Target Selection, Homokaryotic Isolation, and Screening Methods for Gene Editing in the Destructive Global Pathogen, Phytophthora cinnamomi

Phytophthora cinnamomi is a major plant pathogen that affects economically important crops and natural ecosystems, posing a threat to global biodiversity. While gene editing has emerged as a powerful tool for functional genomics in various Phytophthora species, its application in P. cinnamomi remains underexplored. To address this gap, our study investigated the challenges of implementing CRISPR/Cas9-mediated gene editing in P. cinnamomi, with the insights gained applicable to other gene editing platforms. We designed guide RNAs (gRNAs) targeting β-cinnamomin, a highly basic elicitin expressed by the pathogen during early infection stages, known for its role in sterol recruitment. Using an “all-in-one” plasmid containing the gRNA, Cas9, and an antibiotic resistance gene as a selectable marker, we transformed P. cinnamomi protoplasts via PEG/CaCl2-mediated methods. The successful integration of the nptII gene, which confers geneticin (G418) resistance, was confirmed in heterokaryotic regenerants. To isolate pure mutants and eliminate wild-type dominance, we derived homokaryotic colonies from nptII-positive transformants. Mutation screening was performed using mismatch detection assays, T7 endonuclease 1 (T7E1), and restriction fragment length polymorphism (RFLP), followed by Sanger sequencing. Despite the integration of the nptII gene, the β-cinnamomin sequence in the transformants remained identical to the wild-type sequence, indicating challenges in detecting and achieving targeted gene disruption. This study identifies critical steps for optimising mutagenesis in P. cinnamomi, highlighting the importance of thorough preliminary screening, effective separation of heterokaryotic populations, and the isolation of homokaryotic colonies.