SHERLOCK: A CRISPR-based diagnostic test to detect COVID-19
CRISPR-Cas is a well-known biotechnological tool for gene editing but recently scientists all over the globe are trying to harness this technology to develop diagnostic kits for rapid COVID-19 diagnosis. In order to understand the CRISPR-based COVID-19 diagnostic method, it is important to know what is the CRISPR-Cas system and how it works?
What is the CRISPR-Cas system?
The CRISPR-Cas system was originally adapted from a naturally occurring genome editing system in bacteria to fight against invading viruses. CRISPR stands for Clustered Regularly-Interspaced Short Palindromic Repeats and Cas stands for CRISPR-associated genes. The full form of CRISPR suggests that in the genome of bacteria, there are palindromic repetitive sequences that are short segments of DNA- 20–40 bp in length, and the spacer DNA is present between these repeat sequences. Between the repeat sequences, spacers are present.
Each segment of the spacer is unique which matches the DNA found in the viruses. These spacer sequences are some kind of acquired immune system in bacteria that protect the bacteria from getting infected with viruses. In other words, these spacers function as an immunological memory bank, storing sequences from previous encounters with invading viruses. During infection with the virus, these spacers are used by the bacteria as recognition elements to find the matching virus genome and destroy it.
After infection, the CRISPR locus is transcribed into a precursor-CRISPR RNA and cas genes are translated to form Cas proteins including Cas9. The pre-crRNAs are processed to form mature crRNAs, each containing the spacer DNA.
The mature crRNAs then form complexes with a second RNA called tracrRNA and CRISPR associated protein Cas9 to form a complete search complex. These RNAs together are called guide RNA as they guide the Cas9 to interact with the viral DNA sequences matching the 20 nucleotides in the guide RNA. Thus allowing the Cas9 to make a double-stranded break in the target viral DNA, eventually degrading the viral DNA and thus preventing the bacteria from viral infection.
In 2011, Jennifer Doudna, Emmanuel Charpentier, and their team figured out that they can harness this system to cut, not just viral DNA, but any DNA at a specific location by changing the guide RNA to match the target of interest. The simplified engineered CRISPR-Cas9 system was developed by linking the two RNAs crRNA and tracrRNA to form a single guide RNA (sgRNA) or simply called guide RNA. The guide RNA consists of a 20 letter guide sequence that is complementary to a small sequence of target DNA of the host that is required to be edited.
After the guide RNA binds at the desired position in the genome, the Cas9 enzyme introduces double-stranded breaks in the desired DNA. Once the DNA is cut, researchers could use the cell’s own DNA repair machinery to add or delete pieces of genetic material, to make changes to the target DNA.
Cas 9 is referred to as a “programmable endonuclease” because one can program it to cut a specific DNA by providing a unique guide RNA. After the discovery, gene editing by the CRISPR-Cas9 system is being explored in the treatment of various diseases like sickle cell disease, hemophilia, cystic fibrosis, cancer, heart disease, HIV infection, etc.
Cas12/Cas13 enzymes for diagnostic purposes
After establishing the importance of the CRISPR-Cas9 system in gene editing, the scientists are harnessing this technology for diagnosing infectious diseases. A team of scientists found that two cousins of Cas9 called Cas12 and Cas13 can be harnessed to actually detect human disease. While Cas12 targets DNA, Cas13 targets the RNA. Part of the mechanism of their action is similar to that of Cas9. A guide RNA that is complementary to the target sequence is required for specific binding of Cas enzymes and then the Cas12 or Cas13 proteins cleave at the target site. An additional interesting feature of Cas12 and Cas13 enzymes is that they show “trans” or “collateral” cutting activity. It means that on finding the target, the cleavage activity of Cas12 and Cas13 enzymes is not just restricted to the target DNA or RNA, but they can also cut any single-stranded non-targeted nucleic acid molecules in the vicinity.
Although this collateral DNAse and RNase activity might appear to be a disadvantage in terms of specific gene editing, it has made these enzymes a powerful tool for the development of CRISPR-based diagnostics.
The collateral cutting activity of these enzymes has attracted scientists to build reporter systems in CRISPR diagnostics.
Sherlock CRISPR-Cas13 based diagnostic kit
Sherlock Biosciences with its Sherlock CRISPR diagnostic kit is the first company that received the approval for CRISPR- based COVID-19 diagnostic test. SHERLOCK is an acronym for “Specific High-sensitivity Enzymatic Reporter unLOCKing. This kit uses the Cas13 enzyme, which targets the RNA. For the diagnosis, a guide RNA (gRNA) is designed that recognizes a specific RNA sequence found in the SARS-CoV-2 genome. Cas13 and the gRNA then forms a complex and searches for the sequence match in the RNA of the SARS-CoV-2. When the gRNA binds to the programmed sequence of the target RNA, the Cas13 enzyme cuts the target SARS-CoV-2 RNA. After cleaving the target RNA, Cas13 doesn’t get inactivated rather it gets further activated to cut the surrounding unrelated single-stranded RNA reporter molecules that may be nearby in the reaction solution. In the test, the cleavage of these single-stranded RNA reporter molecules as a result of the collateral activity of the Cas13 enzyme is determined.
Let’s study in detail how the CRISPR diagnostic kit “SHERLOCK” works for the detection of COVID-19
Diagnosing COVID-19 requires taking a sample from the patient. Sample taken is usually a nasopharyngeal or oral pharyngeal swab. In order to execute CRISPR-based diagnosis, the viral RNA is extracted from the patient’s sample. The extracted RNA may contain other viral, bacterial, or patient’s own RNA as well. To increase the sensitivity of the test, the SARS-CoV-2 RNA is amplified with the reverse transcription- recombinase polymerase amplification process, abbreviated as RT-RPA.
In the first step, the reverse transcriptase enzyme converts the SARS-CoV-2 RNA into complementary DNA, simply referred to as cDNA. This cDNA is then amplified using the RPA process. Let’s understand how the RPA process is done and its advantages over traditional PCR.
Recombinase Polymerase Amplification (RPA)
The recombinase polymerase amplification is an isothermal nucleic acid amplification. Unlike polymerase chain reaction PCR, the RPA reaction occurs at a single temperature, so there’s no need for a thermocycler. This makes the RPA an excellent candidate for developing low-cost, rapid, point-of-care diagnosis. And is ideally suited to fields and other settings with minimal resources for diagnosing infectious diseases, food contaminations, etc. RPA is as specific as PCR amplification but is much, much faster. Results are typically generated within 3–10 minutes.
The RPA process occurs by 3 enzymes: recombinase, single-stranded DNA binding proteins (SSB), and strand displacing polymerase.
- In the case of traditional RCR, the denaturation step in which double-stranded template DNA is separated into two single strands is performed at 94°C. But in the case of RPA, this step is substituted by the 2 enzymes: recombinase and single-stranded DNA binding protein. The recombinase enzymes form complexes with the oligonucleotide primers, then this complex scans the ds viral cDNA target, searching for the homologous sequences. Once the homologous sequences are found, the recombinase primers complex invades the dsDNA, causing the separation of DNA strands.
- When the primers are paired to their complementary sequences, the single-stranded DNA binding proteins bind to the exposed DNA strand in order to stabilize it. The local separation of these DNA strands forms a D-loop-like structure.
- Finally, the strand displacing DNA polymerase enzyme extends the primer, eventually generating the amplicons from the original strands of template DNA. These newly generated DNA strands are then used for another round of RPA for exponential viral cDNA amplification.
Typically, the RPA reactions are executed at a single temperature ranging from 37°C to 42°C. At optimal temperature, the reaction progresses rapidly and results in specific DNA amplification from just a few target copies to detectable levels, typically within 3–10 minutes. No other sample manipulation such as thermal or chemical melting is required to initiate amplification.
Cas13 collateral activity
1. After the amplification of SARS-CoV-2 cDNA is done, it is then transcribed into RNA because the kit uses detection by the Cas13 enzyme which targets the RNA.
2. This amplified RNA is then mixed with the Cas13 enzyme, guide RNA, and single-stranded RNA reporter molecules in a reaction tube.
The guide RNA matches a specific RNA sequence found in the SARS-CoV-2 genome. On the other hand, each of the single-stranded RNA reporter molecules is coupled with the fluorescent FAM molecule at one end and a biotin molecule at the other end. When the reporter molecules are intact, biotin acts as a quencher which suppresses the fluorescence emitted by the FAM molecule.
3. On the other hand, if the patient sample contains the SARS-CoV-2 virus, then the Cas13 cleaves the viral RNA and also shows collateral activity. Because of collateral activity, Cas13 cleaves the surrounding FAM-biotinylated ssRNA reporter molecules, leading to the separation of the FAM molecules from the biotin molecules. The freed fluorescent FAM molecules when excited with light, produce a quantifiable fluorescence that can be detected by a detector, indicating the presence of SARS-CoV-2 RNA in the sample.
Lateral Flow Assay
Visualization of Cas13a detection can also be achieved using lateral flow strips that are designed to capture labeled nucleic acids. The interpretation of lateral flow strips is very intuitive and easy, similar to a pregnancy test the test changes color if the virus is detected.
Typically, lateral flow test strips are composed of a sample pad, a conjugate pad, a testing area, and an absorbent pad.
- The sample pad is where the test strip receives samples in the form of liquid drops.
- The absorbent pad generates a suction force, pulling the sample from the sample pad, towards the conjugate pad and then to the test area.
- The conjugate pad is covered with a large number of colloidal gold particles, conjugated with anti-FAM antibodies.
- The testing area is marked with the letters T and C. T stands for test area and C stands for control area. The generation of colored lines on T or C areas indicates whether the result is positive or negative. In the control area, the streptavidin is immobilized and in the test area, the antibodies specific for the anti-FAM antibodies are immobilized.
Take the scenario where the sample is negative which means SARS-CoV-2 is not present in the patient’s sample. In this case, the Cas13 enzyme remains inactive and thus no collateral cleavage of the FAM-biotinylated ssRNA reporter molecules. When the sample containing the intact reporter molecule is loaded on the sample pad at one end of the strip, it flows to the conjugate pad. The conjugate pad is covered with a large number of colloidal gold particles, conjugated with anti-FAM antibodies. Thus, on reaching the conjugate pad, the intact reporter molecules by their FAM labels get attached to the anti-FAM antibodies conjugated to the gold particles.
The colloidal gold molecules are then pulled into the testing area by the suction force generated by the absorbent pad. When the gold particles reach the control(C) area, the attached intact reporter molecules accumulate at the C area by the interaction of the biotin label with immobilized streptavidin. The interaction leads to the appearance of a visible colored line in the C area. The bound gold particles are unable to move further into the test area, thus a single-colored line appears in the C area on the strip, indicating the test is negative.
On the other hand, if the patient sample contains SARS-CoV-2, then the Cas13 cleaves the viral RNA and also shows collateral activity. Because of collateral activity, it cleaves the FAM-biotinylated ssRNA reporter molecules, leading to the separation of biotin molecules and FAM molecules. Therefore, when the sample containing the cleaved reporter molecules is loaded on the sample pad at one end of the strip, it flows to the conjugate pad, where the separated FAM labels get attached to the anti-FAM antibodies conjugated to the gold particles.
The bound colloidal gold molecules and unbound biotin label molecules are then pulled into the test area by the suction force generated by the absorbent pad. When they reach the C area, the biotin labels bind to the C area by the interaction with the immobilized streptavidin. The interaction leads to colored line formation in the C area. On the other hand, the FAM label bound colloidal particles are not captured on the C area, and therefore they continue to move to the next location, which is the test area of the lateral flow strip containing the anti-rabbit antibody, which binds anti-FAM antibodies. This generates a second visible colored line in the test area.
Thus the appearance of 2 lines at the T and C area indicate the positive test, in other words, the patient is infected with the SARS-CoV-2 virus. On the other hand, a negative sample shows only a single colored line at the C area of the lateral flow strip.
The diagnostic test produces results within an hour.
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