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Background Deactivated Cas9 (dCas9) led to significant improvement of CRISPR/Cas9-based techniques because it can be fused with a variety of functional groups to form diverse molecular devices, which can manipulate or modify target DNA cassettes. One important metabolic engineering strategy is to localize the enzymes in proximity of their substrates for improved catalytic efficiency. In this work, we developed a novel molecular device to manipulate the cellular location of specific DNA cassettes either on plasmids or on the chromosome, by fusing location tags to dCas9 (Cas9-Lag), and applied the technique for synthetic biology applications. Carotenoids like β-carotene serve as common intermediates for the synthesis of derivative compounds, which are hydrophobic and usually accumulate in the membrane compartment. Results Carotenoids like β-carotene serve as common intermediates for the synthesis of derivative compounds, which are hydrophobic and usually accumulate in the membrane components. To improve the functional expression of membrane-bound enzymes and localize them in proximity to the substrates, Cas9-Lag was used to pull plasmids or chromosomal DNA expressing carotenoid enzymes onto the cell membrane. For this purpose, dCas9 was fused to the E. coli membrane docking tag GlpF, and gRNA was designed to direct this fusion protein to the DNA expression cassettes. With Cas9-Lag, the zeaxanthin and astaxanthin titer increased by 29.0% and 26.7% respectively. Due to experimental limitations, the electron microscopy images of cells expressing Cas9-Lag vaguely indicated that GlpF-Cas9 might have pulled the target DNA cassettes in close proximity to membrane. Similarly, protein mass spectrometry analysis of membrane proteins suggested an increased expression of carotenoid-converting enzymes in the membrane components. Conclusion This work therefore provides a novel molecular device, Cas9-Lag, which was proved to increase zeaxanthin and astaxanthin production and might be used to manipulate DNA cassette location.

Biomaterials offer unique properties that make them irreplaceable for next-generation applications. Fibrous proteins, such as various caterpillar silks and especially spider silk, have strength and toughness not found in human-made materials. In early studies, proteins containing long tandem repeats, such as major ampullate spidroin 1 (MaSp1) and flagelliform silk protein (FSLP), were produced using a large DNA template composed of many tandem repeats. The hierarchical DNA assembly of the DNA template is very time-consuming and labor-intensive, which makes the fibrous proteins difficult to study and engineer. In this study, we designed a circularized mRNA (cmRNA) employing the RNA cyclase ribozyme mechanism. cmRNAs encoding spider silk protein MaSp1 and FSLP were designed based on only one unit of the template sequence but provide ribosomes with a circular and infinite translation template for production of long peptides containing tandem repeats. Using this technique, cmRNAs of MaSp1 and FSLP were successfully generated with circularization efficiencies of 8.5% and 36.7%, respectively, which supported the production of recombinant MaSp1 and FSLP larger than 110 and 88 kDa, containing tens of repeat units. Western blot analysis and mass spectrometry confirmed the authenticity of MaSp1 and FSLP, which were produced at titers of 22.1 and 81.5 mg · liter(−1), respectively. IMPORTANCE Spider silk is a biomaterial with superior properties. However, its heterologous expression template is hard to construct. The cmRNA technique simplifies the construction and expression strategy by proving the ribosome a circular translation template for expression of long peptides containing tandem repeats. This revolutionary technique will allow researchers to easily build, study, and experiment with any fiber proteins with sequences either from natural genes or artificial designs. We expect a significantly accelerated development of fibrous protein-based biomaterials with the cmRNA technique.

Background Natural life systems can be significantly modified at the genomic scale by human intervention, demonstrating the great innovation capacity of genome engineering. Large epi-chromosomal DNA structures were established in Escherichia coli cells, but some of these methods were inconvenient, using heterologous systems, or relied on engineered E. coli strains. Results The wild-type model bacterium E. coli has a single circular chromosome. In this work, a novel method was developed to split the original chromosome of wild-type E. coli. With this method, novel E. coli strains containing two chromosomes of 0.10 Mb and 4.54 Mb, and 2.28 Mb and 2.36 Mb were created respectively, designated as E. coli0.10/4.54 and E. coli2.28/2.36. The new chromosomal arrangement was proved by PCR amplification of joint regions as well as a combination of Nanopore and Illumina sequencing analysis. While E. coli0.10/4.54 was quite stable, the two chromosomes of E. coli2.28/2.36 population recombined into a new chromosome (Chr.4.64MMut), via recombination. Both engineered strains grew slightly slower than the wild-type, and their cell shapes were obviously elongated. Conclusion Finally, we successfully developed a simple CRISPR-based genome engineering technique for the construction of multi-chromosomal E. coli strains with no heterologous genetic parts. This technique might be applied to other prokaryotes for synthetic biology studies and applications in the future.

The 14α-hydroxysteroids have specific anti-gonadotropic and carcinolytic biological activities and can be produced by microbial biotransformation. The steroid 11β-/14α-hydroxylase P-450lun from Cochliobolus lunatus is the only fungal cytochrome P450 enzyme identified to date with steroid C14 hydroxylation ability. Previous work has mainly revealed the 11β-hydroxylation activity of the P-450lun towards cortexolone (RSS) substrate; however, the potential steroid 14α-hydroxylation activity of this enzyme, especially for androstenedione (AD) substrate, has not yet conducted in-depth testing. In this work, we further tested the steroid 14α-hydroxylation activity of the P-450lun towards RSS and AD in the Saccharomyces cerevisiae system. We demonstrated that P-450lun functions as the specific 14α-hydroxylase towards the AD substrate (regiospecificity > 99%); however, it showed a poor C14-hydroxylation regiospecificity (around 40%) for the RSS substrate. In addition, through transcriptome analysis combined with gene functional characterizations, we also identified and cloned the gene for the P-450lun-associated redox partner CPRlun. Finally, through codon optimization, knockout of genes for the side reactions related enzymes GCY1 and YPR1, and increasing copies of the P-450lun and CPRlun, we developed a recombinant S. cerevisiae biocatalyst based on the C. lunatus steroid 14α-hydroxylation system to produce 14α-hydroxysteroids. Initial production of 14α-OH-AD (150 mg/L day productivity, 99% regioisomeric purity, and 60% w/w yield) and 14α-OH-RSS (64 mg/L day productivity, 40% regioisomeric purity, and 26% w/w yield) were separately achieved in shake flasks; these results represent the highest level of 14α-hydroxysteroid production in the current yeast system.

In this study, the FAD synthesis pathway was manipulated to increase its cellular concentration and thereby improve the electroactivity of E. coli. In a microbial electrosynthesis (MES) system with neutral red as electron carrier and fumarate as the sole electron acceptor, the engineered strains derived from three E. coli lines displayed increased electric current in the reaction system, indicating improved electroactivity. Furthermore, the production of succinate from fumarate increased by around 60% compared with that of the parent strains, confirming the improvement of E. coli electroactivity by manipulating the FAD synthesis pathway. An MES reaction was performed with engineered E. coli 8739, and an altered metabolic profile with more reductive fermentation products was obtained. When the electroactive succinate-producing strain E. coli T110 was used in the MES, a yield of 0.97 ± 0.02 mol/mol glucose was achieved, which corresponds to an approximately 1.4-fold increase compared with the fermentation with no electricity supply or non-electroactive T110. In addition, a carbon concentration mechanism (CCM) was employed to further improve succinate production and yield in the MES, which produced a succinate yield of 1.16 mol/mol glucose, a 1.7-fold increase compared with that of the parent strain T110, indicating that the electroactive E. coli could be used in MES to produce specific fermentation products with improved yield.

Massive emission of CO2 into atmosphere from consumption of carbon deposit is causing climate change. Researchers have applied metabolic engineering and synthetic biology techniques for improving CO2 fixation efficiency in many species. One solution might be the utilization of autotrophic bacteria, which have great potential to be engineered into microbial cell factories for CO2 fixation and the production of chemicals, independent of fossil resources. In this work, several pathways of Ralstonia eutropha H16 were modulated by manipulation of heterologous and endogenous genes related to fatty acid synthesis. The resulting strain B2(pCT, pFP) was able to produce 124.48 mg/g (cell dry weight) free fatty acids with fructose as carbon source, a fourfold increase over the parent strain H16. To develop a truly autotrophic fermentation technique with H2, CO2 and O2 as substrates, we assembled a relatively safe, continuous, lab-scale gas fermentation system using micro-fermentation tanks, H2 supplied by a hydrogen generator, and keeping the H2 to O2 ratio at 7:1. The system was equipped with a H2 gas alarm, rid of heat sources and placed into a fume hood to further improve the safety. With this system, the best strain B2(pCT, pFP) produced 60.64 mg free fatty acids per g biomass within 48 h, growing in minimal medium supplemented with 9 × 103 mL/L/h hydrogen gas. Thus, an autotrophic fermentation technique to produce fatty acids was successfully established, which might inspire further research on autotrophic gas fermentation with a safe, lab-scale setup, and provides an alternative solution for environmental and energy problems.

Glycolate is a bulk chemical with wide applications in the textile, food processing, and pharmaceutical industries. Glycolate can be produced from glucose via the glycolysis and glyoxylate shunt pathways, followed by reduction to glycolate. However, two problems limit the productivity and yield of glycolate when using glucose as the sole carbon source. The first is a cofactor imbalance in the production of glycolate from glucose via the glycolysis pathway, since NADPH is required for glycolate production, while glycolysis generates NADH. To rectify this imbalance, the NADP+ -dependent glyceraldehyde 3-phosphate dehydrogenase GapC from Clostridium acetobutylicum was introduced to generate NADPH instead of NADH in the oxidation of glyceraldehyde 3-phosphate during glycolysis. The soluble transhydrogenase SthA was further eliminated to conserve NADPH by blocking its conversion into NADH. The second problem is an unfavorable carbon flux distribution between the tricarboxylic acid cycle and the glyoxylate shunt. To solve this problem, isocitrate dehydrogenase (ICDH) was eliminated to increase the carbon flux of glyoxylate and thereby improve the glycolate titer. After engineering through the integration of gapC, combined with the inactivation of ICDH, SthA, and by-product pathways, as well as the upregulation of the two key enzymes isocitrate lyase (encoding by aceA), and glyoxylate reductase (encoding by ycdW), the glycolate titer increased to 5.3 g/L with a yield of 1.89 mol/mol glucose. Moreover, an optimized fed-batch fermentation reached a titer of 41 g/L with a yield of 1.87 mol/mol glucose after 60 h.

CRISPR-enabled deaminase base editing has become a powerful tool for precisely editing nucleotides on the chromosome. In this study DNA helicases, such as Escherichia coli DnaB, were fused to activation-induced cytidine deaminase (AID) to form enzyme complexes which randomly introduces edited bases throughout the chromosome. DnaB-AID was found to increase 2.5 × 103 fold relative to the mutagenesis frequency of wildtype. 97.9% of these edits were observed on the leading strand during DNA replication suggesting deamination to be highly coordinated with DNA replication. Using DnaB-AID, a 371.4% increase in β-carotene production was obtained following four rounds of editing. In Saccharomyces cerevisiae Helicase-AID was constructed by fusing AID to one of the subunits of eukaryotic helicase Mcm2-7 complex, MCM5. Using MCM5-AID, the average editing efficiency of five strains was 2.1 ± 0.4 × 103 fold higher than the native genomic mutation rate. MCM5-AID was able to improve β-carotene production of S. cerevisiae 4742crt by 75.4% following eight rounds of editing. The S. cerevisiae MCM5-AID technique is the first biological tool for generating and accumulating single base mutations in eukaryotic chromosomes. Since the helicase complex is highly conservative in all eukaryotes, Helicase-AID could be adapted for various applications and research in all eukaryotic cells.

Background CO2 is fixed by all living organisms with an autotrophic metabolism, among which the Calvin–Benson–Bassham (CBB) cycle is the most important and widespread carbon fixation pathway. Thus, studying and engineering the CBB cycle with the associated energy providing pathways to increase the CO2 fixation efficiency of cells is an important subject of biological research with significant application potential. Results In this work, the autotrophic microbe Ralstonia eutropha (Cupriavidus necator) was selected as a research platform for CBB cycle optimization engineering. By knocking out either CBB operon genes on the operon or mega-plasmid of R. eutropha, we found that both CBB operons were active and contributed almost equally to the carbon fixation process. With similar knock-out experiments, we found both soluble and membrane-bound hydrogenases (SH and MBH), belonging to the energy providing hydrogenase module, were functional during autotrophic growth of R. eutropha. SH played a more significant role. By introducing a heterologous cyanobacterial RuBisCO with the endogenous GroES/EL chaperone system(A quality control systems for proteins consisting of molecular chaperones and proteases, which prevent protein aggregation by either refolding or degrading misfolded proteins) and RbcX(A chaperone in the folding of Rubisco), the culture OD600 of engineered strain increased 89.2% after 72 h of autotrophic growth, although the difference was decreased at 96 h, indicating cyanobacterial RuBisCO with a higher activity was functional in R. eutropha and lead to improved growth in comparison to the host specific enzyme. Meanwhile, expression of hydrogenases was optimized by modulating the expression of MBH and SH, which could further increase the R. eutropha H16 culture OD600 to 93.4% at 72 h. Moreover, the autotrophic yield of its major industrially relevant product, polyhydroxybutyrate (PHB), was increased by 99.7%. Conclusions To our best knowledge, this is the first report of successfully engineering the CBB pathway and hydrogenases of R. eutropha for improved activity, and is one of only a few cases where the efficiency of CO2 assimilation pathway was improved. Our work demonstrates that R. eutropha is a useful platform for studying and engineering the CBB for applications.

Current base editors (BEs) catalyze only base transitions (C to T and A to G) and cannot produce base transversions. Here we present BEs that cause C-to-A transversions in Escherichia coli and C-to-G transversions in mammalian cells. These glycosylase base editors (GBEs) consist of a Cas9 nickase, a cytidine deaminase and a uracil-DNA glycosylase (Ung). Ung excises the U base created by the deaminase, forming an apurinic/apyrimidinic (AP) site that initiates the DNA repair process. In E. coli, we used activation-induced cytidine deaminase (AID) to construct AID-nCas9-Ung and found that it converts C to A with an average editing specificity of 93.8% ± 4.8% and editing efficiency of 87.2% ± 6.9%. For use in mammalian cells, we replaced AID with rat APOBEC1 (APOBEC-nCas9-Ung). We tested APOBEC-nCas9-Ung at 30 endogenous sites, and we observed C-to-G conversions with a high editing specificity at the sixth position of the protospacer between 29.7% and 92.2% and an editing efficiency between 5.3% and 53.0%. APOBEC-nCas9-Ung supplements the current adenine and cytidine BEs (ABE and CBE, respectively) and could be used to target G/C disease-causing mutations. New base editors change C to A in bacteria and C to G in mammalian cells.

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.

Background Electrochemical energy is a key factor of biosynthesis, and is necessary for the reduction or assimilation of substrates such as CO2. Previous microbial electrosynthesis (MES) research mainly utilized naturally electroactive microbes to generate non-specific products. Results In this research, an electroactive succinate-producing cell factory was engineered in E. coli T110(pMtrABC, pFccA-CymA) by expressing mtrABC, fccA and cymA from Shewanella oneidensis MR-1, which can utilize electricity to reduce fumarate. The electroactive T110 strain was further improved by incorporating a carbon concentration mechanism (CCM). This strain was fermented in an MES system with neutral red as the electron carrier and supplemented with HCO3+, which produced a succinate yield of 1.10 mol/mol glucose—a 1.6-fold improvement over the parent strain T110. Conclusions The strain T110(pMtrABC, pFccA-CymA, pBTCA) is to our best knowledge the first electroactive microbial cell factory engineered to directly utilize electricity for the production of a specific product. Due to the versatility of the E. coli platform, this pioneering research opens the possibility of engineering various other cell factories to utilize electricity for bioproduction. Electronic supplementary material The online version of this article (10.1186/s12934-019-1067-3) contains supplementary material, which is available to authorized users.

Carotenoids are a class of terpenes of commercial interest that exert important biological functions. While various strategies have been applied to engineer β-carotene production in microbial cell factories, no work has been done to study and improve the storage of hydrophobic terpene products inside the heterologous host cells. Although the membrane is thought to be the cell compartment that accumulates hydrophobic terpenes such as β-carotene, direct evidence is still lacking. In this work, we engineered the membrane of Escherichia coli in both its morphological and biosynthetic aspects, as a means to study and improve its storage capacity for β-carotene. Engineering the membrane morphology by overexpressing membrane-bending proteins resulted in a 28% increase of β-carotene specific producton value, while engineering the membrane synthesis pathway led to a 43% increase. Moreover, the combination of these two strategies had a synergistic effect, which caused a 2.9-fold increase of β-carotene specific production value (from 6.7 to 19.6mg/g DCW). Inward membrane stacks were observed in electron microscopy images of the engineered E. coli cells, which indicated that morphological changes were associated with the increased β-carotene storage capacity. Finally, membrane separation and analysis confirmed that the increased β-carotene was mainly accumulated within the cell membrane. This membrane engineering strategy was also applied to the β-carotene hyperproducing strain CAR025, which led to a 39% increase of the already high β-carotene specific production value (from 31.8 to 44.2mg/g DCW in shake flasks), resulting in one of the highest reported specific production values under comparable culture conditions. The membrane engineering strategy developed in this work opens up a new direction for engineering and improving microbial terpene producers. It is quite possible that a wide range of strains used to produce hydrophobic compounds can be further improved using this novel engineering strategy.

One of the most important research subjects of metabolic engineering is the pursuit of balanced metabolic pathways, which requires the modulation of expression of many genes. However, simultaneously modulating multiple genes on the chromosome remains challenging in prokaryotic organisms, including the industrial workhorse - Escherichia coli. In this work, the CRISPR/Cas9-facilitated multiplex pathway optimization (CFPO) technique was developed to simultaneously modulate the expression of multiple genes on the chromosome. To implement it, two plasmids were employed to target Cas9 to regulatory sequences of pathway genes, and a donor DNA plasmid library was constructed containing a regulator pool to modulate the expression of these genes. A modularized plasmid construction strategy was used to enable the assembly of a complex donor DNA plasmid library. After genome editing using this technique, a combinatorial library was obtained with variably expressed pathway genes. As a demonstration, the CFPO technique was applied to the xylose metabolic pathway genes in E. coli to improve xylose utilization. Three transcriptional units containing a total of four genes were modulated simultaneously with 70% efficiency, and improved strains were selected from the resulting combinatorial library by growth enrichment. The best strain, HQ304, displayed a 3-fold increase of the xylose-utilization rate. Finally, the xylose-utilization pathway of HQ304 was analyzed enzymologically to determine the optimal combination of enzyme activities.

Glycolate is a bulk chemical which has been widely used in textile, food processing, and pharmaceutical industries. Glycolate can be produced from sugars by microbial fermentation. However, when using glucose as the sole carbon source, the theoretical maximum carbon molar yield of glycolate is 0.67 mol/mol due to the loss of carbon as CO2. In this study, a synergetic system for simultaneous utilization of acetate and glucose was designed to increase the carbon yield. The main function of glucose is to provide NADPH while acetate to provide the main carbon backbone for glycolate production. Theoretically, 1 glucose and 5 acetate can produce 6 glycolate, and the carbon molar yield can be increased to 0.75 mol/mol. The whole synthetic pathway was divided into two modules, one for converting acetate to glycolate and another to utilize glucose to provide NADPH. After engineering module I through activation of acs, gltA, aceA and ycdW, glycolate titer increased from 0.07 to 2.16 g/L while glycolate yields increased from 0.04 to 0.35 mol/mol-acetate and from 0.03 to 1.04 mol/mol-glucose. Module II was then engineered to increase NADPH supply. Through deletion of pfkA, pfkB, ptsI and sthA genes as well as upregulating zwf, pgl and tktA, glycolate titer increased from 2.16 to 4.86 g/L while glycolate yields increased from 0.35 to 0.82 mol/mol-acetate and from 1.04 to 6.03 mol/mol-glucose. The activities of AceA and YcdW were further increased to pull the carbon flux to glycolate, which increased glycolate yield from 0.82 to 0.92 mol/mol-acetate. Fed-batch fermentation of the final strain NZ-Gly303 produced 73.3 g/L glycolate with a productivity of 1.04 g/(L·h). The acetate to glycolate yield was 0.85 mol/mol (1.08 g/g), while glucose to glycolate yield was 6.1 mol/mol (2.58 g/g). The total carbon molar yield was 0.60 mol/mol, which reached 80% of the theoretical value.

Escherichia coli BL21 is arguably the most popular host for industrial production of proteins, and industrial fermentations are often plagued by phage infections. The CRISPR/Cas system is guided by a gRNA to cleave a specific DNA cassette, which can be developed into a highly efficient programable phage defense system. In this work, we constructed a CRISPR/Cas system targeting multiple positions on the genome of T7 phage and found that the system increased the BL21’s defense ability against phage infection. Furthermore, the targeted loci on phage genome played a critical role. For better control of expression of CRISPR/Cas9, various modes were tested, and the OD of the optimized strain BL21(pT7cas9, pT7-3gRNA, prfp) after 4 hours of phage infection was significantly improved, reaching 2.0, which was similar to the control culture without phage infection. Although at later time points, the defensive ability of CRISPR/Cas9 systems were not as obvious as that at early time points. The viable cell count of the engineered strain in the presence of phage was only one order of magnitude lower than that of the strain with no infection, which further demonstrated the effectiveness of the CRISPR/Cas9 phage defense system. Finally, the engineered BL21 strain under phage attack expressed RFP protein at about 60% of the un-infected control, which was significantly higher than the parent BL21. In this work, we successfully constructed a programable CRISPR/Cas9 system to increase the ability of E. coli BL21’s to defend against phage infection, and created a resistant protein expression host. This work provides a simple and feasible strategy for protecting industrial E. coli strains against phage infection.

Unlike eukaryotes, bacteria are less proficient in homologous recombination (HR) and non-homologous end joining (NHEJ). All existing genomic editing methods for Escherichia coli rely on exogenous HR or NHEJ systems to repair DNA double-strand breaks (DSBs). Although an E. coli native end-joining (ENEJ) system has been reported, its potential in chromosomal engineering has not yet been explored. Here, we present a CRISPR-Cas9-assisted native end-joining editing and show that ENEJ-dependent DNA repair can be used to conduct rapid and efficient knocking-out of E. coli genomic sequence of up to 83 kb. Moreover, the positive rate and editing efficiency is independent of high-efficiency competent cells. The method requires neither exogenous DNA repair systems nor introduced editing template. The Cas9 complex is the only foreign element in this method. This study is the first successful engineering effort to utilize ENEJ mechanism in genomic editing and provides an effective strategy for genetic engineering in bacteria that are inefficient in HR and NHEJ.SignificanceThe application in prokaryotes is difficult because of the weak homologous recombination and non-homologous end joining systems. E. coli, as the most-used prokaryote in metabolic engineering, has no NHEJ system. All existing genomic editing methods for E. coli rely on exogenous HR or NHEJ systems to repair double-strand breaks introduced by CRISPR/Cas9. In this report, we firstly demonstrate that the weak and previously ignored end-joining mechanism in E. coli can be used for efficient large-scale genetic engineering assisted by CRISPR/Cas9. Our efforts greatly simplify the genomic editing procedure of E. coli and provide an effective strategy for genetic engineering in bacteria that are inefficient in HR and NHEJ.

H2 is extensively available from geological reservoirs, industrial byproducts or natural processes, which makes H2 an ideal supplemental substrate providing additional reductive equivalents for increasing anaerobic fermentation efficiency. While it has been applied broadly in industrial and pharmaceutical fileds, Escherichia coli was not shown to efficiently utilize hydrogen. To construct a versatile biological platform for efficient utilization of H2, the endogenous hydrogenase genes of E. coli were investigated. Consequently, overexpression of the endogenous hydrogenases Hyd-1 and Hyd-2 induced respective 26.5 % and 36.7 % increases of succinate yield, indicating that both Hyd-1 and Hyd-2 contributed to the increased pool of reductive equivalents. Additionally, simultaneously modulating the expression of hya and hyb genes by introducing the strong promoter P10 resulted in the higher yield of succinate. To further improve the utilization of H2, pathway of rTCA cycle and CO2 supplementation were engineered in the strain ZL014(pBTCA), which produced the highest succinate yield of 1.50 mol per mol glucose. Our work successfully engineered a novel E. coli platform that could efficiently utilize H2, which could be employed to construct a variety of microbial cell factories requiring substantial amounts of reductive equivalents.

Background Crocin is a carotenoid-derived natural product found in the stigma of Crocus spp., which has great potential in medicine, food and cosmetics. In recent years, microbial production of crocin has drawn increasing attention, but there were no reports of successful implementation. Escherichia coli has been engineered to produce various carotenoids, including lycopene, β-carotene and astaxanthin. Therefore, we intended to construct E. coli cell factories for crocin biosynthesis. Results In this study, a heterologous crocetin and crocin synthesis pathway was first constructed in E. coli. Firstly, the three different zeaxanthin-cleaving dioxygenases CsZCD, CsCCD2 from Crocus sativus, and CaCCD2 from Crocus ancyrensis, as well as the glycosyltransferases UGT94E5 and UGT75L6 from Gardenia jasminoides, were introduced into zeaxanthin-producing E. coli cells. The results showed that CsCCD2 catalyzed the synthesis of crocetin dialdehyde. Next, the aldehyde dehydrogenases ALD3, ALD6 and ALD9 from Crocus sativus and ALD8 from Neurospora crassa were tested for crocetin dialdehyde oxidation, and we were able to produce 4.42 mg/L crocetin using strain YL4(pCsCCD2-UGT94E5-UGT75L6,pTrc-ALD8). Glycosyltransferases from diverse sources were screened by in vitro enzyme activity assays. The results showed that crocin and its various derivatives could be obtained using the glycosyltransferases YjiC, YdhE and YojK from Bacillus subtilis, and the corresponding genes were introduced into the previously constructed crocetin-producing strain. Finally, crocin-5 was detected among the fermentation products of strain YL4(pCsCCD2-UGT94E5-UGT75L6,pTrc-ALD8,pET28a-YjiC-YdhE-YojK) using HPLC and LC–ESI–MS. Conclusions A heterologous crocin synthesis pathway was constructed in vitro, using glycosyltransferases from the Bacillus subtilis instead of the original plant glycosyltransferases, and a crocetin and crocin-5 producing E. coli cell factory was obtained. This research provides a foundation for the large-scale production of crocetin and crocin in E. coli cell factories. Electronic supplementary material The online version of this article (10.1186/s12934-019-1166-1) contains supplementary material, which is available to authorized users.

The goal of this work was to improve the lycopene storage capacity of the E. coli membrane by engineering both morphological and biosynthetic aspects. First, Almgs, a protein from Acholeplasma laidlawii that is involved in membrane bending is overexpressed to expand the storage space for lycopene, which resulted in a 12% increase of specific lycopene production. Second, several genes related to the membrane-synthesis pathway in E. coli, including plsb, plsc, and dgka, were also overexpressed, which led to a further 13% increase. In addition, membrane separation and component analysis confirmed that the increased amount of lycopene was mainly accumulated within the cell membranes. Finally, by integrating both aforementioned modification strategies, a synergistic effect could be observed which caused a 1.32-fold increase of specific lycopene production, from the 27.5 mg/g of the parent to 36.4 mg/g DCW in the engineered strain. This work demonstrates that membrane engineering is a feasible strategy for increasing the production and accumulation of lycopene in E. coli.

Background Geranyl acetate is widely used in the fragrance and cosmetic industries, and thus has great economic value. However, plants naturally produce a mixture of hundreds of esters, and geranyl acetate is usually only present in trace amounts, which makes its economical extraction from plant sources practically impossible. As an ideal host for heterologous production of fragrance compound, the Saccharomyces cerevisiae has never been engineered to produce the esters, such as geranyl acetate. Results In this study, a heterologous geranyl acetate synthesis pathway was constructed in S. cerevisiae for the first time, and a titer of 0.63 mg/L geranyl acetate was achieved. By expressing an Erg20 mutant to divert carbon flux from FPP to GPP, the geranyl acetate production increased to 2.64 mg/L. However, the expression of heterologous GPP had limited effect. The highest production of 13.27 mg/L geranyl acetate was achieved by additional integration and expression of tHMG1, IDI1 and MAF1. Furthermore, through optimizing fermentation conditions, the geranyl acetate titer increased to 22.49 mg/L. Conclusions We constructed a monoterpene ester producing cell factory in S. cerevisiae for the first time, and demonstrated the great potential of this system for the heterologous production of a large group of economically important fragrance compounds.

A major challenge in producing chemicals and biofuels is to increase the tolerance of the host organism to toxic products or byproducts. An Escherichia coli strain with superior ethanol and more generally alcohol tolerance was identified by screening a library constructed by randomly integrating Lactobacillus plantarum genomic DNA fragments into the E. coli chromosome via Cre-lox recombination. Sequencing identified the inserted DNA fragment as the murA2 gene and its upstream intergenic 973-bp sequence, both coded on the negative genomic DNA strand. Overexpression of this murA2 gene and its upstream 973-bp sequence significantly enhanced ethanol tolerance in both E. coli EC100 and wild type E. coli MG1655 strains by 4.1-fold and 2.0-fold compared to control strains, respectively. Tolerance to n-butanol and i-butanol in E. coli MG1655 was increased by 1.85-fold and 1.91-fold, respectively. We show that the intergenic 973-bp sequence contains a native promoter for the murA2 gene along with a long 5' UTR (286 nt) on the negative strand, while a noncoding, small RNA, named MurA2S, is expressed off the positive strand. MurA2S is expressed in E. coli and may interact with murA2, but it does not affect murA2's ability to enhance alcohol tolerance in E. coli. Overexpression of murA2 with its upstream region in the ethanologenic E. coli KO11 strain significantly improved ethanol production in cultures that simulate the industrial Melle-Boinot fermentation process.

Massive emission of CO2 into atmosphere from consumption of carbon deposit is causing climate change. Researchers have applied metabolic engineering and synthetic biology techniques for improving CO2 fixation efficiency in many species. One solution might be the utilization of autotrophic bacteria, which have great potential to be engineered into microbial cell factories for CO2 fixation and the production of chemicals, independent of fossil resources. In this work, several pathways of Ralstonia eutropha H16 were modulated by manipulation of heterologous and endogenous genes related to fatty acid synthesis. The resulting strain B2(pCT, pFP) was able to produce 124.48 mg/g (cell dry weight) free fatty acids with fructose as carbon source, a fourfold increase over the parent strain H16. To develop a truly autotrophic fermentation technique with H2, CO2 and O2 as substrates, we assembled a relatively safe, continuous, lab-scale gas fermentation system using micro-fermentation tanks, H2 supplied by a hydrogen generator, and keeping the H2 to O2 ratio at 7:1. The system was equipped with a H2 gas alarm, rid of heat sources and placed into a fume hood to further improve the safety. With this system, the best strain B2(pCT, pFP) produced 60.64 mg free fatty acids per g biomass within 48 h, growing in minimal medium supplemented with 9 × 103 mL/L/h hydrogen gas. Thus, an autotrophic fermentation technique to produce fatty acids was successfully established, which might inspire further research on autotrophic gas fermentation with a safe, lab-scale setup, and provides an alternative solution for environmental and energy problems.

Ralstonia eutropha is a facultatively chemolithoautotrophic bacterium able to grow with organic substrates or H 2 and CO 2 under aerobic conditions. Under conditions of nutrient imbalance, R. eutropha produces copious amounts of poly[( R )-3-hydroxybutyrate] (PHB). Its ability to utilize CO 2 as a sole carbon source renders it an interesting new candidate host for the production of renewable liquid transportation fuels. We engineered R. eutropha for the production of fatty acid-derived, diesel-range methyl ketones. Modifications engineered in R. eutropha included overexpression of a cytoplasmic version of the TesA thioesterase, which led to a substantial (>150-fold) increase in fatty acid titer under certain conditions. In addition, deletion of two putative β-oxidation operons and heterologous expression of three genes (the acyl coenzyme A oxidase gene from Micrococcus luteus and fadB and fadM from Escherichia coli ) led to the production of 50 to 65 mg/liter of diesel-range methyl ketones under heterotrophic growth conditions and 50 to 180 mg/liter under chemolithoautotrophic growth conditions (with CO 2 and H 2 as the sole carbon source and electron donor, respectively). Induction of the methyl ketone pathway diverted substantial carbon flux away from PHB biosynthesis and appeared to enhance carbon flux through the pathway for biosynthesis of fatty acids, which are the precursors of methyl ketones.

Background One of the most important research subjects of metabolic engineering is pursuing a balanced metabolic pathway, which is the basis of an efficient cell factory. In this work, we dedicated to develop a simple and efficient technique to modulate expression of multiple genes simultaneously, and select for the optimal regulation pattern. Results A Type IIs restriction based combinatory modulation (TRCM) technique was designed and established in the research. With this technique, a plasmid library containing variably regulated mvaE, mvaS, mvaK 1, mvaD and mvaK 2 of the mevalonate (MVA) pathway were obtained and transformed into E. coli DXS37-IDI46 to obtain a β-carotene producer library. The ratio of successfully assembled plasmids was determined to be 35%, which was increased to 100% when color based pre-screening was applied. Representative strains were sequenced to contain diverse RBSs as designed to regulate expression of MVA pathway genes. A relatively balanced MVA pathway was achieved in E. coli cell factory to increase the β-carotene yield by two fold. Furthermore, the approximate regulation pattern of this optimal MVA pathway was illustrated. Conclusions A TRCM technique for metabolic pathway optimization was designed and established in this research, which can be applied to various applications in terms of metabolic pathway regulation and optimization. Electronic supplementary material The online version of this article (doi:10.1186/s12934-017-0659-z) contains supplementary material, which is available to authorized users.

Background Corynebacterium glutamicum was used as a metabolic engineering chassis for production of crude violacein (mixture of violacein and deoxyviolacein) due to Corynebacterium’s GRAS status and advantages in tryptophan fermentation. The violacein is a commercially potential compound with various applications derived from l-tryptophan. Results Corynebacterium glutamicum ATCC 21850 that could produce 162.98 mg L−1 tryptophan was employed as a novel host for metabolic engineering chassis. Heterologous vio operon from Chromobacterium violaceum was over-expressed in ATCC 21850 strain with constitutive promoter to have obtained 532 mg L−1 violacein. Considering toxicity of violacein, vio operon was expressed with inducible promoter and 629 mg L−1 violacein was obtained in batch culture. Due to the economical coding nature of vio operon, the compressed RBS of vio genes were replaced with complete strong C. glutamicum ones. And extended expression units were assembled to form a synthetic operon. With this strategy, 1116 mg L−1 violacein in batch culture was achieved. Fermentation process was then optimized by studying induction time, induction concentration, culture composition and fermentation temperature. as a result, a titer of 5436 mg L−1 and a productivity of 47 mg L−1 h−1 were achieved in 3 L bioreactor. Conclusions With metabolic engineering and fermentation optimization practice, C. glutamicum 21850 (pEC-C-vio1) was able to produce violacein with both titer and productivity at the highest level ever reported. Due to advantages of mature C. glutamicum fermentation industry, this work has built basis for commercial production of violacein. Electronic supplementary material The online version of this article (doi:10.1186/s12934-016-0545-0) contains supplementary material, which is available to authorized users.

Background Escherichia coli suffer from osmotic stress during succinic acid (SA) production, which reduces the performance of this microbial factory. Results Here, we report that a point mutation leading to a single amino acid change (D654Y) within the β-subunit of DNA-dependent RNA polymerase (RpoB) significantly improved the osmotolerance of E. coli. Importation of the D654Y mutation of RpoB into the parental strain, Suc-T110, increased cell growth and SA production by more than 40% compared to that of the control under high glucose osmolality. The transcriptome profile, determined by RNA-sequencing, showed two distinct stress responses elicited by the mutated RpoB that counterbalanced the osmotic stress. Under non-stressed conditions, genes involved in the synthesis and transport of compatible solutes such as glycine-betaine, glutamate or proline were upregulated even without osmotic stimulation, suggesting a “pre-defense” mechanism maybe formed in the rpoB mutant. Under osmotic stressed conditions, genes encoding diverse sugar transporters, which should be down-regulated in the presence of high osmotic pressure, were derepressed in the rpoB mutant. Additional genetic experiments showed that enhancing the expression of the mal regulon, especially for genes that encode the glycoporin LamB and maltose transporter, contributed to the osmotolerance phenotype. Conclusions The D654Y single amino acid substitution in RpoB rendered E. coli cells resistant to osmotic stress, probably due to improved cell growth and viability via enhanced sugar uptake under stressed conditions, and activated a potential “pre-defense” mechanism under non-stressed conditions. The findings of this work will be useful for bacterial host improvement to enhance its resistance to osmotic stress and facilitate bio-based organic acids production. Electronic supplementary material The online version of this article (doi:10.1186/s12896-017-0337-6) contains supplementary material, which is available to authorized users.

The MEP pathway genes were modulated to investigate whether there were new rate-limiting steps and toxic intermediates in this pathway. Activating IspG led to significant decrease of cell growth and β-carotene production. It was found that ispG overexpression led to accumulation of intermediate HMBPP, which seriously interfered with synthesis machinery of nucleotide and protein in Escherichia coli. Activation of the downstream enzyme IspH could solve HMBPP accumulation problem and eliminate the negative effects of ispG overexpression. In addition, intermediate MECPP accumulated in the starting strain, while balanced activation of IspG and IspH could push the carbon flux away from MECPP and led to 73% and 77% increase of β-carotene and lycopene titer respectively. Our work for the first time identified HMBPP to be a cytotoxic intermediate in MEP pathway and demonstrated that balanced activation of IspG and IspH could eliminate accumulation of HMBPP and MECPP and improve isoprenoids production.

Dilute acid pretreatment is an established method for hydrolyzing the methylglucuronoxylans of hemicellulose to release fermentable xylose. In addition to xylose, this process releases the aldouronate methylglucuronoxylose, which cannot be metabolized by current ethanologenic biocatalysts. Enterobacter asburiae JDR-1, isolated from colonized wood, was found to efficiently ferment both methylglucuronoxylose and xylose in acid hydrolysates of sweet gum xylan, producing predominantly ethanol and acetate. Transformation of E. asburiae JDR-1 with pLOI555 or pLOI297, each containing the PET operon containing pyruvate decarboxylase ( pdc ) and alcohol dehydrogenase B ( adhB ) genes derived from Zymomonas mobilis , replaced mixed-acid fermentation with homoethanol fermentation. Deletion of the pyruvate formate lyase ( pflB ) gene further increased the ethanol yield, resulting in a stable E. asburiae E1(pLOI555) strain that efficiently utilized both xylose and methylglucuronoxylose in dilute acid hydrolysates of sweet gum xylan. Ethanol was produced from xylan hydrolysate by E. asburiae E1(pLOI555) with a yield that was 99% of the theoretical maximum yield and at a rate of 0.11 g ethanol/g (dry weight) cells/h, which was 1.57 times the yield and 1.48 times the rate obtained with the ethanologenic strain Escherichia coli KO11. This engineered derivative of E. asburiae JDR-1 that is able to ferment the predominant hexoses and pentoses derived from both hemicellulose and cellulose fractions is a promising subject for development as an ethanologenic biocatalyst for production of fuels and chemicals from agricultural residues and energy crops.

In the dilute acid pretreatment of lignocellulose, xylose substituted with alpha-1,2-methylglucuronate is released as methylglucuronoxylose (MeGAX), which cannot be fermented by biocatalysts currently used to produce biofuels and chemicals. Enterobacter asburiae JDR-1, isolated from colonized wood, efficiently fermented both MeGAX and xylose in acid hydrolysates of sweetgum xylan. Deletion of pflB and als genes in this bacterium modified the native mixed acid fermentation pathways to one for homolactate production. The resulting strain, Enterobacter asburiae L1, completely utilized both xylose and MeGAX in a dilute acid hydrolysate of sweetgum xylan and produced lactate approximating 100% of the theoretical maximum yield. Enterobacter asburiae JDR-1 offers a platform to develop efficient biocatalysts for production of fuels and chemicals from hemicellulose hydrolysates of hardwood and agricultural residues.

Background Synthetic biology aims to engineer biological systems for desired behaviors. The construction of these systems can be complex, often requiring genetic reprogramming, extensive de novo DNA synthesis, and functional screening. Results Herein, we present a programmable, multipurpose microfluidic platform and associated software and apply the platform to major steps of the synthetic biology research cycle: design, construction, testing, and analysis. We show the platform’s capabilities for multiple automated DNA assembly methods, including a new method for Isothermal Hierarchical DNA Construction, and for Escherichia coli and Saccharomyces cerevisiae transformation. The platform enables the automated control of cellular growth, gene expression induction, and proteogenic and metabolic output analysis. Conclusions Taken together, we demonstrate the microfluidic platform’s potential to provide end-to-end solutions for synthetic biology research, from design to functional analysis. Electronic supplementary material The online version of this article (doi:10.1186/s13036-016-0024-5) contains supplementary material, which is available to authorized users.

Background: The chemoautotrophic bacterium Ralstonia eutropha can utilize H2/CO2 for growth under aerobic conditions. While this microbial host has great potential to be engineered to produce desired compounds (beyond polyhydroxybutyrate) directly from CO2, little work has been done to develop genetic part libraries to enable such endeavors. Results: We report the development of a toolbox for the metabolic engineering of Ralstonia eutropha H16. We have constructed a set of broad-host-range plasmids bearing a variety of origins of replication, promoters, 5' mRNA stem-loop structures, and ribosomal binding sites. Specifically, we analyzed the origins of replication pCM62 (IncP), pBBR1, pKT (IncQ), and their variants. We tested the promoters PBAD, T7, Pxyls/PM, PlacUV5, and variants thereof for inducible expression. We also evaluated a T7 mRNA stem-loop structure sequence and compared a set of ribosomal binding site (RBS) sequences derived from Escherichia coli, R. eutropha, and a computational RBS design tool. Finally, we employed the toolbox to optimize hydrocarbon production in R. eutropha and demonstrated a 6-fold titer improvement using the appropriate combination of parts.Conclusion: We constructed and evaluated a versatile synthetic biology toolbox for Ralstonia eutropha metabolic engineering that could apply to other microbial hosts as well. © 2013 Bi et al.; licensee BioMed Central Ltd.

Acid pretreatment is commonly used to release pentoses from the hemicellulose fraction of cellulosic biomass for bioconversion. The predominant pentose in the hemicellulose fraction of hardwoods and crop residues is xylose in the polysaccharide methylglucuronoxylan, in which as many as one in six of the β-1,4-linked xylopyranose residues is substituted with α-1,2-linked 4- O -methylglucuronopyranose. Resistance of the α-1,2-methylglucuronosyl linkages to acid hydrolysis results in release of the aldobiuronate 4- O -methylglucuronoxylose, which is not fermented by bacterial biocatalysts currently used for bioconversion of hemicellulose. Enterobacter asburiae strain JDR-1, isolated from colonized hardwood (sweetgum), efficiently ferments both methylglucuronoxylose and xylose, producing predominantly ethanol and acetate. 13 C-nuclear magnetic resonance studies defined the Embden-Meyerhof pathway for metabolism of glucose and the pentose phosphate pathway for xylose metabolism. Rates of substrate utilization, product formation, and molar growth yields indicated methylglucuronoxylose is transported into the cell and hydrolyzed to release methanol, xylose, and hexauronate. Enterobacter asburiae strain JDR-1 is the first microorganism described that ferments methylglucuronoxylose generated along with xylose during the acid-mediated saccharification of hemicellulose. Genetic definition of the methylglucuronoxylose utilization pathway may allow metabolic engineering of established gram-negative bacterial biocatalysts for complete bioconversion of acid hydrolysates of methylglucuronoxylan. Alternatively, Enterobacter asburiae strain JDR-1 may be engineered for the efficient conversion of acid hydrolysates of hemicellulose to biofuels and chemical feedstocks.

Background Plasmid expression is a popular method in studies of MVA pathway for isoprenoid production in Escherichia coli. However, heterologous gene expression with plasmid is often not stable and might burden growth of host cells, decreases cell mass and product yield. In this study, MVA pathway was divided into three modules, and two heterologous modules were integrated into the E. coli chromosome. These modules were individually modulated with regulatory parts to optimize efficiency of the pathway in terms of downstream isoprenoid production. Results MVA pathway modules Hmg1-erg12 operon and mvaS-mvaA-mavD1 operon were integrated into E. coli chromosome followed by modulation with promoters with varied strength. Along with activation of atoB, a 26% increase of β-carotene production with no effect on cell growth was obtained. With a combinatory modulation of two key enzymes mvas and Hmg1 with degenerate RBS library, β-carotene showed a further increase of 51%. Conclusions Our study provides a novel strategy for improving production of a target compound through integration and modulation of heterologous pathways in both transcription and translation level. In addition, a genetically hard-coded chassis with both efficient MEP and MVA pathways for isoprenoid precursor supply was constructed in this work. Electronic supplementary material The online version of this article (doi:10.1186/s12934-016-0607-3) contains supplementary material, which is available to authorized users.

Background Microbial genome editing is a powerful tool to modify chromosome in way of deletion, insertion or replacement, which is one of the most important techniques in metabolic engineering research. The emergence of CRISPR/Cas9 technique inspires various genomic editing methods. Results In this research, the goal of development of a fast and easy method for Escherichia coli genome editing with high efficiency is pursued. For this purpose, we designed modular plasmid assembly strategy, compared effects of different length of homologous arms for recombination, and tested different sets of recombinases. The final technique we developed only requires one plasmid construction and one transformation of practice to edit a genomic locus with 3 days and minimal lab work. In addition, the single temperature sensitive plasmid is easy to eliminate for another round of editing. Especially, process of the modularized editing plasmid construction only takes 4 h. Conclusion In this study, we developed a fast and easy genome editing procedure based on CRISPR/Cas9 system that only required the work of one plasmid construction and one transformation, which allowed modification of a chromosome locus within 3 days and could be performed continuously for multiple loci. Electronic supplementary material The online version of this article (doi:10.1186/s12934-016-0605-5) contains supplementary material, which is available to authorized users.

Saccharomyces cerevisiae has already been used for heterologous production of fuel chemicals and valuable natural products. The establishment of complicated heterologous biosynthetic pathways in S. cerevisiae became the research focus of Synthetic Biology and Metabolic Engineering. Thus, simple and efficient genomic integration techniques of large number of transcription units are demanded urgently. An efficient DNA assembly and chromosomal integration method was created by combining homologous recombination (HR) in S. cerevisiae and Golden Gate DNA assembly method, designated as modularized two-step (M2S) technique. Two major assembly steps are performed consecutively to integrate multiple transcription units simultaneously. In Step 1, Modularized scaffold containing a head-to-head promoter module and a pair of terminators was assembled with two genes. Thus, two transcription units were assembled with Golden Gate method into one scaffold in one reaction. In Step 2, the two transcription units were mixed with modules of selective markers and integration sites and transformed into S. cerevisiae for assembly and integration. In both steps, universal primers were designed for identification of correct clones. Establishment of a functional β-carotene biosynthetic pathway in S. cerevisiae within 5 days demonstrated high efficiency of this method, and a 10-transcriptional-unit pathway integration illustrated the capacity of this method. Modular design of transcription units and integration elements simplified assembly and integration procedure, and eliminated frequent designing and synthesis of DNA fragments in previous methods. Also, by assembling most parts in Step 1 in vitro, the number of DNA cassettes for homologous integration in Step 2 was significantly reduced. Thus, high assembly efficiency, high integration capacity, and low error rate were achieved.