Mycobacterium tuberculosis and Mycobacterium marinum non-homologous end-joining proteins can function together to join DNA ends in Escherichia coli

Comparative proteomic investigation unravels the pathobiology of Mycobacterium fortuitum biofilm

Biofilm formation by Mycobacterium fortuitum causes serious threats to human health due to its increased contribution to nosocomial infections. In this study, the first comprehensive global proteome analysis of M. fortuitum was reported under planktonic and biofilm growth states. A label-free Q Exactive Quadrupole-Orbitrap tandem mass spectrometry analysis was performed on the protein lysates. The differentially abundant proteins were functionally characterized and re-annotated using Blast2GO and CELLO2GO. Comparative analysis of the proteins among two growth states provided insights into the phenotypic switch, and fundamental pathways associated with pathobiology of M. fortuitum biofilm, such as lipid biosynthesis and quorum-sensing. Interaction network generated by the STRING database revealed associations between proteins that endure M. fortuitum during biofilm growth state. Hypothetical proteins were also studied to determine their functional alliance with the biofilm phenotype. CARD, VFDB, and PATRIC analysis further showed that the proteins upregulated in M. fortuitum biofilm exhibited antibiotic resistance, pathogenesis, and virulence. Heatmap and correlation analysis provided the biomarkers associated with the planktonic and biofilm growth of M. fortuitum. Proteome data was validated by qPCR analysis. Overall, the study provides insights into previously unexplored biochemical pathways that can be targeted by novel inhibitors, either for shortened treatment duration or for eliminating biofilm of M. fortuitum and related nontuberculous mycobacterial pathogens. KEY POINTS: • Proteomic analyses of M. fortuitum reveals novel biofilm markers. • Acetyl-CoA acetyltransferase acts as the phenotype transition switch. • The study offers drug targets to combat M. fortuitum biofilm infections.

CRISPR-Cas12a-Assisted Genome Editing in Amycolatopsis mediterranei

Amycolatopsis mediterranei U32 is an industrial producer of rifamycin SV, whose derivatives have long been the first-line antimycobacterial drugs. In order to perform genetic modification in this important industrial strain, a lot of efforts have been made in the past decades and a homologous recombination-based method was successfully developed in our laboratory, which, however, requires the employment of an antibiotic resistance gene for positive selection and did not support convenient markerless gene deletion. Here in this study, the clustered regularly interspaced short palindromic repeat (CRISPR) system was employed to establish a genome editing system in A. mediterranei U32. Specifically, the Francisella tularensis subsp. novicida Cas12a (FnCas12a) gene was first integrated into the U32 genome to generate target-specific double-stranded DNA (dsDNA) breaks (DSBs) under the guidance of CRISPR RNAs (crRNAs). Then, the DSBs could be repaired by either the non-homologous DNA end-joining (NHEJ) system or the homology-directed repair (HDR) pathway, generating inaccurate or accurate mutations in target genes, respectively. Besides of A. mediterranei, the present work may also shed light on the development of CRISPR-assisted genome editing systems in other species of the Amycolatopsis genus.

CRISPR-Cas9-assisted native end-joining editing offers a simple strategy for efficient genetic engineering in Escherichia coli

Unlike eukaryotes, prokaryotes are less proficient in homologous recombination (HR) and non-homologous end-joining (NHEJ). All existing genomic editing methods for Escherichia coli (E. coli) rely on exogenous HR or NHEJ systems to repair DNA double-strand breaks (DSBs). Although an E. coli native end-joining (ENEJ) system has been reported, its potential in genetic engineering has not yet been explored. Here, we present a CRISPR-Cas9-assisted native end-joining editing and show that ENEJ-dependent DNA repair can be used to conduct rapid and efficient deletion of chromosome fragments up to 83 kb or gene inactivation. Moreover, the positive rate and editing efficiency are independent of high-efficiency competent cells. The method requires neither exogenous DNA repair systems nor introduced editing template. The Cas9-sgRNA complex is the only foreign element in this method. This study is the first successful engineering effort to utilize ENEJ mechanism in genomic editing and provides an effective strategy for genetic engineering in bacteria that are inefficient in HR and NHEJ.

Single-Strand Annealing Plays a Major Role in Double-Strand DNA Break Repair following CRISPR-Cas9 Cleavage in Leishmania

CRISPR-Cas9 genome editing relies on an efficient double-strand DNA break (DSB) and repair. Contrary to mammalian cells, the protozoan parasite Leishmania lacks the most efficient nonhomologous end-joining pathway and uses microhomology-mediated end joining (MMEJ) and, occasionally, homology-directed repair to repair DSBs. Here, we reveal that Leishmania predominantly uses single-strand annealing (SSA) (>90%) instead of MMEJ (<10%) for DSB repair (DSBR) following CRISPR targeting of the miltefosine transporter gene, resulting in 9-, 18-, 20-, and 29-kb sequence deletions and multiple gene codeletions. Strikingly, when targeting the Leishmania donovani LdBPK_241510 gene, SSA even occurred by using direct repeats 77 kb apart, resulting in the codeletion of 15 Leishmania genes, though with a reduced frequency. These data strongly indicate that DSBR is not efficient in Leishmania, which explains why more than half of DSBs led to cell death and why the CRISPR gene-targeting efficiency is low compared with that in other organisms. Since direct repeat sequences are widely distributed in the Leishmania genome, we predict that many DSBs created by CRISPR are repaired by SSA. It is also revealed that DNA polymerase theta is involved in both MMEJ and SSA in Leishmania Collectively, this study establishes that DSBR mechanisms and their competence in an organism play an important role in determining the outcome and efficacy of CRISPR gene targeting. These observations emphasize the use of donor DNA templates to improve gene editing specificity and efficiency in Leishmania In addition, we developed a novel Staphylococcus aureus Cas9 constitutive expression vector (pLdSaCN) for gene targeting in Leishmania IMPORTANCE Due to differences in double-strand DNA break (DSB) repair mechanisms, CRISPR-Cas9 gene editing efficiency can vary greatly in different organisms. In contrast to mammalian cells, the protozoan parasite Leishmania uses microhomology-mediated end joining (MMEJ) and, occasionally, homology-directed repair (HDR) to repair DSBs but lacks the nonhomologous end-joining pathway. Here, we show that Leishmania predominantly uses single-strand annealing (SSA) instead of MMEJ for DSB repairs (DSBR), resulting in large deletions that can include multiple genes. This strongly indicates that the overall DSBR in Leishmania is inefficient and therefore can influence the outcome of CRISPR-Cas9 gene editing, highlighting the importance of using a donor DNA to improve gene editing fidelity and efficiency in Leishmania.

Multiple Stepwise Gene Knockout Using CRISPR/Cas9 in Escherichia coli

With the recent implementation of the CRISPR/Cas9 technology as a standard tool for genome editing, laboratories all over the world are undergoing one of the biggest advancements in molecular biology since PCR. The key advantage of this method is its simplicity and universal applicability for species of any phylum. Of particular interest is the extensively studied Gram-negative bacterium Escherichia coli, as it is considered as the workhorse for both research and industrial purposes. Here, we present a simple, robust and effective protocol using the CRISPR/Cas9 system in combination with the λ Red machinery for gene knockout in E. coli. Crucial in our procedure is the use of a double-stranded donor DNA and a curing strategy for removal of the guide RNA encoding plasmid that allows starting a new mutation after only two working days. Our protocol allows multiple, stepwise gene knockout strains with high mutagenesis efficiencies applicable for high-throughput approaches.

Evidence for double-strand break mediated mitochondrial DNA replication in Saccharomyces cerevisiae

The mechanism of mitochondrial DNA (mtDNA) replication in Saccharomyces cerevisiae is controversial. Evidence exists for double-strand break (DSB) mediated recombination-dependent replication at mitochondrial replication origin ori5 in hypersuppressive ρ⁻ cells. However, it is not clear if this replication mode operates in ρ⁺ cells. To understand this, we targeted bacterial Ku (bKu), a DSB binding protein, to the mitochondria of ρ⁺ cells with the hypothesis that bKu would bind persistently to mtDNA DSBs, thereby preventing mtDNA replication or repair. Here, we show that mitochondrial-targeted bKu binds to ori5 and that inducible expression of bKu triggers petite formation preferentially in daughter cells. bKu expression also induces mtDNA depletion that eventually results in the formation of ρ⁰ cells. This data supports the idea that yeast mtDNA replication is initiated by a DSB and bKu inhibits mtDNA replication by binding to a DSB at ori5, preventing mtDNA segregation to daughter cells. Interestingly, we find that mitochondrial-targeted bKu does not decrease mtDNA content in human MCF7 cells. This finding is in agreement with the fact that human mtDNA replication, typically, is not initiated by a DSB. Therefore, this study provides evidence that DSB-mediated replication is the predominant form of mtDNA replication in ρ⁺ yeast cells.

Large scale validation of an efficient CRISPR/Cas-based multi gene editing protocol in Escherichia coli

BACKGROUND

The exploitation of the CRISPR/Cas9 machinery coupled to lambda (λ) recombinase-mediated homologous recombination (recombineering) is becoming the method of choice for genome editing in E. coli. First proposed by Jiang and co-workers, the strategy has been subsequently fine-tuned by several authors who demonstrated, by using few selected loci, that the efficiency of mutagenesis (number of mutant colonies over total number of colonies analyzed) can be extremely high (up to 100%). However, from published data it is difficult to appreciate the robustness of the technology, defined as the number of successfully mutated loci over the total number of targeted loci. This information is particularly relevant in high-throughput genome editing, where repetition of experiments to rescue missing mutants would be impractical. This work describes a "brute force" validation activity, which culminated in the definition of a robust, simple and rapid protocol for single or multiple gene deletions.

RESULTS

We first set up our own version of the CRISPR/Cas9 protocol and then we evaluated the mutagenesis efficiency by changing different parameters including sequence of guide RNAs, length and concentration of donor DNAs, and use of single stranded and double stranded donor DNAs. We then validated the optimized conditions targeting 78 "dispensable" genes. This work led to the definition of a protocol, featuring the use of double stranded synthetic donor DNAs, which guarantees mutagenesis efficiencies consistently higher than 10% and a robustness of 100%. The procedure can be applied also for simultaneous gene deletions.

CONCLUSIONS

This work defines for the first time the robustness of a CRISPR/Cas9-based protocol based on a large sample size. Since the technical solutions here proposed can be applied to other similar procedures, the data could be of general interest for the scientific community working on bacterial genome editing and, in particular, for those involved in synthetic biology projects requiring high throughput procedures.