ABSTRACT
The CRISPR/Cas9 system offers an extremely versatile genome editing technology by employing an RNA-
guided endonuclease, Cas9. Originally isolated from Streptococcus pyogenes, this system has been adapted
for use in a wide range of organisms and can be programmed to manipulate almost any site in the genome.
While Cas9 is easy to reprogram, the efficiency of Cas9-mediated editing varies considerably across different
genomic targets. Unlike other common bacterial endonucleases, Cas9 exhibits single-turnover kinetics where it
forms a stable product complex and requires an external force to release the cut DNA. This persistent product
state impairs access to the double-strand break by repair machinery and contributes to reduced genome editing
efficiency. Despite an abundance of Cas9 structures, the structure of the product complex remains
uncharacterized as previous studies were carried out under conditions that prevent DNA cleavage. As a result,
our structural understanding of the entire Cas9 reaction cycle remains incomplete and insufficient to explain why
Cas9 exhibits single-turnover kinetics. Our lab recently used cryo-EM to determine the structure of the
catalytically active Cas9-sgRNA-dsDNA ternary complex and captured three distinct conformational states (pre-
and post-catalytic, and product states). These structures provide new insight into the coordination of Cas9
domains throughout catalysis and reveal persistent Cas9-nucleic acid interactions in the product state. Guided
by this new insight, we will expand upon these studies to define the major structural elements contributing to
Cas9 catalysis and single-turnover kinetics. Using innovative in vitro kinetic assays and established structural
analyses, we will investigate the structural features that control the rate at which Cas9 cuts and releases the
targeted DNA. Using rational design in conjunction with random mutagenesis, we will engineer a multi-turnover
enzyme capable of spontaneously releasing its cleaved DNA product. Using an in vitro genome editing assay
and an in vivo ¿-galactosidase assay, we will screen Cas9 mutants for an increased rate of DNA cleavage. We
anticipate the proposed studies will not only advance our molecular understanding of the Cas9 reaction cycle,
but also support the development of more efficient CRISPR/Cas9 technologies.
