Bacillus subtilis genome editing using ssDNA with short homology regions

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.

Conjugation-Mediated Plasmid Transfer Enables Genetic Modification of Diverse Bacillus Species

Performing genetic manipulations in Bacillus strains is often hindered by difficulty in identifying conditions appropriate for DNA uptake. This shortcoming limits our understanding of the functional diversity within this genus and the practical application of new strains. We have developed a simple method for increasing the genetic tractability of Bacillus spp. through conjugation-mediated plasmid transfer via a diaminopimelic acid (DAP) auxotrophic Escherichia coli donor strain. We observe transfer into representatives of the Bacillus clades subtilis, cereus, galactosidilyticus, and Priestia megaterium and successfully applied this protocol to 9 out of 12 strains attempted. We utilized the BioBrick 2.0 plasmids pECE743 and pECE750, as well as the CRISPR plasmid pJOE9734.1, to generate a xylose-inducible green-fluorescent protein (GFP)-expressing conjugal vector, pEP011. The use of xylose-inducible GFP ensures ease of confirming transconjugants, which enables users to quickly rule out false positives. Additionally, our plasmid backbone offers the flexibility to be used in other contexts, including transcriptional fusions and overexpression, with only a few modifications. IMPORTANCE Bacillus species are widely used to produce proteins and to understand microbial differentiation. Unfortunately, outside a few lab strains, genetic manipulation is difficult and can prevent thorough dissection of useful phenotypes. We developed a protocol that utilizes conjugation (plasmids that initiate their own transfer) to introduce plasmids into a diverse range of Bacillus spp. This will facilitate a deeper study of wild isolates for both industrial and pure research uses.

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.

Recombineering in Non-Model Bacteria

The technology of recombineering, in vivo genetic engineering, was initially developed in Escherichia coli and uses bacteriophage-encoded homologous recombination proteins to efficiently recombine DNA at short homologies (35 to 50 nt). Because the technology is homology driven, genomic DNA can be modified precisely and independently of restriction site location. Recombineering uses linear DNA substrates that are introduced into the cell by electroporation; these can be PCR products, synthetic double-strand DNA (dsDNA), or single-strand DNA (ssDNA). Here we describe the applications, challenges, and factors affecting ssDNA and dsDNA recombineering in a variety of non-model bacteria, both Gram-negative and -positive, and recent breakthroughs in the field. We list different microbes in which the widely used phage λ Red and Rac RecET recombination systems have been used for in vivo genetic engineering. New homologous ssDNA and dsDNA recombineering systems isolated from non-model bacteria are also described. The Basic Protocol outlines a method for ssDNA recombineering in the non-model species of Shewanella. The Alternate Protocol describes the use of CRISPR/Cas as a counter-selection system in conjunction with recombineering to enhance recovery of recombinants. We provide additional background information, pertinent considerations for experimental design, and parameters critical for success. The design of ssDNA oligonucleotides (oligos) and various internet-based tools for oligo selection from genome sequences are also described, as is the use of oligo-mediated recombination. This simple form of genome editing uses only ssDNA oligo(s) and does not require an exogenous recombination system. The information presented here should help researchers identify a recombineering system suitable for their microbe(s) of interest. If no system has been characterized for a specific microbe, researchers can find guidance in developing a recombineering system from scratch. We provide a flowchart of decision-making paths for strategically applying annealase-dependent or oligo-mediated recombination in non-model and undomesticated bacteria. © 2022 Wiley Periodicals LLC. This article has been contributed to by U.S. Government employees and their work is in the public domain in the USA. Basic Protocol: ssDNA recombineering in Shewanella species Alternate Protocol: ssDNA recombineering coupled to CRISPR/Cas9 in Shewanella species.

Simultaneous Multiplex Genome Engineering via Accelerated Natural Transformation in Bacillus subtilis

Multiplex engineering at the scale of whole genomes has become increasingly important for synthetic biology and biotechnology applications. Although several methods have been reported for engineering microbe genomes, their use is limited by their complex procedures using multi-cycle transformations. Natural transformation, involving in species evolution by horizontal gene transfer in many organisms, indicates its potential as a genetic tool. Here, we aimed to develop simultaneous multiplex genome engineering (SMGE) for the simple, rapid, and efficient design of bacterial genomes via one-step of natural transformation in Bacillus subtilis. The transformed DNA, competency factors, and recombinases were adapted to improved co-editing frequencies above 27-fold. Single to octuplet variants with genetic diversity were simultaneously generated using all-in-one vectors harboring multi-gene cassettes. To demonstrate its potential application, the tyrosine biosynthesis pathway was further optimized for producing commercially important resveratrol by high-throughput screening of variant pool in B. subtilis. SMGE represents an accelerated evolution platform that generates diverse multiplex mutations for large-scale genetic engineering and synthetic biology in B. subtilis.

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.

Lox'd in translation: contradictions in the nomenclature surrounding common lox-site mutants and their implications in experiments

The Cre-Lox system is a highly versatile and powerful DNA recombinase mechanism, mainly used in genetic engineering to insert or remove desired DNA sequences. It is widely utilized across multiple fields of biology, with applications ranging from plants, to mammals, to microbes. A key feature of this system is its ability to allow recombination between mutant lox sites. Two of the most commonly used mutant sites are named lox66 and lox71, which recombine to create a functionally inactive double mutant lox72 site. However, a large portion of the published literature has incorrectly annotated these mutant lox sites, which in turn can lead to difficulties in replication of methods, design of proper vectors and confusion over the proper nomenclature. Here, we demonstrate common errors in annotations, the impacts they can have on experimental viability, and a standardized naming convention. We also show an example of how this incorrect annotation can induce toxic effects in bacteria that lack optimal DNA repair systems, exemplified by Mycoplasma pneumoniae.

Positioning Bacillus subtilis as terpenoid cell factory

Increasing demands for bioactive compounds have motivated researchers to employ micro‐organisms to produce complex natural products. Currently, Bacillus subtilis has been attracting lots of attention to be developed into terpenoids cell factories due to its generally recognized safe status and high isoprene precursor biosynthesis capacity by endogenous methylerythritol phosphate (MEP) pathway. In this review, we describe the up‐to‐date knowledge of each enzyme in MEP pathway and the subsequent steps of isomerization and condensation of C5 isoprene precursors. In addition, several representative terpene synthases expressed in B. subtilis and the engineering steps to improve corresponding terpenoids production are systematically discussed. Furthermore, the current available genetic tools are mentioned as along with promising strategies to improve terpenoids in B. subtilis, hoping to inspire future directions in metabolic engineering of B. subtilis for further terpenoid cell factory development.

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.

A regulated synthetic operon facilitates stable overexpression of multigene terpenoid pathway in Bacillus subtilis

The creation of microbial cell factories for sustainable production of natural products is important for medical and industrial applications. This requires stable expression of biosynthetic pathways in a host organism with favorable fermentation properties such as Bacillus subtilis. The aim of this study is to construct B. subtilis strains that produce valuable terpenoid compounds by overexpressing the innate methylerythritol phosphate (MEP) pathway. A synthetic operon allowing the concerted and regulated expression of multiple genes was developed. Up to 8 genes have been combined in this operon and a stably inherited plasmid-based vector was constructed resulting in a high production of C30 carotenoids. For this, two vectors were examined, one with rolling circle replication and another with theta replication. Theta-replication constructs were clearly superior in structural and segregational stability compared to rolling circle constructs. A strain overexpressing all eight genes of the MEP pathway on a theta-replicating plasmid clearly produced the highest level of carotenoids. The level of transcription for each gene in the operon was similar as RT-qPCR analysis indicated. Hence, that corresponding strain can be used as a stable cell factory for production of terpenoids. This is the first report of merging and stably expressing this large-size operon (eight genes) from a plasmid-based system in B. subtilis enabling high C30 carotenoid production.

Development and characterization of a CRISPR/Cas9n-based multiplex genome editing system for Bacillus subtilis

BACKGROUND

Metabolic engineering has expanded from a focus on designs requiring a small number of genetic modifications to increasingly complex designs driven by advances in multiplex genome editing technologies. However, simultaneously modulating multiple genes on the chromosome remains challenging in Bacillus subtilis. Thus, developing an efficient and convenient method for B. subtilis multiplex genome editing is imperative.

RESULTS

Here, we developed a CRISPR/Cas9n-based multiplex genome editing system for iterative genome editing in B. subtilis. This system enabled us to introduce various types of genomic modifications with more satisfying efficiency than using CRISPR/Cas9, especially in multiplex gene editing. Our system achieved at least 80% efficiency for 1-8 kb gene deletions, at least 90% efficiency for 1-2 kb gene insertions, near 100% efficiency for site-directed mutagenesis, 23.6% efficiency for large DNA fragment deletion and near 50% efficiency for three simultaneous point mutations. The efficiency for multiplex gene editing was further improved by regulating the nick repair mechanism mediated by ligD gene, which finally led to roughly 65% efficiency for introducing three point mutations on the chromosome. To demonstrate its potential, we applied our system to simultaneously fine-tune three genes in the riboflavin operon and significantly improved the production of riboflavin in a single cycle.

CONCLUSIONS

We present not only the iterative CRISPR/Cas9n system for B. subtilis but also the highest efficiency for simultaneous modulation of multiple genes on the chromosome in B. subtilis reported to date. We anticipate this CRISPR/Cas9n mediated system to greatly enhance the optimization of diverse biological systems via metabolic engineering and synthetic biology.

Highly Efficient Genome Engineering in Bacillus anthracis and Bacillus cereus Using the CRISPR/Cas9 System

Genome editing is an effective tool for the functional examination of bacterial genes and for live attenuated vaccine construction. Here, we report a method to edit the genomic DNA of Bacillus anthracis and Bacillus cereus using the clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein (Cas)9 system. Using two prophages in B. anthracis as targets, large-fragment deletion mutants were achieved with rates of 100 or 20%. In B. cereus, we successfully introduced precise point mutations into plcR, with phenotypic assays showing that the resulting mutants lost hemolytic and phospholipase enzyme activities similar to B. anthracis, which is a natural plcR mutant. Our study indicates that CRISPR/Cas9 is a powerful genetic tool for genome editing in the Bacillus cereus group, and can efficiently modify target genes without the need for residual foreign DNA such as antibiotic selection markers. This system could be developed for use in the generation of marker-free live anthrax vaccines or for safer construction of microbiological candidate-based recombinant B. cereus.

Development of a novel strategy for robust synthetic bacterial promoters based on a stepwise evolution targeting the spacer region of the core promoter in Bacillus subtilis

Background

Promoter evolution by synthetic promoter library (SPL) is a powerful approach to development of functional synthetic promoters to synthetic biology. However, it requires much tedious and time-consuming screenings because of the plethora of different variants in SPL. Actually, a large proportion of mutants in the SPL are significantly lower in strength, which contributes only to fabrication of a promoter library with a continuum of strength. Thus, to effectively obtain the evolved synthetic promoter exhibiting higher strength, it is essential to develop novel strategies to construct mutant library targeting the pivotal region rather than the arbitrary region of the template promoter. In this study, a strategy termed stepwise evolution targeting the spacer of core promoter (SETarSCoP) was established in Bacillus subtilis to effectively evolve the strength of bacterial promoter.

Results

The native promoter, P_srfA, from B. subtilis, which exhibits higher strength than the strong promoter P43, was set as the parental template. According to the comparison of conservation of the spacer sequences between − 35 box and − 10 box among a set of strong and weak native promoter, it revealed that 7-bp sequence immediately upstream of the − 10 box featured in the regulation of promoter strength. Based on the conservative feature, two rounds of consecutive evolution were performed targeting the hot region of P_srfA. In the first round, a primary promoter mutation library (pPML) was constructed by mutagenesis targeting the 3-bp sequence immediately upstream of the − 10 box of the P_srfA. Subsequently, four evolved mutants from pPML were selected to construction of four secondary promoter mutation libraries (sPMLs) based on mutagenesis of the 4-bp sequence upstream of the first-round target. After the consecutive two-step evolution, the mutant P_BH4 was identified and verified to be a highly evolved synthetic promoter. The strength of P_BH4 was higher than P_srfA by approximately 3 times. Moreover, P_BH4 also exhibited broad suitability for different cargo proteins, such as β-glucuronidase and nattokinase. The proof-of-principle test showed that SETarSCoP successfully evolved both constitutive and inducible promoters.

Conclusion

Comparing with the commonly used SPL strategy, SETarSCoP facilitates the evolution process to obtain strength-evolved synthetic bacterial promoter through fabrication and screening of small-scale mutation libraries. This strategy will be a promising method to evolve diverse bacterial promoters to expand the toolbox for synthetic biology.

Electronic supplementary material

The online version of this article (10.1186/s12934-019-1148-3) contains supplementary material, which is available to authorized users.

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.

An alternative genome-integrated method for undomesticated Bacillus subtilis and related species

Given their applicability in genetic engineering, undomesticated Bacillus strains are extensively used as non-natural hosts for chemical production due to their high tolerance of toxic substrates or products. However, they are difficult to genomically modify due to their low transformation efficiencies. In this study, the Bacillus-E. coli shuttle vector pHY300PLK, which is widely used in gram-positive bacteria, was adopted for genome integration in organic solvent-tolerant Bacillus isolates. The Bacillus-replicative vector was used to deliver homologous recombinant DNA and propagate itself inside the host cell, increasing the likelihood of genome integration of the recombinant DNA. Then, the unintegrated vectors were cured by cell cultivation in antibiotic-free medium with facilitation of nickel ions. The developed protocol was successfully demonstrated and validated by the disruption of amyE gene in B. subtilis 168. With an improved clonal selection protocol, the probability of clonal selection of the amyE::cat genome-integrated mutants was increased up to 42.0 ± 10.2%. Genome integration in undomesticated, organic solvent tolerant Bacillus strains was also successfully demonstrated with amyE as well as proB gene creating the gene-disrupted mutants with the corresponding phenotype and genotype. Not only was this technique effectively applied to several strains of undomesticated B. subtilis, but it was also successfully applied to B. cereus. This study validates the possibility of the application of Bacillus-replicative vector as well as the developed protocol in a variety of genome modification of undomesticated Bacillus species.

A rapid and versatile tool for genomic engineering in Lactococcus lactis

BACKGROUND

Lactococcus lactis is one of the most extensively characterized lactic acid bacteria, from physiological traits to industrial exploitation. Since last decade, L. lactis has been developed into cell factories for the production of bioactive compounds such as enzymes, vaccine antigens and natural products. However, its precise and efficient genome editing tools is still required to make L. lactis more suitable candidate for engineered functionality.

RESULTS

A high active recombinase, RecT of Enterococcus faecalis ATCC14506, was selected from six candidates and mediated homologous recombination between single-stranded DNA (ssDNA) and the L. lactis chromosomal rpoB locus with an efficiency of 100% after rifampin selection. To screen mutants without an externally selectable phenotype, the CRISPR/Cas9 system was used for counterselection, yielding an upp mutant with an efficiency of 46%. By optimization of the copy number of plasmid carrying the CRISPR/Cas9 system and the length of spacer sequence, the off-target efficiency of the recA, galK, hemN and noxD genes were eliminated. The ability of this optimized tool to perform sequential point mutation was demonstrated using the upp and galK gene loci as targets with improved efficiencies > 75%. Moreover, seamless genomic DNA deletions (50/100 bp) or insertion (a loxP site, 34 bp) was efficiently accomplished within 72 h.

CONCLUSIONS

The work provided a rapid, versatile and precise tool for L. lactis genomic engineering by combination of ssDNA recombineering with improved CRISPR/Cas9 counterselection. This tool will simplify the production of isogenic strains for assessment of gene function or construction of biosynthetic host.

Targeted Nucleotide Editing Technologies for Microbial Metabolic Engineering

Since the emergence of programmable RNA-guided nucleases based on clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein (Cas) systems, genome editing technologies have become a simplified and versatile tool for genome editing in various organisms and cell types. Although genome editing enables efficient genome manipulations, such as gene disruptions, gene knockins, and chromosomal translocations via DNA double-strand break (DSB) repair in eukaryotes, DSBs induced by the CRISPR/Cas system are lethal or severely toxic to many microorganisms. Therefore, in many prokaryotes, including industrially useful microbes, the CRISPR/Cas system is often used as a negative selection component in combination with recombineering or other related strategies. Novel and revolutionary technologies have been recently developed to re-write targeted nucleotides (C:G to T:A and A:T to G:C) without DSBs and donor DNA templates. These technologies rely on the recruitment of deaminases at specific target loci using the nuclease-deficient CRISPR/Cas system. Here, the authors review and compare CRISPR-based genome editing, current base editing platforms and their spectra. The authors discuss how these technologies can be applied in various aspects of microbial metabolic engineering to overcome barriers to cellular regulation in prokaryotes.

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.

Deaminase-mediated multiplex genome editing in Escherichia coli

In eukaryotes, the CRISPR-Cas9 system has now been widely used as a revolutionary genome engineering tool1, 2. However, in prokaryotes, the use of nuclease-mediated genome editing tools has been limited to negative selection for the already modified cells because of its lethality3, 4. Here, we report on deaminase-mediated targeted nucleotide editing (Target-AID) 5 adopted in Escherichia coli. Cytidine deaminase PmCDA1 fused to the nuclease-deficient CRISPR-Cas9 system achieved specific point mutagenesis at the target sites in E. coli by introducing cytosine mutations without compromising cell growth. The cytosine-to-thymine substitutions were induced mainly within an approximately five-base window of target sequences on the protospacer adjacent motif-distal side, which can be shifted depending on the length of the single guide RNA sequence. Use of a uracil DNA glycosylase inhibitor 6 in combination with a degradation tag (LVA tag) 7 resulted in a robustly high mutation efficiency, which allowed simultaneous multiplex editing of six different genes. The major multi-copy transposase genes that consist of at least 41 loci were also simultaneously edited by using four target sequences. As this system does not rely on any additional or host-dependent factors, it may be readily applicable to a wide range of bacteria.

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.

Construction of a high-efficiency cloning system using the Golden Gate method and I-SceI endonuclease for targeted gene replacement in Bacillus anthracis

To investigate gene function in Bacillus anthracis, a high-efficiency cloning system is required with an increased rate of allelic exchange. Golden Gate cloning is a molecular cloning strategy allowing researchers to simultaneously and directionally assemble multiple DNA fragments to construct target plasmids using type IIs restriction enzymes and T4 DNA ligase in the same reaction system. Here, a B. anthracis S-layer protein EA1 allelic exchange vector was successfully constructed using the Golden Gate method. No new restriction sites were introduced into this knockout vector, and seamless assembly of the DNA fragments was achieved. To elevate the efficiency of homologous recombination between the allelic exchange vector and chromosomal DNA, we introduced an I-SceI site into the allelic exchange vector. The eag gene was successfully knocked out in B. anthracis using this vector. Simultaneously, the allelic exchange vector construction method was developed into a system for generating B. anthracis allelic exchange vectors. To verify the effectiveness of this system, some other allelic exchange vectors were constructed and gene replacements were performed in B. anthracis. It is speculated that this gene knockout vector construction system and high-efficiency targeted gene replacement using I-SceI endonuclease can be applied to other Bacillus spp.

Toward Genome-Based Metabolic Engineering in Bacteria

Prokaryotes modified stably on the genome are of great importance for production of fine and commodity chemicals. Traditional methods for genome engineering have long suffered from imprecision and low efficiencies, making construction of suitable high-producer strains laborious. Here, we review the recent advances in discovery and refinement of molecular precision engineering tools for genome-based metabolic engineering in bacteria for chemical production, with focus on the λ-Red recombineering and the clustered regularly interspaced short palindromic repeats/Cas9 nuclease systems. In conjunction, they enable the integration of in vitro-synthesized DNA segments into specified locations on the chromosome and allow for enrichment of rare mutants by elimination of unmodified wild-type cells. Combination with concurrently developing improvements in important accessory technologies such as DNA synthesis, high-throughput screening methods, regulatory element design, and metabolic pathway optimization tools has resulted in novel efficient microbial producer strains and given access to new metabolic products. These new tools have made and will likely continue to make a big impact on the bioengineering strategies that transform the chemical industry.

Development of a CRISPR-Cas9 Tool Kit for Comprehensive Engineering of Bacillus subtilis

The establishment of a clustered regularly interspaced short palindromic repeat (CRISPR)-Cas9 system for strain construction in Bacillus subtilis is essential for its progression toward industrial utility. Here we outline the development of a CRISPR-Cas9 tool kit for comprehensive genetic engineering in B. subtilis In addition to site-specific mutation and gene insertion, our approach enables continuous genome editing and multiplexing and is extended to CRISPR interference (CRISPRi) for transcriptional modulation. Our tool kit employs chromosomal expression of Cas9 and chromosomal transcription of guide RNAs (gRNAs) using a gRNA transcription cassette and counterselectable gRNA delivery vectors. Our design obviates the need for multicopy plasmids, which can be unstable and impede cell viability. Efficiencies of up to 100% and 85% were obtained for single and double gene mutations, respectively. Also, a 2.9-kb hyaluronic acid (HA) biosynthetic operon was chromosomally inserted with an efficiency of 69%. Furthermore, repression of a heterologous reporter gene was achieved, demonstrating the versatility of the tool kit. The performance of our tool kit is comparable with those of systems developed for Escherichia coli and Saccharomyces cerevisiae, which rely on replicating vectors to implement CRISPR-Cas9 machinery.

IMPORTANCE

In this paper, as the first approach, we report implementation of the CRISPR-Cas9 system in Bacillus subtilis, which is recognized as a valuable host system for biomanufacturing. The study enables comprehensive engineering of B. subtilis strains with virtually any desired genotypes/phenotypes and biochemical properties for extensive industrial application.

Novel Routes for Improving Biocontrol Activity of Bacillus Based Bioinoculants

Biocontrol (BC) formulations prepared from plant-growth-promoting bacteria are increasingly applied in sustainable agriculture. Especially inoculants prepared from endospore-forming Bacillus strains have been proven as efficient and environmental-friendly alternative to chemical pesticides due to their long shelf life, which is comparable with that of agrochemicals. However, these formulations of the first generation are sometimes hampered in their action and do not fulfill in each case the expectations of the appliers. In this review we use the well-known plant-associated Bacillus amyloliquefaciens type strain FZB42 as example for the successful application of different techniques offered today by comparative, evolutionary and functional genomics, site-directed mutagenesis and strain construction including marker removal, for paving the way for preparing a novel generation of BC agents.

Genomes by design

Next-generation DNA sequencing has revealed the complete genome sequences of numerous organisms, establishing a fundamental and growing understanding of genetic variation and phenotypic diversity. Engineering at the gene, network and whole-genome scale aims to introduce targeted genetic changes both to explore emergent phenotypes and to introduce new functionalities. Expansion of these approaches into massively parallel platforms establishes the ability to generate targeted genome modifications, elucidating causal links between genotype and phenotype, as well as the ability to design and reprogramme organisms. In this Review, we explore techniques and applications in genome engineering, outlining key advances and defining challenges.

Differential requirements of singleplex and multiplex recombineering of large DNA constructs

Recombineering is an in vivo genetic engineering technique involving homologous recombination mediated by phage recombination proteins. The use of recombineering methodology is not limited by size and sequence constraints and therefore has enabled the streamlined construction of bacterial strains and multi-component plasmids. Recombineering applications commonly utilize singleplex strategies and the parameters are extensively tested. However, singleplex recombineering is not suitable for the modification of several loci in genome recoding and strain engineering exercises, which requires a multiplex recombineering design. Defining the main parameters affecting multiplex efficiency especially the insertion of multiple large genes is necessary to enable efficient large-scale modification of the genome. Here, we have tested different recombineering operational parameters of the lambda phage Red recombination system and compared singleplex and multiplex recombineering of large gene sized DNA cassettes. We have found that optimal multiplex recombination required long homology lengths in excess of 120 bp. However, efficient multiplexing was possible with only 60 bp of homology. Multiplex recombination was more limited by lower amounts of DNA than singleplex recombineering and was greatly enhanced by use of phosphorothioate protection of DNA. Exploring the mechanism of multiplexing revealed that efficient recombination required co-selection of an antibiotic marker and the presence of all three Red proteins. Building on these results, we substantially increased multiplex efficiency using an ExoVII deletion strain. Our findings elucidate key differences between singleplex and multiplex recombineering and provide important clues for further improving multiplex recombination efficiency.

A high-efficiency recombineering system with PCR-based ssDNA in Bacillus subtilis mediated by the native phage recombinase GP35

Bacillus subtilis and its closely related species are important strains for industry, agriculture, and medicine. However, it is difficult to perform genetic manipulations using the endogenous recombination machinery. In many bacteria, phage recombineering systems have been employed to improve recombineering frequencies. To date, an efficient phage recombineering system for B. subtilis has not been reported. Here, we, for the first time, identified that GP35 from the native phage SPP1 exhibited a high recombination activity in B. subtilis. On this basis, we developed a high-efficiency GP35-meditated recombineering system. Taking single-stranded DNA (ssDNA) as a recombineering substrate, ten recombinases from diverse sources were investigated in B. subtilis W168. GP35 showed the highest recombineering frequency (1.71 ± 0.15 × 10(-1)). Besides targeting the purine nucleoside phosphorylase gene (deoD), we also demonstrated the utility of GP35 and Beta from Escherichia coli lambda phage by deleting the alpha-amylase gene (amyE) and uracil phosphoribosyltransferase gene (upp). In all three genetic loci, GP35 exhibited a higher frequency than Beta. Moreover, a phylogenetic tree comparing the kinship of different recombinase hosts with B. subtilis was constructed, and the relationship between the recombineering frequency and the kinship of the host was further analyzed. The results suggested that closer kinship to B. subtilis resulted in higher frequency in B. subtilis. In conclusion, the recombinase from native phage or prophage can significantly promote the genetic recombineering frequency in its host, providing an effective genetic tool for constructing genetically engineered strains and investigating bacterial physiology.

Genome engineering using a synthetic gene circuit in Bacillus subtilis

Genome engineering without leaving foreign DNA behind requires an efficient counter-selectable marker system. Here, we developed a genome engineering method in Bacillus subtilis using a synthetic gene circuit as a counter-selectable marker system. The system contained two repressible promoters (B. subtilis xylA (P_xyl) and spac (P_spac)) and two repressor genes (lacI and xylR). P_xyl-lacI was integrated into the B. subtilis genome with a target gene containing a desired mutation. The xylR and P_spac–chloramphenicol resistant genes (cat) were located on a helper plasmid. In the presence of xylose, repression of XylR by xylose induced LacI expression, the LacIs repressed the P_spac promoter and the cells become chloramphenicol sensitive. Thus, to survive in the presence of chloramphenicol, the cell must delete P_xyl-lacI by recombination between the wild-type and mutated target genes. The recombination leads to mutation of the target gene. The remaining helper plasmid was removed easily under the chloramphenicol absent condition. In this study, we showed base insertion, deletion and point mutation of the B. subtilis genome without leaving any foreign DNA behind. Additionally, we successfully deleted a 2-kb gene (amyE) and a 38-kb operon (ppsABCDE). This method will be useful to construct designer Bacillus strains for various industrial applications.

Oligonucleotide recombination in corynebacteria without the expression of exogenous recombinases

Brevibacterium lactofermentum and Corynebacterium glutamicum are important biotechnology species of the genus Corynebacterium. The single-strand DNA annealing protein (SSAP)-independent oligonucleotide-mediated recombination procedure was successfully applied to the commonly used wild-type strains B. lactofermentum AJ1511 and C. glutamicum ATCC13032. When the rpsL gene was used as a target, the optimized protocol yielded up to (1.4±0.3)×10(3) and (6.7±1.3)×10(3) streptomycin-resistant colonies per 10(8) viable cells for the corresponding strains. We tested the influence of several parameters that are known to enhance the efficiency of oligonucleotide-mediated recombination in other bacterial species. Among them, increasing the concentration of oligonucleotides and targeting the lagging strand of the chromosome have proven to have positive effects on both of the tested species. No difference in the efficiency of recombination was observed between the oligonucleotides phosphorothiorated at the 5' ends and the unmodified oligonucleotides or between the oligonucleotides with four mutated nucleotides and those with one mutated nucleotide. The described approach demonstrates that during the adaptation of the recombineering technique, testing SSAP-independent oligonucleotide-mediated recombination could be a good starting point. Such testing could decrease the probability of an incorrect interpretation of the effect of exogenous protein factors (such as SSAP and/or corresponding exonucleases) due to non-optimal experimental conditions. In addition, SSAP-independent recombination itself could be useful in combination with suitable selection/enrichment methods.

Current development in genetic engineering strategies of Bacillus species

The complete sequencing and annotation of the genomes of industrially-important Bacillus species has enhanced our understanding of their properties, and allowed advances in genetic manipulations in other Bacillus species. Post-genomic studies require simple and highly efficient tools to enable genetic manipulation. Here, we summarize the recent progress in genetic engineering strategies for Bacillus species. We review the available genetic tools that have been developed in Bacillus species, as well as methods developed in other species that may also be applicable in Bacillus. Furthermore, we address the limitations and challenges of the existing methods, and discuss the future research prospects in developing novel and useful tools for genetic modification of Bacillus species.

Bacterial cellular engineering by genome editing and gene silencing

Genome editing is an important technology for bacterial cellular engineering, which is commonly conducted by homologous recombination-based procedures, including gene knockout (disruption), knock-in (insertion), and allelic exchange. In addition, some new recombination-independent approaches have emerged that utilize catalytic RNAs, artificial nucleases, nucleic acid analogs, and peptide nucleic acids. Apart from these methods, which directly modify the genomic structure, an alternative approach is to conditionally modify the gene expression profile at the posttranscriptional level without altering the genomes. This is performed by expressing antisense RNAs to knock down (silence) target mRNAs in vivo. This review describes the features and recent advances on methods used in genomic engineering and silencing technologies that are advantageously used for bacterial cellular engineering.

Establishment of a markerless mutation delivery system in Bacillus subtilis stimulated by a double-strand break in the chromosome

Bacillus subtilis has been a model for gram-positive bacteria and it has long been exploited for industrial and biotechnological applications. However, the availability of facile genetic tools for physiological analysis has generally lagged substantially behind traditional genetic models such as Escherichia coli and Saccharomyces cerevisiae. In this work, we have developed an efficient, precise and scarless method for rapid multiple genetic modifications without altering the chromosome of B. subtilis. This method employs upp gene as a counter-selectable marker, double-strand break (DSB) repair caused by exogenous endonuclease I-SceI and comK overexpression for fast preparation of competent cell. Foreign dsDNA can be simply and efficiently integrated into the chromosome by double-crossover homologous recombination. The DSB repair is a potent inducement for stimulating the second intramolecular homologous recombination, which not only enhances the frequency of resolution by one to two orders of magnitude, but also selects for the resolved product. This method has been successfully and reiteratively used in B. subtilis to deliver point mutations, to generate in-frame deletions, and to construct large-scale deletions. Experimental results proved that it allowed repeated use of the selectable marker gene for multiple modifications and could be a useful technique for B. subtilis.

Promoter analysis and transcription regulation of fus gene cluster responsible for fusaricidin synthesis of Paenibacillus polymyxa SQR-21

Fusaricidins produced by Paenibacillus polymyxa are lipopeptide antibiotics with outstanding antifungal activity. In this study, the whole gene cluster responsible for fusaricidin biosynthesis (fusA) was isolated and identified from the cDNA library of one biocontrol agent P. polymyxa SQR-21 (SQR-21). MALDI-TOF MS analysis confirmed that SQR-21 could produce four kinds of fusaricidins: A, B, C, and D. A central promoter that drove the transcription of fusGFEDCBA was revealed by mapping of the fus promoter region by 5' deletions. The disruption of fusA in SQR-21 led to the abolishment of fusaricidin production and antifungal activity. The direct interaction between a potential regulator, AbrB, and the promoter region of fus gene cluster was confirmed by electrophoretic mobility shift assays. One abrB disruption mutant showed significantly higher antifungal activity compared with the wild type. These results revealed a pathway for the transcriptional regulation of the fus gene cluster in P. polymyxa.

A versatile mini-mazF-cassette for marker-free targeted genetic modification in Bacillus subtilis

There are some drawbacks for MazF-cassette constructed in previous reports for marker-free genetic manipulation in Bacillus subtilis, including cloning-dependent methodology and non-strictly controlled expression system. In our study, the modifications on mazF-cassette are carried out, such as using mini Zeocin resistance gene as positive-selectable marker and strictly controlled xyl promoter from the B. subtilis to replace non-strictly controlled IPTG-inducible Pspac or xyl promoter from Bacillus megaterium. Then the mini-mazF-cassette was successfully applied to knock-out the amyE gene, to delete a 90-kb gene cluster, and to knock-in a green fluorescent protein expression cassette employing a cloning-independent methodology, without introducing undesirable redundant sequences at the modified locus in the B. subtilis 1A751. Besides, the mini-mazF-cassette could be used repeatedly to delete multiple genes or gene clusters with only a 2- to 2.5-kb PCR-fused fragment, which largely reduced the frequency of nucleic acid mutations generated by PCR compared to previous reports. We further demonstrated that the frequency of spontaneous mazF-resistant mutants was lower, and the frequency of generating desired clones was nearly 100%. The entire procedure for marker-free genetic manipulation using the mini-mazF-cassette can be finished in about 3days. This modified cassette has remarkable improvement compared to existing approaches and is applicable for available manipulating Bacillus species chromosomes.

Recombineering in Corynebacterium glutamicum combined with optical nanosensors: a general strategy for fast producer strain generation

Recombineering in bacteria is a powerful technique for genome reconstruction, but until now, it was not generally applicable for development of small-molecule producers because of the inconspicuous phenotype of most compounds of biotechnological relevance. Here, we establish recombineering for Corynebacterium glutamicum using RecT of prophage Rac and combine this with our recently developed nanosensor technology, which enables the detection and isolation of productive mutants at the single-cell level via fluorescence-activated cell sorting (FACS). We call this new technology RecFACS, which we use for genomic site-directed saturation mutagenesis without relying on pre-constructed libraries to directly isolate L-lysine-producing cells. A mixture of 19 different oligonucleotides was used targeting codon 81 in murE of the wild-type, at a locus where one single mutation is known to cause L-lysine production. Using RecFACS, productive mutants were screened and isolated. Sequencing revealed 12 different amino acid exchanges in the targeted murE codon, which caused different L-lysine production titers. Apart from introducing a rapid genome construction technology for C. glutamicum, the present work demonstrates that RecFACS is suitable to simply create producers as well as genetic diversity in one single step, thus establishing a new general concept in synthetic biology.

Generalized bacterial genome editing using mobile group II introns and Cre-lox

Efficient bacterial genetic engineering approaches with broad-host applicability are rare. We combine two systems, mobile group II introns ('targetrons') and Cre/lox, which function efficiently in many different organisms, into a versatile platform we call GETR (Genome Editing via Targetrons and Recombinases). The introns deliver lox sites to specific genomic loci, enabling genomic manipulations. Efficiency is enhanced by adding flexibility to the RNA hairpins formed by the lox sites. We use the system for insertions, deletions, inversions, and one-step cut-and-paste operations. We demonstrate insertion of a 12-kb polyketide synthase operon into the lacZ gene of Escherichia coli, multiple simultaneous and sequential deletions of up to 120 kb in E. coli and Staphylococcus aureus, inversions of up to 1.2 Mb in E. coli and Bacillus subtilis, and one-step cut-and-pastes for translocating 120 kb of genomic sequence to a site 1.5 Mb away. We also demonstrate the simultaneous delivery of lox sites into multiple loci in the Shewanella oneidensis genome. No selectable markers need to be placed in the genome, and the efficiency of Cre-mediated manipulations typically approaches 100%.

Enhanced root colonization and biocontrol activity of Bacillus amyloliquefaciens SQR9 by abrB gene disruption

Root colonization by antagonistic bacteria is a prerequisite for successful biological control, and the instability of colonization under varying environmental conditions has accentuated the need to improve the colonization activity. Root colonization by Bacillus spp. is mainly determined by chemotaxis and biofilm formation, and both functions are negatively controlled by the global transcription regulator AbrB. Here, we disrupted the gene abrB in Bacillus amyloliquefaciens SQR9, which has been proven to be a promising biocontrol agent of cucumber and watermelon wilt disease. Chemotaxis, biofilm formation, and colonization activities as well as biocontrol efficiency were measured and compared between the wild-type strain of SQR9 and the abrB mutant. The data presented in this article demonstrate that the colonization and biocontrol activity of B. amyloliquefaciens SQR9 could be significantly improved by abrB gene disruption. The results offer a new strategy to enhance the biocontrol efficacy of B. amyloliquefaciens SQR9.

A genetic replacement system for selection-based engineering of essential proteins

BACKGROUND

Essential genes represent the core of biological functions required for viability. Molecular understanding of essentiality as well as design of synthetic cellular systems includes the engineering of essential proteins. An impediment to this effort is the lack of growth-based selection systems suitable for directed evolution approaches.

RESULTS

We established a simple strategy for genetic replacement of an essential gene by a (library of) variant(s) during a transformation.The system was validated using three different essential genes and plasmid combinations and it reproducibly shows transformation efficiencies on the order of 107 transformants per microgram of DNA without any identifiable false positives. This allowed for reliable recovery of functional variants out of at least a 105-fold excess of non-functional variants. This outperformed selection in conventional bleach-out strains by at least two orders of magnitude, where recombination between functional and non-functional variants interfered with reliable recovery even in recA negative strains.

CONCLUSIONS

We propose that this selection system is extremely suitable for evaluating large libraries of engineered essential proteins resulting in the reliable isolation of functional variants in a clean strain background which can readily be used for in vivo applications as well as expression and purification for use in in vitro studies.