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Frequently Asked Questions

Frequently Asked Questions

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The glossary is available should you get stuck on any terminology.

Why choose NatureMetrics for your GCN eDNA surveys?

1. 100 % GCN eDNA proficiency scores for 5 years running

Natural England will only accept GCN eDNA results from laboratories that have passed an annual proficiency test, where laboratories are required to correctly identify ‘blind’ samples as positive, negative or inconclusive.

NatureMetrics are proud to have scored 100% proficiency for five years running, including in 2022, for our GCN eDNA service which we have provided since 2015.

We follow Natural England’s approved protocol (WC1067), ensuring our tests meet regulatory requirements. We offer standard (10 working days), fast (5 working days) or super-fast (2 working days) turnaround to accommodate all timeframes and budgets.

Our user-friendly GCN eDNA service includes your iDNAture Great Crested Newt eDNA Kits and convenient tracking of orders with my.naturemetrics. To adhere to the WC1067 protocol, field sampling must be carried out by a licensed great crested newt surveyor if the results will be used for planning and licence applications.

2. Convenience at every step

  • Our NatureMetrics app lets you collect and submit your field data on the go.
  • We offer next-day UK delivery for orders placed by midday.
  • We get results to you on time, even going above and beyond the regular service.
  • Whether it is last minute kits or super-fast turnaround, we will do our very best to meet your needs this GCN season.

3. World-leading eDNA facilities

We have dedicated, ultra-clean laboratories for DNA extraction and qPCR analysis to guard against contamination.

NatureMetrics are a leader in eDNA research and standardisation. In addition to GCN eDNA surveys, we analyse a range of samples (water, soil, faeces, and invertebrates) for other priority species or whole communities. Some of this work has been published in peer-reviewed journals or as technical reports. We can design bespoke sampling strategies for your species or community of interest and offer training in eDNA sampling.

What are the UK guidelines around great crested newts (GCN)?

Legal protection was afforded to the GCN at all life stages in response to ongoing declines, primarily due to habitat loss and degradation. Legislation states it is an offence to kill, injure, or take great crested newt individuals. Disturbance is prohibited, and breeding sites and hibernacula are protected. As a protected species, developers are required to survey for GCN and if found, mitigation must be proposed for GCN and their habitat in order to obtain a mitigation licence from the relevant government regulatory agency (e.g. Natural England, NatureScot, Natural Resources Wales) before proceeding with development. Mitigation may relate to methods or timings of work and installation of mitigation strategies.

GCN are listed as an European Protected Species under the Conservation of Habitats and Species Regulations (2017). There are also protected under Schedule 5 of the Wildlife and Countryside Act (1981) and as a rare and most threatened species under Section 41 of the Natural Environment and Rural Communities Act (2006) (NERC).

Do I need a GCN survey?

GCN and other protected species are likely to be impacted by development work, including but not limited to; pond maintenance or infilling, building demolition, changes to land use, wind turbines, removal of trees/hedgerows, and road building or maintenance. GCN surveys become more important where land has favourable features for GCN, such as a waterbody within 500 metres, damp patches or bogs, nearby woodland, hedgerows, trees, unmanaged grass and moss. GCN surveys will likely be needed for changes to ancient woodland, large land with good growth, land with damp mossy areas, land with unmanaged grass, hedgerows or trees, and land close to a Site of Special Scientific Interest (SSSI) or local nature reserve. A GCN survey will also be needed if there are historical records of newts on the land or close to the land proposed for development.

Can I build on land where there are Great Crested Newts (GCN)?

You will need a mitigation plan and licence from Natural England to show how you will avoid, reduce or manage any negative effects to GCN in order to continue building.

Why do I need a GCN survey?

Development can have a large impact on GCN. Without performing GCN surveys, you could be breaking the law and incur an unlimited fine or be given a prison sentence of up to 6 months for each offence.

What are the UK regulations around Great Crested Newts (GCN)?

GCN are strictly protected by British and European law at all life stages which makes it an offence to kill, injure, capture or disturb them, damage or destroy their habitat, and to possess, sell or trade them.

Are Great Crested Newts (GCN) a protected species?

GCN are fully protected by UK and European legislation. GCN are listed as a European Protected Species under Conservation of Habitats and Species Regulations (2017). UK legislation protecting GCN includes the Wildlife & Countryside Act (as Amended) 1981: Schedule 5, the EC Habitats Directive 1992: annex 11 and 1V, the Conservation (Natural Habitats etc.) Regulations 1994: Schedule 2, and the Countryside Rights of Way Act 2000 (CRoW 2000). GCN are also listed as a rare and most threatened species under Section 41 of Natural Environment and Rural Communities Act (2006).

What should I do if I have found a great crested newt (GCN)?

If a newt or evidence of newt habitat is found before development has begun, you will need to ensure minimal detrimental impact to the GCN habitat, shelter, breeding or resting site and potentially produce a mitigation scheme. If GCN are discovered after development has commenced, all works should stop until GCN surveys have been performed and appropriate measures have been taken to protect the newts on site. Speak to an ecological consultant to discuss your development plans, the surveys required and mitigation measures.

How do you carry out GCN surveys?

GCN are typically monitored using a combination of torchlight surveys, bottle trapping, egg searches and netting. Many of these activities will require a licence as they involve disturbance or capture of GCN. Four site visits using three of these methods must be undertaken, with at least two visits during mid-April to mid-May to determine presence/likely absence. If GCN are found to be a present during these surveys a further two visits using three methods must be undertaken to establish the relative abundance or population size.. eDNA analysis is a non-invasive, time-efficient and cost-effective alternative that can be used by both professionals and volunteers. This involves the capture and analysis of DNA left behind by GCN in the water column (e.g. mucus, skin cells, eggs, faeces). eDNA survey only requires a single visit to determine GCN presence/likely absence, but cannot be used to provide estimates of relative abundance.

How much does a GCN survey typically cost?

Conventional GCN surveys can range from between £300 – £1000 depending on ecologist fees, location of the development, and size or complexity of the site. The NatureMetrics GCN eDNA service (including kit and analysis) ranges from £160 to £300 (exc. VAT) per sample depending on turnaround time selected. GCN eDNA Kits alone cost £35 (exc. VAT).

How accurate are the results produced by GCN surveys?

In a comparison of eDNA analysis and conventional methods for GCN survey by professionals across 35 ponds, eDNA analysis achieved a detection rate of 99.3% (139 of 140 samples) compared to bottle trapping (76%), torchlight counts (75%) and egg searches (44%) or torchlight surveys and bottle trapping combined (95%). Across 239 ponds known to support GCN that were sampled for eDNA by volunteers, eDNA analysis achieved a detection rate of 91.3%. For more details, see Biggs et al. (2014).

How do I know if I need a Great Crested Newt (GCN) mitigation license?

A mitigation licence permits actions that are prohibited under current legislation for GCN. If survey information and specialist knowledge indicate that the proposed activity is likely to result in an offence (i.e. killing, damage to breeding sites etc.), then a mitigation licence should be obtained.

What is a GCN mitigation license, do I need one and how do I get one?

You can obtain a mitigation licence by contacting a licensed ecologist who will survey your site and surrounding ponds to inform the mitigation licence and complete an application form, method statement, work schedule and where necessary a reasoned statement on your behalf. This should be submitted to Natural England who will evaluate the documents and issue the mitigation licence. The mitigation licence lawfully permits activities to proceed which would otherwise be unlawful.

When is the GCN season?

The GCN season runs from mid-March to mid-June, but eDNA surveys can only be performed from 15th April to 30th June inclusively.

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).

Is there any update on 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. eg 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.

Are reference libraries shared between institutes, i.e. is there a large shared global database (publically) 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 (BOLD), 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).

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 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 (there is some really nice work being carried out on this by researchers in Belgium at the moment using cage experiments).

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. 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. For GCN it’s okay of course, but fish or other species that don’t frequent that part of the water?

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.

For the 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 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? Like you might 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 the 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 frozen or left in direct sunlight.

How much is there a risk that the eDNA collection can be compromised 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!

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.

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.

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.

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 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.

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. We treat each OTU as a species-level entity and 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 we are able to assign a species label.

Will there be a move to filtration sample kits for GCN eDNA sampling in the future?

We have assessed the efficacy of filtration versus precipitation methods and shown (in common with other researchers, e.g. Spens et al. 2016) that detection probabilities are higher using filtration. This is likely because the standard GCN eDNA Kits only process 90 ml of water, while our Aquatic eDNA Kits that use filters have been designed to deal with the high turbidity of ponds and typically process an order of magnitude more water. Filters pose less logistical challenges because they do not involve ethanol, are easier to process in the laboratory, and are much less susceptible to contamination. Unfortunately, a move to filtration is not up to us, but the evidence has been made available to Natural England along with contact details of independent scientists who they can consult. We hope that results derived from these kits may be accepted in coming years.

What about the risk of false positives e.g. from a Heron eating a GCN and pooping in the pond or ducks moving between ponds and carrying DNA from one to the other?

eDNA technology is very sensitive and theoretically there is potential for this type of environmental contamination. However, we would expect that this is very rare. When clients have speculated to us that they believe this may explain a surprising positive result, subsequent surveys have usually confirmed GCN presence.

How many kits do I need?

For ponds, a single kit is usually enough as long as subsamples have been merged from around the perimeter of the pond.

As the size of the waterbody increases, so should your sampling effort.

The precise number of samples will usually be a trade-off between your budget and the amount of spatial resolution you need or the importance of detecting rare species.

For most lakes, 5-10 samples are sufficient, given appropriate merging of subsamples but you may want to take more if you are targeting rare species. In rivers and streams, it depends on the size and flow rate of the waterbody and the area that you need to survey.

We are always happy to advise on sampling design for specific projects.

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?

NatureMetrics has detected European otter (Lutra lutra), neotropical otter (Lontra longicaudis), and giant river otter (Pteronura brasiliensis) from water samples using our vertebrate or mammal metabarcoding assays. We have also identified otters from their spraints, and can analyse spraints to understand otter diet (including vertebrate and invertebrate prey) using metabarcoding.

However, detection rates for these species are low relative to how much time they spend in water. Otters deposit spraint outside of water on riverbanks, logs and rocks, and use latrines associated with caves and dens. Spraint may get washed into water if heavy rainfall occurs and/or water levels rise, but the likelihood of this is unknown. Furthermore, otter fur is very dense so they may not shed much hair into the water either. Although otters hunt in water, they tend to take their prey back to shore to eat, so DNA from saliva/prey remains is also unlikely to be deposited in water. Finally, these mammals have large territories and tend to be solitary apart from mating and rearing pups.

High sampling effort across a number of waterbodies and in different seasons is needed to maximise chances of otter detection.

Can you analyze 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 analyze coral eDNA?

We have not yet worked on coral eDNA and it is a complex group. However we do think this is an exciting application with great potential and would love to develop it as part of a collaborative research project.

Can you analyze 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.

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)!

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.

Do you think that traditional taxonomic skills and natural 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.

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 also 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 type 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?

Please visit our careers page to see our latest job vacancies.

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.

How do I order GCN kits?

NatureMetrics has an online ordering portal that enables you to easily place your orders, track their progress and view the results of all your projects.

To set up an account, simply email gcn@naturemetrics.co.uk and we will get your account set up.

If you are an existing customer and have forgotten your login details, just go to the portal and click on ‘forgot password’.

What is the latest time I can place an order for next day delivery?

Orders placed by midday will be eligible for next-day UK delivery.

How do I pay for the kits?

Once we have dispatched your kits, we will send you an invoice with 30-day payment terms. If you have a PO, please add this to the appropriate field when placing your order.

When can I expect to receive my results?

We offer three different turnaround times for GCN testing: standard (10 working days), fast (5 working days), and super-fast (2 working days). We begin to process the samples as soon as they are received by our lab team but, as we cannot control the delivery time, analysis begins the day after samples are delivered to the lab.

How many kits do I need for a GCN Survey?

Natural England states that you should use “…one kit per pond up to an area of 1 hectare. Beyond this, use an additional kit per hectare”. For example, two kits should be used for a pond between 1-2 hectares in size, and each kit should be used to sample one half of the pond in accordance with the Natural England protocol (i.e. one kit used to collect 20 subsamples from one half of the pond, and the other kit used to collect 20 subsamples from the other half of the pond).

When can GCN eDNA samples be taken?

Natural England states that samples should be taken between 15th April and 30th June.

How do I return my GCN eDNA kits once I have collected the samples?

Collection of Great Crested Newt eDNA Kits is included in the service price and a member of our team will book the collection on the date you chose when you placed your order. Once booked, it is vital that you follow the shipping instructions given with your order to ensure you are complying with shipping regulations. Please bear in mind that we do not arrange collections for Fridays in order to avoid samples being held in a depot over the weekend in an uncontrolled temperature environment.

How should the GCN eDNA sampling kits be stored?

Following Natural England approved protocols, NatureMetrics Great Crested Newt eDNA Kits have a shelf life of 3 months, meaning you can store your kits for most of the GCN season before use. Kits can be stored at ambient temperature before use.

How do I change my delivery or collection date?

Simply email gcn@naturemetrics.co.uk with the kit ID, project name and new date.

How long can a GCN sampling kit be kept for?

Our Great Crested Newt eDNA Kits have a shelf life of 3 months. Kits can be stored at ambient temperature after use if couriered on the day of collection. If samples are stored overnight (or longer) before shipping, they should be refrigerated at 2-4°C. Samples can be stored for up to 1 month in the refrigerator before analysis but should not be frozen. We advise that samples are returned to our lab as soon as possible after collection.

We have unused kits from a previous order/project, how do we use them for a new separate project?

You can move an unused kit to another project. Simply email gcn@naturemetrics.co.uk with the kit ID and which projects you are moving from and to.

What will my results look like?

Following analysis, NatureMetrics will send you a report through the order portal that summarises the results and provides you with the relevant information on methods used and quality control.

We have some ‘inconclusive’ results and would like to know why?

Inconclusive results may arise because DNA is degraded or samples are inhibited. DNA degradation is a process by which DNA has been damaged to the point that it cannot be amplified and subsequently detected. The use of preservatives in our kits minimises the risk of DNA damage, but inappropriate storage conditions may still lead to DNA degradation. PCR inhibition occurs when compounds present in the sample (e.g. acids) interfere with the qPCR reaction at a molecular level, preventing or reducing target DNA amplification and subsequent detection. Inhibitors may interact directly with target DNA, or interfere with the enzyme that drives the PCR reaction.

A synthetic control is included in kits to test for degradation, and a synthetic control added to samples after DNA extraction to test for inhibition. A qPCR assay specific to each control is used to test the original DNA extract. If the signal from either synthetic control is lower than expected or absent, then degradation and/or inhibition are present. If degradation is identified, there is no way to resolve this and the sample will be reported as inconclusive. If inhibition is identified, the sample is diluted twice prior to testing for GCN presence in accordance with the Natural England protocol. If dilution fails to resolve inhibition, then the sample will be reported as inconclusive.

Ponds with an inconclusive result should be resampled, adhering strictly to the Natural England protocol, and new samples returned to the lab for analysis as soon as possible.

How long does GCN eDNA take to degrade?

eDNA is typically detectable for several days from when the organism leaves behind the biological material. eDNA has been reported to last for up to 300 hours in aquatic systems, but there is no definitive answer for this yet. The degradation of DNA is slowest when it is cold, dark, or when the DNA is bound to sediment.

Is bleach an effective method for cleaning of equipment?

Ideally, sampling equipment should be ordered sterile and be individually wrapped and single-use to minimise the risk of introducing contaminant DNA from sampling equipment to environmental samples. Sampling kits purchased from NatureMetrics only contain sterile, single-use components. We are continually striving to reduce plastics in our kits and actively promote recycling. If any equipment must be decontaminated and reused (note that no equipment for GCN eDNA survey should be reused between sites apart from surveyor PPE), then bleach sterilisation is the accepted method.

Bleach must be purchased at a stock concentration of 3-6% sodium hypochlorite (higher is better) from a supermarket, cleaning service or laboratory supplier. The stock bleach should be diluted 1:10 with purified or deionised water (e.g. 1 L of stock bleach to 9 L of purified water). Where purified water is unavailable, mineral water purchased from a supermarket should be used. This dilution creates a 10% bleach solution. You should submerge non-metal and non-electrical equipment in the 10% bleach solution for at least 10 minutes but ideally 3 hours or overnight. After this time, remove equipment from bleach and thoroughly rinse (at least 3 times) with purified or deionised water. After rinsing, place equipment on fresh paper towel and leave to dry overnight. Metal equipment should be submerged for 5 minutes before rinsing and drying overnight. Surfaces of electrical equipment should only be wiped with 10% bleach, followed by 70% ethanol.

Is there any chance of cross contamination at the lab?

NatureMetrics has strict policies and procedures in place to minimise any risk of contamination. Our staff undergo rigorous training prior to GCN season and we routinely run quality checks with every test to ensure the risk of contamination is minimised. Extraction is performed in a dedicated laboratory for GCN samples and separate to where qPCR is carried out.

Is it possible to detect other species from the sample?

While GCN samples could be analysed for other species, this is not recommended due to the small volume of water processed (90 ml) which limits the amount of eDNA that can be captured and will likely result in lower detection rates. Instead, we recommend collecting a larger sample (up to 3 L) and filtering the water using our Aquatic eDNA Kits. Email eDNA-lab@naturemetrics.co.uk for more information on these kits and available analyses.

What does the ‘score’ in my results mean?

This is the number of qPCR replicates that successfully amplified GCN eDNA. A total of 12 replicates are performed on each sample during qPCR. A low score can be interpreted as a sample containing a low amount of GCN eDNA, whereas a high score can be interpreted as a sample containing a high amount of GCN eDNA. Samples with a score of 1 are still classed as positive for GCN in accordance with the Natural England protocol, but we would recommend that these ponds are resurveyed for GCN using conventional survey or eDNA survey to confirm species presence. Note that a negative result does not preclude the presence of GCN eDNA at a concentration below that which can be detected with qPCR. If GCN individuals are present in low abundance, they may not be shedding enough DNA into the water column to enable it to be captured and amplified.

What is eDNA?

Environmental DNA (eDNA) is DNA that has been left behind by organisms in their environment in the form of skin cells, hair, mucus, faeces, urine, blood, saliva, gametes, and deceased remains. eDNA accumulates in water, sediment, soil and air, but persists for different times in these environments. eDNA can persist for hours to weeks in water compared to months to years in sediment. eDNA can be captured from environmental samples and used to survey single species or whole communities.

What is in a GCN eDNA Kit?
  • 2 x pairs of nitrile gloves
  • 1 x 1.6 L sampling bag
  • 1 x ladle dipper (30 mL)
  • 1 x plastic pipette
  • 6 x 50 mL tubes containing 35 mL of ethanol and preservative inside cardboard box with insert
  • 1 x resealable bag
  • 1 x sampling datasheet
How many kits do I need?

Natural England states that you should use “…one kit per pond up to an area of 1 hectare. Beyond this, use an additional kit per hectare”. For example, two kits should be used for a pond between 1-2 hectares in size, and each kit should be used to sample one half of the pond in accordance with the Natural England protocol (i.e. one kit used to collect 20 subsamples from one half of the pond, and the other kit used to collect 20 subsamples from the other half of the pond).

Can I collect water samples outside the GCN season?

Although water samples can be collected outside the GCN season and analysed for GCN, the results cannot be used for planning or licence applications.

How do I collect my samples?

Detailed instructions for sample collection are available to download from our website, which follow the steps stated in the WC1067 Technical Advice Note. Detailed instructions are also available on the NatureMetrics app, and a simplified pictorial protocol is provided with our GCN eDNA Kit.

Can I get a refund for unused kits?

Unused kits and their analysis are non-refundable. Unused kits can be returned to NatureMetrics to safely dispose of chemicals.

Where can I find my results?

NatureMetrics has an online ordering portal that enables you to easily place your orders, track their progress and view the results of all your projects.

Why were GCN detected with conventional methods but not eDNA analysis?

If GCN are in low abundance, they may not be shedding enough DNA into the water column to enable it to be captured and amplified. Alternatively, if a pond has greater habitat complexity, then higher sampling effort may be required than what is stated in the Natural England protocol. Dilution is the prescribed strategy to resolve inhibition when identified in samples to dilute inhibitors present in the sample. However, any target DNA present will also be diluted, possibly to concentrations too low to be detected.

Where can I find more information about eDNA analysis for GCN?

Several scientific studies using eDNA analysis for GCN have been published, including:

Rees, H.C., Bishop, K., Middleditch, D.J., Patmore, J.R.M., Maddison, B.C. & Gough, K.C. (2014) The application of eDNA for monitoring of the Great Crested Newt in the UK. Ecology and Evolution, 4, 4023–4032. https://doi.org/10.1002/ece3.1272

Biggs, J., Ewald, N., Valentini, A., Gaboriaud, C. & Griffiths, R.A. (2014) Analytical and methodological development for improved surveillance of the Great Crested Newt. Defra Project WC1067.

Biggs, J., Ewald, N., Valentini, A., Gaboriaud, C., Dejean, T., Griffiths, R.A., Foster, J., Wilkinson, J.W., Arnell, A., Brotherton, P., Williams, P. & Dunn, F. (2015) Using eDNA to develop a national citizen science-based monitoring programme for the great crested newt (Triturus cristatus). Biological Conservation, 183, 19–28. https://doi.org/10.1016/j.biocon.2014.11.029

Rees, H.C., Baker, C.A., Gardner, D.S., Maddison, B.C. & Gough, K.C. (2017) The detection of great crested newts year round via environmental DNA analysis. BMC Research Notes, 10, 327. https://doi.org/10.1186/s13104-017-2657-y

Buxton, A.S., Groombridge, J.J. & Griffiths, R.A. (2017a) Is the detection of aquatic environmental DNA influenced by substrate type? PLoS ONE, 12, e0183371. https://doi.org/10.1371/journal.pone.0183371

Buxton, A.S., Groombridge, J.J., Zakaria, N.B. & Griffiths, R.A. (2017b) Seasonal variation in environmental DNA in relation to population size and environmental factors. Scientific Reports, 7, 46294. https://doi.org/10.1038/srep46294

Buxton, A., Groombridge, J. & Griffiths, R. (2018a) Comparison of Two Citizen Scientist Methods for Collecting Pond Water Samples for Environmental DNA Studies. Citizen Science: Theory and Practice, 3. http://doi.org/10.5334/cstp.151

Buxton, A.S., Groombridge, J.J. & Griffiths, R.A. (2018b) Seasonal variation in environmental DNA detection in sediment and water samples. PLoS ONE, 13, e0191737. https://doi.org/10.1371/journal.pone.0191737

Griffin, J.E., Matechou, E., Buxton, A.S., Bormpoudakis, D. & Griffiths, R.A. (2019) Modelling environmental DNA data; Bayesian variable selection accounting for false positive and false negative errors. Journal of the Royal Statistical Society: Applied Statistics Series C, 93, 372. https://doi.org/10.1111/rssc.12390

Buxton, A.S. (2021) How reliable is the habitat suitability index as a predictor of great crested newt presence or absence? The Herpetological Journal, 111–117. https://doi.org/10.33256/31.2.111117

Buxton, A., Matechou, E., Griffin, J., Diana, A. & Griffiths, R.A. (2021) Optimising sampling and analysis protocols in environmental DNA studies. Scientific Reports, 11, 11637. https://doi.org/10.1038/s41598-021-91166-7

Buxton, A., Diana, A., Matechou, E., Griffin, J. & Griffiths, R.A. (2022) Reliability of environmental DNA surveys to detect pond occupancy by newts at a national scale. Scientific Reports, 12, 1295. https://doi.org/10.1038/s41598-022-05442-1

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