Novel fluorescence-assisted whole-cell assay for engineering and characterization of proteases and their substrates.

We have developed a sensitive and highly efficient whole-cell methodology for quantitative analysis and screening of protease activity in vivo. The method is based on the ability of a genetically encoded protease to rescue a coexpressed short-lived fluorescent substrate reporter from cytoplasmic degradation and thereby confer increased whole-cell fluorescence in proportion to the protease's apparent activity in the Escherichia coli cytoplasm. We demonstrated that this system can reveal differences in the efficiency with which tobacco etch virus (TEV) protease processes different substrate peptides. In addition, when analyzing E. coli cells expressing TEV protease variants that differed in terms of their in vivo solubility, cells containing the most-soluble protease variant exhibited the highest fluorescence intensity. Furthermore, flow cytometry screening allowed for enrichment and subsequent identification of an optimal substrate peptide and protease variant from a large excess of cells expressing suboptimal variants (1:100,000). Two rounds of cell sorting resulted in a 69,000-fold enrichment and a 22,000-fold enrichment of the superior substrate peptide and protease variant, respectively. Our approach presents a new promising path forward for high-throughput substrate profiling of proteases, engineering of novel protease variants with desired properties (e.g., altered substrate specificity and improved solubility and activity), and identification of protease inhibitors.