A Highly Efficient CRISPR-Cas9-Mediated Large Genomic Deletion in Bacillus subtilis

Targeted knockout and plasmid-based transfer of NRPS/PKS for improving lipopeptide iturin A synthesis

The nonribosomal peptide synthetase (NRPS)/polyketide synthase (PKS) assembly lines are a large enzymatic machinery that facilitates the transfer and synthesis of intermediates between multimodular megasynthases through complex protein-protein interactions. Although NRPS/PKS systems display a highly sophisticated biosynthetic pathway, a similar strategy for the holistic editing of these gene clusters has not been described. In this study, practical gene-editing tools were developed for long gene clusters, allowing the efficient knockout of entire NRPS/PKS gene clusters. In addition, a strategy for the genome cleavage and plasmid capture of NRPS/PKS genes was developed. Using iturin A synthesis as a case study, the competing lipopeptide biosynthesis gene cluster was knocked out, and the NRPS/PKS gene cluster responsible for its production were transferred into a plasmid. The engineered strain exhibited significantly enhanced expression of iturin A NRPS/PKS genes and increased production of iturin A. This approach facilitates the optimization of NRPS/PKS systems and other long-gene cluster-derived natural product biosynthesis.

Exploring the Probiotic Potential of Bacteroides spp. Within One Health Paradigm

Probiotics are pivotal in maintaining or restoring the balance of human intestinal microbiota, a crucial factor in mitigating diseases and preserving the host's health. Exploration into Bacteroides spp. reveals substantial promise in their development as next-generation probiotics due to their profound interaction with host immune cells and capability to regulate the microbiome's metabolism by significantly impacting metabolite production. These beneficial bacteria exhibit potential in ameliorating various health issues such as intestinal disorders, cardiovascular diseases, behavioral disorders, and even cancer. Though it's important to note that a high percentage of them are as well opportunistic pathogens, posing risks under certain conditions. Studies highlight their role in modifying immune responses and improving health conditions by regulating lymphocytes, controlling metabolism, and preventing inflammation and cancer. The safety and efficacy of Bacteroides strains are currently under scrutiny by the European Commission for authorization in food processing, marking a significant step towards their commercialization. The recent advancements in bacterial isolation and sequencing methodologies, coupled with the integration of Metagenome-Assembled Genomes (MAGs) binning from metagenomics data, continue to unveil the potential of Bacteroides spp., aiding in the broader understanding and application of these novel probiotics in health and disease management.

Improving Bacillus subtilis as Biological Chassis Performance by the CRISPR Genetic Toolkit

Bacillus subtilis is the model Gram-positive and industrial chassis bacterium; it has blossomed as a robust and promising host for enzyme, biochemical, or bioflocculant production. However, synthetic biology and metabolic engineering technologies of B. subtilis have lagged behind the most widely used industrial chassis Saccharomyces cerevisiae and Escherichia coli. CRISPR (an acronym for clustered regularly interspaced short palindromic repeats) enables efficient, site-specific, and programmable DNA cleavage, which has revolutionized the manner of genome editing. In 2016, CRISPR technology was first introduced into B. subtilis and has been intensely upgraded since then. In this Review, we discuss recently developed key additions to CRISPR toolkit design in B. subtilis with gene editing, transcriptional regulation, and enzyme modulation. Second, advances in the B. subtilis chassis of efficient biochemicals and proteins with CRISPR engineering are discussed. Finally, we conclude with perspectives on the challenges and opportunities of CRISPR-based biotechnology in B. subtilis, wishing that B. subtilis can be comparable to traditional industrial microorganisms such as E. coli and S. cerevisiae someday soon.

Bacterial genome reduction for optimal chassis of synthetic biology: a review

Bacteria with streamlined genomes, that harbor full functional genes for essential metabolic networks, are able to synthesize the desired products more effectively and thus have advantages as production platforms in industrial applications. To obtain streamlined chassis genomes, a large amount of effort has been made to reduce existing bacterial genomes. This work falls into two categories: rational and random reduction. The identification of essential gene sets and the emergence of various genome-deletion techniques have greatly promoted genome reduction in many bacteria over the past few decades. Some of the constructed genomes possessed desirable properties for industrial applications, such as: increased genome stability, transformation capacity, cell growth, and biomaterial productivity. The decreased growth and perturbations in physiological phenotype of some genome-reduced strains may limit their applications as optimized cell factories. This review presents an assessment of the advancements made to date in bacterial genome reduction to construct optimal chassis for synthetic biology, including: the identification of essential gene sets, the genome-deletion techniques, the properties and industrial applications of artificially streamlined genomes, the obstacles encountered in constructing reduced genomes, and the future perspectives.

Dual-sgRNA CRISPRa System for Enhanced MK-7 Production and Salmonella Infection Mitigation in Bacillus subtilis natto Applied to Caco-2 Cells

This study optimized the menaquinone-7 (MK-7) synthetic pathways in Bacillus subtilis (B. subtilis) natto NB205, a strain that originated from natto, to enhance its MK-7 production. Utilizing mutation breeding, we developed NBMK308, a mutant strain that demonstrated a significant 117.23% increase in MK-7 production. A comprehensive transcriptome analysis identified two key genes, ispA and ispE, as being critical in MK-7 synthesis. The dual-sgRNA CRISPRa system was utilized to achieve precise regulation of ispA and ispE in the newly engineered strain, A3E3. This strategic modulation resulted in a significant enhancement of MK-7 production, achieving increases of 20.02% and 201.41% compared to traditional overexpression systems and the original strain NB205, respectively. Furthermore, the fermentation supernatant from A3E3 notably inhibited Salmonella invasion in Caco-2 cells, showcasing its potential for combating such infections. The safety of the dual-sgRNA CRISPRa system was confirmed through cell assays. The utilization of the dual-sgRNA CRISPRa system in this study was crucial for the precise regulation of key genes in MK-7 synthesis, leading to a remarkable increase in production and demonstrating additional therapeutic potential in inhibiting pathogenic infections. This approach effectively combined the advantages of microbial fermentation and biotechnology, addressing health and nutritional challenges.

CRISPR genetic toolkits of classical food microorganisms: Current state and future prospects

Production of food-related products using microorganisms in an environmentally friendly manner is a crucial solution to global food safety and environmental pollution issues. Traditional microbial modification methods rely on artificial selection or natural mutations, which require time for repeated screening and reproduction, leading to unstable results. Therefore, it is imperative to develop rapid, efficient, and precise microbial modification technologies. This review summarizes recent advances in the construction of gene editing and metabolic regulation toolkits based on the clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated proteins (CRISPR-Cas) systems and their applications in reconstructing food microorganism metabolic networks. The development and application of gene editing toolkits from single-site gene editing to multi-site and genome-scale gene editing was also introduced. Moreover, it presented a detailed introduction to CRISPR interference, CRISPR activation, and logic circuit toolkits for metabolic network regulation. Moreover, the current challenges and future prospects for developing CRISPR genetic toolkits were also discussed.

CRISPR-Cas-Based Engineering of Probiotics

Probiotics are the treasure of the microbiology fields. They have been widely used in the food industry, clinical treatment, and other fields. The equivocal health-promoting effects and the unknown action mechanism were the largest obstacles for further probiotic's developed applications. In recent years, various genome editing techniques have been developed and applied to explore the mechanisms and functional modifications of probiotics. As important genome editing tools, CRISPR-Cas systems that have opened new improvements in genome editing dedicated to probiotics. The high efficiency, flexibility, and specificity are the advantages of using CRISPR-Cas systems. Here, we summarize the classification and distribution of CRISPR-Cas systems in probiotics, as well as the editing tools developed on the basis of them. Then, we discuss the genome editing of probiotics based on CRISPR-Cas systems and the applications of the engineered probiotics through CRISPR-Cas systems. Finally, we proposed a design route for CRISPR systems that related to the genetically engineered probiotics.

Antimicrobial peptides from Bacillus spp. and strategies to enhance their yield

Antibiotic resistance is a growing concern that is affecting public health globally. The search for alternative antimicrobial agents has become increasingly important. Antimicrobial peptides (AMPs) produced by Bacillus spp. have emerged as a promising alternative to antibiotics, due to their broad-spectrum antimicrobial activity against resistant pathogens. In this review, we provide an overview of Bacillus-derived AMPs, including their classification into ribosomal (bacteriocins) and non-ribosomal peptides (lipopeptides and polyketides). Additionally, we delve into the molecular mechanisms of AMP production and describe the key biosynthetic gene clusters involved. Despite their potential, the low yield of AMPs produced under normal laboratory conditions remains a challenge to large-scale production. This review thus concludes with a comprehensive summary of recent studies aimed at enhancing the productivity of Bacillus-derived AMPs. In addition to medium optimization and genetic manipulation, various molecular strategies have been explored to increase the production of recombinant antimicrobial peptides (AMPs). These include the selection of appropriate expression systems, the engineering of expression promoters, and metabolic engineering. Bacillus-derived AMPs offer great potential as alternative antimicrobial agents, and this review provides valuable insights on the strategies to enhance their production yield, which may have significant implications for combating antibiotic resistance. KEY POINTS: • Bacillus-derived AMP is a potential alternative therapy for resistant pathogens • Bacillus produces two main classes of AMPs: ribosomal and non-ribosomal peptides • AMP yield can be enhanced using culture optimization and molecular approaches.

A recombineering system for Bacillus subtilis based on the native phage recombinase pair YqaJ/YqaK

Bacillus subtilis plays an important role in fundamental and applied research, and it has been widely used as a cell factory for the production of enzymes, antimicrobial materials, and chemicals for agriculture, medicine, and industry. However, genetic manipulation tools for B. subtilis have low efficiency. In this work, our goal was to develop a simple recombineering system for B. subtilis. We showed that genome editing can be achieved in B. subtiliis 1A751 through co-expression of YqaJ/YqaK, a native phage recombinase pair found in B. subtilis 168, and the competence master regulator ComK using a double-stranded DNA substrate with short homology arms (100 bp) and a phosphorothioate modification at the 5'-end. Efficient gene knockouts and large DNA insertions were achieved using this new recombineering system in B. subtilis 1A751. As far as we know, this is the first recombineering system using the native phage recombinase pair YqaJ/YqaK in B. subtilis. In conclusion, this new recombineering system provides a simple and fast tool for genetic manipulation of B. subtilis, and it will promote studies of genome function, construction of production strains, and genome mining in B. subtilis.

A Programmable CRISPR/Cas9 Toolkit Improves Lycopene Production in Bacillus subtilis

Bacillus subtilis has been widely used and generally recognized as a safe host for the production of recombinant proteins, high-value chemicals, and pharmaceuticals. Thus, its metabolic engineering attracts significant attention. Nevertheless, the limited availability of selective markers makes this process difficult and time-consuming, especially in the case of multistep biosynthetic pathways. Here, we employ CRISPR/Cas9 technology to build an easy cloning toolkit that addresses commonly encountered obstacles in the metabolic engineering of B. subtilis, including the chromosomal integration locus, promoter, terminator, and guide RNA (gRNA) target. Six promoters were characterized, and the promoter strengths ranged from 0.9- to 23-fold that of the commonly used strong promoter P43. We characterized seven terminators in B. subtilis, and the termination efficiencies (TEs) of the seven terminators are all more than 90%. Six gRNA targets were designed upstream of the promoter and downstream of the terminator. Using a green fluorescent protein (GFP) reporter, we confirmed integration efficiency with the single-locus integration site is up to 100%. We demonstrated the applicability of this toolkit by optimizing the expression of a challenging but industrially important product, lycopene. By heterologous expression of the essential genes for lycopene synthesis on the B. subtilis genome, a total of 13 key genes involved in the lycopene biosynthetic pathway were manipulated. Moreover, our findings showed that the gene cluster ispG-idi-dxs-ispD could positively affect the production of lycopene, while the cluster dxr-ispE-ispF-ispH had a negative effect on lycopene production. Hence, our multilocus integration strategy can facilitate the pathway assembly for production of complex chemicals and pharmaceuticals in B. subtilis. IMPORTANCE We present a toolkit that allows for rapid cloning procedures and one-step subcloning to move from plasmid-based expression to stable chromosome integration and expression in a production strain in less than a week. The utility of the customized tool was demonstrated by integrating the MEP (2C-methyl-d-erythritol-4-phosphate) pathway, part of the pentose phosphate pathway (PPP), and the hetero-lycopene biosynthesis genes by stable expression in the genome. The tool could be useful to engineer B. subtilis strains through diverse recombination events and ultimately improve its potential and scope of industrial application as biological chassis.

Improvement of Bacilysin Production in Bacillus subtilis by CRISPR/Cas9-Mediated Editing of the 5'-Untranslated Region of the bac Operon

Bacilysin is a dipeptide antibiotic composed of L-alanine and L-anticapsin produced by certain strains of Bacillus subtilis. Bacilysin is gaining increasing attention in industrial agriculture and pharmaceutical industries due to its potent antagonistic effects on various bacterial, fungal, and algal pathogens. However, its use in industrial applications is hindered by its low production in the native producer. The biosynthesis of bacilysin is mainly based on the bacABCDEF operon. Examination of the sequence surrounding the upstream of the bac operon did not reveal a clear, strong ribosome binding site (RBS). Therefore, in this study, we aimed to investigate the impact of RBS as a potential route to improve bacilysin production. For this, the 5' untranslated region (5'UTR) of the bac operon was edited using the CRISPR/Cas9 approach by introducing a strong ribosome binding sequence carrying the canonical Shine-Dalgarno sequence (TAAGGAGG) with an 8 nt spacing from the AUG start codon. Strong RBS substitution resulted in a 2.87-fold increase in bacilysin production without affecting growth. Strong RBS substitution also improved the mRNA stability of the bac operon. All these data revealed that extensive RBS engineering is a promising key option for enhancing bacilysin production in its native producers.

Simplified Construction of Engineered Bacillus subtilis Host for Improved Expression of Proteins Harboring Noncanonical Amino Acids

The UAG-based genetic code expansion (GCE) enables site-specific incorporation of noncanonical amino acids (ncAAs) harboring novel chemical functionalities in specific target proteins. However, most GCE studies were done in several whole-genome engineered chassis cells whose hundreds of UAG stop codons were systematically edited to UAA to avoid readthrough in protein synthesis in the presence of GCE. The huge workload of removing all UAG limited the application of GCE in other microbial cell factories (MCF) such as Bacillus subtilis, which has 607 genes ended with UAG among its 4245 coding genes. Although the 257 essential genes count only 6.1% of the genes in B. subtilis, they transcribe 12.2% of the mRNAs and express 52.1% of the proteins under the exponential phase. Here, we engineered a strain named Bs-22 in which all 22 engineerable UAG stop codons in essential genes were edited to UAA via CRISPR/Cas9-mediated multiple-site engineering to minimize the negative effect of GCE on the expression of essential genes. Besides the process of constructing GCE-compatible B. subtilis was systematically optimized. Compared with wild-type B. subtilis (Bs-WT), the fluorescence signal of the eGFP expression could enhance 2.25-fold in Bs-22, and the production of protein tsPurple containing l-(7-hydroxycoumarin-4-yl) ethylglycine (Cou) was increased 2.31-fold in Bs-22. We verified that all purified tsPurple proteins from Bs-22 contained Cou, indicating the excellent fidelity of the strategy. This proof-of-concept study reported efficient overexpression of ncAA-rich proteins in MCF with minimized engineering, shedding new light on solving the trade-off between efficiency and workload.

Barriers to simultaneous multilocus integration in Bacillus subtilis tumble down: development of a straightforward screening method for the colorimetric detection of one-step multiple gene insertion using the CRISPR-Cas9 system

BACKGROUND

Despite recent advances in genetic engineering tools for effectively regulating and manipulating genes, efficient simultaneous multigene insertion methods have not been established in Bacillus subtilis. To date, multilocus integration systems in B. subtilis, which is one of the main industrial enzyme producers and a GRAS (generally regarded as safe) microbial host, rely on iterative rounds of plasmid construction for sequential insertions of genes into the B. subtilis chromosome, which is tedious and time consuming.

RESULTS

In this study, we present development and proof-of-concept of a novel CRISPR-Cas9-based genome-editing strategy for the colorimetric detection of one-step multiple gene insertion in B. subtilis. First, up to three copies of the crtMN operon from Staphylococcus aureus, encoding a yellow pigment, were incorporated at three ectopic sites within the B. subtilis chromosome, rendering engineered strains able to form yellow colonies. Second, a single CRISPR-Cas9-based plasmid carrying a highly specific single guide RNA (sgRNA) targeting crtMN operon and a changeable editing template was constructed to facilitate simultaneous insertion of multiple gene-copies through homology-directed repair (HDR). Upon transformation of engineered strains with engineered plasmids, strains harboring up to three gene copies integrated into the chromosome formed white colonies because of the removal of the crtMN operon, clearly distinguishable from yellow colonies harboring undesired genetic modifications. As a result, construction of a plasmid-less, marker-free, high-expression stable producer B. subtilis strain can be completed in only seven days, demonstrating the potential that the implementation of this technology may bring for biotechnology purposes.

CONCLUSIONS

The novel technology expands the genome-editing toolset for B. subtilis and means a substantial improvement over current methodology, offering new application possibilities that we envision should significantly boost the development of B. subtilis as a chassis in the field of synthetic biology.

In Vivo Genome Editing in Type I and II Methanotrophs Using a CRISPR/Cas9 System

Methanotrophic bacteria are Gram-negative, aerobic organisms that use methane as their sole source of carbon and energy. In this study, we constructed and exemplified a CRISPR/Cas9 genome editing system and used it to successfully make gene deletions and insertions in the type I methanotroph Methylococcus capsulatus Bath and the type II methanotroph Methylocystis parvus OBBP. High frequencies of gene deletions and insertions were achieved in combination with homology-directed repair. In M. parvus OBBP, we also investigated the impact of several parameters on the CRISPR/Cas9 genome editing, where the ligD gene was targeted with various PAM sequences and guide RNA spacer sequences, homology arms of variable length, differences in the duration of mating during conjugation, and exploiting promoters of different strengths to control the expression of cas9 and sgRNA. Although not the first attempt to develop a CRISPR/Cas system in methanotrophs, this work demonstrated for the first time an efficient CRISPR/Cas9 system generating scarless clean gene deletions and insertions in methanotroph genomes.

Bacillus integrative plasmid system combining a synthetic gene circuit for efficient genetic modifications of undomesticated Bacillus strains

BACKGROUND

Owing to CRISPR-Cas9 and derivative technologies, genetic studies on microorganisms have dramatically increased. However, the CRISPR-Cas9 system is still difficult to utilize in many wild-type Bacillus strains owing to Cas9 toxicity. Moreover, less toxic systems, such as cytosine base editors, generate unwanted off-target mutations that can interfere with the genetic studies of wild-type strains. Therefore, a convenient alternative system is required for genetic studies and genome engineering of wild-type Bacillus strains. Because wild-type Bacillus strains have poor transformation efficiencies, the new system should be based on broad-host-range plasmid-delivery systems.

RESULTS

Here, we developed a Bacillus integrative plasmid system in which plasmids without the replication initiator protein gene (rep) of Bacillus are replicated in a donor Bacillus strain by Rep proteins provided in trans but not in Bacillus recipients. The plasmids were transferred to recipients through a modified integrative and conjugative element, which is a wide host range plasmid-delivery system. Genetic mutations were generated in recipients through homologous recombination between the transferred plasmid and the genome. The system was improved by adding a synthetic gene circuit for efficient screening of the desired mutations by double crossover recombination in recipient strains. The improved system exhibited a mutation efficiency of the target gene of approximately 100% in the tested wild-type Bacillus strains.

CONCLUSION

The Bacillus integrative plasmid system developed in this study can generate target mutations with high efficiency when combined with a synthetic gene circuit in wild-type Bacillus strains. The system is free of toxicity and unwanted off-target mutations as it generates the desired mutations by traditional double crossover recombination. Therefore, our system could be a powerful tool for genetic studies and genome editing of Cas9-sensitive wild-type Bacillus strains.

An efficient CRISPR/Cas9-based genome editing system for alkaliphilic Bacillus sp. N16-5 and application in engineering xylose utilization for D-lactic acid production

Alkaliphiles are considered more suitable chassis than traditional neutrophiles due to their excellent resistance to microbial contamination. Alkaliphilic Bacillus sp. N16-5, an industrially interesting strain with great potential for the production of lactic acid and alkaline polysaccharide hydrolases, can only be engineered genetically by the laborious and time-consuming homologous recombination. In this study, we reported the successful development of a CRISPR/Cas9-based genome editing system with high efficiency for single-gene deletion, large gene fragment deletion and exogenous DNA chromosomal insertion. Moreover, based on a catalytically dead variant of Cas9 (dCas9), we also developed a CRISPRi system to efficiently regulate gene expression. Finally, this efficient genome editing system was successfully applied to engineer the xylose metabolic pathway for the efficient bioproduction of D-lactic acid. Compared with the wild-type Bacillus sp. N16-5, the final engineered strain with XylR deletion and AraE overexpression achieved 34.3% and 27.7% increases in xylose consumption and D-lactic acid production respectively. To our knowledge, this is the first report on the development and application of CRISPR/Cas9-based genome editing system in alkaliphilic Bacillus, and this study will significantly facilitate functional genomic studies and genome manipulation in alkaliphilic Bacillus, laying a foundation for the development of more robust microbial chassis.

SURE editing: combining oligo-recombineering and programmable insertion/deletion of selection markers to efficiently edit the Mycoplasma pneumoniae genome

The development of advanced genetic tools is boosting microbial engineering which can potentially tackle wide-ranging challenges currently faced by our society. Here we present SURE editing, a multi-recombinase engineering rationale combining oligonucleotide recombineering with the selective capacity of antibiotic resistance via transient insertion of selector plasmids. We test this method in Mycoplasma pneumoniae, a bacterium with a very inefficient native recombination machinery. Using SURE editing, we can seamlessly generate, in a single step, a wide variety of genome modifications at high efficiencies, including the largest possible deletion of this genome (30 Kb) and the targeted complementation of essential genes in the deletion of a region of interest. Additional steps can be taken to remove the selector plasmid from the edited area, to obtain markerless or even scarless edits. Of note, SURE editing is compatible with different site-specific recombinases for mediating transient plasmid integration. This battery of selector plasmids can be used to select different edits, regardless of the target sequence, which significantly reduces the cloning load associated to genome engineering projects. Given the proven functionality in several microorganisms of the machinery behind the SURE editing logic, this method is likely to represent a valuable advance for the synthetic biology field.

Development and application of CRISPR-based genetic tools in Bacillus species and Bacillus phages

Recently, the clustered regularly interspaced short palindromic repeats (CRISPR) system has been developed into a precise and efficient genome editing tool. Since its discovery as an adaptive immune system in prokaryotes, it has been applied in many different research fields including biotechnology and medical sciences. The high demand for rapid, highly efficient and versatile genetic tools to thrive in bacteria-based cell factories accelerates this process. This review mainly focuses on significant advancements of the CRISPR system in Bacillus subtilis, including the achievements in gene editing, and on problems still remaining. Next, we comprehensively summarize this genetic tool's up-to-date development and utilization in other Bacillus species, including B. licheniformis, B. methanolicus, B. anthracis, B. cereus, B. smithii and B. thuringiensis. Furthermore, we describe the current application of CRISPR tools in phages to increase Bacillus hosts' resistance to virulent phages and phage genetic modification. Finally, we suggest potential strategies to further improve this advanced technique and provide insights into future directions of CRISPR technologies for rendering Bacillus species cell factories more effective and more powerful.

Genome Editing by CRISPR/Cas12 Recognizing AT-Rich PAMs in Shewanella oneidensis MR-1

Homologous recombination-mediated genomic editing is urgently needed to obtain high-performance chassis of electroactive microorganisms. However, the existing tools cannot meet the requirement of genome-wide editing in Shewanella oneidensis. Here, we develop different CRISPR-Cas systems that are ideal to be employed in AT-rich sequences as the supplements to Cas9. AsCpf1 and BhCas12b show low cell toxicity and superior ability to target sequences and are thus screened out in S. oneidensis MR-1. The PAMs of AsCpf1 and BhCas12b are 5'-TTTV-3' and 5'-ATTN-3'. For gene deletion, ∼1-kb gene is knocked out and the editing efficiency is 41.67% by BhCas12b-mediated system. For gene replacement, endogenous promoter of nagK was substituted to a constitutive promoter with the efficiency of 25% through BhCas12b system. For gene insertion, the integration efficiency was up to 94.4% and 83.9% via CRISPR-BhCas12b and AsCpf1 tools. This study implies a great potential of CRISPR-BhCas12b/AsCpf1 systems recognizing AT-rich PAMs for genomic editing in S. oneidensis to facilitate multifaceted gene manipulation.

Bacterial genome reductions: Tools, applications, and challenges

Bacterial cells are widely used to produce value-added products due to their versatility, ease of manipulation, and the abundance of genome engineering tools. However, the efficiency of producing these desired biomolecules is often hindered by the cells’ own metabolism, genetic instability, and the toxicity of the product. To overcome these challenges, genome reductions have been performed, making strains with the potential of serving as chassis for downstream applications. Here we review the current technologies that enable the design and construction of such reduced-genome bacteria as well as the challenges that limit their assembly and applicability. While genomic reductions have shown improvement of many cellular characteristics, a major challenge still exists in constructing these cells efficiently and rapidly. Computational tools have been created in attempts at minimizing the time needed to design these organisms, but gaps still exist in modelling these reductions in silico. Genomic reductions are a promising avenue for improving the production of value-added products, constructing chassis cells, and for uncovering cellular function but are currently limited by their time-consuming construction methods. With improvements to and the creation of novel genome editing tools and in silico models, these approaches could be combined to expedite this process and create more streamlined and efficient cell factories.

Development and application of a rapid all-in-one plasmid CRISPR-Cas9 system for iterative genome editing in Bacillus subtilis

BACKGROUND

Bacillus subtilis, an important industrial microorganism, is commonly used in the production of industrial enzymes. Genome modification is often necessary to improve the production performance of cell. The dual-plasmid CRISPR-Cas9 system suitable for iterative genome editing has been applied in Bacillus subtilis. However, it is limited by the selection of knockout genes, long editing cycle and instability.

RESULTS

To address these problems, we constructed an all-in-one plasmid CRISPR-Cas9 system, which was suitable for iterative genome editing of B. subtilis. The PEG4000-assisted monomer plasmid ligation (PAMPL) method greatly improved the transformation efficiency of B. subtilis SCK6. Self-targeting sgRNA_rep transcription was tightly controlled by rigorous promoter P_acoR, which could induce the elimination of plasmids after genome editing and prepare for next round of genome editing. Our system achieved 100% efficiency for single gene deletions and point mutations, 96% efficiency for gene insertions, and at least 90% efficiency for plasmid curing. As a proof of concept, two extracellular protease genes epr and bpr were continuously knocked out using this system, and it only took 2.5 days to complete one round of genome editing. The engineering strain was used to express Douchi fibrinolytic enzyme DFE27, and its extracellular enzyme activity reached 159.5 FU/mL.

CONCLUSIONS

We developed and applied a rapid all-in-one plasmid CRISPR-Cas9 system for iterative genome editing in B. subtilis, which required only one plasmid transformation and curing, and accelerated the cycle of genome editing. To the best of our knowledge, this is the rapidest iterative genome editing system for B. subtilis. We hope that the system can be used to reconstruct the B. subtilis cell factory for the production of various biological molecules.

Elicitation of Innate Immunity by a Bacterial Volatile 2-Nonanone at Levels below Detection Limit in Tomato Rhizosphere

Bacterial volatile compounds (BVCs) exert beneficial effects on plant protection both directly and indirectly. Although BVCs have been detected in vitro, their detection in situ remains challenging. The purpose of this study was to investigate the possibility of BVCs detection under in situ condition and estimate the potentials of in situ BVC to plants at below detection limit. We developed a method for detecting BVCs released by the soil bacteria Bacillus velezensis strain GB03 and Streptomyces griseus strain S4-7 in situ using solid-phase microextraction coupled with gas chromatography-mass spectrometry (SPME-GC-MS). Additionally, we evaluated the BVC detection limit in the rhizosphere and induction of systemic immune response in tomato plants grown in the greenhouse. Two signature BVCs, 2-nonanone and caryolan-1-ol, of GB03 and S4-7 respectively were successfully detected using the soil-vial system. However, these BVCs could not be detected in the rhizosphere pretreated with strains GB03 and S4-7. The detection limit of 2-nonanone in the tomato rhizosphere was 1 µM. Unexpectedly, drench application of 2-nonanone at 10 nM concentration, which is below its detection limit, protected tomato seedlings against Pseudomonas syringae pv. tomato. Our finding highlights that BVCs, including 2-nonanone, released by a soil bacterium are functional even when present at a concentration below the detection limit of SPME-GC-MS.

Development of an efficient iterative genome editing method in Bacillus subtilis using the CRISPR-AsCpf1 system

Bacillus subtilis is a useful chassis in the fields of synthetic biology and metabolic engineering for chemical production. Here, we constructed CRISPR-AsCpf1-based expression plasmids with the temperature-sensitive replicon for iterative genome editing in B. subtilis. This method allowed gene insertion and large genomic deletion with an editing efficiency of up 80%-100% and rapid plasmid curing to facilitate the iterative genome editing in B. subtilis 168. Using the customized CRISPR-AsCpf1 system, we successfully and efficiently implemented the related gene editing in B. subtilis 168 for hyaluronic acid (HA) biosynthesis, HA synthase gene (hasA) insertion, UDP-glucose-dehydrogenase gene (tuaD) insertion, and eps gene cluster (epsA-O) deletion. The heterologous production of HA was realized by the engineered strain with a yield of 1.39 g/L. These results support the finding that the CRISPR-AsCpf1 system is highly efficient in bacteria genome editing and provide valuable guidance and essential references for genome engineering in B. subtilis using the CRISPR-AsCpf1 system.

Development and application of a fast and efficient CRISPR-based genetic toolkit in Bacillus amyloliquefaciens LB1ba02

BACKGROUND

Bacillus amyloliquefaciens is generally recognized as food safe (GRAS) microbial host and important enzyme-producing strain in the industry. B.amyloliquefaciens LB1ba02 is a production strain suitable for secreting mesophilic α-amylase in the industry. Nevertheless, due to the low transformation efficiency and restriction-modification system, the development of its CRISPR tool lags far behind other species and strains from the genus Bacillus. This work was undertaken to develop a fast and efficient gene-editing tool in B.amyloliquefaciens LB1ba02.

RESULTS

In this study, we fused the nuclease-deficient mutant Cas9n (D10A) of Cas9 with activation-induced cytidine deaminase (AID) and developed a fast and efficient base editing system for the first time in B. amyloliquefaciens LB1ba02. The system was verified by inactivating the pyrF gene coding orotidine 5'-phosphate decarboxylase and the mutant could grow normally on M9 medium supplemented with 5-fluoroorotic acid (5-FOA) and uridine (U). Our base editing system has a 6nt editing window consisting of an all-in-one temperature-sensitive plasmid that facilitates multiple rounds of genome engineering in B. amyloliquefaciens LB1ba02. The total editing efficiency of this method reached 100% and it achieved simultaneous editing of three loci with an efficiency of 53.3%. In addition, based on the base editing CRISPR/Cas9n-AID system, we also developed a single plasmid CRISPR/Cas9n system suitable for rapid gene knockout and integration. The knockout efficiency for a single gene reached 93%. Finally, we generated 4 genes (aprE, nprE, wprA, and bamHIR) mutant strain, LB1ba02△4. The mutant strain secreted 1.25-fold more α-amylase into the medium than the wild-type strain.

CONCLUSIONS

The CRISPR/Cas9n-AID and CRISPR/Cas9n systems developed in this work proved to be a fast and efficient genetic manipulation tool in a restriction-modification system and poorly transformable strain.

Cell Factory Engineering of Undomesticated Bacillus Strains Using a Modified Integrative and Conjugative Element for Efficient Plasmid Delivery

A large number of Bacillus strains have been isolated from various environments and many of them have great potential as cell factories. However, they have been rarely developed as cell factories due to their poor transformation efficiency. In this study, we developed a highly efficient plasmid delivery system for undomesticated Bacillus strains using a modified integrative and conjugative element (MICE), which was designed to be activated by an inducer, prevent self-transfer, and deliver desired plasmids to the recipient cells. The MICE system was demonstrated to successfully introduce a gfp-containing plasmid into all 41 undomesticated Bacillus subtilis strains tested and eight other Bacillus species. The MICE was used to deliver a cytosine base editor (CBE)-based multiplex genome-editing tool for the cell factory engineering of the Bacillus species. The introduced CBE enabled one-step inactivation of the major extracellular protease genes of the tested strains. The engineered strains were used as hosts for heterologous expression of nattokinase, which resulted in various enzyme expression levels. The results suggested that the MICE and CBE systems can be powerful tools for genetic engineering of undomesticated Bacillus strains, and greatly contribute to the expansion of the Bacillus cell factory.

Efficient Large-Scale and Scarless Genome Engineering Enables the Construction and Screening of Bacillus subtilis Biofuel Overproducers

Bacillus subtilis is a versatile microbial cell factory that can produce valuable proteins and value-added chemicals. Long fragment editing techniques are of great importance for accelerating bacterial genome engineering to obtain desirable and genetically stable host strains. Herein, we develop an efficient CRISPR-Cas9 method for large-scale and scarless genome engineering in the Bacillus subtilis genome, which can delete up to 134.3 kb DNA fragments, 3.5 times as long as the previous report, with a positivity rate of 100%. The effects of using a heterologous NHEJ system, linear donor DNA, and various donor DNA length on the engineering efficiencies were also investigated. The CRISPR-Cas9 method was then utilized for Bacillus subtilis genome simplification and construction of a series of individual and cumulative deletion mutants, which are further screened for overproducer of isobutanol, a new generation biofuel. These results suggest that the method is a powerful genome engineering tool for constructing and screening engineered host strains with enhanced capabilities, highlighting the potential for synthetic biology and metabolic engineering.

Microbial chassis engineering drives heterologous production of complex secondary metabolites

The cryptic secondary metabolite biosynthetic gene clusters (BGCs) far outnumber currently known secondary metabolites. Heterologous production of secondary metabolite BGCs in suitable chassis facilitates yield improvement and discovery of new-to-nature compounds. The two juxtaposed conventional model microorganisms, Escherichia coli, Saccharomyces cerevisiae, have been harnessed as microbial chassis to produce a bounty of secondary metabolites with the help of certain host engineering. In last decade, engineering non-model microbes to efficiently biosynthesize secondary metabolites has received increasing attention due to their peculiar advantages in metabolic networks and/or biosynthesis. The state-of-the-art synthetic biology tools lead the way in operating genetic manipulation in non-model microorganisms for phenotypic optimization or yields improvement of desired secondary metabolites. In this review, we firstly discuss the pros and cons of several model and non-model microbial chassis, as well as the importance of developing broader non-model microorganisms as alternative programmable heterologous hosts to satisfy the desperate needs of biosynthesis study and industrial production. Then we highlight the lately advances in the synthetic biology tools and engineering strategies for optimization of non-model microbial chassis, in particular, the successful applications for efficient heterologous production of multifarious complex secondary metabolites, e.g., polyketides, nonribosomal peptides, as well as ribosomally synthesized and post-translationally modified peptides. Lastly, emphasis is on the perspectives of chassis cells development to access the ideal cell factory in the artificial intelligence-driven genome era.

CRISPR-Cas9-mediated Large Cluster Deletion and Multiplex Genome Editing in Paenibacillus polymyxa

The use of molecular tools based on the clustered regularly interspaced short palindromic repeats-Cas (CRISPR-Cas) systems has rapidly advanced genetic engineering. These molecular biological tools have been applied for different genetic engineering purposes in multiple organisms, including the quite rarely explored Paenibacillus polymyxa. However, only limited studies on large cluster deletion and multiplex genome editing have been described for this highly interesting and versatile bacterium. Here, we demonstrate the utilization of a Cas9-based system to realize targeted deletions of four biosynthetic gene clusters in the range of 12-41 kb by the use of a single targeting sgRNA. Furthermore, we also harnessed the system for multiplex editing of genes and large genomic regions. Multiplex deletion was achieved with more than 80% efficiency, while simultaneous integration at two distantly located sites was obtained with 58% efficiency. The findings reported in this study are anticipated to accelerate future research in P. polymyxa and related species.

Enzymatic deamination of the epigenetic nucleoside N6-methyladenosine regulates gene expression

N⁶-methyladenosine (m⁶A) modification is the most extensively studied epigenetic modification due to its crucial role in regulating an array of biological processes. Herein, Bsu06560, formerly annotated as an adenine deaminase derived from Bacillus subtilis 168, was recognized as the first enzyme capable of metabolizing the epigenetic nucleoside N⁶-methyladenosine. A model of Bsu06560 was constructed, and several critical residues were putatively identified via mutational screening. Two mutants, F91L and Q150W, provided a superiorly enhanced conversion ratio of adenosine and N⁶-methyladenosine. The CRISPR-Cas9 system generated Bsu06560-knockout, F91L, and Q150W mutations from the B. subtilis 168 genome. Transcriptional profiling revealed a higher global gene expression level in BS-F91L and BS-Q150W strains with enhanced N⁶-methyladenosine deaminase activity. The differentially expressed genes were categorized using GO, COG, KEGG and verified through RT-qPCR. This study assessed the crucial roles of Bsu06560 in regulating adenosine and N⁶-methyladenosine metabolism, which influence a myriad of biological processes. This is the first systematic research to identify and functionally annotate an enzyme capable of metabolizing N⁶-methyladenosine and highlight its significant roles in regulation of bacterial metabolism. Besides, this study provides a novel method for controlling gene expression through the mutations of critical residues.

The CRISPR toolbox for the gram-positive model bacterium Bacillus subtilis

CRISPR has revolutionized the way we engineer genomes. Its simplicity and modularity have enabled the development of a great number of tools to edit genomes and to control gene expression. This powerful technology was first adapted to Bacillus subtilis in 2016 and has been intensely upgraded since then. Many tools have been successfully developed to build a CRISPR toolbox for this Gram-positive model and important industrial chassis. The toolbox includes tools, such as double-strand and single-strand cutting CRISPR for point mutation, gene insertion, and gene deletion up to 38 kb. Moreover, catalytic dead Cas proteins have been used for base editing, as well as for the control of gene expression (CRISPRi and CRISPRa). Many of these tools have been used for multiplex CRISPR with the most successful one targeting up to six loci simultaneously for point mutation. However, tools for efficient multiplex CRISPR for other functionalities are still missing in the toolbox. CRISPR engineering has already resulted in efficient protein and metabolite-producing strains, demonstrating its great potential. In this review, we cover all the important additions made to the B. subtilis CRISPR toolbox since 2016, and strain developments fomented by the technology.

A Simplified Method for CRISPR-Cas9 Engineering of Bacillus subtilis

The clustered regularly interspaced short palindromic repeat (CRISPR)-Cas9 system from Streptococcus pyogenes has been widely deployed as a tool for bacterial strain construction. Conventional CRISPR-Cas9 editing strategies require design and molecular cloning of an appropriate guide RNA (gRNA) to target genome cleavage and a repair template for introduction of the desired site-specific genome modification. Here, we present a streamlined method that leverages the existing collection of nearly 4,000 Bacillus subtilis strains (the BKE collection) with individual genes replaced by an integrated erythromycin (erm) resistance cassette. A single plasmid (pAJS23) with a gRNA targeted to erm allows cleavage of the genome at any nonessential gene and at sites nearby to many essential genes. This plasmid can be engineered to include a repair template, or the repair template can be cotransformed with the plasmid as either a PCR product or genomic DNA. We demonstrate the utility of this system for generating gene replacements, site-specific mutations, modification of intergenic regions, and introduction of gene-reporter fusions. In sum, this strategy bypasses the need for gRNA design and allows the facile transfer of mutations and genetic constructions with no requirement for intermediate cloning steps. IMPORTANCE Bacillus subtilis is a well-characterized Gram-positive model organism and a popular platform for biotechnology. Although many different CRISPR-based genome editing strategies have been developed for B. subtilis, they generally involve the design and cloning of a specific guide RNA (gRNA) and repair template for each application. By targeting the erm resistance cassette with an anti-erm gRNA, genome editing can be directed to any of nearly 4,000 gene disruptants within the existing BKE collection of strains. Repair templates can be engineered as PCR products, or specific alleles and constructions can be transformed as chromosomal DNA, thereby bypassing the need for plasmid construction. The described method is rapid and facilitates a wide range of genome manipulations.

Genome editing for resistance against plant pests and pathogens

The conventional breeding of crops struggles to keep up with increasing food needs and ever-adapting pests and pathogens. Global climate changes have imposed another layer of complexity to biological systems, increasing the challenge to obtain improved crop cultivars. These dictate the development and application of novel technologies, like genome editing (GE), that assist targeted and fast breeding programs in crops, with enhanced resistance to pests and pathogens. GE does not require crossings, hence avoiding the introduction of undesirable traits through linkage in elite varieties, speeding up the whole breeding process. Additionally, GE technologies can improve plant protection by directly targeting plant susceptibility (S) genes or virulence factors of pests and pathogens, either through the direct edition of the pest genome or by adding the GE machinery to the plant genome or to microorganisms functioning as biocontrol agents (BCAs). Over the years, GE technology has been continuously evolving and more so with the development of CRISPR/Cas. Here we review the latest advancements of GE to improve plant protection, focusing on CRISPR/Cas-based genome edition of crops and pests and pathogens. We discuss how other technologies, such as host-induced gene silencing (HIGS) and the use of BCAs could benefit from CRISPR/Cas to accelerate the development of green strategies to promote a sustainable agriculture in the future.

Cytosine Base Editor-Mediated Multiplex Genome Editing to Accelerate Discovery of Novel Antibiotics in Bacillus subtilis and Paenibacillus polymyxa

Genome-based identification of new antibiotics is emerging as an alternative to traditional methods. However, uncovering hidden antibiotics under the background of known antibiotics remains a challenge. To over this problem using a quick and effective genetic approach, we developed a multiplex genome editing system using a cytosine base editor (CBE). The CBE system achieved simultaneous double, triple, quadruple, and quintuple gene editing with efficiencies of 100, 100, 83, and 75%, respectively, as well as the 100% editing efficiency of single targets in Bacillus subtilis. Whole-genome sequencing of the edited strains showed that they had an average of 8.5 off-target single-nucleotide variants at gRNA-independent positions. The CBE system was used to simultaneously knockout five known antibiotic biosynthetic gene clusters to leave only an uncharacterized polyketide biosynthetic gene cluster in Paenibacillus polymyxa E681. The polyketide showed antimicrobial activities against gram-positive bacteria, but not gram-negative bacteria and fungi. Therefore, our findings suggested that the CBE system might serve as a powerful tool for multiplex genome editing and greatly accelerating the unraveling of hidden antibiotics in Bacillus and Paenibacillus species.

Engineering of Multiple Modules to Improve Amorphadiene Production in Bacillus subtilis Using CRISPR-Cas9

Engineering strategies to improve terpenoids' production in Bacillus subtilis mainly focus on 2C-methyl-d-erythritol-4-phosphate (MEP) pathway overexpression. To systematically engineer the chassis strain for higher amorphadiene (precursor of artemisinin) production, a clustered regularly interspaced short palindromic repeat-Cas9 (CRISPR-Cas9) system was established in B. subtilis to facilitate precise and efficient genome editing. Then, this system was employed to engineer three more modules to improve amorphadiene production, including the terpene synthase module, the branch pathway module, and the central metabolic pathway module. Finally, our combination of all of the useful strategies within one strain significantly increased extracellular amorphadiene production from 81 to 116 mg/L after 48 h flask fermentation without medium optimization. For the first time, we attenuated the FPP-derived competing pathway to improve amorphadiene biosynthesis and investigated how the TCA cycle affects amorphadiene production in B. subtilis. Overall, this study provides a universal strategy for further increasing terpenoids' production in B. subtilis by comprehensive and systematic metabolic engineering.

CRISPR-Cas system, antibiotic resistance and virulence in bacteria: Through a common lens

CRISPR-Cas system, antibiotic resistance and virulence are different modes of survival for the bacteria. CRISPR-Cas is an adaptive immune system that can degrade foreign DNA, antibiotic resistance helps bacteria to evade drugs that can threaten their existence and virulence determinants are offensive tools that can facilitate the establishment of infection by pathogens. This chapter focuses on these three aspects, providing insights about the CRISPR system and resistance mechanisms in brief, followed by understanding the synergistic or antagonistic relationship of resistance and virulence determinants in connection to the CRISPR system. We have addressed the discussion of this evolving topic through specific examples and studies. Different approaches for successful detection of this unique defense system in bacteria and various applications of the CRISPR-Cas systems to show how it can be harnessed to tackle the increasing problem of antibiotic resistance have been put forth. World Health Organization has declared antibiotic resistance as a serious global problem of the 21st century. As antibiotic-resistant bacteria increase their footprint across the globe, newer tools such as the CRISPR-Cas system hold immense promise to tackle this problem.

In-situ generation of large numbers of genetic combinations for metabolic reprogramming via CRISPR-guided base editing

Reprogramming complex cellular metabolism requires simultaneous regulation of multigene expression. Ex-situ cloning-based methods are commonly used, but the target gene number and combinatorial library size are severely limited by cloning and transformation efficiencies. In-situ methods such as multiplex automated genome engineering (MAGE) depends on high-efficiency transformation and incorporation of heterologous DNA donors, which are limited to few microorganisms. Here, we describe a Base Editor-Targeted and Template-free Expression Regulation (BETTER) method for simultaneously diversifying multigene expression. BETTER repurposes CRISPR-guided base editors and in-situ generates large numbers of genetic combinations of diverse ribosome binding sites, 5’ untranslated regions, or promoters, without library construction, transformation, and incorporation of DNA donors. We apply BETTER to simultaneously regulate expression of up to ten genes in industrial and model microorganisms Corynebacterium glutamicum and Bacillus subtilis. Variants with improved xylose catabolism, glycerol catabolism, or lycopene biosynthesis are respectively obtained. This technology will be useful for large-scale fine-tuning of multigene expression in both genetically tractable and intractable microorganisms.

To obtain optimal yield and productivity in bioproduction, expression of pathway genes must be appropriately coordinated. Here, the authors report repurposing of base editors for simultaneous regulation of multiple gene expression and demonstrate its application in industrially important and model microorganisms.

Fragment Exchange Plasmid Tools for CRISPR/Cas9-Mediated Gene Integration and Protease Production in Bacillus subtilis

We complemented a cloning platform with new editing plasmids that allow a quick transition from high-throughput cloning and the expression of new enzymes to the stable integration of genes for the production of enzymes through B. subtilis fermentation. We present two systems for the effective assembly cloning of any genome-editing cassette that shortens the engineering procedure to obtain the final editing constructs. The utility of the customized tools is demonstrated by disrupting Bacillus’ capacity to sporulate and by introducing the stable expression of subtilisin. The tools should be useful to engineer B. subtilis strains by a variety of recombination events to ultimately improve the application range of this industry-relevant host.

Since its discovery as part of the bacterial adaptative immune system, CRISPR/Cas has emerged as the most promising tool for targeted genome editing over the past few years. Various tools for genome editing in Bacillus subtilis have recently been developed, expanding and simplifying its potential development as an industrial species. A collection of vectors compatible with high-throughput (HTP) fragment exchange (FX) cloning for heterologous expression in Escherichia coli and Bacillus was previously developed. This vector catalogue was through this work supplemented with editing plasmids for genome engineering in Bacillus by adapting two CRISPR/Cas plasmids to the cloning technology. The customized tools allow versatile editing at any chosen genomic position (single-plasmid strategy) or at a fixed genomic locus (double-plasmid strategy). The single-plasmid strategy was validated by deleting the spoIIAC gene, which has an essential role in sporulation. Using the double-plasmid strategy, we demonstrate the quick transition from plasmid-based subtilisin expression to the stable integration of the gene into the amyE locus of a seven-protease-deficient KO7 strain. The newly engineered B. subtilis strain allowed the successful production of a functional enzyme. The customized tools provide improvements to the cloning procedure, should be useful for versatile genomic engineering, and contribute to a cloning platform for a quick transition from HTP enzyme expression to production through the fermentation of industrially relevant B. subtilis and related strains.

IMPORTANCE We complemented a cloning platform with new editing plasmids that allow a quick transition from high-throughput cloning and the expression of new enzymes to the stable integration of genes for the production of enzymes through B. subtilis fermentation. We present two systems for the effective assembly cloning of any genome-editing cassette that shortens the engineering procedure to obtain the final editing constructs. The utility of the customized tools is demonstrated by disrupting Bacillus’ capacity to sporulate and by introducing the stable expression of subtilisin. The tools should be useful to engineer B. subtilis strains by a variety of recombination events to ultimately improve the application range of this industry-relevant host.

CRISPR-based metabolic pathway engineering

A highly effective metabolic pathway is the key for an efficient cell factory. However, the engineered homologous or heterologous multi-gene pathway may be unbalanced, inefficient and causing the accumulation of potentially toxic intermediates. Therefore, pathways must be constructed optimally to minimize these negative effects and maximize catalytic efficiency. With the development of CRISPR technology, some of the problems of previous pathway engineering and genome editing techniques were resolved, providing higher efficiency, lower cost, and easily customizable targets. Moreover, CRISPR was demonstrated as robust and effective in various organisms including both prokaryotes and eukaryotes. In recent years, researchers in the field of metabolic engineering and synthetic biology have exploited various CRISPR-based pathway engineering approaches, which are both effective and convenient, as well as valuable from a theoretical standpoint. In this review, we systematically summarize novel pathway engineering techniques and strategies based on CRISPR nucleases system, CRISPR interference (CRISPRi), and CRISPR activation (CRISPRa), including figures and descriptions for easy understanding, with the aim to facilitate their broader application among fellow researchers.

Mining New Plipastatins and Increasing the Total Yield Using CRISPR/Cas9 in Genome-Modified Bacillus subtilis 1A751

CRISPR/Cas9 is one of the robust and effective gene manipulation tools which has been widely applied in various organisms. In this study, the plipastatin gene cluster was successfully expressed in genome-modified Bacillus subtilis 1A751 by disrupting the surfactin operon (srf) through CRISPR/Cas9 technology. The presumed plipastatin biosynthetic pathway was proposed based on the analysis of its biosynthetic gene cluster. Two new plipastatins were identified by a combination of ultra-high performance liquid chromatography-coupled electron spray ionization-tandem mass spectrometry and gas chromatography-mass spectrometry analyses, together with nine known plipastatins or their derivatives. The yield of plipastatin was as high as 1600 mg/L which is the highest reported to date. Antimicrobial experiments revealed that its methanolic extracts exhibited powerful inhibitory effects on the growth of the tested pathogens and fungi. The results from this investigation highlight the remarkable utility of CRISPR/Cas9 in mining new plipastatins and increasing the total plipastatin yield, providing a new pipeline for the industrial application of plipastatin.

Bacillus subtilis: a universal cell factory for industry, agriculture, biomaterials and medicine

Due to its clear inherited backgrounds as well as simple and diverse genetic manipulation systems, Bacillus subtilis is the key Gram-positive model bacterium for studies on physiology and metabolism. Furthermore, due to its highly efficient protein secretion system and adaptable metabolism, it has been widely used as a cell factory for microbial production of chemicals, enzymes, and antimicrobial materials for industry, agriculture, and medicine. In this mini-review, we first summarize the basic genetic manipulation tools and expression systems for this bacterium, including traditional methods and novel engineering systems. Secondly, we briefly introduce its applications in the production of chemicals and enzymes, and summarize its advantages, mainly focusing on some noteworthy products and recent progress in the engineering of B. subtilis. Finally, this review also covers applications such as microbial additives and antimicrobials, as well as biofilm systems and spore formation. We hope to provide an overview for novice researchers in this area, offering them a better understanding of B. subtilis and its applications.

Design and Construction of Portable CRISPR-Cpf1-Mediated Genome Editing in Bacillus subtilis 168 Oriented Toward Multiple Utilities

Bacillus subtilis is an important Gram-positive bacterium for industrial biotechnology, which has been widely used to produce diverse high-value added chemicals and industrially and pharmaceutically relevant proteins. Robust and versatile toolkits for genome editing in B. subtilis are highly demanding to design higher version chassis. Although the Streptococcus pyogenes (Sp) CRISPR-Cas9 has been extensively adapted for genome engineering of multiple bacteria, it has many defects, such as higher molecular weight which leads to higher carrier load, low deletion efficiency and complexity of sgRNA construction for multiplex genome editing. Here, we designed a CRISPR-Cpf1-based toolkit employing a type V Cas protein, Cpf1 from Francisella novicida. Using this platform, we precisely deleted single gene and gene cluster in B. subtilis with high editing efficiency, such as sacA, ganA, ligD & ligV, and bac operon. Especially, an extremely large gene cluster of 38 kb in B. subtilis genome was accurately deleted from the genome without introducing any unexpected mutations. Meanwhile, the synthetic platform was further upgraded to a version for multiplex genome editing, upon which two genes sacA and aprE were precisely and efficiently deleted using only one plasmid harboring two targeting sequences. In addition, we successfully inserted foreign genes into the genome of the chassis using the CRISPR-Cpf1 platform. Our work highlighted the availability of CRISPR-Cpf1 to gene manipulation in B. subtilis, including the flexible deletion of a single gene and multiple genes or a gene cluster, and gene knock-in. The designed genome-editing platform was easily and broadly applicable to other microorganisms. The novel platforms we constructed in this study provide a promising tool for efficient genome editing in diverse bacteria.

Genome editing of Francisella tularensis using (CRISPR-Cas9)

The clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) system is a powerful tool for gene editing in eukaryotic genomes but is still being developed for editing bacterial genomes. Here we describe the construction of an all-in-one vector for generating potentially scarless deletion mutants in Francisella tularensis LVS using a CRISPR-Cas9-based system.

CRISPR-dCas9 Mediated Cytosine Deaminase Base Editing in Bacillus subtilis

Base editing technology based on clustered regularly interspaced short palindromic repeats/associated protein 9 (CRISPR/Cas9) is a recent addition to the family of CRISPR technologies. Compared with the traditional CRISPR/Cas9 technology, it does not rely on DNA double strand break and homologous recombination, and can realize gene inactivation and point mutation more quickly and simply. Herein, we first developed a base editing method for genome editing in Bacillus subtilis utilizing CRISPR/dCas9 (a fully nuclease-deficient mutant of Cas9 from S. pyogenes) and activation-induced cytidine deaminase (AID). This method achieved three and four loci simultaneous editing with editing efficiency up to 100% and 50%, respectively. Our base editing system in B. subtilis has a 5 nt editing window, which is similar to previously reported base editing in other microorganisms. We demonstrated that the plasmid curing rate is almost 100%, which is advantageous for multiple rounds of genome engineering in B. subtilis. Finally, we applied multiplex genome editing to generate a B. subtilis 168 mutant strain with eight inactive extracellular protease genes in just two rounds of base editing and plasmid curing, suggesting that it is a powerful tool for gene manipulation in B. subtilis and industrial applications in the future.

Antibiotic Resistance Profiles, Molecular Mechanisms and Innovative Treatment Strategies of Acinetobacter baumannii

Antibiotic resistance is one of the biggest challenges for the clinical sector and industry, environment and societal development. One of the most important pathogens responsible for severe nosocomial infections is Acinetobacter baumannii, a Gram-negative bacterium from the Moraxellaceae family, due to its various resistance mechanisms, such as the β-lactamases production, efflux pumps, decreased membrane permeability and altered target site of the antibiotic. The enormous adaptive capacity of A. baumannii and the acquisition and transfer of antibiotic resistance determinants contribute to the ineffectiveness of most current therapeutic strategies, including last-line or combined antibiotic therapy. In this review, we will present an update of the antibiotic resistance profiles and underlying mechanisms in A. baumannii and the current progress in developing innovative strategies for combating multidrug-resistant A. baumannii (MDRAB) infections.

Targeting Plasmids to Limit Acquisition and Transmission of Antimicrobial Resistance

Antimicrobial resistance (AMR) is a significant global threat to both public health and the environment. The emergence and expansion of AMR is sustained by the enormous diversity and mobility of antimicrobial resistance genes (ARGs). Different mechanisms of horizontal gene transfer (HGT), including conjugation, transduction, and transformation, have facilitated the accumulation and dissemination of ARGs in Gram-negative and Gram-positive bacteria. This has resulted in the development of multidrug resistance in some bacteria. The most clinically significant ARGs are usually located on different mobile genetic elements (MGEs) that can move intracellularly (between the bacterial chromosome and plasmids) or intercellularly (within the same species or between different species or genera). Resistance plasmids play a central role both in HGT and as support elements for other MGEs, in which ARGs are assembled by transposition and recombination mechanisms. Considering the crucial role of MGEs in the acquisition and transmission of ARGs, a potential strategy to control AMR is to eliminate MGEs. This review discusses current progress on the development of chemical and biological approaches for the elimination of ARG carriers.

Recent Advances in Recombinant Protein Production byBacillus subtilis

Bacillus subtilis has become a widely used microbial cell factory for the production of recombinant proteins, especially those associated with foods and food processing. Recent advances in genetic manipulation and proteomic analysis have been used to greatly improve protein production in B. subtilis. This review begins with a discussion of genome-editing technologies and application of the CRISPR–Cas9 system to B. subtilis. A summary of the characteristics of crucial legacy strains is followed by suggestions regarding the choice of origin strain for genetic manipulation. Finally, the review analyzes the genes and operons of B. subtilis that are important for the production of secretory proteins and provides suggestions and examples of how they can be altered to improve protein production. This review is intended to promote the engineering of this valuable microbial cell factory for better recombinant protein production.

Recent Advances of CRISPR/Cas9-Based Genetic Engineering and Transcriptional Regulation in Industrial Biology

Industrial biology plays a crucial role in the fields of medicine, health, food, energy, and so on. However, the lack of efficient genetic engineering tools has restricted the rapid development of industrial biology. Recently, the emergence of clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) system brought a breakthrough in genome editing technologies due to its high orthogonality, versatility, and efficiency. In this review, we summarized the barriers of CRISPR/Cas9 and corresponding solutions for efficient genetic engineering in industrial microorganisms. In addition, the advances of industrial biology employing the CRISPR/Cas9 system were compared in terms of its application in bacteria, yeast, and filamentous fungi. Furthermore, the cooperation between CRISPR/Cas9 and synthetic biology was discussed to help build complex and programmable gene circuits, which can be used in industrial biotechnology.

Restructured Lactococcus lactis strains with emergent properties constructed by a novel highly efficient screening system

Background

After 2.83% genome reduction in Lactococcus lactis NZ9000, a good candidate host for proteins production was obtained in our previous work. However, the gene deletion process was time consuming and laborious. Here, we proposed a convenient gene deletion method suitable for large-scale genome reduction in L. lactis NZ9000.

Results

Plasmid pNZ5417 containing a visually selectable marker P_nisZ-lacZ was constructed, which allowed more efficient and convenient screening of gene deletion mutants. Using this plasmid, two large nonessential DNA regions, L-4A and L-5A, accounting for 1.25% of the chromosome were deleted stepwise in L. lactis 9k-3. When compared with the parent strain, the mutant L. lactis 9k-5A showed better growth characteristics, transformability, carbon metabolic capacity, and amino acids biosynthesis.

Conclusions

Thus, this study provides a convenient and efficient system for large-scale genome deletion in L. lactis through application of visually selectable marker, which could be helpful for rapid genome streamlining and generation of restructured L. lactis strains that can be used as cell factories.