A new mutation delivery system for genome-scale approaches in Bacillus subtilis

Bacillus subtilis as a host for natural product discovery and engineering of biosynthetic gene clusters

Covering: up to October 2023Many bioactive natural products are synthesized by microorganisms that are either difficult or impossible to cultivate under laboratory conditions, or that produce only small amounts of the desired compound. By transferring biosynthetic gene clusters (BGCs) into alternative host organisms that are more easily cultured and engineered, larger quantities can be obtained and new analogues with potentially improved biological activity or other desirable properties can be generated. Moreover, expression of cryptic BGCs in a suitable host can facilitate the identification and characterization of novel natural products. Heterologous expression therefore represents a valuable tool for natural product discovery and engineering as it allows the study and manipulation of their biosynthetic pathways in a controlled setting, enabling innovative applications. Bacillus is a genus of Gram-positive bacteria that is widely used in industrial biotechnology as a host for the production of proteins from diverse origins, including enzymes and vaccines. However, despite numerous successful examples, Bacillus species remain underexploited as heterologous hosts for the expression of natural product BGCs. Here, we review important advantages that Bacillus species offer as expression hosts, such as high secretion capacity, natural competence for DNA uptake, and the increasing availability of a wide range of genetic tools for gene expression and strain engineering. We evaluate different strain optimization strategies and other critical factors that have improved the success and efficiency of heterologous natural product biosynthesis in B. subtilis. Finally, future perspectives for using B. subtilis as a heterologous host are discussed, identifying research gaps and promising areas that require further exploration.

Establishment of low-cost production platforms of polyhydroxyalkanoate bioplastics from Halomonas cupida J9

Microbial production of polyhydroxyalkanoate (PHA) is greatly restricted by high production cost arising from high-temperature sterilization and expensive carbon sources. In this study, a low-cost PHA production platform was established from Halomonas cupida J9. First, a marker-less genome-editing system was developed in H. cupida J9. Subsequently, H. cupida J9 was engineered to efficiently utilize xylose for PHA biosynthesis by introducing a new xylose metabolism module and blocking xylonate production. The engineered strain J9UΔxylD-P8xylA has the highest PHA yield (2.81 g/L) obtained by Halomonas with xylose as the sole carbon source so far. This is the first report on the production of short- and medium-chain-length (SCL-co-MCL) PHA from xylose by Halomonas. Interestingly, J9UΔxylD-P8xylA was capable of efficiently utilizing glucose and xylose as co-carbon sources for PHA production. Furthermore, fed-batch fermentation of J9UΔxylD-P8xylA coupled to a glucose/xylose co-feeding strategy reached up to 12.57 g/L PHA in a 5-L bioreactor under open and unsterile condition. Utilization of corn straw hydrolysate as the carbon source by J9UΔxylD-P8xylA reached 7.0 g/L cell dry weight (CDW) and 2.45 g/L PHA in an open fermentation. In summary, unsterile production in combination with inexpensive feedstock highlights the potential of the engineered strain for the low-cost production of PHA from lignocellulose-rich agriculture waste.

Engineered bacterial orthogonal DNA replication system for continuous evolution

Continuous evolution can generate biomolecules for synthetic biology and enable evolutionary investigation. The orthogonal DNA replication system (OrthoRep) in yeast can efficiently mutate long DNA fragments in an easy-to-operate manner. However, such a system is lacking in bacteria. Therefore, we developed a bacterial orthogonal DNA replication system (BacORep) for continuous evolution. We achieved this by harnessing the temperate phage GIL16 DNA replication machinery in Bacillus thuringiensis with an engineered error-prone orthogonal DNA polymerase. BacORep introduces all 12 types of nucleotide substitution in 15-kilobase genes on orthogonally replicating linear plasmids with a 6,700-fold higher mutation rate than that of the host genome, the mutation rate of which is unchanged. Here we demonstrate the utility of BacORep-based continuous evolution by generating strong promoters applicable to model bacteria, Bacillus subtilis and Escherichia coli, and achieving a 7.4-fold methanol assimilation increase in B. thuringiensis. BacORep is a powerful tool for continuous evolution in prokaryotic cells.

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.

Establishing genetic manipulation for novel strains of human gut bacteria

Recent years have seen the development of high-accuracy and high-throughput genetic manipulation techniques, which have greatly improved our understanding of genetically tractable microbes. However, challenges remain in establishing genetic manipulation techniques in novel organisms, owing largely to exogenous DNA defence mechanisms, lack of selectable markers, lack of efficient methods to introduce exogenous DNA and an inability of genetic vectors to replicate in their new host. In this review, we describe some of the techniques that are available for genetic manipulation of novel microorganisms. While many reviews exist that focus on the final step in genetic manipulation, the editing of recipient DNA, we particularly focus on the first step in this process, the transfer of exogenous DNA into a strain of interest. Examples illustrating the use of these techniques are provided for a selection of human gut bacteria in which genetic tractability has been established, such as Bifidobacterium, Bacteroides and Roseburia. Ultimately, this review aims to provide an information source for researchers interested in developing genetic manipulation techniques for novel bacterial strains, particularly those of the human gut microbiota.

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 of a Markerless Deletion Mutagenesis System in Nitrate-Reducing Bacterium Rhodanobacter denitrificans

Rhodanobacter has been found as the dominant genus in aquifers contaminated with high concentrations of nitrate and uranium in Oak Ridge, TN, USA. The in situ stimulation of denitrification has been proposed as a potential method to remediate nitrate and uranium contamination. Among the Rhodanobacter species, Rhodanobacter denitrificans strains have been reported to be capable of denitrification and contain abundant metal resistance genes. However, due to the lack of a mutagenesis system in these strains, our understanding of the mechanisms underlying low-pH resistance and the ability to dominate in the contaminated environment remains limited. Here, we developed an in-frame markerless deletion system in two R. denitrificans strains. First, we optimized the growth conditions, tested antibiotic resistance, and determined appropriate transformation parameters in 10 Rhodanobacter strains. We then deleted the upp gene, which encodes uracil phosphoribosyltransferase, in R. denitrificans strains FW104-R3 and FW104-R5. The resulting strains were designated R3_Δupp and R5_Δupp and used as host strains for mutagenesis with 5-fluorouracil (5-FU) resistance as the counterselection marker to generate markerless deletion mutants. To test the developed protocol, the narG gene encoding nitrate reductase was knocked out in the R3_Δupp and R5_Δupp host strains. As expected, the narG mutants could not grow in anoxic medium with nitrate as the electron acceptor. Overall, these results show that the in-frame markerless deletion system is effective in two R. denitrificans strains, which will allow for future functional genomic studies in these strains furthering our understanding of the metabolic and resistance mechanisms present in Rhodanobacter species. IMPORTANCE Rhodanobacter denitrificans is capable of denitrification and is also resistant to toxic heavy metals and low pH. Accordingly, the presence of Rhodanobacter species at a particular environmental site is considered an indicator of nitrate and uranium contamination. These characteristics suggest its future potential application in bioremediation of nitrate or concurrent nitrate and uranium contamination in groundwater ecosystems. Due to the lack of genetic tools in this organism, the mechanisms of low-pH and heavy metal resistance in R. denitrificans strains remain elusive, which impedes its use in bioremediation strategies. Here, we developed a genome editing method in two R. denitrificans strains. This work marks a crucial step in developing Rhodanobacter as a model for studying the diverse mechanisms of low-pH and heavy metal resistance associated with denitrification.

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.

An overview and future prospects of recombinant protein production in Bacillus subtilis

Bacillus subtilis is a well-characterized Gram-positive bacterium and a valuable host for recombinant protein production because of its efficient secretion ability, high yield, and non-toxicity. Here, we comprehensively review the recent studies on recombinant protein production in B. subtilis to update and supplement other previous reviews. We have focused on several aspects, including optimization of B. subtilis strains, enhancement and regulation of expression, improvement of secretion level, surface display of proteins, and fermentation optimization. Among them, optimization of B. subtilis strains mainly involves undirected chemical/physical mutagenesis and selection and genetic manipulation; enhancement and regulation of expression comprises autonomous plasmid and integrated expression, promoter regulation and engineering, and fine-tuning gene expression based on proteases and molecular chaperones; improvement of secretion level predominantly involves secretion pathway and signal peptide screening and optimization; surface display of proteins includes surface display of proteins on spores or vegetative cells; and fermentation optimization incorporates medium optimization, process condition optimization, and feeding strategy optimization. Furthermore, we propose some novel methods and future challenges for recombinant protein production in B. subtilis.Key points• A comprehensive review on recombinant protein production in Bacillus subtilis.• Novel techniques facilitate recombinant protein expression and secretion.• Surface display of proteins has significant potential for different applications.

Advances in Accurate Microbial Genome-Editing CRISPR Technologies

Previous studies have modified microbial genomes by introducing gene cassettes containing selectable markers and homologous DNA fragments. However, this requires several steps including homologous recombination and excision of unnecessary DNA regions, such as selectable markers from the modified genome. Further, genomic manipulation often leaves scars and traces that interfere with downstream iterative genome engineering. A decade ago, the CRISPR/Cas system (also known as the bacterial adaptive immune system) revolutionized genome editing technology. Among the various CRISPR nucleases of numerous bacteria and archaea, the Cas9 and Cas12a (Cpf1) systems have been largely adopted for genome editing in all living organisms due to their simplicity, as they consist of a single polypeptide nuclease with a target-recognizing RNA. However, accurate and fine-tuned genome editing remains challenging due to mismatch tolerance and protospacer adjacent motif (PAM)-dependent target recognition. Therefore, this review describes how to overcome the aforementioned hurdles, which especially affect genome editing in higher organisms. Additionally, the biological significance of CRISPR-mediated microbial genome editing is discussed, and future research and development directions are also proposed.

Advances in Enhanced Menaquinone-7 Production From Bacillus subtilis

The production of nutraceutical compounds through biosynthetic approaches has received considerable attention in recent years. For example, Menaquinone-7 (MK-7), a sub-type of Vitamin K2, biosynthesized from Bacillus subtilis (B. subtilis), proved to be more efficiently produced than the conventional chemical synthesis techniques. This is possible due to the development of B. subtilis as a chassis cell during the biosynthesis stages. Hence, it is imperative to provide insights on the B. subtilis membrane permeability modifications, biofilm reactors, and fermentation optimization as advanced techniques relevant to MK-7 production. Although the traditional gene-editing method of homologous recombination improves the biosynthetic pathway, CRISPR-Cas9 could potentially resolve the drawbacks of traditional genome editing techniques. For these reasons, future studies should explore the applications of CRISPRi (CRISPR interference) and CRISPRa (CRISPR activation) system gene-editing tools in the MK-7 anabolism pathway.

Development of an efficient pathway construction strategy for rapid evolution of the biodegradation capacity of Pseudomonas putida KT2440 and its application in bioremediation

In this work, we developed an efficient pathway construction strategy, consisting of DNA assembler-assisted pathway assembly and counterselection system-based chromosomal integration, for the rapid and efficient integration of synthetic biodegradation pathways into the chromosome of Pseudomonas putida KT2440. Using this strategy, we created a novel degrader capable of complete mineralization of γ-hexachlorocyclohexane (γ-HCH) and 1,2,3-trichloropropane (TCP) by integrating γ-HCH and TCP biodegradation pathways into the chromosome of P. putida KT2440. Furthermore, the chromosomal integration efficiencies of γ-HCH and TCP biodegradation pathways were improved to 50% and 41.6% in P. putida KT2440, respectively, by the inactivation of a type I DNA restriction-modification system. The currently developed pathway construction strategy coupled with the mutant KTUΔhsdRMS will facilitate implantation of heterologous catabolic pathways into the chromosome for rapid evolution of the biodegradation capacity of P. putida. More importantly, the successful removal of γ-HCH (10 mg/kg soil) and TCP (0.2 mM) from soil and wastewater within 14 days, respectively, highlighted the potential of the novel degrader for in situ bioremediation of γ-HCH- and TCP-contaminated sites. Moreover, chromosomal integration of gfp made the degrader to be monitored easily during bioremediation. In the future, this strategy can be expanded to a broad range of bacterial species for widespread applications in bioremediation.

Development and application of a highly efficient CRISPR-Cas9 system for genome engineering in Bacillus megaterium

Bacillus megaterium has become increasingly important for the biotechnological production of valuable compounds of industrial and pharmaceutical importance. Despite recent advances in rational strain design of B. megaterium, these studies have been largely impaired by the lack of molecular tools that are not state-of-the-art for comprehensive genome engineering approaches. In the current work, we describe the adaptation of the CRISPR-Cas9 vector pJOE8999 to enable efficient genome editing in B. megaterium. Crucial modifications comprise the exchange of promoter elements and associated ribosomal binding sites as well as the implementation of a 5-fluorouracil based counterselection system to facilitate proper plasmid curing. In addition, the functionality and performance of the new CRISPR-Cas9 vector pMOE was successfully evaluated by chromosomal disruption studies of the endogenous β-galactosidase gene (BMD_2126) and demonstrated an outstanding efficiency of 100 % based on combinatorial pheno- and genotype analyses. Furthermore, pMOE was applied for the genomic deletion of a steroid esterase gene (BMD_2256) that was identified among several other candidates as the gene encoding the esterase, which prevented accumulation of pharmaceutically important glucocorticoid esters. Recombinant expression of the bacterial chloramphenicol acetyltransferase 1 gene (cat1) in the resulting esterase deficient B. megaterium strain ultimately yielded C21-acetylated as well as novel C21-esterified derivates of cortisone.

Approaches to genetic tool development for rapid domestication of non-model microorganisms

Non-model microorganisms often possess complex phenotypes that could be important for the future of biofuel and chemical production. They have received significant interest the last several years, but advancement is still slow due to the lack of a robust genetic toolbox in most organisms. Typically, "domestication" of a new non-model microorganism has been done on an ad hoc basis, and historically, it can take years to develop transformation and basic genetic tools. Here, we review the barriers and solutions to rapid development of genetic transformation tools in new hosts, with a major focus on Restriction-Modification systems, which are a well-known and significant barrier to efficient transformation. We further explore the tools and approaches used for efficient gene deletion, DNA insertion, and heterologous gene expression. Finally, more advanced and high-throughput tools are now being developed in diverse non-model microbes, paving the way for rapid and multiplexed genome engineering for biotechnology.

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.

Bacterial Genetic Engineering by Means of Recombineering for Reverse Genetics

Serving a robust platform for reverse genetics enabling the in vivo study of gene functions primarily in enterobacteriaceae, recombineering -or recombination-mediated genetic engineering-represents a powerful and relative straightforward genetic engineering tool. Catalyzed by components of bacteriophage-encoded homologous recombination systems and only requiring short ∼40–50 base homologies, the targeted and precise introduction of modifications (e.g., deletions, knockouts, insertions and point mutations) into the chromosome and other episomal replicons is empowered. Furthermore, by its ability to make use of both double- and single-stranded linear DNA editing substrates (e.g., PCR products or oligonucleotides, respectively), lengthy subcloning of specific DNA sequences is circumvented. Further, the more recent implementation of CRISPR-associated endonucleases has allowed for more efficient screening of successful recombinants by the selective purging of non-edited cells, as well as the creation of markerless and scarless mutants. In this review we discuss various recombineering strategies to promote different types of gene modifications, how they are best applied, and their possible pitfalls.

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.

Advances on systems metabolic engineering of Bacillus subtilis as a chassis cell

The Gram-positive model bacterium Bacillus subtilis, has been broadly applied in various fields because of its low pathogenicity and strong protein secretion ability, as well as its well-developed fermentation technology. B. subtilis is considered as an attractive host in the field of metabolic engineering, in particular for protein expression and secretion, so it has been well studied and applied in genetic engineering. In this review, we discussed why B. subtilis is a good chassis cell for metabolic engineering. We also summarized the latest research progress in systematic biology, synthetic biology and evolution-based engineering of B. subtilis, and showed systemic metabolic engineering expedite the harnessing B. subtilis for bioproduction.

Relative contributions of non-essential Sec pathway components and cell envelope-associated proteases to high-level enzyme secretion by Bacillus subtilis

Background

Bacillus subtilis is an important industrial workhorse applied in the production of many different commercially relevant proteins, especially enzymes. Virtually all of these proteins are secreted via the general secretion (Sec) pathway. Studies from different laboratories have demonstrated essential or non-essential contributions of various Sec machinery components to protein secretion in B. subtilis. However, a systematic comparison of the impact of each individual Sec machinery component under conditions of high-level protein secretion was so far missing.

Results

In the present study, we have compared the contributions of non-essential Sec pathway components and cell envelope-associated proteases on the secretion efficiency of three proteins expressed at high level. This concerned the α-amylases AmyE from B. subtilis and AmyL from Bacillus licheniformis, and the serine protease BPN’ from Bacillus amyloliquefaciens. We compared the secretion capacity of mutant strains in shake flask cultures, and the respective secretion kinetics by pulse-chase labeling experiments. The results show that secDF, secG or rasP mutations severely affect AmyE, AmyL and BPN’ secretion, but the actual effect size depends on the investigated protein. Additionally, the chaperone DnaK is important for BPN’ secretion, while AmyE or AmyL secretion are not affected by a dnaK deletion. Further, we assessed the induction of secretion stress responses in mutant strains by examining AmyE- and AmyL-dependent induction of the quality control proteases HtrA and HtrB. Interestingly, the deletion of certain sip genes revealed a strong differential impact of particular signal peptidases on the magnitude of the secretion stress response.

Conclusions

The results of the present study highlight the importance of SecDF, SecG and RasP for protein secretion and reveal unexpected differences in the induction of the secretion stress response in different mutant strains.

Identification of critical residues for the catalytic activity of ComQ, a Bacillus prenylation enzyme for quorum sensing, by using a simple bioassay system

Bacillus ComQ participates in the biosynthesis of a quorum-sensing signaling molecule (ComX pheromone) through catalyzing the prenylation at a Trp residue of the precursor peptide (pre-ComX) with geranyl diphosphate (C10 type) or farnesyl diphosphate (C15 type). We hypothesized that several residues specifically conserved among either type of ComQs are important for their substrate specificities. Using a simple bioassay, we revealed that Phe63, Asn186, and Gly190 in ComQRO-E-2 (C10 type) were nondisplaceable to Ser63, Gly186, and Val190, the corresponding residues in the C15-type ComQ, respectively. A three-dimensional model suggested that the 186th and 190th residues are involved in the pre-ComX binding. In vitro analysis showed that substitution of Phe63 with Ser in ComQRO-E-2 significantly reduced the geranylation activity but substantially enhanced the farnesylation activity, whereas substitution of Ser63 with Phe in ComQ168 (C15 type) reduced the farnesylation activity. Therefore, the 63rd residue was found to be significant for the prenyl-substrate preference.

Abbreviations: GPP: geranyl diphosphate; FPP: farnesyl diphosphate; IPP: isopentenyl diphosphate; GGPP: geranylgeranyl diphosphate; FARM: first aspartate-rich motif; SARM: second aspartate-rich motif; β-Gal: β-galactosidase; TBABG: tryptose blood agar base supplemented with glucose; X-gal: 5-bromo-4-chloro-3-indolyl-β-D-galactoside

Regulation of hyaluronic acid molecular weight and titer by temperature in engineered Bacillus subtilis

Hyaluronic acid (HA) is a biopolymer used in several industries. There is increasing global demand. HA is normally produced on a large scale using attenuated strains of group C streptococci that are pathogenic and fastidious. Accordingly, it is of interest to use a "generally recognized as safe" (GRAS) organism such as Bacillus subtilis for HA production. Here, we report an engineered B. subtilis strain named WmB that produces different molecular weights (MW) and titers of HA at different temperatures. The faster the bacteria grew, the lower the MW of HA produced and the higher the titer. The MW of HA obtained ranged from 6.937 MDa at 47 °C to 0.392 MDa at 32 °C. At 32 °C, the HA titer reached 3.65 ± 0.13 g/L. We have engineered a strain that can produce high-molecular-weight and medium-molecular-weight HA at different growth temperatures. This GRAS B. subtilis strain can be applied in industry and provides a new strategy for production of HA with different molecular weights.

Programmed gRNA Removal System for CRISPR-Cas9-Mediated Multi-Round Genome Editing in Bacillus subtilis

CRISPR/Cas9 has become a simple and powerful genome editing tool for many organisms. However, multi-round genome editing should replace single-guide RNA (sgRNA) every round, which is laborious and time-consuming. Here, we have developed a multi-round genome editing system in which genome editing and the programmed removal of the sgRNA have sequentially occurred in a growth-dependent manner in Bacillus subtilis. The system contains two plasmids, one containing a cas9 gene and the other containing two sgRNAs and a donor DNA for homology directed repair (HDR). The two sgRNAs are chromosome-targeting (sgRNA_ct) and self-targeting (sgRNA_st) under the control of a constitutive promoter and sporulation-specific promoter, respectively. In the growth phase, the sgRNA_ct is transcribed and complexed with the Cas9 to edit the chromosomal target, while the sgRNA_st is transcribed in the sporulation phase and complexed with the Cas9 to attack its own plasmid. Therefore, the system automatically makes the cell ready for next-round genome editing during cultivation. The system was approved through the sequential deletion of eight extracellular protease genes in the B. subtilis, suggesting that it can be used for versatile applications in multi-round genome editing.

Modular Pathway Engineering of Bacillus subtilis To Promote De Novo Biosynthesis of Menaquinone-7

Menaquinone-7 (MK-7), a valuable vitamin K₂, plays an important role in the prevention of osteoporosis and cardiovascular calcification. We chose B. subtilis 168 as the chassis for the modular metabolic engineering design to promote the biosynthesis of MK-7. The biosynthetic pathway of MK-7 was categorized into four modules, namely, the MK-7 pathway (Module I), the shikimate (SA) pathway (Module II), the methylerythritol phosphate (MEP) pathway (Module III), and the glycerol metabolism pathway (Module IV). Overexpression of menA (Module I) resulted in 6.6 ± 0.1 mg/L of MK-7 after 120 h fermentation, which was 2.1-fold that of the starting strain BS168NU (3.1 ± 0.2 mg/L). Overexpression of aroA, aroD, and aroE (Module II) had a negative effect on the synthesis of MK-7. Simultaneous overexpression of dxs, dxr, yacM, and yacN (Module III) enabled the yield of MK-7 to 12.0 ± 0.1 mg/L. Moreover, overexpression of glpD (Module IV) resulted in an increase of the yield of MK-7 to 13.7 ± 0.2 mg/L. Furthermore, deletion of dhbB reduced the consumption of the intermediate metabolite isochorismate, thus promoting the yield of MK-7 to 15.4 ± 0.6 mg/L. Taken together, the final resulting strain MK3-MEP123-Gly2-Δ dhbB with simultaneous overexpression of menA, dxs, dxr, yacM-yacN, glpD and deletion of dhbB enabled the yield of MK-7 to 69.5 ± 2.8 mg/L upon 144 h fermentation in a 2 L baffled flask.

Genome Editing Method for the Anaerobic Magnetotactic Bacterium Desulfovibrio magneticus RS-1

Magnetotactic bacteria (MTB) are a group of organisms that form intracellular nanometer-scale magnetic crystals though a complex process involving lipid and protein scaffolds. These magnetic crystals and their lipid membranes, termed magnetosomes, are model systems for studying bacterial cell biology and biomineralization and are potential platforms for biotechnological applications. Due to a lack of genetic tools and unculturable representatives, the mechanisms of magnetosome formation in phylogenetically deeply branching MTB remain unknown. These MTB contain elongated bullet-/tooth-shaped magnetite and greigite crystals that likely form in a manner distinct from that of the cubooctahedral-shaped magnetite crystals of the genetically tractable MTB within the Alphaproteobacteria. Here, we present a method for genome editing in Desulfovibrio magneticus RS-1, a cultured representative of the deeply branching MTB of the class Deltaproteobacteria. This marks a crucial step in developing D. magneticus as a model for studying diverse mechanisms of magnetic particle formation by MTB.

Magnetosomes are complex bacterial organelles that serve as model systems for studying bacterial cell biology, biomineralization, and global iron cycling. Magnetosome biogenesis is primarily studied in two closely related Alphaproteobacteria of the genus Magnetospirillum that form cubooctahedral-shaped magnetite crystals within a lipid membrane. However, chemically and structurally distinct magnetic particles have been found in physiologically and phylogenetically diverse bacteria. Due to a lack of molecular genetic tools, the mechanistic diversity of magnetosome formation remains poorly understood. Desulfovibrio magneticus RS-1 is an anaerobic sulfate-reducing deltaproteobacterium that forms bullet-shaped magnetite crystals. A recent forward genetic screen identified 10 genes in the conserved magnetosome gene island of D. magneticus that are essential for its magnetic phenotype. However, this screen likely missed mutants with defects in crystal size, shape, and arrangement. Reverse genetics to target the remaining putative magnetosome genes using standard genetic methods of suicide vector integration have not been feasible due to the low transconjugation efficiency. Here, we present a reverse genetic method for targeted mutagenesis in D. magneticus using a replicative plasmid. To test this method, we generated a mutant resistant to 5-fluorouracil by making a markerless deletion of the upp gene that encodes uracil phosphoribosyltransferase. We also used this method for targeted marker exchange mutagenesis by replacing kupM, a gene identified in our previous screen as a magnetosome formation factor, with a streptomycin resistance cassette. Overall, our results show that targeted mutagenesis using a replicative plasmid is effective in D. magneticus and may also be applied to other genetically recalcitrant bacteria.

IMPORTANCE Magnetotactic bacteria (MTB) are a group of organisms that form intracellular nanometer-scale magnetic crystals though a complex process involving lipid and protein scaffolds. These magnetic crystals and their lipid membranes, termed magnetosomes, are model systems for studying bacterial cell biology and biomineralization and are potential platforms for biotechnological applications. Due to a lack of genetic tools and unculturable representatives, the mechanisms of magnetosome formation in phylogenetically deeply branching MTB remain unknown. These MTB contain elongated bullet-/tooth-shaped magnetite and greigite crystals that likely form in a manner distinct from that of the cubooctahedral-shaped magnetite crystals of the genetically tractable MTB within the Alphaproteobacteria. Here, we present a method for genome editing in Desulfovibrio magneticus RS-1, a cultured representative of the deeply branching MTB of the class Deltaproteobacteria. This marks a crucial step in developing D. magneticus as a model for studying diverse mechanisms of magnetic particle formation by MTB.

Enhanced Biosynthesis of 2-Deoxy-scyllo-inosose in Metabolically Engineered Bacillus subtilis Recombinants

2-Deoxy-scyllo-inosose (DOI) has been a valuable starting natural product for the manufacture of pharmaceuticals or chemical engineering resources such as pyranose catechol. DOI synthase, which uses glucose-6-phosphate (Glc6P) as a substrate for DOI biosynthesis, is indispensably involved in the initial stage of the biosynthesis of 2-deoxystreptamine-containing aminoglycoside antibiotics including butirosin, gentamicin, kanamycin, and tobramycin. A number of metabolically engineered recombinant strains of Bacillus subtilis were constructed here; either one or both genes pgi and pgcA that encode Glc6p isomerase and phosphoglucomutase, respectively, was (or were) disrupted in the sugar metabolic pathway of the host. After that, three different DOI synthase–encoding genes, which were artificially synthesized according to the codon preference of the B. subtilis host, were separately introduced into the engineered recombinants. The expression of a natural btrC gene, encoding DOI synthase in butirosin-producing B. circulans, in the heterologous host B. subtilis (BSDOI-2) generated approximately 2.3 g/L DOI, whereas expression of an artificially codon-optimized tobC gene, derived from tobramycin-producing Streptomyces tenebrarius, into the recombinant of B. subtilis (BSDOI-15) in which both genes pgi and pgcA are disrupted significantly enhanced the DOI titer: up to 37.2 g/L. Fed-batch fermentation by the BSDOI-15 recombinant using glycerol and glucose as a dual carbon source yielded the highest DOI titer (38.0 g/L). The development of engineered microbial cell factories empowered through convergence of metabolic engineering and synthetic biology should enable mass production of DOI. Thus, strain BSDOI-15 will surely be a useful contributor to the industrial manufacturing of various kinds of DOI-based pharmaceuticals and fine chemicals.

Restriction-deficient mutants and marker-less genomic modification for metabolic engineering of the solvent producer Clostridium saccharobutylicum

BACKGROUND

Clostridium saccharobutylicum NCP 262 is a solventogenic bacterium that has been used for the industrial production of acetone, butanol, and ethanol. The lack of a genetic manipulation system for C. saccharobutylicum currently limits (i) the use of metabolic pathway engineering to improve the yield, titer, and productivity of n-butanol production by this microorganism, and (ii) functional genomics studies to better understand its physiology.

RESULTS

In this study, a marker-less deletion system was developed for C. saccharobutylicum using the codBA operon genes from Clostridium ljungdahlii as a counterselection marker. The codB gene encodes a cytosine permease, while codA encodes a cytosine deaminase that converts 5-fluorocytosine to 5-fluorouracil, which is toxic to the cell. To introduce a marker-less genomic modification, we constructed a suicide vector containing: the catP gene for thiamphenicol resistance; the codBA operon genes for counterselection; fused DNA segments both upstream and downstream of the chromosomal deletion target. This vector was introduced into C. saccharobutylicum by tri-parental conjugation. Single crossover integrants are selected on plates supplemented with thiamphenicol and colistin, and, subsequently, double-crossover mutants whose targeted chromosomal sequence has been deleted were identified by counterselection on plates containing 5-fluorocytosine. Using this marker-less deletion system, we constructed the restriction-deficient mutant C. saccharobutylicum ΔhsdR1ΔhsdR2ΔhsdR3, which we named C. saccharobutylicum Ch2. This triple mutant exhibits high transformation efficiency with unmethylated DNA. To demonstrate its applicability to metabolic engineering, the method was first used to delete the xylB gene to study its role in xylose and arabinose metabolism. Furthermore, we also deleted the ptb and buk genes to create a butyrate metabolism-negative mutant of C. saccharobutylicum that produces n-butanol at high yield.

CONCLUSIONS

The plasmid vectors and the method introduced here, together with the restriction-deficient strains described in this work, for the first time, allow for efficient marker-less genomic modification of C. saccharobutylicum and, therefore, represent valuable tools for the genetic and metabolic engineering of this industrially important solvent-producing organism.

Unmarked genetic manipulation in Bacillus subtilis by natural co-transformation

Bacillus subtilis is well known as both a model organism and as a microbial cell factory. Simple and scarless gene modification is a desirable tool for basic research and industrial applications of B. subtilis. It has been demonstrated that naturally competent strains of B. subtilis can uptake multiple different DNA molecules, a phenomenon called co-transformation. Here, we describe a co-transformation-based method for generating unmarked mutants of B. subtilis. The PCR product containing the desired mutant allele is introduced into B. subtilis through co-transformation of the plasmid pUS20, which harbours a spectinomycin-resistant marker (Spc^r). The target mutation is acquired by screening transformants for integration of pUS20 by resistance to spectinomycin. Due to its unstable replication in B. subtilis, pUS20 is easily cured from transformants in the absence of spectinomycin. This method allows for point mutation delivery at frequencies of approximately 30%. Deletions and insertions of long DNA fragments can also be carried out efficiently using this method. Moreover, this method is also successful in Bacillus velezensis, indicating that it may be extended to other Bacillus species that can form natural competence.

Bacterial Genome Editing via a Designed Toxin-Antitoxin Cassette

Manipulating the bacterial genomes in an efficient manner is essential to biological and biotechnological research. Here, we reprogrammed the bacterial TA systems as the toxin counter-selectable cassette regulated by an antitoxin switch (TCCRAS) for genetic modifications in the extensively studied and utilized Gram-positive bacteria, B. subtilis and Corynebacterium glutamicum. In the five characterized type II TA systems, the RelBE complex can specifically and efficiently regulate cell growth and death by the conditionally controlled antitoxin RelB switch, thereby serving as a novel counter-selectable cassette to establish the TCCRAS system. Using a single vector, such a system has been employed to perform in-frame deletion, functional knock-in, gene replacement, precise point mutation, large-scale insertion, and especially, deletion of the fragments up to 194.9 kb in B. subtilis. In addition, the biosynthesis of lycopene was first achieved in B. subtilis using TCCRAS to integrate a 5.4-kb fusion cluster ( P _spac- crtI- crtE- crtB). The system was further adapted for gene knockdown and replacement, and large-scale deletion of the fragments up to 179.8 kb in C. glutamicum, with the mutation efficiencies increased by 0.8-1.0-fold compared to the conventional SacB method. TCCRAS thus holds promise as an effective and versatile genome-scale engineering technology for metabolic engineering and synthetic genomics research in a broad range of the Gram-positive bacteria.

DNA gyrase activity regulates DnaA-dependent replication initiation in Bacillus subtilis

In bacteria, initiation of DNA replication requires the DnaA protein. Regulation of DnaA association and activity at the origin of replication, oriC, is the predominant mechanism of replication initiation control. One key feature known to be generally important for replication is DNA topology. Although there have been some suggestions that topology may impact replication initiation, whether this mechanism regulates DnaA-mediated replication initiation is unclear. We found that the essential topoisomerase, DNA gyrase, is required for both proper binding of DnaA to oriC as well as control of initiation frequency in Bacillus subtilis. Furthermore, we found that the regulatory activity of gyrase in initiation is specific to DnaA and oriC. Cells initiating replication from a DnaA-independent origin, oriN, are largely resistant to gyrase inhibition by novobiocin, even at concentrations that compromise survival by up to four orders of magnitude in oriC cells. Furthermore, inhibition of gyrase does not impact initiation frequency in oriN cells. Additionally, deletion or overexpression of the DnaA regulator, YabA, significantly modulates sensitivity to gyrase inhibition, but only in oriC and not oriN cells. We propose that gyrase is a negative regulator of DnaA-dependent replication initiation from oriC, and that this regulatory mechanism is required for cell survival.

Engineering peptidoglycan degradation related genes of Bacillus subtilis for better fermentation processes

In this study, Bacillus subtilis 168 Δupp was engineered to change the bacterial shapes. Namely, some peptidoglycan hydrolase related genes were inactivated individually or in different combinations, including sigD, lytE, lytF, lytC, lytD and lytG. Inactivations of these genes resulted in various intensities of blockages on cell division, leading to elongation of bacterial cells. The resulted fiber phenotypes showed different lengths ranging from tens of microns to several millimeters. Mutants with multiple gene inactivations such as ΔsigDΔlytEΔlytD showed more easily precipitated phenomenon, obviously increased growth rate, more sensitive to antibiotics and improved α-amylase production compared with that of B. subtilis 168 Δupp. Mutants ΔsigDΔlytEΔlytD and ΔsigDΔlytEΔlytCΔlytD also showed an increased tolerance to high osmotic pressure of sodium chloride, allowing unsterile fermentation, all of which contributes to reduced processing cost.

The restriction modification system of Bacillus licheniformis MS1 and generation of a readily transformable deletion mutant

Restriction modification systems (R-M systems), consisting of a restriction endonuclease and a cognate methyltransferase, constitute an effective means of a cell to protect itself from foreign DNA. Identification, characterization, and deletion of the restriction modification system BliMSI, a putative isoschizomer of ClaI from Caryophanon latum, were performed in the wild isolate Bacillus licheniformis MS1. BliMSI was produced as recombinant protein in Escherichia coli, purified, and in vitro analysis demonstrated identical restriction endonuclease activity as for ClaI. A recombinant E. coli strain, expressing the heterologous bliMSIM gene, was constructed and used as the host for in vivo methylation of plasmids prior to their introduction into B. licheniformis to improve transformation efficiencies. The establishment of suicide plasmids in the latter was rendered possible. The subsequent deletion of the restriction endonuclease encoding gene, bliMSIR, caused doubled transformation efficiencies in the respective mutant B. licheniformis MS2 (∆bliMSIR). Along with above in vivo methylation, the establishment of further gene deletions (∆upp, ∆yqfD) was performed. The constructed triple mutant (∆bliMSIR, ∆upp, ∆yqfD) enables rapid genome manipulation, a requirement for genetic engineering of industrially important strains.

The contribution of bacterial genome engineering to sustainable development

The United Nations’ Sustainable Development Goals define important challenges for the prosperous development of mankind. To reach several of these goals, among them the production of value‐added compounds, improved economic and ecologic balance of production processes, prevention of climate change and protection of ecosystems, the use of engineered bacteria can make valuable contributions. We discuss the strategies for genome engineering and how they can be applied to meet the United Nations’ goals for sustainable development.

Conversion of Glycerol to 3-Hydroxypropanoic Acid by Genetically Engineered Bacillus subtilis

3-Hydroxypropanoic acid (3-HP) is an important biomass-derivable platform chemical that can be converted into a number of industrially relevant compounds. There have been several attempts to produce 3-HP from renewable sources in cell factories, focusing mainly on Escherichia coli, Klebsiella pneumoniae, and Saccharomyces cerevisiae. Despite the significant progress made in this field, commercially exploitable large-scale production of 3-HP in microbial strains has still not been achieved. In this study, we investigated the potential of Bacillus subtilis as a microbial platform for bioconversion of glycerol into 3-HP. Our recombinant B. subtilis strains overexpress the two-step heterologous pathway containing glycerol dehydratase and aldehyde dehydrogenase from K. pneumoniae. Genetic engineering, driven by in silico optimization, and optimization of cultivation conditions resulted in a 3-HP titer of 10 g/L, in a standard batch cultivation. Our findings provide the first report of successful introduction of the biosynthetic pathway for conversion of glycerol into 3-HP in B. subtilis. With this relatively high titer in batch, and the robustness of B. subtilis in high density fermentation conditions, we expect that our production strains may constitute a solid basis for commercial production of 3-HP.

Study of the tensile properties of individual multicellular fibres generated by Bacillus subtilis

Multicellular fibres formed by Bacillus subtilis (B. subtilis) are attracting interest because of their potential application as degradable biomaterials. However, mechanical properties of individual fibres remain unknown because of their small dimensions. Herein, a new approach is developed to investigate the tensile properties of individual fibres with an average diameter of 0.7 μm and a length range of 25.7–254.3 μm. Variations in the tensile strengths of fibres are found to be the result of variable interactions among pairs of microbial cells known as septa. Using Weibull weakest-link model to study this mechanical variability, we predict the length effect of the sample. Moreover, the mechanical properties of fibres are found to depend highly on relative humidity (RH), with a brittle–ductile transition occurring around RH = 45%. The elastic modulus is 5.8 GPa in the brittle state, while decreases to 62.2 MPa in the ductile state. The properties of fibres are investigated by using a spring model (RH < 45%) for its elastic behaviour, and the Kelvin–Voigt model (RH > 45%) for the time-dependent response. Loading-unloading experiments and numerical calculations demonstrate that necking instability comes from structural changes (septa) and viscoelasticity dominates the deformation of fibres at high RH.

Intramembrane protease RasP boosts protein production in Bacillus

Background

The microbial cell factory Bacillus subtilis is a popular industrial platform for high-level production of secreted technical enzymes. Nonetheless, the effective secretion of particular heterologous enzymes remains challenging. Over the past decades various studies have tackled this problem, and major improvements were achieved by optimizing signal peptides or removing proteases involved in product degradation. On the other hand, serious bottlenecks in the protein export process per se remained enigmatic, especially for protein secretion at commercially significant levels by cells grown to high density. The aim of our present study was to assess the relevance of the intramembrane protease RasP for high-level protein production in B. subtilis.

Results

Deletion of the rasP gene resulted in reduced precursor processing and extracellular levels of the overproduced α-amylases AmyE from B. subtilis and AmyL from Bacillus licheniformis. Further, secretion of the overproduced serine protease BPN’ from Bacillus amyloliquefaciens was severely impaired in the absence of RasP. Importantly, overexpression of rasP resulted in threefold increased production of a serine protease from Bacillus clausii, and 2.5- to 10-fold increased production of an AmyAc α-amylase from Paenibacillus curdlanolyticus, depending on the culture conditions. Of note, growth defects due to overproduction of the two latter enzymes were suppressed by rasP-overexpression.

Conclusion

Here we show that an intramembrane protease, RasP, sets a limit to high-level production of two secreted heterologous enzymes that are difficult to produce in the B. subtilis cell factory. This finding was unexpected and suggests that proteolytic membrane sanitation is key to effective enzyme production in Bacillus.

CRISPR/Cas9 Editing of the Bacillus subtilis Genome

A fundamental procedure for most modern biologists is the genetic manipulation of the organism under study. Although many different methods for editing bacterial genomes have been used in laboratories for decades, the adaptation of CRISPR/Cas9 technology to bacterial genetics has allowed researchers to manipulate bacterial genomes with unparalleled facility. CRISPR/Cas9 has allowed for genome edits to be more precise, while also increasing the efficiency of transferring mutations into a variety of genetic backgrounds. As a result, the advantages are realized in tractable organisms and organisms that have been refractory to genetic manipulation. Here, we describe our method for editing the genome of the bacterium Bacillus subtilis. Our method is highly efficient, resulting in precise, markerless mutations. Further, after generating the editing plasmid, the mutation can be quickly introduced into several genetic backgrounds, greatly increasing the speed with which genetic analyses may be performed.

pheS * , an effective host-genotype-independent counter-selectable marker for marker-free chromosome deletion in Bacillus amyloliquefaciens

Aside from applications in the production of commercial enzymes and metabolites, Bacillus amyloliquefaciens is also an important group of plant growth-promoting rhizobacteria that supports plant growth and suppresses phytopathogens. A host-genotype-independent counter-selectable marker would enable rapid genetic manipulation and metabolic engineering, accelerating the study of B. amyloliquefaciens and its development as both a microbial cell factory and plant growth-promoting rhizobacteria. Here, a host-genotype-independent counter-selectable marker pheS ^* was constructed through a point mutation of the gene pheS, which encodes the α-subunit of phenylalanyl-tRNA synthetase in Bacillus subtilis strain 168. In the presence of 5 mM p-chloro-phenylalanine, 100 % of B. amyloliquefaciens strain SQR9 cells carrying pheS ^* were killed, whereas the wild-type strain SQR9 showed resistance to p-chloro-phenylalanine. A simple pheS ^* and overlap-PCR-based strategy was developed to create the marker-free deletion of the amyE gene as well as a 37-kb bmy cluster in B. amyloliquefaciens SQR9. The effectiveness of pheS ^* as a counter-selectable marker in B. amyloliquefaciens was further confirmed through the deletion of amyE genes in strains B. amyloliquefaciens FZB42 and NJN-6. In addition, the potential use of pheS ^* in other Bacillus species was preliminarily assessed. The expression of PheS^* in B. subtilis strain 168 and B. cereus strain ATCC 14579 caused pronounced sensitivity of both hosts to p-chloro-phenylalanine, indicating that pheS ^* could be used as a counter-selectable marker (CSM) in these strains.

The Blueprint of a Minimal Cell: MiniBacillus

Bacillus subtilis is one of the best-studied organisms. Due to the broad knowledge and annotation and the well-developed genetic system, this bacterium is an excellent starting point for genome minimization with the aim of constructing a minimal cell. We have analyzed the genome of B. subtilis and selected all genes that are required to allow life in complex medium at 37°C. This selection is based on the known information on essential genes and functions as well as on gene and protein expression data and gene conservation. The list presented here includes 523 and 119 genes coding for proteins and RNAs, respectively. These proteins and RNAs are required for the basic functions of life in information processing (replication and chromosome maintenance, transcription, translation, protein folding, and secretion), metabolism, cell division, and the integrity of the minimal cell. The completeness of the selected metabolic pathways, reactions, and enzymes was verified by the development of a model of metabolism of the minimal cell. A comparison of the MiniBacillus genome to the recently reported designed minimal genome of Mycoplasma mycoides JCVI-syn3.0 indicates excellent agreement in the information-processing pathways, whereas each species has a metabolism that reflects specific evolution and adaptation. The blueprint of MiniBacillus presented here serves as the starting point for a successive reduction of the B. subtilis genome.

Substrate Specificity of the Bacillus subtilis BY-Kinase PtkA Is Controlled by Alternative Activators: TkmA and SalA

Bacterial protein-tyrosine kinases (BY-kinases) are known to regulate different aspects of bacterial physiology, by phosphorylating cellular protein substrates. Physiological cues that trigger BY-kinases activity are largely unexplored. In Proteobacteria, BY-kinases contain a cytosol-exposed catalytic domain and a transmembrane activator domain in a single polypeptide chain. In Firmicutes, the BY-kinase catalytic domain and the transmembrane activator domain exist as separate polypeptides. We have previously speculated that this architecture might enable the Firmicutes BY-kinases to interact with alternative activators, and thus account for the observed ability of these kinases to phosphorylate several distinct classes of protein substrates. Here, we present experimental evidence that supports this hypothesis. We focus on the model Firmicute-type BY-kinase PtkA from Bacillus subtilis, known to phosphorylate several different protein substrates. We demonstrate that the transcriptional regulator SalA, hitherto known as a substrate of PtkA, can also act as a PtkA activator. In doing so, SalA competes with the canonical PtkA activator, TkmA. Our results suggest that the respective interactions of SalA and TkmA with PtkA favor phosphorylation of different protein substrates in vivo and in vitro. This observation may contribute to explaining how specificity is established in the seemingly promiscuous interactions of BY-kinases with their cellular substrates.

Editing of the Bacillus subtilis Genome by the CRISPR-Cas9 System

The clustered regularly interspaced short palindromic repeat (CRISPR)-associated (Cas) systems are adaptive immune systems of bacteria. A type II CRISPR-Cas9 system from Streptococcus pyogenes has recently been developed into a genome engineering tool for prokaryotes and eukaryotes. Here, we present a single-plasmid system which allows efficient genome editing of Bacillus subtilis The plasmid pJOE8999 is a shuttle vector that has a pUC minimal origin of replication for Escherichia coli, the temperature-sensitive replication origin of plasmid pE194(ts) for B. subtilis, and a kanamycin resistance gene working in both organisms. For genome editing, it carries the cas9 gene under the control of the B. subtilis mannose-inducible promoter PmanP and a single guide RNA (sgRNA)-encoding sequence transcribed via a strong promoter. This sgRNA guides the Cas9 nuclease to its target. The 20-nucleotide spacer sequence at the 5' end of the sgRNA sequence, responsible for target specificity, is located between BsaI sites. Thus, the target specificity is altered by changing the spacer sequences via oligonucleotides fitted between the BsaI sites. Cas9 in complex with the sgRNA induces double-strand breaks (DSBs) at its target site. Repair of the DSBs and the required modification of the genome are achieved by adding homology templates, usually two PCR fragments obtained from both sides of the target sequence. Two adjacent SfiI sites enable the ordered integration of these homology templates into the vector. The function of the CRISPR-Cas9 vector was demonstrated by introducing two large deletions in the B. subtilis chromosome and by repair of the trpC2 mutation of B. subtilis 168.

IMPORTANCE

In prokaryotes, most methods used for scarless genome engineering are based on selection-counterselection systems. The disadvantages are often the lack of a suitable counterselection marker, the toxicity of the compounds needed for counterselection, and the requirement of certain mutations in the target strain. CRISPR-Cas systems were recently developed as important tools for genome editing. The single-plasmid system constructed for the genome editing of B. subtilis overcomes the problems of counterselection methods. It allows deletions and introduction of point mutations. It is easy to handle and very efficient, and it may be adapted for use in other firmicutes.