CRISPRi loss-of-function mutations revealed through experimental evolution in Burkholderia cenocepacia

Essential genes encode proteins required for cell growth and survival, which makes them ideal targets for controlling bacterial growth during infection or in industrial applications. A gene silencing approach, CRISPR interference (CRISPRi), enables transcriptional control of essential genes, allowing phenotypic characterization of knockdown mutants. We previously developed a CRISPRi system for the Gram-negative bacteria Burkholderia, consisting of a chromosomally encoded rhamnose-inducible dCas9 and a plasmid encoding constitutively expressed synthetic guide RNAs (sgRNAs) targeting essential genes. Despite the system's utility, the CRISPRi mutants often reverted to wild-type growth phenotypes, hindering their use in gene-to-function studies that require many generations. To understand the reasons for the CRISPRi mutants’ phenotypic reversions, we performed experimental evolution of 15 CRISPRi mutants under dCas9-inducing and non-inducing conditions, followed by quantitative gene expression analysis and whole-genome sequencing (WGS). While the CRISPRi machinery remained genetically stable in the CRISPRi-evolved mutants, reverse transcription quantitative PCR revealed a loss of CRISPRi repression in several mutants. Immunoblotting analysis further demonstrated that dCas9 expression was abolished, suggesting a defect in rhamnose uptake. One evolved CRISPRi mutant showed single-nucleotide polymorphisms (SNPs) affecting putative ABC sugar transporters. This work highlights the strengths and limitations of CRISPRi-based downregulation of essential genes for extended control of bacterial growth.