Mycobacterium abscessus ATCC 19977 was used as the parental strain for all genetic manipulations. Cultures were grown in Middlebrook 7H9 broth or on Middlebrook 7H10 agar (BD Biosciences, United States) supplemented with 10% OADC (oleic acid, albumin, dextrose, and catalase), 0.2% glycerol (Sangon Biotech), and 0.05% Tween-80 (Sangon Biotech, broth only) at 37°C. Plasmid-containing strains were maintained with the following antibiotics: kanamycin (MCE, United States, 100 μg/mL) for pCas9-mScarlet-containing strains and zeocin (Invitrogen^™, 20 μg/mL) for pQL033-X-sg-harboring strains. Anhydrotetracycline (aTc; MCE, United States, 500 ng/mL) was used to induce CRISPR components. Escherichia coli DH5α (Tsingke; transformation efficiency >10⁹ CFU/μg) was used for plasmid construction and propagation in LB medium (Sangon Biotech) with kanamycin (100 μg/mL) or zeocin (20 μg/mL) as required.

To minimize toxicity, the CRISPR/Cas9 system was implemented using two separate plasmids. The Cas9 expression plasmid was derived from pLJR962 (Addgene #115162) via site-directed mutagenesis to introduce A9D and A599H mutations, restoring nuclease activity and generating pCas9. The mScarlet fluorescent reporter was inserted into the EcoRV (New England Biolabs, United States; 10,000 U/mL) site of pCas9 generate pCas9-mScarlet plasmid, allowing visual selection of transformants.

For targeted gene deletion, we developed a dual-sgRNA system based on pKM461 (Addgene #108320). As shown in Supplementary Figure S1, the anhydrotetracycline-inducible promoter (P_tet), kanamycin resistance gene, and Sth1 sgRNA scaffold (Sth1SC) were amplified from pLJR962 and assembled into pKM461through SapI (New England Biolabs, United States; 10,000 U/mL) and EcoRv sites, generating the pQL033-SapItwin-sgSC plasmid, which incorporated SapI restriction sites for Golden Gate assembly of target-specific sgRNAs. The pUC19-CS-ZeoR plasmid was constructed by inserting the Sth1SC-ZeoR-Ptet cassette into the pUC19 backbone. Target-specific protospacer sequences were designed to flank the genomic region of interest (e.g., Mab_0673/0674) and were incorporated into primers used for amplifying the Sth1SC-ZeoR-Ptet cassette, which contained SapI recognition sites. The resulting PCR product was then cloned into pQL033-SapItwin-sgSC to generate the final dual-sgRNA expression plasmid. All plasmids were confirmed by Sanger sequencing (Tsingke) before use.

M. abscessus cells were grown from 1 mL of frozen stock inoculated into 100 mL of Middlebrook 7H9 broth supplemented with 0.2% glycerol, 0.01% Tween-80, and 10% OADC. Cultures were incubated at 37°C, 180 rpm until mid-log phase (OD₆₀₀ = 0.6–0.8), followed by 0.2 M glycine induction for 3 h. Cultures were then pelleted by centrifugation (room temperature, 4,000 rpm, 10 min), and washed sequentially with 30 mL, 20 mL, and 10 mL of 10% glycerol (in deionized water). Cells were finally resuspended in 1 mL of 10% glycerol and ready for electroporation.

Genetic manipulation of M. abscessus was performed using optimized electroporation protocols. Competent cells were prepared as described above. For initial transformation, 1–2 μg of pCas9-mScarlet (or pCas9-mScarlet-noint) plasmid DNA was electroporated using a Bio-Rad Gene Pulser Xcell with parameters set to 2.5 kV, 25 μF, and 1,000 Ω. Transformants were recovered in 7H9/OADC medium for 4 h before plating on selective 7H11/OADC agar containing 100 μg/mL kanamycin. After 4–6 days incubation at 37°C, pink colonies expressing mScarlet were picked for verification by PCR and sequencing of the Cas9 cassette.

To generate knockout mutants, competent cells of the pCas9-mScarlet (or pCas9-mScarlet-noint) strain were transformed with 1 μg of the appropriate pQL033-X-sg plasmid (expressing dual sgRNAs). Following electroporation and recovery, transformants were selected on 7H11 plates containing both kanamycin (100 μg/mL) and zeocin (20 μg/mL). For gene deletion, positive clones were grown to OD600 ~ 0.8 in 7H9 medium with antibiotics, then split into two cultures—one induced with 500 ng/mL aTc and one uninduced control. After overnight induction, serial dilutions were plated on selective media with or without aTc to quantify survival rates. Potential knockout mutants were screened by PCR using primers flanking the target region, with successful deletions identified by the appearance of a smaller amplicon compared to wild-type. The deletion boundaries were confirmed by Sanger sequencing of PCR products. The pQL033-X-sg plasmid was cured by streaking mutant cells onto 7H11/OADC agar supplemented with 5% sucrose in the absence of antibiotics. To remove the pCas9-mScarlet (or pCas9-mScarlet-noint) plasmid from mutant strains, cells were electroporated with plasmid pQL027 (ZeoR, sacB), which expresses the L5 integrase and excisionase (L5gp36). Transformants were selected on 7H11/OADC agar containing 20 μg/mL zeocin. White colonies indicated loss of the pCas9-mScarlet (or pCas9-mScarlet-noint) plasmid. The pQL027 plasmid was subsequently cured by streaking cells onto 7H11/OADC agar supplemented with 5% sucrose in the absence of antibiotics.

To establish a CRISPR-based gene knockout system in M. abscessus, we engineered a dual-plasmid system comprising a Cas9 expression vector (pCas9-mScarlet) and a dual-sgRNA expressing plasmid (pKMZeoR-sg). The pCas9-mScarlet construct integrates into the L5 attB locus of the M. abscessus genome and carries an anhydrotetracycline (aTc)-inducible Sth1Cas9 gene, a kanamycin resistance marker, and an mScarletfluorescent reporter for visual selection of transformants (Figures 1A,B). Following electroporation into wild-type M. abscessus, approximately 30% of kanamycin-resistant colonies lacked fluorescence, indicating a high false-positive rate on kanamycin plates.

The sgRNA expression vector (pKMZeoR-sg) is a replicative plasmid conferring zeocin resistance, with sgRNA transcription under tetracycline repressor control. To validate whether dual-sgRNA design could mediate gene knockout in M. abscessus, we targeted the Mab_0673-Mab_0674 (phoP/phoR) gene cluster which is of particular interest due to its potential role in virulence regulation, designing two sgRNAs flanking the locus (Supplementary Figures S2A,B). Following electroporation into M. abscessus harboring pCas9-mScarlet and aTc induction, no knockout mutants were detected among 69 colonies screened (Supplementary Figure S2C). Sequencing of PCR amplicons from five clones confirmed incomplete cleavage, suggesting inefficient editing. We hypothesized that tandem arrangement of the two sgRNA cassettes might promote intermolecular or intramolecular recombination, leading to sgRNA cassette loss. Indeed, PCR and sequencing of the sgRNA region in three randomly selected clones confirmed extensive deletions in the sgRNA cassettes (Supplementary Figure S2D).

To overcome this limitation, we redesigned the sgRNA expression vector (pQL033-Mab_0673/0674sg), by placing the two sgRNA cassettes either side of the ZeoR resistance gene (Supplementary Figures S1, S2E). Electroporation of this modified construct into M. abscessus carrying pCas9-mScarlet resulted in significantly improved editing efficiency. PCR analysis 72 colonies showed that 52 colonies had successful Mab_0673/0674 deletion, 8 lacked deletions, 1 exhibited partial deletion, and 11 produced no PCR amplicon—suggesting larger-than-expected genomic deletions (Supplementary Figure S2F). These findings establish dual-sgRNA CRISPR/Cas9 as an effective strategy for precise gene deletion in M. abscessus.

To systematically assess the gene deletion efficiency of our dual-sgRNA CRISPR–Cas9 system in M. abscessus, we targeted four additional loci (Ms1 ncRNA, nucS, Mab_2999c, and Mab_2300/2301) distributed across different genomic regions and varying in size (220 bp to 3.5 kb; Figures 2A,D,G,J). Deletion efficiency was consistently high but varied based on genomic context rather than fragment size.

For Ms1ncRNA and nucS (~200 bp and ~480 bp deletions, respectively), correct knockout rates reached 96% (22/23 clones each) with sequencing confirming precise edits (Figures 2B–F). In contrast, the ~400-bp Mab_2999c deletion showed a lower success rate (65%, 15/23 clones), with some exhibiting additional deletions (+140 bp or +264 bp) or loss of PCR amplification (Figures 2G–I). Similarly, the ~3.5 kb deletion of Mab_2300/2301 was successful in 71% (17/24) of clones, though some exhibited larger deletions (+269 bp) or CRISPR escape events (Figures 2J–L). These results confirm the robustness of the dual-sgRNA system while highlighting potential context-dependent variability in editing efficiency.

During attempts to delete the methionine synthase gene metH, the cyclic di-AMP riboswitch (cdAMPRibo, the promoter region of MAB_0869c), and SRP_RNA coding sequence in M. abscessus, we observed high rates of pCas9-mScarlet plasmid loss following aTc induction, with white colonies appearing on selective plates (57.1, 67.8, and 20.7%, respectively) (Figure 3A). PCR verification of 23 red colonies revealed no correct knockouts for metH and SRP_RNA, while only 11/23 clones showed successful cdAMPRibo deletion (Figures 3B–D).

Although L5 integration vectors are generally stable in M. smegmatis (Saviola, 2009), rare excision events can occur in the presence of L5 integrase. To prevent plasmid loss, we constructed pCas9-mScarlet-noint by removing the L5 integrase gene. Co-electroporation with pBSint (expressing transient L5 integrase) enabled integration of pCas9-mScarlet-noint without persistent integrase expression. This strategy significantly reduced plasmid loss, eliminating white colonies in metH and SRP_RNA knockouts and reducing loss to 15.5% for cdAMPRibo (Figure 3A). PCR verification of randomly selected red surviving clones showed markedly improved knockout efficiency for metH and cdAMPRibo, reaching 87% (20/23) and 73.9% (17/23) respectively (Figures 3B,C), demonstrating the importance of stabilizing Cas9 expression in some M. abscessus genes editing.

To evaluate the system’s capability for large-scale deletions, we targeted the mps1-mps2 glycopeptidolipid biosynthetic cluster (18.1 kb), a non-essential region for in vitro growth (Figure 4A). Following aTc induction, all surviving clones exhibited rough colony morphology, indicative of mps1/mps2 inactivation (Figure 4C). PCR screening of 23 rough colonies confirmed successful deletions in 43.5% (10/23), with sequencing verifying an average excision of 16.7 kb (Figures 4B,D). One clone exhibited an additional 370 bp deletion, while eight yielded no PCR products, suggesting larger deletions affecting primer binding sites. Notably, four clones retained wild-type PCR band sizes, and sequencing revealed small deletions near sgRNA target sites (Figure 4D). These findings establish the efficacy of dual-sgRNA CRISPR/Cas9 for ultra-large fragment deletions in M. abscessus.

Msp-like porins serve as the principal hydrophilic diffusion channels in the outer membrane of rapidly growing mycobacteria (Niederweis, 2003). Partial or complete deletions of the corresponding genes are recurrent in clinical M. abscessus isolates and have been linked to increased virulence (Everall et al., 2017; de Moura et al., 2021). de Moura et al. (2021) individually disrupted the two msp homologues, mmpA (Mab_1080) and mmpB (Mab_1081), in strain CIP108297. Both ∆mmpA and ∆mmpB mutants grew more slowly in vitro than wild type, consistent with a role in nutrient uptake, and each mutant exhibited heightened pathogenicity in SCID mice. Repeated attempts to generate a double knockout with Che9c RecET recombineering failed, leading those authors to propose that at least one functional mmp gene is required for in-vitro viability.

Taking advantage of the high efficiency of our dual-sgRNA CRISPR–Cas9 system, we targeted the two adjacent msp-like outer membrane channel genes, Mab_1080 and Mab_1081, for simultaneous deletion in M. abscessus strain ATCC 19977. Unexpectedly, we were able to isolate double-knockout (ΔMab_1080ΔMab_1081) mutants on 7H11/OADC agar supplemented with 500 ng/μl aTc (Figures 5A–C). The double-knockout strain exhibited markedly slower growth than the wild type on solid medium, forming visibly smaller colonies over the same incubation period. In 7H9/OADC broth, ΔMab_1080ΔMab_1081 mutants displayed a pronounced growth delay relative to wild-type cells. Complementation with either Mab_1080 or Mab_1081, expressed from the P_hsp60 promoter via the integrative plasmid pMV306, fully restored growth to wild-type levels (Figure 5D). In LB broth, wild-type cells proliferated robustly, whereas the double mutant showed almost no growth; complementation with Mab_1080 fully rescued proliferation, while Mab_1081 afforded only partial rescue (Figure 5E).

The ability of the ΔMab_1080ΔMab_1081 strain to grow in 7H9/OADC medium, albeit at a reduced rate, indicates that Mab_1080 and Mab_1081 are not essential under these conditions. This finding also points to the likely presence of additional outer membrane channel proteins in M. abscessus, at least in the ATCC 19977 strain. More broadly, the successful generation of this double knockout underscores the high efficiency of our dual-sgRNA CRISPR system in disrupting genes that strongly impair in vitro growth—mutants that are typically difficult to isolate using conventional recombineering approaches.