Detection and discrimination of single nucleotide polymorphisms by quantification of CRISPR-Cas catalytic efficiency
The specificity of CRISPR-Cas12 assays is attractive for the detection of single nucleotide polymorphisms (SNPs) implicated in, e.g., SARS-CoV-2 variants. Such assays often employ endpoint measurements of SNP or wild type (WT) activated Cas12 trans-cleavage activity; however, the fundamental kinetic effects of SNP versus WT activation remain unknown. We here show that endpoint-based assays are limited by arbitrary experimental choices (like used reporter concentration and assay duration) and work best for known target concentrations. More importantly, we show that SNP (versus WT) activation results in measurable shifts in the Cas12 trans-cleavage substrate affinity (KM) and apparent catalytic efficiency [Formula]. To address endpoint-based assay limitations, we then develop an assay based on the quantification of Michalis-Menten parameters and apply this assay to a 20-base pair WT target of the SARS-CoV-2 E gene. We find that the [Formula] measured for WT is 130-fold greater than the lowest [Formula] among all 60 measured SNPs (compared to a 4.8-fold for endpoint fluorescence of the same SNP). KM also offers strong ability to distinguish SNPs, varies 27-fold over all the cases, and is insensitive to target concentration. Lastly, we point out trends among kinetic rates and SNP base and location within the CRISPR-Cas12 targeted region.