The specificity of CRISPR-Cas12 assays is attractive for the detection of single nucleotide polymorphisms (SNPs) implicated in, e.g., cancer and 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 kinetic shifts in the Cas12 trans-cleavage substrate affinity (K_M) and apparent catalytic efficiency (k_cat^*/K_M). To address endpoint-based assay limitations, we then develop an assay based on the quantification of Michaelis-Menten parameters and apply this assay to a 20 base pair WT target of the SARS-CoV-2 E gene. We find that the k_cat^*/K_M measured for WT is 130-fold greater than the lowest k_cat^*/K_M among all 60 measured SNPs (compared to a 4.8-fold for endpoint fluorescence of the same SNP). K_M also offers a strong ability to distinguish SNPs, varies 27-fold over all the cases, and, importantly, is insensitive to the target concentration. Last, we point out trends among kinetic rates and SNP base and location within the CRISPR-Cas12 targeted region.

Charles Blanluet

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