A systematic optimization of medium chain fatty acid biosynthesis via the reverse beta-oxidation cycle in Escherichia coli

Medium-chain fatty acids (MCFAs, 6–10 carbons) are valuable precursors to many industrial biofuels and chemicals, recently engineered reversal of the β-oxidation (r-BOX) cycle has been proposed as a potential platform for efficient synthesis of MCFAs. Previous studies have made many exciting achievements on functionally characterizing four core enzymes of this r-BOX cycle. However, the information about bottleneck nodes in this cycle is elusive. Here, a quantitative assessment of the inherent limitations of this cycle was conducted to capitalize on its potential. The selection of the core β-oxidation reversal enzymes in conjunction with acetyl-CoA synthetase endowed the ability to synthesize about 1 g/L MCFAs. Furthermore, a gene dosage experiment was developed to identify two rate-limiting enzymes (acetyl-CoA synthetase and thiolase). The de novo pathway was then separated into two modules at thiolase and MCFA production titer increased to 2.8 g/L after evaluating different construct environments. Additionally, the metabolism of host organism was reprogrammed to the desired biochemical product by the clustered regularly interspaced short palindromic repeats interference system, resulted in a final MCFA production of 3.8 g/L. These findings described here identified the inherent limitations of r-BOX cycle and further unleashed the lipogenic potential of this cycle, thus paving the way for the development of a bacterial platform for microbial production of high-value oleo-chemicals from low-value carbons in a sustainable and environmentally friendly manner.     

Engineering Escherichia coli for the synthesis of short- and medium-chain α,β-unsaturated carboxylic acids

Concerns over sustained availability of fossil resources along with environmental impact of their use have stimulated the development of alternative methods for fuel and chemical production from renewable resources. In this work, we present a new approach to produce α,β-unsaturated carboxylic acids (α,β-UCAs) using an engineered reversal of the β-oxidation (r-BOX) cycle. To increase the availability of both acyl-CoAs and enoyl-CoAs for α,β-UCA production, we use an engineered Escherichia coli strain devoid of mixed-acid fermentation pathways and known thioesterases. Core genes for r-BOX such as thiolase, hydroxyacyl-CoA dehydrogenase, enoyl-CoA hydratase, and enoyl-CoA reductase were chromosomally overexpressed under the control of a cumate inducible phage promoter. Native E. coli thioesterase YdiI was used as the cycle-terminating enzyme, as it was found to have not only the ability to convert trans-enoyl-CoAs to the corresponding α,β-UCAs, but also a very low catalytic efficiency on acetyl-CoA, the primer and extender unit for the r-BOX pathway. Coupling of r-BOX with YdiI led to crotonic acid production at titers reaching 1.5 g/L in flask cultures and 3.2 g/L in a controlled bioreactor. The engineered r-BOX pathway was also used to achieve for the first time the production of 2-hexenoic acid, 2-octenoic acid, and 2-decenoic acid at a final titer of 0.2 g/L. The superior nature of the engineered pathway was further validated through the use of in silico metabolic flux analysis, which showed the ability of r-BOX to support growth-coupled production of α,β-UCAs with a higher ATP efficiency than the widely used fatty acid biosynthesis pathway. Taken together, our findings suggest that r-BOX could be an ideal platform to implement the biological production of α,β-UCAs.     

Engineering of carboligase activity reaction in Candida glabrata for acetoin production

Utilization of Candida glabrata overproducing pyruvate is a promising strategy for high-level acetoin production. Based on the known regulatory and metabolic information, acetaldehyde and thiamine were fed to identify the key nodes of carboligase activity reaction (CAR) pathway and provide a direction for engineering C. glabrata. Accordingly, alcohol dehydrogenase, acetaldehyde dehydrogenase, pyruvate decarboxylase, and butanediol dehydrogenase were selected to be manipulated for strengthening the CAR pathway. Following the rational metabolic engineering, the engineered strain exhibited increased acetoin biosynthesis (2.24 g/L). In addition, through in silico simulation and redox balance analysis, NADH was identified as the key factor restricting higher acetoin production. Correspondingly, after introduction of NADH oxidase, the final acetoin production was further increased to 7.33 g/L. By combining the rational metabolic engineering and cofactor engineering, the acetoin-producing C. glabrata was improved stepwise, opening a novel pathway for rational development of microorganisms for bioproduction.     

Engineering cofactor and transport mechanisms in Saccharomyces cerevisiae for enhanced acetyl-CoA and polyketide biosynthesis

Synthesis of polyketides at high titer and yield is important for producing pharmaceuticals and biorenewable chemical precursors. In this work, we engineered cofactor and transport pathways in Saccharomyces cerevisiae to increase acetyl-CoA, an important polyketide building block. The highly regulated yeast pyruvate dehydrogenase bypass pathway was supplemented by overexpressing a modified Escherichia coli pyruvate dehydrogenase complex (PDHm) that accepts NADP⁺ for acetyl-CoA production. After 24 h of cultivation, a 3.7-fold increase in NADPH/NADP⁺ ratio was observed relative to the base strain, and a 2.2-fold increase relative to introduction of the native E. coli PDH. Both E. coli pathways increased acetyl-CoA levels approximately 2-fold relative to the yeast base strain. Combining PDHm with a ZWF1 deletion to block the major yeast NADPH biosynthesis pathway resulted in a 12-fold NADPH boost and a 2.2-fold increase in acetyl-CoA. At 48 h, only this coupled approach showed increased acetyl-CoA levels, 3.0-fold higher than that of the base strain. The impact on polyketide synthesis was evaluated in a S. cerevisiae strain expressing the Gerbera hybrida 2-pyrone synthase (2-PS) for the production of the polyketide triacetic acid lactone (TAL). Titers of TAL relative to the base strain improved only 30% with the native E. coli PDH, but 3.0-fold with PDHm and 4.4-fold with PDHm in the Δzwf1 strain. Carbon was further routed toward TAL production by reducing mitochondrial transport of pyruvate and acetyl-CoA; deletions in genes POR2, MPC2, PDA1, or YAT2 each increased titer 2–3-fold over the base strain (up to 0.8 g/L), and in combination to 1.4 g/L. Combining the two approaches (NADPH-generating acetyl-CoA pathway plus reduced metabolite flux into the mitochondria) resulted in a final TAL titer of 1.6 g/L, a 6.4-fold increase over the non-engineered yeast strain, and 35% of theoretical yield (0.16 g/g glucose), the highest reported to date. These biological driving forces present new avenues for improving high-yield production of acetyl-CoA derived compounds.     

Engineering high-level production of fatty alcohols by Saccharomyces cerevisiae from lignocellulosic feedstocks

Fatty alcohols in the C12-C18 range are used in personal care products, lubricants, and potentially biofuels. These compounds can be produced from the fatty acid pathway by a fatty acid reductase (FAR), yet yields from the preferred industrial host Saccharomyces cerevisiae remain under 2% of the theoretical maximum from glucose. Here we improved titer and yield of fatty alcohols using an approach involving quantitative analysis of protein levels and metabolic flux, engineering enzyme level and localization, pull-push-block engineering of carbon flux, and cofactor balancing. We compared four heterologous FARs, finding highest activity and endoplasmic reticulum localization from a Mus musculus FAR. After screening an additional twenty-one single-gene edits, we identified increasing FAR expression; deleting competing reactions encoded by DGA1, HFD1, and ADH6; overexpressing a mutant acetyl-CoA carboxylase; limiting NADPH and carbon usage by the glutamate dehydrogenase encoded by GDH1; and overexpressing the Δ9-desaturase encoded by OLE1 as successful strategies to improve titer. Our final strain produced 1.2 g/L fatty alcohols in shake flasks, and 6.0 g/L in fed-batch fermentation, corresponding to ~ 20% of the maximum theoretical yield from glucose, the highest titers and yields reported to date in S. cerevisiae. We further demonstrate high-level production from lignocellulosic feedstocks derived from ionic-liquid treated switchgrass and sorghum, reaching 0.7 g/L in shake flasks. Altogether, our work represents progress towards efficient and renewable microbial production of fatty acid-derived products.     

Selection of an endogenous 2,3-butanediol pathway in Escherichia coli by fermentative redox balance

Fermentative redox balance has long been utilized as a metabolic evolution platform to improve efficiency of NADH-dependent pathways. However, such system relies on the complete recycling of NADH and may become limited when the target pathway results in excess NADH stoichiometrically. In this study, endogenous capability of Escherichia coli for 2,3-butanediol (2,3-BD) synthesis was explored using the anaerobic selection platform based on redox balance. To address the issue of NADH excess associated with the 2,3-BD pathway, we devised a substrate-decoupled system where a pathway intermediate is externally supplied in addition to the carbon source to decouple NADH recycling ratio from the intrinsic pathway stoichiometry. In this case, feeding of the 2,3-BD precursor acetoin effectively restored anaerobic growth of the mixed-acid fermentation mutant that remained otherwise inhibited even in the presence of a functional 2,3-BD pathway. Using established 2,3-BD dehydrogenases as model enzyme, we verified that the redox-based selection system is responsive to NADPH-dependent reactions but with lower sensitivity. Based on this substrate-decoupled selection scheme, we successfully identified the glycerol/1,2-propanediol dehydrogenase (Ec-GldA) as the major enzyme responsible for the acetoin reducing activity (k_cat/K_m≈0.4 mM⁻¹ s⁻¹) observed in E. coli. Significant shift of 2,3-BD configuration upon withdrawal of the heterologous acetolactate decarboxylase revealed that the endogenous synthesis of acetoin occurs via diacetyl. Among the predicted diacetyl reductase in E. coli, Ec-UcpA displayed the most significant activity towards diacetyl reduction into acetoin (V_max≈6 U/mg). The final strain demonstrated a meso-2,3-BD production titer of 3 g/L without introduction of foreign genes. The substrate-decoupled selection system allows redox balance regardless of the pathway stoichiometry thus enables segmented optimization of different reductive pathways through enzyme bioprospecting and metabolic evolution.     

Harnessing the respiration machinery for high-yield production of chemicals in metabolically engineered Lactococcus lactis

When modifying the metabolism of living organisms with the aim of achieving biosynthesis of useful compounds, it is essential to ensure that it is possible to achieve overall redox balance. We propose a generalized strategy for this, based on fine-tuning of respiration. The strategy was applied on metabolically engineered Lactococcus lactis strains to optimize the production of acetoin and (R,R)-2,3-butanediol (R-BDO). In the absence of an external electron acceptor, a surplus of two NADH per acetoin molecule is produced. We found that a fully activated respiration was able to efficiently regenerate NAD⁺, and a high titer of 371 mM (32 g/L) of acetoin was obtained with a yield of 82% of the theoretical maximum. Subsequently, we extended the metabolic pathway from acetoin to R-BDO by introducing the butanediol dehydrogenase gene from Bacillus subtilis. Since one mole of NADH is consumed when acetoin is converted into R-BDO per mole, only the excess of NADH needs to be oxidized via respiration. Either by fine-tuning the respiration capacity or by using a dual-phase fermentation approach involving a switch from fully respiratory to non-respiratory conditions, we obtained 361 mM (32 g/L) R-BDO with a yield of 81% or 365 mM (33 g/L) with a yield of 82%, respectively. These results demonstrate the great potential in using finely-tuned respiration machineries for bio-production.     

Efficient production of 2,3-butanediol in Saccharomyces cerevisiae by eliminating ethanol and glycerol production and redox rebalancing

2,3-Butanediol is a promising valuable chemical that can be used in various areas as a liquid fuel and a platform chemical. Here, 2,3-butanediol production in Saccharomyces cerevisiae was improved stepwise by eliminating byproduct formation and redox rebalancing. By introducing heterologous 2,3-butanediol biosynthetic pathway and deleting competing pathways producing ethanol and glycerol, metabolic flux was successfully redirected to 2,3-butanediol. In addition, the resulting redox cofactor imbalance was restored by overexpressing water-forming NADH oxidase (NoxE) from Lactococcus lactis. In a flask fed-batch fermentation with optimized conditions, the engineered adh1Δadh2Δadh3Δadh4Δadh5Δgpd1Δgpd2Δ strain overexpressing Bacillus subtilis α-acetolactate synthase (AlsS) and α-acetolactate decarboxylase (AlsD), S. cerevisiae 2,3-butanediol dehydrogenase (Bdh1), and L. lactis NoxE from a single multigene-expression vector produced 72.9 g/L 2,3-butanediol with the highest yield (0.41 g/g glucose) and productivity (1.43 g/(L·h)) ever reported in S. cerevisiae.     

Combining metabolic engineering and biocompatible chemistry for high-yield production of homo-diacetyl and homo-(S,S)-2,3-butanediol

Biocompatible chemistry is gaining increasing attention because of its potential within biotechnology for expanding the repertoire of biological transformations carried out by enzymes. Here we demonstrate how biocompatible chemistry can be used for synthesizing valuable compounds as well as for linking metabolic pathways to achieve redox balance and rescued growth. By comprehensive rerouting of metabolism, activation of respiration, and finally metal ion catalysis, we successfully managed to convert the homolactic bacterium Lactococcus lactis into a homo-diacetyl producer with high titer (95 mM or 8.2 g/L) and high yield (87% of the theoretical maximum). Subsequently, the pathway was extended to (S,S)-2,3-butanediol (S-BDO) through efficiently linking two metabolic pathways via chemical catalysis. This resulted in efficient homo-S-BDO production with a titer of 74 mM (6.7 g/L) S-BDO and a yield of 82%. The diacetyl and S-BDO production rates and yields obtained are the highest ever reported, demonstrating the promising combination of metabolic engineering and biocompatible chemistry as well as the great potential of L. lactis as a new production platform.     

Introduction of a bacterial acetyl-CoA synthesis pathway improves lactic acid production in Saccharomyces cerevisiae

Acid-tolerant Saccharomyces cerevisiae was engineered to produce lactic acid by expressing heterologous lactate dehydrogenase (LDH) genes, while attenuating several key pathway genes, including glycerol-3-phosphate dehydrogenase1 (GPD1) and cytochrome-c oxidoreductase2 (CYB2). In order to increase the yield of lactic acid further, the ethanol production pathway was attenuated by disrupting the pyruvate decarboxylase1 (PDC1) and alcohol dehydrogenase1 (ADH1) genes. Despite an increase in lactic acid yield, severe reduction of the growth rate and glucose consumption rate owing to the absence of ADH1 caused a considerable decrease in the overall productivity. In Δadh1 cells, the levels of acetyl-CoA, a key precursor for biologically applicable components, could be insufficient for normal cell growth. To increase the cellular supply of acetyl-CoA, we introduced bacterial acetylating acetaldehyde dehydrogenase (A-ALD) enzyme (EC 1.2.1.10) genes into the lactic acid-producing S. cerevisiae. Escherichia coli-derived A-ALD genes, mhpF and eutE, were expressed and effectively complemented the attenuated acetaldehyde dehydrogenase (ALD)/acetyl-CoA synthetase (ACS) pathway in the yeast. The engineered strain, possessing a heterologous acetyl-CoA synthetic pathway, showed an increased glucose consumption rate and higher productivity of lactic acid fermentation. The production of lactic acid was reached at 142 g/L with production yield of 0.89 g/g and productivity of 3.55 g L⁻¹ h⁻¹ under fed-batch fermentation in bioreactor. This study demonstrates a novel approach that improves productivity of lactic acid by metabolic engineering of the acetyl-CoA biosynthetic pathway in yeast.     

Production of butyric acid by a cellulolytic actinobacterium Thermobifida fusca on cellulose

Thermobifida fusca not only produces cellulases, hemicellulases and xylanases, but also excretes butyric acid. In order to achieve a high yield of butyric acid, the effect of different carbon sources: mannose, xylose, lactose, cellobiose, glucose, sucrose and acetates, on butyric acid production was studied. The highest yield of butyric acid was 0.67 g/g C (g-butyric acid/g-carbon input) on cellobiose. The best stir speed and aeration rate for butyric acid production were found to be 400 rpm and 2 vvm in a 5-L fermentor. The maximum titer of 2.1 g/L butyric acid was achieved on 9.66 g/L cellulose. In order to test the production of butyric acid on lignocellulosic biomass, corn stover was used as the substrate, on which there was 2.37 g/L butyric acid produced under the optimized conditions. In addition, butyric acid synthesis pathway was identified involving five genes that catalyzed reactions from acetyl-CoA to butanoyl-CoA in T. fusca.     

Metabolic engineering of Saccharomyces cerevisiae to improve 1-hexadecanol production

Fatty alcohols are important components of a vast array of surfactants, lubricants, detergents, pharmaceuticals and cosmetics. We have engineered Saccharomyces cerevisiae to produce 1-hexadecanol by expressing a fatty acyl-CoA reductase (FAR) from barn owl (Tyto alba). In order to improve fatty alcohol production, we have manipulated both the structural genes and the regulatory genes in yeast lipid metabolism. The acetyl-CoA carboxylase gene (ACC1) was over-expressed, which improved 1-hexadecanol production by 56% (from 45 mg/L to 71 mg/L). Knocking out the negative regulator of the INO1 gene in phospholipid metabolism, RPD3, further enhanced 1-hexadecanol production by 98% (from 71 mg/L to 140 mg/L). The cytosolic acetyl-CoA supply was next engineered by expressing a heterologous ATP-dependent citrate lyase, which increased the production of 1-hexadecanol by an additional 136% (from 140 mg/L to 330 mg/L). Through fed-batch fermentation using resting cells, over 1.1 g/L 1-hexadecanol can be produced in glucose minimal medium, which represents the highest titer reported in yeast to date.     

Effects of redox potential control on succinic acid production by engineered Escherichia coli under anaerobic conditions

The effects of oxido-reduction potential (ORP) control on succinic acid production have been investigated in Escherichia coli LL016. In LL016, two CO₂ fixation pathways were achieved and NAD⁺ supply was enhanced by co-expression of heterologous pyruvate carboxylase (PYC) and nicotinic acid phosphoribosyltransferase (NAPRTase). During anaerobic fermentation, cell growth and metabolite distribution were changed with redox potential levels in the range of −200 to −400 mV. From the results, the ORP level of −400 mV was preferable, which resulted in the high succinic acid concentration (28.6 g/L) and high succinic acid productivity (0.33 g/L/h). Meanwhile, the yield of succinic acid at the ORP level of −400 mV was 39% higher than that at the ORP level of −200 mV. In addition, a higher NADH/NAD⁺ ratio and increased enzyme activities were also achieved by regulating the culture to a more reductive environment, which further enhanced the succinic acid production.     

Modular optimization of multi-gene pathways for fumarate production

Microbial fumarate production from renewable feedstock is a promising and sustainable alternative to petroleum-based chemical synthesis. Here, we report a modular engineering approach that systematically removed metabolic pathway bottlenecks and led to significant titer improvements in a multi-gene fumarate metabolic pathway. On the basis of central pathway architecture, yeast fumarate biosynthesis was re-cast into three modules: reduction module, oxidation module, and byproduct module. We targeted reduction module and oxidation module to the cytoplasm and the mitochondria, respectively. Combinatorially tuning pathway efficiency by constructing protein fusions RoMDH-P160A and KGD2-SUCLG2 and optimizing metabolic balance by controlling genes RoPYC, RoMDH-P160A, KGD2-SUCLG2 and SDH1 expression strengths led to significantly improved fumarate production (20.46 g/L). In byproduct module, synthetizing DNA-guided scaffolds and designing sRNA switchs enabled further production improvement up to 33.13 g/L. These results suggest that modular pathway engineering can systematically optimize biosynthesis pathways to enable an efficient production of fumarate.     

Metabolic evolution of two reducing equivalent-conserving pathways for high-yield succinate production in Escherichia coli

Reducing equivalents are an important cofactor for efficient synthesis of target products. During metabolic evolution to improve succinate production in Escherichia coli strains, two reducing equivalent-conserving pathways were activated to increase succinate yield. The sensitivity of pyruvate dehydrogenase to NADH inhibition was eliminated by three nucleotide mutations in the lpdA gene. Pyruvate dehydrogenase activity increased under anaerobic conditions, which provided additional NADH. The pentose phosphate pathway and transhydrogenase were activated by increased activities of transketolase and soluble transhydrogenase SthA. These data suggest that more carbon flux went through the pentose phosphate pathway, thus leading to production of more reducing equivalent in the form of NADPH, which was then converted to NADH through soluble transhydrogenase for succinate production. Reverse metabolic engineering was further performed in a parent strain, which was not metabolically evolved, to verify the effects of activating these two reducing equivalent-conserving pathways for improving succinate yield. Activating pyruvate dehydrogenase increased succinate yield from 1.12 to 1.31 mol/mol, whereas activating the pentose phosphate pathway and transhydrogenase increased succinate yield from 1.12 to 1.33 mol/mol. Activating these two pathways in combination led to a succinate yield of 1.5 mol/mol (88% of theoretical maximum), suggesting that they exhibited a synergistic effect for improving succinate yield.     

Metabolic engineering of Thermoanaerobacterium saccharolyticum for n-butanol production

The thermophilic anaerobe Thermoanaerobacterium saccharolyticum JW/SL-YS485 was investigated as a host for n-butanol production. A systematic approach was taken to demonstrate functionality of heterologous components of the clostridial n-butanol pathway via gene expression and enzymatic activity assays in this organism. Subsequently, integration of the entire pathway in the wild-type strain resulted in n-butanol production of 0.85 g/L from 10 g/L xylose, corresponding to 21% of the theoretical maximum yield. We were unable to integrate the n-butanol pathway in strains lacking the ability to produce acetate, despite the theoretical overall redox neutrality of n-butanol formation. However, integration of the n-butanol pathway in lactate deficient strains resulted in n-butanol production of 1.05 g/L from 10 g/L xylose, corresponding to 26% of the theoretical maximum.     

Effects of culture redox potential on succinic acid production by Corynebacterium crenatum under anaerobic conditions

During mixed-acid fermentation by Corynebacterium crenatum under anaerobic conditions, two moles of NADH are required to synthesize 1 mol of succinic acid. In this work, four controlled culture redox potentials and different carbon sources with different oxidation states were used to investigate the possibility of enhancing the succinic acid production by increasing the availability of NADH. When the culture redox potential was ?300 mV, the yield of succinic acid was 0.31 g/g, representing a 72% increase compared with the yield when the culture redox potential was ?40 mV. Meanwhile, the molar ratio of succinic acid/lactic acid increased from 0.27 to 0.48. When 0.1% neutral red was added to the acid production medium, the yield of succinic acid was 0.25 g/g, and the molar ratio of succinic acid/lactic acid was 0.38. Both values were higher than those obtained from glucose only (0.19 g/g, 0.26) or gluconate (0.05 g/g, 0.18). A higher NADH/NAD⁺ ratio and increased enzymatic activity could be achieved to enhance the succinic acid production by manipulating the culture to a more reductive environment.     

High titer mevalonate fermentation and its feeding as a building block for isoprenoids (isoprene and sabinene) production in engineered Escherichia coli

Isoprenoids are important fine chemicals as material monomers, advanced fuels and pharmaceuticals. A variety of natural isoprenoids can be synthesized by engineered microbial strains. This work established a process by dividing the current isoprenoid pathway into the upstream fermentation process, from sugar to mevalonate (MVA), and the downstream process, from MVA to the target isoprenoids. The results showed that significant differences existed in the process conditions between the upstream and downstream fermentations. After individually optimizing the process conditions, the upstream MVA production (84.0 g/L, 34.0% and 1.8 g/ L/h) and downstream isoprene production (11.0 g/L and 0.23 g/L/h) were greatly improved in this two-step process. Flask fermentation experiments also confirmed that two-step route can significantly improve the sabinene titer to 150 mg/L (6.5-fold of the sabinene titer in an earlier flask study of our lab). Therefore, the two-step route proposed in this study may have potential benefits towards the current isoprenoids production directly from glucose. The high titer and yield of MVA indicate that MVA has great potential to be more broadly utilized as starting precursor in synthetic biology.     

Metabolic engineering of Clostridium tyrobutyricum for enhanced butyric acid production from glucose and xylose

Clostridium tyrobutyricum is a promising microorganism for butyric acid production. However, its ability to utilize xylose, the second most abundant sugar found in lignocellulosic biomass, is severely impaired by glucose-mediated carbon catabolite repression (CCR). In this study, CCR in C. tyrobutyricum was eliminated by overexpressing three heterologous xylose catabolism genes (xylT, xylA and xlyB) cloned from C. acetobutylicum. Compared to the parental strain, the engineered strain Ct-pTBA produced more butyric acid (37.8 g/L vs. 19.4 g/L) from glucose and xylose simultaneously, at a higher xylose utilization rate (1.28 g/L·h vs. 0.16 g/L·h) and efficiency (94.3% vs. 13.8%), resulting in a higher butyrate productivity (0.53 g/L·h vs. 0.26 g/L·h) and yield (0.32 g/g vs. 0.28 g/g). When the initial total sugar concentration was ~120 g/L, both glucose and xylose utilization rates increased with increasing their respective concentration or ratio in the co-substrates but the total sugar utilization rate remained almost unchanged in the fermentation at pH 6.0. Decreasing the pH to 5.0 significantly decreased sugar utilization rates and butyrate productivity, but the effect was more pronounced for xylose than glucose. The addition of benzyl viologen (BV) as an artificial electron carrier facilitated the re-assimilation of acetate and increased butyrate production to a final titer of 46.4 g/L, yield of 0.43 g/g sugar consumed, productivity of 0.87 g/L·h, and acid purity of 98.3% in free-cell batch fermentation, which were the highest ever reported for butyric acid fermentation. The engineered strain with BV addition thus can provide an economical process for butyric acid production from lignocellulosic biomass.