CRISPR-based diagnosis of infectious diseases (Part 56- CRISPR in Gene Editing and Beyond)
Welcome to the 56th part of the multi-part series on applications of CRISPR in gene editing and beyond.
The CRISPR-Cas system has recently emerged as a significant tool in the field of infectious disease diagnosis. With the emergence and reemergence of new viral diseases compromising healthcare worldwide, there is a critical need for reliable and rapid point-of-care (POC) testing kits to diagnose infected patients. Unlike RT-PCR, which necessitates a specialized central laboratory infrastructure, POC tests can be performed at or near the point of care, such as clinics and hospitals. This eliminates the need for sample transportation and allows communities without specialized laboratory infrastructure to detect infected patients quickly. This can help prevent delays in diagnosis and enable timely intervention, particularly in regions where laboratory facilities or highly trained personnel are not readily available. Several respiratory viruses, including SARS CoV-1, MERS, and SARS CoV-2 (COVID-19), as well as other viruses like Influenza A H1N1 (swine flu), Ebola, and Zika, have caused major viral outbreaks globally, emphasizing the need for reliable and rapid POC testing kits. The CRISPR-Cas system has shown promise in developing such POC kits, making it a valuable tool in the fight against infectious diseases and a crucial asset in the field of healthcare.
Cas enzymes for diagnosing infectious diseases
A team of scientists found that two cousins of Cas9, called, Cas12 and Cas13, can be harnessed to detect human disease. While Cas12 targets DNA, Cas13 targets 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 (Fig 1). 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; 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 developing CRISPR-based diagnostics. The collateral cutting activity of these enzymes has attracted scientists to build reporter systems in CRISPR diagnostics.
Before the COVID-19 pandemic, the field of CRISPR diagnostics was in its infancy. However, with the outbreak of the pandemic and the widespread impact of the virus, there has been a significant surge in this area of research to develop rapid and accurate diagnostic tools for detecting SARS-CoV-2, the virus that causes COVID-19.
1. Sherlock (CRISPR-Cas13 based) diagnostic test
On 6th May 2020, Massachusetts-based biotech firm Sherlock Biosciences with its “Sherlock CRISPR diagnostic kit,” became the first company that received FDA approval for the 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 (Li et al., 2018). Cas13 and the gRNA then form a complex and search 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; instead, HEPN motifs of Cas13 get 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 resulting from the collateral activity of the Cas13 enzyme is determined (Kellner et al., 2019).
Mechanism of action
Diagnosing COVID-19 requires taking a sample from the patient. The sample taken is usually a nasopharyngeal or oral pharyngeal swab. 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. Thus, to increase the test’s sensitivity, the SARS-CoV-2 RNA is amplified with the Reverse Transcription- Recombinase Polymerase Amplification process, abbreviated as RT-RPA (Zhang et al., 2020; Joung et al., 2020).
In the first step, the reverse transcriptase enzyme converts the SARS-CoV-2 RNA into complementary DNA, referred to as cDNA. The cDNA is then amplified by the RPA process using primers specific to the SARS-CoV-2 genes, for example, S gene, Orf1ab gene, N gene, etc (Fig 2).
Recombinase Polymerase Amplification (RPA) process: 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, making the RPA an excellent candidate for developing low-cost, rapid, point-of-care diagnosis (POC). And is ideally suited to fields and other settings with minimal resources for diagnosing infectious diseases like COVID-19, 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 three enzymes: recombinase, single-stranded DNA binding proteins (SSB), and strand displacing polymerase (Fig 3).
(i) In traditional PCR, the denaturation step in which double-stranded template DNA is separated into two single strands is performed at 94°C. But in RPA, this step is substituted by the two enzymes: recombinase and single-stranded DNA binding proteins. The recombinase enzymes form complexes with the oligonucleotide primers (Fig 3a), then these complexes scan the ds viral cDNA target, searching for the homologous sequences. And once the homologous sequences are found, the recombinase primer complexes invade the dsDNA, causing the separation of DNA strands (Fig 3b).
(ii) When the primers are paired to their complementary sequences, the single-stranded DNA binding proteins bind to the exposed DNA strand to stabilize it. The local separation of these DNA strands forms a D-loop structure (Fig 3c).
(iii) Finally, the strand displacing DNA polymerase enzyme extends the primer, eventually generating the amplicons from the original strands of template DNA (Fig 3d). These newly generated DNA strands are then used for another round of RPA for exponential viral cDNA amplification (Fig 3e).
Typically, the RPA reactions are executed at a single temperature ranging from 37°C-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 activity: After the amplification of SARS-CoV-2 cDNA is done, it is then transcribed into RNA because the kit uses detection by Cas-13 enzyme, which targets the RNA. 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.
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 (Fig 4).
Imagine, if the patient sample contains the SARS-CoV-2 virus, then the Cas13 cleaves the viral RNA and 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 (Fig 5a). When excited with light, the freed fluorescent FAM molecules produce a quantifiable fluorescence that can be detected by a detector, indicating the presence of SARS-CoV-2 RNA in the sample (Fig 5b).
Lateral Flow Assay: Visualization of Cas13 activity 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 strip changes color if the virus is detected (Fig 6).
Typically, lateral flow test strips are composed of a sample pad, a conjugate pad, a testing area, and an absorbent pad (Fig 7).
· 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 the test area, and C stands for the 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.
Negative Test: 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 occurs. 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 particles are then pulled into the control (C) area by the suction force generated by the absorbent pad. When the gold particles reach the C area, the attached intact reporter molecules accumulate at the C area by the interaction of the biotin labels with immobilized streptavidin. The interaction leads to the appearance of a visible colored line in the C area (Fig 8). 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 that the test is negative.
Note: This process does not involve the use of dyes or enzymes to generate the visible signal. Instead, the gold particles themselves create a colored line on the strip because they absorb and scatter light. When many of these tiny gold particles gather in one spot, their colors combine, making a visible line that’s easy to see.
Positive Test: On the other hand, if the patient sample contains SARS-CoV-2, then the Cas13 cleaves the viral RNA and shows collateral activity. Because of the collateral activity, Cas13 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 particles and unbound biotin label molecules are then pulled into the control 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 interacting with the immobilized streptavidin (Fig 9). On the other hand, the FAM label bound gold particles are not captured in the C area, but their presence creates a visible colored line in the C area, indicating that the test has worked correctly.
Then, they continue to move to the next location, the test (T) area of the lateral flow strip containing the anti-rabbit antibody, which binds anti-FAM antibodies. This results in the accumulation of gold particles and the appearance of a second visible colored line in the T area, confirming the positive test result.
Thus, the appearance of 2 lines in the Test (T) and Control © area indicates the positive test; in other words, the patient is infected with the SARS-CoV-2. On the other hand, the appearance of a single colored line in the C area of the lateral flow strip indicates the negative test; in other words, the patient is not infected with SARS-CoV-2.
The Sherlock CRISPR-Cas13 based diagnostic test produces results within an hour.
2. DETECTR (CRISPR-Cas12 based) diagnostic test
Another CRISPR-based diagnostic kit is “DETECTR,” developed by Mammoth Biosciences. DETECTR is an acronym for DNA Endonuclease Targeted CRISPR Trans Reporter. This kit works similarly to SHERLOCK. The difference is in the types of Cas enzymes used. While SHERLOCK uses the Cas13 enzyme, which targets RNA, DETECTR uses the Cas12a enzyme (Chen et al., 2018), which targets DNA. For the diagnosis, a guide RNA (gRNA) is designed that recognizes a specific DNA sequence, for example, one of the genes found in the SARS-CoV-2 genome.
Like the Cas13 enzyme, Cas12a also possesses collateral activity, which means the Cas12a enzyme first cuts the specific target DNA. The enzyme is then further activated to chop the surrounding unrelated single-stranded DNA molecules regardless of their complementarity with these ssDNA molecules. In other words, Cas12a acts like a paper shredder that shreds the surrounding DNA irrespective of the complementarity (Broughton et al., 2020a; Broughton et al., 2020b; Mustafa and Makhawi, 2020).
Loop Mediated Isothermal Amplification (LAMP): Viral RNA is first extracted from the patient’s sample to perform the CRISPR-based diagnosis. This extracted RNA might include other viral, bacterial, or the patient’s own RNA. In order to enhance the test’s sensitivity, the SARS-CoV-2 RNA is amplified with the Reverse Transcription Loop-Mediated Amplification process, abbreviated as RT-LAMP.
In the first step, the reverse transcriptase enzyme converts the SARS-CoV-2 RNA into complementary DNA (cDNA). The LAMP process then amplifies the cDNA. The reaction is performed at 62 °C for 20–30 min. Like RPA, LAMP is also an isothermal DNA amplification method. It also occurs at a single temperature, so there’s no need for a thermocycler.
For performing the LAMP process, four different primers are used that can specifically recognize six distinct regions of the target DNA i.e., SARS-CoV-2 cDNA. In the Fig 10, suppose two lines represent the target DNA. The six regions on template strand 3’-5’ that are required to be recognized by primers are represented as F3c, F2c, F1c, B1, B2, and B3. Therefore, on the complementary coding strand 5’-3’, these regions are represented as F3, F2, F1, B1c, B2c, and B3c.
Primers required for the amplification process:
- The first primer is the forward inner primer designated as FIP. It consists of an F2 region at the 3' end and an F1c region at the 5'end. The F2 region of the primer is complementary to the template sequence’s F2c region. And the F1c region is identical to the F1c region of the template sequence (Fig 11).
2. The second primer is the forward outer primer, also called the F3 primer, because it consists of only one region, i.e., an F3 region complementary to the F3c region of the template sequence (Fig 12).
3. The third primer is the backward inner primer designated as BIP. It consists of a B2 region at the 3' end and a B1c region at the 5' end. The B2 region is complementary to the template sequence’s B2c region. And the B1c region is identical to the B1c region of the template sequence (Fig 13).
4. The fourth primer is the backward outer primer, also called as B3 primer. It consists of a B3 region, which is complementary to the B3c region of the template sequence (Fig 14).
DNA Amplification
The amplification reaction is initiated from the strand invasion by the forward inner primer (FIP).
- F2 region of FIP hybridizes to the F2c region of the target DNA, and then the strand displacing DNA polymerase extends the primer leading to the synthesis of the complementary strand. This leads to the separation of ds target DNA (Fig 15).
2. To this first product, outer primer F3 binds to the F3c region of the target DNA. Then strand displacing DNA polymerase extends the F3 primer, which leads to displacing the FIP linked complementary strand. As the strand is displaced, it forms a self-hybridizing loop at the 5' end due to the inclusion of a reverse complementary sequence F1c in the primer sequence (Fig 15).
3. This displaced single-stranded DNA with a loop at the 5' end now serves as a template for backward inner primer (BIP) that consists of a B2 region at the 3' end and a B1c region at the 5' end. B2 region of the primer hybridizes to the B2c region of the template DNA. DNA synthesis is then initiated, leading to the creation of a complementary strand and opening of the 5' end loop (Fig 16).
4. Now, the outer backward primer B3 hybridizes to the B3c region of the target DNA and is extended by the DNA polymerase, displacing the backward inner primer-linked complementary strand (Fig 16).
As the backward inner primer (BIP)-linked strand is displaced, it results in the formation of a dumbbell-shaped DNA because of the presence of reverse complementary sequences B1c and F1. The dumbbell structure DNA then forms a seed for exponential LAMP amplification, which is the second stage of the LAMP reaction. This dumbbell structure contains multiple sites for the initiation of DNA synthesis (Fig 17).
(i) One is from the 3’ end of the F1 region. The nucleotides are added to the 3' end of the F1 region by DNA polymerase, extending and opening up the loop at the 5' end. The dumbbell-shaped DNA now gets converted to a stem-loop structure (Fig 18).
(ii) To initiate LAMP cycling, the forward inner primer (FIP) hybridizes to the loop of the stem-loop DNA structure. Strand synthesis is initiated here. As the strand synthesis occurs, the F1 strand is displaced and forms a new loop at the 3' end (Fig 19).
Now, nucleotides are added to the 3' end of B1, leading to a long concatemer formation. The extension leads to the displacement of the FIP linked strand (Fig 20).
This displaced strand again forms a dumbbell-shaped DNA. Now the dumbbell-shaped DNA and the concatemers formed have multiple sites for annealing of primers, leading to DNA synthesis initiation (Fig 21).
The final products obtained due to amplification are a mixture of stem-loop DNA with various stem lengths and different dumbbell structures with loops (Fig 22).
The LAMP reaction occurs in a single tube containing target DNA, buffer, primers and DNA polymerase enzyme. The tube is incubated at 64°C in a regular laboratory water bath or heat block that maintains a constant temperature.
Cas12 activity: For the detection of SARS-CoV-2 in the sample, the amplified cDNA is then mixed with the Cas12a enzyme, guide RNA, and single-stranded DNA reporter molecules in a reaction tube. The guide RNA matches a specific DNA sequence found in the SARS-CoV-2 genome, for example, one of the genes found in the SARS-CoV-2 genome. Each of the single-stranded DNA reporter molecules are 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. If the patient sample contains SARS-CoV-2, then the Cas12a cleaves the amplified viral cDNA and shows collateral activity. Because of the collateral activity, Cas12a cleaves the surrounding FAM-biotinylated ssDNA 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 which can be detected by a detector, indicating the presence of SARS-CoV-2 cDNA in the sample.
Detection by Lateral Flow Strip: Detection of Cas12a activity and the viral cDNA can also be done via a lateral flow strip, similar to the Sherlock test (Broughton et al., 2020c). Lateral flow test strips are composed of a sample pad, a conjugate pad, a testing area, and an absorbent pad. The testing area contains two types of lines or areas: The test area designated as T and the control area designated as C. In the control area, the streptavidin is immobilized, and in the test area, the antibodies specific for the anti-FAM antibodies are immobilized.
Negative Test: If the sample is negative, i.e., SARS-CoV-2 is not present in the patient’s sample, the Cas12a enzyme remains inactive, and thus, no collateral cleavage of the FAM-biotinylated ssDNA reporter molecules occurs. When the sample containing the intact reporter molecule is loaded on the sample pad of the strip, on reaching the conjugate pad, the intact reporter molecules get attached to the anti-FAM antibodies conjugated to the gold particles by their FAM labels. When the gold particles reach the C area, the attached intact reporter molecules accumulate at the C area by the interaction of biotin labels with immobilized streptavidin. The bound gold particles are unable to move further into the T area. Thus, a single-colored line appears in the C area on the strip, indicating that the test is negative (Fig 23).
Positive Test: On the other hand, if the patient sample contains SARS-CoV-2, then the Cas12a cleaves the viral cDNA and shows collateral activity; thus cleaves the FAM-biotinylated ssDNA 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, 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 C area. After reaching the C area, the biotin labels bind to the C area by interacting with the immobilized streptavidin. Whereas the FAM label bound gold particles are not captured in the C area, but their presence creates a visible colored line in the C area, indicating that the test has worked correctly.
Then, they continue to move to the T area of the lateral flow strip, which contains anti-rabbit antibodies that bind to anti-FAM antibodies. This interaction causes gold particles to accumulate, forming a second visible colored line in the T area (Fig 24).
Thus the appearance of 2 lines in the T and C area indicates a positive test, i.e., the patient is infected with SARS-CoV-2. On the other hand, the appearance of a single colored line in the C area of the lateral flow strip indicates a negative test, indicating the patient is not infected with SARS-CoV-2.
The DETECTR CRISPR-Cas12a-based diagnostic test produces results within 30–40 minutes. The LAMP process takes 20–30 minutes, and the Cas detection reaction takes another 10 minutes; thus, one can get the result within 30–40 minutes.
3. FELUDA (CRISPR-Cas9 based) diagnostic test
The other type of CRISPR-based diagnostic test is FELUDA, made by Indian researchers at CSIR Institute of Genomics and Integrative Biology (CSIR-IGIB). FELUDA is an acronym for FnCas9 Editor Linked Uniform Detection Assay. FELUDA test is different from other CRISPR based tests SHERLOCK and DETECTR in the types of Cas enzymes used. SHERLOCK and DETECTR use the Cas13 and Cas12a enzymes, which show collateral activity. On the other hand, the FELUDA test uses specially adapted Cas9 protein, derived from Francisella novicida bacteria, and it does not show collateral activity like Cas12 and Cas13 enzymes (Acharya et al., 2019). It means Cas9 is incapable of cleaving the surrounding ssFAM-biotinylated reporter molecules like Cas12 and Cas13.
In the FELUDA test, because of the inability to cleave the surrounding labeled reporter molecules, the Cas9 activity readout on the test strip can be done by labeling the target DNA with biotin molecule and the guide RNA with FAM molecule (Azhar et al., 2020a; Azhar et al., 2020b; Suvvari et al., 2020).
Cas9 activity: First, the patient sample is collected, and then the RNA is extracted from the sample. This is followed by reverse transcription wherein SARS-CoV-2 RNA is converted into cDNA, which is further amplified by the PCR or RPA. The amplification is performed using a normal forward primer and a reverse biotinylated primer. The biotinylated reverse primers label the amplified SARS-CoV-2 cDNA at the 5’ end (Fig 25).
On the other hand, the guide RNA is labeled with FAM molecules (Fig 26). In the next step, the biotin-labeled amplified cDNA molecules are then mixed with Cas9 protein and FAM-labeled guide RNA. The Cas9 and FAM-labeled gRNA forms a search complex, which searches for the sequence match in the amplified cDNA of SARS-CoV-2. After the binding of gRNA, Cas9 protein introduces double-stranded breaks in the SARS-CoV-2 cDNA.
Detection by Lateral Flow Strip:
Detection of Cas9 activity is also done via a lateral flow paper strip, similar to SHERLOCK and DETECTR assays. In the paper strip used for the FELUDA test, positions of the test and control lines are swapped compared to paper strips used for DETECTR and SHERLOCK (Fig 27a). It leads to a completely different assessment of the resulting signals. On the test line, streptavidin is immobilized, and on the control line, antibodies specific for anti-FAM antibodies are immobilized (Fig 27b).
Negative Test: Take the scenario where the patient’s sample does not contain SARS-CoV-2. In this case, FAM-labeled gRNA cannot find the matching sequence; thus, the Cas9 enzyme remains inactive. It also means that the biotin label is not present in the sample. When the sample that doesn’t contain SARS-CoV-2 is loaded on the sample pad, it flows to the conjugate pad which is covered with a large number of colloidal gold particles conjugated with anti-FAM antibodies. Thus, on reaching the conjugate pad, the search complex by the FAM labels (gRNA is FAM-labeled) gets attached to the anti-FAM antibodies conjugated to the gold particles.
The FAM-labeled search complex bound colloidal gold molecules are then pulled into the test area by the suction force generated by the absorbent pad. When the gold particles reach the test area, they are unable to bind to immobilized streptavidin. Thus, they are pulled further into the control area, where anti-FAM antibodies are immobilized. On reaching the C area, the FAM-labeled search complex bound gold molecules accumulate, thereby interacting with anti-FAM antibodies, leading to the appearance of a visible colored line in the control area. It indicates that the patient sample does not contain SARS-CoV-2; in other words, the test is negative (Fig 28).
Positive Test: On the other hand, if the patient’s sample contains SARS-CoV-2, then FAM-labeled gRNA can find the matching sequence; thus, the Cas9 enzyme gets activated and cleaves the biotinylated SARS-CoV-2 cDNA. It also means that both the FAM and biotin labels are present in the sample. When the sample containing cleaved SARS-CoV-2 cDNA and FAM-labeled search complex is loaded on the sample pad at one end of the strip, it flows to the conjugate pad containing colloidal gold particles, conjugated with anti-FAM antibodies. Thus, on reaching the conjugate pad, the FAM-labeled search complex gets attached to the anti-FAM antibodies conjugated to the gold particles.
The FAM-labeled search complex bound gold molecules, and cleaved biotinylated cDNA are then pulled into the test area by the suction force generated by the absorbent pad. When they reach the test area, the cleaved biotinylated cDNA accumulates at the test area because of the interaction of the biotin labels with immobilized streptavidin. Some of the FAM-labeled search complexes recognize the cleaved cDNA and bind to it through complementary sequence interactions. This binding leads to the accumulation of gold nanoparticles at the T area, resulting in the appearance of a colored line (Fig 29).
On the other hand, the FAM-labeled search complex gold particles that were not captured at the T area move to the control area of the lateral flow strip. On reaching the C area, these complexes encounter immobilized antibodies that specifically recognize and bind to anti-FAM antibodies. The interaction leads to the accumulation of gold nanoparticles at the C area, producing a second visible colored line. It indicates that the patient sample contains SARS-CoV-2; in other words, the test is positive.
Thus, the appearance of 2 lines in the T and C area indicate the positive test; in other words, the patient is infected with the SARS-CoV-2. On the other hand, the appearance of a single-colored line in the C area of the lateral flow strip indicates the negative test; in other words, the patient is not infected with SARS-CoV-2.
This diagnostic test kit can deliver test results in 40–45 minutes.
CRISPR-based diagnostic tests meet high-quality benchmarks with 96% sensitivity and 98% specificity for detecting the SARS-CoV-2. Sensitivity can be defined as the ability of a test to correctly identify individuals with the disease, while specificity is the assay’s ability to accurately identify those without the disease.
Additionally, the CRISPR-based diagnostic tests do not require RT-PCR-based confirmation for samples that are confirmed as positives or negatives by the CRISPR SARS-CoV-2 test.
Pros and Cons of CRISPR technology in diagnostics
The CRISPR-based diagnostic assay is as accurate, specific, and sensitive as real-time RT–PCR but is a more rapid approach for Point of Care diagnosis. Like RT-PCR, the requirement of personal protective equipment, extraction kits, reagents, sample collection, and RNA extraction also exists for the CRISPR-based diagnosis. However, the CRISPR-based approach has some more advantages over real-time RT–PCR.
Pros of CRISPR-based diagnosis
(i) Point of care applications: Besides their accuracy is on par with RT-PCR, CRISPR-based diagnostic tests can perform the test at one temperature using a type of isothermal nucleic acid amplification process called LAMP (loop-mediated isothermal amplification) or RPA (Recombinase Polymerase Amplification) methods. For performing amplification, CRISPR-based diagnostic tests do not require a thermocycler (PCR machine). The reaction can occur in a single tube containing buffer, target DNA, DNA polymerase, and primers. The tube is then incubated at the required temperature in a regular laboratory water bath or heat block to maintain a constant temperature. Because of this, the CRISPR based tests can be used for point of care (POC).
On the contrary, RT-PCR uses thermocyclers, expensive laboratory equipment, and well-trained personnel, which are hard to adapt for point of care (POC).
Point of care diagnostic tests are extremely important for diagnosing diseases in countries or settings where laboratory facilities or extremely trained personnel are unavailable. CRISPR-based tests can be just performed in a test tube, and then the results can be interpreted on the paper strip, similar to pregnancy tests. Because of the ease of performing the test and simple interpretation of results, CRISPR-based assay can be used for diagnosis at airports, ports, clinics, schools, etc., for better disease diagnosis, monitoring, management, and containment of diseases like COVID-19.
(ii) Speed: Another critical benefit of CRISPR-based diagnostics is the speed at which it generates test results compared to RT-PCR. The CRISPR-based diagnostic test provides results within 45 minutes compared to RT-PCR, which provides results within 3–4 hours. Laboratory-based RT-PCR testing takes around 3–4 hours, but when transportation of clinical samples is required, the diagnosis time often exceeds 24–48 hours, resulting in delays in diagnosis. But the CRISPR based diagnostic kits can diagnose the samples even in minimal settings; the test can be performed at the hospital or clinic, where the sample is collected. Thus transportation of samples to specific laboratories is not required, because of which delay in diagnosis can be avoided. Therefore, it can help healthcare professionals to decide treatment or course of action for a patient as soon as the patient is diagnosed.
Additionally, with an efficiency equivalent to RT-PCR, CRISPR-based diagnostics can run more tests in roughly the same amount of time.
(iii) Cost-effective: The third benefit of CRISPR-based diagnostic tests is that they provide a more cost-effective approach to diagnosis than the RT-PCR technique.
Cons of CRISPR-based diagnosis
(i) Off-target effects: However, one of the main disadvantages of the CRISPR-Cas system is the non-specific binding of the guide RNA to the target genome, which in turn, can lead to poor signaling and misinterpretation of the results.
Therefore, to reduce the off-target effects of CRISPR-Cas complex
· Guide RNAs must be carefully designed using specialized bioinformatics tools to select the best ones.
· Or two guide RNAs can be used instead of 1 guide RNA to detect the viral genome.
(ii) Separate nucleic acid-preamplification steps: SHERLOCK, DETECTR, and FELUDA tests require separate nucleic acid pre-amplification steps by RPA or LAMP processes and several manual handling steps, which undoubtedly complicates the testing procedures and potentially increases the risk of carry-over contaminations due to amplification products transferring.
Therefore, to minimize these limitations, the researchers have developed more efficient POC CRISPR-based diagnostic tests like:
4. All-in-one dual CRISPR-Cas12a assay
CRISPR-based nucleic acid detection methods like SHERLOCK and DETECTR typically require separate nucleic acid pre-amplification by RPA or LAMP processes and multiple manual operations. To overcome these limitations, a team of researchers led by associate professor Changchun Liu, University of Connecticut, developed an all-in-one dual CRISPR-Cas12a (AIOD-CRISPR) assay, in which two crRNAs are used instead of 1 crRNA for the detection of SARS-CoV-2. The two individual crRNAs are without PAM sequence limitation and are designed to bind two different sites that are close to the recognition sites of primers in the target sequence (Ding et al., 2020a; Ding et al., 2020b; Ding et al., 2020c).
Suppose two lines represent target DNA, and the region that is to be recognized by the first Cas12a-crRNA complex is represented as T1 on the 5’-3’ strand. Thus on the complementary strand 3’-5’, this region is represented as T1c. On the other hand, the region that is to be recognized by the second Cas12a-crRNA is represented as T2 on the 3’-5’ strand. Thus, on the complementary strand 5’-3’, this region is represented as T2c (Fig 30).
As already discussed, in the case of RPA reactions, denaturation of the target DNA is performed by the two enzymes: recombinase and single-stranded DNA binding protein. The recombinase enzymes form complexes with the forward (F) and reverse (R) oligonucleotide primers, then these complexes scan the ds viral cDNA target, searching for the homologous sequences. The region to which the forward primer-recombinase enzyme complex binds is represented as Fc on the 3’-5’ strand. Therefore, on the complementary strand 5’-3’, this region is labeled as F. Likewise, the region to which the reverse primer-recombinase enzyme complex pairs is represented as Rc on the 5’-3’ strand. Therefore on the complementary strand 3’-5’, this region is labeled as R.
To initiate the RPA reaction, the two Cas12a-crRNA complexes are then mixed with RPA primers, FAM-biotinylated ssDNA reporter molecules, RPA enzymes (recombinase, single-stranded DNA binding proteins, strand-displacement DNA polymerase), and the viral cDNA in a single reaction tube (Ding et al., 2020c).
(i) When the reaction tube is incubated at 37 °C, the RPA amplification is initiated. The recombinase enzymes form complexes with the forward (F) and reverse (R) primers; then, these complexes scan the ds viral cDNA target. Once the homologous sequences are found, the recombinase primer complex invades the dsDNA, causing the separation of DNA strands (Fig 31a).
(ii) When the primers are paired to their complementary sequences, the single-stranded DNA binding proteins bind to the exposed DNA strand to stabilize it (Fig 31b). The local separation of these DNA strands forms a D-loop like structure, which exposes the binding sites for the attachment of Cas12a-crRNA complexes (Fig 31c).
(iii) When the Cas12a-crRNA complexes bind the target sites, the Cas12a endonuclease is activated and cleaves the FAM-biotinylated ssDNA reporter molecules, generating strong fluorescence signals (Fig 32).
(iv) When strand displacing DNA polymerase enzyme extends the primer, it eventually generates the amplicons from the original strands of template DNA.
(v) The newly generated DNA strands are then used for another round of RPA and the binding of Cas12a-guide RNA complexes leading to the activation of Cas12a endonuclease. Activated Cas12a enzyme then again shows collateral activity, causing the cleavage of the surrounding FAM-biotinylated ssDNA reporter molecules, leading to the separation of the fluorescent FAM molecules from the biotin molecules. This same process is repeated continuously on amplified products of RPA. This gives leverage to amplify the fluorescence signal. This process is possible in a single tube that eliminates the need for separate preamplification and amplified product transferring steps.
(vi) After 40 minutes, the fluorescence signal can be detected by keeping the tube under a blue LED or UV light illuminator. The presence of fluorescence signals indicates the patient is infected with SARS-CoV-2. Because of the simplicity and accuracy, the AIOD-CRISPR assay can be used as POC molecular diagnostic technology for the rapid detection of COVID-19 in small clinics, schools, and other settings with minimal resources.
5. Electric field-driven microfluidics for CRISPR-based diagnostics
The CRISPR-based diagnostic tests require a considerable volume of reagents and certain manual steps like viral RNA extraction and nucleic acid amplification. Such reagents many times become scarce during the pandemic because of supply chain issues. Keeping this in mind, scientists at Stanford developed a CRISPR- Cas12 assay using electric field and microfluidics technologies to extract nucleic acids, thus, reducing time, manual steps, and reagents for the assay (Ramachandran et al., 2020).
The assay involves three main steps.
(i) First, the human sample is placed on the microfluidic chip, and SARS-CoV-2 RNA is purified from the patient’s sample with the help of a technique called isotachophoresis (ITP). Isotachophoresis is a form of electrophoresis that separates and concentrates charged molecules in solution, solely based on their electrophoretic mobility or the speed at which the molecules move within the applied electric field. Electric fields are applied across the sample to selectively extract and then concentrate viral RNA (Fig 33).
(ii) Next, the extracted RNA is manually pipetted off the chip and put into a standard test tube. The test tube contains the reverse transcriptase enzyme that converts RNA to cDNA, primers, and the DNA polymerase enzyme that amplifies it through the isothermal amplification process (LAMP or RPA).
(iii) Finally, the amplified cDNA is pipetted back onto the microfluidic chip, where the CRISPR enzymatic assay is performed. In this step, electric fields are again applied to concentrate critical components, such as the viral cDNA, guide RNA-Cas12 complex, and ssDNA reporter molecules into a tiny well. This increases the chances that the guide RNA and Cas12 enzyme will interact with the target cDNA, speeding up the reaction. Activated Cas12 enzyme shows collateral activity, and because of collateral activity, it cleaves the FAM-biotinylated ssDNA reporter molecules, leading to the separation of fluorescent FAM molecules. The freed fluorescent FAM molecules, when excited with light, produce a quantifiable fluorescence which can be detected by a detector, indicating the presence of SARS-CoV-2 RNA in the sample.
Additionally, this method consumes a minimal volume of reagents (around 100-fold lower than other CRISPR based methods).
6. CRISPR-based diagnostic tests use smartphone cameras
Scientists at Gladstone Institutes and UC Berkeley have developed a CRISPR-based COVID-19 diagnostic test that can provide the test results with a smartphone’s help in 15–30 minutes. The test eliminates the need to convert RNA to DNA steps and then amplify the DNA segments, thus reducing the time needed to complete the analysis. The diagnostic test utilizes three guide RNAs CRISPR-Cas13 protein complexes, which bind and cleave viral RNA. CRISPR Cas13 proteins are “programmed” to recognize three different regions of SARS-CoV-2 viral RNA and then combined with reporter molecules that become fluorescent when cleaved. Cas13 proteins get activated upon binding of guide RNAs to SARS-CoV-2 RNA, and the activated Cas13 proteins then start to cleave the reporter molecules, leading to the separation of the fluorescent label from the quencher (Fig 34).
The mobile phone camera converted into a fluorescent microscope can detect the fluorescence emitted by the cleaved reporter molecules. The higher the fluorescence detected, the higher the number of virus particles in the sample, and vice versa.
Besides rapid testing, the high sensitivity of mobile phone cameras, along with the connectivity, GPS, and data-processing capabilities of the smartphone, has the potential to make the CRISPR-based test portable and an attractive tool for point-of-care disease (POC) diagnosis in low-resource regions.
Additionally, the test can detect tiny amounts of the virus, in under 30 minutes and much larger concentrations of the virus, such as those in highly contagious people, in under 5 minutes.
How is a smartphone converted into a fluorescent microscope?
Fluorescence microscope is a type of microscope that works on the principle of fluorescence. A substance is said to be fluorescent when it absorbs the energy of invisible shorter wavelength radiation (such as UV light) and emits longer wavelength radiation of visible light (such as green or red light). High-energy and short-wavelength light (such as UV or blue light) is generated from a light source and passed through an excitation filter (Fig 35). This filter allows only specific short-wavelength light to pass through and removes non-specific wavelengths. The filtered light is reflected by a dichroic filter and illuminates the fluorophore-labeled sample. The fluorophore absorbs the short-wavelength light and emits longer-wavelength light, which passes through an emission filter. The emission filter blocks any residual excitation light and allows the desired longer-wavelength emission to be detected, resulting in glowing images of the fluorophore-labeled molecules against a dark background.
The smartphone is converted into a fluorescent microscope by clipping the lens onto the phone or tablet camera to enable magnification (Fig 36). The lens collects and focuses the fluorescent light emitted by the sample, which is then captured by the camera. The emission filter is placed over the lens to filter out unwanted wavelengths of light, allowing only the fluorescent light emitted by the specimen to pass through. This helps to enhance the contrast and clarity of the fluorescent image, making it easier to visualize and analyze.
Simple LEDs can be used to excite the fluorescent reporter molecules. The excitation light can scatter or reflect off the sample and create background noise, which can interfere with the accuracy and quality of the images obtained. Thus a simple plastic filter is used to reject the scattered excitation light, creating a dark field background that is necessary for viewing fluorescent images.
Moreover, the CRISPR-based diagnostic kits that we have discussed are not just for diagnosing SARS-CoV-2 but can also be used to detect other infectious diseases. All needed is to design guide RNA specific to the genome of the organism that is required to be detected.
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