The TRIPLELOCK™ Test Strips deliver COVID-19 results within 60 minutes The First and Only Approved Canadian-Made Solution to Support Canada’s Response to the Pandemic
GUELPH, Ontario, November 4, 2020 – Precision Biomonitoring announced today it has received Health Canada approval for its ready-to-use thermostable SARS-CoV-2 TRIPLELOCKTM Test Strips for use across Canada. The TRIPLELOCK™ Test Strips are a rapid, point-of-need diagnostic RT-PCR test able to provide accurate results for 9 samples in just 60 minutes. This cost-effective, Canadian-manufactured test will help support the government and industry as Canada focuses on safely re-opening the economy amidst a second wave of the virus. Precision Biomonitoring announced in June that it received approval and funding from Next Generation Manufacturing Canada (NGen) to support manufacturing of its ready-to-use thermostable test.
“We are proud to be adding another Health Canada approval to the list of testing devices offered by Precision Biomonitoring. We have been working closely with various industries across Canada to help bring their workforce back safely, and are thrilled to be expanding that capability,” says Dr. Mario Thomas, CEO, Precision Biomonitoring. “This approval also means we can strengthen our ongoing support for federal and provincial governments, as well as Canadian industries.”
The SARS-CoV-2 TRIPLELOCK™ Test Strips are designed for accurate RT-PCR point-of-need diagnostics, are stable at room temperature, and to be used by qualified laboratory personnel only. The lyophilized products combine the highest accuracy performance of RT-PCR with convenience of use and stability, which are crucial for remote parts of the country where adequate access to precise testing may be limited.
“We are excited by today’s news as this approval is a huge step forward for Canadians as we continue our battle with COVID-19,” says Eric Hoskins, former Minister of Health and Long-Term Care of Ontario and Precision Biomonitoring Board Member. “The healthcare community has come together in incredible ways to help Canadians manage their way through this global pandemic. The thermostable TRIPLELOCK™ test that is now available will be crucial in workplaces across the country, schools, and rural and remote areas, where answers are needed fast.”
Precision Biomonitoring also recently received CE Mark approval in Europe for its TRIPLELOCKTM SARS-CoV-2 test in 96-Well Plate format. Made available for immediate use in labs across Europe, the CE mark will help to address the rising demand for testing in various countries, including hotspot regions.
About Precision Biomonitoring Rapid SARS-CoV-2 TRIPLELOCKTM Test Strips
Precision Biomonitoring’s easy-to-use Test Strips are a mobile solution for the healthcare community in Ontario and across Canada. The portable TRIPLELOCK™ Test Strips can be transported without refrigeration and when used by qualified lab personnel, are ideal when results are needed immediately in workplaces and more rural and remote regions. The SARS-CoV-2 TRIPLELOCKTM Test Strips detect the RNA of the severe acute respiratory syndrome, COVID-19. The COVID-19 RNA targets are multiplexed together with an RNA positive control. Early identification and diagnosis of COVID-19 is crucial to ensure a rapid response, thus mitigating the possible additional negative consequences of the virus.
About Precision Biomonitoring
Founded in 2016 by a team of scientists from the University of Guelph’s Biodiversity Institute of Ontario, Precision Biomonitoring provides TRIPLELOCK™ onsite eDNA surveillance platform solutions that give customers earlier detection of organisms for a more rapid response. Customers are any organizations that need onsite surveillance and rapid identification of any organism in any environment. The Precision Biomonitoring team is at the forefront of technological innovations in the genomics industry. Our vision is a world where we can identify any organism on the spot, in an instant, anywhere on the planet.
The fight against COVID-19 is taking a new turn. There are now more testing solutions today than just a few months ago. Due to the high demand for many services, goods and commodities, a large number of companies and businesses must keep their worksite fully operational while ensuring the health and safety of their employees, contractors, and clients.
Many companies are now opting to use the newly available and approved RT-PCR testing solutions at their facilities, which shows their commitment and seriousness about the health and safety of their employees, contractors and clients. These solutions offer on-site actionable results much faster than a traditional lab-based test while helping Public Health with positive contact tracing more efficiently.
As a decision-maker in a company, you understand that testing is the best way to show your colleagues and business partners that you are serious about their safety and keeping a virus-free work environment. Now that you decided to implement testing into the business, what are the next steps in navigating this implementation?
If you are looking at actualizing a testing program or protocol in your business, you most likely already have a screening and cleaning strategy in place. But you want to be more proactive about potential positive case detection than relying only on cleaning and screening methods.
You probably did some research on RT-PCR units and now you are convinced that it would be the best option for testing. You know it effectively and efficiently tests the people that need to come on your premises, get results quickly and be able to react accordingly.
The most crucial part of the implementation process needs to happen before the mobile device is at your site. If not done previously, you will now have to establish your relations with your regional Public Health office and develop your testing strategy.
It is critical to inform Public Health about your testing plans because they control COVID testing across all the Canadian provinces. Before buying an RT-PCR machine, make sure that they approve your use the device on your premises since it is outside a recognized medical lab facility.
You now realize that you must have a proper site to perform testing. The RT-PCR mobile device is small enough to be operated in a very small environment, but it must be clean. The setting should have lots of lighting and be free of clutter. Counters should also be non-porous for easy disinfection. Proper cleaning protocols must be implemented wherever the mobile device is to be used. We will post a blog specifically dedicated to cleaning and disinfecting.
Our testing device is licenced by Health Canada as a Class IV medical device. As such, it must be operated by a licenced lab technician who is trained in handling sample processing to avoid contamination, vial labelling and organization of collected samples, and have experience performing the types of sensitive manipulations involved in executing this test.
Your lab technician will be quite busy preparing the samples, interpreting the results, recording data and maintaining the lab. Also, someone has to give the results back to the donors, which implies time on the phone or computer. It is helpful to enlist a nurse to assist with these duties, as well as to collect the swab samples.
At this point, you have figured out the logistics, and you are happy about the way it looks on paper. You now must think about the most crucial part of your screening program: the testing strategy.
The testing strategy will determine the workload on your lab team, and most importantly, impact your productivity. These are some questions that you will need to answer to have a clear idea of your testing protocol:
Who will you test?
Which criteria will you use to determine who will be tested?
What is the frequency of testing?
Is this frequency the same for all people being tested?
Do people have to wait to have their results before proceeding to their workspace?
If they do have to wait, where will they wait?
Will you include contractors in our program?
What do we do if we have a positive case?
Where will the testing facility be located?
If you have a multi-location site, will a fixed testing unit be suitable, or will you need a mobile unit?
How will the testing strategy fit into my overall pandemic fighting strategy?
At which level do we get Public Health involved in the process?
Who will communicate with Public Health? What are their communication expectations?
Your testing strategy will also determine your budget allocation. The mobile devices have a limited sample processing capacity. Your testing strategy will dictate the number of devices and personnel you need to meet your testing targets.
You will also have to prepare to source disposable lab consumables that you will need, based on the number of tests you will perform. These include throat or nasal swabs, viral transfer media, PPE (disposable gloves, masks, etc.), and cleaning agents.
Once you have answered most of these questions, you will then have to figure out a time frame to set-up and implement the testing program. With this time frame you will also have to develop a cadence for the frequency of testing andre-testing individuals. This is important because over the first 4 days of infection before typical symptom on-set, the probability of false-negative results in an infected person are very high. On day 1 of the infection the probability of a false-negative is 100%, whereas on day 4 the probability decrease to 67%. Therefore, it is important to create a morefrequent testing strategy for workers who think they may have come into contact with someone who is ill, or for shift workers who spend extended periods of time on-and off-sites, like a mining or oil-drilling site. (Source: Annals of Internal Medicine available at: https://www.acpjournals.org/doi/10.7326/M20-1495)
Setting up and implementing these protocols is not, for most businesses, a simple task. It will require a dedicated person, preferably with some knowledge of this type of process, to lead the implementation and guide the senior management in their corporate decisions. At this stage, you have different options. You can DIY or outsource your program to a third party that will manage the entire testing process for you.
Finally, you will have to communicate the enhanced screening process you are putting in place with your employees and stakeholders. It is important to communicate this as soon as possible to ensure they understand that you are implementing a process to create a virus-free workplace.
All the stakeholders and the senior leadership team must understand that the testing process involving an RT-PCR mobile unit is just one more tool in your screening process to ensure a virus-free environment. Having an RT-PCR device at your site does not mean that you can let your guard down on the other important pieces of the screening and cleaning/disinfecting strategy that you have in place.
A Virus-Free Environment
As mentioned earlier, this is not an easy process to put in place. However, it is very rewarding to see all the pieces of your screening and cleaning/disinfecting strategy working together seamlessly to prevent positive cases at your workplace.
How do we know it is so rewarding? Because we have helped many companies achieve their goal of implementing a world-class screening process, and we see how rewarding it has been for them to give their employees a safe environment to work in and peace of mind.
Environmental DNA or eDNA can improve the biomonitoring process in three central areas when compared to conventional biomonitoring methods involving cost, accuracy, and time. The infographic below highlights these three benefits.
Environmental DNA (eDNA) methods offer several advantages
over conventional species survey methods, especially when carried out with the
right amount of expertise at each step of the workflow. Such advantages include
time and cost savings, especially when on-site detection is used. eDNA offers
higher sensitivity and specificity reducing observer bias compared to
conventional methods. eDNA also reduces disturbances to the species and its
Although eDNA science is relatively young, it is maturing rapidly, with many aspects of survey design, sampling methods, and laboratory analysis being accepted as best practices – and one day industry standards [1,2]. However, eDNA surveys that fail to comply with such a set of best practices can quickly fall into a trap of poor detection probability and erroneous results. While eDNA surveys can appear relatively simple to conduct, one cannot simply grab a water sample in a Nalgene bottle and go to a lab, hoping to achieve optimal detection probability or have high confidence in their results. Quality eDNA surveys are conducted with careful consideration of all aspects of survey design and sample collection and processing.
Here at Precision Biomonitoring Inc. we developed the TripleLock™ Platform which at its core is based on the widely accepted best practices for eDNA surveys, placing us at the forefront of the eDNA survey industry. Conducting robust eDNA work is especially important at this early stage if eDNA methods are to be accepted so that the advantages of eDNA can be employed for environmental assessments and conservation biology across the world. The focus of this blog will be to outline the advantages of the TripleLock™ Platform and how they compare to other methods in the industry.
Platform Core I: TripleLock™ qPCR Assays
The first core of the TripleLock™ platform consists of
high-quality, rigorously validated qPCR assays for the detection of a target
species. At Precision Biomonitoring Inc. we have conceived and developed
a proprietary method for stringent assay development, which allows us to consistently
design and validate species specific and highly sensitive qPCR assays. We
developed our method meeting and exceeding The Minimum Information for
Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines. The
MIQE guidelines outline several standards that should be adhered to for reliable
and interpretable qPCR analyses, and by building our platform based on such
rigorous standards we ensure replicable results while minimizing errors. Our
assays are also verified with highest level of international standards by a 3rd
party ISO 17025 accredited laboratory. The advantages of following standardized
methods are clear as they produce consistency in results. Assays need to
conform to such rigorous standards and cannot be simply collected from different
sources of literature and 3rd party reporting, since that would result
in inconsistent standardization across the catalogue.
We take pride in developing our own assays in-house, as
opposed to obtaining published assays in literature, as it allows us to
optimize assays using our trusted methods and equipment. While the quality of
assays in the eDNA literature may vary, one must remember that an assay
consists of more than just a primer and probe set, it encompasses the exact
type of qPCR reagents and instrumentation with which it was validated. This
means that replicating the exact sensitivity of published assays on different
equipment can entail extra costs and time, while also moving away from the MIQE
standards if the published assay parameters can’t be replicated in vitro.
Additionally, our assays are designed with an internal
positive control (IPC) to detect PCR inhibition, an absolute necessity to avoid
false negative results. Many experts in eDNA science concur that checking for
PCR inhibition is part of the foundation of good eDNA work .
Platform Core II:
Optimal Survey Design
The second core of the TripleLock™ Platform is providing optimal survey designs to maximize the probability of detection for a target species. It is widely understood that the distribution of eDNA is not homogenous and depends on several variables including species ecology, water quality, pH, and turbidity, to name a few. It is for this reason that understanding the ecology of eDNA, and how best to sample for it, is so crucial. Our platform brings together optimal sampling designs, together with sophisticated sampling methods to conduct eDNA surveys with the utmost confidence.
We take into account the critical environmental variables,
species biology and ecology, and site specific considerations that directly relate
to the purpose of the study. At Precision Biomonitoring we use a proprietary method
based on statistical analysis, including habitat occupancy modelling, to
determine sampling optimality for a given eDNA survey. We are currently
developing a software tool (patent pending) that will allow for customized survey
designs based on the most up to date eDNA data available. This survey design
tool adds another level of consistency and standardization to our workflow.
The foundation of a good sampling scheme is how the samples are collected to begin with. Our field sampling methods are far more advanced compared to other offerings. Precision Biomonitoring makes use of the ANDe™ sampling backpack, the first purpose-built aquatic eDNA sampling system . The ANDe™ affords increased sampling versatility, allowing us to sample more effectively at depth, across transects, with increased throughput, and higher water volumes. For example, we can sample for species that typically reside on the bottom of a lake, at 30 feet deep, very easily using the ANDe™ system, which greatly increases the effectiveness of such a survey. This is something that cannot be done using small plastic bottles to sample surface water, a low fidelity practice offered by others in the industry. Our survey design methods along with our advanced sampling techniques let us conduct superior eDNA surveys compared to others in the eDNA industry.
Platform Core III:
The third core of the TripleLock™ platform is on-site detection which enables us to obtain rapid results in the field. On-site detection brings together several components of our field sampling methods including the ANDe™ filtration system, as well as on-site DNA extraction and qPCR analysis. The advantages of on-site detection come when obtaining results is time sensitive and a decision is pending. By making use of the Biomeme eDNA sample prep kit and Biomeme Three9™, a portable qPCR device, we can obtain results in the field and communicate them rapidly, via built in data connectivity, to a decision maker. This means no waiting on samples to be transported and analysed back at a central lab. Another advantage of these methods is that by filtering, extracting, and eluting DNA into a pH and thermostable buffer, in the field we minimize the potential for sample degradation during transit, thus decreasing the possibility of false negatives.
Removing the potential for sample degradation is an inherent
advantage of our field methods compared to methodologies offered by others. By
quickly extracting DNA from a filter into a stable buffer we preserve those
valuable fragments of eDNA for analysis.
There are several eDNA analysis companies that offer
sampling “kits” consisting of plastic bottles which are used to grab a small
surface sample which then must be shipped all the way back to a lab before any analysis
can start. Using handheld plastic bottles places the technician right where
they need to sample, causing disturbances in the water as well as increasing
the potential for contaminant transmission by said technician between sampling
points, not to mention contamination stemming from the wet exterior of a bottle.
By using the ANDe™ system we enable our technicians to be up to 3.6 meters
from the point where the sample is being taken, and contain our sample in an
enclosed and sterile, single-use filter cartridge, greatly reducing the
potential for contamination.
Besides the serious disadvantages that stem from being
limited to surface samples due to using bottles, other disadvantages and
concerns arise from transporting water samples back to a lab for analysis.
Transporting water samples introduces significant opportunity for eDNA to
degrade. Transporting water samples on ice can potentially decrease degradation
but does not eliminate it. Additionally, delays in shipment, especially during
warm weather, make keeping water samples cold logistically difficult and at
times impossible. By extracting and analysing DNA in the field we remove the
potential for DNA to decay, and eliminate these significant logistical
Additionally by minimizing DNA degradation we eliminate the
need for superfluous, ambiguous, and costly additional qPCR tests for so called
total viable eDNA. These tests are often based on plant/algal DNA. Plant/algal
cells are surrounded by resilient cell walls that prevent osmotic lysis, but
animal cells are not. In theory this means that more plant/algal DNA may be
preserved inside cells caught on filters prior to eDNA extraction, while animal
cells easily lose integrity and release DNA, making it less long-lived compared
to plant based eDNA. Inferences about the integrity of a target animal
species DNA that are based upon the results of a plant based test are likely
suspect. Furthermore, tests for total eDNA of a family of organisms, all fish for
example, are also superfluous. Target DNA can be in extremely low abundance
compared to total eDNA, shown to be > 0.0004% of total eDNA in some cases .
In theory, tests for total fish or amphibian eDNA most likely don’t give
pertinent information about the integrity of a single target species eDNA, especially
in cases where samples are taken from an isolated location or depth where other
organisms in a family may not inhabit. A theoretical example of such a case is
when an anthropogenically constructed habitat, i.e. a remediated river, is home
to only a select few pioneer or reintroduced species. In these cases, a test
for total fish eDNA may be interpreted as a false negative due to low
abundance, not low integrity.
Precision Biomonitoring’s TripleLock™ Platform brings together
multiple aspects of molecular biology, environmental survey design and
sampling, and leading innovations to produce gold standard eDNA surveys. The
development of our platform is anchored in the best practices for eDNA surveys
as well as standards such as the MIQE guidelines, making Precision
Biomonitoring an industry leader for eDNA surveys.
1. Wilcox TM, Carim KJ, Young MK,
McKelvey KS, Franklin TW, Schwartz MK. Comment: The Importance of Sound
Methodology in Environmental DNA Sampling. North Am J Fish Manag. 2018; 1–5.
2. Goldberg CS, Turner CR, Deiner K,
Klymus KE, Thomsen PF, Murphy MA, et al. Critical considerations for the
application of environmental DNA methods to detect aquatic species. Methods
Ecol Evol. 2016;7: 1299–1307. doi:10.1111/2041-210X.12595
3. Bustin SA, Benes V, Garson JA,
Hellemans J, Huggett J, Kubista M, et al. The MIQE Guidelines: Minimum
Information for Publication of Quantitative Real-Time PCR Experiments. Clin
Chem. 2009;55: 611–622. doi:10.1373/clinchem.2008.112797
4. Thomas AC, Goldberg CS, Howard J,
Nguyen PL, Seimon TA, Thomas AC. ANDe TM : A fully integrated
environmental DNA sampling system. 2018;2018: 1379–1385.
5. Turner CR, Barnes MA, Xu CCY, Jones
SE, Jerde CL, Lodge DM. Particle size distribution and optimal capture of
aqueous macrobial eDNA. Gilbert M, editor. Methods Ecol Evol. 2014;5: 676–684.
Exploring MIQE-Like Standards for qPCR-Based Molecular Detection of eDNA
In the previous
blog, we briefly touched on the MIQE standards – guidelines published to promote
minimal standards of reporting qPCR data from clinical trials, used to detect small relative changes in gene expression
within cells or tissues, or to quantify the amount of pathogen(s). As much of
contemporary environmental DNA work is conducted using qPCR approaches, it is
only sensible that similar standards be developed for eDNA work, that should –
it will be argued – extend beyond what is acceptable for clinical applications.
This is somewhat
of a reverse of convention in scientific circles, as it is normally the case
that clinical applications need an elevated burden of evidence to support any
conclusions drawn from experimental data. However, because of the ecology of
eDNA, its distribution in natural systems with myriad potential sources of
generation and decay, allied to the hierarchical nature of its sampling, I believe
eDNA presents a special case whereby more rigorous standards should be applied than
clinical settings so that confidence in our results are trustworthy.
What are the
chief differences in using qPCR to detect changes in gene expression or viral
load, for instance, with using qPCR to detect eDNA? Figure outlines the
workflow for A) performing a gene expression qPCR analysis alongside B) a
generic eDNA workflow. In A) an investigator has a sample of tissue, within
which there is guaranteed nucleic acid content. Imagine that they want to test
for the expression levels of a gene that is hypothesized to play a positive
role in alleviating stress in plants subject to an environmental stressor. In
such studies, expression levels are compared against so-called reference genes,
which are always expressed, so that any changes in the target gene can be
compared after normalization of expression levels. Although plants in a control
plot should show little expression of the target gene, it would still be
expected to be found, albeit at reduced levels, in the plant tissue subject to
nucleic acid extractions (in this case, messenger RNA, which is converted to
DNA (complementary DNA or cDNA)) in a process called reverse transcription, so
that the target is amenable to DNA-based qPCR. As such, one expects a lot of
cDNA, if it were to be visualized using standard laboratory gel techniques. There
will always be much more – and stable – expression of the reference gene’s
mRNA, and by extension cDNA. In short: there is a reliable source of cDNA,
which forms the template for the qPCR assays for both the target gene and the
environmental DNA. Leaving aside the complex ecology of eDNA to one side
(subject to another blog post but see excellent review by Barnes and Turner ),
the distribution of eDNA in a water body is largely ephemeral and at much lower
concentrations, with no guarantee that sampling will entrain DNA molecules or
cellular debris into sampling tubes or onto filter papers. There is a
significant source of observational error at this stage, which is largely
absent in gene expression studies, although both share procedural errors that
can impact downstream qPCR success. Depending on factors including the volume
of water sampled and pore size of the filters, the amount of total eDNA
collected may vary substantially, although if visualized on a gel, is less
likely to contain as much target as tissue-extracted mRNA turned cDNA.
for either A) or B) requires taking an amount of total cDNA or total eDNA for
use as template for the reactions. The probability of subsampling this extract
and not getting detection is higher for eDNA, due to the extra rarity of the
target in both the water sample – which may not have been collected at all –
and by the relative rarity of the target molecule in the soup of total eDNA
molecules. For eDNA we have two
uncertainties in sampling due to the two sampling events – of the water body
and of the total eDNA – that act to increase underlying statistical error.
eDNA needs to adopt more sensitive and, arguably, more specific assays than
clinical applications. More specific? Well, whilst many genes evolve during
evolution by way of duplication and subsequent divergence, these events are
localized to gene families so whilst when developing a qPCR assay for a gene with
a number of evolutionary similar orthologs and paralogs, the nucleotide
divergence between these genes and all others in the genome of a single tissue
type is huge, a dn thus non-specific amplification of a non-target gene
attenuated. However, when using an eDNA marker, that same stretch of DNA (e.g.,
COI locus) is present in all non-targets, as it has been inherited by a common
ancestor. However, evolution will increase the number of nucleotide differences
between species as generations pass. However, more recently diverged taxa may
not display enough between species variation to design an appropriate assay –
but see the previous blog for an in-depth treatment of assay design and target
In the next
blog, I shall discuss exactly how we estimate LOD, LOQ – further adopting MIQE
standards – and how we optimize an assay to befit a rigorous, repeatable and
reproducible eDNA assay. To do so, not only do we need to optimize the target,
but also identify and countenance factors that input variance into the system,
most nefariously PCR inhibitors. I shall describe how we use MIQE-like guidance
and synthetic internal positive control elements to determine the reliability
of eDNA results. We shall also discuss in the future, how the ecology of eDNA
and assay performance in pilot trials can be used to optimize detection (and
the potential quantification) of targeted eDNA detection studies.
 Bustin et al. (2009). The MIQE guidelines: minimum information for
publication of qPCR experiments. Clinical Chemistry55, doi.org/41373/clinch.2008.112797.
 Barnes and Turner (2016). The
ecology of environmental DNA and implications for conservation genetics. Conservation Genetics, 17, 1-17.
 Doi et al. (2017). Environmental
DNA analysis for estimating the abundance and biomass of stream fish. Freshwater Biology, 62, doi.org/10.1111/fwb.12846.
 Nevers et al. (2018). Environmental DNA
(eDNA): a tool for quantifying the abundant but elusive round goby (Neogobius
melanostomus). PLoS One, 13, doi.org/10.1371/journal.pone.0191720.
 Evans et al. (2016).
Quantification of mesocosm fish and amphibian species diversity via eDNA
metabarcoding. Molecular Ecology
 Hunter et al. (2016). Detection limits
of quantitative and digital PCR assays and their influence in presence-absence
surveys of eDNA. Molecular Ecology
 Forootan et al. (2017). Methods
to determine limit of detection and limit of quantification in quantitative
real-time PCR (qPCR). Biomolecular Detection
Quantification, 12, 1-6.
***cDNA vs eDNA
(diagram of how each is made and detected)? – largely deterministic range of
signal vs. stochastic ephemeral signal; # orthologs/paralogs intragenomically
vs. interspecifically; targets are constant within cells vs. ephemeral and
temporal-spatial of species distributions and predictors of shedding rate and
eDNA decomposition in ecosystems. ***
In 2009, Bustin and colleagues codified a set of minimal requirements for the publication of quantitative real-time polymerase chain reaction (qPCR) data, ostensibly for use in the fields of clinical medicine and molecular biology, as a means of ensuring rigorous datasets that enable investigators to quantify subtle changes in the expression of particular genes or in the estimation of viral load, for instance. In gene expression studies, fractional changes in the production of intracellular messenger molecules, called mRNAs, which convey genetic information encoded in genes to their proteinaceous or final form, need to be perceived so as to test crucial hypotheses of physiological, medical and even evolutionary and ecological import. Bustin et al. address some of the minimal mandatory standards that need to be reported to ensure continuing robust detection of nucleic acids at low concentrations. These standards were termed the MIQE guidelines: Minimal Information for the publication of Quantitative PCR Experiments.
We will discuss these guidelines and how they compare with eDNA standards in future posts, but for now we will divulge the chief difference between the studies that invoke MIQE and those that involve environmental DNA quantification using qPCR: that concentrations of eDNA, depending upon the circumstances in which it is collected, is likely to be several orders of magnitude lower than many observed changes in the level of mRNA expression or absolute levels of viral genomes present within tissues. Consequently, we need to modify MIQE standards to ensure that eDNA surveys are protected from accusations of too lax standards that may result in the significant incurrence of Types I and II error (false positive and negatives, respectively). Like ancient DNA (aDNA) before, eDNA detection has to elevate these standards to a higher level given the much more ephemeral nature of the target molecules in contemporaneous natural systems.
The Importance of Specificity and Sensitivity
Shared with aDNA and biomedical applications of qPCR assays, is a vital dependency on two touchstones of molecular-based detections: specificity (or: what is the likelihood of incorrectly detecting a non-target, which may lead to Type I error if not wholly specific to the target(s)?) and sensitivity (or: what is the likelihood of detecting extremely low concentrations of the target species, which, if unquantified, may lead to Type II error?). The very first step to minimize these potential sources of error is to design extremely robust assays in silico, which depends, crucially, on having substantial genomic data from target and sympatric (co-distributed) non-target organisms with which to design highly discriminative assays. To do so, genomic information is paramount; one must collect and curate a significant body of genetic sequence information.
Information is Key – An Evolutionary and Population Genetics Rationale for Data Generation and Curation
How much genomic information is enough? And how can we account for high levels of within-species genetic variation, and low levels of genomic sequence divergence between closely-related, sister and/or cryptic species? All excellent questions, and each needs to be answered satisfactorily, or one cannot with good conscience publish or make available a generic assay for the target species in question.
In this post, we have broadly discussed gathering and curating data for designing a species-specific assay to be generally deployed across a species’ range, or at least in the populations from which sequence data were generated. Here at PBI, we are also developing novel ways to tease apart even extremely closely-related species with low levels of nucleotide variation. In a similar vein, we also hope to design population-specific assays that may be able to target individual evolutionary significant units (ESUs) and management or conservation units (M/CUs).
In the next blog, we shall develop further the required minimal standards of a reliable, robust and reputable eDNA qPCR assay, and how these requirements extend and embellish those of the MIQE guidelines. We shall treat in some detail the concepts LOD (limit of detection) and LOQ (limit of quantification) and ask: what are the crucial assay requisite parameters (CARP) for designing, optimizing and validating eDNA assays?
 Bustin et al. (2009). The MIQE guidelines: minimum information for publication of qPCR experiments. Clinical Chemistry55, doi.org/41373/clinch.2008.112797.
 Lahoz-Monfort et al. (2015). Statistical approaches to account for false positive errors in environmental DNA samples. Molecular Ecology Resources, 16, doi: 10.1111/1755-0998.12486.
 Hale et al. (2012). Sampling for Microsatellite-Based Population Genetic Studies: 25 to 30 individuals per population Is enough to accurately estimate allele frequencies. PloS One, doi: 10.1371/journal.pone/0045170
 Luo et al. (2018). Biparental inheritance of Mitochondrial DNA in humans. Proceedings of the National Academy of Sciences, doi: 10.1073/pnas.1810946115
 Benasson et al. (2001). Mitochondrial pseudogenes: evolution’s misplaced witnesses. Trends in Ecology and Evolution, 16, 314-321.
 Smith & Smith (1996). Synonymous nucleotide divergence: what is “saturation”? Genetics, 142, 1033-1036.
Environmental DNA monitoring – eDNA – is at the vanguard of a new wave of technologically advanced monitoring efforts. With roots in soil and paleoecology, eDNA was first used to detect a multicellular aquatic organism– the invasive American bullfrog Lithobates catesbeianus– in a landmark scientific paper by Ficetola et al. in 2008. By applying the widely used, yet highly sensitive, polymerase chain reaction (PCR) – a reaction that ‘amplifies’ a specific DNA sequence in a sample, only if the species’ DNA is present in however little amount – Ficetola and colleagues detected the frogs’ DNA in sediment filtered out of the water column. To understand why, a current working definition of eDNA – adopted by most ecologists – will illuminate: eDNA is “…genetic material obtained directly from environmental samples (soil, sediment, water, etc.) without any obvious signs of biological source material” (Thomsen & Willerslev 2015). Examples of how eDNA is shed by an organism – e.g., here a midland painted turtle Chrysemys picta – are illustrated in the figure below.
In the figure, DNA-containing cells are constantly or periodically shed from the internal linings of the turtle’s gut, reproductive system, through regurgitation of food, the replacement of skin cells and mucus, the egress of waste materials, and through the release of sex cells (i.e., sperm and eggs). Once in the water, DNA is somewhat protected within cells. Eventually, cells are broken down and DNA is released into the aqueous environment whereupon it is to be found in solution. Although eDNA is depleted through a number of biological, chemical and physical processes, it will keep being replenished if the organism is still to be found living in the vicinity. That is to say, the signal of eDNA will be stronger the closer it is to its source, and will also increase when there are more individuals in a local population, if the volume of water remains the same. Therefore, an eDNA signal can be a reliable indicator of the target species’ presence in a given habitat.
What are the chief benefits of eDNA monitoring? First of all, there is no need to physically sample the species, thus minimising disturbance associated by the targeted monitoring of live creatures that are sensitive to stress. Furthermore, as only water samples are taken, and by few people, there will be a decrease in the environmental footprint associated with monitoring efforts perse. Because PCR is a highly sensitive molecular biological assay, eDNA surveys tend to have much higher sensitivities to be able to detect rare or cryptic species than conventional methods (e.g., Schmelzle & Kinziger 2016). As a result, species-specific eDNA surveys are also potentially much more cost-effective than conventional techniques. Furthermore, anyone can take a water sample, following simple instructions, democratising and facilitating citizen science projects across the globe. Indeed, the current crop of companies that offer eDNA detection services are predicated on a model of water samples being collected by lay and technical personnel for processing back in a central laboratory.
However, as eDNA is a nascent technology, uncertainty exists over some of the conclusions drawn from early eDNA studies. However, these issues (i.e., sources of ‘error’) are under ongoing scrutiny by scientists, including here at Precision Biomonitoring, to minimise their impacts. As such, all potential sources of error, as they are currently understood, must be acknowledged and incorporated into any technical development (i.e., design, production and validation of species-specific PCR assays) or standard operating protocols for field surveys. For example, organisms are liable to move throughout their lifetimes. Seasonality has shown to be a strong factor in eDNA detection success (de Souza et al. 2016). It is imperative that surveys are conducted with a thorough knowledge of a species’ ecology, including insight into current distributions and habitat preferences, otherwise inadequate surveying will lead to a false negative result, i.e., inferring a target to be absent when it actually is present; just undetected. Failure to account for false negatives can result in severe financial repercussions if infrastructure projects are subsequently halted, put on-hold or abandoned due to the rediscovery of the target by an intrepid ecologist or member of the public. False negatives can also result from improper assay development, the underestimation of within-species genetic diversity at PCR amplification sites, and by the current disjointed process by which eDNA samples are processed by the majority of eDNA practitioners.
As noted previously, eDNA will decay if left exposed to natural world processes. Therefore, collected eDNA is at risk of post-sampling decay, as there would be no mechanism for eDNA replenishment in the collection vessel, reducing the eDNA signal and potentially failing to garner a positive PCR result. Therefore the risk of eDNA degradation during sampling – particularly on hot, sunny days – and in transit from the field to the laboratory, is highly significant. Inappropriate storage may also destroy eDNA (e.g., water crystal formation during freezing may ‘shred’ DNA molecules). To compound the status quo further, despite the best efforts of contemporaneous laboratories, there remains a significant risk of false positive PCR results mediated by the transportation of aerosolised DNA particles among labs within buildings through ventilation pathways. Most eDNA practitioners seek to physically separate the processing of eDNA samples (e.g., filter papers or precipitated water samples) with downstream PCR detection, but even that is far from fool-poof.
Here at Precision Biomonitoring, we are set to unveil a state-of-the-art platform that will seek to eliminate, or minimise, these sources of eDNA analytical and sampling error, through the eradication of transit stages to a central laboratory and the application of standard operating procedures. Moreover, we will further the cause of a democratised biomonitoring field in which no technical specialty is required to conduct sophisticated PCR-based species-specific assays. Our system, using bespoke PCR assays, will yield PCR eDNA results in real-time (< 2 hours from water sampling to PCR read-out), which can then be immediately disseminated to colleagues via the cloud.
It is our aim to give those working at the coalface of biodiversity monitoring (from professional ecologists to local citizen science projects), the power to conduct highly rigorous, and potentially highly coordinated, targeted eDNA surveys to better vouchsafe our world’s biodiversity heritage for all generations to come.
 Ficetola et al. (2008). Species detection using environmental DNA from water samples. Biology Letters, 4, 423-425.
 Thomsen & Willerslev (2015). Environmental DNA – An emerging tool in conservation for monitoring past and present biodiversity. Biological Conservation, 183, 4-18.
 Schmelzle & Kinziger (2016). Using occupancy modelling to compare environmental DNA to traditional field methods for regional-scale monitoring of an endangered aquatic species. Molecular Ecology Resources, 16, 895-908.
 de Souza et al. (2016). Environmental DNA (eDNA) detection probability is influenced by seasonal activity of organisms. PLoS One, doi: 10.1371/journal.pone.0165273