RNA-guided single/double gene repressions in Corynebacterium glutamicum using an efficient CRISPR interference and its application to industrial strain

CRISPR-dCas13a system for programmable small RNAs and polycistronic mRNA repression in bacteria

Bacterial small RNAs (sRNAs) function in post-transcriptional regulatory responses to environmental changes. However, the lack of eukaryotic RNA interference-like machinery in bacteria has limited the systematic engineering of RNA repression. Here, we report the development of clustered regularly interspaced short palindromic repeats (CRISPR)-guided dead CRIPSR-associated protein 13a (dCas13a) ribonucleoprotein that utilizes programmable CRISPR RNAs (crRNAs) to repress trans-acting and cis-acting sRNA as the target, altering regulatory mechanisms and stress-related phenotypes. In addition, we implemented a modular loop engineering of the crRNA to promote modular repression of the target gene with 92% knockdown efficiency and a single base-pair mismatch specificity. With the engineered crRNAs, we achieved targetable single-gene repression in the polycistronic operon. For metabolic application, 102 crRNAs were constructed in the biofoundry and used for screening novel knockdown sRNA targets to improve lycopene (colored antioxidant) production in Escherichia coli. The CRISPR-dCas13a system will assist as a valuable systematic tool for the discovery of novel sRNAs and the fine-tuning of bacterial RNA repression in both scientific and industrial applications.

Recent advances in the metabolic engineering and physiological opportunities for microbial synthesis of L-aspartic acid family amino acids: A review

L-aspartic acid, L-threonine, L-isoleucine, l-lysine, and L-methionine constitute the l-aspartate amino acids (AFAAs). Except for L-aspartic acid, these are essential amino acids that cannot be synthesized by humans or animals themselves. E. coli and C. glutamicum are the main model organisms for AFAA production. It is necessary to reconstitute microbial cell factories and the physiological state of industrial fermentation cells for in-depth research into strains with higher AFAA production levels and optimal growth states. Considering that the anabolic pathways of the AFAAs and engineering modifications have rarely been reviewed in the latest progress, this work reviews the central metabolic pathways of two strains and strategies for the metabolic engineering of AFAA synthetic pathways. The challenges posed by microbial physiology in AFAA production and possible strategies to address them, as well as future research directions for constructing strains with high AFAA production levels, are discussed in this review article.

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

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

CRISPR-Based Gene Editing in Acinetobacter baumannii to Combat Antimicrobial Resistance

Antimicrobial resistance (AMR) poses a significant threat to the health, social, environment, and economic sectors on a global scale and requires serious attention to addressing this issue. Acinetobacter baumannii was given top priority among infectious bacteria because of its extensive resistance to nearly all antibiotic classes and treatment options. Carbapenem-resistant A. baumannii is classified as one of the critical-priority pathogens on the World Health Organization (WHO) priority list of antibiotic-resistant bacteria for effective drug development. Although available genetic manipulation approaches are successful in A. baumannii laboratory strains, they are limited when employed on newly acquired clinical strains since such strains have higher levels of AMR than those used to select them for genetic manipulation. Recently, the CRISPR-Cas (Clustered regularly interspaced short palindromic repeats/CRISPR-associated protein) system has emerged as one of the most effective, efficient, and precise methods of genome editing and offers target-specific gene editing of AMR genes in a specific bacterial strain. CRISPR-based genome editing has been successfully applied in various bacterial strains to combat AMR; however, this strategy has not yet been extensively explored in A. baumannii. This review provides detailed insight into the progress, current scenario, and future potential of CRISPR-Cas usage for AMR-related gene manipulation in A. baumannii.

Recent advances in microbial production of diamines, aminocarboxylic acids, and diacids as potential platform chemicals and bio-based polyamides monomers

Recently, bio-based manufacturing processes of value-added platform chemicals and polymers in biorefineries using renewable resources have extensively been developed for sustainable and carbon dioxide (CO₂) neutral-based industry. Among them, bio-based diamines, aminocarboxylic acids, and diacids have been used as monomers for the synthesis of polyamides having different carbon numbers and ubiquitous and versatile industrial polymers and also as precursors for further chemical and biological processes to afford valuable chemicals. Until now, these platform bio-chemicals have successfully been produced by biorefinery processes employing enzymes and/or microbial host strains as main catalysts. In this review, we discuss recent advances in bio-based production of diamines, aminocarboxylic acids, and diacids, which has been developed and improved by systems metabolic engineering strategies of microbial consortia and optimization of microbial conversion processes including whole cell bioconversion and direct fermentative production.

Recent Advances in CRISPR-Cas Technologies for Synthetic Biology

With developments in synthetic biology, "engineering biology" has emerged through standardization and platformization based on hierarchical, orthogonal, and modularized biological systems. Genome engineering is necessary to manufacture and design synthetic cells with desired functions by using bioparts obtained from sequence databases. Among various tools, the CRISPR-Cas system is modularly composed of guide RNA and Cas nuclease; therefore, it is convenient for editing the genome freely. Recently, various strategies have been developed to accurately edit the genome at a single nucleotide level. Furthermore, CRISPR-Cas technology has been extended to molecular diagnostics for nucleic acids and detection of pathogens, including disease-causing viruses. Moreover, CRISPR technology, which can precisely control the expression of specific genes in cells, is evolving to find the target of metabolic biotechnology. In this review, we summarize the status of various CRISPR technologies that can be applied to synthetic biology and discuss the development of synthetic biology combined with CRISPR technology in microbiology.

High-Fidelity Cytosine Base Editing in a GC-Rich Corynebacterium glutamicum with Reduced DNA Off-Target Editing Effects

Genome editing technology is a powerful tool for programming microbial cell factories. However, rat APOBEC1-derived cytosine base editor (CBE) that converts C•G to T•A at target genes induced DNA off-targets, regardless of single-guide RNA (sgRNA) sequences. Although the high efficiencies of the bacterial CBEs have been developed, a risk of unidentified off-targets impeded genome editing for microbial cell factories. To address the issues, we demonstrate the genome engineering of Corynebacterium glutamicum as a GC-rich model industrial bacterium by generating premature termination codons (PTCs) in desired genes using high-fidelity cytosine base editors (CBEs). Through this CBE-STOP approach of introducing specific cytosine conversions, we constructed several single-gene-inactivated strains for three genes (ldh, idsA, and pyc) with high base editing efficiencies of average 95.6% (n = 45, C6 position) and the highest success rate of up to 100% for PTCs and ultimately developed a strain with five genes (ldh, actA, ackA, pqo, and pta) that were inactivated sequentially for enhancing succinate production. Although these mutant strains showed the desired phenotypes, whole-genome sequencing (WGS) data revealed that genome-wide point mutations occurred in each strain and further accumulated according to the duration of CBE plasmids. To lower the undesirable mutations, high-fidelity CBEs (pCoryne-YE1-BE3 and pCoryne-BE3-R132E) was employed for single or multiplexed genome editing in C. glutamicum, resulting in drastically reduced sgRNA-independent off-targets. Thus, we provide a CRISPR-assisted bacterial genome engineering tool with an average high efficiency of 90.5% (n = 76, C5 or C6 position) at the desired targets. IMPORTANCE Rat APOBEC1-derived cytosine base editor (CBE) that converts C•G to T•A at target genes induced DNA off-targets, regardless of single-guide RNA (sgRNA) sequences. Although the high efficiencies of bacterial CBEs have been developed, a risk of unidentified off-targets impeded genome editing for microbial cell factories. To address the issues, we identified the DNA off-targets for single and multiple genome engineering of the industrial bacterium Corynebacterium glutamicum using whole-genome sequencing. Further, we developed the high-fidelity (HF)-CBE with significantly reduced off-targets with comparable efficiency and precision. We believe that our DNA off-target analysis and the HF-CBE can promote CRISPR-assisted genome engineering over conventional gene manipulation tools by providing a markerless genetic tool without need for a foreign DNA donor.

Co-production of l-Lysine and Heterologous Squalene in CRISPR/dCas9-Assisted Corynebacterium glutamicum

Corynebacterium glutamicum is widely used for a large-scale industrial producer of feed additive amino acids, such as l-lysine. Moreover, C. glutamicum has been engineered for producing various non-native chemicals, including terpenes. For the first time, C. glutamicum was engineered for co-production of l-lysine and heterologous squalene. To control metabolic fluxes for either the l-lysine biosynthesis pathway or the squalene biosynthesis pathway, pyruvate, an intermediate in the central metabolism, a node was regulated by a clustered regularly interspaced short palindromic repeat (CRISPR) interference system. Repressing pyc encoding for pyruvate carboxylase in the l-lysine producer (DM1919) and its derivatives resulted in 99.24 ± 7.63 mg/L total squalene and 6.25 ± 0.20 g/L extracellular lysine at 120 h. Furthermore, various oil overlays were tested for efficient co-productions. In situ extraction with corn oil (10%, v/v) exhibited a separation of 99.75% (w/v) of total squalene (intra- and extracellular squalene), while l-lysine can be secreted in the medium. This co-production strategy will help a potential bioprocess of amino acid production with various terpenes.

Identification of key genes through the constructed CRISPR-dcas9 to facilitate the efficient production of O-acetylhomoserine in Corynebacterium glutamicum

O-Acetylhomoserine (OAH) is an important platform chemical for the synthesis of L-methamidophos and l-methionine. It has been produced efficiently in Corynebacterium glutamicum. However, a wider range of key factors had not been identified, limiting further increases in OAH production. This study successfully identified some limiting factors and regulated them to improve OAH titer. Firstly, an efficient clustered regularly interspaced short palindromic repeats/dead CRISPR associated protein 9 (CRISPR-dCas9) system was constructed and used to identify the key genes in central metabolism and branch pathways associated with OAH biosynthesis. Then, the gltA gene involved in TCA cycle was identified as the most critical gene. A sequential promoter P_NCgl2698, which showed different transcriptional intensity in different strain growth periods, was used to control the expression of gltA gene, resulting in OAH production of 7.0 g/L at 48 h. Finally, the OAH titer of the engineered strain reached 25.9 g/L at 72 h in a 5-L bioreactor. These results show that the identification and regulation of key genes are critical for OAH biosynthesis, which would provide a better research basis for the industrial production of OAH in C. glutamicum.

Optimum flux rerouting for efficient production of naringenin from acetate in engineered Escherichia coli

Background

Microbial production of naringenin has received much attention owing to its pharmaceutical applicability and potential as a key molecular scaffold for various flavonoids. In the microbial fermentation, a cheap and abundant feedstock is required to achieve an economically feasible bioprocess. From this perspective, utilizing acetate for naringenin production could be an effective strategy, with the advantages of both low-cost and abundant feedstock. For the efficient production of naringenin using acetate, identification of the appropriate regulatory node of carbon flux in the biosynthesis of naringenin from acetate would be important. While acetyl-CoA is a key precursor for naringenin production, carbon flux between the TCA cycle and anaplerosis is effectively regulated at the isocitrate node through glyoxylate shunt in acetate metabolism. Accordingly, appropriate rerouting of TCA cycle intermediates from anaplerosis into naringenin biosynthesis via acetyl-CoA replenishment would be required.

Results

This study identified the isocitrate and oxaloacetate (OAA) nodes as key regulatory nodes for the naringenin production using acetate. Precise rerouting at the OAA node for enhanced acetyl-CoA was conducted, avoiding extensive loss of OAA by fine-tuning the expression of pckA (encoding phosphoenolpyruvate carboxykinase) with flux redistribution between naringenin biosynthesis and cell growth at the isocitrate node. Consequently, the flux-optimized strain exhibited a significant increase in naringenin production, a 27.2-fold increase (with a 38.3-fold increase of naringenin yield on acetate) over that by the unoptimized strain, producing 97.02 mg/L naringenin with 21.02 mg naringenin/g acetate, which is a competitive result against those in previous studies on conventional substrates, such as glucose.

Conclusions

Collectively, we demonstrated efficient flux rerouting for maximum naringenin production from acetate in E. coli. This study was the first attempt of naringenin production from acetate and suggested the potential of biosynthesis of various flavonoids derived from naringenin using acetate.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13068-022-02188-w.

The application of CRISPR /Cas mediated gene editing in synthetic biology: Challenges and optimizations

Clustered regularly interspaced short palindromic repeats (CRISPR) and its associated enzymes (Cas) is a simple and convenient genome editing tool that has been used in various cell factories and emerging synthetic biology in the recent past. However, several problems, including off-target effects, cytotoxicity, and low efficiency of multi-gene editing, are associated with the CRISPR/Cas system, which have limited its application in new species. In this review, we briefly describe the mechanisms of CRISPR/Cas engineering and propose strategies to optimize the system based on its defects, including, but not limited to, enhancing targeted specificity, reducing toxicity related to Cas protein, and improving multi-point editing efficiency. In addition, some examples of improvements in synthetic biology are also highlighted. Finally, future perspectives of system optimization are discussed, providing a reference for developing safe genome-editing tools for new species.

CRISPR-Based Approaches for Gene Regulation in Non-Model Bacteria

CRISPR interference (CRISPRi) and CRISPR activation (CRISPRa) have become ubiquitous approaches to control gene expression in bacteria due to their simple design and effectiveness. By regulating transcription of a target gene(s), CRISPRi/a can dynamically engineer cellular metabolism, implement transcriptional regulation circuitry, or elucidate genotype-phenotype relationships from smaller targeted libraries up to whole genome-wide libraries. While CRISPRi/a has been primarily established in the model bacteria Escherichia coli and Bacillus subtilis, a growing numbering of studies have demonstrated the extension of these tools to other species of bacteria (here broadly referred to as non-model bacteria). In this mini-review, we discuss the challenges that contribute to the slower creation of CRISPRi/a tools in diverse, non-model bacteria and summarize the current state of these approaches across bacterial phyla. We find that despite the potential difficulties in establishing novel CRISPRi/a in non-model microbes, over 190 recent examples across eight bacterial phyla have been reported in the literature. Most studies have focused on tool development or used these CRISPRi/a approaches to interrogate gene function, with fewer examples applying CRISPRi/a gene regulation for metabolic engineering or high-throughput screens and selections. To date, most CRISPRi/a reports have been developed for common strains of non-model bacterial species, suggesting barriers remain to establish these genetic tools in undomesticated bacteria. More efficient and generalizable methods will help realize the immense potential of programmable CRISPR-based transcriptional control in diverse bacteria.

Double-Gene Copromoting Expression Analysis in tPA/GH Transgenic Goat Mammary Epithelial Cells and Thrombolytic Activity of tPA In Vitro

Human tissue-plasminogen activator (tPA) is a thrombolytic drug widely used in the treatment of stroke, pulmonary thrombosis, acute myocardial infarction, and other thrombotic diseases. The double genes cointegrated into the organisms and cells can produce a synergistic effect, which will improve the expression level of the target gene. However, the study of the integration of the GH and tPA genes to improve the expression level of tPA has not yet been reported. In order to elucidate this, we generated monoclonal goat mammary epithelial cell lines with tPA/GH double-gene integration and analyzed the tPA expression level in single- and double-gene integrated cells. We selected the mammary gland-specific expressing vectors BLC14/tPA and BLC14/GH with the β-lactoglobulin gene as a regulatory sequence in our previous research. The tPA and GH genes were electronically cotransfected into goat mammary epithelial cells. Resistant cell lines were screened by G418, and transgenic monoclonal cell lines were confirmed by PCR. The tPA expression was induced by prolactin and detected in the cell induction solution after 48 h by ELISA and Western blotting. We detected the tPA biological activity in vitro by fibrin agarose plate assay (FAPA). The results showed that a total of 207 resistant monoclonal cells were obtained, including 126 cell lines with tPA monogenic integration and 51 cell lines with tPA/GH double-gene integration. The rate of double-gene integration was 24.6% (51/207). A total of 48 cells expressed tPA, of which 25.3% (19/75) cells expressed single gene, and 56.9% (29/51) cells expressed double genes. The concentration of tPA in single-gene-expressing cells was 8.0-64.0 μg/mL, and the tPA level in double-gene-expressing cells was significantly higher (200-7200 μg/mL). In addition, the tPA had a relatively strong in vitro thrombolytic activity determined by FAPA. The results showed that goat mammary epithelial cell lines with tPA/GH gene integration were successfully established by electrotransfection, and the expression level of tPA in double-gene integrated cell lines was significantly increased. This study provided a new way for the preparation of a transgenic goat and other animal with high tPA expression by somatic cell nuclear transfer. The findings also laid a foundation for efficient production of pharmaceutical proteins in transgenic animal mammary gland bioreactors in the future.

CRISPR-Cas system in microbial hosts for terpenoid production

Terpenoids represent the largest group of secondary metabolites with variable structures and functions. Terpenoids are well known for their beneficial application in human life, such as pharmaceutical products, vitamins, hormones, anticancer drugs, cosmetics, flavors and fragrances, foods, agriculture, and biofuels. Recently, engineering microbial cells have been provided with a sustainable approach to produce terpenoids with high yields. Noticeably, the clustered regularly interspaced short palindromic repeats (CRISPR)-associated (Cas) system has emerged as one of the most efficient genome-editing technologies to engineer microorganisms for improving terpenoid production. In this review, we summarize the application of the CRISPR-Cas system for the production of terpenoids in microbial hosts such as Escherichia coli, Saccharomyces cerevisiae, Corynebacterium glutamicum, and Pseudomonas putida. CRISPR-Cas9 deactivated Cas9 (dCas9)-based CRISPR (CRISPRi), and the dCas9-based activator (CRISPRa) have been used in either individual or combinatorial systems to control the metabolic flux for enhancing the production of terpenoids. Finally, the prospects of using the CRISPR-Cas system in terpenoid production are also discussed.

Unravelling the promise and limitations of CRISPR/Cas system in natural product research: Approaches and challenges

An incredible array of natural products are produced by plants that serve several ecological functions, including protecting them from herbivores and microbes, attracting pollinators, and dispersing seeds. In addition to their obvious medical applications, natural products serve as flavouring agents, fragrances and many other uses by humans. With the increasing demand for natural products and the development of various gene engineering systems, researchers are trying to modify the plant genome to increase the biosynthetic pathway of the compound of interest or blocking the pathway of unwanted compound synthesis. The clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 has had widespread success in genome editing due to the system's high efficiency, ease of use, and accuracy which revolutionized the genome editing system in living organisms. This article highlights the method of the CRISPR/Cas system, its application in different organisms including microbes, algae, fungi and also higher plants in natural product research, its shortcomings and future prospects. This article is protected by copyright. All rights reserved.

Engineering of microbial cells for L-valine production: challenges and opportunities

L-valine is an essential amino acid that has wide and expanding applications with a suspected growing market demand. Its applicability ranges from animal feed additive, ingredient in cosmetic and special nutrients in pharmaceutical and agriculture fields. Currently, fermentation with the aid of model organisms, is a major method for the production of L-valine. However, achieving the optimal production has often been limited because of the metabolic imbalance in recombinant strains. In this review, the constrains in L-valine biosynthesis are discussed first. Then, we summarize the current advances in engineering of microbial cell factories that have been developed to address and overcome major challenges in the L-valine production process. Future prospects for enhancing the current L-valine production strategies are also discussed.

Recent advances in tuning the expression and regulation of genes for constructing microbial cell factories

To overcome environmental problems caused by the use of fossil resources, microbial cell factories have become a promising technique for the sustainable and eco-friendly development of valuable products from renewable resources. Constructing microbial cell factories with high titers, yields, and productivity requires a balance between growth and production; to this end, tuning gene expression and regulation is necessary to optimise and precisely control complicated metabolic fluxes. In this article, we review the current trends and advances in tuning gene expression and regulation and consider their engineering at each of the three stages of gene regulation: genomic, mRNA, and protein. In particular, the technological approaches utilised in a diverse range of genetic-engineering-based tools for the construction of microbial cell factories are reviewed and representative applications of these strategies are presented. Finally, the prospects for strategies and systems for tuning gene expression and regulation are discussed.

CRISPRi-Library-Guided Target Identification for Engineering Carotenoid Production by Corynebacterium glutamicum

Corynebacterium glutamicum is a prominent production host for various value-added compounds in white biotechnology. Gene repression by dCas9/clustered regularly interspaced short palindromic repeats (CRISPR) interference (CRISPRi) allows for the identification of target genes for metabolic engineering. In this study, a CRISPRi-based library for the repression of 74 genes of C. glutamicum was constructed. The chosen genes included genes encoding enzymes of glycolysis, the pentose phosphate pathway, and the tricarboxylic acid cycle, regulatory genes, as well as genes of the methylerythritol phosphate and carotenoid biosynthesis pathways. As expected, CRISPRi-mediated repression of the carotenogenesis repressor gene crtR resulted in increased pigmentation and cellular content of the native carotenoid pigment decaprenoxanthin. CRISPRi screening identified 14 genes that affected decaprenoxanthin biosynthesis when repressed. Carotenoid biosynthesis was significantly decreased upon CRISPRi-mediated repression of 11 of these genes, while repression of 3 genes was beneficial for decaprenoxanthin production. Largely, but not in all cases, deletion of selected genes identified in the CRISPRi screen confirmed the pigmentation phenotypes obtained by CRISPRi. Notably, deletion of pgi as well as of gapA improved decaprenoxanthin levels 43-fold and 9-fold, respectively. The scope of the designed library to identify metabolic engineering targets, transfer of gene repression to stable gene deletion, and limitations of the approach were discussed.

CRISPR-based metabolic pathway engineering

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

Application of different types of CRISPR/Cas-based systems in bacteria

As important genome editing tools, CRISPR/Cas systems, especially those based on type II Cas9 and type V Cas12a, are widely used in genetic and metabolic engineering of bacteria. However, the intrinsic toxicity of Cas9 and Cas12a-mediated CRISPR/Cas tools can lead to cell death in some strains, which led to the development of endogenous type I and III CRISPR/Cas systems. However, these systems are hindered by complicated development and limited applications. Thus, further development and optimization of CRISPR/Cas systems is needed. Here, we briefly summarize the mechanisms of different types of CRISPR/Cas systems as genetic manipulation tools and compare their features to provide a reference for selecting different CRISPR/Cas tools. Then, we show the use of CRISPR/Cas technology for bacterial strain evolution and metabolic engineering, including genome editing, gene expression regulation and the base editor tool. Finally, we offer a view of future directions for bacterial CRISPR/Cas technology.

CRISPR Tools To Control Gene Expression in Bacteria

CRISPR-Cas systems have been engineered as powerful tools to control gene expression in bacteria. The most common strategy relies on the use of Cas effectors modified to bind target DNA without introducing DNA breaks. These effectors can either block the RNA polymerase or recruit it through activation domains. Here, we discuss the mechanistic details of how Cas effectors can modulate gene expression by blocking transcription initiation or acting as transcription roadblocks. CRISPR-Cas tools can be further engineered to obtain fine-tuned control of gene expression or target multiple genes simultaneously. Several caveats in using these tools have also been revealed, including off-target effects and toxicity, making it important to understand the design rules of engineered CRISPR-Cas effectors in bacteria. Alternatively, some types of CRISPR-Cas systems target RNA and could be used to block gene expression at the posttranscriptional level. Finally, we review applications of these tools in high-throughput screens and the progress and challenges in introducing CRISPR knockdown to other species, including nonmodel bacteria with industrial or clinical relevance. A deep understanding of how CRISPR-Cas systems can be harnessed to control gene expression in bacteria and build powerful tools will certainly open novel research directions.

Establishment of CRISPR interference in Methylorubrum extorquens and application of rapidly mining a new phytoene desaturase involved in carotenoid biosynthesis

The methylotrophic bacterium Methylorubrum extorquens AM1 holds a great potential of a microbial cell factory in producing high value chemicals with methanol as the sole carbon and energy source. However, many gene functions remain unknown, hampering further rewiring of metabolic networks. Clustered regularly interspaced short palindromic repeat interference (CRISPRi) has been demonstrated to be a robust tool for gene knockdown in diverse organisms. In this study, we developed an efficient CRISPRi system through optimizing the promoter strength of Streptococcus pyogenes-derived deactivated cas9 (dcas9). When the dcas9 and sgRNA were respectively controlled by medium PR/tetO and strong PmxaF-g promoters, dynamic repression efficacy of cell growth through disturbing a central metabolism gene glyA was achieved from 41.9 to 96.6% dependent on the sgRNA targeting sites. Furthermore, the optimized CRISPRi system was shown to effectively decrease the abundance of exogenous fluorescent protein gene mCherry over 50% and to reduce the expression of phytoene desaturase gene crtI by 97.7%. We then used CRISPRi technology combined with 26 sgRNAs pool to rapidly discover a new phytoene desaturase gene META1_3670 from 2470 recombinant mutants. The gene function was further verified through gene deletion and complementation as well as phylogenetic tree analysis. In addition, we applied CRISPRi to repress the transcriptional level of squalene-hopene cyclase gene shc involved in hopanoid biosynthesis by 64.9%, which resulted in enhancing 1.9-fold higher of carotenoid production without defective cell growth. Thus, the CRISPRi system developed here provides a useful tool in mining functional gene of M. extorquens as well as in biotechnology for producing high-valued chemicals from methanol. KEY POINTS: Developing an efficient CRISPRi to knockdown gene expression in C1-utilizing bacteria CRISPRi combined with sgRNAs pool to rapidly discover a new phytoene desaturase gene Improvement of carotenoid production by repressing a competitive pathway.

Efficient Multiplex Gene Repression by CRISPR-dCpf1 in Corynebacterium glutamicum

Corynebacterium glutamicum is an important workhorse for industrial production of diversiform bioproducts. Multiplex control of metabolic pathway genes is crucial for maximizing biosynthesis of desired products. However, few tools for simultaneously regulating multiple genes in C. glutamicum have been reported. Here, a CRISPR-dCpf1-based multiplex gene repression system was developed for C. glutamicum. This system successfully repressed two fluorescent reporter genes simultaneously by expressing a dCpf1 (E1006A, D917A) and a designed single crRNA array. To demonstrate applications of this CRISPR-dCpf1 system in metabolic engineering, we applied this system to repress four genes involved in lysine biosynthesis (gltA, pck, pgi, and hom) with a single array, which increased the lysine titer and yield for over 4.0-fold. Quantitative PCR demonstrated that transcription of all the four endogenous target genes were repressed by over 90%. Thus, the CRISPR-dCpf1 system is a simple and effective technique for multiplex gene repression in C. glutamicum and holds promise for metabolic engineering of C. glutamicum to produce valuable chemicals and fuels.

Case study of xylose conversion to glycolate in Corynebacterium glutamicum: Current limitation and future perspective of the CRISPR-Cas systems

RNA-guided genome engineering technologies have been developed for the advanced metabolic engineering of microbial cells to enhance the production of value-added chemicals in Corynebacterium glutamicum as an industrial host. Here, we described the biotransformation of xylose to glycolate using engineered Corynebacterium glutamicum, a well-known industrial amino acid producer. A synthetic pathway involving heterologous D-tagatose 3-epimerase and L-fuculose kinase/aldolase reactions was introduced in C. glutamicum, resulting in 9.9 ± 0.01 g/L glycolate from 20 g/L xylose at a yield of 0.51 g/g (equal to 1.0 mol/mol). Additional glyoxylate reduction pathway developed by CRISPR-Cas12a recombineering has been introduced and attempted to increase the maximum theoretical molar yield of 2.0 (mol/mol). Due to the limitation of the CRISPR-Cas12a recombineering with TTTV PAM sites, advanced CRISPR-Cas systems were suggested for the next-round metabolic engineering for improving the glycolate yield to overcome the current genome-editing tool for metabolic engineering in C. glutamicum.

Bioconversion of Xylose to Ethylene Glycol and Glycolate in Engineered Corynebacterium glutamicum

The biological production of two-carbon compounds (ethylene glycol (EG) and glycolate) has been studied for the sustainable supply of the compounds to the polymer, cosmetic, textile, and medical industries. Here, we demonstrated the bioconversion of xylose to either ethylene glycol (EG) or glycolate using engineered Corynebacterium glutamicum, a well-known industrial amino acid producer. A synthetic ribulose 1-phosphate (Ru1P) pathway involving heterologous d-tagatose 3-epimerase and l-fuculose kinase/aldolase reactions was introduced in C. glutamicum. Subsequently, heterologous expression of Escherichia coli YqhD reductase with the synthetic Ru1P pathway led to ethylene glycol production from xylose. Additional pathway engineering in C. glutamicum by mutating ald, which encodes an aldehyde dehydrogenase, abolished the by-product formation of glycolate during xylose conversion to EG at a yield of 0.75 mol per mol. In addition, the bioconversion of xylose to glycolate was achieved, and the almost maximum molar yield was 0.99 mol per mol xylose in C. glutamicum via the Ru1P pathway. Thus, the synthetic Ru1P pathway in C. glutamicum led bioconversion of xylose to either ethylene glycol or glycolate with high molar yields.

Combined genome editing and transcriptional repression for metabolic pathway engineering in Corynebacterium glutamicum using a catalytically active Cas12a

Corynebacterium glutamicum is a versatile workhorse for producing industrially important commodities. The design of an optimal strain often requires the manipulation of metabolic and regulatory genes to different levels, such as overexpression, downregulation, and deletion. Unfortunately, few tools to achieve multiple functions simultaneously have been reported. Here, a dual-functional clustered regularly interspaced short palindromic repeats (CRISPR) (RE-CRISPR) system that combined genome editing and transcriptional repression was designed using a catalytically active Cas12a (a.k.a. Cpf1) in C. glutamicum. Firstly, gene deletion was achieved using Cas12a under a constitutive promoter. Then, via engineering of the guide RNA sequences, transcriptional repression was successfully achieved using a catalytically active Cas12a with crRNAs containing 15 or 16 bp spacer sequences, whose gene repression efficiency was comparable to that of the canonical system (deactivated Cas12a with full-length crRNAs). Finally, RE-CRISPR was developed to achieve genome editing and transcriptional repression simultaneously by transforming a single crRNA plasmid and Cas12a plasmid. The application of RE-CRISPR was demonstrated to increase the production of cysteine and serine for ~ 3.7-fold and 2.5-fold, respectively, in a single step. This study expands the application of CRISPR/Cas12a-based genome engineering and provides a powerful synthetic biology tool for multiplex metabolic engineering of C. glutamicum.

A simple dual-inducible CRISPR interference system for multiple gene targeting in Corynebacterium glutamicum

The development of CRISPR interference (CRISPRi) technology has dramatically increased the pace and the precision of target identification during platform strain development. In order to develop a simple, reliable, and dual-inducible CRISPRi system for the industrially relevant Corynebacterium glutamicum, we combined two different inducible repressor systems in a single plasmid to separately regulate the expression of dCas9 (anhydro-tetracycline-inducible) and a given single guide RNA (IPTG-inducible). The functionality of the resulting vector was demonstrated by targeting the l-arginine biosynthesis pathway in C. glutamicum. By co-expressing dCas9 and a specific single guide RNA targeting the 5'-region of the argininosuccinate lyase gene argH, the specific activity of the target enzyme was down-regulated and in a l-arginine production strain, l-arginine formation was shifted towards citrulline formation. The system was also employed for down-regulation of multiple genes by concatenating sgRNA sequences encoded on one plasmid. Simultaneous down-regulated expression of both argH and the phosphoglucose isomerase gene pgi proved the potential of the system for multiplex targeting. The system can be a promising tool for further pathway engineering in C. glutamicum. Cumulative effects on targeted genes can be rapidly evaluated avoiding tedious and time-consuming traditional gene knockout approaches.

Metabolic engineering advances and prospects for amino acid production

Amino acid fermentation is one of the major pillars of industrial biotechnology. The multi-billion USD amino acid market is rising steadily and is diversifying. Metabolic engineering is no longer focused solely on strain development for the bulk amino acids L-glutamate and L-lysine that are produced at the million-ton scale, but targets specialty amino acids. These demands are met by the development and application of new metabolic engineering tools including CRISPR and biosensor technologies as well as production processes by enabling a flexible feedstock concept, co-production and co-cultivation schemes. Metabolic engineering advances are exemplified for specialty proteinogenic amino acids, cyclic amino acids, omega-amino acids, and amino acids functionalized by hydroxylation, halogenation and N-methylation.

Heterologous Production of Squalene from Glucose in Engineered Corynebacterium glutamicum Using Multiplex CRISPR Interference and High-Throughput Fermentation

The sustainable production of squalene has driven the development of microbial cell factories due to the limitation of low-yielding bioprocesses from plants and illegal harvesting shark liver. We report the metabolic engineering of Corynebacterium glutamicum to produce squalene from glucose. Combinatorial metabolic engineering strategies for precursor rebalancing, redox balancing, and blocking the competing pathway for the isopentenyl diphosphate availabilities were applied by repressing the target genes using the CRISPR interference. The best engineered strain using high-throughput fermentation produced squalene from glucose at 5.4 ± 0.3 mg/g dry cell weight (DCW) and 105.3 ± 3.0 mg/L, which was a 5.2-fold increase over the parental strain. In addition, flask cultivation of C. glutamicum overexpressing the dxs and idi genes with squalene synthase gene and repressing the idsA gene resulted in production of squalene at 5.8 ± 0.4 mg/g DCW and 82.8 ± 6.2 mg/L, which was a 3.4-fold increase over the parental strain.

CRISPR/Cas-based genome engineering in natural product discovery

This review summarizes the current state of the art of CRISPR/Cas-based genome editing technologies for natural product producers.

Triple deletion of clpC, porB, and mepA enhances production of small ubiquitin-like modifier-N-terminal pro-brain natriuretic peptide in Corynebacterium glutamicum

In our previous work, a two-plasmid CRISPR/Cas9 system was constructed for genome editing in Corynebacterium glutamicum. To increase the transformation efficiency and simplify the plasmid curing steps, an all-in-one CRISPR/Cas9 system was constructed for efficient genome editing. In addition, to research proteolysis during the production of recombinant proteins and generate a host for enhanced expression of recombinant proteins, the system was used to delete three genes, clpC, porB, and mepA in C. glutamicum CGMCC1.15647, which encoded the Clp protease subunit ClpC, anion selective channel protein B, and metallopeptidase A, respectively. After the evaluation of different plasmids and hosts, small ubiquitin-like modifier-N-terminal pro-brain natriuretic peptide (SUMO-NT-proBNP), an important protein used for the diagnosis of mild heart failure was successfully expressed in the triple mutant ΔclpCΔporBΔmepA, which exhibit threefold higher levels of protein expression compared with the wild-type. In conclusion, we created a simplified CRISPR tool for genome editing in C. glutamicum, provided a method to generate a host for enhanced expression of recombinant proteins and successfully expressed SUMO-NT-proBNP in C. glutamicum. This tool and method will greatly facilitate genetic engineering and metabolic optimization of this important platform organism.

CRISPR/Cas-based genome engineering in natural product discovery

This review summarizes the current state of the art of CRISPR/Cas-based genome editing technologies for natural product producers.

CRISPR-Cas9/Cas12a biotechnology and application in bacteria

CRISPR-Cas technologies have greatly reshaped the biology field. In this review, we discuss the CRISPR-Cas with a particular focus on the associated technologies and applications of CRISPR-Cas9 and CRISPR-Cas12a, which have been most widely studied and used. We discuss the biological mechanisms of CRISPR-Cas as immune defense systems, recently-discovered anti-CRISPR-Cas systems, and the emerging Cas variants (such as xCas9 and Cas13) with unique characteristics. Then, we highlight various CRISPR-Cas biotechnologies, including nuclease-dependent genome editing, CRISPR gene regulation (including CRISPR interference/activation), DNA/RNA base editing, and nucleic acid detection. Last, we summarize up-to-date applications of the biotechnologies for synthetic biology and metabolic engineering in various bacterial species.

Rapid identification of unknown carboxyl esterase activity in Corynebacterium glutamicum using RNA-guided CRISPR interference

RNA-guided genome engineering technologies have been developed for the advanced metabolic engineering of microbial cells to enhance production of value-added chemicals in Corynebacterium glutamicum as an industrial host. In this study, the RNA-guided CRISPR interference (CRISPRi) was applied to rapidly identify of unknown genes for native esterase activity in C. glutamicum. Combining with the carboxyl esterase (MekB) protein sequence alignment, two target genes (the cg0961 and cg0754) were selected for the CRISPRi application to investigate the possible native esterase in C. glutamicum. The recombinant strain with repressed expression of the cg0961 gene exhibited almost no capability on degradation of methyl acetate as a substrate of carboxyl esterase. This result was also confirmed in the cg0961 gene deletion mutant. Thus, we concluded that Cg0961 plays a major role of the native carboxyl esterase activity in C. glutamicum. In addition, CRISPRi demonstrated an application for gene identification and its function as another genetic tool for metabolic engineering in C. glutamicum.

CRISPR interference-mediated metabolic engineering of Corynebacterium glutamicum for homo-butyrate production

Combinatorial metabolic engineering enabled the development of efficient microbial cell factories for modulating gene expression to produce desired products. Here, we report the combinatorial metabolic engineering of Corynebacterium glutamicum to produce butyrate by introducing a synthetic butyrate pathway including phosphotransferase and butyrate kinase reactions and repressing the essential acn gene-encoding aconitase, which has been targeted for downregulation in a genome-scale model. An all-in-one clustered regularly interspaced short palindromic repeats interference system for C. glutamicum was used for tunable downregulation of acn in an engineered strain, where by-product-forming reactions were deleted and the synthetic butyrate pathway was inserted, resulting in butyrate production (0.52 ± 0.02 g/L). Subsequently, biotin limitation enabled the engineered strain to produce butyrate (0.58 ± 0.01 g/L) without acetate formation for the entire duration of the culture. These results demonstrate the potential homo-production of butyrate using engineered C. glutamicum. This method can also be applied to other industrial microorganisms.

Applications of CRISPR/Cas System to Bacterial Metabolic Engineering

The clustered regularly interspaced short palindromic repeats/CRISPR-associated (CRISPR/Cas) adaptive immune system has been extensively used for gene editing, including gene deletion, insertion, and replacement in bacterial and eukaryotic cells owing to its simple, rapid, and efficient activities in unprecedented resolution. Furthermore, the CRISPR interference (CRISPRi) system including deactivated Cas9 (dCas9) with inactivated endonuclease activity has been further investigated for regulation of the target gene transiently or constitutively, avoiding cell death by disruption of genome. This review discusses the applications of CRISPR/Cas for genome editing in various bacterial systems and their applications. In particular, CRISPR technology has been used for the production of metabolites of high industrial significance, including biochemical, biofuel, and pharmaceutical products/precursors in bacteria. Here, we focus on methods to increase the productivity and yield/titer scan by controlling metabolic flux through individual or combinatorial use of CRISPR/Cas and CRISPRi systems with introduction of synthetic pathway in industrially common bacteria including Escherichia coli. Further, we discuss additional useful applications of the CRISPR/Cas system, including its use in functional genomics.

Engineering CRISPR interference system in Klebsiella pneumoniae for attenuating lactic acid synthesis

BACKGROUND

Klebsiella pneumoniae is a promising industrial species for bioproduction of bulk chemicals such as 1,3-propanediol, 2,3-butanediol and 3-hydroxypropionic acid (3-HP). However, lactic acid is a troublesome by-product when optimizing for 3-HP production. Therefore, it is highly desirable to minimize lactic acid.

RESULTS

Here, we show that lactic acid synthesis can be largely blocked by an engineered CRISPR interference (CRISPRi) system in K. pneumoniae. EGFP was recruited as a reporter of this CRISPRi system. Fluorescence assay of this CRISPRi system showed that enhanced green fluorescent protein (EGFP) expression level was repressed by 85-90%. To further test this CRISPRi system, guide RNAs were designed to individually or simultaneously target four lactate-producing enzyme genes. Results showed that all lactate-producing enzyme genes were significantly repressed. Notably, D-lactate dehydrogenase (ldhA) was shown to be the most influential enzyme for lactic acid formation in micro-aerobic conditions, as inhibiting ldhA alone led to lactic acid level similar to simultaneously repressing four genes. In shake flask cultivation, the strain coexpressing puuC (an aldehyde dehydrogenase catalyzing 3-hydroxypropionaldehyde to 3-HP) and dCas9-sgRNA inhibiting ldhA produced 1.37-fold 3-HP relative to the reference strain. Furthermore, in bioreactor cultivation, this CRISPRi strain inhibiting ldhA produced 36.7 g/L 3-HP, but only generated 1 g/L lactic acid. Clearly, this engineered CRISPRi system largely simplified downstream separation of 3-HP from its isomer lactic acid, an extreme challenge for 3-HP bioprocess.

CONCLUSIONS

This study offers a deep understanding of lactic acid metabolism in diverse species, and we believe that this CRISPRi system will facilitate biomanufacturing and functional genome studies of K. pneumoniae or beyond.