Using Environmental DNA (eDNA) to Monitor Aquatic Ecosystems

Environmental DNA monitoring – eDNA – is at the vanguard of a new wave of technologically advanced monitoring efforts. With roots in soil and paleoecology, eDNA was first used to detect a multicellular aquatic organism– the invasive American bullfrog Lithobates catesbeianus– in a landmark scientific paper by Ficetola et al. in 2008[1]. By applying the widely used, yet highly sensitive, polymerase chain reaction (PCR) – a reaction that ‘amplifies’ a specific DNA sequence in a sample, only if the species’ DNA is present in however little amount – Ficetola and colleagues detected the frogs’ DNA in sediment filtered out of the water column. To understand why, a current working definition of eDNA – adopted by most ecologists – will illuminate: eDNA is “…genetic material obtained directly from environmental samples (soil, sediment, water, etc.) without any obvious signs of biological source material” (Thomsen & Willerslev 2015[2]). Examples of how eDNA is shed by an organism – e.g., here a midland painted turtle Chrysemys picta – are illustrated in the figure below.

Blog pic 1

In the figure, DNA-containing cells are constantly or periodically shed from the internal linings of the turtle’s gut, reproductive system, through regurgitation of food, the replacement of skin cells and mucus, the egress of waste materials, and through the release of sex cells (i.e., sperm and eggs). Once in the water, DNA is somewhat protected within cells. Eventually, cells are broken down and DNA is released into the aqueous environment whereupon it is to be found in solution. Although eDNA is depleted through a number of biological, chemical and physical processes, it will keep being replenished if the organism is still to be found living in the vicinity. That is to say, the signal of eDNA will be stronger the closer it is to its source, and will also increase when there are more individuals in a local population, if the volume of water remains the same. Therefore, an eDNA signal can be a reliable indicator of the target species’ presence in a given habitat.

What are the chief benefits of eDNA monitoring? First of all, there is no need to physically sample the species, thus minimising disturbance associated by the targeted monitoring of live creatures that are sensitive to stress. Furthermore, as only water samples are taken, and by few people, there will be a decrease in the environmental footprint associated with monitoring efforts per se. Because PCR is a highly sensitive molecular biological assay, eDNA surveys tend to have much higher sensitivities to be able to detect rare or cryptic species than conventional methods (e.g., Schmelzle & Kinziger 2016[3]). As a result, species-specific eDNA surveys are also potentially much more cost-effective than conventional techniques. Furthermore, anyone can take a water sample, following simple instructions, democratising and facilitating citizen science projects across the globe. Indeed, the current crop of companies that offer eDNA detection services are predicated on a model of water samples being collected by lay and technical personnel for processing back in a central laboratory.

However, as eDNA is a nascent technology, uncertainty exists over some of the conclusions drawn from early eDNA studies. However, these issues (i.e., sources of ‘error’) are under ongoing scrutiny by scientists, including here at Precision Biomonitoring, to minimise their impacts. As such, all potential sources of error, as they are currently understood, must be acknowledged and incorporated into any technical development (i.e., design, production and validation of species-specific PCR assays) or standard operating protocols for field surveys. For example, organisms are liable to move throughout their lifetimes. Seasonality has shown to be a strong factor in eDNA detection success (de Souza et al. 2016[4]). It is imperative that surveys are conducted with a thorough knowledge of a species’ ecology, including insight into current distributions and habitat preferences, otherwise inadequate surveying will lead to a false negative result, i.e., inferring a target to be absent when it actually is present; just undetected. Failure to account for false negatives can result in severe financial repercussions if infrastructure projects are subsequently halted, put on-hold or abandoned due to the rediscovery of the target by an intrepid ecologist or member of the public. False negatives can also result from improper assay development, the underestimation of within-species genetic diversity at PCR amplification sites, and by the current disjointed process by which eDNA samples are processed by the majority of eDNA practitioners.

As noted previously, eDNA will decay if left exposed to natural world processes. Therefore, collected eDNA is at risk of post-sampling decay, as there would be no mechanism for eDNA replenishment in the collection vessel, reducing the eDNA signal and potentially failing to garner a positive PCR result. Therefore the risk of eDNA degradation during sampling – particularly on hot, sunny days – and in transit from the field to the laboratory, is highly significant. Inappropriate storage may also destroy eDNA (e.g., water crystal formation during freezing may ‘shred’ DNA molecules). To compound the status quo further, despite the best efforts of contemporaneous laboratories, there remains a significant risk of false positive PCR results mediated by the transportation of aerosolised DNA particles among labs within buildings through ventilation pathways. Most eDNA practitioners seek to physically separate the processing of eDNA samples (e.g., filter papers or precipitated water samples) with downstream PCR detection, but even that is far from fool-poof.

Here at Precision Biomonitoring, we are set to unveil a state-of-the-art platform that will seek to eliminate, or minimise, these sources of eDNA analytical and sampling error, through the eradication of transit stages to a central laboratory and the application of standard operating procedures. Moreover, we will further the cause of a democratised biomonitoring field in which no technical specialty is required to conduct sophisticated PCR-based species-specific assays. Our system, using bespoke PCR assays, will yield PCR eDNA results in real-time (< 2 hours from water sampling to PCR read-out), which can then be immediately disseminated to colleagues via the cloud.

It is our aim to give those working at the coalface of biodiversity monitoring (from professional ecologists to local citizen science projects), the power to conduct highly rigorous, and potentially highly coordinated, targeted eDNA surveys to better vouchsafe our world’s biodiversity heritage for all generations to come.

[1] Ficetola et al. (2008). Species detection using environmental DNA from water samples. Biology Letters, 4, 423-425.

[2] Thomsen & Willerslev (2015). Environmental DNA – An emerging tool in conservation for monitoring past and present biodiversity. Biological Conservation, 183, 4-18.

[3] Schmelzle & Kinziger (2016). Using occupancy modelling to compare environmental DNA to traditional field methods for regional-scale monitoring of an endangered aquatic species. Molecular Ecology Resources, 16, 895-908.

[4] de Souza et al. (2016). Environmental DNA (eDNA) detection probability is influenced by seasonal activity of organisms. PLoS One, doi: 10.1371/journal.pone.0165273

Author: precisionbiomonitoring

Precision Biomonitoring provides the environmental and aquaculture sectors with molecular point-of-need tools that detect environmental DNA (eDNA) for the identification of invasive, keystone and at-risk species, and pathogens in aquatic systems. The molecular point-of-need tools are simple to use, give more definitive species identification, under 2 hours and at 25% of the costs of traditional methods, and without having to send samples to a central lab.

Leave a Reply

Fill in your details below or click an icon to log in: Logo

You are commenting using your account. Log Out /  Change )

Google photo

You are commenting using your Google account. Log Out /  Change )

Twitter picture

You are commenting using your Twitter account. Log Out /  Change )

Facebook photo

You are commenting using your Facebook account. Log Out /  Change )

Connecting to %s