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Home > Resources > COVID-19 > Molecular-based Tests

Molecular-based Tests for COVID-19

The diagnostic testing field for COVID-19 is rapidly evolving and improving in quality every day, with many tests focused on diagnosing patients with active viral infections. Diagnostics able to detect current, active infections are typically molecular-based diagnostics, which inform researchers of the presence of the pathogen, either by identifying its genetic material or identifying unique markers of the pathogen itself. The viral genomic material for SARS-CoV-2 is ribonucleic acid (RNA), which remains in the body only while the virus is still replicating. There are also rapid antigen tests in development that act by detecting specific surface markers on the outside of the virus; none of these tests have received FDA emergency use authorization (EUA), nor are any available on the market at this time.

Molecular diagnostics usually require samples from the patient that are likely to contain virus, such as nasopharyngeal swabs or sputum samples. Some pathogens may also be detected in feces, urine, or blood. For respiratory diseases like COVID-19, nasopharyngeal swabs have been considered to be the most reliable, as they sample an area of the respiratory tract where the virus appears to first infect an individual. This site is relatively easily accessed, compared to the final site of viral infection: the lower respiratory tract. Consequently, the nasopharyngeal tract likely has (1) active virus replication and (2) sufficient amounts of virus to be detected in kits. Many tests now available or in development can use saliva or nasal swabs that facilitate easier sampling procedures for healthcare providers and patients.

If you are looking for information on serology (antibody) tests, please visit our serology tracker or visit our recent report outlining the needs of a national serology strategy.

 

Disclaimer

This website is updated twice weekly and includes only tests with EUA status, either from commercial manufacturers or laboratory-developed tests. In the Assays in Development section, novel methodologies for direct detection of SARS-CoV-2 are described. While the described methodologies are appropriately cited, exact specifications of these tests will not be directly incorporated into the tracker tables until they receive FDA EUA status. This website is not intended to be used as a reference for funding or grant proposals. Non-inclusion in this list should not be interpreted as a judgment on the validity or legitimacy of tests.

If you are a manufacturer or research institution that has an EUA-approved molecular test that is not yet listed and you would like to contact us, please submit your information here. Submission of this form does not guarantee inclusion on the website, but it will allow us to verify your test's information so that we can accurately describe the test if it is included.

 

Note on sensitivity and specificity data

Where available, we list the manufacturer-reported sensitivity and specificity data. A highly sensitive test should capture all true positive results. A highly specific test should rule out all true negative results. These measures are not independently validated by the Johns Hopkins Center for Health Security. If sensitivity or specificity is not listed, it was not available from the manufacturer at the time of posting. When available, the number of samples used for sensitivity/specificity definitions are listed in the product description.

It should also be noted that the terms “sensitivity” and “specificity” may not appear in the manufacturers’ information sheets, but rather these values are often reported as “positive percent agreement” and “negative percent agreement.“ Sensitivity may also be measured by calculating the limit of detection (LOD), which is the lowest detectable number of virus copies in a sample at which the test will return a positive result at least 95% of the time. Essentially, a lower limit of detection indicates a more sensitive test, with fewer viral copies per sample necessary to elicit a positive test result. The FDA recommends that manufacturers use these terms to indicate that a nonreference standard was used when evaluating the test.

This resource was created and is updated by Amanda Kobokovich, MPH, Rachel West, PhD, and Gigi Gronvall, PhD.

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How Do Molecular Tests Detect SARS-CoV-2?

Most molecular tests for SARS-CoV-2 use the process of real-time reverse transcriptase quantitative polymerase chain reaction (rRT-PCR). Throughout this site, the majority of molecular kits are labeled as rRT-PCR. The tests included have been referred to as their manufacturers have listed them, though in some cases they use qPCR. If a test provides quantitative information, and not merely qualitative (yes/no), this requires quantitative (q)PCR and not PCR alone. These tests rely on the same basic steps:

  1. detecting genetic material (DNA or RNA) specific to the pathogen;
     
  2. amplifying (making more copies of) detected region of the genetic material of the pathogen; and
     
  3. producing an output measurement of the amount of amplified genetic material, if present in the sample.
 

RT PCR figure


In step 1, researchers design small pieces of single-stranded DNA called “primers,” which precisely match a specific area of the viral genome. Primers then attach or “anneal” to the specific areas of the viral genome and provide the backbone for amplification of that region. When building primers, researchers seek specific parts of a viral genome that are unique to the virus in question. Because the viral RNA is too small to visualize and detect in such small quantities, signal amplification is needed.

For most viral RNA-based genomes, another step called reverse transcription is needed. The SARS-CoV-2 genome is made of RNA, which is less stable and more sensitive to UV radiation and breakdown by enzymes than DNA. Therefore, RNA extraction and use in testing must be done carefully to preserve the genetic material. Reverse transcription uses proteins called reverse transcriptase enzymes to translate RNA into DNA, which is a more stable molecule.

In step 2, the area in which the primers attach or anneal is amplified in repeated cycles. These cycles are designed to closely mimic the natural DNA replication processes in all human cells. In most PCR assays, amplification cycles rely on programmed temperature changes that encourage the double-stranded DNA to split apart, allow replication enzymes to create a new copy of the DNA, and then close the newly formed strands back together. For this reason, most PCR assays must take place in machines called “thermocyclers,” which allow for adjustments in cycle timing, temperature, and number of iterations. Exceptions to this process are isothermal methods, such as loop-mediated isothermal amplification (LAMP), which do not require heating cycles to amplify the target DNA. Step 2 continues until the researchers have synthesized enough genetic material for them to be able to read.

In step 3, the output from the amplification process is studied, and researchers are able to visualize the virus within the sample. In real-time RT-qPCR machines, the readable output is shown in the form of fluorescence that the amplified material gives off as its quantity increases after multiple amplification cycles.

While not all tests listed below are rRT-qPCR tests, all molecular tests are developed to inform researchers of the presence of the pathogen, either by identifying its genetic material or identifying unique markers of the pathogen itself. An amplification step is crucial for these tests because otherwise researchers would be unable to easily and rapidly detect the presence of such small molecules.

 

Common Types of Molecular and Antigen Tests Being Developed for SARS-CoV-2

For a concise review of these tests and methodologies, please see the table below.

  1. Reverse transcriptase quantitative polymerase chain reaction (rRT-qPCR): Identifies and quantifies the presence of infectious agents in a sample through the process of detection, amplification, and output measurement.

    Pros: Highly specific, can be modified as the virus evolves to fit new iterations, hundreds of samples can be run at once, and is extremely sensitive (low limit of detection).

    Cons: Requires trained personnel and special equipment, primer and probe design must be exact, samples may need to be transported to the lab, and the test itself takes 1 to 3 hours. Samples must undergo RNA extraction before the test can be run, adding another 1 to 2 hours, depending on lab capacity.
     

    It can involve 1 or 2 steps, depending on the reagents and kits used. The process, described above, transcribes viral RNA into DNA, if present in the sample, for amplification and visualization. These tests typically take 1 to 3 hours, and hundreds of samples can be processed at once. Results can be read quantitatively or, more simply, can be used to indicate the presence or absence of infection. In either quantitative or qualitative iterations, rRT-qPCR tests require special equipment and trained lab technicians to correctly obtain and interpret results. In addition to laboratory personnel and equipment needs, specialized reagents called primers and probes are necessary for the test to be run. These primers and probes must be specifically designed to bind only to viral RNA of interest. The primers allow for amplification of the RNA while the probes allow the amplified RNA to give off a fluorescent signal that is read and quantified by the PCR machine.

     
     
  2. Reverse transcription loop-mediated isothermal amplification (RT-LAMP): Rapid amplification of viral genomic material coupled with a color- or light-based readout, and it can be performed at a single temperature, unlike rRT-PCR.

    Pros: Requires a single temperature only, rapid (minutes to results), point-of-care appropriate, highly sensitive and specific to defined SARS-CoV-2 sequences.

    Cons: Primer design must be exact, incorrectly folded primers can cause “debris” that can interfere with the reaction, difficult to quantify virus. Samples must undergo RNA extraction before the test can be run, adding another 1 to 2 hours, depending on lab capacity.
     

    This method is a more recent development in point-of-care diagnostics. In contrast to rRT-qPCR, which requires rapid cycling of multiple temperatures to amplify nucleic acids, LAMP reactions occur at a single isothermal temperature, between 63°C and 65°C. This makes the reaction much simpler, faster, and easier in a point-of-care setting. In addition, LAMP reactions typically produce a color or cloudiness change in the reaction mixture that are often visible to the eye. Again, this simplifies the protocol for use in a variety of settings.

    The main principle behind RT-LAMP is a reverse transcription step (RNA into DNA), followed by the addition of 6 primers that bind to the gene of interest. At the correct temperature, these 6 primers bind the target DNA, loop around to create circular constructs, and extend the DNA. Each “loop” opens up a new site for primers to bind, amplifying the gene further. This process allows for rapid, exponential increases in the gene of interest. If probes, dyes, or a fluorophore are included in the mixture, there can be a visible change during the reaction that can be measured by eye or by special equipment.

    Thus, an RT-LAMP reaction on a patient sample containing the virus will elicit a visible change in the reaction within minutes. RT-LAMP can be hundreds of times more sensitive than RT-qPCR, meaning it can pick up on even smaller concentrations of virus within the sample than some RT-qPCR assays. Software is available to design the primer sets for RT-LAMP, as the sensitive and complicated process can easily be thrown off by poorly designed primers. The debris that can interfere with reactions includes hairpin loops and primer-dimers, which can form if the primers accidentally bind themselves. Researchers have also identified ways to tag patient samples with “barcodes” and then amplify them as one pooled sample. This process, called LAMP-Seq, cuts down on equipment needs by pooling many patient samples into 1 reaction tube that can later be identified and separated for analysis.

     
     
  3. Recombinase polymerase amplification (RPA): Detects DNA sequences through precise matches of an enzyme called recombinase that can pull apart (displace) DNA strands and then amplify specific viral genes.

    Pros: Requires a single temperature only, rapid, less equipment required, highly sensitive and specific to defined SARS-CoV-2 sequences.

    Cons: Primer design must be exact, can be difficult to quantify virus, and debris can interfere with reaction. Samples must undergo RNA extraction before the test can be run, adding another 1 to 2 hours, depending on lab capacity.
     

    In the case of SARS-CoV-2, this would need to be coupled with a reverse transcriptase step to take a viral gene from RNA to DNA. Like RT-LAMP, this method is also isothermal, meaning only 1 temperature is necessary to carry out the reaction. The reaction causes exponential amplification of DNA, is rapid, easy, and requires few reagents.

    The main idea behind RPA depends on primer binding to a DNA sequence of interest, where a recombinase enzyme can then bind. This recombinase splits apart the 2 strands of DNA and is then stabilized by special proteins. With its job done, the recombinase leaves the DNA open for easier amplification. In RPA, primers are designed to be opposing over the same stretch of DNA, so that every time the extension is completed, there are 2 resulting copies of DNA. This contributes to the rapid amplification. The constant binding of primers, and opening by recombinase, also contributes to the rapid, exponential amplification.

    This rapid amplification method is very sensitive, requiring very little DNA starting material. However, primers must be designed carefully and temperature controlled, so that the enzymes can properly assemble and disassemble the DNA.

     
     
  4. CRISPR-based diagnostics: Utilizes the highly specific targeting and cleaving action of CRISPR-Cas systems to locate and cut a specific part of SARS-CoV-2 RNA sequence. The cleaving action results in a visual signal that indicates the presence of the virus.

    Pros: Highly specific to defined sequences of SARS-CoV-2, less equipment is required, very rapid results (minutes), very sensitive.

    Cons: Requires exact primer design, can require troubleshooting and specific design of all components (enzymes, primers, reporters). Samples must undergo RNA extraction before the test can be run, adding another 1 to 2 hours, depending on lab capacity.
     

    CRISPR-based tests can be more rapid than PCR, if coupled with LAMP, and do not require the specialized equipment that PCR does.

    The system has 2 main components: the CRISPR (clustered regularly interspaced palindromic repeats) sequence, which is designed to include guide RNAs that match parts of the viral genome, and the Cas enzyme, which cuts the RNA where the CRISPR sequence matches. The Cas enzyme is like a construction crew, ready to demolish a certain site. The guide RNAs, which the researcher designs, are the GPS for the crew, telling the enzyme where to cut. These systems were first discovered in bacteria, as a sort of bacterial immune response to viral infections. More recently, CRISPR has been adapted for a wide range of uses, particularly in gene editing, because of its ease of use, quick turnaround time, and very specific cleavage of nucleic acid sequences by the Cas enzyme. Different Cas enzymes (Cas 9, Cas 13, etc) cleave different types of nucleic acids. In CRISPR diagnostics for COVID-19, they must use a Cas that can recognize and cleave RNA (rather than DNA). Cas12 is such an enzyme, and it was recently used in the DETECTR system for rapid diagnosis of SARS-CoV-2, with limited cross reactivity. Cas13 has been used in field detection of dengue virus, using the SHERLOCK system.

    In order to use CRISPR-based diagnostics, researchers create the following:

    1. Guide RNAs that are designed to be complementary to viral RNA. These direct the Cas enzyme to the viral gene, where it can cleave the RNA. This activates the Cas enzyme.
    2. A special reporter that has fluorescent molecules or color, and an anchor molecule, like biotin, or a quenching molecule that inhibits light readout. If the reporter stays intact, then the anchor/quencher will prevent the reporter from being detected. If the reporter is cleaved by the Cas enzyme, then the signal can be emitted.
    3. The reporter is mixed with enzymes, guide RNAs, and patient sample material. The reporter is then cleaved only upon the guide RNAs’ binding to the proper target in viral RNA. Once the Cas enzyme has recognized the viral target, it can also cleave the bystander reporter sequence. The cleaved reporter can then bind the “test” strip, while any non-cleaved reporter remains at the control strip anchored by the biotin. In the case of a quenching molecule, once the reporter is cleaved, the fluorescence can be emitted.
    4. The researcher can read the color- or fluorescence-based result, which is sometimes on a lateral flow strip (similar to a pregnancy test or RDT serology test).

     
     
  5. Rapid antigen test: Detects easy-to-find surface markers on the outside of the virus and avoids extraction and amplification steps. Researchers or clinicians collect samples from easy-to-reach areas (like the nasal passage) where the virus tends to replicate the most.

    Pros: Can detect active production of viral proteins, fairly rapid tests (minutes to results), RNA extraction and amplification steps not needed.

    Cons: Requires extensive design and troubleshooting of test antibodies that will react to viral proteins in the sample, deep knowledge of viral proteins produced in various tissue environments, and the test may yield false negatives if the viral protein production is low or if there is not enough virus replication in the sampled area.
     

    These tests can be fairly rapid, using lateral flow assays. These types of tests are common in diagnosing illnesses such as strep throat. They may seem similar to serology tests, but they are different in 3 important ways: (1) they only detect active infection by the virus; (2) they do not detect antibodies, but rather viral proteins that the virus is producing within the human host; and (3) they are typically not blood- or serum-based.

    The main principle behind antigen detection tests is the use of designed, synthetic antibodies to probe a patient sample for the presence of viral proteins. During an infection, viral proteins are often produced and can be detected in the blood or in other tissues or secretions. A patient sample is mixed with buffer, reagents, and synthetic antibodies. Importantly, these antibodies are designed so that they bind only to certain viral proteins; they will not bind to viral proteins from other viruses. If the proteins are present in the patient sample, then the synthetic antibodies will bind, and these complexes can be detected on a lateral flow strip with a color- or light-based readout. The synthetic antibodies used must be very carefully designed so that they properly bind the protein produced by the virus during a natural infection, including any special folding or protein modifications that could be present. This method also depends on there being sufficient viral proteins in the sample tested for them to be detected. In other words, these tests sometimes have sensitivity issues and can be less sensitive than RT-qPCR or RT-LAMP methods. These tests also must be very specific, so that similar proteins produced by other pathogens (eg, other human coronaviruses, influenza, etc) are not detected by these synthetic antibodies. While there are some shortcomings to the sensitivity of this method, it is rapid and could be used in a point-of-care setting.

     
 

Summary of Types of Molecular and Antigen Tests Being Developed for SARS-CoV-2

There are five common types of molecular tests:

Type of TestTime to ResultsWhat It Tells UsLimitationsLink
rRT-qPCR2-4 hoursThe presence of an active infection, by targeting specific gene sequences of SARS-CoV-2. This can be quantitative but is usually qualitative (yes/no). It typically has a very low limit of detection, around 100 viruses/mL.The time needed to complete the test; trained personnel and special equipment to analyze the resultsrRT-qPCR
LAMP15-60 minutesThe presence of an active infection, by targeting specific gene sequences of SARS-CoV-2. This is typically qualitative (yes/no). It relies on specially designed primers that help create the looped structures needed for amplification. It is very rapid and does not always require special equipment (can be measured by eye in some cases). It has a very low limit of detection of 125 viruses/mL.Designing the primers needed can be complex, and debris can interfere with the reaction. It is also difficult to quantify the results (level of viral infection).LAMP
RPA15-60 minutesThe presence of an active infection, by targeting specific gene sequences of SARS-CoV-2. This is typically qualitative (yes/no). It relies on the recombinase enzyme. It is very rapid and does not always require special equipment. It has a very low limit of detection of 125 viruses/mL.Designing the necessary primers can be complex, and debris can interfere with the reaction. It is also difficult to quantitfy the results (level of viral infection).RPA
CRISPR-based diagnostics15-60 minutesThe presence of an active infection, by targeting specific gene sequences of SARS-CoV-2. This is typically coupled with LAMP, but this is not always necessary. Results are typically visible to the eye, not requiring special equipment.Requires expert, specific design of components (enzymes, primers, reporters).CRISPR
Rapid antigen test (RAT)15-30 minutesThe presence of an active infection, by detecting specific viral proteins present in a patient sample. This is typically coupled with lateral flow assays to display results that can be read by eye.Requires very careful design of synthetic antibodies, deep knowledge of viral proteins produced in various tissue environments, and may yield false negatives if the viral protein production is low.RAT
 

Current Molecular and Antigen Tests with FDA EUA Status

We have compiled a list of commercial and laboratory-developed tests that have received FDA Emergency Use Authorization. There are 2 main sections:

  1. Commercial tests: These tests (sometimes referred to as kits) have been developed by industry professionals in biotechnology. These kits are typically available for purchase by healthcare providers or by researchers, undergo internal validation, and researchers manufacture the tests themselves. These tests can be purchased and completed at labs and facilities other than the manufacturing facility.
  2. Laboratory-developed tests: These tests have been developed by medical and research professionals. They are for use only in the institution in which they were developed. For instance, if a test was developed in a hospital, the test can only be used in that hospital. The laboratory developing and performing the test must be certified under Clinical Laboratory Improvement Amendments of 1988 (CLIA), 42 U.S.C. §263a, to perform high-complexity tests. These kits are not for commercial sale; they undergo internal validation and list the needed reagents in the protocols.
 

Trends in Development

The field of molecular diagnostics is rapidly expanding in response to the need for rapid, large-scale testing for SARS-CoV-2. The development of novel assays or technologies is important for finding new and creative ways to increase testing capacity. The information presented in this section is meant to illuminate industry trends in response to the evolving needs of the COVID-19 pandemic. The trends presented in this section are not intended to be comprehensive, but rather demonstrate some techniques or technologies that may move into the traditional development pipeline in the future.

It should be noted that many of the sources in this section are from preprints, meaning the research has not yet undergone peer review and may contain major or minor errors that might invalidate certain findings. The results of this research should not be presented as established fact or be used to guide clinical or health-based decision making.

As of 5/5/2020
  1. RT-LAMP: Making RT-LAMP accessible at point-of-care settings (including making reagents lyophylized, using dried blood spots instead of venipuncture). Because RT-LAMP is so rapid and requires less specialized equipment than PCR, this is an attractive option for point-of-care testing. However, traditional blood samples require very careful processing and storage. The ability to use a finger prick on a dried blood spot, stored at room temperature for later use, would allow for more extensive sample collection.
     
  2. Barcoding and pooling PCR, including technologies such as LAMP-Seq. This could allow for analysis of many more samples at once, which would expand our ability to test more patients per day.
     
  3. Using CRISPR-based methods, such as SHERLOCK and DETECTR, to detect RNA present in a variety of types of samples and in 1-step formats. These systems are as specific as PCR but far more rapid—taking minutes rather than hours. They are also much more malleable to adaptation if the virus mutates.
     
  4. Using microfluidics-based cartridge systems for all-in-one testing systems, seen here and here. These systems improve ease of use by removing the sample transportation and RNA extraction steps that are often time-consuming and risky. RNA extraction can be easily contaminated if stringent protocols are not followed, leading to false-negative results. A cartridge system allows for a sample collection, followed by rapid extraction, amplification, and detection. The microfluidics can improve reagent mixing automatically, so that each reaction is uniformly processed.

 

 

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