Patent Application
20240043945
The present invention relates to the detection of nucleic acids using enzyme-assisted nanotechnology. More specifically, the present invention provides a molecular nanotechnology in the form of a transition-state DNA-enzyme molecular switch and methods of use that enables direct and sensitive detection of viral RNA targets in native clinical samples.
The rapid global spread of coronavirus disease 2019 (COVID-19) has stretched the limits of healthcare resources [Huang H, et al., ACS Nano. 2020, 14: 3747-3754]. Person-to-person transmissions from infected individuals with no or mild symptoms have been widely reported [Bai W, et al., JAMA. 2020, 323(14): 1406-1407; Rothe C et al., N Engl J Med. 2020, 382: 970-871]. Aggressive testing for SARS-CoV-2, the causal pathogen of COVID-19 [Gorbalenya A E et al., Nat Microbiol. 2020, 5: 536-544], is important in controlling the disease spread and devising safety measures. To date, quantitative reverse transcription polymerase chain reaction (RT-qPCR) remains the primary assay for detecting SARS-CoV-2 [Carter L J et al., ACS Cent Sci. 2020, 6: 591-605]. Albeit its sensitive performance, the technology requires extensive sample preparation (e.g., RNA extraction), exquisite primer design, specialized instrument and trained personnel [Centers for Disease Control and Prevention, 2020, fdadotgov/media/134922/download]. These limitations not only result in a long assay turnaround time, but also hinder its large-scale implementation and adaptation in a rapidly evolving pandemic. Indeed, these shortcomings are particularly apparent, when challenged under the severe pressure of COVID-19; a global shortage of reagents and the emergence of new mutations and false negatives pose critical challenges for RT-qPCR-based detection [Bruce E A, et al., bioRxiv. 2020 doi.org/10.1101/2020.03.20.001008; Li Y et al., J Med Virol. 2020, 92: 903-908; Toyoshima Y et al., J Hum Genet. 2020, 65(12): 1075-1082]. Accurate, rapid and easy-to-use molecular diagnostic tests for SARS-CoV-2 are crucially needed across the globe [Weissleder R et al., Sci Transl Med. 2020, 12, eabc1931].
Molecular nanotechnology offers unparalleled precision and programmability to construct a variety of self-assembled functional nanostructures [Li J et al., Nat Chem. 2017, 9: 1056-1067; Li Y et al., Nat Biotechnol. 2005, 23: 885-889; Jones M R et al., Science. 2005, 347: 1260901; Sundah N R et al., Nat Biomed Eng. 2019, 3: 684-694]. These nanostructures can be designed as versatile, multi-function machines, which can not only recognize external stimuli, but also respond and actuate various activities [Wilner 01 et al., Nat Nanotechnol. 2009, 4: 239-254; Song P et al., Nat Commun. 2020, 11: 838]. We have previously developed a molecular nanotechnology platform for rapid detection of nucleic acids [Ho N R Y et al., Nat Commun. 2018, 9: 3238]. Instead of relying on the traditional approach of target amplification (as in conventional RT-qPCR), the technology detects through target hybridization. It leverages enzyme-DNA hybrid nanocomplexes as molecular switches; upon the direct binding of specific nucleic acids (even RNA targets), the nanocomplexes dissociate to activate strong enzymatic activity. Importantly, the technology is highly programmable; new assays can be readily developed by modifying the highly configurable nanocomplexes, without needing complex design of PCR primers and dedicated fluorescent probes (e.g., Taqman probes). Due to this unique sensing mechanism and high programmability, we thus envision that the technology could enable direct detection of SARS-CoV-2, bypassing many steps and challenges of PCR detection (e.g., reverse transcription and thermal cycling). Nevertheless, given that a significant proportion of COVID-19 patients are reported to have a very low viral load [Pan Y et al., Lancet Infect Dis. 2020, 20: 411-412], our previously developed assay, with a limit of detection of ˜10 amol, would have a limited sensitivity to diagnose a broad spectrum of COVID-19 patients.
There is a need to bridge this gap in detection sensitivity.
Motivated by the multi-component nature of individual nanocomplexes, we reason that they can be tuned to establish highly responsive molecular switches. Specifically, the nanocomplex switches are self-assembled from multiple molecular constituents—Taq polymerase and distinct DNA strands—which exist in a dynamic equilibrium and exert different effects on overall switch characteristics. Through ratiometric tuning of these molecular constituents, we found that the most responsive state is a metastable state, where even trace amounts of target nucleic acids can readily activate the molecular switches to induce strong enzymatic activity. Leveraging molecular switches in this hyper-responsive state, which we call the transition state, we developed a highly sensitive and direct nucleic acid detection assay for SARS-CoV-2. The technology, termed catalytic amplification by transition-state molecular switch (CATCH), benefits from dual catalytic amplification: its transition-state molecular switches are readily activated upon the direct binding of even sparse amounts of viral RNA targets to liberate substantial enzymatic activity; this switch activation further recruits additional enzymatic cascades to transduce strong signal output.
Harnessing its hyper-responsiveness, CATCH achieves superior performance. It enables sensitive and specific detection of RNA targets, against a complex biological background, and reports a limit of detection (LOD) of ˜8 copies of target per μl, which is >10,000-fold more sensitive than our previous platform. The detection is also direct and rapid; the entire assay can be completed in <1 hour at room temperature and can be applied to a variety of sample types (e.g., purified RNA as well as complex clinical samples), bypassing all steps of conventional RT-qPCR (i.e., RNA extraction, reverse transcription and thermal cycling amplification). Importantly, CATCH enables versatile assay implementation. To support different diagnostic needs, the assay can be implemented in a 96-well format for high-throughput analysis and as a miniaturized microfluidic cartridge for portable smartphone-based measurement. When applied for clinical detection of SARS-CoV-2, CATCH demonstrated accurate and sensitive detection in both extracted RNA samples as well as inactivated patient swabs.
In a first aspect of the invention there is provided a method of detecting target polynucleotides in a sample, comprising the steps of:
In some embodiments the signalling nanostructure in d) comprises:
It would be understood that detection of target nucleic acid in the sample using signalling nanostructure i) is indicated by an increase in signal intensity, whereas the signalling nanostructure in ii) comprises a signal capacity that is reduced in the presence of activated DNA polymerase enzyme.
In some embodiments, the method further comprises;
In some embodiments, the DNA polymerase inhibitor conserved sequence region comprises the nucleic acid sequence set forth in SEQ ID NO: 14; 5′-CAATGTACAGTATTG-3′.
In some embodiments, the amount of inhibitory complex in the composition is in the range of 20 nM to 60 nM and/or the enhancer to inhibitor ratio in the composition is less than 1:1, preferably in the range of 0.3:1 to 0.6:1.
In some embodiments, the enhancer is at least one nucleotide longer than the inhibitor duplex region.
In some embodiments, the enhancer is about 35 to 45, preferably about 40, nucleotides in length.
In some embodiments, about half of the length of the enhancer oligonucleotide forms the inhibitor-enhancer duplex and about half forms an overhang segment.
In some embodiments, the self-priming portion of the signalling nanostructure comprises the nucleic acid sequence set forth in SEQ ID NO: 5: 5′-CGGCGTACGTAGAGCGTTGAGCAGGATGCCAACAGTCGATCAGGACGAGTGCTAACG CATTGTCGATAGCTCAGCTGTCTGAGCTATCGACAATGCGTT-3′.
In some embodiments, the dNTP label is biotin.
In some embodiments, the signalling dumbbell nanostructure comprises the nucleic acid sequence set forth in SEQ ID NO: 6: 5′-GTGCGTACATAGATCGTTATCTGTC TAACGATCTATGTACGCACTCACTCAGCTAACGCATTGTCGATAGCTCAGCTGTCTGAG CTATCGACAATGCGTT-3′.
In some embodiments, the signal development reagents comprise a fusion protein comprising avidin or a derivative thereof and an enzyme, selected from a group comprising but not limited to HRP, beta-lactamase, amylase, beta-galactosidase, and respective substrates selected from a group comprising but not limited to DAB, TMB, ABTS, ADHP, nitrocefin, luminol, starch and iodine, wherein signals can be measured and quantified as but not limited to colour, fluorescence, luminescence or electrochemical changes.
In some embodiments, the target is at least one nucleic acid associated with a non-human or human disease, genetic variants, forensic, strain identification, environmental and/or food contamination.
In some embodiments, the target is at least one pathogen polynucleotide.
In some embodiments, the target is a SARS-CoV-2 polynucleotide, preferably wherein the inhibitor and enhancer polynucleotides are selected from those listed in Table 1. In some embodiments, the inhibitor and enhancer polynucleotides are selected from the group comprising SEQ ID NO: 1, SEQ ID NO: 2, SEQ ID NO: 3 and SEQ ID NO: 4.
In some embodiments, the method according to any aspect of the invention is performed in a multi-well format, a microfluidic device or lateral flow device.
In some embodiments, the method steps are performed at a temperature in the range from 16° C. to 40° C., preferably at room temperature.
A third aspect of the invention provides a device comprising:
In some embodiments, the device is selected from a group comprising a multi-well plate, a microfluidic device and a lateral flow device.
In some embodiments, the device is a microfluidic device comprising:
Examples of a suitable microfluidic devices are shown in
A fourth aspect of the invention provides a nucleic acid detection kit comprising;
In some embodiments the signalling nanostructure in b) comprises:
In some embodiments, components (a) to (c) are as defined according to any aspect of the invention.
In some embodiments, the nucleic acid detection kit is configured into a device according to any aspect of the invention.
In some embodiments, at least one of the inhibitor polynucleotides and/or enhancer polynucleotides is structurally and/or chemically modified from its natural nucleic acid.
In some embodiments, said structural and/or chemical modification is selected from the group comprising the addition of tags, such as fluorescent tags, radioactive tags, biotin, a 5′ tail, the addition of phosphorothioate (PS) bonds, 2′-O-Methyl modifications and/or phosphoramidite C3 Spacers during synthesis.
Bibliographic references mentioned in the present specification are for convenience listed in the form of a list of references and added at the end of the examples. The whole content of such bibliographic references is herein incorporated by reference. Any discussion about prior art is not an admission that the prior art is part of the common general knowledge in the field of the invention.
Unless defined otherwise, all technical and scientific terms used herein have the same meaning as is commonly understood by one of skill in the art to which the invention belongs. Certain terms employed in the specification, examples and appended claims are collected here for convenience.
It must be noted that as used herein and in the appended claims, the singular forms “a,” “an,” and “the” include plural reference unless the context clearly dictates otherwise. Thus, for example, a reference to “a target sequence” includes a plurality of such target sequences, and a reference to “an enzyme” is a reference to one or more enzymes and equivalents thereof known to those skilled in the art, and so forth.
The phrases “nucleic acid” or “nucleic acid sequence,” as used herein, refer to an oligonucleotide, nucleotide, polynucleotide, or any fragment thereof, to DNA or RNA of genomic or synthetic origin which may be single-stranded or double-stranded and may represent the sense or the antisense strand, to peptide nucleic acid (PNA), or to any DNA-like or RNA-like material.
As used herein, the term “inhibitory complex” refers to a duplex of inhibitor polynucleotide and enhancer polynucleotide which inactivates DNA polymerase bound to it.
As used herein, the term “DNA polymerase inhibitor” or “inhibitor” is a polynucleotide comprising a conserved region and a variable region, wherein the conserved region is recognized and bound by the DNA polymerase enzyme, and the variable region is complementary to a portion of the enhancer polynucleotide.
As used herein, the term “enhancer” refers to a polynucleotide comprising a sequence that is complementary to a target polynucleotide sequence, of which a portion is involved in forming a duplex with complementary sequences of the variable region of the DNA polymerase inhibitor and a portion is involved in an overhang. Preferably, the enhancer is about 40 nucleotides in length, that is complementary to a target polynucleotide sequence, and 20 of the 40 nucleotides form the duplex with the inhibitor and 20 nucleotides of the 40 nucleotides form an overhang. Thus, the inhibitor and enhancer form a duplexed inhibitory DNA complex which inhibits DNA polymerase activity until such time as the enhancer is displaced upon duplex formation with target polynucleotide sequence.
As used herein, the term “transition state” refers to an optimum inhibitor complex:DNA polymerase ratio which is further optimized in respect of enhancer:inhibitor ratio (see, for example
The term “sample,” as used herein, is used in its broadest sense. For example, a biological sample suspected of containing SARS-CoV-2 genome sequences may comprise a bodily fluid; an extract from a cell, chromosome, organelle, or membrane isolated from a cell; a cell; genomic DNA, RNA, or cDNA (in solution or bound to a solid support); a tissue; a tissue print; and the like.
It would be understood that oligonucleotides used in the present invention may be structurally and/or chemically modified to, for example, prolong their activity in samples potentially containing nucleases, during performance of methods of the invention, or to improve shelf-life in a kit. Thus, the inhibitor and/or enhancer and/or signalling nanostructure or any oligonucleotide primers or probes used according to the invention may be chemically modified. In some embodiments, said structural and/or chemical modifications include the addition of tags, such as fluorescent tags, radioactive tags, biotin, a 5′ tail, the addition of phosphorothioate (PS) bonds, 2′-O-Methyl modifications and/or phosphoramidite C3 spacers during synthesis.
For example, the signalling oligonucleotide was modified for attachment chemistry with a 5′ thiol group. Other attachment modifications can be made on the 5′ end such as amino, acryldite, azide, etc.
The term “comprising” as used in the context of the invention refers to where the various components, ingredients, or steps, can be conjointly employed in practicing the present invention. Accordingly, the term “comprising” encompasses the more restrictive terms “consisting essentially of” and “consisting of.” With the term “consisting essentially of” it is understood that the phenotypic features of the present invention “substantially” comprise the indicated features as “essential” element.
Material and Methods
All oligonucleotide sequences can be found in Table 1 and were purchased from Integrated DNA Technologies (IDT). Genome sequences of SARS-CoV-2 (NC_045512), SARS-CoV (FJ882957), MERS (NC_019843), dengue virus (NC_001477) and influenza A subtype H1N1 virus (strain A/California/07/2009(H1N1), NC_026431-NC_026438) were obtained from NCBI RefSeq. Multiple sequence alignment was performed using the UGENE suite of tools [Okonechnikov K et al., Bioinformatics. 2012, 28:1166-1167]. To prepare molecular switches, we mixed inhibitor and enhancer oligonucleotides (Table 1, IDT) in a reaction buffer made up of 50 mM NaCl, 1.5 mM MgCl2, and 50 mM Tris-HCl (pH 8.5). The mixture was incubated at 95° C. for 5 min and slowly cooled at 0.1° C./s until the reaction reached 25° C. to form the inhibitory complex. Taq DNA polymerase (Promega) was then added to form the complete molecular switch.
GTGCGTACATAGATCGTTATCTGTCTAACGATCTATGTACGCACT
Bolded nucleotides indicate possible sites of biotin incorporation/removal.
Underlined nucleotides indicate mismatches.
To identify various states of the molecular switch, we varied the ratio of its constituents, first with the inhibitor and enhancer strand at 1:1 ratio, then at varying ratios of these two components. The resultant polymerase activity was measured through 5′ exonuclease degradation of fluorescent signalling probe. Briefly, equimolar amounts of fluorescent probe, template and primer (IDT) were mixed with deoxynucleotide triphosphates (dNTPs, Thermo Scientific) in the reaction buffer. The mixture was incubated at 95° C. for 5 min and slowly cooled to 25° C. at 0.1° C./s. Molecular switches were then added to the probe mixture and incubated at 25° C. while fluorescence readings were taken. Based on the observed changes in polymerase activity, we defined the different states of molecular switches: the open state is where the inhibitory complex is lacking (<20 nM), the closed state is where the inhibitory complex is in excess (>60 nM) and the transition state is the most responsive state (i.e., the vertex of the first derivative of the inhibition curve, where a small change in the switch composition would result in the largest change in polymerase activity). To characterize the responsiveness of the different switch states to nucleic acid targets, we prepared switches at the following representative composition and incubated the switches with target oligonucleotides: open state, 1 nM of inhibitor strand and 1 nM of enhancer strand; closed state, 100 nM of inhibitor strand and 100 nM of enhancer strand; and the transition state, 36 nM of inhibitor strand and 24 nM of enhancer strand. All experiments were also performed with scrambled oligonucleotides to determine background off-target signal.
Signalling oligonucleotides were immobilized on an ELISA plate as illustrated in
Two forms of signalling oligonucleotides were used and evaluated to amplify and transduce different types of polymerase activity, namely exonuclease- and elongation-based activity. In both approaches, control wells containing no polymerase were run concurrently to provide the baseline signal. For the exonuclease-based strategy, we immobilized dumbbell DNA signalling structures on the plate and measured the polymerase activity (5′ exonuclease activity) through the catalytic removal of biotin-modified nucleotides from the immobilized dumbbells. Briefly, we mixed sample targets with transition-state molecular switches and directly incubated the reaction with immobilized oligonucleotides, in the presence of dNTPs (Thermo Scientific), for 30 min at room temperature. Following washing steps with PBST and incubation with streptavidin-conjugated horseradish peroxidase (HRP, Thermo Scientific), we applied QuantaRed chemifluorescence substrate (Thermo Scientific) and measured the fluorescence intensity (Tecan) to evaluate the removal of biotin-modified nucleotides.
For the elongation-based strategy, polymerase activity was measured through the incorporation of biotin-modified nucleotides to self-priming, hairpin DNA signalling structures immobilized on the plate. Sample and molecular switches were added to the signalling structures and incubated in the presence of biotin-modified dNTPs mixture (TriLink BioTechnologies). Following incubation for 30 min at room temperature and washing with PBST, we incubated streptavidin-conjugated HRP (Thermo Scientific). After washing, we applied QuantaRed chemifluorescence substrate (Thermo Scientific) and measured the fluorescence intensity (Tecan) to evaluate the addition of biotin-modified nucleotides.
Transition-state molecular switches were prepared as previously described. Sample containing target was mixed with the prepared molecular switches to a final volume of 50 μl. The mixture was added to the self-priming DNA signalling structures, immobilized on the plate, in the presence of biotin-modified dNTP mixture. The reaction mixture was incubated for 30 min at room temperature. Following washing steps with PBST and incubation with streptavidin-conjugated horseradish peroxidase (HRP, Thermo Scientific), we applied QuantaRed chemifluorescence substrate (Thermo Scientific) and measured the fluorescence intensity (Tecan). For each sample, sample-matched positive (containing polymerase without inhibitory complex) and negative (scrambled molecular switch) controls were run concurrently for data normalization.
A prototype microfluidic device was fabricated through standard soft lithography as previously described [X. Wu, et al., Sci Adv 6, eaba2556 (2020)]. Briefly, 50-μm-thick cast molds were patterned with SU-8 photoresist and silicon wafers using a cleanroom mask aligner (SUSS MicroTec) and developed after ultraviolet (UV) exposure. Polydimethylsiloxane (PDMS, Dow Corning) and cross-linker were mixed at a ratio of 10:1 and casted on the SU-8 mold. The polymer was first cured at 75° C. for 30 min. Then, multiple nylon screws and hex nuts (RS Components) were positioned on the PDMS film over their respective channels and embedded in the PDMS, before a final curing step.
To immobilize the signalling oligonucleotides on the device, we treated the device's glass surface with (3-aminopropyl)triethoxysilane (APTES, 2% v/v, Sigma) in 95% ethanol for 1 h at room temperature. The chambers were then flushed with ethanol to remove excess APTES and dried. Separately, thiol-modified signalling oligonucleotides were activated as previously described. The activated oligonucleotides were then flowed in and incubated for 2 h at room temperature. After flushing with PBST to remove excess oligonucleotides, the chambers were blocked with 2% BSA for 1 h at room temperature. The chambers were then washed with PBST and the reaction buffer. To prepare the device for operation, we lyophilized the assay reagents within the device. The reagent mixture, containing inhibitor strand, enhancer strand, polymerase and biotin-modified dNTP mixture, was flowed into the device and lyophilized overnight (Labconco).
Operation steps of the microfluidic device are illustrated in
To enable smartphone analysis of the microfluidic CATCH assay, we developed a sensor that comprised a LED source, an optical filter and a magnification lens within a 3D-printed optical cage as previously described [X. Wu, et al., Sci Adv 6, eaba2556 (2020)]. The optical cage was fabricated from a UV-curable resin (HTM 140) using a desktop 3D printer (Aureus). The central wavelengths of the LED light source (Chaoziran S&T) and optical filter (Thorlabs) were 500 and 600 nm, respectively. The magnification lens (Thorlabs) was placed before the smartphone camera to improve the image quality. The assembled system measured 45 mm (width) by 45 mm (length) by 50 mm (height) in dimension and was equipped with two sliding slots for quick attachment to smartphones (Apple). Sensor performance was evaluated against a commercial microplate reader (Tecan) for different fluorescent dyes and intensities.
I
norm=(Itarget−Icontrol)/(Ipol−Icontrol)
where Inorm is the normalized fluorescence intensity, Itarget is the fluorescence intensity of the sample incubated with molecular switches against the target, Icontrol is the fluorescence intensity of the sample-matched negative control, incubated with scrambled control molecular switches, and Ipol is the fluorescence intensity of the sample-matched positive control, incubated with active polymerase.
To evaluate the specificity of the transition state compared to that of the closed state, molecular switches were mixed with targets with varying number of mismatches at positions that would most drastically affect the signal produced by the molecular switches [Ho N R Y et al., Nat Commun. 2018, 9: 3238] (Table 1). The resultant polymerase activity was measured using the assay on the plate as previously described. To characterize the sensitivity of the assay, we prepared serial ten-fold dilutions of the target and mixed the target samples with molecular switches in distinct states (e.g., transition vs. closed states) to evaluate changes in polymerase activity. To investigate the incubation time required to recover the functionality of lyophilized switches, we reconstituted the lyophilized reagents with the reaction buffer and incubated the mixture for less than 1 min, 5 min, 10 min, and 30 min before mixing with target and transferring to the functionalized plate for signalling. To evaluate the performance of lyophilized switches, we mixed lyophilized and non-lyophilized switches with target and the resultant polymerase activity was measured through 5′ exonuclease degradation of fluorescent signalling probe as previously described.
Human lung epithelial cell line (PC9) was obtained from American Type Culture Collection (ATCC) and grown in RPMI-1640 medium (HyClone) supplemented with 10% fetal bovine serum (FBS, HyClone) and 1% penicillin-streptomycin (Gibco) in a humidified 37° C. incubator with 5% CO2. The cell line was tested and free of mycoplasma contamination (MycoAlert Mycoplasma Detection Kit, Lonza, LT07-418). To evaluate the performance of the assay in biological samples, we prepared cell lysates through different protocols and spiked in synthetic target oligonucleotides, before testing the samples with molecular switches. RNase inhibitor was added to all lysate mixtures. Specifically, we lysed cell pellets through heating or incubating with detergent buffer. For heat treatment, cell pellets were resuspended in the reaction buffer and heated at 56° C. for 30 min, 70° C. for 5 min, or 90° C. for 5 min [Chin A W H, et al., The Lancet Microbe. 2020, 1: e10; Ladha A et al., medRxiv (2020). Doi.org/10.1101/2020.05.07.20055947]. For chemical lysis, we prepared lysis buffers, by mixing the reaction buffer with varying amounts of single or a mixture of detergents: Triton X-100, sodium dodecyl sulfate (SDS), Saponin, Tween-20, Igepal CA-630, NP-40 (Sigma). To optimize the chemical lysis composition and incubation duration, we evaluated various lysis conditions for their ability to rapidly lyse cells while maintaining good polymerase activity. To assess cell lysis efficiency, cells were incubated with the lysis buffers and the resultant cell numbers were counted using Countess II Automated Cell Counter (Thermo Scientific). Polymerase activity was measured through 5′ exonuclease degradation of fluorescent signalling probe, as described above.
RNA extraction was performed with a commercially available kit (RNeasy Mini, Qiagen) per manufacturer's protocol. Extracted RNA was quantified with Nanodrop spectrophotometer (Thermo Scientific). To detect specific RNA targets through gold-standard RT-qPCR analysis, extracted RNA was first reverse-transcribed to generate first-strand cDNA (MultiScribe Reverse Transcriptase, Thermo Scientific). For PCR analysis, to detect housekeeping genes (i.e., GAPDH and beta-actin), we used Taqman Fast Advanced Master Mix (Thermo Scientific) and primer sets (Taqman gene expression assays, Thermo Scientific) as recommended by the manufacturer. Amplification conditions consisted of 1 cycle of 95° C. for 2 min, 45 cycles of 95° C. for 1 s and 60° C. for 20 s. All thermal cycling was performed on a QuantStudio 5 real-time PCR system (Applied Biosystems).
A total of 48 clinical samples consisting of extracted RNA and heat-inactivated swabs were evaluated in this study. To determine the diagnostic performance of the CATCH assay, extracted RNA samples (positive, n=20; negative, n=9) were used directly on the CATCH assay, while swab lysates (positive, n=9; negative, n=10) were prepared through heating at 70° C. for 30 min, before measurement by the CATCH assay. SARS-CoV-2 clinical diagnoses were generated by commercial RT-qPCR assay (Fortitude Kit, MiRXES). Amplification conditions consisted of 1 cycle of 48° C. for 15 min, 1 cycle of 95° C. for 150 s, 42 cycles of 95° C. for 10 s and 59° C. for 42 s. Ct value <40 was determined as positive as per CDC's guidelines [Centers for Disease Control and Prevention, CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel. (2020), available at worldwidewebdotfda.gov/media/134922/download]. All measurements on clinical samples were performed in an anonymized and blinded fashion and finalized before comparison with clinical Ct value.
Unless otherwise stated, all measurements were performed in biological triplicate, and the data are presented as mean±standard deviation. For inter-sample comparisons, multiple pairs of samples were each tested via Student's t-test, and the resulting P values were adjusted for multiple hypothesis testing using Bonferroni correction. An adjusted P<0.05 was determined as significant. Receiver operating characteristic (ROC) curves were generated from patient profiling data and constructed by plotting sensitivity versus (1—specificity), and the values of area under the curve (AUC) were computed using the trapezoidal rule. The clinical reports were used as classifiers (true positives and true negatives). Detection sensitivity, specificity and accuracy were calculated using standard formulas. Statistical analyses were performed using GraphPad Prism software (version 7.0c).
The working principle of the CATCH assay is illustrated in
By ratiometric tuning of various switch components, we found the transition-state switches to be hyper-responsive to RNA targets, and leveraged this state to develop the CATCH assay for rapid, sensitive detection of SARS-CoV-2. To further enhance the detection signal, we measured the changes in polymerase activity through additional enzymatic amplification (
To develop the CATCH assay for detecting SARS-CoV-2, we designed molecular switches as specific probes against the viral RNA. We chose regions of the spike (S) gene [Yan C et al., Clin Microbiol Infect. 2020, 26: 773-779] and the nucleocapsid (N) gene [Broughton J P et al., Nat Biotechnol. 2020, 38: 870-874] of the virus as specific targets and constructed distinct molecular switches based on these sequences (
To establish the transition state, we further tuned the responsive-state molecular switches by titrating the amount of enhancer strand (i.e., through which target hybridizes and activates the switch) while keeping constant the amount of inhibitor strand (
Next, we devised a signalling mechanism to enzymatically amplify and measure the switch-induced polymerase activity. Specifically, we designed two signalling oligonucleotide structures to leverage different types of polymerase activity (i.e., elongation vs. exonuclease activity) and recruit additional enzymatic cascades (i.e., horseradish peroxidase, HRP) for signal amplification (
Motivated by the signalling performance, we developed the CATCH assay workflow to utilize transition-state molecular switches for responsive target recognition, and elongation-based multi-enzyme cascade for signal enhancement. Specifically, we mixed RNA targets with transition-state switches and directly incubated the reaction with immobilized oligonucleotides (30 minutes at room temperature) for signal transduction and enhancement. As compared to a similar assay using closed-state molecular switches (i.e., fully inactivated molecular switches and HRP-based signal enhancement), the CATCH assay demonstrated comparable specificity against target mismatches, even when the mismatches were introduced against the most sensitive segment of the switches (
To address the need for extensive sample preparation in conventional qPCR (i.e., RNA extraction), we next determined if the CATCH assay could be developed to bypass this crucial and limiting step. Using specific molecular switches designed for SARS-CoV-2 RNA targets (i.e., S-gene and N-gene switches), which demonstrated specific detection and minimal activity against sequences of other closely-related human coronaviruses (SARS-CoV and MERS-CoV) as well as other viruses causing diseases with similar symptoms (dengue virus and influenza A subtype H1N1 virus) (
Specifically, we explored two modes of direct lysis, namely thermal (
With these selected thermal and chemical lysis protocols, we first validated the ability of these methods to release and preserve endogenous RNA targets (i.e., GAPDH and beta-actin) in human lung epithelial cells. We demonstrated that for both endogenous targets tested, when assayed via RT-qPCR, all lysates generated similar cycle threshold (Ct) values as compared with the gold-standard extracted RNA samples (
To test the clinical utility of the CATCH platform for SARS-CoV-2 detection, we conducted a feasibility study with patient samples. We aimed at addressing the following questions: (1) if the CATCH assay can be applied directly to detect extracted RNA of nasopharyngeal swab samples (i.e., bypassing RT-qPCR), (2) if the CATCH platform can be used for direct detection of swab lysates (i.e., bypassing RNA extraction), and (3) the accuracy of CATCH in COVID-19 diagnosis.
We first tested swab-extracted RNA samples (n=49) using the CATCH assay. RNA samples were extracted through commercial columns and incubated directly with the CATCH mixture for 30 minutes at room temperature. Of the 49 extracted RNA samples, 24 were determined by gold-standard RT-qPCR assay as positive for COVID-19 infection and 25 as negative. The positive and negative diagnostic prediction of CATCH relative to the clinical RT-qPCR outcome were 100% and 92%, respectively (
To evaluate the clinical performance, across all tested clinical samples, we correlated the CATCH assay with the matched RT-qPCR Ct values (
Amongst current COVID-19 testing protocols, nucleic acid detection, particularly RT-qPCR, remains the gold standard. Nevertheless, the approach is almost exclusively performed in large, centralized clinical laboratories, due to its extensive processing, high complexity and need for trained personnel; reliance on RT-qPCR has thus placed much pressure on public health systems (Huang H, et al., ACS Nano. 2020, 14: 3747-3754; Weissleder R et al., Sci Transl Med. 2020, 12, eabc1931), leading to a significant global supply shortage and delayed diagnoses. For prompt detection and efficient management, rapid and accurate diagnostic assays are urgently needed [Ong C W M et al., Eur Respir J. 2020, 56: 2001727; Ong C W M et al., Int J Tubc Lung Dis. 2020, 24:547-548]. We developed the CATCH assay as an alternative nucleic acid detection method to complement the current gold standard. Specifically, the CATCH assay demonstrates distinct advantages, through its unique assay mechanism and facile clinical adaptation, to address multiple challenges of COVID-19 diagnostics.
From the assay perspective, CATCH leverages DNA-enzyme hybrid complexes as hyper-responsive molecular switches. By tuning their molecular composition, the multi-component molecular switches are prepared in a hyper-responsive state—the transition state—that can be readily activated upon the direct hybridization of even sparse RNA targets to turn on substantial enzymatic activity. CATCH thus achieves an enhanced response that that is not only bigger in magnitude, but also faster in kinetics. Yet, CATCH retains all key advantages inherent to molecular switching: 1) it is highly specific and activates only when complementary targets bind to the switches; 2) it can be readily integrated with other enzyme cascades (e.g., HRP) for further signal enhancement; and 3) it enables programmable design and rapid new assay prototyping. CATCH achieved a LOD of 8 RNA copies per μl (>10,000-fold more sensitive than our previous platform), could be completed in <1 hour at room temperature and applied directly to a variety of sample types (e.g., swab lysates). Its superior performance enables CATCH to accurately detect SARS-CoV-2 even in patient samples with a low viral load.
For clinical adaptation, CATCH detects through target hybridization, instead of conventional target amplification (as in RT-qPCR). This enables the technology to bypass essentially all critical steps of RT-qPCR (i.e., RNA extraction, reverse transcription and thermal cycling amplification). Importantly, CATCH supports versatile assay implementation to accommodate the different diagnostic needs of COVID-19. In its 96-well format, the assay configuration closely resembles conventional ELISA in terms of assay workflow and readout, and can be readily adapted for high-throughput analysis, using existing infrastructure of clinical laboratories (e.g., plate reader and trained personnel). In its portable format, CATCH is implemented through a miniaturized microfluidic cartridge, where assay reagents are lyophilized within the device for user-friendly application and smartphone-based detection [Yelleswarapu V et al., Proc Nat Acad Sci USA. 2019, 116: 4489-4495; Xu H et al., Sci Adv. 2020, 6: eaaz7445; Wu X et al., Sci Adv. 2020, 6: eaba2556]. For different clinical applications, the CATCH assay threshold should be adjusted with respect to the proposed application. This threshold setting presents a trade-off between assay sensitivity vs. specificity. For example, considering the potential application of CATCH as a preliminary screening test, we prioritized assay sensitivity when setting the current detection threshold (100% sensitivity, minimal false negatives and maximal false positives); even at this assay threshold, we determined a low incidence of false positives (<8%), which is within the range reported of existing assays (0-16.7%) [Surkova E et al., Lancet Respir Med. 2020, 8: 1167-1168; Cohen A N et al., medRxiv (2020). doi.org/10.1101/2020.04.26.20080911]. The cause of false results in nucleic acid tests could be assay-associated or PCR-based misclassification, both of which have been reported [Cohen A N et al., medRxiv (2020). doi.org/10.1101/2020.04.26.20080911; Vogels C B F et al., Nat Microbiol. 2020, 5: 1299-1305].
The technology has the potential to be expanded further. For COVID-19 diagnostics, in view of the rapidly evolving pandemic, we envision the integration of multiple CATCH switches, designed to recognize different genetic loci of SARS-CoV-2, to not only enhance the detection coverage of the infection, but also enable subtype differentiation and mutation identification [Toyoshima Y et al., J Hum Genet. 2020, 65(12): 1075-1082]. With its robust performance in minimally-processed clinical lysates, CATCH could be readily expanded to investigate other more accessible sample types (e.g., saliva and sputum) [Garg N et al., Lab Chip. 2019, 19: 1524-1533; Jeong J H et al., J Med Virol. 2014, 86: 2122-2127]. To further improve user-friendliness, the microfluidic CATCH platform could be integrated with automated liquid handling systems (e.g., computer-programmed fluidics and pumps for compact liquid handling) [Shaffer S M et al., Lab Chip. 2015, 15: 3170-3182; Yeh E C et al., Sci Adv. 2017, 3: e1501645]. Such sample expansion and system automation could facilitate new clinical opportunities for repeat-testing as well as self-testing. Finally, beyond the current COVID-19 pandemic, CATCH can be further developed to discover and measure new biomarker signatures. The platform could be applied across a spectrum of diseases (e.g., infectious diseases, cancers and neurodegenerative diseases) to facilitate sensitive detection of nucleic acid targets and composite signatures [Lim C Z J et al., Nat Commun. 2019, 10: 1144]. Further technical improvements, such as multiplexed microfluidic compartmentalization [Duncombe T A et al., Nat Rev Mol Cell Biol. 2015, 16: 554-567; Tokeshi M et al., Anal Chem. 2002, 74: 1565-1571; Shao H et al., Nat Commun. 2015, 6: 6999], could enable microarray-type assay implementation for highly-parallel biomarker discovery and large-scale clinical validation.
| Number | Date | Country | Kind |
|---|---|---|---|
| 10202101092X | Feb 2021 | SG | national |
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/SG2022/050031 | 1/24/2022 | WO |
