The specific targeting and editing of site-specific recombinases (SSRs) provide an appealing option for gene editing therapies where precision is crucial. Recent developments to engineer and evolve Cre/loxP-type SSRs, have made it possible to shift the specificity of Cre from its native target preference, the loxP site, to catalyze recombination of a range of DNA substrates. Effort in developing these designer-SSRs rely on the novel target selection and ability to incorporate mutations for successful retargeting of the SSR. In order to improve rational redesign of SSRs, it is critical to first understand Cre’s specificity for DNA, allowing for a more strategic target selection and identify specificity-altering mutations.

Here we developed a high-throughput assay for assessing the DNA specificity of Cre and evolved Cre variants. This method measures the activity of natural and designer-SSRs on a large set of over 5,000 rationally designed, non-randomized DNA target sequences, to profile the recombinase/DNA specificity. The assay is carried out in E. coli to minimize experimentation and eliminate artifacts, and employs widely accessible Illumina sequencing as the final readout.

We demonstrated the capabilities of this assay by profiling specificity and relative tolerance to base substitutions of Cre and an evolved Cre variant. By comparing the changes in DNA specificity of the complexes, we identified specificity-altering mutations and used molecular modelling and dynamics simulations to elucidate the mechanism behind their specificity switch. Furthermore, we used the approach to investigate the therapeutic potential of these novel recombinases and characterize their general specificity to detect potential off-targets.