An update of the suicide plasmid-mediated genome editing system in Corynebacterium glutamicum

Fine-Regulating the Carbon Flux of l-Isoleucine Producing Corynebacterium glutamicum WM001 for Efficient l-Threonine Production

l-Threonine, an essential amino acid, is widely used in various industries, with an annually growing demand. However, the present Corynebacterium glutamicum strains are difficult to achieve industrialization of l-threonine due to low yield and purity. In this study, we engineered an l-isoleucine-producing C. glutamicum WM001 to efficiently produce l-threonine by finely regulating the carbon flux. First, the threonine dehydratase in WM001 was mutated to lower the level of l-isoleucine production, then the homoserine dehydrogenase and aspartate kinase were mutated to release the feedback inhibition of l-threonine, and the resulting strain TWZ006 produced 14.2 g/L l-threonine. Subsequently, aspartate ammonia-lyase and aspartate transaminase were overexpressed to accumulate the precursor l-aspartate. Next, phosphoenolpyruvate carboxylase, pyruvate carboxylase and pyruvate kinase were overexpressed, and phosphoenolpyruvate carboxykinase, oxaloacetate decarboxylase were inactivated to fine-regulate the carbon flux among oxaloacetate, pyruvate and phosphoenolpyruvate. The resulting strain TWZ017 produced 21.5 g/L l-threonine. Finally, dihydrodipicolinate synthase was mutated with strong allosteric inhibition from l-lysine to significantly decrease byproducts accumulation, l-threonine export was optimized, and the final engineered strain TWZ024/pXTuf-thrE produced 78.3 g/L of l-threonine with the yield of 0.33 g/g glucose and the productivity of 0.82 g/L/h in a 7 L bioreactor. To the best of our knowledge, this represents the highest l-threonine production in C. glutamicum, providing possibilities for industrial-scale production.

Metabolic Engineering of Corynebacterium glutamicum for the High-Level Production of l-Valine under Aerobic Conditions

l-Valine, an essential amino acid, serves as a valuable compound in various industries. However, engineering strains with both high yield and purity are yet to be delivered for microbial l-valine production. We engineered a Corynebacterium glutamicum strain capable of highly efficient production of l-valine. We initially introduced an acetohydroxy acid synthase mutant from an industrial l-valine producer and optimized a cofactor-balanced pathway, followed by the activation of the nonphosphoenolpyruvate-dependent carbohydrate phosphotransferase system and the introduction of an exogenous Entner-Doudoroff pathway. Subsequently, we weakened anaplerotic pathways, and attenuated the tricarboxylic acid cycle via start codon substitution in icd, encoding isocitrate dehydrogenase. Finally, to balance bacterial growth and l-valine production, an l-valine biosensor-dependent genetic circuit was established to dynamically repress citrate synthase expression. The engineered strain Val19 produced 103 g/L of l-valine with a high yield of 0.35 g/g glucose and a productivity of 2.67 g/L/h. This represents the highest reported l-valine production in C. glutamicum via direct fermentation and exhibits potential for its industrial-scale production, leveraging the advantages of C. glutamicum over other microbes.

Advancements in gene editing technologies for probiotic-enabled disease therapy

Probiotics typically refer to microorganisms that have been identified for their health benefits, and they are added to foods or supplements to promote the health of the host. A growing number of probiotic strains have been identified lately and developed into valuable regulatory pharmaceuticals for nutritional and medical applications. Gene editing technologies play a crucial role in addressing the need for safe and therapeutic probiotics in disease treatment. These technologies offer valuable assistance in comprehending the underlying mechanisms of probiotic bioactivity and in the development of advanced probiotics. This review aims to offer a comprehensive overview of gene editing technologies applied in the engineering of both traditional and next-generation probiotics. It further explores the potential for on-demand production of customized products derived from enhanced probiotics, with a particular emphasis on the future of gene editing in the development of live biotherapeutics.

One-step genome engineering in bee gut bacterial symbionts

Mechanistic understanding of interactions in many host-microbe systems, including the honey bee microbiome, is limited by a lack of easy-to-use genome engineering approaches. To this end, we demonstrate a one-step genome engineering approach for making gene deletions and insertions in the chromosomes of honey bee gut bacterial symbionts. Electroporation of linear or non-replicating plasmid DNA containing an antibiotic resistance cassette flanked by regions with homology to a symbiont genome reliably results in chromosomal integration. This lightweight approach does not require expressing any exogenous recombination machinery. The high concentrations of large DNAs with long homology regions needed to make the process efficient can be readily produced using modern DNA synthesis and assembly methods. We use this approach to knock out genes, including genes involved in biofilm formation, and insert fluorescent protein genes into the chromosome of the betaproteobacterial bee gut symbiont Snodgrassella alvi. We are also able to engineer the genomes of multiple strains of S. alvi and another species, Snodgrassella communis, which is found in the bumble bee gut microbiome. Finally, we use the same method to engineer the chromosome of another bee symbiont, Bartonella apis, which is an alphaproteobacterium. As expected, gene knockout in S. alvi using this approach is recA-dependent, suggesting that this straightforward procedure can be applied to other microbes that lack convenient genome engineering methods.

IMPORTANCE

Honey bees are ecologically and economically important crop pollinators with bacterial gut symbionts that influence their health. Microbiome-based strategies for studying or improving bee health have utilized wild-type or plasmid-engineered bacteria. We demonstrate that a straightforward, single-step method can be used to insert cassettes and replace genes in the chromosomes of multiple bee gut bacteria. This method can be used for investigating the mechanisms of host-microbe interactions in the bee gut community and stably engineering symbionts that benefit pollinator health.

Path to bacteriotherapy: From bacterial engineering to therapeutic perspectives

The major reason for the failure of conventional therapies is the heterogeneity and complexity of tumor microenvironments (TMEs). Many malignant tumors reprogram their surface antigens to evade the immune surveillance, leading to reduced antigen-presenting cells and hindered T-cell activation. Bacteria-mediated cancer immunotherapy has been extensively investigated in recent years. Scientists have ingeniously modified bacteria using synthetic biology and nanotechnology to enhance their biosafety with high tumor specificity, resulting in robust anticancer immune responses. To enhance the antitumor efficacy, therapeutic proteins, cytokines, nanoparticles, and chemotherapeutic drugs have been efficiently delivered using engineered bacteria. This review provides a comprehensive understanding of oncolytic bacterial therapies, covering bacterial design and the intricate interactions within TMEs. Additionally, it offers an in-depth comparison of the current techniques used for bacterial modification, both internally and externally, to maximize their therapeutic effectiveness. Finally, we outlined the challenges and opportunities ahead in the clinical application of oncolytic bacterial therapies.

Development of SacB-based Counterselection for Efficient Allelic Exchange in Fusobacterium nucleatum

Fusobacterium nucleatum , prevalent in the oral cavity, is significantly linked to overall human health. Our molecular comprehension of its role in oral biofilm formation and its interactions with the host under various pathological circumstances has seen considerable advancements in recent years, primarily due to the development of various genetic tools for DNA manipulation in this bacterium. Of these, counterselection-based unmarked in-frame mutation methods have proved notably effective. Under suitable growth conditions, cells carrying a counterselectable gene die, enabling efficient selection of rare defined allelic exchange mutants. The sacB gene from Bacillus subtilis , encoding levansucrase, is a widely used counterselective marker partly due to the easy availability of sucrose. Yet, its potential application in F. nucleatum genetic study remains untested. We demonstrated that F. nucleatum cells expressing sacB in either a shuttle or suicide plasmid exhibit a lethal sensitivity to supplemental sucrose. Utilizing sucrose counterselection, we created an in-frame deletion of the F. nucleatum tonB gene, a critical gene for energy-dependent transport processes in Gram-negative bacteria, and a precise knockin of the luciferase gene immediately following the stop codon of the hslO gene, the last gene of a five-gene operon possible related to the natural competence of F. nucleatum . Post counterselection with 5% sucrose, chromosomal plasmid loss occurred in all colonies, leading to gene alternations in half of the screened isolates. This sacB -based counterselection technique provides a reliable method for isolating unmarked gene mutations in wild-type F. nucleatum , enriching the toolkit for fusobacterial research.

IMPORTANCE

Investigations into Fusobacterium nucleatum 's role in related diseases significantly benefit from the strategies of creating unmarked gene mutations, which hinge on using a counterselective marker. Previously, the galk -based allelic exchange method, while effective, faced an inherent limitation - the need for a modified host. This study aims to surmount this limitation by substituting galK with sacB for gene modification in F. nucleatum . Our application of the sacB -based methodology successfully yielded a tonB in-frame deletion mutant and a luciferase gene knockin at the precise chromosomal location in the wild-type background. The new method augments the existing toolkit for F. nucleatum research and has far-reaching implications due to the easy accessibility to the counterselection compound sucrose. We anticipate its broader adoption in further exploration, thereby reinforcing its critical role in propelling our understanding of F. nucleatum .

Dual genetic level modification engineering accelerate genome evolution of Corynebacterium glutamicum

High spontaneous mutation rate is crucial for obtaining ideal phenotype and exploring the relationship between genes and phenotype. How to break the genetic stability of organisms and increase the mutation frequency has become a research hotspot. Here, we present a practical and controllable evolutionary tool (oMut-Cgts) based on dual genetic level modification engineering for Corynebacterium glutamicum. Firstly, the modification engineering of transcription and replication levels based on RNA polymerase α subunit and DNA helicase Cgl0854 as the 'dock' of cytidine deaminase (pmCDA1) significantly increased the mutation rate, proving that the localization of pmCDA1 around transient ssDNA is necessary for genome mutation. Then, the combined modification and optimization of engineering at dual genetic level achieved 1.02 × 104-fold increased mutation rate. The genome sequencing revealed that the oMut-Cgts perform uniform and efficient C:G→T:A transitions on a genome-wide scale. Furthermore, oMut-Cgts-mediated rapid evolution of C. glutamicum with stress (acid, oxidative and ethanol) tolerance proved that the tool has powerful functions in multi-dimensional biological engineering (rapid phenotype evolution, gene function mining and protein evolution). The strategies for rapid genome evolution provided in this study are expected to be applicable to a variety of applications in all prokaryotic cells.

Efficient Fermentative Production of d-Alanine and Other d-Amino Acids by Metabolically Engineered Corynebacterium glutamicum

d-Amino acids (d-AAs) have wide applications in industries such as pharmaceutical, food, and cosmetics due to their unique properties. Currently, the production of d-AAs has relied on chemical synthesis or enzyme catalysts, and it is challenging to produce d-AAs via direct fermentation from glucose. We observed that Corynebacterium glutamicum exhibits a remarkable tolerance to high concentrations of d-Ala, a crucial characteristic for establishing a successful fermentation process. By optimizing meso-diaminopilmelate dehydrogenases in different C. glutamicum strains and successively deleting l-Ala biosynthetic pathways, we developed an efficient d-Ala fermentation system. The d-Ala titer was enhanced through systems metabolic engineering, which involved strengthening glucose assimilation and pyruvate supply, reducing the formation of organic acid byproducts, and attenuating the TCA cycle. During fermentation in a 5-L bioreactor, a significant accumulation of l-Ala was observed in the broth, which was subsequently diminished by introducing an l-amino acid deaminase. Ultimately, the engineered strain DA-11 produced 85 g/L d-Ala with a yield of 0.30 g/g glucose, accompanied by an optical purity exceeding 99%. The fermentation platform has the potential to be extended for the synthesis of other d-AAs, as demonstrated by the production of d-Val and d-Glu.

Construction of the flagellin F mutant of Vibrio parahaemolyticus and its toxic effects on silver pomfret (Pampus argenteus) cells

Vibrio parahaemolyticus causes diseases in aquatic organisms, leading to substantial financial losses to the aquaculture industry; its flagellin F (flaF) protein triggers severe inflammation in host cells. To enhance the understanding of the function of flaF in V. parahaemolyticus infection, in this study, a flaF-deficient mutant was constructed by employing two-step homologous recombination. The flaF-deficient mutant induced a significantly lower toll-like receptor 5 (TLR5) expression and apoptosis in fish intestinal epithelial cells than the wild-type V. parahaemolyticus. Furthermore, fluorescence labelling and microscopy analysis of TLR5 showed that V. parahaemolyticus and its mutant strain significantly enhanced TLR5 expression. Additionally, the findings suggest that flaF deletion did not significantly affect the expression of myeloid differentiation factor 88 (MyD88) and interleukin-8 (IL-8) induced by V.parahaemolyticus. In summary, V. parahaemolyticus induced a TLR5-dependent inflammatory response and apoptosis through MyD88, which was observed to be influenced by flaF deletion. In this study, we obtained stable mutants of V. parahaemolyticus via target gene deletion-which is a rapid and effective approach-and compared the induction of inflammatory response and apoptosis by V. parahaemolyticus and its mutant strain, providing novel perspectives for functional gene research in V. parahaemolyticus.

Review of knockout technology approaches in bacterial drug resistance research

Gene knockout is a widely used method in biology for investigating gene function. Several technologies are available for gene knockout, including zinc-finger nuclease technology (ZFN), suicide plasmid vector systems, transcription activator-like effector protein nuclease technology (TALEN), Red homologous recombination technology, CRISPR/Cas, and others. Of these, Red homologous recombination technology, CRISPR/Cas9 technology, and suicide plasmid vector systems have been the most extensively used for knocking out bacterial drug resistance genes. These three technologies have been shown to yield significant results in researching bacterial gene functions in numerous studies. This study provides an overview of current gene knockout methods that are effective for genetic drug resistance testing in bacteria. The study aims to serve as a reference for selecting appropriate techniques.

Establishment of CRISPR-Cpf1-assisted gene editing tool and engineering of 4-hydroxyisoleucine biosynthesis in Corynebacterium glutamicum

Corynebacterium glutamicum, an important industrial producer, is a model microorganism. However, the limited gene editing methods and their defects limit the efficient genome editing of C. glutamicum. To improve the screening efficiency of second-cross-over strains of traditional SacB editing system, a universal pCS plasmid which harbors CRISPR-Cpf1 system targeting kan gene of SacB system was designed and established to kill the false positive single-cross-over strains remained abundantly after the second-cross-over events. The lethality of pCS plasmid to C. glutamicum carrying kan gene on its genome was as high as 98.6%. In the example of PodhA::PilvBNC replacement, pCS plasmid improved the screening efficiency of second-cross-over bacteria from 5% to over 95%. Then this pCS-assisted gene editing system was applied to improve the supply of precursors and reduce the generation of by-products in the production of 4-hydroxyisoleucine (4-HIL). The 4-HIL titer of one edited strain SC01-TD5IM reached 137.0 ± 33.9 mM, while the weakening of lysE by promoter engineering reduced Lys content by 19.0-47.7% and 4-HIL titer by 16.4-64.5%. These editing demonstrates again the efficiency of this novel CRISPR-Cpf1-assisted gene editing tool, suggesting it as a useful tool for improving the genome editing and metabolic engineering in C. glutamicum.

Enhanced production of D-pantothenic acid in Corynebacterium glutamicum using an efficient CRISPR-Cpf1 genome editing method

BACKGROUND

Corynebacterium glutamicum has industrial track records for producing a variety of valuable products such as amino acids. Although CRISPR-based genome editing technologies have undergone immense developments in recent years, the suicide-plasmid-based approaches are still predominant for C. glutamicum genome manipulation. It is crucial to develop a simple and efficient CRISPR genome editing method for C. glutamicum.

RESULTS

In this study, we developed a RecombinAtion Prior to Induced Double-strand-break (RAPID) genome editing technology for C. glutamicum, as Cpf1 cleavage was found to disrupt RecET-mediated homologous recombination (HR) of the donor template into the genome. The RAPID toolbox enabled highly efficient gene deletion and insertion, and notably, a linear DNA template was sufficient for gene deletion. Due to the simplified procedure and iterative operation ability, this methodology could be widely applied in C. glutamicum genetic manipulations. As a proof of concept, a high-yield D-pantothenic acid (vitamin B5)-producing strain was constructed, which, to the best of our knowledge, achieved the highest reported titer of 18.62 g/L from glucose only.

CONCLUSIONS

We developed a RecET-assisted CRISPR-Cpf1 genome editing technology for C. glutamicum that harnessed CRISPR-induced DSBs as a counterselection. This method is of great importance to C. glutamicum genome editing in terms of its practical applications, which also guides the development of CRISPR genome editing tools for other microorganisms.

Synthetic biology tools for engineering Corynebacterium glutamicum

Corynebacterium glutamicum is a promising organism for the industrial production of amino acids, fuels, and various value-added chemicals. From the whole genome sequence release, C. glutamicum has been valuable in the field of industrial microbiology and biotechnology. Continuous discovery of genetic manipulations and regulation mechanisms has developed C. glutamicum as a synthetic biology platform chassis. This review summarized diverse genomic manipulation technologies and gene expression tools for static, dynamic, and multiplex control at transcription and translation levels. Moreover, we discussed the current challenges and applicable tools to C. glutamicum for future advancements.

Combining metabolite doping and metabolic engineering to improve 2-phenylethanol production by engineered cyanobacteria

2-Phenylethanol (2-PE) is a rose-scented aromatic compound, with broad application in cosmetic, pharmaceutical, food and beverage industries. Many plants naturally synthesize 2-PE via Shikimate Pathway, but its extraction is expensive and low-yielding. Consequently, most 2-PE derives from chemical synthesis, which employs petroleum as feedstock and generates unwanted by products and health issues. The need for "green" processes and the increasing public demand for natural products are pushing biotechnological production systems as promising alternatives. So far, several microorganisms have been investigated and engineered for 2-PE biosynthesis, but a few studies have focused on autotrophic microorganisms. Among them, the prokaryotic cyanobacteria can represent ideal microbial factories thanks to their ability to photosynthetically convert CO2 into valuable compounds, their minimal nutritional requirements, high photosynthetic rate and the availability of genetic and bioinformatics tools. An engineered strain of Synechococcus elongatus PCC 7942 for 2-PE production, i.e., p120, was previously published elsewhere. The strain p120 expresses four heterologous genes for the complete 2-PE synthesis pathway. Here, we developed a combined approach of metabolite doping and metabolic engineering to improve the 2-PE production kinetics of the Synechococcus elongatus PCC 7942 p120 strain. Firstly, the growth and 2-PE productivity performances of the p120 recombinant strain were analyzed to highlight potential metabolic constraints. By implementing a BG11 medium doped with L-phenylalanine, we covered the metabolic burden to which the p120 strain is strongly subjected, when the 2-PE pathway expression is induced. Additionally, we further boosted the carbon flow into the Shikimate Pathway by overexpressing the native Shikimate Kinase in the Synechococcus elongatus PCC 7942 p120 strain (i.e., 2PE_aroK). The combination of these different approaches led to a 2-PE yield of 300 mg/gDW and a maximum 2-PE titer of 285 mg/L, 2.4-fold higher than that reported in literature for the p120 recombinant strain and, to our knowledge, the highest recorded for photosynthetic microorganisms, in photoautotrophic growth condition. Finally, this work provides the basis for further optimization of the process aimed at increasing 2-PE productivity and concentration, and could offer new insights about the use of cyanobacteria as appealing microbial cell factories for the synthesis of aromatic compounds.

Analysis of the Plasmid-Based ts Allele of PA0006 Reveals Its Function in Regulation of Cell Morphology and Biosynthesis of Core Lipopolysaccharide in Pseudomonas aeruginosa

Over 300 essential genes are predicted using transposon sequencing in the genome of Pseudomonas aeruginosa. However, methods for reverse genetic analysis of essential genes are scarce. To address this issue, we developed a three-step protocol consisting of integration of deletion plasmid, introduction of temperature-sensitive rescue plasmid, and excision of integrated-deletion plasmid to construct the plasmid-based temperature-sensitive allele of essential genes. Using PA0006 as an example, we showed that PA0006(Ts) exhibited wild-type cell morphology at permissive temperature but filamentous form at restrictive temperatures. We further showed that the glycerol-mannoheptose-bisphosphate phosphatase GmhB in Escherichia coli shared 32.4% identity with that of PA0006p and functionally complemented the defect of PA0006(Ts) at 42°C. SDS-PAGE and Western blotting indicated the presence and absence of the complete core lipopolysaccharide (LPS) and B-band O-antigen in PA0006(Ts) at 30 and 42°C, respectively. An isolated suppressor sup displayed wild-type-like cell morphology but no complete core LPS or O-antigen. Genome resequencing together with comparative transcriptomic profiling identified a candidate suppressor fructose-bisphosphate phosphatase in which the promoter harbored a SNP and the transcription level was not downregulated at 42°C compared to 30°C in sup. It was further validated that fbp overexpression suppressed the lethality of PA0006(Ts) at 42°C. Taken together, our results demonstrate that PA0006 plays a role in regulation of cell morphology and biosynthesis of core LPS. This three-step protocol for construction of conditional lethal allele in P. aeruginosa should be widely applicable for genetic analysis of other essential genes of interest, including analysis of bypass suppressibility. IMPORTANCE Microbial essential genes encode nondispensable function for cell growth and therefore are ideal targets for the development of new drugs. Essential genes are readily identified using transposon-sequencing technology at the genome scale. However, genetic analysis of essential genes of interest was hampered by limited methodologies. To address this issue, we developed a three-step protocol for construction of conditional allele of essential genes in the opportunistic pathogen Pseudomonas aeruginosa. Using PA0006 as an example, we demonstrated that the plasmid-based PA0006(Ts) mutant exhibited defects in regulation of cell morphology, formation of intact core LPS, and attachment of the O-antigen at restrictive temperatures but not at permissive temperatures. A suppressor of PA0006(Ts) was isolated through spontaneous mutations and showed restored cell morphology but not core oligosaccharide or O-antigen. This method should be widely applicable for phenotype and suppressibility analyses of other essential genes of interest in P. aeruginosa.

Engineering Citrobacter freundii using CRISPR/Cas9 system

The CRISPR/Cas9 (clustered regularly interspaced short palindromic repeats/CRISPR associated proteins) system is a useful tool to edit genomes quickly and efficiently. However, the use of CRISPR/Cas9 to edit bacterial genomes has been limited to select microbial chassis primarily used for bioproduction of high value products. Thus, expansion of CRISPR/Cas9 tools to other microbial organisms is needed. Here, our aim was to assess the suitability of CRISPR/Cas9 for genome editing of the Citrobacter freundii type strain ATCC 8090. We evaluated the commonly used two plasmid pCas/pTargetF system to enable gene deletions and insertions in C. freundii and determined editing efficiency. The CRISPR/Cas9 based method enabled high editing efficiency (~91%) for deletion of galactokinase (galk) and enabled deletion with various single guide RNA (sgRNA) sequences. To assess the ability of CRISPR/Cas9 tools to insert genes, we used the fluorescent reporter mNeonGreen, an endopeptidase (yebA), and a transcriptional regulator (xylS) and found successful insertion with high efficiency (81-100%) of each gene individually. These results strengthen and expand the use of CRISPR/Cas9 genome editing to C. freundii as an additional microbial chassis.

Bacterially mediated drug delivery and therapeutics: Strategies and advancements

It was already clinically apparent 150 years ago that bacterial therapy could alleviate diseases. Recently, a burgeoning number of researchers have been using bacterial regimens filled with microbial therapeutic leads to diagnose and treat a wide range of disorders and diseases, including cancers, inflammatory diseases, metabolic disorders and viral infections. Some bacteria that were designed to have low toxicity and high efficiency in drug delivery have been used to treat diseases successfully, especially in tumor therapy in animal models or clinical trials, thanks to the progress of genetic engineering and synthetic bioengineering. Therefore, genetically engineered bacteria can serve as efficient drug delivery vehicles, carrying nucleic acids or genetic circuits that encode and regulate therapeutic payloads. In this review, we summarize the development and applications of this approach. Strategies for genetically modifying strains are described in detail, along with their objectives. We also describe some controlled strategies for drug delivery and release using these modified strains as carriers. Furthermore, we discuss treatment methods for various types of diseases using engineered bacteria. Tumors are discussed as the most representative example, and other diseases are also briefly described. Finally, we discuss the challenges and prospects of drug delivery systems based on these bacteria.

AutoESD: a web tool for automatic editing sequence design for genetic manipulation of microorganisms

Advances in genetic manipulation and genome engineering techniques have enabled on-demand targeted deletion, insertion, and substitution of DNA sequences. One important step in these techniques is the design of editing sequences (e.g. primers, homologous arms) to precisely target and manipulate DNA sequences of interest. Experimental biologists can employ multiple tools in a stepwise manner to assist editing sequence design (ESD), but this requires various software involving non-standardized data exchange and input/output formats. Moreover, necessary quality control steps might be overlooked by non-expert users. This approach is low-throughput and can be error-prone, which illustrates the need for an automated ESD system. In this paper, we introduce AutoESD (https://autoesd.biodesign.ac.cn/), which designs editing sequences for all steps of genetic manipulation of many common homologous-recombination techniques based on screening-markers. Notably, multiple types of manipulations for different targets (CDS or intergenic region) can be processed in one submission. Moreover, AutoESD has an entirely cloud-based serverless architecture, offering high reliability, robustness and scalability which is capable of parallelly processing hundreds of design tasks each having thousands of targets in minutes. To our knowledge, AutoESD is the first cloud platform enabling precise, automated, and high-throughput ESD across species, at any genomic locus for all manipulation types.

Genomic Iterative Replacements of Large Synthetic DNA Fragments in Corynebacterium glutamicum

Synthetic genomics will advance our understanding of life and allow us to rebuild the genomes of industrial microorganisms for enhancing performances. Corynebacterium glutamicum, a Gram-positive bacterium, is an important industrial workhorse. However, its genome synthesis is impeded by the low efficiencies in DNA delivery and in genomic recombination/replacement. In the present study, we describe a genomic iterative replacement system based on RecET recombination for C. glutamicum, involving the successive integration of up to 10 kb DNA fragments obtained in vitro, and the transformants are selected by the alternative use of kanR and speR selectable markers. As a proof of concept, we systematically redesigned and replaced a 54.3 kb wild-type sequence of C. glutamicumATCC13032 with its 55.1 kb synthetic counterpart with several novel features, including decoupled genes, the standard PCRTags, and 20 loxPsym sites, which was for the first time incorporated into a bacterial genome. The resulting strain semi-synCG-A1 had a phenotype and fitness similar to the wild-type strain under various stress conditions. The stability of the synthetic genome region faithfully maintained over 100 generations of nonselective growth. Genomic deletions, inversions, and translocations occurred in the synthetic genome region upon induction of synthetic chromosome rearrangement and modification by loxP-mediated evolution (SCRaMbLE), revealing potential genetic flexibility for C. glutamicum. This strategy can be used for the synthesis of a larger region of the genome and facilitate the endeavors for metabolic engineering and synthetic biology of C. glutamicum.

Physiological Responses of Ribosomal Protein S12 K43 Mutants of Corynebacterium glutamicum

Bacterial resistance to streptomycin is often acquired as a consequence of mutations in rpsL, the gene encoding ribosomal protein S12. Corynebacterium glutamicum is a non-pathogenic Gram-positive soil bacterium that has been widely used in industry. In a previous study, we screened several streptomycin-resistant rpsL K43 mutants of C. glutamicum, and surprisingly found that two of them also confer chloramphenicol and/or kanamycin resistance. In order to understand whether or not a single mutation of rpsL^K43 could confer resistance to multiple antibiotics, in this study we attempted to construct saturation mutagenesis of rpsL K43 by rational genetic manipulation. Despite many efforts had been made, only nine mutants were successfully constructed. They were indeed resistant to streptomycin, but not to other antibiotics. This suggested that other mutations should be acquired, contributing to multiple antibiotics in the screened strains. The growth and enhanced green fluorescent protein (eGFP) expression of these nine mutants were then investigated. The results showed that they grew differently in CGXII minimal medium, but not in BHI medium. When cultured in the absence of streptomycin, the expression of eGFP was positively proportional to the growth, approximately, while in the presence of streptomycin, the expression of eGFP was proportional to the ability of streptomycin resistance.

Creation of Markerless Genome Modifications in a Nonmodel Bacterium by Fluorescence-Aided Recombineering

Metabolic engineering of nonmodel bacteria is often challenging because of the paucity of genetic tools for iterative genome modification necessary to equip bacteria with pathways to produce high-value products. Here, we outline a homologous recombination-based method developed to delete or add genes to the genome of a nonmodel bacterium, Zymomonas mobilis, at the desired locus using a suicide plasmid that contains gfp as a fluorescence marker to track its presence in cells. The suicide plasmid is engineered to contain two 500 bp regions homologous to the DNA sequence immediately flanking the target locus. A single crossover event at one of the two homologous regions facilitates insertion of the plasmid into the genome and subsequent homologous recombination events excise the plasmid from the genome, leaving either the original genotype or the desired modified genotype. A key feature of this plasmid is that Green Fluorescent Protein (GFP) expressed from the suicide plasmid allows easy identification and sorting of cells that have lost the plasmid by use of a fluorescence activated cell sorter. Subsequent PCR amplification of genomic DNA from strains lacking GFP allows rapid identification of the desired genotype, which is confirmed by DNA sequencing. This method provides an efficient and flexible platform for improved genetic engineering of Z. mobilis, which can be easily adapted to other nonmodel bacteria.

GEDpm-cg: Genome Editing Automated Design Platform for Point Mutation Construction in Corynebacterium glutamicum

Advances in robotic system-assisted genome editing techniques and computer-aided design tools have significantly facilitated the development of microbial cell factories. Although multiple separate software solutions are available for vector DNA assembly, genome editing, and verification, by far there is still a lack of complete tool which can provide a one-stop service for the entire genome modification process. This makes the design of numerous genetic modifications, especially the construction of mutations that require strictly precise genetic manipulation, a laborious, time-consuming and error-prone process. Here, we developed a free online tool called GEDpm-cg for the design of genomic point mutations in C. glutamicum. The suicide plasmid-mediated counter-selection point mutation editing method and the overlap-based DNA assembly method were selected to ensure the editability of any single nucleotide at any locus in the C. glutamicum chromosome. Primers required for both DNA assembly of the vector for genetic modification and sequencing verification were provided as design results to meet all the experimental needs. An in-silico design task of over 10,000 single point mutations can be completed in 5 min. Finally, three independent point mutations were successfully constructed in C. glutamicum guided by GEDpm-cg, which confirms that the in-silico design results could accurately and seamlessly be bridged with in vivo or in vitro experiments. We believe this platform will provide a user-friendly, powerful and flexible tool for large-scale mutation analysis in the industrial workhorse C. glutamicum via robotic/software-assisted systems.

A SacB-based system for diverse and multiple genome editing in Gluconobacter oxydans

Gluconobacter oxydans is an important industrial bacterial strain widely used to produce a lot of useful products. However, very few gene editing tools are available for G. oxydans. This study aimed to develop an efficient genome editing method for G. oxydans using SacB as a counter-selectable marker. A plasmid that could express the kanamycin resistance gene in both E. coli and G. oxydans was constructed using the screened shuttle promoter P₁₁₆. After optimizing the genome editing conditions, the derivative plasmids could be effectively utilized for diverse genome editing, such as gene deletion, insertion, replacement, and in situ modification in G. oxydans WSH-003. In addition, the SacB-based system also achieved multiple gene editing in G. oxydans. Moreover, the genome of the industrial strain G. oxydans WSH-003 was modified and the growth rate and substrate conversion rate of the strain successfully increased using this system. The system could also have potential to be applied in different G. oxydans strains. The process established in this study also provides a reference for constructing genetic tools for many other genetically recalcitrant bacteria.

Genome Editing in Bacteria: CRISPR-Cas and Beyond

Genome editing in bacteria encompasses a wide array of laborious and multi-step methods such as suicide plasmids. The discovery and applications of clustered regularly interspaced short palindromic repeats (CRISPR)-Cas based technologies have revolutionized genome editing in eukaryotic organisms due to its simplicity and programmability. Nevertheless, this system has not been as widely favored for bacterial genome editing. In this review, we summarize the main approaches and difficulties associated with CRISPR-Cas-mediated genome editing in bacteria and present some alternatives to circumvent these issues, including CRISPR nickases, Cas12a, base editors, CRISPR-associated transposases, prime-editing, endogenous CRISPR systems, and the use of pre-made ribonucleoprotein complexes of Cas proteins and guide RNAs. Finally, we also address fluorescent-protein-based methods to evaluate the efficacy of CRISPR-based systems for genome editing in bacteria. CRISPR-Cas still holds promise as a generalized genome-editing tool in bacteria and is developing further optimization for an expanded application in these organisms. This review provides a rarely offered comprehensive view of genome editing. It also aims to familiarize the microbiology community with an ever-growing genome-editing toolbox for bacteria.

Advances and Perspectives for Genome Editing Tools of Corynebacterium glutamicum

Corynebacterium glutamicum has been considered a promising synthetic biological platform for biomanufacturing and bioremediation. However, there are still some challenges in genetic manipulation of C. glutamicum. Recently, more and more genetic parts or elements (replicons, promoters, reporter genes, and selectable markers) have been mined, characterized, and applied. In addition, continuous improvement of classic molecular genetic manipulation techniques, such as allelic exchange via single/double-crossover, nuclease-mediated site-specific recombination, RecT-mediated single-chain recombination, actinophages integrase-mediated integration, and transposition mutation, has accelerated the molecular study of C. glutamicum. More importantly, emerging gene editing tools based on the CRISPR/Cas system is revolutionarily rewriting the pattern of genetic manipulation technology development for C. glutamicum, which made gene reprogramming, such as insertion, deletion, replacement, and point mutation, much more efficient and simpler. This review summarized the recent progress in molecular genetic manipulation technology development of C. glutamicum and discussed the bottlenecks and perspectives for future research of C. glutamicum as a distinctive microbial chassis.

Recent Advances in Genetic Tools for Acinetobacter baumannii

Acinetobacter baumannii is classified as a top priority pathogen by the World Health Organization (WHO) because of its widespread resistance to all classes of antibiotics. This makes the need for understanding the mechanisms of resistance and virulence critical. Therefore, tools that allow genetic manipulations are vital to unravel the mechanisms of multidrug resistance (MDR) and virulence in A. baumannii. A host of current strategies are available for genetic manipulations of A. baumannii laboratory-strains, including ATCC® 17978TM and ATCC® 19606T, but depending on susceptibility profiles, these strategies may not be sufficient when targeting strains newly obtained from clinic, primarily due to the latter's high resistance to antibiotics that are commonly used for selection during genetic manipulations. This review highlights the most recent methods for genetic manipulation of A. baumannii including CRISPR based approaches, transposon mutagenesis, homologous recombination strategies, reporter systems and complementation techniques with the spotlight on those that can be applied to MDR clinical isolates.

De Novo Engineering of Corynebacterium glutamicum for l-Proline Production

l-Proline is an important amino acid that has various industrial applications. Industrial l-proline-producing strains are obtained by the mutagenesis of Corynebacterium glutamicum. In this study, the optimized C. glutamicum genome-editing tools were further applied in the de novo construction of a hyper-l-proline-producing strain. Overexpression of a feedback inhibition-resistant γ-glutamic kinase mutant ProBG149K, deletion of a proline dehydrogenase to block l-proline degradation, overexpression of glutamate dehydrogenase to increase glutamate synthesis flux, the mutation of 6-phosphate gluconate dehydrogenase and glucose-6-phosphate-dehydrogenase in the pentose phosphate pathway to enhance NADPH supply, the deletion of pyruvate aminotransferase to decrease the byproduct l-alanine synthesis, and weakening of α-ketoglutarate dehydrogenase to regulate the TCA cycle were combined to obtain ZQJY-9. ZQJY-9 produced 19.68 ± 0.22 g/L of l-proline in flask fermentation and was also demonstrated at the 3 L bioreactor level by fed-batch fermentation producing 120.18 g/L of l-proline at 76 h with the highest productivity of 1.581 g/L/h.

Efficient dual-negative selection for bacterial genome editing

Background

Gene editing is key for elucidating gene function. Traditional methods, such as consecutive single-crossovers, have been widely used to modify bacterial genomes. However, cumbersome cloning and limited efficiency of negative selection often make this method slower than other methods such as recombineering.

Results

Here, we established a time-effective variant of consecutive single-crossovers. This method exploits rapid plasmid construction using Gibson assembly, a convenient E. coli donor strain, and efficient dual-negative selection for improved suicide vector resolution. We used this method to generate in-frame deletions, insertions and point mutations in Salmonella enterica with limited hands-on time. Adapted versions enabled efficient gene editing also in Pseudomonas aeruginosa and multi-drug resistant (MDR) Escherichia coli clinical isolates.

Conclusions

Our method is time-effective and allows facile manipulation of multiple bacterial species including MDR clinical isolates. We anticipate that this method might be broadly applicable to additional bacterial species, including those for which recombineering has been difficult to implement.

Homing endonuclease I-SceI-mediated Corynebacterium glutamicum ATCC 13032 genome engineering

Corynebacterium glutamicum is widely used to produce amino acids and is a chassis for the production of value-added compounds. Effective genome engineering methods are crucial to metabolic engineering and synthetic biology studies of C. glutamicum. Herein, a homing endonuclease I-SceI-mediated genome engineering strategy was established for the model strain C. glutamicum ATCC 13032. A vegetative R6K replicon-based, suicide plasmid was employed. The plasmid, pLS3661, contains both tightly regulated, IPTG (isopropyl-β-D-1-thiogalactopyranoside)-inducible I-SceI expression elements and two I-SceI recognition sites. Following cloning of the homologous arms into pLS3661 and transfer the recombinant vector into C. glutamicum ATCC 13032, through the homologous recombination between the cloned fragment and its chromosomal allele, a merodiploid was selected under kanamycin selection. Subsequently, a merodiploid was resolved by double-stranded break repair stimulated by IPTG-stimulated I-SceI expression, generating desired mutants. The protocol obviates a pre-generated strain, transfer of a second I-SceI expression plasmid, and there is not any strain, medium, and temperature restrictions. We validated the approach via deletions of five genes (up to ~ 13.0 kb) and knock-in of one DNA fragment. Furthermore, through kanamycin resistance repair, the ssDNA recombineering parameters were optimized. We hope the highly efficient method will be helpful for the studies of C. glutamicum, and potentially, to other bacteria. KEY POINTS: • Counterselection marker I-SceI-mediated C. glutamicum genome engineering • A suicide vector contains I-SceI expression elements and its recognition sites • Gene deletions and knock-in were conducted; efficiency was as high as 90% • Through antibiotic resistance repair, ssDNA recombineering parameters were optimized.

Efficient fermentative production of L-theanine by Corynebacterium glutamicum

L-Theanine is a unique non-protein amino acid found in tea plants that has been shown to possess numerous functional properties relevant to food science and human nutrition. L-Theanine has been commercially developed as a valuable additive for use in food and beverages, and its market is expected to expand substantially if the production cost can be lowered. Although the enzymatic approach holds considerable potential for use in L-theanine production, demand exists for developing more tractable methods (than those currently available) that can be implemented under mild conditions and will reduce operational procedures and cost. Here, we sought to engineer fermentative production of L-theanine in Corynebacterium glutamicum, an industrially safe host. For L-theanine synthesis, we used γ-glutamylmethylamide synthetase (GMAS), which catalyzes the ATP-dependent ligation of L-glutamate and ethylamine. First, distinct GMASs were expressed in C. glutamicum wild-type ATCC 13032 strain and GDK-9, an L-glutamate overproducing strain, to produce L-theanine upon ethylamine addition to the hosts. Second, the L-glutamate exporter in host cells was disrupted, which markedly increased the L-theanine titer in GDK-9 cells and almost eliminated the accumulation of L-glutamate in the culture medium. Third, a chromosomally gmas_Mm-integrated L-alanine producer was constructed and used, attempting to synthesize ethylamine endogenously by expressing plant-derived L-serine/L-alanine decarboxylases; however, these enzymes showed no L-alanine decarboxylase activity under our experimental conditions. The optimal engineered strain that we ultimately created produced ~ 42 g/L L-theanine, with a yield of 19.6%, in a 5-L fermentor. This is the first report of fermentative production of L-theanine achieved using ethylamine supplementation.