Metabolic Engineering Communications最新文献

L-Tryptophan (Trp) is a high-value aromatic amino acid with diverse applications in food and pharmaceutical industries. Although production of Trp by engineered Escherichia coli has been extensively studied, the need of multiple precursors for its synthesis and the complex regulations of the biosynthetic pathways make the achievement of a high product yield still very challenging. Metabolic flux analysis suggests that the use of a phosphoenolpyruvate:sugar phosphotransferase system (PTS) independent glucose uptake system, i.e. the galactose permease/glucokinase (GalP/Glk) system, can theoretically double the Trp yield from glucose. To explore this possibility, a PTS⁻ and GalP/Glk-dependent E. coli strain was constructed from a previously rationally developed Trp producer strain S028. However, the growth rate of the S028 mutant was severely impaired. To overcome this problem, promoter screening for modulated gene expression of GalP/Glk was carried out, following by a batch mode of adaptive laboratory evolution (ALE) which resulted in a strain K3 with a similar Trp yield and concentration as S028. In order to obtain a more efficient Trp producer, a novel continuous ALE system was developed by combining CRISPR/Cas9-facilitated in vivo mutagenesis with real-time measurement of cell growth and online monitoring of Trp-mediated fluorescence intensity. With the aid of this automatic system (auto-CGSS), a promising strain T5 was obtained and fed-batch fermentations showed an increase of Trp yield by 19.71% with this strain compared with that obtained by the strain K3 (0.164 vs. 0.137 ​g/g). At the same time, the specific production rate was increased by 52.93% (25.28 vs. 16.53 ​mg/g _DCW/h). Two previously engineered enzyme variants AroG^D6G−D7A and AnTrpC^R378F were integrated into the strain T5, resulting in a highly productive strain T5AA with a Trp yield of 0.195 ​g/g and a specific production rate of 28.83 ​mg/g _DCW/h.

The necessity to decrease our fossil energy dependence requests bioprocesses based on biomass degradation. Cellobiose is the main product released by cellulases when acting on the major plant cell wall polysaccharide constituent, the cellulose. Escherichia coli, one of the most common model organisms for the academy and the industry, is unable to metabolize this disaccharide. In this context, the remodeling of E. coli to catabolize cellobiose should thus constitute an important progress for the design of such applications. Here, we developed a robust E. coli strain able to metabolize cellobiose by integration of a small set of modifications in its genome. Contrary to previous studies that use adaptative evolution to achieve some growth on this sugar by reactivating E. coli cryptic operons coding for cellobiose metabolism, we identified easily insertable modifications impacting the cellobiose import (expression of a gene coding a truncated variant of the maltoporin LamB, modification of the expression of lacY encoding the lactose permease) and its intracellular degradation (genomic insertion of a gene encoding either a cytosolic β-glucosidase or a cellobiose phosphorylase). Taken together, our results provide an easily transferable set of mutations that confers to E. coli an efficient growth phenotype on cellobiose (doubling time of 2.2 ​h in aerobiosis) without any prior adaptation.

Cyanobacteria are one of the most promising microorganisms to produce biofuels and renewable chemicals due to their oxygenic autotrophic growth properties. However, to rely on photosynthesis, which is one of the main reasons for slow growth, low carbon assimlation rate and low production, is a bottleneck. To address this challenge, optimizing the Calvin-Benson-Bassham (CBB) cycle is one of the strategies since it is the main carbon fixation pathway. In a previous study, we showed that overexpression of either aldolase (FBA), transketolase (TK), or fructose-1,6/sedoheptulose-1,7-bisphosphatase (FBP/SBPase), enzymes responsible for RuBP regeneration and vital for controlling the CBB carbon flux, led to higher production rates and titers in ethanol producing strains of Synechocystis PCC 6803. In the present study, we investigated the combined effects of the above enzymes on ethanol production in Synechocystis PCC 6803.

The ethanol production of the strains overexpressing two CBB enzymes (FBA ​+ ​TK, FBP/SBPase ​+ ​FBA or FBP/SBPase ​+ ​TK) was higher than the respective control strains, overexpressing either FBA or TK. The co-overexpression of FBA and TK led to more than 9 times higher ethanol production compared to the overexpression of FBA. Compared to TK the respective increase is 4 times more ethanol production. Overexpression of FBP/SBPase in combination with FBA showed 2.5 times higher ethanol production compared to FBA. Finally, co-overexpression of FBP/SBPase and TK reached about twice the production of ethanol compared to overexpression of only TK. This study clearly demonstrates that overexpression of two selected CBB enzymes leads to significantly increased ethanol production compared to overexpression of a single CBB enzyme.

Genome-scale stoichiometric models (GSMs) have been widely utilized to predict and understand cellular metabolism. GSMs and the flux predictions resulting from them have proven indispensable to fields ranging from metabolic engineering to human disease. Nonetheless, it is challenging to parse these flux predictions due to the inherent size and complexity of the GSMs. Several previous approaches have reduced this complexity by identifying key pathways contained within the genome-scale flux predictions. However, a reduction method that overlays carbon atom transitions on stoichiometry and flux predictions is lacking. To fill this gap, we developed NetFlow, an algorithm that leverages genome-scale carbon mapping to extract and quantitatively distinguish biologically relevant metabolic pathways from a given genome-scale flux prediction. NetFlow extends prior approaches by utilizing both full carbon mapping and context-specific flux predictions. Thus, NetFlow is uniquely able to quantitatively distinguish between biologically relevant pathways of carbon flow within the given flux map. NetFlow simulates ¹³C isotope labeling experiments to calculate the extent of carbon exchange, or carbon yield, between every metabolite in the given GSM. Based on the carbon yield, the carbon flow to or from any metabolite or between any pair of metabolites of interest can be isolated and readily visualized. The resulting pathways are much easier to interpret, which enables an in-depth mechanistic understanding of the metabolic phenotype of interest. Here, we first demonstrate NetFlow with a simple network. We then depict the utility of NetFlow on a model of central carbon metabolism in E. coli. Specifically, we isolated the production pathway for succinate synthesis in this model and the metabolic mechanism driving the predicted increase in succinate yield in a double knockout of E. coli. Finally, we describe the application of NetFlow to a GSM of lycopene-producing E. coli, which enabled the rapid identification of the mechanisms behind the measured increases in lycopene production following single, double, and triple knockouts.

Reconstitution of prenylflavonoids using the flavonoid biosynthetic pathway and prenyltransferases (PTs) in microbes can be a promising attractive alternative to plant-based production or chemical synthesis. Here, we demonstrate that promiscuous microbial PTs can be a substitute for regiospecific but mostly unidentified botanical PTs. To test the prenylations of naringenin, we constructed a yeast strain capable of producing naringenin from l-phenylalanine by genomic integration of six exogenous genes encoding components of the naringenin biosynthetic pathway. Using this platform strain, various microbial PTs were tested for prenylnaringenin production. In vitro screening demonstrated that the fungal AnaPT (a member of the tryptophan dimethylallyltransferase family) specifically catalyzed C-3′ prenylation of naringenin, whereas SfN8DT-1, a botanical PT, specifically catalyzed C-8 prenylation. In vivo, the naringenin-producing strain expressing the microbial AnaPT exhibited heterologous microbial production of 3′-prenylnaringenin (3′-PN), in contrast to the previously reported in vivo production of 8-prenylnaringenin (8-PN) using the botanical SfN8DT-1. These findings provide strategies towards expanding the production of a variety of prenylated compounds, including well-known prenylnaringenins and novel prenylflavonoids. These results also suggest the opportunity for substituting botanical PTs, both known and unidentified, that display relatively strict regiospecificity of the prenyl group transfer.

Mannosylerythritol lipids (MELs) are surface active glycolipids secreted by various fungi. MELs can be used as biosurfactants and are a biodegradable resource for the production of detergents or pharmaceuticals. Different fungal species synthesize a unique mixture of MELs differing in acetyl- and acyl-groups attached to the sugar moiety. Here, we report the construction of a toolbox for production of glycolipids with predictable fatty acid side chains in the basidiomycete Ustilago maydis. Genes coding for acyl-transferases involved in MEL production (Mac1 and Mac2) from different fungal species were combined to obtain altered MEL variants with distinct physical properties and altered antimicrobial activity. We also demonstrate that a U. maydis paralog of the acyltransferase Mac2 with a different substrate specificity can be employed for the biosynthesis of modified MEL variants. In summary, our data showcase how the fungal repertoire of Mac enzymes can be used to engineer tailor-made MELs according to specific biotechnological or pharmaceutical requirements.

Terpenoids are a wide class of organic compounds with industrial relevance. The natural ability of cyanobacteria to produce terpenoids via the methylerythritol 4-phosphate (MEP) pathway makes these organisms appealing candidates for the generation of light-driven cell factories for green chemistry. Here we address the improvement of the production of (E)-α-bisabolene, a valuable biofuel feedstock, in Synechocystis sp. PCC 6803 via sequential heterologous expression of bottleneck enzymes of the native pathway. Expression of the bisabolene synthase is sufficient to complete the biosynthetic pathway of bisabolene. Expression of a farnesyl-pyrophosphate synthase from Escherichia coli did not influence production of bisabolene, while enhancement of the MEP pathway via additional overexpression of 1-deoxy-D-xylulose-5-phosphate synthase (DXS) and IPP/DMAPP isomerase (IDI) significantly increased production per cell. However, in the absence of a carbon sink, the overexpression of DXS and IDI leads to significant growth impairment. The final engineered strain reached a volumetric titre of 9 ​mg ​L⁻¹ culture of bisabolene after growing for 12 days. When the cultures were grown in a high cell density (HCD) system, we observed an increase in the volumetric titres by one order of magnitude for all producing-strains. The strain with improved MEP pathway presented an increase twice as much as the remaining engineered strains, yielding more than 180 ​mg ​L⁻¹ culture after 10 days of cultivation. Furthermore, the overexpression of these two MEP enzymes prevented the previously reported decrease in the bisabolene specific titres when grown in HCD conditions, where primary metabolism is usually favoured. We conclude that fine-tuning of the cyanobacterial terpenoid pathway is crucial for the generation of microbial platforms for terpenoid production on industrial-scale.

Cyanobacteria play an important role in photobiotechnology. Yet, one of their key central metabolic pathways, the tricarboxylic acid (TCA) cycle, has a unique architecture compared to most heterotrophs and still remains largely unexploited. The conversion of 2-oxoglutarate to succinate via succinyl-CoA is absent but is by-passed by several other reactions. Overall, fluxes under photoautotrophic growth conditions through the TCA cycle are low, which has implications for the production of chemicals. In this study, we investigate the capacity of the TCA cycle of Synechocystis sp PCC 6803 for the production of trans-4-hydroxy-L-proline (Hyp), a valuable chiral building block for the pharmaceutical and cosmetic industries. For the first time, photoautotrophic Hyp production was achieved in a cyanobacterium expressing the gene for the L-proline-4-hydroxylase (P4H) from Dactylosporangium sp. strain RH1. Interestingly, while elevated intracellular Hyp concentrations could be detected in the recombinant Synechocystis strains under all tested conditions, detectable Hyp secretion into the medium was only observed when the pH of the medium exceeded 9.5 and mostly in the late phases of the cultivation. We compared the rates obtained for autotrophic Hyp production with published sugar-based production rates in E. coli. The land-use efficiency (space-time yield) of the phototrophic process is already in the same order of magnitude as the heterotrophic process considering sugar farming as well. But, the remarkable plasticity of the cyanobacterial TCA cycle promises the potential for a 23–55 fold increase in space-time yield when using Synechocystis. Altogether, these findings contribute to a better understanding of bioproduction from the TCA cycle in photoautotrophs and broaden the spectrum of chemicals produced in metabolically engineered cyanobacteria.

Foot-and-mouth disease virus (FMDV) 2A constructs have been successfully used for the production of “Golden Rice”, a β-carotene producing rice strain. However, to allay public fears and opposition to plants carrying a mammalian pathogenic viral sequence, 2A-like synthetic sequences from Thosea asigna virus and Infectious myonecrosis virus were used to coordinate the coexpression of carotenoid biosynthetic genes. Here, up to four carotenogenic genes encoding PSY, CRTI, BCH and BKT were concatenated and produced β-carotene, zeaxanthin, and ketocarotenoids (astaxanthin and adonixanthin) in transgenic rice seeds displaying color variation due to the difference in carotenoid content and composition.

An engineered B. subtilis 1A1 strain (BsADH2) expressing a secondary alcohol dehydrogenase (CpSADH) was co-cultured with C. beijerinckii G117 under an aerobic condition. During the fermentation on glucose, B. subtilis BsADH2 depleted oxygen in culture media completely and created an anaerobic environment for C. beijerinckii G117, an obligate anaerobe, to grow. Meanwhile, lactate produced by B. subtilis BsADH2 was re-assimilated by C. beijerinckii G117. In return, acetone produced by C. beijerinckii G117 was reduced into isopropanol by B. subtilis BsADH2 via expressing the CpSADH, which helped maintain the redox balance of the engineered B. subtilis. In the symbiotic system consisting of two strains, 1.7 ​g/L of acetone, 4.8 ​g/L of butanol, and 0.9 ​g/L of isopropanol (with an isopropanol/acetone ratio of 0.53) was produced from 60 ​g/L of glucose. This symbiotic system also worked when oxygen was supplied to the culture, although less isopropanol was produced (0.9 ​g/L of acetone, 4.9 ​g/L of butanol, and 0.2 ​g/L of isopropanol). The isopropanol titer was increased substantially to 2.5 ​g/L when we increased the inoculum size of B. subtilis BsADH2 and optimized other process parameters. With the Bacillus-Clostridium co-culture, switching from the original acetone-butanol (AB) fermentation to an aerobic acetone-butanol-isopropanol (ABI) fermentation can be easily achieved without genetic engineering of Clostridium. This strategy of employing a recombinant Bacillus to co-culture with Clostridium should be potentially useful to modify traditional acetone-butanol-ethanol fermentation for the production of other value-added chemicals.