Michael A. Cotta

Dilute acid pretreatment, enzymatic saccharification, and fermentation of rice hulls to ethanol.

Rice hulls, a complex lignocellulosic material with high lignin (15.38 ± 0.2%) and ash (18.71 ± 0.01%) content, contain 35.62 ± 0.12% cellulose and 11.96 ± 0.73% hemicellulose and has the potential to serve as a low‐cost feedstock for production of ethanol. Dilute H2SO4 pretreatments at varied temperature (120–190 °C) and enzymatic saccharification (45 °C, pH 5.0) were evaluated for conversion of rice hull cellulose and hemicellulose to monomeric sugars. The maximum yield of monomeric sugars from rice hulls (15%, w/v) by dilute H2SO4 (1.0%, v/v) pretreatment and enzymatic saccharification (45 °C, pH 5.0, 72 h) using cellulase, β‐glucosidase, xylanase, esterase, and Tween 20 was 287 ± 3 mg/g (60% yield based on total carbohydrate content). Under this condition, no furfural and hydroxymethyl furfural were produced. The yield of ethanol per L by the mixed sugar utilizing recombinant Escherichia coli strain FBR 5 from rice hull hydrolyzate containing 43.6 ± 3.0 g fermentable sugars (glucose, 18.2 ± 1.4 g; xylose, 21.4 ± 1.1 g; arabinose, 2.4 ± 0.3 g; galactose, 1.6 ± 0.2 g) was 18.7 ± 0.6 g (0.43 ± 0.02 g/g sugars obtained; 0.13 ± 0.01 g/g rice hulls) at pH 6.5 and 35 °C. Detoxification of the acid‐ and enzyme‐treated rice hull hydrolyzate by overliming (pH 10.5, 90 °C, 30 min) reduced the time required for maximum ethanol production (17 ± 0.2 g from 42.0 ± 0.7 g sugars per L) by the E. coli strain from 64 to 39 h in the case of separate hydrolysis and fermentation and increased the maximum ethanol yield (per L) from 7.1 ± 2.3 g in 140 h to 9.1 ± 0.7 g in 112 h in the case of simultaneous saccharification and fermentation.

Ethanol Production from Alkaline Peroxide Pretreated Enzymatically Saccharified Wheat Straw

Wheat straw used in this study contained 44.24 ± 0.28% cellulose and 25.23 ± 0.11% hemicellulose. Alkaline H2O2 pretreatment and enzymatic saccharification were evaluated for conversion of wheat straw cellulose and hemicellulose to fermentable sugars. The maximum yield of monomeric sugars from wheat straw (8.6%, w/v) by alkaline peroxide pretreatment (2.15% H2O2, v/v; pH 11.5; 35 °C; 24 h) and enzymatic saccharification (45 °C, pH 5.0, 120 h) by three commercial enzyme preparations (cellulase, β‐glucosidase, and xylanase) using 0.16 mL of each enzyme preparation per g of straw was 672 ± 4 mg/g (96.7% yield). During the pretreatment, no measurable quantities of furfural and hydroxymethyl furfural were produced. The concentration of ethanol (per L) from alkaline peroxide pretreated enzyme saccharified wheat straw (66.0 g) hydrolyzate by recombinant Escherichia coli strain FBR5 at pH 6.5 and 37 °C in 48 h was 18.9 ± 0.9 g with a yield of 0.46 g per g of available sugars (0.29 g/g straw). The ethanol concentration (per L) was 15.1 ± 0.1 g with a yield of 0.23 g/g of straw in the case of simultaneous saccharification and fermentation by the E. coli strain at pH 6.0 and 37 °C in 48 h.

Butanol Production from Corn Fiber Xylan Using Clostridium acetobutylicum

Acetone, butanol, and ethanol (ABE) were produced from corn fiber arabinoxylan (CFAX) and CFAX sugars (glucose, xylose, galactose, and arabinose) using Clostridium acetobutylicum P260. In mixed sugar (glucose, xylose, galactose, and arabinose) fermentation, the culture preferred glucose and arabinose over galactose and xylose. Under the experimental conditions, CFAX (60 g/L) was not fermented until either 5 g/L xylose or glucose plus xylanase enzyme were added to support initial growth and fermentation. In this system, C. acetobutylicum produced 9.60 g/L ABE from CFAX and xylose. This experiment resulted in a yield and productivity of 0.41 and 0.20 g/L·h, respectively. In the integrated hydrolysis, fermentation, and recovery process, 60 g/L CFAX and 5 g/L xylose produced 24.67 g/L ABE and resulted in a higher yield (0.44) and a higher productivity (0.47 g/L·h). CFAX was hydrolyzed by xylan‐hydrolyzing enzymes, and ABE were recovered by gas stripping. This investigation demonstrated that integration of hydrolysis of CFAX, fermentation to ABE, and recovery of ABE in a single system is an economically attractive process. It is suggested that the culture be further developed to hydrolyze CFAX and utilize all xylan sugars simultaneously. This would further increase productivity of the reactor.

Expression of a heterologous xylose transporter in a Saccharomyces cerevisiae strain engineered to utilize xylose improves aerobic xylose consumption

The goal of this investigation was to determine the effect of a xylose transport system on glucose and xylose co-consumption as well as total xylose consumption in Saccharomyces cerevisiae. We expressed two heterologous transporters from Arabidopsis thaliana in recombinant xylose-utilizing S. cerevisiae cells. Strains expressing the heterologous transporters were grown on glucose and xylose mixtures. Sugar consumption rates and ethanol concentrations were determined and compared to an isogenic control strain lacking the A. thaliana transporters. Expression of the transporters increased xylose uptake and xylose consumption up to 46% and 40%, respectively. Xylose co-consumption rates (prior to glucose depletion) were also increased by up to 2.5-fold compared to the control strain. Increased xylose consumption correlated with increased ethanol concentration and productivity. During the xylose/glucose co-consumption phase, strains expressing the transporters had up to a 70% increase in ethanol production rate. It was concluded that in these strains, xylose transport was a limiting factor for xylose utilization and that increasing xylose/glucose co-consumption is a viable strategy for improving xylose fermentation.

Comparison of pretreatment strategies for enzymatic saccharification and fermentation of barley straw to ethanol

Barley straw used in this study contained 34.3% cellulose, 23.0% hemicellulose and 13.3% lignin (moisture, 6.5%). Several pretreatments (dilute acid, lime and alkaline peroxide) and enzymatic saccharification procedures were evaluated for the conversion of barley straw to monomeric sugars. The maximum release of sugars (glucose, 384 mg; xylose, 187 mg; arabinose, 32 mg; total sugars, 604 mg/g; 94% of maximum theoretical sugar yield) from barley straw (10%, w/v) was obtained by alkaline peroxide (2.5% H(2)O(2), pH 11.5) pretreatment (35 degrees C, 24 hours) and enzymatic saccharification (45 degrees C, pH 5.0, 120 hours) after diluting 2 times before adding a cocktail of three commercial enzyme preparations (cellulase, beta-glucosidase and hemicellulase) each at the dose level of 0.15 ml/g of straw. Dilute acid and lime pretreatments followed by enzymatic saccharification generated 566 mg (88% yield) and 582 mg (91% yield) total sugars/g of barley straw, respectively. The yield of ethanol from the dilute acid pretreated and enzymatically saccharified barley straw hydrolyzate (23.7g sugars/L) was 11.4g/L (0.48g/g available sugars, 0.26g/g straw) by the mixed sugar utilizing recombinant Escherichia coli strain FBR5 in 17 hours. The ethanol yields were 11.4 and 11.9g/L from 24.4 and 26.2g sugars/L obtained from lime and alkaline peroxide pretreated barley straw, respectively. No inhibition of fermentation occurred by any of the three pretreatments under the conditions used.

Carbon dioxide addition to microbial fuel cell cathodes maintains sustainable catholyte pH and improves anolyte pH, alkalinity, and conductivity.

Bioelectrochemical system (BES) pH imbalances develop due to anodic proton-generating oxidation reactions and cathodic hydroxide-ion-generating reduction reactions. Until now, workers added unsustainable buffers to reduce the pH difference between the anode and cathode because the pH imbalance contributes to BES potential losses and, therefore, power losses. Here, we report that adding carbon dioxide (CO(2)) gas to the cathode, which creates a CO(2)/bicarbonate buffered catholyte system, can diminish microbial fuel cell (MFC) pH imbalances in contrast to the CO(2)/carbonate buffered catholyte system by Torres, Lee, and Rittmann [Environ. Sci. Technol. 2008, 42, 8773]. We operated an air-cathode and liquid-cathode MFC side-by-side. For the air-cathode MFC, CO(2) addition resulted in a stable catholyte film pH of 6.61 +/- 0.12 and a 152% increase in steady-state power density. By adding CO(2) to the liquid-cathode system, we sustained a steady catholyte pH (pH = 5.94 +/- 0.02) and a low pH imbalance (DeltapH = 0.65 +/- 0.18) over a 2-week period without external salt buffer addition. By migrating bicarbonate ions from the cathode to the anode (with an anion-exchange membrane), we increased the anolyte pH (DeltapH = 0.39 +/- 0.31), total alkalinity (494 +/- 6 to 582 +/- 6 as mg CaCO(3)/L), and conductivity (1.53 +/- 0.49 to 2.16 +/- 0.03 mS/cm) relative to the feed properties. We also verified with a phosphate-buffered MFC that our reaction rates were limited mainly by the reactor configuration rather than limitations due to the bicarbonate buffer.

Aerated Shewanella oneidensis in continuously fed bioelectrochemical systems for power and hydrogen production.

We studied the effects of aeration of Shewanella oneidensis on potentiostatic current production, hydrogen production in a microbial electrolysis cell, and electric power generation in a microbial fuel cell (MFC). The potentiostatic performance of aerated S. oneidensis was considerably enhanced to a maximum current density of 0.45 A/m2 or 80.3 A/m3 (mean: 0.34 A/m2, 57.2 A/m3) compared to anaerobically grown cultures. Biocatalyzed hydrogen production rates with aerated S. oneidensis were studied within the applied potential range of 0.3–0.9 V and were highest at 0.9 V with 0.3 m3 H2/m3 day, which has been reported for mixed cultures, but is ∼10 times higher than reported for an anaerobic culture of S. oneidensis. Aerated MFC experiments produced a maximum power density of 3.56 W/m3 at a 200‐Ω external resistor. The main reasons for enhanced electrochemical performance are higher levels of active biomass and more efficient substrate utilization under aerobic conditions. Coulombic efficiencies, however, were greatly reduced due to losses of reducing equivalents to aerobic respiration in the anode chamber. The next challenge will be to optimize the aeration rate of the bacterial culture to balance between maximization of bacterial activation and minimization of aerobic respiration in the culture. Biotechnol. Bioeng. 2010;105: 880–888.

Fermentation of bioenergy crops into ethanol using biological abatement for removal of inhibitors

Biological abatement was used to condition dilute acid-pretreated hydrolysates of three perennial herbaceous crops that are potential bioenergy feedstocks: switchgrass, reed canarygrass, and alfalfa stems. Fungal isolate Coniochaeta ligniaria was inoculated into the hydrolysates to metabolize and remove inhibitory compounds prior to yeast fermentation of glucose. Switchgrass, reed canarygrass, and alfalfa stem samples were pretreated with dilute acid at 10% w/w biomass loading and subjected to bioabatement with strain NRRL30616, to prepare the material for simultaneous saccharification of cellulose and fermentation by Saccharomyces cerevisiae. Bioabatement eliminated the extended fermentation lag times associated with inhibitory compounds and observed for the unconditioned biomass hydrolysates controls. Bioabatement was as effective as lime conditioning at reducing fermentation lag times. Prolonged incubations with the bioabatement microbe resulted in consumption of some glucose and reduced production of ethanol.

Response surface optimization of corn stover pretreatment using dilute phosphoric acid for enzymatic hydrolysis and ethanol production.

Dilute H(3)PO(4) (0.0-2.0%, v/v) was used to pretreat corn stover (10%, w/w) for conversion to ethanol. Pretreatment conditions were optimized for temperature, acid loading, and time using central composite design. Optimal pretreatment conditions were chosen to promote sugar yields following enzymatic digestion while minimizing formation of furans, which are potent inhibitors of fermentation. The maximum glucose yield (85%) was obtained after enzymatic hydrolysis of corn stover pretreated with 0.5% (v/v) acid at 180°C for 15min while highest yield for xylose (91.4%) was observed from corn stover pretreated with 1% (v/v) acid at 160°C for 10min. About 26.4±0.1g ethanol was produced per L by recombinant Escherichia coli strain FBR5 from 55.1±1.0g sugars generated from enzymatically hydrolyzed corn stover (10%, w/w) pretreated under a balanced optimized condition (161.81°C, 0.78% acid, 9.78min) where only 0.4±0.0g furfural and 0.1±0.0 hydroxylmethyl furfural were produced.

Shewanella oneidensis in a lactate-fed pure-culture and a glucose-fed co-culture with Lactococcus lactis with an electrode as electron acceptor

Bioelectrochemical systems (BESs) employing mixed microbial communities as biocatalysts are gaining importance as potential renewable energy, bioremediation, or biosensing devices. While we are beginning to understand how individual microbial species interact with an electrode as electron donor, little is known about the interactions between different microbial species in a community: sugar fermenting bacteria can interact with current producing microbes in a fashion that is either neutral, positively enhancing, or even negatively affecting. Here, we compare the bioelectrochemical performance of Shewanella oneidensis in a pure-culture and in a co-culture with the homolactic acid fermenter Lactococcus lactis at conditions that are pertinent to conventional BES operation. While S. oneidensis alone can only use lactate as electron donor for current production, the co-culture is able to convert glucose into current with a comparable coulombic efficiency of ∼17%. With (electro)-chemical analysis and transcription profiling, we found that the BES performance and S. oneidensis physiology were not significantly different whether grown as a pure- or co-culture. Thus, the microbes worked together in a purely substrate based (neutral) relationship. These co-culture experiments represent an important step in understanding microbial interactions in BES communities with the goal to design complex microbial communities, which specifically convert target substrates into electricity.