High-yield and plasmid-free biocatalytic production of 5-methylpyrazine-2-carboxylic acid by combinatorial genetic elements engineering and genome engineering of Escherichia coli

Engineering Escherichia coli for high-yielding 2,5-Dimethylpyrazine synthesis from L-Threonine by reconstructing metabolic pathways and enhancing cofactors regeneration

2,5-Dimethylpyrazine (2,5-DMP) is important pharmaceutical raw material and food flavoring agent. Recently, engineering microbes to produce 2,5-DMP has become an attractive alternative to chemical synthesis approach. In this study, metabolic engineering strategies were used to optimize the modified Escherichia coli BL21 (DE3) strain for efficient synthesis of 2,5-DMP using L-threonine dehydrogenase (EcTDH) from Escherichia coli BL21, NADH oxidase (EhNOX) from Enterococcus hirae, aminoacetone oxidase (ScAAO) from Streptococcus cristatus and L-threonine transporter protein (EcSstT) from Escherichia coli BL21, respectively. We further optimized the reaction conditions for synthesizing 2,5-DMP. In optimized conditions, the modified strain can convert L-threonine to obtain 2,5-DMP with a yield of 2897.30 mg/L. Therefore, the strategies used in this study contribute to the development of high-level cell factories for 2,5-DMP.

Metabolic Engineering of Escherichia coli for Production of 2,5-Dimethylpyrazine

2,5-Dimethylpyrazine (2,5-DMP) is a high-value-added alkylpyrazine compound with important applications in both the food and pharmaceutical fields. In response to the increasing consumer preference for natural products over chemically synthesized ones, efforts have been made to develop efficient microbial cell factories for the production of 2,5-DMP. However, the previously reported recombinant strains have exhibited low yields and relied on expensive antibiotics and inducers. In this study, we employed metabolic engineering strategies to develop an Escherichia coli strain capable of producing 2,5-DMP at high levels without the need for inducers or antibiotics. Initially, the biosynthesis pathway of 2,5-DMP was constructed that realized 2,5-DMP production from glucose. Subsequently, efforts focused on enhancing 2,5-DMP production by improving the availability of the cofactor NAD+ and precursor l-threonine. Additionally, the supply and conversion of l-threonine were balanced by optimizing the copy number of the key gene tdh on the chromosome and by modifying the l-threonine transport system. The final engineering strain D19 produced 3.1 g/L of 2,5-DMP, which is the highest titer for fermentative production of 2,5-DMP using glucose as the carbon source up to date. The strategies used in this study lay a good foundation for the production of 2,5-DMP on a large scale.

Toward bioproduction of oxo chemicals from C1 feedstocks using isobutyraldehyde as an example

Oxo chemicals are valuable chemicals for synthesizing a wide array of industrial and consumer products. However, producing of oxo chemicals is predominately through the chemical process called hydroformylation, which requires petroleum-sourced materials and generates abundant greenhouse gas. Current concerns on global climate change have renewed the interest in reducing greenhouse gas emissions and recycling the plentiful greenhouse gas. A carbon–neutral manner in this regard is producing oxo chemicals biotechnologically using greenhouse gas as C1 feedstocks. Exemplifying isobutyraldehyde, this review demonstrates the significance of using greenhouse gas for oxo chemicals production. We highlight the current state and the potential of isobutyraldehyde synthesis with a special focus on the in vivo and in vitro scheme of C1-based biomanufacturing. Specifically, perspectives and scenarios toward carbon– and nitrogen–neutral isobutyraldehyde production are proposed. In addition, key challenges and promising approaches for enhancing isobutyraldehyde bioproduction are thoroughly discussed. This study will serve as a reference case in exploring the biotechnological potential and advancing oxo chemicals production derived from C1 feedstocks.

Constructing a methanol-dependent Bacillus subtilis by engineering the methanol metabolism

Methanol is a promising green feedstock for producing fuels and chemicals because it is inexpensive, clean, environmentally friendly, and easily prepared. Thus, many studies have been devoted to engineering non-native methylotrophic platform microorganisms to utilize methanol. This study adopted a series of strategies to develop a synthetic methylotrophic Bacillus subtilis that can use methanol as the carbon source, including the heterologous expression of methanol dehydrogenase (Mdh), enhancement of the expressions of 3-hexulose-6-phosphate synthase (Hps) and 6-phospho-3-hexuloisomerase (Phi), regulation of the expressions of key enzymes at both the translational and transcriptional levels, stabilization of the key enzyme expression through a dual-system for expressing the target genes on both the plasmid and genome, and improvement of the catalytic activity of Mdh with a recycling strategy for NAD⁺. As a result, the methanol consumption of the synthetic methylotrophic B. subtilis reached 4.09 g/L, with the maximum OD₆₀₀ showing a 2.21-fold increase compared with the wild-type B. subtilis, which cannot use methanol. We further deleted the phosphoglucose isomerase (Pgi) and added co-substrates to increase the supply of ribulose-5-phosphate (Ru-5-P), and the specific methanol consumption rate increased by an additional 27.54%. Finally, we successfully constructed two strains that cannot grow in M9 medium with xylose or ribose unless methanol is utilized. The strategies used in this study are generally applicable to other studies on synthetic methylotrophy.

High level production of diacetylchitobiose deacetylase by refactoring genetic elements and cellular metabolism

Diacetylchitobiose deacetylase (Dac) from Pyrococcus horikoshii can realize the one-step production of glucosamine (GlcN). The efficient expression and secretion of Dac play a central role in the green production of GlcN. In this study, Bacillus subtilis WB600 was used as the expression host. Firstly, we screened 12 signal peptides, among which signal peptide NprB had the strongest ability of guiding Dac secretion. Further optimization of the functional region showed that the extracellular Dac activity of NprB mutant was increased to 3682.2 U/mL. Next, the extracellular Dac activity was increased to 4807.6 U/mL by RBS sequence optimization. Then we got a new recombinant B. subtilis C6 for plasmid-free expression of Dac by integrating comK gene and silencing bpr, nprB, aprE, mpr and nprE genes. Finally, the extracellular Dac activity of genome-integrating strain reached 6357.38 U/mL, which was the highest level reported so far.

Intelligent host engineering for metabolic flux optimisation in biotechnology

Optimising the function of a protein of length N amino acids by directed evolution involves navigating a 'search space' of possible sequences of some 20N. Optimising the expression levels of P proteins that materially affect host performance, each of which might also take 20 (logarithmically spaced) values, implies a similar search space of 20P. In this combinatorial sense, then, the problems of directed protein evolution and of host engineering are broadly equivalent. In practice, however, they have different means for avoiding the inevitable difficulties of implementation. The spare capacity exhibited in metabolic networks implies that host engineering may admit substantial increases in flux to targets of interest. Thus, we rehearse the relevant issues for those wishing to understand and exploit those modern genome-wide host engineering tools and thinking that have been designed and developed to optimise fluxes towards desirable products in biotechnological processes, with a focus on microbial systems. The aim throughput is 'making such biology predictable'. Strategies have been aimed at both transcription and translation, especially for regulatory processes that can affect multiple targets. However, because there is a limit on how much protein a cell can produce, increasing kcat in selected targets may be a better strategy than increasing protein expression levels for optimal host engineering.

Semi-rational design of L-amino acid deaminase for production of pyruvate and D-alanine by Escherichia coli whole-cell biocatalyst

In our previous study, one-step pyruvate and D-alanine production from D,L-alanine by a whole-cell biocatalyst Escherichia coli expressing L-amino acid deaminase (Pm1) derived from Proteus mirabilis was investigated. However, due to the low catalytic efficiency of Pm1, the pyruvate titer was relatively low. Here, semi-rational design based on site-directed saturation mutagenesis was carried out to improve the catalytic efficiency of Pm1. A novel high-throughput screening (HTS) method for pyruvate based on 2,4-dinitrophenylhydrazine indicator was then established. The catalytic efficiency (k_cat/K_m) of the mutant V437I screened out by this method was 1.88 times higher than wild type. Next, to improve the growth of the engineered strain BLK07, the genes encoding for Xpk and Fbp were integrated into its genome to construct non-oxidative glycolysis (NOG) pathway. Finally, the CRISPR/Cas9 system was used to integrate the N6-pm1-V437I gene into the genome of BLK07. Pyruvic acid titer of the plasmid-free strain reached 42.20 g/L with an L-alanine conversion rate of 77.62% and a D-alanine resolution of 82.4%. This work would accelerate the industrial production of pyruvate and D-alanine by biocatalysis, and the HTS method established here could be used to screen other Pm1 mutants with high pyruvate titers.

Microbial production of chemicals driven by CRISPR-Cas systems

Microorganisms have provided an attractive route for biosynthesis of various chemicals from renewable resources. CRISPR-Cas systems have served as powerful mechanisms for generating cell factories with desirable properties by manipulating nucleic acids quickly and efficiently. The CRISPR-Cas system provides a toolbox with excellent opportunities for identifying better biocatalysts, multiplexed fine-tuning of metabolic flux, efficient utilization of low-cost substrates, and improvement of metabolic robustness. The overall goal of this review highlights recent advances in the development of microbial cell factories for chemical production using various CRISPR-Cas systems. The perspectives for further development or applications of CRISPR-Cas systems for strain improvement are also discussed.

Biotransformation and chiral resolution of d,l-alanine into pyruvate and d-alanine with a whole-cell biocatalyst expressing l-amino acid deaminase

Pyruvate is an important pharmaceutical intermediate and is widely used in food, nutraceuticals, and pharmaceuticals. However, high environmental pollution caused by chemical synthesis or complex separation process of microbial fermentation methods constrain the supply of pyruvate. Here, one-step pyruvate and d-alanine production from d,l-alanine by whole-cell biocatalysis was investigated. First, l-amino acid deaminase (Pm1) from Proteus mirabilis was expressed in Escherichia coli, resulting in pyruvate titer of 12.01 g/L. Then, N-terminal coding sequences were introduced to the 5'-end of the pm1 gene to enhance the expression of Pm1 and the pyruvate titer increased to 15.13 g/L. Next, product utilization by the biocatalyst was prevented by knocking out the pyruvate uptake transporters (cstA, btsT) and the pyruvate metabolic pathway genes pps, poxB, pflB, ldhA, and aceEF using CRISPR/Cas9, yielding 30.88 g/L pyruvate titer. Finally, by optimizing the reaction conditions, the pyruvate titer was further enhanced to 43.50 g/L in 8 H with a 79.99% l-alanine conversion rate; meanwhile, the resolution of d-alanine reached 84.0%. This work developed a whole-cell biocatalyst E. coli strain for high-yield, high-efficiency, and low-pollution pyruvate and d-alanine production, which has great potential for the commercial application in the future.