Genome engineering and gene expression control for bacterial strain development

High-throughput genetic engineering of nonmodel and undomesticated bacteria via iterative site-specific genome integration

Efficient genome engineering is critical to understand and use microbial functions. Despite recent development of tools such as CRISPR-Cas gene editing, efficient integration of exogenous DNA with well-characterized functions remains limited to model bacteria. Here, we describe serine recombinase-assisted genome engineering, or SAGE, an easy-to-use, highly efficient, and extensible technology that enables selection marker-free, site-specific genome integration of up to 10 DNA constructs, often with efficiency on par with or superior to replicating plasmids. SAGE uses no replicating plasmids and thus lacks the host range limitations of other genome engineering technologies. We demonstrate the value of SAGE by characterizing genome integration efficiency in five bacteria that span multiple taxonomy groups and biotechnology applications and by identifying more than 95 heterologous promoters in each host with consistent transcription across environmental and genetic contexts. We anticipate that SAGE will rapidly expand the number of industrial and environmental bacteria compatible with high-throughput genetics and synthetic biology.

Development and Applications of CRISPR/Cas9-Based Genome Editing in Lactobacillus

Lactobacillus, a genus of lactic acid bacteria, plays a crucial function in food production preservation, and probiotics. It is particularly important to develop new Lactobacillus strains with superior performance by gene editing. Currently, the identification of its functional genes and the mining of excellent functional genes mainly rely on the traditional gene homologous recombination technology. CRISPR/Cas9-based genome editing is a rapidly developing technology in recent years. It has been widely applied in mammalian cells, plants, yeast, and other eukaryotes, but less in prokaryotes, especially Lactobacillus. Compared with the traditional strain improvement methods, CRISPR/Cas9-based genome editing can greatly improve the accuracy of Lactobacillus target sites and achieve traceless genome modification. The strains obtained by this technology may even be more efficient than the traditional random mutation methods. This review examines the application and current issues of CRISPR/Cas9-based genome editing in Lactobacillus, as well as the development trend of CRISPR/Cas9-based genome editing in Lactobacillus. In addition, the fundamental mechanisms of CRISPR/Cas9-based genome editing are also presented and summarized.

Rational development of mycobacteria cell factory for advancing the steroid biomanufacturing

Steroidal resource occupies a vital proportion in the pharmaceutical industry attributing to their important therapeutic effects on fertility, anti-inflammatory and antiviral activities. Currently, microbial transformation from phytosterol has become the dominant strategy of steroidal drug intermediate synthesis that bypasses the traditional chemical route. Mycobacterium sp. serve as the main industrial microbial strains that are capable of introducing selective functional modifications of steroidal intermediate, which has become an indispensable platform for steroid biomanufacturing. By reviewing the progress in past two decades, the present paper concentrates mainly on the microbial rational modification aspects that include metabolic pathway editing, key enzymes engineering, material transport pathway reinforcement, toxic metabolic intermediates removal and byproduct reconciliation. In addition, progress on omics analysis and direct genetic manipulation are summarized and classified that may help reform the industrial hosts with more efficiency. The paper provides an insightful present for steroid biomanufacturing especially on the current trends and prospects of mycobacteria.

Development and optimization of Lysis gene E as a counter-selection marker with high stringency

BACKGROUND

Seamless modification of bacterial chromosomes is widely performed in both theoretical and practical research. For this purpose, excellent counter-selection marker genes with high stringency are needed.

MAIN METHODS AND MAJOR RESULTS

The lysis gene E was first constructed under the control of the P_L promoter and the cI857 repressor. At 42°C, it could effectively kill Escherichia coli and seamless modification in this bacterium using E as a counter-selection marker was successfully conducted. It also works in another gram-negative strain, Serratia marcescens, under the control of the Arac/P_BAD regulatory system. By combining lysis gene E and kil, the counter-selection frequencies of the P_L -kil-sd-E cassette in E. coli reached 4.9 × 10^-8 and 3.2 × 10^-8 at two test loci, which are very close to frequencies observed with the best counter-selection systems reported, the inducible toxin systems. Under the control of the Arac/P_BAD , the counter-selection frequency of P_BAD -kil-sd-E in S. marcescens reached the level of 10^-7 at four test loci. By expressing the araC gene from plasmid pKDsg-ack, 5- to 17-fold improvements in counter-selection stringency were observed at these loci. A surprisingly low counter-selection frequency of 4.9 × 10^-9 was obtained at the marR-1 locus, which reflects the highest stringency for a counter-selection cassette reported thus far. Similarly, at the araB locus of E. coli, the counter-selection frequency of P_BAD -kil-sd-E was 3 × 10^-9 after introducing plasmid pKDsg-ack.

CONCLUSIONS AND IMPLICATIONS

We have developed and optimized a new universal counter-selection marker based on lysis gene E. The best counter-selection stringency of this new marker exceeds the inducible toxin system several fold. Our work can also provide inspiration for improving counter-selection stringency based on existing markers. This article is protected by copyright. All rights reserved.

Improved dsDNA recombineering enables versatile multiplex genome engineering of kilobase-scale sequences in diverse bacteria

Recombineering assisted multiplex genome editing generally uses single-stranded oligonucleotides for site directed mutational changes. It has proven highly efficient for functional screens and to optimize microbial cell factories. However, this approach is limited to relatively small mutational changes. Here, we addressed the challenges involved in the use of double-stranded DNA substrates for multiplex genome engineering. Recombineering is mediated by phage single-strand annealing proteins annealing ssDNAs into the replication fork. We apply this insight to facilitate the generation of ssDNA from the dsDNA substrate and to alter the speed of replication by elevating the available deoxynucleoside triphosphate (dNTP) levels. Intracellular dNTP concentration was elevated by ribonucleotide reductase overexpression or dNTP addition to establish double-stranded DNA Recombineering-assisted Multiplex Genome Engineering (dReaMGE), which enables rapid and flexible insertional and deletional mutagenesis at multiple sites on kilobase scales in diverse bacteria without the generation of double-strand breaks or disturbance of the mismatch repair system. dReaMGE can achieve combinatorial genome engineering works, for example, alterations to multiple biosynthetic pathways, multiple promoter or gene insertions, variations of transcriptional regulator combinations, within a few days. dReaMGE adds to the repertoire of bacterial genome engineering to facilitate discovery, functional genomics, strain optimization and directed evolution of microbial cell factories.

CRISPR/Cas9 mediated T7 RNA polymerase gene knock-in in E. coli BW25113 makes T7 expression system work efficiently

T7 Expression System is a common method of ensuring tight control and high-level induced expression. However, this system can only work in some bacterial strains in which the T7 RNA Polymerase gene resides in the chromosome. In this study, we successfully introduced a chromosomal copy of the T7 RNA Polymerase gene under control of the lacUV5 promoter into Escherichia coli BW25113. The T7 Expression System worked efficiently in this mutant strain named BW25113-T7. We demonstrated that this mutant strain could satisfactorily produce 5-Aminolevulinic Acid via C5 pathway. A final study was designed to enhance the controllability of T7 Expression System in this mutant strain by constructing a T7 Promoter Variants Library. These efforts advanced E. coli BW25113-T7 to be a practical host for future metabolic engineering efforts.

Current perspectives on integrated approaches to enhance lipid accumulation in microalgae

In recent years, research initiatives on renewable bioenergy or biofuels have been gaining momentum, not only due to fast depletion of finite reserves of fossil fuels but also because of the associated concerns for the environment and future energy security. In the last few decades, interest is growing concerning microalgae as the third-generation biofuel feedstock. The CO₂ fixation ability and conversion of it into value-added compounds, devoid of challenging food and feed crops, make these photosynthetic microorganisms an optimistic producer of biofuel from an environmental point of view. Microalgal-derived fuels are currently being considered as clean, renewable, and promising sustainable biofuel. Therefore, most research targets to obtain strains with the highest lipid productivity and a high growth rate at the lowest cultivation costs. Different methods and strategies to attain higher biomass and lipid accumulation in microalgae have been extensively reported in the previous research, but there are fewer inclusive reports that summarize the conventional methods with the modern techniques for lipid enhancement and biodiesel production from microalgae. Therefore, the current review focuses on the latest techniques and advances in different cultivation conditions, the effect of different abiotic and heavy metal stress, and the role of nanoparticles (NPs) in the stimulation of lipid accumulation in microalgae. Techniques such as genetic engineering, where particular genes associated with lipid metabolism, are modified to boost lipid synthesis within the microalgae, the contribution of "Omics" in metabolic pathway studies. Further, the contribution of CRISPR/Cas9 system technique to the production of microalgae biofuel is also briefly described.

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

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

Genome-Scale Metabolic Modeling of Escherichia coli and Its Chassis Design for Synthetic Biology Applications

Genome-scale metabolic modeling is and will continue to play a central role in computational systems metabolic engineering and synthetic biology applications for the productions of chemicals and antibiotics. To that end, a survey and workflows of methods used for the development of high-quality genome-scale metabolic models (GEMs) and chassis design for synthetic biology are described here. The chapter consists of two parts (a) the methods of development of GEMs (Escherichia coli as a case study) and (b) E. coli chassis design for synthetic production of 1,4-butanediol (BDO). The methods described here can guide existing and future development of GEMs coupled with host chassis design for synthetic productions of novel antibiotics.

Microbial Base Editing: A Powerful Emerging Technology for Microbial Genome Engineering

Genome engineering is crucial for answering fundamental questions about, and exploring practical applications of, microorganisms. Various microbial genome-engineering tools, including CRISPR/Cas-enhanced homologous recombination (HR), have been developed, with ever-improving simplicity, efficiency, and applicability. Recently, a powerful emerging technology based on CRISPR/Cas-nucleobase deaminase fusions, known as base editing, opened new avenues for microbial genome engineering. Base editing enables nucleotide transition without inducing lethal double-stranded (ds)DNA cleavage, adding foreign donor DNA, or depending on inefficient HR. Here, we review ongoing efforts to develop and apply base editing to engineer industrially and clinically relevant microorganisms. We also summarize bioinformatics tools that would greatly facilitate guide (g)RNA design and sequencing data analysis and discuss the future challenges and prospects associated with this technology.

CRISPR interference (CRISPRi) as transcriptional repression tool for Hungateiclostridium thermocellum DSM 1313

Hungateiclostridium thermocellum DSM 1313 has biotechnological potential as a whole-cell biocatalyst for ethanol production using lignocellulosic renewable sources. The full exploitation of H. thermocellum has been hampered due to the lack of simple and high-throughput genome engineering tools. Recently in our research group, a thermophilic bacterial CRISPR-Cas9-based system has been developed as a transcriptional suppression tool for regulation of gene expression. We applied ThermoCas9-based CRISPR interference (CRISPRi) to repress the H. thermocellum central metabolic lactate dehydrogenase (ldh) and phosphotransacetylase (pta) genes. The effects of repression on target genes were studied based on transcriptional expression and product formation. Single-guide RNA (sgRNA) under the control of native intergenic 16S/23S rRNA promoter from H. thermocellum directing the ThermodCas9 to the promoter region of both pta and ldh silencing transformants reduced expression up to 67% and 62% respectively. This resulted in 24% and 17% decrease in lactate and acetate production, correspondingly. Hence, the CRISPRi approach for H. thermocellum to downregulate metabolic genes can be used for remodelling of metabolic pathways without the requisite for genome engineering. These data established for the first time the feasibility of employing CRISPRi-mediated gene repression of metabolic genes in H. thermocellum DSM 1313.

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.

Biomass and lipid induction strategies in microalgae for biofuel production and other applications

The use of fossil fuels has been strongly related to critical problems currently affecting society, such as: global warming, global greenhouse effects and pollution. These problems have affected the homeostasis of living organisms worldwide at an alarming rate. Due to this, it is imperative to look for alternatives to the use of fossil fuels and one of the relevant substitutes are biofuels. There are different types of biofuels (categories and generations) that have been previously explored, but recently, the use of microalgae has been strongly considered for the production of biofuels since they present a series of advantages over other biofuel production sources: (a) they don’t need arable land to grow and therefore do not compete with food crops (like biofuels produced from corn, sugar cane and other plants) and; (b) they exhibit rapid biomass production containing high oil contents, at least 15 to 20 times higher than land based oleaginous crops. Hence, these unicellular photosynthetic microorganisms have received great attention from researches to use them in the large-scale production of biofuels. However, one disadvantage of using microalgae is the high economic cost due to the low-yields of lipid content in the microalgae biomass. Thus, development of different methods to enhance microalgae biomass, as well as lipid content in the microalgae cells, would lead to the development of a sustainable low-cost process to produce biofuels. Within the last 10 years, many studies have reported different methods and strategies to induce lipid production to obtain higher lipid accumulation in the biomass of microalgae cells; however, there is not a comprehensive review in the literature that highlights, compares and discusses these strategies. Here, we review these strategies which include modulating light intensity in cultures, controlling and varying CO₂ levels and temperature, inducing nutrient starvation in the culture, the implementation of stress by incorporating heavy metal or inducing a high salinity condition, and the use of metabolic and genetic engineering techniques coupled with nanotechnology.

Advances in engineered trans-acting regulatory RNAs and their application in bacterial genome engineering

Small noncoding RNAs, a large class of ancient posttranscriptional regulators, are increasingly recognized and utilized as key modulators of gene expression in a broad range of microorganisms. Owing to their small molecular size and the central role of Watson-Crick base pairing in defining their interactions, structure and function, numerous diverse types of trans-acting RNA regulators that are functional at the DNA, mRNA and protein levels have been experimentally characterized. It has become increasingly clear that most small RNAs play critical regulatory roles in many processes and are, therefore, considered to be powerful tools for genetic engineering and synthetic biology. The trans-acting regulatory RNAs accelerate this ability to establish potential framework for genetic engineering and genome-scale engineering, which allows RNA structure characterization, easier to design and model compared to DNA or protein-based systems. In this review, we summarize recent advances in engineered trans-acting regulatory RNAs that are used in bacterial genome-scale engineering and in novel cellular capabilities as well as their implementation in wide range of biotechnological, biological and medical applications.

Metabolic engineering of Corynebacterium glutamicum by synthetic small regulatory RNAs

Corynebacterium glutamicum is an important platform strain that is wildly used in industrial production of amino acids and various other biochemicals. However, due to good genomic stability, C. glutamicum is more difficult to engineer than genetically tractable hosts. Herein, a synthetic small regulatory RNA (sRNA)-based gene knockdown strategy was developed for C. glutamicum. The RNA chaperone Hfq from Escherichia coli and a rationally designed sRNA consisting of the E. coli MicC scaffold and a target binding site were proven to be indispensable for repressing green fluorescent protein expression in C. glutamicum. The synthetic sRNA system was applied to improve glutamate production through knockdown of pyk, ldhA, and odhA, resulting almost a threefold increase in glutamate titer and yield. Gene transcription and enzyme activity were down-regulated by up to 80%. The synthetic sRNA system developed holds promise to accelerate C. glutamicum metabolic engineering for producing valuable chemicals and fuels.

Multiple-step chromosomal integration of divided segments from a large DNA fragment via CRISPR/Cas9 in Escherichia coli

Although CRISPR/Cas9-mediated gene editing technology has developed vastly in Escherichia coli, the chromosomal integration of large DNA fragment is still challenging compared with gene deletion and small fragment integration. Moreover, to guarantee sufficient Cas9-induced double-strand breaks, it is usually necessary to design several gRNAs to select the appropriate one. Accordingly, we established a practical daily routine in the laboratory work, involving multiple-step chromosomal integration of the divided segments from a large DNA fragment. First, we introduced and optimized the protospacers from Streptococcus pyogenes in E. coli W3110. Next, the appropriate fragment size for each round of integration was optimized to be within 3-4 kb. Taking advantage of the optimized protospacer/gRNA pairs, a DNA fragment with a total size of 15.4 kb, containing several key genes for uridine biosynthesis, was integrated into W3110 chromosome, which produced 5.6 g/L uridine in shake flask fermentation. Using this strategy, DNA fragments of virtually any length can be integrated into a suitable genomic site, and two gRNAs can be alternatively used, avoiding the tedious construction of gRNA-expressing plasmids. This study thus presents a useful strategy for large DNA fragment integration into the E. coli chromosome, which can be easily adapted for use in other bacteria.

Regulatory non-coding sRNAs in bacterial metabolic pathway engineering

Non-coding RNAs (ncRNAs) are versatile and powerful controllers of gene expression that have been increasingly linked to cellular metabolism and phenotype. In bacteria, identified and characterized ncRNAs range from trans-acting, multi-target small non-coding RNAs to dynamic, cis-encoded regulatory untranslated regions and riboswitches. These native regulators have inspired the design and construction of many synthetic RNA devices. In this work, we review the design, characterization, and impact of ncRNAs in engineering both native and exogenous metabolic pathways in bacteria. We also consider the opportunities afforded by recent high-throughput approaches for characterizing sRNA regulators and their corresponding networks to showcase their potential applications and impact in engineering bacterial metabolism.

Metabolic engineering of Escherichia coli for high-yield uridine production

Uridine is a kind of pyrimidine nucleoside that has been widely applied in the pharmaceutical industry. Although microbial fermentation is a promising method for industrial production of uridine, an efficient microbial cell factory is still lacking. In this study, we constructed a metabolically engineered Escherichia coli capable of high-yield uridine production. First, we developed a CRISPR/Cas9-mediated chromosomal integration strategy to integrate large DNA into the E. coli chromosome, and a 9.7 kb DNA fragment including eight genes in the pyrimidine operon of Bacillus subtilis F126 was integrated into the yghX locus of E. coli W3110. The resultant strain produced 3.3 g/L uridine and 4.5 g/L uracil in shake flask culture for 32 h. Subsequently, five genes involved in uridine catabolism were knocked out, and the uridine titer increased to 7.8 g/L. As carbamyl phosphate, aspartate, and 5'-phosphoribosyl pyrophosphate are important precursors for uridine synthesis, we further modified several metabolism-related genes and synergistically improved the supply of these precursors, leading to a 76.9% increase in uridine production. Finally, nupC and nupG encoding nucleoside transport proteins were deleted, and the extracellular uridine accumulation increased to 14.5 g/L. After 64 h of fed-batch fermentation, the final engineered strain UR6 produced 70.3 g/L uridine with a yield and productivity of 0.259 g/g glucose and 1.1 g/L/h, respectively. To the best of our knowledge, this is the highest uridine titer and productivity ever reported for the fermentative production of uridine.

Unmarked genetic manipulation in Bacillus subtilis by natural co-transformation

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

CRISPR/Cas9 editing genome of extremophile Halomonas spp

Extremophiles are suitable chassis for developing the next generation industrial biotechnology (NGIB) due to their resistance to microbial contamination. However, engineering extremophiles are not an easy task. Halomonas, an industrially interesting halophile able to grow under unsterile and continuous conditions in large-scale processes, can only be engineered using suicide plasmid-mediated two-step homologous recombination which is very laborious and time-consuming (up to half a year). A convenient approach for the engineering of halophiles that can possibly be extended to other extremophiles is therefore urgently required. To meet this requirement, a rapid, efficient and scarless method via CRISPR/Cas9 system was developed in this study for genome editing in Halomonas. The method achieved the highest efficiency of 100%. When eight different mutants were constructed via this special CRISPR/Cas9 method to study the combinatorial influences of four different genes on the glucose catabolism in H. bluephagenesis TD01, it took only three weeks to complete the deletion and insertion of up to 4.5 kb DNA. H. bluephagenesis was designed to produce a microbial copolymer P(3HB-co-3HV) consisting of 3-hydroxybutyrate (3HB) and 3-hydroxyvalerate (3HV). The CRISPR/Cas9 was employed to delete the prpC gene in H. bluephagenesis TD01. Shake flask studies showed that the 3HV fraction in the copolymers increased approximately 16-folds, demonstrating enhanced effectiveness of the ΔprpC mutant to synthesize PHBV. This genome engineering strategy significantly speeds up the studies on Halomonas engineering, opening up a wide area for developing NGIB.

L-Cysteine Production in Escherichia coli Based on Rational Metabolic Engineering and Modular Strategy

L-cysteine is an amino acid with important physiological functions and has a wide range of applications in medicine, food, animal feed, and cosmetics industry. In this study, the L-cysteine synthesis in Escherichia coliEscherichia coli is divided into four modules: the transport module, sulfur module, precursor module, and degradation module. The engineered strain LH03 (overexpression of the feedback-insensitive cysE and the exporter ydeD in JM109) accumulated 45.8 mg L^-1 of L-cysteine in 48 hr with yield of 0.4% g/g glucose. Further modifications of strains and culture conditions which based on the rational metabolic engineering and modular strategy improved the L-cysteine biosynthesis significantly. The engineered strain LH06 (with additional overexpression of serA, serC, and serB and double mutant of tnaA and sdaA in LH03) produced 620.9 mg L^-1 of L-cysteine with yield of 6.0% g/g glucose, which increased the production by 12 times and the yield by 14 times more than those of LH03 in the original condition. In fed-batch fermentation performed in a 5-L reactor, the concentration of L-cysteine achieved 5.1 g L^-1 in 32 hr. This work demonstrates that the combination of rational metabolic engineering and module strategy is a promising approach for increasing the L-cysteine production in E. coli.

A Novel Tool for Microbial Genome Editing Using the Restriction-Modification System

Scarless genetic manipulation of genomes is an essential tool for biological research. The restriction-modification (R-M) system is a defense system in bacteria that protects against invading genomes on the basis of its ability to distinguish foreign DNA from self DNA. Here, we designed an R-M system-mediated genome editing (RMGE) technique for scarless genetic manipulation in different microorganisms. For bacteria with Type IV REase, an RMGE technique using the inducible DNA methyltransferase gene, bceSIIM (RMGE-bceSIIM), as the counter-selection cassette was developed to edit the genome of Escherichia coli. For bacteria without Type IV REase, an RMGE technique based on a restriction endonuclease (RMGE-mcrA) was established in Bacillus subtilis. These techniques were successfully used for gene deletion and replacement with nearly 100% counter-selection efficiencies, which were higher and more stable compared to conventional methods. Furthermore, precise point mutation without limiting sites was achieved in E. coli using RMGE-bceSIIM to introduce a single base mutation of A128C into the rpsL gene. In addition, the RMGE-mcrA technique was applied to delete the CAN1 gene in Saccharomyces cerevisiae DAY414 with 100% counter-selection efficiency. The effectiveness of the RMGE technique in E. coli, B. subtilis, and S. cerevisiae suggests the potential universal usefulness of this technique for microbial genome manipulation.

CRISPRi mediated phosphoenolpyruvate carboxylase regulation to enhance the production of lipid in Chlamydomonas reinhardtii

In this study, CRISPRi (clustered regularly interspaced short palindromic repeats interference) was used for the first time to regulate expression of exogenously supplied rfp gene as a proof-of-concept, and endogenous PEPC1 gene as a proof-of-function in Chlamydomonas reinhardtii. The efficiency of 94% and stability of 7 generations via CRISPRi mediated gene regulation in C. reinhardtii have been demonstrated by RFP. Gene PEPC1 encoding proteins are essential for controlling the carbon flux that enters the TCA cycle and plays a crucial role in carbon partitioning of substrates in competition with lipid synthesis. All CrPEPC1 down-regulated strains have lower chlorophyll color, but higher biomass concentration and lipid accumulation rate. The present results revealed that CRISPRi based transcriptional silencing was applicable in C. reinhardtii and expanded the way to improve the yield, titer and productivity of microalgae-based products.

A novel and versatile dual fluorescent reporter tool for the study of gene expression and regulation in multi- and single copy number

To unravel intricate mechanisms of gene regulation it is imperative to work in physiologically relevant conditions and therefore preferentially in single copy constructs, which are not always easy to manipulate. Such in vivo studies are generally based on enzymatic assays, microarrays, RNA-seq, qRT-PCR, or multicopy reporter gene systems, frequently with β-galactosidase, luciferase or a fluorescent protein as reporter. Each method has its advantages and shortcomings and may require validation. Enzyme assays are generally reliable but may be quite complex, time consuming, and require a (expensive) substrate. Microarrays and RNA-seq provide a genome wide view of gene expression but may rapidly become expensive and time consuming especially for detailed studies with large numbers of mutants, different growth conditions and multiple time points. Multicopy reporter gene systems are handy to generate numerous constructs but may not provide accurate information due to titration effects of trans-acting regulatory elements. Therefore and in spite of the existence of various reporter systems, there is still need for an efficient and user-friendly tool for detailed studies and high throughput screenings. Here we develop and validate a novel and versatile fluorescent reporter tool to study gene regulation in single copy mode that enables real-time measurement. This tool bears two independent fluorescent reporters that allow high throughput screening and standardization, and combines modern efficient cloning methods (multicopy, in vitro manipulation) with classical genetics (in vivo homologous recombination with a stable, self-transmissible episome) to generate multi- and single copy reporter systems. We validate the system with constitutive and differentially regulated promoters and show that the tool can equally be used with heterologous transcription factors. The flexibility and versatility of this dual reporter tool in combination with an easy conversion from a multicopy plasmid to a stable, single copy reporter system makes this system unique and attractive for a variety of applications. Examples are in vivo studies of DNA-binding transcription factors (single copy) or screening of promoter and RBS libraries (multicopy) for synthetic biology purposes.

Challenges and Advances for Genetic Engineering of Non-model Bacteria and Uses in Consolidated Bioprocessing

Metabolic diversity in microorganisms can provide the basis for creating novel biochemical products. However, most metabolic engineering projects utilize a handful of established model organisms and thus, a challenge for harnessing the potential of novel microbial functions is the ability to either heterologously express novel genes or directly utilize non-model organisms. Genetic manipulation of non-model microorganisms is still challenging due to organism-specific nuances that hinder universal molecular genetic tools and translatable knowledge of intracellular biochemical pathways and regulatory mechanisms. However, in the past several years, unprecedented progress has been made in synthetic biology, molecular genetics tools development, applications of omics data techniques, and computational tools that can aid in developing non-model hosts in a systematic manner. In this review, we focus on concerns and approaches related to working with non-model microorganisms including developing molecular genetics tools such as shuttle vectors, selectable markers, and expression systems. In addition, we will discuss: (1) current techniques in controlling gene expression (transcriptional/translational level), (2) advances in site-specific genome engineering tools [homologous recombination (HR) and clustered regularly interspaced short palindromic repeats (CRISPR)], and (3) advances in genome-scale metabolic models (GSMMs) in guiding design of non-model species. Application of these principles to metabolic engineering strategies for consolidated bioprocessing (CBP) will be discussed along with some brief comments on foreseeable future prospects.

Preliminary Transcriptome Analysis of Mature Biofilm and Planktonic Cells of Salmonella Enteritidis Exposure to Acid Stress

Salmonella has emerged as a well-recognized food-borne pathogen, with many strains able to form biofilms and thus cause cross-contamination in food processing environments where acid-based disinfectants are widely encountered. In the present study, RNA sequencing was employed to establish complete transcriptome profiles of Salmonella Enteritidis in the forms of planktonic and biofilm-associated cells cultured in Tryptic Soytone Broth (TSB) and acidic TSB (aTSB). The gene expression patterns of S. Enteritidis significantly differed between biofilm-associated and planktonic cells cultivated under the same conditions. The assembled transcriptome of S. Enteritidis in this study contained 5,442 assembled transcripts, including 3,877 differentially expressed genes (DEGs) identified in biofilm and planktonic cells. These DEGs were enriched in terms such as regulation of biological process, metabolic process, macromolecular complex, binding and transferase activity, which may play crucial roles in the biofilm formation of S. Enteritidis cultivated in aTSB. Three significant pathways were observed to be enriched under acidic conditions: bacterial chemotaxis, porphyrin-chlorophyll metabolism and sulfur metabolism. In addition, 15 differentially expressed novel non-coding small RNAs (sRNAs) were identified, and only one was found to be up-regulated in mature biofilms. This preliminary study of the S. Enteritidis transcriptome serves as a basis for future investigations examining the complex network systems that regulate Salmonella biofilm in acidic environments, which provide information on biofilm formation and acid stress interaction that may facilitate the development of novel disinfection procedures in the food processing industry.

Controlling Citrate Synthase Expression by CRISPR/Cas9 Genome Editing for n-Butanol Production in Escherichia coli

Genome editing using CRISPR/Cas9 was successfully demonstrated in Esherichia coli to effectively produce n-butanol in a defined medium under microaerobic condition. The butanol synthetic pathway genes including those encoding oxygen-tolerant alcohol dehydrogenase were overexpressed in metabolically engineered E. coli, resulting in 0.82 g/L butanol production. To increase butanol production, carbon flux from acetyl-CoA to citric acid cycle should be redirected to acetoacetyl-CoA. For this purpose, the 5'-untranslated region sequence of gltA encoding citrate synthase was designed using an expression prediction program, UTR designer, and modified using the CRISPR/Cas9 genome editing method to reduce its expression level. E. coli strains with decreased citrate synthase expression produced more butanol and the citrate synthase activity was correlated with butanol production. These results demonstrate that redistributing carbon flux using genome editing is an efficient engineering tool for metabolite overproduction.

Genome engineering Escherichia coli for L-DOPA overproduction from glucose

Genome engineering has become a powerful tool for creating useful strains in research and industry. In this study, we applied singleplex and multiplex genome engineering approaches to construct an E. coli strain for the production of L-DOPA from glucose. We first used the singleplex genome engineering approach to create an L-DOPA-producing strain, E. coli DOPA-1, by deleting transcriptional regulators (tyrosine repressor tyrR and carbon storage regulator A csrA), altering glucose transport from the phosphotransferase system (PTS) to ATP-dependent uptake and the phosphorylation system overexpressing galactose permease gene (galP) and glucokinase gene (glk), knocking out glucose-6-phosphate dehydrogenase gene (zwf) and prephenate dehydratase and its leader peptide genes (pheLA) and integrating the fusion protein chimera of the downstream pathway of chorismate. Then, multiplex automated genome engineering (MAGE) based on 23 targets was used to further improve L-DOPA production. The resulting strain, E. coli DOPA-30N, produced 8.67 g/L of L-DOPA in 60 h in a 5 L fed-batch fermentation. This titer is the highest achieved in metabolically engineered E. coli having PHAH activity from glucose.

A BioBrick™-Compatible Vector for Allelic Replacement Using the XylE Gene as Selection Marker

Background

Circular plasmid-mediated homologous recombination is commonly used for marker-less allelic replacement, exploiting the endogenous recombination machinery of the host. Common limitations of existing methods include high false positive rates due to mutations in counter-selection genes, and limited applicability to specific strains or growth media. Finally, solutions compatible with physical standards, such as the BioBrick™, are not currently available, although they proved to be successful in the design of other replicative or integrative plasmids.

Findings

We illustrate pBBknock, a novel BioBrick™-compatible vector for allelic replacement in Escherichia coli. It includes a temperature-sensitive replication origin and enables marker-less genome engineering via two homologous recombination events. Chloramphenicol resistance allows positive selection of clones after the first event, whereas a colorimetric assay based on the xylE gene provides a simple way to screen clones in which the second recombination event occurs. Here we successfully use pBBknock to delete the lactate dehydrogenase gene in E. coli W, a popular host used in metabolic engineering.

Conclusions

Compared with other plasmid-based solutions, pBBknock has a broader application range, not being limited to specific strains or media. We expect that pBBknock will represent a versatile solution both for practitioners, also among the iGEM competition teams, and for research laboratories that use BioBrick™-based assembly procedures.

Electronic supplementary material

The online version of this article (doi:10.1186/s12575-016-0036-z) contains supplementary material, which is available to authorized users.

Metabolic engineering with systems biology tools to optimize production of prokaryotic secondary metabolites

This Highlight examines current status of metabolic engineering and systems biology tools deployed for the optimal production of prokaryotic secondary metabolites.

Antisense-RNA-Mediated Gene Downregulation in Clostridium pasteurianum

Clostridium pasteurianum is receiving growing attention for its unique metabolic properties, particularly its ability to convert waste glycerol and glycerol-rich byproducts into butanol, a prospective biofuel. Genetic tool development and whole genome sequencing have recently been investigated to advance the genetic tractability of this potential industrial host. Nevertheless, methodologies for tuning gene expression through plasmid-borne expression and chromosomal gene downregulation are still absent. Here we demonstrate plasmid-borne heterologous gene expression and gene knockdown using antisense RNA in C. pasteurianum. We first employed a common thermophilic β-galactosidase (lacZ) gene reporter system from Thermoanaerobacterium thermosulfurogenes to characterize two promoters involved in the central fermentative metabolism of C. pasteurianum. Due to a higher level of constitutive lacZ expression compared to the ferredoxin gene (fdx) promoter, the thiolase (thl) promoter was selected to drive expression of asRNA. Expression of a lacZ asRNA resulted in 52%–58% downregulation of β-galactosidase activity compared to the control strain throughout the duration of culture growth. Subsequent implementation of our asRNA approach for downregulation of the native hydrogenase I gene (hydA) in C. pasteurianum resulted in altered end product distribution, characterized by an increase in production of reduced metabolites, particularly butyrate (40% increase) and ethanol (25% increase). Knockdown of hydA was also accompanied by increased acetate formation and lower levels of 1,3-propanediol, signifying a dramatic shift in cellular metabolism in response to inhibition of the hydrogenase enzyme. The methodologies described herein for plasmid-based heterologous gene expression and antisense-RNA-mediated gene knockdown should promote rational metabolic engineering of C. pasteurianum for enhanced production of butanol as a prospective biofuel.

Rapid and high-throughput construction of microbial cell-factories with regulatory noncoding RNAs

Due to global crises such as pollution and depletion of fossil fuels, sustainable technologies based on microbial cell-factories have been garnering great interest as an alternative to chemical factories. The development of microbial cell-factories is imperative in cutting down the overall manufacturing cost. Thus, diverse metabolic engineering strategies and engineering tools have been established to obtain a preferred genotype and phenotype displaying superior productivity. However, these tools are limited to only a handful of genes with permanent modification of a genome and significant labor costs, and this is one of the bottlenecks associated with biofactory construction. Therefore, a groundbreaking rapid and high-throughput engineering tool is needed for efficient construction of microbial cell-factories. During the last decade, copious small noncoding RNAs (ncRNAs) have been discovered in bacteria. These are involved in substantial regulatory roles like transcriptional and post-transcriptional gene regulation by modulating mRNA elongation, stability, or translational efficiency. Because of their vulnerability, ncRNAs can be used as another layer of conditional control over gene expression without modifying chromosomal sequences, and hence would be a promising high-throughput tool for metabolic engineering. Here, we review successful design principles and applications of ncRNAs for high-throughput metabolic engineering or physiological studies of diverse industrially important microorganisms.

Directed evolution combined with synthetic biology strategies expedite semi-rational engineering of genes and genomes

Owing to our limited understanding of the relationship between sequence and function and the interaction between intracellular pathways and regulatory systems, the rational design of enzyme-coding genes and de novo assembly of a brand-new artificial genome for a desired functionality or phenotype are difficult to achieve. As an alternative approach, directed evolution has been widely used to engineer genomes and enzyme-coding genes. In particular, significant developments toward DNA synthesis, DNA assembly (in vitro or in vivo), recombination-mediated genetic engineering, and high-throughput screening techniques in the field of synthetic biology have been matured and widely adopted, enabling rapid semi-rational genome engineering to generate variants with desired properties. In this commentary, these novel tools and their corresponding applications in the directed evolution of genomes and enzymes are discussed. Moreover, the strategies for genome engineering and rapid in vitro enzyme evolution are also proposed.

Developments in the tools and methodologies of synthetic biology

Synthetic biology is principally concerned with the rational design and engineering of biologically based parts, devices, or systems. However, biological systems are generally complex and unpredictable, and are therefore, intrinsically difficult to engineer. In order to address these fundamental challenges, synthetic biology is aiming to unify a “body of knowledge” from several foundational scientific fields, within the context of a set of engineering principles. This shift in perspective is enabling synthetic biologists to address complexity, such that robust biological systems can be designed, assembled, and tested as part of a biological design cycle. The design cycle takes a forward-design approach in which a biological system is specified, modeled, analyzed, assembled, and its functionality tested. At each stage of the design cycle, an expanding repertoire of tools is being developed. In this review, we highlight several of these tools in terms of their applications and benefits to the synthetic biology community.