Obtaining the best igRNAs for bystander-less correction of all ABE-reversible pathogenic SNVs using high-throughput screening.

Imperfect -gRNA (igRNA) provides a simple strategy for single-base editing of a base editor. However, a significant number of igRNAs need to be generated and tested for each target locus to achieve efficient single-base reversion of pathogenic single nucleotide variations (SNVs), which hinders the direct application of this technology. To provide ready-to-use igRNAs for single-base and bystander-less correction of all the adenine base editor (ABE)-reversible pathogenic SNVs, we employed a high-throughput method to edit all 5,253 known ABE-reversible pathogenic SNVs, each with multiple systematically designed igRNAs, and two libraries of 96,000 igRNAs were tested. A total of 1,988 SNV loci could be single-base reversed by igRNA with a >30% efficiency. Among these 1,988 loci, 378 SNV loci exhibited an efficiency of more than 90%. At the same time, the bystander editing efficiency of 76.62% of the SNV loci was reduced to 0%, while remaining below 1% for another 18.93% of the loci. These ready-to-use igRNAs provided the best solutions for a substantial portion of the 4,657 pathogenic/likely pathogenic SNVs. In this work, we overcame one of the most significant obstacles of base editors and provide a ready-to-use platform for the genetic treatment of diseases caused by ABE-reversible SNVs.

Imperfect guide-RNA (igRNA) enables CRISPR single-base editing with ABE and CBE.

CRISPR base editing techniques tend to edit multiple bases in the targeted region, which is a limitation for precisely reverting disease-associated single-nucleotide polymorphisms (SNPs). We designed an imperfect gRNA (igRNA) editing methodology, which utilized a gRNA with one or more bases that were not complementary to the target locus to direct base editing toward the generation of a single-base edited product. Base editing experiments illustrated that igRNA editing with CBEs greatly increased the single-base editing fraction relative to normal gRNA editing with increased editing efficiencies. Similar results were obtained with an adenine base editor (ABE). At loci such as DNMT3B, NSD1, PSMB2, VIATA hs267 and ANO5, near-perfect single-base editing was achieved. Normally an igRNA with good single-base editing efficiency could be selected from a set of a few igRNAs, with a simple protocol. As a proof-of-concept, igRNAs were used in the research to construct cell lines of disease-associated SNP causing primary hyperoxaluria construction research. This work provides a simple strategy to achieve single-base base editing with both ABEs and CBEs and overcomes a key obstacle that limits the use of base editors in treating SNP-associated diseases or creating disease-associated SNP-harboring cell lines and animal models.

Targeted C-to-T and A-to-G dual mutagenesis system for RhtA transporter in vivo evolution.

T7 RNA polymerase (T7RNAP) has been fused with cytosine or adenine deaminase individually, enabling in vivo C-to-T or A-to-G transitions on DNA sequence downstream of T7 promoter, and greatly accelerated directed protein evolution. However, its base conversion type is limited. In this study, we created a dual-functional system for simultaneous C-to-T and A-to-G in vivo mutagenesis, called T7-DualMuta, by fusing T7RNAP with both cytidine deaminase (PmCDA1) and a highly active adenine deaminase (TadA-8e). The C-to-T and A-to-G mutagenesis frequencies of T7-DualMuta were 4.02 × 10-3 and 1.20 × 10-2, respectively, with 24 h culturing and distributed mutations evenly across the target gene. The T7-DualMuta system was used to in vivo directed evolution of L-homoserine transporter RhtA, resulting in efficient variants that carried the four types of base conversions by T7-DualMuta. The evolved variants greatly increased the host growth rates at L-homoserine concentrations of 8 g/L, which was not previously achieved, and demonstrated the great in vivo evolution capacity. The novel molecular device T7-DualMuta efficiently provides both C/G-to-T/A and A/T-to-G/C mutagenesis on target regions, making it useful for various applications and research in Enzymology and Synthetic Biology studies. It also represents an important expansion of the base editing toolbox.ImportanceA T7-DualMuta system for simultaneous C-to-T and A-to-G in vivo mutagenesis was created. The mutagenesis frequency was 4.02 × 107 fold higher than the spontaneous mutation, which was reported to be approximately 10-10 bases per nucleotide per generation. This mutant system can be utilized for various applications and research in Enzymology and Synthetic Biology studies.

Reconstructed glycosylase base editors GBE2.0 with enhanced C-to-G base editing efficiency and purity.

Base editing techniques were developed for precise base conversion on cellular genomic DNA, which has great potential for the treatment of human genetic diseases. The glycosylase base editor (GBE) recently developed in our lab was used to perform C-to-G transversions in mammalian cells. To improve the application prospects of GBE, it is necessary to further increase its performance. With this aim, we replaced the human Ung in GBE with Ung1 from Saccharomyces cerevisiae. The resulting editor APOBEC-nCas9-Ung1 was tested at 17 chromosomal loci and was found to have an increased C-to-G editing efficiency ranging from 2.63% to 52.3%, with an average of 23.48%, which was a significant improvement over GBE, with an average efficiency of 15.54%, but with a decreased purity. For further improvement, we constructed APOBEC(R33A)-nCas9-Rad51-Ung1 with two beneficial modifications adapted from previous reports. This base editor was able to achieve even higher editing efficiency ranging from 8.70% to 72.1%, averaging 30.88%, while also exhibiting high C-to-G purity ranging from 35.57% to 92.92%, and was designated GBE2.0. GBE2.0 provides high C-to-G editing efficiency and purity in mammalian cells, making it a powerful genetic tool for scientific research or potential genetic therapies for disease-causing G/C mutations.

Engineering Circularized mRNAs for the Production of Spider Silk Proteins.

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.

A CRISPR-based chromosomal-separation technique for Escherichia coli.

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.

CRISPR-based metabolic pathway engineering.

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

Helicase-AID: A novel molecular device for base editing at random genomic loci.

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 ß-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 ß-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.

Multiple strategies for metabolic engineering of Escherichia coli for efficient production of glycolate.

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.

Identification of Absidia orchidis steroid 11ß-hydroxylation system and its application in engineering Saccharomyces cerevisiae for one-step biotransformation to produce hydrocortisone.

Hydrocortisone is an effective anti-inflammatory drug and also an important intermediate for synthesis of other steroid drugs. The filamentous fungus Absidia orchidis is renowned for biotransformation of acetylated cortexolone through 11ß-hydroxylation to produce hydrocortisone. However, due to the presence of 11α-hydroxylase in A. orchidis, the 11α-OH by-product epi-hydrocortisone is always produced in a 1:1 M ratio with hydrocortisone. In order to decrease epi-hydrocortisone production, Saccharomyces cerevisiae was engineered in this work as an alternative way to produce hydrocortisone through biotransformation. Through transcriptomic analysis coupled with genetic verification in S. cerevisiae, the A. orchidis steroid 11ß-hydroxylation system was characterized, including a cytochrome P450 enzyme CYP5311B2 and its associated redox partners cytochrome P450 reductase and cytochrome b5. CYP5311B2 produces a mix of stereoisomers containing 11ß- and 11α-hydroxylation derivatives in a 4:1 M ratio. This fungal steroid 11ß-hydroxylation system was reconstituted in S. cerevisiae for hydrocortisone production, resulting in a productivity of 22 mg/L·d. Protein engineering of CYP5311B2 generated a R126D/Y398F variant, which had 3 times higher hydrocortisone productivity compared to the wild type. Elimination of C20-hydroxylation by-products and optimization of the expression of A. orchidis 11ß-hydroxylation system genes further increased hydrocortisone productivity by 238% to 223 mg/L·d. In addition, a novel steroid transporter ClCDR4 gene was identified from Cochliobolus lunatus, overexpression of which further increased hydrocortisone productivity to 268 mg/L·d in S. cerevisiae. Through increasing cell mass, 1060 mg/L hydrocortisone was obtained in 48 h and the highest productivity reached 667 mg/L·d. This is the highest hydrocortisone titer reported for yeast biotransformation system so far.

Construction of a carbon-conserving pathway for glycolate production by synergetic utilization of acetate and glucose in Escherichia coli.

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.

A novel gene expression system for Ralstonia eutropha based on the T7 promoter.

BACKGROUND: Ralstonia eutropha (syn. Cupriavidus necator) is a model microorganism for studying metabolism of polyhydroxyalkanoates (PHAs) and a potential chassis for protein expression due to various advantages. Although current plasmid systems of R. eutropha provide a basic platform for gene expression, the performance of the expression-inducing systems is still limited. In addition, the sizes of the cloned genes are limited due to the large sizes of the plasmid backbones. RESULTS: In this study, an R. eutropha T7 expression system was established by integrating a T7 RNA polymerase gene driven by the PBAD promoter into the genome of R. eutropha, as well as adding a T7 promoter into a pBBR1-derived plasmid for gene expression. In addition, the essential DNA sequence necessary for pBBR1 plasmid replication was identified, and the redundant parts were deleted reducing the expression plasmid size to 3392 bp, which improved the electroporation efficiency about 4 times. As a result, the highest expression level of RFP was enhanced, and the L-arabinose concentration for expression induction was decreased 20 times. CONCLUSIONS: The R. eutropha T7 expression system provides an efficient platform for protein production and synthetic biology applications.

Engineering the Calvin-Benson-Bassham cycle and hydrogen utilization pathway of Ralstonia eutropha for improved autotrophic growth and polyhydroxybutyrate production.

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.

Manipulating the position of DNA expression cassettes using location tags fused to dCas9 (Cas9-Lag) to improve metabolic pathway efficiency.

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 ß-carotene serve as common intermediates for the synthesis of derivative compounds, which are hydrophobic and usually accumulate in the membrane compartment. RESULTS: Carotenoids like ß-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.

A programmable CRISPR/Cas9-based phage defense system for Escherichia coli BL21(DE3).

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 h 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.

Engineering an electroactive Escherichia coli for the microbial electrosynthesis of succinate from glucose and CO2.

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.

Construction of Escherichia coli cell factories for crocin biosynthesis.

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, ß-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.

Production of 14α-hydroxysteroids by a recombinant Saccharomyces cerevisiae biocatalyst expressing of a fungal steroid 14α-hydroxylation system.

The 14α-hydroxysteroids have specific anti-gonadotropic and carcinolytic biological activities and can be produced by microbial biotransformation. The steroid 11ß-/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 11ß-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.

CRISPR-Cas9-assisted native end-joining editing offers a simple strategy for efficient genetic engineering in Escherichia coli.

Unlike eukaryotes, prokaryotes are less proficient in homologous recombination (HR) and non-homologous end-joining (NHEJ). All existing genomic editing methods for Escherichia coli (E. 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 genetic 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 deletion of chromosome fragments up to 83 kb or gene inactivation. Moreover, the positive rate and editing efficiency are independent of high-efficiency competent cells. The method requires neither exogenous DNA repair systems nor introduced editing template. The Cas9-sgRNA 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.

Development of an autotrophic fermentation technique for the production of fatty acids using an engineered Ralstonia eutropha cell factory.

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.