Environmental DNA Applications

Could this collection method be used to track invasive species and their spread?

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 (e.g. Jerde et al., 2013).

For the marine environment, qPCR assays are continually being developed for the early detection of invasive non-native species (e.g. Holman et al. 2019; LeBlanc et al. 2020).

Is there any update of regulatory acceptance of eDNA methods for fish by the Environment Agency in England?

The Environment Agency is itself using eDNA for monitoring fish communities, and is working on a tool that would generate a Water Framework Directive (WFD)-compliant index score for lake fish communities, based on the work carried out in collaboration with the University of Hull (e.g. Lawson-Handley et al., 2019). It is best to check directly with the agencies with regard to specific projects.

I’m interested to know what potential eDNA has in the deep sea. E.g. for EIA for deep sea mining.

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.

Find out more

Are reference libraries shared between institutes? I.e. is there a large shared global database (publicly) available so conservation/research all over the world can be shared and help other studies in remote areas?

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, and these are used as the basis for our species identification pipelines. However, Genbank in particular (which is the most extensive database) is known to contain many errors, so we have applied our own careful curation and quality control measures to a downloaded version and it is this that is used in our pipelines. 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).

DNA in the Environment

How long does DNA last for in the environment? Is there a risk of finding something that is no longer there?

The 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.

How do water currents affect the results? How does it work with movement of water downstream in rivers or in currents in the sea?

While 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 our own experiments, based on fast flowing European rivers, we have found that DNA transport distances are typically less than 1 km.

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.

Do you have examples of where you can demonstrate that eDNA can differentiate between different points in a fast flowing river?

In our Amazon baseline study we found that samples from consecutive sampling locations (c. 10 km apart from one another) were quite independent of one another even in a large river that is several hundred metres wide and flowing quickly. Shoaling species were useful here because you would see a large amount of DNA from them at one point and then no detection at the next location downriver. We did see some transfer of DNA where a natural barrier (a steep gorge) caused a significant change in species composition, and the sample taken just 1km or so downstream of the gorge still contained the DNA of the species in the upriver section. The river was flowing very fast here. In smaller, lowland rivers eDNA might integrate information over an area of up to around 1km upstream. Recent work from our collaborators in Belgium using cage experiments in river systems, revealed that eDNA quantities exhibited a steep drop off in concentration after 5 m from the source (Brys et al. 2020).

Is there a requirement for multiple kits for large water bodies?

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. In a lake typically one kit will suffice for 400m of shoreline, where subsamples are collected every 20m. Where budgets may constrain the number of samples that can be collected, we work with our clients to help design a survey that will maximise the amount of information given the constrained number of samples.

What methods should I use to collect water samples without risking contamination?

The best way to avoid contamination is to use our sampling kits and follow the instructions provided. We have a great set of resources for clients, and you can watch our aquatic eDNA sampling video on YouTube here.

NatureMetrics filters are fairly robust to contamination because the filter membrane is enclosed within a plastic housing. However, you need to take care not to introduce contamination via the vessel you use to collect the water from the waterbody or to hold it while you filter. For this reason, the NatureMetrics sampling kits contain a sterile single-use collecting bag as well as gloves to prevent the introduction of your own DNA into the sample. If you are using a bucket to collect and/or hold the water, this will need to be cleaned with a 10% solution of household bleach to remove any traces of DNA, the bleach should be disposed of responsibly and then rinsed thoroughly with clean, distilled water before sampling.

How small of a contamination is required to show in results?

A very small amount! So be careful about eating fish for lunch if you’re going sampling, we recently managed to tell one of our clients what he had eaten for lunch before taking the sample! 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.

Are there any potential issues with only collecting samples from the boundaries of a large water body?

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.

Regarding marine research, how can we be convinced that the sampling point can represent the wide range of study area? Do we have to take a lot of samples in proportion to the size of the area?

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. In one pilot study we did in the North Sea, just three eDNA samples detected ⅔ of the species that had been recorded in a 2-year netting survey that had cost £150k.

What criteria do you use to set a sampling point?

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.

Can you detect abundance?

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.

Does each DNA sequence represent 1 individual? For example, if I have 12 DNA sequences of the same animal species, does it mean 12 different individuals of that same species were recorded?

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.

Is there potential, now or in the future, to identify individuals within a population and calculate/estimate the size of a species population?

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.

How long can a filter kit be kept?

NatureMetrics filter kits don’t have a shelf-life so can be kept until you need to use them. Once a sample has been taken and the preservative solution added as described in the kit instructions, it is stable at ambient temperature for several weeks. Samples should not be left in direct sunlight.

Can your sample be compromised by remnant sources of DNA (e.g. traces of DNA held in sediment)?

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.

Is it better to take water samples just after it has rained in the UK to get more species?

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!

DNA in the Lab

Is there any chance of cross-contamination at the lab?

Contamination in the lab is a risk that we take extremely seriously. This is one of the advantages of being a commercial laboratory where we have full control over the use of our space and movement of people and equipment within the system, but it is one of the reasons that our costs are higher than those sometimes reported by academic research institutions. We operate a unidirectional workflow from kit preparation → DNA extraction → pre-PCR → post-PCR. DNA extractions from filters are carried out in a dedicated cleanroom facility where tissue samples are never handled, and which features positive air pressure with HEPA filters. Regular disinfectant schedules are in operation across all our labs, which includes a minimum of two daily cleans of all surfaces using chemicals that remove DNA before and after operations start. Surfaces are regularly cleaned between procedures to avoid cross sample contamination. For equipment (e.g. laminar flow hoods and pipettes) additional cleansing is carried out using DNA removal wipes. High intensity UV lights provide overnight irradiation in our laboratories, and UV light is also used to irradiate the flow hoods for 30 minutes prior to every PCR set-up. In addition to these steps, we operate a robust quality control system where negative controls are integrated throughout the workflow to check for contamination. If any of these negative controls show signs of contamination, the analysis is repeated. As a result of these measures we very rarely experience issues related to contamination in the laboratory.

You can find out more about our Facilities and the standards we’re proud to represent and uphold, when you step Inside NatureMetrics.

If you’re testing whether a single species is present or not, what is the advantage of using qPCR over PCR? And qPCR versus Metabarcoding?

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.

How does metabarcoding compare with classic DNA barcoding?

Although we use classic DNA barcoding for some things, for example when building reference databases, it is of limited use for biodiversity surveys for two main reasons. First, classic barcoding is too slow and expensive for generating large-scale data across diverse groups (e.g. arthropods and other invertebrates) because it requires a separate sequencing reaction for each specimen. Second, classic barcoding can’t be used on samples that contain DNA from a mixture of related species (e.g. environmental samples). This is why we use metabarcoding to bypass these limitations when generating large scale biodiversity data.

What type of preservation solution do you use?

For our eDNA filter kits we use a salt and detergent-based lysis solution. This solution is non-hazardous, stable at room temperature, and doesn’t have the logistical difficulties associated with ethanol, which is another type of preservative which can be used. Ethanol is required to be used currently for Great Crested Newt tests in the UK. We have a range of preservatives that we use for other types of sample e.g. soil and insects, which we chose depending on the project, location, speed from sampling to processing etc.

DNA Data Processing & Bioinformatics

What about species that are not (yet) in the reference library – species yet unknown to science?

Species not yet in the reference database cannot be identified at this time. We therefore strongly encourage clients to share lists of priority species with us during project planning so that we can assess whether reference sequences are already available. In some cases, this will inform which assay is used to process the samples. We encourage our clients to consider barcoding campaigns alongside eDNA sampling where practical to fill any important gaps in the database and improve the quality of the identifications we can provide.

Where species-level identifications are not possible, there may nevertheless be sufficient references from congeners 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 with reference to available reference data, but we cannot distinguish between gaps in the reference database and gaps in taxonomic knowledge.

Do you ever find new species while doing these surveys? How do you go about identifying them?

This is closely related to the previous question. We frequently encounter taxa where we are unable to make a definitive species-level assignment. This can happen where several species have identical sequences in the region targeted by the assay, or where there are gaps in the reference data. In these cases we make a taxonomic assignment at the lowest level at which we are confident, given the available reference data.

Can you give more details about the probabilistic species identification algorithm you use and are your methods public?

Our identification pipeline uses a published probabilistic algorithm. The algorithm accounts for gaps in the sequence reference databases by comparing the content of that database with expected taxonomic diversity, allowing the probability of a query sequence arising from an unreferenced species to be calculated correctly. This reduces the likelihood of overconfident assignments to species level due to database gaps. As the taxonomy is a key input to the algorithm, the probability of assignment can be estimated at each level and an acceptance threshold can be applied to ensure only high confidence assignments are retained.

Do you pick up sub-species variation and if not, is this a possibility in the future?

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.

What is your ‘species’ definition for species that lack matches in the reference database?

We generate Operational Taxonomic Units (OTUs) as part of our 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. OTUs are delimited at a level approximately equivalent to species in each case, however rates of sequence evolution vary between lineages. This is especially true where a very broad assay is used, so the clustering threshold used will more closely approximate species for some amplified taxa than others.

We treat each OTU as a species-level entity and attempt to make a taxonomic assignment for each. Taxonomic assignments will be to the lowest level confidently supported by the available reference data. These identifications can be revisited as new reference data are accumulated, potentially improving the achieved resolution for many OTUs over time. All OTUs are included in estimates of diversity, regardless of whether we can assign a species label.

DNA for Different Taxonomic Groups

Can water sampling detect chytrid fungus that affects amphibians?

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

Can water sampling detect otters?

Yes. We’ve detected otters (Lutra lutra) with both single-species qPCR assays and metabarcoding assays. We’ve even detected Neotropical otters (Lontra longicaudis) and giant river otters (Pteronura brasiliensis) in Peru. Otters do seem to be slightly underrepresented in metabarcoding datasets given how much time they spend in the water. This may be because a lot of eDNA originates from faeces, and otter latrines are on land rather than in the water. We can also ID otters from their spraints (and even use metabarcoding of the spraints to understand their diets).

Can you analyse algae eDNA?

We have made some initial forays into analysing algal eDNA, but the group is such a diverse and informally named group that a single assay for so many different evolutionary lineages makes it difficult. Nevertheless we have managed to amplify and sequence algal eDNA, but this pipeline is still in its infancy and the reference databases very incomplete, so we would regard this as being during the R&D phase.

Can you analyse coral eDNA?

Coral eDNA is a complex group with exciting applications and great potential. We are currently collaborating with a partner lab to validate coral specific primers for their rapid and accurate characterisation. The project is still in its infancy, but watch this space.

Can you analyse DNA from the soil?

We do analyse DNA from soil, from which we typically generate data on soil fauna, bacteria and fungi. These groups are incredibly diverse, which gives them great power to indicate even fine scale ecological changes. This is a very active area of research for us.

Meet our Soil and Sediment Expert; Dr Hayley Craig and our lead scientist, Dr Cuong Tang

Do you use leeches to collect vertebrate eDNA?

This is classic invertebrate-derived DNA (iDNA). While we’ve never directly handled leeches within our immediate lab team, we’ve analysed leech iDNA data before and our co-founder Prof. Douglas Yu has processed over 30,000 leeches in his lab in China in the hunt for a possibly-extinct antelope (Ji et al., 2020)! On top of this, we have processed invertebrates from Malaise traps to get a handle on the wider vertebrate communities that they interacted with.

Which other taxonomic groups can you look for other than vertebrates?

Currently, in addition to vertebrates, we can also analyse for Insects, Bacteria, Fungi, Crustaceans and Mussels. We have made some forays into detecting algae and diatoms – but these assays are still being developed and optimised. Diet analyses can also be performed on bird and bat faeces.

Explore our Services.

DNA-based Methods & Traditional Taxonomy

Do you think that traditional taxonomic skills and history research are still important? Should we be concerned that the use of eDNA could lead to a loss of taxonomic skills in the scientific community?

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.

NatureMetrics Kits, Products & Business

If I use a NatureMetrics kit, what results will I get back?

NatureMetrics will send you a report that summarises your results primarily in a species by sample table. We also provide quality control tests and checks that have been carried out. We can also send you the species-by-sample table in Excel format. This will give taxonomic identification at multiple levels and tell you how many sequences each species was represented by in each sample. For large and complex projects, we can also offer more extensive reporting and ecological analysis.

For non EU samples, how do you cope with Nagoya Protocol obligations on access to genetic resources? Are they a barrier?

They are an important consideration in many projects and one of the reasons that we have begun to form partnerships with in-country labs in many cases where we have opportunities for large projects in non-EU countries. Every country implements the Nagoya protocol in a different way so some places are easier to work in than others and we have to take it on a case-by-case basis. We invest in building up strong networks of governmental and non-governmental stakeholders in the regions where we work, and supporting the development of key resources such as national reference libraries.

Interesting to hear there is a Peruvian lab that you collaborate with and that has the capacity/equipment to do these types of analysis. Do you work with labs in other parts of the world?

Yes, we are in discussions with labs in Liberia, Ghana, Mozambique, South Africa, Singapore, Malaysia and Indonesia. We also have contacts in labs in Brazil and Colombia. Quality control is really important when working with partner labs so we have to make sure that we have sufficient resources to cover this when we enter into new partnerships.

Please can you share any journal article references you have from your work to date?

You can find these on the Publications section of our website.

What costs are associated with the lab processing and how do overall costs of these types of surveys compare with conventional surveys, for example fish nettings?

Analysis of an eDNA sample costs £275 + VAT and we provide discounts for conservation NGOs and researchers. This includes the sampling kit and full laboratory analysis with QC testing and reporting. Electrofishing is an order of magnitude more expensive and often detects fewer species than a single eDNA sample (small, bottom-dwelling fish such as bullheads and sticklebacks are routinely missed by electrofishing). Cost comparison with netting depends very much on what kind of boat you would need to use for the netting and whether you were in an environment where you could collect eDNA samples from the shoreline (e.g. in a lake). Because eDNA is more sensitive for fish surveys than netting is, even if you have to use a boat to collect the eDNA samples, you will need less field effort to capture the same amount of data (see Hänfling et al., 2019 for comparison of catch per unit effort in Lake Windermere).

Where can I find out more about a career in eDNA?

Step Inside NatureMetrics and find out more about life in eDNA roles, you can also see what vacancies we have available in our Careers section.

Future Research & Development

Are there options being tested that skip the PCR step that may bias some results due to primer bias?

PCR, the stage that you amplify the target region of the genome, relies on designing accurate primers that can bind to a complementary part of the target genome. If there are any mismatches between the primers and the target region then the binding is less efficient and these inefficiencies are carried through the process. These binding inefficiencies will differ among taxa and this is what’s called primer bias. At best, primer bias will result in slightly fewer sequences being detected for that taxon, but at worse it could result in false negatives. PCR free methods (i.e. that don’t have that amplification step) avoid the need for primers and are based around the sequencing of genomic DNA and then the subsequent stitching together of that information for the purposes of identification (among other things). Our co-founder, Prof. Douglas Yu, recently published a paper on PCR-free metagenomics of pollen (Peel et al., 2019), but for the moment this remains too expensive to be a commercially viable option for routine monitoring. Moreover it is not a good solution for eDNA samples as the concentration of target DNA in these samples is just too low for PCR-free methods to provide useful data. We are keeping an eye on this exciting area of research and are sure it will progress quickly.

eDNA from Water

What’s the shelf life of the aquatic eDNA kit before taking the sample?

There is no shelf life for aquatic eDNA kits, but we do recommend that you store them at ambient temperature and out of direct sunlight.

How long after taking aquatic eDNA samples is the DNA safe before it starts degrading?

Analysis of our filters have returned comparable data after 6 months of storage (after collection) at room temperature. We have shown that longer storage up to 3 months has little impact on pond/river samples, but a bigger impact on marine samples. While the integrity and robustness of the samples is generally high, we would recommend that filters are sent back to NatureMetrics as soon as possible so that they can be processed in a controlled environment.

How many kits do I need?

This will depend on the system sampled and the target taxa, but generally speaking we recommend that triplicate or at minimum duplicate samples are taken at sites in rivers and marine environments for best results. Increased sampling effort, as with more conventional methods, 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. In a lake typically one kit will suffice for 400m of shoreline, where subsamples are collected every 20m. Where budgets may constrain the number of samples that can be collected, we work with our clients to help design a survey that will maximise the amount of information given the constrained number of samples.

How quickly can I get results?

We aim to return results for single species qPCR assays within 10 working days, and metabarcoding results within 6-8 working weeks. We will return results when we have them processed – even if this is earlier than the pre-defined due date. Fast track services are available and can be discussed.

How much does it cost?

The cost of kit will depend on the number and type of analyses you require. This can range from one single species analysis to multiple multi-species analyses depending on your focus questions. Please to discuss your project and we can provide you with a quote.

How quickly can I receive the kits? And how do I store them safely?

Domestic orders placed before 12 will be sent out for next day delivery. The kits can be effectively stored at room temperature. For international delivery we aim to ship straight away, but depending on the country, delivery can take up to a week. Please contact us (eDNA-lab@naturemetrics.co.uk) for more information.

Does eDNA from Water comply with the Water Framework Directive (WFD)?

Integration of DNA-based methods for the WFD is the sole focus of EU COST Action DNAqua-net (www.dnaqua.net 2017-2020), which NatureMetrics have been actively involved in, working collaboratively with academics, other service providers and end users. Activities within the DNAqua-net program have made important steps in building consensus among the scientific community with regards to molecular biodiversity assessment tools in aquatic habitats, but as yet they are not compliant with the WFD and so should be used in conjunction with approved methods.

Why should I use eDNA compared to other traditional methods?

eDNA has been shown to be more sensitive than traditional methods for several different organisms (e.g. GCN, fish, etc.). in head-to-head comparisons, eDNA surveys typically return more diversity per unit effort. Additionally, the methods deployed are non-invasive, user-friendly, and the data is auditable and objective.

Soils & Sediments

How much will it cost to do soil metabarcoding?

The cost of kits will depend on the number and type of analyses you require. This can vary depending on your focus questions. Please contact us to discuss your project and we can provide you with a quote.

What can I measure from soil and sediment samples? 

We can generate data on the fauna, bacteria, and fungi present in soils and sediment samples. This data can be used to assess the richness and diversity of your soil/sediment communities and can also be used to assess differences in community composition between sites or changes in composition with time. 

How do I preserve my soil/sediment sample for transportation to the laboratory?

There are two options for sample preservation. The best option is to keep samples in a coolbox with ice packs while in the field and for express transport to us. If not shipping the same day, samples can be kept in a fridge for a few days before transport.  

If cold storage is not logistically feasible for your project, we can provide a preservation buffer which is added to the samples during collection in the field. These samples can be kept at room temperature in the dark and gives you a few weeks to get the samples to us. 

What are the soil and sediment metabarcoding results going to tell me?

The metabarcoding results can be used to assess the richness and diversity of your soil/sediment communities. It can also be used to assess differences in community composition and diversity between sites/treatments or changes with time. 


What’s the shelf life of the GCN eDNA kit before taking the sample?

Natural England state that the kits are stable for 3 months. The kit contains ethanol, a salt, and a DNA marker used to monitor degradation and inhibition. With the correct storage (cold and dark), these components are stable for long periods of time and we have returned positive GCN detections from kits that our clients have forgotten about for 6 months!

Will my sample degrade?

The sample is preserved with ethanol, which acts to slow, but not eliminate, degradation of any DNA. Refrigerating samples and keeping them in the dark will also reduce the rate of decay. We monitor DNA degradation by use of a DNA marker that is already in the kits before you use them to sample. We recommend that you return the kits to us as soon as possible so that we can process/store in a controlled environment. When stored correctly, the timeframes associated with GCN season are not a huge concern.

Macroinvertebrate & Invertebrate Services

What can it detect?

We use a universal primer set to amplify and sequence all of the DNA present in the sample. We estimate that up to 75% of the UK’s macroinvertebrate biodiversity has a reference, and so we expect that most macroinvertebrate can be identified to species-level. There are known biases associated with certain taxa (e.g. soft bodied taxa will tend to be over-represented compared to species with hard carapaces), so please get in touch if you want to discuss these further.

What can the data be used for?

Our invertebrate services overcome the taxonomic bottleneck associated with analysing bulk invertebrate samples. Rather than focusing on one target group at a time, we use universal primers and sequencing technology to characterize whole community samples. We provide species-by-sample tables that can converted to generate presence-absence data, which can be used to baseline or assess changes in diversity across different treatments. The sampling design of the experiment can be tailored from the outset to help answer any number of ecological questions, so please get in touch and we can help.

Is it good for the Water Framework Directive (WFD)?

Invertebrate Metabarcoding returns presence-absence data, rather than abundance data, and can’t currently be used for the Water Framework Directive (WFD) monitoring of benthic macroinvertebrates. It is, however, an excellent option for processing large numbers of benthic macroinvertebrate samples for projects outside the scope of the WFD.

What samples do you accept?

A sample constitutes a well preserved pot (< 250 ml) of invertebrates. Samples larger than this will either be split or subsampled.

How do I sample?

The methods we use are blind to the sampling method and SOP.

What preservation can I use?

We recommend you use ethanol. Some conventional collecting fluids (e.g. ethylene glycol), aren’t great at preserving DNA, and some should be avoided at all costs (e.g. formalin). So if you’re not sure what to use, please get in touch and we’ll help you find an alternative.

What traps can I use?

You can use whatever trap you like. To date, we have successfully processed kick net samples, Malaise traps, pitfall traps, CWAC samples, vane traps, interceptor traps, pan traps.

Species from Faeces

From what species do you analyse?

We have a well validated bespoke pipelines that can identify all 18 native bat species and that can confirm otter spraints. Other species can be identified using more universal sequencing assays, so please get in touch with us so that we can help you (eDNA-lab@naturemetrics.co.uk). We currently only accept domestic samples.

How much faeces do I need to collect?

Generally speaking, more faeces = more DNA. The degraded nature of these samples, especially by the time they are collected, leads to poorer quality and lower quantity DNA. Having said that, we have returned conclusive identifications from a single pipistrelle pellet.

How do I store the faeces?

Dry and in an airtight container. For bat pellets we recommend you use 1.5 ml tubes, which we can provide if needed.

How quickly can I get my results?

We aim to return results to you within 10 working days.

Citizen Science

Where can I find out more about eDNA citizen science?

Visit our eDNA Discovery Lab website for all our information on this topic, and get in touch if you would like to discuss becoming a citizen scientist or organising a larger project.

Who can get involved?

We’ve had great success helping Citizen Scientists to leverage eDNA methods to discover animal communities in water samples, using the NatureMetrics eDNA Discovery Lab. This can be used by individuals or families looking to explore their local wildlife, or as community activities and for team-building exercises.