Summary Pseudomonas species have become reliable platforms for bioproduction due to their capability to tolerate harsh conditions imposed by large‐scale bioprocesses and their remarkable resistance to diverse physicochemical stresses. The last few years have brought forth a variety of synthetic biology tools for the genetic manipulation of pseudomonads, but most of them are either applicable only to obtain certain types of mutations, lack efficiency, or are not easily accessible to be used in different Pseudomonas species (e.g. natural isolates). In this work, we describe a versatile, robust and user‐friendly procedure that facilitates virtually any kind of genomic manipulation in Pseudomonas species in 3–5 days. The protocol presented here is based on DNA recombination forced by double‐stranded DNA cuts (through the activity of the I‐SceI homing meganuclease from yeast) followed by highly efficient counterselection of mutants (aided by a synthetic CRISPR‐Cas9 device). The individual parts of the genome engineering toolbox, tailored for knocking genes in and out, have been standardized to enable portability and easy exchange of functional gene modules as needed. The applicability of the procedure is illustrated both by eliminating selected genomic regions in the platform strain P. putida KT2440 (including difficult‐to‐delete genes) and by integrating different reporter genes (comprising novel variants of fluorescent proteins) into a defined landing site in the target chromosome.

Summary Owing to its wide metabolic versatility and physiological robustness, together with amenability to genetic manipulations and high resistance to stressful conditions, Pseudomonas putida is increasingly becoming the organism of choice for a range of applications in both industrial and environmental applications. However, a range of applied synthetic biology and metabolic engineering approaches are still limited by the lack of specific genetic tools to effectively and efficiently regulate the expression of target genes. Here, we present a single‐plasmid CRISPR‐interference (CRISPRi) system expressing a nuclease‐deficient cas9 gene under the control of the inducible XylS/Pm expression system, along with the option of adopting constitutively expressed guide RNAs (either sgRNA or crRNA and tracrRNA). We showed that the system enables tunable, tightly controlled gene repression (up to 90%) of chromosomally expressed genes encoding fluorescent proteins, either individually or simultaneously. In addition, we demonstrate that this method allows for suppressing the expression of the essential genes pyrF and ftsZ, resulting in significantly low growth rates or morphological changes respectively. This versatile system expands the capabilities of the current CRISPRi toolbox for efficient, targeted and controllable manipulation of gene expression in P. putida.

CRISPR/Cas technologies constitute a powerful tool for genome engineering, yet their use in non-traditional bacteria depends on host factors or exogenous recombinases, which limits both efficiency and throughput. Here we mitigate these practical constraints by developing a widely-applicable genome engineering toolset for Gram-negative bacteria. The challenge is addressed by tailoring a CRISPR base editor that enables single-nucleotide resolution manipulations (C·G → T·A) with >90% efficiency. Furthermore, incorporating Cas6-mediated processing of guide RNAs in a streamlined protocol for plasmid assembly supports multiplex base editing with >85% efficiency. The toolset is adopted to construct and deconstruct complex phenotypes in the soil bacterium Pseudomonas putida. Single-step engineering of an aromatic-compound production phenotype and multi-step deconstruction of the intricate redox metabolism illustrate the versatility of multiplex base editing afforded by our toolbox. Hence, this approach overcomes typical limitations of previous technologies and empowers engineering programs in Gram-negative bacteria that were out of reach thus far.