Developing multiplexed genome-edit platforms in yeast (Saccharomyces cerevisiae) using a single gRNA-mediated CRISPR-Cas9

Abstract

The budding yeast Saccharomyces cerevisiae is a versatile microbial platform to build synthetic metabolic pathways for the production of a diverse array of chemicals. To expedite the construction of synthetic pathways in yeast, a novel multiplex genome-editing platform was developed where CRISPR-Cas9 can be applied to simultaneously integrate up to five foreign genes by a single transformation with one gRNA. To choose optimal loci for integration and expression of transgenes, eight desirable intergenic loci, located adjacent to highly expressed genes, were identified. The eight intergenic loci were fully characterized for different parameters after integrating a green fluorescent protein (GFP) cassette – CRISPR-mediated GFP integration efficiency, degree of GFP expression, growth rates of GFP-integrated strains, and genomic stability of the GFP integration. From these analyses, five loci were selected to build the multiplex platform where a common 23-bp DNA comprised of 20-bp synthetic DNA and 3-bp protospacer adjacent motif (PAM) was seamlessly placed in the five loci in a sequential manner. This process resulted in five distinct yeast strains harboring one to five copies of the synthetic gRNA-binding site in the genome. Using these pre-engineered yeast strains, simultaneous double, triple, quadruple, and quintuple gene integrations were demonstrated at between 84.8% and 98.0% integration efficiencies using a 3-gene betalain biosynthetic pathway and geneticin and hygromycin B resistance markers. The quadruple and quintuple gene integration platforms were applied to successfully generate yeast strains synthesizing a representative sesquiterpene lactone, costunolide, and its precursor, germacrene A acid. This work demonstrates the utility of the single gRNA-mediated CRISPR-Cas9 platform to build complex metabolic pathways in yeast.