Yes, eDNA is a great surveillance tool for invasive species because it can often detect them at much lower populations levels than conventional surveys would. If the species in question is in the reference library, we will be able to identify it in metabarcoding datasets. Single-species qPCR tests can also be used to screen for the presence of particular species. There are good qPCR tests for species such as zebra and quagga mussels, and signal crayfish, and in the US eDNA has been used extensively for tracking the invasion of Asian carp in waterways around the Great Lakes.

Metabarcoding can be used to generate high-resolution datasets on the meiofaunal invertebrates (nematodes etc) and microorganisms living within the ocean floor sediments of areas earmarked for impact or restoration. These small organisms are numerous and respond quickly to impacts. As such metabarcoding can be used to track species biodiversity and community composition over time in relation to e.g. drilling impacts, and or restoration efforts. eDNA can also provide data on fish and marine mammal communities, and collecting water from different depths in the water column can reveal the different communities at each level.

Large publicly available reference libraries do exist. These include the National Center for Biotechnology Information (NCBI) database, also known as Genbank, and the Barcode of Life Database (BOLD). Although these databases are often incomplete for poorly studied areas, they can be augmented with data from local or private databases and also through barcoding studies (where tissue or swabs from animals identified in the field are sequenced).

Accordion SaThe average half life of eDNA is about 48 hours but this varies depending on environmental conditions and small amounts of DNA have been known to last for weeks. The degradation of the DNA is slowest when it’s cold, dark, or when the DNA is bound to sediment, and faster in more acidic environments. Collins et al., 2018 provides a good overview of eDNA persistence in marine environments, and Li et al., 2019 showed that there was no detectable eDNA signal 48 hours after removal of fish from small lakes. Findings are typically that eDNA analysis gives a good snapshot of contemporary communities and not historical records. mple Description

AccordionWhile eDNA has been known to theoretically travel many kilometres in rivers, its constant deposition and decay makes the probability of detection increasingly small over larger distances and depending on the size of the river and flow rates. In marine environments it was originally thought that water/DNA would be so well mixed that there would be limited spatial resolution. However, this was found not to be the case. In fact DNA from animals in specific habitats can be detected using eDNA in marine environments with surprisingly good spatial resolution, at least in shallow to moderately deep waters (see Port et al., 2016 for an example). In deep water, thermoclines, haloclines and strong currents could affect eDNA and as such multiple samples are recommended at different depths for best results. Sample Description

Yes, this is definitely recommended. We suggest that triplicate or at minimum duplicate samples are taken at sites in rivers and marine environments for best results. This will maximise detection of rare species while also building confidence in the replicability of the approach for recovering the more common species. In still water (ponds and lakes) DNA does not always mix well so a subsampling approach should be adopted whereby subsamples are collected from a section of the shoreline and mixed before filtering.

A very small amount! So be careful about eating fish for lunch if you’re going sampling. Possible environmental sources of contamination include fishing bait and waste water from kitchens, which needs to be taken into account when choosing sampling locations – especially in populated areas.

This will depend on the level of mixing in the water body, which may vary seasonally. For water bodies where there is a lot of mixing (i.e. rivers) eDNA is more homogeneously distributed. In still water (i.e. ponds) then there is much more spatial heterogeneity and so the probability of detection is lower if water is taken from a single point, and appropriate (sub)sampling design is key to cover all microhabitats. However, see Lawson-Handley et al. (2019) for a comprehensive study of spatial dynamics of eDNA in large lakes. This study concluded that shoreline sampling was sufficient to detect all species in Lake Windermere during the winter when more mixing occurred, and only missed one species (Arctic Charr, which lives deep in the middle of the lake) during summer when there was less mixing.

eDNA in the marine environment is much more dilute than in freshwater systems and so the detection probability for any species in a given sample will be lower (note other survey methods are also less sensitive in the open ocean), and means it is important to filter more water and collect a greater number of samples in the marine environment. Generally, the more samples you can collect, the more representative and comprehensive your dataset will be. At the moment it’s very difficult to say how many samples are needed for a comprehensive survey in the ocean, or what depths these should be collected from, and spatial interpretation is also difficult because of the complexity of currents and other aspects of oceanography – there is definitely the opportunity for lots of large-scale research here! However, eDNA does still provide a lot of data in the marine environment and compares very favourably with alternative tools in this regard.

This is dependent on multiple site factors and the study question. In river systems for example, confluence points in rivers represent areas of water and DNA mixing of two potentially distinct fish communities. Therefore a point collected within the tributary as well as upstream and downstream of its confluence with the main stem may be required to determine fish community equivalence in the different parts of the river system. Similarly this logic could be applied to barriers or dams in rivers. Other site factors to consider would be pollution sources, major land use or habitat differences and differences in riparian vegetation.

eDNA provides replicable and meaningful data on relative abundance of aquatic organisms, but not absolute abundance (except in some very specific cases where extensive calibration has taken place – see Levi et al., 2018 for an example of using eDNA to count salmon in Alaska). Some behavioural factors affect the amount of DNA given off by a particular species at a particular time (e.g. spikes of DNA associated with breeding or high levels of activity), and there are some interspecific differences in DNA shedding – for instance, small active fish tend to give off more DNA than large, slow ones. In rivers, if you detect a small trace of a species it is difficult to tell whether this means there are a small number of individuals close to the sampling point or a larger number some distance upstream. That said, overall the rank abundance of species based on eDNA data tends to be a good reflection of the community.

A single sequence does not represent a single individual. This question is linked to the wider discussion on whether sequence count is correlated with abundance. See above.

There is potential. We currently use relatively conserved and high copy regions of the genome to identify taxa, but the inherent properties of these DNA regions, which makes them well suited for species identification, also makes them less than ideal for individual/population level assessment. You would need to look at a more quickly evolving region of the genome, but these tend to be much more difficult to work with for various reasons. We’ve made some initial forays into this and some studies suggest that it may be possible to identify different haplotypes within the same species, but essentially this is still very much in the research phase. See Sigsgaard et al., 2020 for a recent review and synthesis of progress in this area.

The persistence of eDNA is typically short lived and new eDNA will typically overwhelm remnant eDNA. That said, traces of eDNA can theoretically be detected from older sources, but these will likely be trace amounts, present only in very small fragments and screened out following quality control. A bigger risk is environmental contamination from fishing bait or wastewater, which should be taken into account when designing sampling campaigns, especially in populated areas.

In some respects yes, but this will depend on the target groups being studied. Studies we have conducted have indicated that more species are typically detected in the wet season in the tropics. This is likely as a result of more DNA being washed from riparian areas into rivers in the wet season. However increased water volumes following heavy rainfall may also slightly reduce sensitivity by diluting the DNA signal. These finescale spatial dynamics are still being investigated across a number of different projects, although eDNA gives good data in all seasons!

Contamination in the lab is a risk. A unidirectional workflow with spatial seperation of the different steps is important. Also a dedicated cleanroom with UV light and steril benches as well as gloves, face masks and overalls are necessary. Negative controls of all steps can track possible contaminations.

qPCR (probe-based) assay is often more reliable than PCR (end-point PCR visualised on a gel) because the binding of the probe represents a third point (in addition to the two primer sequences) at which the target sequence must be matched, thereby reducing the risk of non-target amplification causing false positive results . qPCR is much faster to carry out in the lab than metabarcoding, which requires a substantial amount more lab work and computational processing. However, because qPCR and other types of single-species screening assay are ‘blind’ tests which give a positive or negative result without providing a sequence to confirm species identity, the assays need to be extremely rigorously validated before you can rely heavily on the results for decision-making in management contexts (Thalinger et al., 2020 gives a comprehensive overview of this). This can take a long time and is an expensive process. Metabarcoding can provide data on many more species than qPCR, and because it generates the DNA sequences that are used for determining species identity, high confidence can be ascribed to detections even early on in the assay validation process.

Species not yet in the reference database cannot be definitively identified without obtaining a reference sequence. However, in many cases there are sufficient references from congeneric species to make a confident identification at genus-level. In a scenario where only one representative of that genus is expected in the sampled community, a putative species label can be associated with the metabarcoding sequence we generate, pending confirmation from reference material. Species unknown to science would similarly be identified as best we can given their similarity with available reference data, but we cannot distinguish between gaps in the reference database and gaps in taxonomic knowledge.

We do sometimes see different sequences assigned to the same species, however our assays are chosen to target species-level variation across a broad taxonomic group. These are therefore relatively slow-evolving markers that do not coincide with regions targeted for conservation genetics. Population genetics is theoretically possible with this sequencing technology but requires significant R&D in the selection and testing of the marker for each target species and is not something we currently offer as a service.

Operational Taxonomic Units (OTUs) are often part of the data processing pipeline. This is a taxonomy-free clustering approach based purely on how similar the sequences generated are to one another. The threshold at which the clusters are defined varies between assays because of the properties of different gene regions that are used. Each OTU is treated as a species-level entity in attempt to make a taxonomic assignment for each but this will not always be possible at species level. All OTUs are included in estimates of diversity, regardless of whether it is possible to assign a species label.

The technology is very sensitive, so theoretically there is potential for this type of natural contamination. We would expect that this is very rare though. If there is doubts about the presence of a specific species, further investigation such as a second sampling can be helpful.

Yes, qPCR tests for both Batrachochytrium dendrobatidis (Bd) and Batrachochytrium salamandrivorans (B-sal) chytrid species exist.

There are many reasons why morphological taxonomy is still important. For a start, molecular taxonomy in the sense that we use it relies on a reference database underpinned by traditional taxonomy – and this will always be the case. Molecular taxonomy can’t on its own discover and describe new species – that will always rely on traditional taxonomic skills. There are simply not enough taxonomists in the world to be able to generate the monitoring data that we need at the scales we need it to underpin decision-making – especially in the tropics – so there is a need for new tools and approaches. We believe that widespread adoption of molecular tools will actually highlight the need for good taxonomists.