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Antigen, Antibody or PCR tests: What is the difference?

Antigen testing, antibody testing, PCR (Polymerase Chain Reaction) tests. . . what’s the difference? Which one is the most reliable? Which is right for my situation?

As the number of COVID-19 cases climb, the need to test more people increases. Even with the hope offered by vaccines, there is no doubt that coronavirus testing is still required and remains a critical part of controlling the spread of this global pandemic. The reason for this is simple – accurate and reliable testing is a proven and effective way to slow the spread of SARS-CoV-2, the virus that causes COVID-19.  Slowing the spread of this deadly virus will save lives, reduce burden on our health care systems and keep our workforce safe. There has been a lot of discussion in the news about the types of tests currently on the market and when you factor in the ones that are in development it can be hard to decide which testing approach is the right fit for given situation.  

Antigen, antibody, and PCR testing all have their pros and cons.  It is important to distinguish that these tests don’t all look for the same things. As new tests come to market, in particular antigen and antibody tests, it is important to understand these differences and learn which test is right for which situation.

For any approved COVID-19 test currently on the market, be it antigen, antibody or PCR based, there should not be any reliability concerns, if these tests are conducted properly, by the appropriate personnel, and using the appropriate space, equipment, and reagents to conduct the test.  The real question should be about which type of testing to use for specific information regarding the COVID-19 situation. Let’s compare the three testing methodologies paying attention to things like what the test detects, how and where the test is typically performed and most importantly what kind of information the test provides and in what situation is it most helpful.

PCR tests are currently the most commonly used type of test to detect whether a person has an active COVID-19 infection.  These tests look for genetic material from the SARS-CoV-2 virus that causes COVID-19.  Once a sample is collected by a health care professional, these tests can be done quickly (1-2 hours) at the place of testing or can be done in a large central lab (anywhere from a few hours to a few days).  These test are the gold standard and can be used to diagnose a COVID-19 infection without any additional confirmatory tests. They often require specialized equipment to run the test (ex. a thermocycler) and with the right set up can be used in a remote setting.

Antigen tests, often referred to as “rapid tests” are also used to detect if a person is currently infected, but tend to perform better if a person has had symptoms for a few days. These tests are designed to detect pieces of the proteins that make up the virus.  Once a sample is collected by a health care provider, antigen tests can often be done quickly while you wait (a few minutes to about an hour or so), but occasionally they are sent to a central lab.  One thing to keep in mind is that for the current antigen tests on the market, an additional PCR test is required to confirm an antigen test result. Since antigen tests typically don’t require a lot of special equipment, they can be well suited to remote settings.

Lastly, antibody tests (also called serology tests, but not to be confused with antigen tests) look for COVID-19 antibodies in the blood and are the least commonly used testing method for population wide COVID-19 testing.  Antibodies are molecules made by the immune system in response to SARS-Cov-2 and so a positive antibody test can indicate that the person was exposed to or infected by the virus in the past. Because antibody tests require a blood draw, samples are almost exclusively taken by a health care professional and sent to a lab for testing.  While antibody tests are not best suited to determine if a person is currently infectious, understanding who has been exposed to the virus in the past can be helpful. 

The following table, adapted from the DC Health publication, summarizes and expands upon the differences between these three types of testing approaches.

TOPICPCR TESTANTIGEN TESTANTIBODY (SEROLOGY) TEST
WHAT IS THE TEST LOOKING FOR?PCR tests look for pieces of genetic material from SARS-CoV-2, using samples from the nose, throat, or other areas in the respiratory tract.Antigen tests look for pieces of proteins that make up the SARS-CoV-2 virus, using samples from the nose, throat, or other areas in the respiratory tract.Serology tests look for antibodies1; molecules made by the immune response against SARS-CoV-2 in the bloodstream.
WHAT DOES THE TEST TELL YOU?Determines if the person has an active (i.e., current) infection.Determines if the person has an active (i.e., current) infection.Determines if there was a past infection.
HOW IS THE TEST PERFORMED ON PEOPLE?In most cases, a nasopharyngeal or nasal swab is taken by a healthcare provider and tested; however, oral swabs and saliva are also acceptable. Sometimes the test can be run while you wait, but most commonly, the swab needs to be sent to a lab for testing. In most cases, a nasopharyngeal or nasal swab is taken by a healthcare provider and tested. Most often the test can be run while you wait, and occasionally the swab needs to be sent to a lab for testing. In most cases, a blood sample is taken by a healthcare provider and is sent to a lab for testing.
WHEN IS IT HELPFUL?It can help determine who has an active infection, regardless of a person’s symptoms.
• It can help identify people who are contagious to others.
• In communities where transmission rates are low and mitigation efforts are effective, PCR testing is more reliable at detecting active infection.
• It accurately identifies people who are or are not infected with SARS-CoV-2.
For someone with symptoms, it can be used as a point-of-care test to quickly determine who has an active infection.
• It can help identify people who are contagious to others.
• It is a less expensive test than PCR.
• It performs best 5-7 days after symptoms onset, in the early stages of infection with SARS-CoV-2.
It can identify people who had an infection in the past, even if they had no symptoms of the illness.
• In some cases, it could help determine when COVID-19 occurred since we know that IgM forms before IgG and that IgM goes away before IgG.
• It can help determine who qualifies to donate convalescent plasma (a blood product that contains antibodies against COVID-19 and can be used as a COVID-19 treatment).
• If many people take the test in a community, it can help public health leaders and researchers know what percentage of the population already had COVID-19.
LIMITATIONSIt does not help determine who had an infection in the past.
• It also gives you a result for the point and time when the specimen was collected and cannot predict if you will remain negative. For example, if you are quarantining after an exposure, a negative test does not allow you to stop quarantining.
• In some people, the virus can be found by PCR in the nose and throat for several weeks, even longer than their infectious period (the time they are contagious to other people).
• This test requires certain kinds of swabs and reagents that may be in short supply.
Best results are achieved with those who are symptomatic, and it will not help determine who had an infection in the past.
• Antigen tests have lower sensitivity3 than PCR tests, so there may be false-negative results.
• In persons with symptoms or known exposure, negative tests must be treated as a preliminary result and confirmed with PCR testing.
If used too close to the beginning of an infection, this may result in a negative test which is why it must not be used to detect active COVID-19 infection.
• In areas where there have not been many cases of COVID-19, many of the positive test results will be false positives (see Positive Predictive Value2). Some antibody tests have low sensitivity3 and specificity4 and thus may not produce reliable results.
• Some antibody tests may cross-react with other coronaviruses that are not SARS-CoV-2, leading to false test results.
• We do not know yet if having antibodies to the virus that causes COVID-19 can protect someone from getting infected again or, if they do, how long this protection might last. Until scientists get more information about whether antibodies protect against reinfection with this virus, everyone must continue to take steps to protect themselves and others. This includes staying at least 2 meters away from other people (social distancing), even if they have had a positive antibody test.
WHAT DOES A POSITIVE TEST RESULT MEAN?A positive PCR test means that the person tested has an active COVID-19 infection.A positive antigen test means that the person tested has an active COVID-19 infection.A positive antibody test means that the person tested was infected with COVID-19 in the past and that their immune system developed antibodies to try to fight it off. Until scientists get more information about whether antibodies protect against reinfection with this virus, everyone must continue to take steps to protect themselves and others. This includes staying at least 2 meters away from other people (social distancing), even if they have had a positive antibody test.
WHAT DOES A NEGATIVE TEST RESULT MEAN?A negative PCR test means that person was probably not infected at the time their sample was collected.
It doesn’t mean that someone won’t get sick – it only means that they didn’t have COVID-19 at the time of testing.
A negative antigen test means that SARS-CoV-2 viral proteins were not detected.
In persons with symptoms or known exposure, a negative test does not rule out COVID-19. The individual must quarantine until a confirmatory PCR test can be completed.
A negative antibody test means that the person may not have had COVID-19 in the past. However, they could still have a current infection, and the antibody test was conducted too soon to give a positive result.

1 The body forms antibodies to fight off infections. Immunoglobulin M (IgM) is the first antibody formed against a germ, so it appears on tests first, usually within 1-2 weeks. The body then forms immunoglobulin G (IgG), which appears on tests about two weeks after the illness starts. IgM usually disappears from the blood within a few months, but IgG can last for years. Some antibody tests test for IgM and IgG, and some only test for IgG.

2 Positive predictive value is a measure of how likely it is that a positive test is a true positive rather than a false positive. This is dependent on how many people in the population have tested to have had the disease. When there are very few people in the population that have had the disease, then there is a higher chance for a false positive. When many people in a population have had the disease, then there is a higher chance that a positive test is a true positive.

3 Sensitivity is sometimes called the “true positive rate.” It measures how frequently the test is positive when the person tested has the disease. For example, when a test has 80% sensitivity, the test detects 80% of patients with the disease (true positives). However, 20% of patients with the disease are not detected (false negatives) by the test.

4 Specificity is sometimes called the “true negative rate.” It measures how frequently the test is negative when the person tested doesn’t have the disease. For example, when a test has 80% specificity, the test correctly reports 80% of patients without the disease as negative (true negatives). However, 20% of patients without the disease are incorrectly identified as testing positive (false positives) by the test.

It’s important to remember that no test is 100% accurate all the time. Some things that may affect the test’s accuracy include:

  • You may have the virus, but the swab might not collect it from your nose or throat.
  • The swab or mucus sample may have been accidentally contaminated with the virus during collection or analysis.
  • The nasal or throat swab may not have been stored at the correct temperature before it was analyzed.
  • The chemicals used to extract the virus genetic material and make copies of the virus DNA may not work correctly.

COVID-19 Testing at Various Infection Stages

It’s clear that each testing approach has its pros and cons, and there is a place for all three in the global effort to control the spread of COVID-19. As per the chart COVID-19 Testing at Various Infection Stages, it is clear that every day of infection counts, and that tests with higher sensitivity allow for earlier detection of the virus, and we can see what technology can be used to detect the virus at earlier stages. When it comes to screening people for infection, PCR testing remains the most common type of test used in Canada and the USA.  According to Dr. Christina Wojewoda, a pathologist at the University of Vermont and the vice-chair of the College of American Pathologists’ microbiology committee, we “should be diagnosing people with PCR tests because they are the most accurate”. Our SARS-CoV-2 real-time RT-PCR test kit can bring PCR testing to workplaces, community settings, and pop-up service centres, even in the most remote communities.  Our approach provides a safe, easy, and accurate way of testing to ensure a safe workplace.

Health Canada Approves Precision Biomonitoring’s Canadian-Made Rapid SARS-CoV-2 Testing Device

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.  

Sales Inquiries:
info@pbidiagnostics.com

Media Contact:
Meredith Adams
416-459-7086
Madams@national.ca

Implementing a COVID-19 testing laboratory at your workplace

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.

The TripleLock™ Advantage

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 natural habitat.

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 TripleLock™ 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 [2].

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 [4]. 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: On-site Detection

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 challenges.

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 [5]. 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.

References:

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. doi:10.1002/nafm.10055

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. doi:10.1111/2041-210X.12994

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. doi:10.1111/2041-210X.12206

Exploring MIQE-Like Standards for qPCR-Based Molecular Detection of eDNA

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 reference gene(s).

Generic workflows for conducting A) gene expression qPCR and B) eDNA qPCR

Consider then, 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 [2]), 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.

Performing qPCR 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.

Bottom-line: 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 specificity.

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.

[1] Bustin et al. (2009). The MIQE guidelines: minimum information for publication of qPCR experiments. Clinical Chemistry 55, doi.org/41373/clinch.2008.112797.

[2] Barnes and Turner (2016). The ecology of environmental DNA and implications for conservation genetics. Conservation Genetics, 17, 1-17.

[3] 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.

 [4] 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.

[5] Evans et al. (2016). Quantification of mesocosm fish and amphibian species diversity via eDNA metabarcoding. Molecular Ecology Resources, 16, doi/10.1111/1755-0998.12433.

[6] Hunter et al. (2016). Detection limits of quantitative and digital PCR assays and their influence in presence-absence surveys of eDNA. Molecular Ecology Resources, 17, doi/10.1111/1755-0998.12619.

[7] 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. ***

Detecting eDNA – The Importance of Assay Specificity and Sensitivity Part I: An Introduction to the MIQE Guidelines and DNA Sequence Database Generation & Curation for qPCR Assay Design

MIQE – a Brief Introduction

In 2009, Bustin and colleagues[1] 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?)[2]. 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.

Coda

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)[9].

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?

[1] Bustin et al. (2009). The MIQE guidelines: minimum information for publication of qPCR experiments. Clinical Chemistry 55doi.org/41373/clinch.2008.112797.

[2] Lahoz-Monfort et al. (2015). Statistical approaches to account for false positive errors in environmental DNA samples. Molecular Ecology Resources16, doi: 10.1111/1755-0998.12486.

[3] 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

[4] Luo et al. (2018). Biparental inheritance of Mitochondrial DNA in humans. Proceedings of the National Academy of Sciences, doi: 10.1073/pnas.1810946115

[5] Benasson et al. (2001). Mitochondrial pseudogenes: evolution’s misplaced witnesses. Trends in Ecology and Evolution16, 314-321.

[6] Smith & Smith (1996). Synonymous nucleotide divergence: what is “saturation”? Genetics142, 1033-1036.

[7] Felsenstein J (2003). Inferring Phylogenies, Sunderland (Sinauer).

[8] Crête-Lafrenière et al. (2012). Framing the Salmonidae family phylogenetic portrait: a more complete pictire from increased taxon sampling. PloS One, doi: 10.1371/journalpone.0046662.

[9] Palsbøll et al. (2006). Identification of management units using population genetic data. Trends in Ecology and Evolution22, 11-16.

Using Environmental DNA (eDNA) to Monitor Aquatic Ecosystems

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

Blog pic 1

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

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

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

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

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

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

[1] Ficetola et al. (2008). Species detection using environmental DNA from water samples. Biology Letters4, 423-425.

[2] Thomsen & Willerslev (2015). Environmental DNA – An emerging tool in conservation for monitoring past and present biodiversity. Biological Conservation183, 4-18.

[3] Schmelzle & Kinziger (2016). Using occupancy modelling to compare environmental DNA to traditional field methods for regional-scale monitoring of an endangered aquatic species. Molecular Ecology Resources16, 895-908.

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