New Gene Editing Tool for Cold-Adapted Bacteria

To develop a functional CRISPR/Cas9 editing system for P. fuliginea, we constructed a recombinant plasmid based on pK18mobsacB-Ery, a shuttle vector that has been successfully used for gene knockout in Pseudoalteromonas via double-crossover homologous recombination, and is capable of conjugative transfer between E. coli and Pseudoalteromonas (15). While pK18mobsacB-Ery has been previously used for gene knockout via double-crossover homologous recombination, in our CRISPR/Cas9 system the, sacB counter-selection marker serves a different purpose: it facilitates plasmid curing after genome editing, thereby ensuring removal of the CRISPR/Cas9 vector and enabling scarless genetic modifications. The CRISPR/Cas9 cassette was assembled by inserting the Cas9 coding sequence and a sgRNA expression module into the pK18mobsacB-Ery backbone. For efficient gene expression in P. fuliginea, we screened suitable promoters to drive both Cas9 and sgRNA expression. As a result, the D4-12 sequence of the Pf1 sRNA promoter was selected, as it had previously been demonstrated to be an active regulatory element in P. fuliginea BSW20308 (28). Its native origin increases the likelihood of strong transcriptional activity in this host. To ensure proper transcription termination, the rrnB T1 sequence was employed as a rho-independent terminator downstream of both expression units. This element has been widely used in prokaryotic expression systems due to its robust and reliable termination efficiency, which prevents transcriptional read-through (29). The Cas9 protein used in this system was derived from Streptococcus pyogenes MGAS5005 (30) and was codon-optimized to match the codon usage preferences of Pseudoalteromonas, thereby improving translation efficiency in the heterologous host. The complete expression cassette, which includes the D4-12 promoter, Cas9, sgRNA module, and rrnB T1 terminator, was cloned into the pK18mobsacB-Ery plasmid to yield the final CRISPR/Cas9 construct (Fig. 1A). To validate the CRISPR/Cas9 system in P. fuliginea, we performed gene editing on the P. fuliginea BSW20308 strain, as detailed in the subsequent sections. In contrast to prior work using CRISPRi in P. luteoviolacea, which achieved transcriptional repression of secondary metabolite and host-associated genes via dCas9 (27), our system implements active Cas9 to produce double-strand breaks, enabling precise, template-directed genome editing (including deletions and insertions) in a cold-adapted species.

The fliJ, integral to the bacterial flagellar structure and pivotal for cell motility and chemotaxis, is present within the genome of P. fuliginea BSW20308 (31). Consequently, we opted to eliminate this gene to assess the efficacy of the CRISPR/Cas9 system in P. fuliginea. A CRISPR/Cas9 suicide plasmid, pK18Ery-fliJ-CRISPR/Cas9, was constructed based on the pK18mobsacB-Ery backbone. This plasmid carried the Cas9 gene, a specific sgRNA targeting fliJ, and two homologous arms (1,000 bp each) flanking the fliJ locus. Conjugative transfer of the plasmid into P. fuliginea BSW20308 was performed, and transconjugants were selected on 2216E agar containing erythromycin. Following successful transfer, the Cas9 protein was expressed under the control of the D4-12 promoter. The Cas9-sgRNA complex induced a double-strand break at the target site adjacent to the PAM sequence (Fig. 1B ), which was repaired via homologous recombination using the provided flanking sequences, resulting in the deletion of the target gene.

To confirm the gene knockout, we performed PCR using a primer pair (fliJ-wS/ fliJ-dA) spanning the upstream homologous region and the 3′ end of the fliJ locus. The deletion of the fliJ gene (447 bp) was confirmed by gel electrophoresis (Fig. 2A) and further validated through DNA sequencing. Out of 17 randomly selected erythromycin-resistant clones, 15 exhibited successful gene deletions (Fig. S1, Data Set S1Supplementary Data 1). In our hands, under identical selection and screening conditions, classical double-crossover recombination yielded no confirmed fliJ mutants among 100 screened colonies (Fig. S2), while CRISPR/Cas9 generated 15 confirmed edits among 17 screened clones. To further validate the successful genetic manipulation via phenotypic analysis, cell motility assays were performed on 0.3% agar plates. The ΔfliJ mutant exhibited a significant reduction in motility compared to the wild-type strain, aligning with the anticipated role of fliJ in flagellar assembly (Fig. 2B). Complementation of the ΔfliJ mutant with a plasmid-borne copy of fliJ under its native promoter fully restored motility to wild-type levels, confirming that the impaired motility phenotype was specifically attributable to the loss of fliJ. This impaired motility may reduce the strain’s ability to respond to environmental stimuli or colonize new niches, highlighting the role of fliJ in microbial adaptation to physically challenging or nutrient-scarce environments such as polar oceans.

P. fuliginea BSW20308 is capable of producing a characteristic dark blue pigment when grown on 2216E medium. Previous research identified a potential indigoidine biosynthetic gene cluster in this strain, which includes the indA gene as a candidate gene likely responsible for pigment production (31). To investigate the function of indA and further validate the CRISPR/Cas9 system, we constructed a deletion mutant targeting this gene. Homologous fragments of 1,000 bp upstream and downstream of indA were inserted into the pK18Ery-indA-CRISPR/Cas9 plasmid. The resulting construct was introduced into P. fuliginea BSW20308 via conjugation, and integration was confirmed by PCR. Subsequently, counter-selectable 2216E media containing 15% sucrose was utilized to produce the ΔindA mutant. The PCR product of ΔindA was 310 bp, while the corresponding wild-type PCR product was 1,255 bp (Fig. 2C). DNA sequencing further confirmed the deletion of the 945 bp of the indA coding region. To assess the phenotypic effects of this deletion, we compared pigment production between wild-type and mutant strains. The ΔindA mutant lost the ability to synthesize the characteristic blue pigment, confirming the functional role of indA in indigoidine biosynthesis (Fig. 2D). Moreover, functional complementation was performed by introducing a plasmid carrying the intact indA gene (pK18-indA) into the ΔindA mutant. The complemented strain regained the wild-type phenotype and restored pigment production, providing further evidence that the loss of pigment was directly attributable to indA deletion. In total, 24 erythromycin-resistant colonies were screened, of which 19 exhibited successful indA gene deletions (Fig. S3).

In a previous study, an attempt was made to delete the small RNA gene Pf1 through double-crossover homologous recombination in P. fuliginea BSW20308. However, only one mutant was recovered from nearly a thousand colonies screened, resulting in an extremely low efficiency of approximately 0.1% (28). To address this limitation, we constructed CRISPR/Cas9-based plasmids targeting three small RNAs (Pf1, Pf2, and Pf3), designated as pK18Ery-Pf1-CRISPR/Cas9, pK18Ery-Pf2-CRISPR/Cas9, and pK18Ery-Pf3-CRISPR/Cas9, respectively. These plasmids were mobilized into P. fuliginea BSW20308 in a sequential manner. The pK18Ery-Pf1-CRISPR/Cas9 plasmid was first introduced into BSW20308 via conjugation. The successful deletion of Pf1 was confirmed by PCR and DNA sequencing (Fig. S4). Using the ΔPf1 mutant as a starting strain, we next knocked out Pf2 to generate the ΔPf12 double mutant, followed by deletion of Pf3 to obtain the ΔPf123 triple mutant. For Pf1, Pf2, and Pf3, the CRISPR/Cas9 system generated mutants in 22/24, 15/17, and 11/15 colonies tested, respectively (Fig. S4, Data Set S1Supplementary Data 1). These values represent a substantial improvement over the homologous recombination method and highlight the effectiveness of our CRISPR/Cas9 platform in targeting non-coding RNA genes in P. fuliginea.

The CsrA protein is a global post-transcriptional regulator involved in diverse cellular processes through RNA binding, and plays a central role in the carbon storage regulator (Csr) system. To investigate its function in P. fuliginea, we initially attempted to generate a complete knockout of the csrA gene. However, repeated attempts failed to produce viable knockout mutants, suggesting that csrA may be essential for cell survival under standard growth conditions (32, 33). To circumvent this limitation, we constructed a modified plasmid, pK18Ery-csrA_1-50-CRISPR/Cas9, which included a donor fragment encoding the first 50 amino acids of CsrA. This design aimed to generate a partial deletion or functional disruption of csrA, while preserving minimal functionality to maintain cell viability. The plasmid was mobilized into P. fuliginea via conjugation, and positive transconjugants were identified using two sets of plasmid-specific primers (Ery-F/Ery-R and SacB-F/SacB-R). Sequencing of the csrA locus in edited clones revealed a 39 bp deletion at the 3′ end of the gene (Fig. S5), confirming the intended truncation. Among the screened colonies, 6 out of 24 exhibited the targeted modification (Data Set S1Supplementary Data 1). This relatively low frequency is likely due to the essential role of CsrA as a global post-transcriptional regulator. Disrupting this gene may impair key cellular functions, making complete or partial edits detrimental to cell survival under standard conditions.

To evaluate the gene knock-in capability of the CRISPR/Cas9 system in P. fuliginea BSW20308, we engineered the insertion of a 3×FLAG epitope tag at the 3′ end of the csrA gene. A recombinant plasmid, pK18Ery-csrA::3×FLAG-CRISPR/Cas9, was constructed by assembling the csrA sequence, the 3×FLAG tag, and homologous arms flanking the csrA locus, along with the guide RNA cassette and Cas9 gene, into the pK18Ery backbone. This plasmid was then introduced into P. fuliginea via conjugation. Entry of the plasmid was confirmed using plasmid-specific primer pairs (Ery-F/Ery-R and SacB-F/SacB-R). Successful knock-in of the 3×FLAG sequence was confirmed by PCR using primers FLAG-KI-dS and FLAG-KI-wA. The resulting 1,141 bp amplicon was detected in edited mutants but not in the wild-type strain (Fig. 3A). DNA sequencing confirmed precise insertion of the 3×FLAG tag at the 3′ terminus of the csrA gene. Protein-level expression of the CsrA-3×FLAG fusion product was validated via western blot using a FLAG-specific antibody. The results showed detectable expression of the CsrA-3×FLAG fusion protein in the knock-in strains (Fig. 3B), further validating the insertion. Editing efficiency was assessed by screening 48 erythromycin-resistant clones through PCR. Of these, 46 showed successful amplification of the 3×FLAG tag (Fig. S6). To eliminate the possibility of false negatives, the two non-amplified clones were retested in triplicate, all of which yielded no detectable bands.

To further demonstrate the versatility of the CRISPR/Cas9 system in P. fuliginea, we performed targeted gene tagging with a fluorescent reporter. A gene encoding a protein with a conserved Tse2 ADP-ribosyltransferase toxin domain was selected as the insertion site for a green fluorescent protein (GFP) reporter (34), providing a useful model for downstream applications. A donor construct was designed by appending the gfp gene to the 3′ end of the tse2 coding sequence within the pK18Ery-tse2::gfp-CRISPR/Cas9 plasmid. This recombinant plasmid, carrying homologous arms flanking the tse2 gene, Cas9, and the sgRNA cassette, was mobilized into P. fuliginea via conjugation. Successful integration of the gfp sequence at the 3′ end of the target gene was verified by PCR and confirmed through DNA sequencing (Fig. 3C). Fluorescence microscopy revealed detectable, albeit weak, GFP signal in the modified strain. This observation was further supported by fluorescence quantification, confirming expression of the Tse2-GFP fusion protein (Fig. 3D). Among 24 erythromycin-resistant clones screened, 20 showed successful insertion of the gfp gene (Fig. S7). These results validate the feasibility of using CRISPR/Cas9 for targeted fluorescent tagging in P. fuliginea, offering a valuable tool for real-time protein tracking and functional analysis in this organism.

This study presents the establishment of a CRISPR/Cas9-based genome editing system for P. fuliginea. By constructing a series of optimized editing plasmids, we demonstrated that this system enables efficient gene knockouts and knock-ins with high reliability and versatility. Across multiple genomic targets—including structural genes (fliJ), biosynthetic genes (indA), regulatory elements (csrA), small RNAs (Pf1–Pf3), and tagged reporter constructs (csrA::3×FLAG and tse2::gfp)—editing efficiencies consistently exceeded 70% except for the relatively low efficiency of the csrA gene. Compared to traditional double-crossover recombination, which showed low success in similar editing attempts, the CRISPR/Cas9 system proved to be more efficient and practical. Overall, this CRISPR/Cas9 platform provides a powerful and flexible genetic toolkit for P. fuliginea, greatly enhancing its potential as a model organism for studying cold adaptation, regulatory networks, and host-microbe interactions. It also lays a technical foundation for future applications in marine biotechnology and synthetic biology involving polar marine bacteria.

The strains and plasmids used in this study are listed in Data Set S1Supplementary Data 1. E. coli WM3064 strains were grown in LB medium with 0.3 mM DAP (diaminopimelic acid) at 37°C (10 g tryptone, 5 g yeast extract, and 10 g NaCl dissolved in 1,000 mL of deionized water), supplemented with kanamycin (50 mg/L) or gentamicin (10 mg/L). The P. fuliginea BSW20308 and its mutation strains were grown in 2216E medium. The plasmids used in this study are listed in Data Set S1Supplementary Data 1. The antibiotics were added at the following concentrations: 50 µg/mL for kanamycin (Kan); 25 µg/mL for erythromycin (Ery).

The plasmids were transferred into P. fuliginea through conjugation, following the established protocols described in previous reports (15, 35). In brief, E. coli WM3064 (donor) and recipient strains were cultured to an OD₆₀₀ of 0.6–1.0. The donor and the recipient strains were collected by centrifugation at 12,000 rpm, and washed twice with MLB medium (modified LB medium containing 10 g tryptone, 5 g yeast extract, and 10 g NaCl dissolved in 500 mL deionized water and 500 mL sterile seawater). The combined strain mixture was resuspended in 100 µL MLB medium and then dropped onto MLB plates supplemented with DAP. The plates were incubated at 25°C for 24 h. The bacterial lawn was scraped off, washed multiple times, and then spread onto erythromycin-containing 2216E plates to select the positive transconjugants.

The design of gRNA was carried out on the website (https://zlab.bio/guide-design-resources↗) in this study. The appropriate PAM type and host genome were selected through the "Cas-Designer" tool within the website. The target gene sequence that needed to be edited was then input into the "Target Sequence" field according to the specified format. After submission, the results were filtered based on requirements. The website featured a built-in scoring function, and typically, the gRNA with a higher score was selected for further use. The donor fragments, gRNA cassette, and Cas9 gene were ligated into the plasmid pK18mobsacB-Ery, digested with BamH I and Hind III using the ClonExpress II one-step cloning kit (Vazyme Co., Ltd., China). Suicide CRISPR/Cas plasmid was transformed into E. coli WM3064 and then mobilized into P. fuliginea via conjugation. The positive transconjugants were screened by 2216E plates containing erythromycin (25 µg/mL) and confirmed by PCR followed by DNA sequencing. The mutant strains were spread onto 2216E solid medium containing 15% sucrose and incubated statically at 25°C until single colonies emerged. This process was repeated for 2–3 generations to ensure stability of the mutation. Subsequently, single colonies were picked and inoculated into 2216E liquid medium containing 15% sucrose for activation. To verify the loss of the plasmid, PCR was performed using two sets of primers specific to the plasmid fragments as amplification primers. The absence of PCR products corresponding to the plasmid-specific fragments would indicate successful loss of the plasmid.

Briefly, a knockout plasmid was constructed by cloning ~1,000 bp fragments upstream and downstream of the target gene into the pK18mobsacB-Ery vector. The recombinant plasmid was introduced into E. coli WM3064 and subsequently mated with P. fuliginea BSW20308 on plates supplemented with DAP. After overnight incubation at 25°C, cells were resuspended from the plates, spread onto 2216E agar containing erythromycin, and incubated for selection of single-crossover. Colonies were screened by PCR to confirm the integration event. For the second crossover, selected colonies were cultured without antibiotics, followed by plating on 2216E agar containing sucrose to counter select for plasmid loss. Resulting colonies were PCR-verified and confirmed by DNA sequencing to identify successful in-frame deletions.

Strains were cultured to the exponential growth phase (OD₆₀₀ of 0.4), centrifuged and resuspended in fresh 2216E medium for motility and biofilm assays. Cell motility was measured using semisolid agar plates in triplicate. For each strain, 5 µL of re-suspended cells were placed in the center of a 2216E plate containing 0.3% agar and incubated at 32°C for over 24 h.

For western blot analysis, samples were transferred from the SDS-polyacrylamide gels onto the polyvinylidene difluoride (PVDF) membranes by electroblotting. The PVDF membranes were blocked for 2 h with 50 mL of 5% (wt/vol) milk powder that dissolved in PBST (PBS with 0.5% Tween-20), and then incubated overnight at 4°C with primary antibody (anti-FLAG Tag monoclonal antibody, Sangon Biotech, #D191041, China). Membranes were then washed with PBST solution three times, followed by incubation with secondary antibody (HRP-conjugated Goat anti-mouse IgM, Sangon Biotech, #D110103, China) for 30 min. After washing with PBST for three times, the blot was developed using High-sensitivity Plus ECL luminescence reagent (Sangon Biotech, #C520045, China) and visualized using ChemiDoc MP Imaging System (Bio-Rad, USA). The strains with native CsrA were used as the negative controls.

The WT and tse2::gfp mutants were grown in 2216E medium, cells were collected by removing the medium, washed three times, and resuspended in 200 µL of sterile distilled water. The WT strain was used as the negative control. Five biological replicates were performed for both the control and experiment groups. The GFP fluorescence intensity was measured using a microplate reader (excitation, 485 nm; emission, 538 nm) and divided by the corresponding OD₆₀₀ value to determine the unit fluorescence intensity.