Answers to Common Questions

A.

In order to develop an assay for a new species, we need the organism’s DNA sequence. In many cases, we can acquire species-specific DNA sequences from GenBank (a publicly accessible database of known genetic sequences) and develop an assay “in silico,” that is, based on information downloaded from the database.

However, because of natural variations in populations, we always seek to validate in silico assays in vivo using authentic, location-specific DNA. If we’re testing for an animal, this would require some kind of tissue sample. This is the only way we can ensure that the assay we develop truly identifies the target species in your location.

Take the cutthroat trout (Onocorhynchus clarki) as an illustration. Even if a cutthroat from Colorado is morphologically identical to a cutthroat from New Mexico, they may have differences in molecular taxonomy. So, if you’re in New Mexico, we’ll want fin clips from your cutthroats so that we can be sure the assay we develop is appropriate to your local animal population.

A.

Almost all eukaryotes (animals) have both a nuclear genome and a mitochondrial genome. The much smaller mitochondrial genome is independent of the nuclear genome and codes for proteins involved in energy metabolism.
In any assay, we’re seeking to determine whether a particular species is present. Each assay uses species-specific DNA markers to make this determination. Most geneticists develop assays for mitochondrial targets, because a single animal cell can contain a hundred mitochondria, but the same cell has only two copies of each nuclear gene (for a diploid organism).
However, because mitochondrial DNA is inherited only maternally, it does not work well for genetic purity assays or species identification assays aimed at differentiating between closely related organisms (such as distinguishing between different types of cutthroat trout).
And of course, because prokaryote target organisms (like bacteria) have neither mitochondria nor nuclei, we must find genetic markers for prokaryotes elsewhere—for example, in nucleoid DNA.

A.

Hybridization or genetic purity assays use a series of genetic markers to determine the relative genetic purity of a sample in relation to reference samples. The more markers are used in the assay, the more sensitive the results. For example, although blue eyes are commonly associated with northern European ancestry, possession of this single trait is not enough to establish a person’s Scandinavian heritage. For example, blue eyes are not unusual among the Tuareg people of North Africa. More traits must be examined.

In fisheries, wildlife, and conservation biology, this type of assay is used to aid wildlife agencies in determining whether fish within their jurisdictions are genetically pure or have been hybridized with invasive or introduced species.

For example, rainbow trout originated in the Mount Shasta region in California, but because humans moved them to numerous other locations, they are now found throughout the United States. Realizing that this kind of activity significantly alters the ecology of streams and rivers, many state wildlife agencies have been working to remove rainbow trout from nonnative habitats and to reintroduce or reestablish native fish species.

For years, Pisces has supported Colorado Parks and Wildlife’s (CPW’s) efforts to reestablish cutthroat trout—including the Colorado state fish, the greenback cutthroat trout—in their native habitats. This has involved checking many cutthroat trout populations for the degree to which they have been hybridized with rainbow trout or other trout species. The current technique used at Pisces (amplified fragment length polymorphism, or AFLP) checks 119 markers.

A.

Assays for determining genetic purity or hybridization are relative; that is, they must compare the samples being tested with samples from a reference population (or reference populations). Because of the natural genetic variation within any given species, a reference population should not be defined by a single sample. We use a program called STRUCTURE to analyze the results of AFLP assays. Because we have found that there is too much statistical noise in the results when fewer samples are used, we generally recommend that clients provide two to three dozen samples of each reference population.

In some cases, conservation biologists are not certain which population represents the most genetically pure lineage of a native species. In this kind of situation, the reference population can be adjusted over time as multiple populations are sampled, tested, and compared.

A.

Yes. Any living organism that has a genome, we can test for—including viruses, bacteria, and plant and animal species.

A.

Yes. To develop this type of assay, genetic markers must be identified on the sex chromosomes of the target species.

It is difficult for humans to identify the sex of many animals, either because their external genitalia are not obvious upon visual inspection or because juveniles of both sexes are indistinguishable. This can create challenges in conservation programs aimed at breeding animals with the goal of reintroducing them in the wild.

For example, at the St. Louis Zoo in Missouri, conservationists are breeding Ozark hellbenders, a threatened species endemic to the southeastern United States and one of the largest salamanders in the world. One clutch of hellbenders may include 200–300 eggs, and hatchlings cannot be sexed visually until they are two years old. Pisces can determine the sex of baby hellbenders using a tiny clip from the end of the tail.

Pisces improved a marginal older assay for determining the sex of juvenile Komodo dragons; the new assay requires just one drop of blood as a sample and works not only for Komodos but also for several other monitor lizard species.

A.

As a rule of thumb, we recommend using a statistic developed years ago by the American Fisheries Society (AFS)—assuming a 5% incidence level of a disease in an infinite population, in order to have a 95% confidence level of detecting the disease, you need to test 60 animals.

More broadly, the factors to consider are (1) What is the (assumed) incidence level of the disease? and (2) How certain do you need to be of detecting the disease?

If you assume a lower incidence level, more samples are needed in order to have a 95% confidence level of detecting the disease. For example, if you assume a 2% incidence level and require a 95% confidence level, 150 samples are needed.

Of course, we can test as many samples as you have the time to collect and the budget to cover. But practical considerations will likely prevail, depending on your situation. If you want to know if chytrid fungus is present in the amphibians in a particular nature preserve, the AFS rule of thumb may be appropriate. However, if you are breeding boreal toads or Panamanian golden frogs for release in the wild, you may decide to test your entire captive population.

A.

While eDNA is a powerful tool in fisheries, wildlife, and conservation biology, it does have limitations. In theory, a higher number of samples leads to higher certainty, but in practice, things are more complicated.

If we test a sample and get a positive for the target species, we know that the target species is (or was) present in the environment in question. However, if we get a negative, we only know that the target species was not present in that sample; we cannot prove absolutely that the species is not present in the environment.

A number of factors affect eDNA testing, including the type of organism, the time (e.g., season) of sample collection, and the environment (stream, reservoir, etc.).

Some organisms shed DNA material more consistently than others. Fish swim around their environment, and mucus on their skin sloughs off, leaving DNA evidence of their passing. But we tested an eDNA sample taken by a biologist right next to a turtle and got a negative for that turtle species—turtles apparently do not shed organic material at the same rate as fish.

In other cases, an organism may shed a lot of DNA in one season but very little at other times of year. Zebra and quagga mussels are stationary and encased in a shell, but when the water warms up in the spring, they release millions of microscopic larvae called veligers. Consequently, it is much easier to detect them via eDNA in warmer weather.

Another complicating factor is the persistence of eDNA; in some cases, DNA can be detected decades after the organism it came from has died. This persistence may be a boon if you’re conducting historical research but a bane if you want to know if you’ve cleared an invasive fish species from a stream.

If you let us know what organism you’re targeting, we’ll give you our recommendations based on our experience and what we know from the literature.

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Yes.

We can test for 3–4 different target species in a single reaction in one assay.

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The typical DNA extraction protocol provides 200 microliters of DNA extract; we need either 2 or 4 microliters to run an assay. Therefore, we have enough DNA from one sample to run at least 50 tests.

A.

Yes.

If we acquire 200 microliters of DNA extract from one sample and only use, for instance, 2, 4, or 25 microliters of it, we archive the remainder by freezing it. We have pulled samples out of storage and successfully tested them for clients years after the samples were taken.

One interesting case involved trout whirling disease, which is caused by a parasitic organism. The parasite lives half its life in fish and half in tubifex worms on lake and stream bottoms. The four existing tubifex lineages in North America have different sensitivities to the parasite. At the request of a CPW researcher, we used six years of archived samples to create a timeline showing the shifts in the prevalence of the tubifex lineages as the incidence of trout whirling disease increased.

A.

No, because the advantages of droplet digital PCR (ddPCR) are generally not relevant in conservation biology.
In qPCR, the entire reaction volume (usually 10 or 20 microliters) goes into one reaction. In ddPCR, the same 10 or 20 microliters are spread out into thousands of separate droplets, and an endpoint PCR reaction is carried out in each droplet. In qPCR, the amount of target DNA in the sample is quantified by measuring the fluorescence of the entire reaction volume and comparing it to a standard curve. In ddPCR, the apparatus tallies the number of droplets that have a positive reaction optically (like pixels on a digital camera sensor) to provide an absolute quantification of the target DNA without the need for a standard curve.
Although ddPCR has some advantages over qPCR, we continue to use qPCR for the following reasons:

A.

Very simply, eDNA samples are collected by filtering as much of the environment as possible—whether that’s air, water, or dirt. The filter catches cellular debris; you send us the filtration media, and we extract DNA from it.

There are ways of collecting samples published in the literature, and there are procedures that we recommend. However, eDNA is a new enough field that there has not been much standardization of sample collection techniques.

Of course, the more of the environment you can filter, the higher the detection sensitivity will be. Some considerations naturally follow from this. How will you channel air, water, or dirt through your filter? Then, how much air, water, or dirt can pass through the filter before the filter is clogged? This is a function of the pore size of your filter and the nature of what you are filtering.

For more detailed instructions, please see the chart on collecting, preserving, and shipping samples on the DISCUSS page.

A.

This is not as important as you may think. Although there are many papers on this topic in the literature, in our opinion the results reported are largely specific to the species and environments considered and are not generalizable.

One important thing to bear in mind is that eDNA sampling does not collect naked strands of DNA, which are only about 10 angstroms across (about a billionth of a meter). Rather, eDNA sampling captures cells and cell debris, which are much, much larger. We usually recommend filters with a 5-micron pore size. The larger the pore size you are able to use, the more air, water, or dirt you can filter before the filter clogs.

For more detailed information, please see the chart on collecting, preserving, and shipping samples on the DISCUSS page.

A.

If you are using Pisces eDNA filters, we usually recommend leaving filtration media in the filter housing, keeping the sample frozen or refrigerated, and shipping it to us overnight. Samples collected using other apparatuses can be dried and preserved with desiccant beads.

For more detailed instructions, please see the chart on collecting, preserving, and shipping samples on the DISCUSS page.

A.

Probably less than you think.

Field biologists are used to thinking in terms of whole animals; molecular biologists, not so much. We need approximately 10 micrograms of tissue in order to be able to extract enough DNA to test a sample.

The National Ecological Observatory Network (NEON) asked us to species identify mosquitoes they were trapping. We were able to extract enough DNA from a single dried mosquito leg.

For more detailed information, please see the chart on collecting, preserving, and shipping samples on the DISCUSS page.

A.

When testing to determine if a species is present or not present in a sample, molecular assays are sensitive down to just three molecules of DNA. Keep this scale in mind; things that are not visible to the naked eye can contaminate your samples!

The Utah Division of Wildlife Resources sent us samples from a tiny, remote pond to be tested for zebra and quagga mussels. Because there is no boat traffic on the pond, there shouldn’t have been any mussels—but the assay returned a positive for quagga mussels. Why? The field biologist collected the sample in a plankton tow net and set the net down on the seat of his truck; a week earlier, he had set a mussel-encrusted length of rope down on the same seat. That was all it took to contaminate the sample!

A.

Although the DNA in your samples can persist through many circumstances, there are some things that will render samples useless:

A.

Genetic samples need to include enough tissue (approximately 10 micrograms) for DNA or RNA extraction. Samples take many different forms; our database of sample types includes more than 200 items.
Some common sample types include:

A.

Individual swab samples (typically for chytrid fungus testing) allow us to test individual animals for a pathogen and, therefore, give the finest resolution. Pre-pooled samples and samples pooled at Pisces provide additional options that can stretch your budget and may be more cost effective if you don’t need the level of detail afforded by individual samples.
Pooling works as follows. Up to eight swabs can be placed in one larger tube with ethanol. We vortex the tube to extract all the chytrid fungus spores into the ethanol. Then, we run an assay to detect whether there was chytrid fungus among the eight swabs. This is an excellent option if you need to screen a large number of animals; it can reduce your costs nearly fourfold over individual testing.
We refer to samples that you pool before sending them to us as pre-pooled. We refer to samples that we pool for you as pooled at Pisces. Pooling at Pisces offers a testing resolution that is intermediate between individual testing and pre-pooling. When we pool at Pisces, we hold back a small volume of the liquid from each individual sample before pooling. This gives you the option to have individual samples from a pool tested if we get a positive for that pool.
For example, suppose you swab 120 amphibians for chytrid fungus in the Amazon rainforest:

A.

Yes. We sell collection tubes, swabs, and filters.

For more detailed information, please see the chart on sample collection supplies on the DISCUSS page. The chart includes links to the supplies we sell as well as those sold by large supply houses like VWR and Fisher. The minimum quantities sold by these vendors are often far more than our clients need; in such cases, you can purchase smaller quantities from us.

A.

Yes. We have positive controls for 6–12 species.

For a list of the positive controls we sell, please see the SUPPLIES page. We are always adding new products, so if you don’t see what you’re looking for, please inquire to find out what’s in development.

A.

Assuming that a sample was not contaminated either in the field or in the laboratory, a positive result from an eDNA assay indicates that the target species was present in the environment. However, a positive result does not provide any indication of when the target species was in the sampled environment; nor does it indicate whether the organism that shed the detected DNA was alive or dead at the time of sample collection.

A negative result provides less information; it only indicates that DNA from the target species was not present in the sample—it cannot prove that the target species was not in the sampled environment.

Consequently, in planning your study and sample collection, it is important to consider what both positive and negative eDNA test results will mean in the context of your work. For example, will a low-level positive result for Dreissenid mussels trigger significant action, like shutting down a lake?

For more discussion of factors involved in eDNA testing, see How many eDNA samples do I need to test to be sure a target species isn’t present in a particular location?

A.

No.

Although the copy number does refer to a quantity—the number of target DNA molecules we detected in a sample’s DNA extract—what we are able to determine from a qPCR eDNA assay is qualitative in nature.

In other words, although a higher copy number probably indicates more individuals of the target species in the environment than a lower copy number, there is no way of determining precisely how many individuals of the target species were present in the environment.

A.

The typical turnaround for test results is two weeks. At the end of summer field season, turnaround time may be longer, but we do offer priority turnaround service (one week or 48 hours) for an additional fee. Please contact us for details.