.1365-294X.2011.05418.x.

 Waits, L.P., and Paetkau, D., 2005, Noninvasive genetic sampling tools for 

wildlife biologists—Review of applications and recommendations for accurate 

data collection: Journal of Wildlife Management, v. 69, iss. 4, p. 1419–1433, 

doi:10.2193/0022-541X(2005)69[1419:NGSTFW]2.0.CO;2.