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Appl Environ Microbiol. Dec 2005; 71(12): 7866–7871.
PMCID: PMC1317483

Cofactor Dependence in Furan Reduction by Saccharomyces cerevisiae in Fermentation of Acid-Hydrolyzed Lignocellulose

Abstract

A decreased fermentation rate due to inhibition is a significant problem for economic conversion of acid-pretreated lignocellulose hydrolysates to ethanol, since the inhibition gives rise to a requirement for separate detoxification steps. Together with acetic acid, the sugar degradation products furfural and 5-hydroxymethyl furfural are the inhibiting compounds found at the highest concentrations in hydrolysates. These aldehydes have been shown to affect both the specific growth rate and the rate of fermentation by yeast. Two strains of Saccharomyces cerevisiae with different abilities to ferment inhibiting hydrolysates were evaluated in fermentations of a dilute acid hydrolysate from spruce, and the reducing activities for furfural and 5-hydroxymethyl furfural were determined. Crude cell extracts of a hydrolysate-tolerant strain (TMB3000) converted both furfural and 5-hydroxymethyl furfural to the corresponding alcohol at a rate that was severalfold higher than the rate observed for cell extracts of a less tolerant strain (CBS 8066), thereby confirming that there is a correlation between the fermentation rate in a lignocellulosic hydrolysate and the bioconversion capacity of a strain. The in vitro NADH-dependent furfural reduction capacity of TMB3000 was three times higher than that of CBS 8066 (1,200 mU/mg protein and 370 mU/mg protein, respectively) in fed-batch experiments. Furthermore, the inhibitor-tolerant strain TMB3000 displayed a previously unknown NADH-dependent reducing activity for 5-hydroxymethyl furfural (400 mU/mg protein during fed-batch fermentation of hydrolysates). No corresponding activity was found in strain CBS 8066 (<2 mU/mg). The ability to reduce 5-hydroxymethyl furfural is an important characteristic for the development of yeast strains with increased tolerance to lignocellulosic hydrolysates.

Ethanol produced from renewable resources, such as lignocellulose, is considered an attractive alternative to partially replace fossil fuels (1). Lignocellulosic materials, including wood, forest, and agricultural residues, can potentially be used for ethanol production (6). Prior to fermentation cellulose and hemicellulose are converted to monomeric sugars by a combination of physical, chemical, and enzymatic processes (6). In these processes inhibitory compounds, including carboxylic acids, furans, and phenolic compounds, are formed (8, 9, 16, 18, 19). The inhibitors can reduce both the growth rate and the ethanol productivity (17).

Saccharomyces cerevisiae is the preferred microorganism in the ethanol fermentation industry. This yeast species is more tolerant to inhibitors, such as acetic acid, furfural, and 5-hydroxymethyl furfural (HMF), than other candidate microorganisms, such as Escherichia coli, Zymomonas mobilis, Pichia stipitis, and Candida shehatae (15). Strains of S. cerevisiae exhibit significant differences in fermentative capacity and tolerance to lignocellulose-derived inhibitors (11). The tolerance to aldehyde compounds is most likely due to the ability of microorganisms to convert these compounds to the corresponding less inhibitory alcohols (20, 23). Dilute-acid hydrolysates are often strongly inhibiting and cannot be fermented in batch mode, but they may be fermented by S. cerevisiae without prior detoxification in fed-batch mode (13, 14, 21). In fed-batch operation inhibitors are maintained at low levels because of their continuous conversion to less toxic compounds. Fed-batch operation also permits the simultaneous uptake of different monomeric sugars because the sugar concentration is maintained at a low level and saturation of uptake systems and/or the glycolytic flux is avoided.

The objective of this study was to quantitatively identify biochemical traits that are responsible for inhibitor tolerance in S. cerevisiae and that allow high specific ethanol productivity in fed-batch fermentation of nondetoxified hydrolysates. The capacity for in situ detoxification of furans is presumed to limit the maximum specific productivity of acid-hydrolyzed lignocellulose conversion. In the present work, two strains of S. cerevisiae, one highly tolerant to lignocellulose hydrolysates and the other a laboratory reference strain, were investigated using batch and fed-batch fermentations of hydrolysates, as well as chemostat cultures with synthetic medium. Identification of characteristics that differ between strains with different types of inhibitor tolerance can be used for the design of genetically modified strains.

MATERIALS AND METHODS

Strains and inoculum cultivation.

Two strains of S. cerevisiae were used. Strain TMB3000 (=ATCC 96581), originally isolated from a spent sulfite liquor fermentation plant (designated “Isolate 3” in reference 10), is known to be robust in lignocellulosic hydrolysates or model inhibiting media (7, 11). The widely used diploid laboratory strain CBS 8066 (22) was used as a reference. The strains were maintained on agar plates containing 10 g/liter yeast extract, 20 g/liter soy peptone, 20 g/liter agar, and 20 g/liter glucose in ultrapure water obtained froma Maxima water purification unit (Elga Labwater, High Wycombe, United Kingdom). Inoculum cultures were grown in 300-ml cotton-plugged unbaffled shake flasks with 100 ml of synthetic medium with 15 g/liter glucose as a carbon and energy source. The medium contained (per liter) 7.5 g (NH4)SO4, 3.5 g KH2PO4, 0.75 g MgSO4 · 7H2O, 30 mg EDTA, 9 mg CaCl2 · 2H2O, 9 mg ZnSO4· 7H2O, 6 mg FeSO4 · 7H2O, 2 mg H3BO3, 1.6 mg MnCl2 · 2H2O, 0.8mgNa2MoO4 · 2H2O, 0.6 mg CoCl2 · 2H2O, 0.6 mg CuSO4 · 5H2O, 0.2 mg KI,50μg d-biotin, 0.2 mg p-aminobenzoic acid, 1 mg nicotinic acid, 1 mg calcium pantothenate, 1 mg pyridoxine HCl, 1 mg thiamine HCl, 25 mg m-inositol, 10 mg ergosterol, and 420 mg Tween 80. The inoculum cultures were grown for 24 h at 30°C on a shaker set at 150 rpm. Twenty milliliters of inoculum was added to a fermentor to start a cultivation.

Hydrolysate medium.

The hydrolysate used was produced from forest residue, primarily spruce, in a two-stage dilute-acid hydrolysis process with sulfuric acid as the catalyst (13). The hydrolysates from the two stages were mixed and stored at 8°C until they were used. The resulting hydrolysate contained 16 g/liter glucose, 13 g/liter mannose, 3.2 g/liter galactose, 6.1 g/liter xylose, 1.1 g/liter arabinose, 1.5 g/liter acetic acid, 1.6 g/liter HMF, and 0.2 g/liter furfural.

Batch and fed-batch fermentations.

Batch and fed-batch fermentations were performed in a 3.3-liter BioFlo III bioreactor (New Brunswick Scientific, Edison, NJ). The stirring rate was 400 rpm, and the fermentor was sparged continuously with 600 or 1,000 ml/min N2 gas [O2 concentration, <5 ppm; ADR class 2, 1(a); AGA, Sundbyberg, Sweden]. The pH was maintained at 5.0 with 2.0 M NaOH. All experiments began with an initial batch phase in 1 liter of synthetic medium containing 50 g glucose as the carbon source. The concentrations of the other medium components were tripled compared to the concentrations used for the inoculum to compensate for dilution during the fed-batch operation. Hydrolysate feeding began when glucose was depleted (i.e., when the CO2 evolution rate [CER] was less than 1 mmol/h).

Two types of fermentation experiments were conducted. In the first type, hydrolysate was added to the reactor using the maximum feed rate of the medium pump (~2 liters/h) after the initial batch cultivation. This protocol is referred to below as “batch” fermentation. In the second type, referred to as “fed-batch” fermentation, the hydrolysate feed rate was controlled using a previously described step-response method (13), in which the feed rate was adjusted in a step-wise manner, where each step was proportional to the derivative of the measured CO2 evolution rate from the previous step. The feed rate was controlled with a peristaltic pump (Watson-Marlow Alitea AB, Stockholm, Sweden). In both types of fermentation the total volume of hydrolysate added was 1.5 liters.

Experiments with nongrowing cells.

Nongrowing cells were studied in fed-batch fermentations. Cycloheximide was added to a final concentration of 10 mg/liter immediately before addition of the hydrolysate. This concentration completely inhibits protein synthesis by S. cerevisiae (5).

Continuous culture.

Continuous culture was used to evaluate the induction of furan-reducing activity. Synthetic medium in which the concentrations of all medium components were increased by 33% was used. The glucose concentration was 20 g/liter. In some experiments, 0.5 g/liter of HMF was added to the medium. The liquid volume in the reactor (Belach BR 0.5 bioreactor; Belach Bioteknik AB, Solna, Sweden) was 500 ml. After complete consumption of glucose in the batch phase, feeding was started at a dilution rate of 0.1 h−1. The reactor was sparged with 300 ml of N2/min. The pH was maintained at 5.0 with 0.75 M NaOH, and the temperature was 30°C. The stirrer speed was set at 500 rpm. To ensure steady-state conditions, samples were not taken until after a minimum of five residence times after the start of feeding or a change in the medium composition.

Off-gas analysis.

A gas monitor (model 1311; Brüel and Kjaer, Naerum, Denmark) was used to measure the CO2 evolution rate in batch and fed-batch experiments. The gas analyzer had three channels for measurement of CO2, O2, and ethanol in the off-gas from the reactor. The ethanol in the gas phase was assumed to be in equilibrium with the ethanol in the broth, and the ethanol signal was calibrated against ethanol concentrations measured in the broth by high-performance liquid chromatography. Calibration for O2 and CO2 was done using a gas containing 20% O2 and 5% CO2.

Biomass.

A flow injection analysis system (2) was used to measure the biomass concentration, which was expressed as the optical density at 610 nm (OD610), hourly. After every fermentation, the flow injection analysis system signal was calibrated against the measured dry weight. Duplicate 10-ml samples of fermentation broth were centrifuged (2,000 × g, 3 min, room temperature) in tared tubes. Cells were washed with distilled water, pelleted again by centrifugation, and dried overnight at 105°C before they were weighed. Dry weight was measured three times during each fermentation (just before addition of hydrolysate, during fermentation of hydrolysate, and at the end of each fermentation).

Metabolite concentrations.

Samples for analysis of metabolite concentrations were taken regularly from the reactor. The samples were centrifuged (10,000 × g,3min, room temperature) and filtered through 0.2-μm filters. The concentrations of glucose, mannose, galactose, and arabinose were measured on an Aminex HPX-87P column (Bio-Rad, Hercules, Calif.) at 80°C. The mobile phase was distilled water at a flow rate of 0.6 ml/min. The precolumns used were Deashing refill cartridges (Bio-Rad). The concentrations of ethanol, HMF, furfural, glycerol, and acetic acid were measured on an Aminex HPX-87H column (Bio-Rad) at 65°C with 5 mM sulfuric acid as the mobile phase (0.6 ml/min) and with Cation-H refill cartridges (Bio-Rad) as precolumns. Most compounds were detected with a refractive index detector; the exceptions were HMF and furfural, which were detected with a UV detector (210 nm).

To compensate for ethanol that evaporated during fermentation, the mole fraction of ethanol in the gas phase was assumed to be proportional to the mole fraction of ethanol in the liquid phase. Thus, the total amount of evaporated ethanol could be estimated by integration of the gas flow leaving the reaction multiplied by the mole fraction of ethanol in the gas (14).

Preparation of cell extracts.

Cell extracts were prepared for measurement of enzyme activities using Y-PER reagent (Pierce, Rockford, IL). Cell extracts were kept at −80°C until they were used. The protein content was determined with Coomassie protein assay reagent using bovine serum albumin as a standard (Pierce). The cell-free preparations had an average protein concentration of 19g/liter.

Measurement of furfural- and HMF-reducing activities.

Furfural- and HMF-reducing activities were measured in cell extracts as previously described (24). The measurements were performed in 100 mM phosphate buffer (50 mM KH2PO4, 50 mM K2HPO4; pH 7.0) with 2 μl of cell extract. Furfural was added to a concentration of 10 mM. The samples were heated to 30°C, and the reaction was started by adding NADH to a concentration of 100 μM. The oxidation of NADH was monitored by determining the change in A340. The same procedure was repeated with NADPH (100 μM) as the cofactor but with 20 μl of cell extract.

HMF-reducing activity was measured by the same procedure. The concentration of HMF was 10 mM, and 20 μl of cell extract was used. The activity was measured with both NADH and NADPH as cofactors.

Measurement of ADH activity.

Alcohol dehydrogenase (ADH) activity was measured in cell extracts as previously described (3). Ethanol was added to a final concentration of 100 mM in 100 mM phosphate buffer (pH 7.0), and 2 μl of cell extract was used. After the mixture was heated to 30°C, the reaction was started by adding NAD+ to a final concentration of 100 μM. The reduction of NAD+ was monitored by determining the change in A340 (3).

RESULTS

Batch fermentation.

Both strains were able to ferment the dilute acid hydrolysate used. However, there were significant differences between the two strains, particularly with respect to fermentation rates. This could be seen from the CER, which was directly coupled to the fermentation rate (Fig. (Fig.1A).1A). For CBS 8066 a gradual decrease in the CER was observed during fermentation of the hydrolysate. In contrast, TMB3000 had a more constant CER during the course of the fermentation. Furthermore, the concentration of HMF decreased much faster for TMB3000 than it did for CBS 8066 (Fig. (Fig.1B).1B). The concentration of furfural was very low in both fermentation experiments (less than 0.05 g/liter). The average specific ethanol productivity, calculated from the formation of ethanol and the amount of biomass in the reactor (Fig. (Fig.1C),1C), for strain CBS 8066 (0.13 g g−1 h−1) was lower than that for TMB3000 (0.36 g g−1 h−1). In addition, the specific ethanol productivity gradually decreased throughout the fermentation for CBS 8066, although the hexose sugars glucose and mannose were completely consumed at the end of the batch fermentation. Neither strain could grow in batch culture on the dilute acid hydrolysate (Fig. (Fig.1C1C).

FIG. 1.
Batch fermentation of dilute acid lignocellulose hydrolysate. After an initial batch cultivation on synthetic medium, 1.5 liters of hydrolysate was added to the reactor at the maximal rate (at time zero). (A) CER for batch fermentation with TMB3000 and ...

Fed-batch fermentation.

The fermentation rates were higher in the fed-batch experiments than in the batch experiments for both strains, as observed from both the CER and ethanol measurements (Fig. (Fig.2A).2A). A gradual decrease in the CER occurred with time for CBS 8066, but not for TMB3000. The average specific ethanol productivities were 0.30 g g−1 h−1 and 0.61 g g−1 h−1 for CBS 8066 and TMB3000, respectively. The concentrations of HMF were maintained at much lower levels for TMB3000 than for CBS 8066 (Fig. (Fig.2A).2A). No furfural was detected in any of the fed-batch fermentation experiments, so the conversion rate for furfural was the same as the addition rate for furfural. TMB3000 could grow in hydrolysate with an average specific growth rate of ~0.12 h−1 during the feed phase, but no growth was observed for CBS 8066 (Fig. (Fig.2A2A).

FIG. 2.
Fed-batch fermentation of dilute-acid lignocellulose hydrolysate with and without addition of cycloheximide. After an initial batch phase on synthetic medium, the fed-batch phase was started (time zero); 1.5 liters of hydrolysate was added according to ...

When cycloheximide (25 mg/liter) was added immediately before the start of the fed-batch phase, the CER curve for TMB3000 changed drastically (Fig. (Fig.2B)2B) compared to the curve for the fed-batch fermentation in which no cycloheximide had been added (Fig. (Fig.2A).2A). The CER for TMB3000 during fed-batch fermentation with cycloheximide was similar to the CER for CBS 8066 during fed-batch fermentation without cycloheximide (Fig. (Fig.2A).2A). The CER profile for the fed-batch fermentation of CBS 8066 was affected less by addition of cycloheximide than the CER profile for the fed-batch fermentation of TMB3000 was (Fig. (Fig.2B).2B). Furan conversion also continued inthe absence of growth. Complete furfural reduction was maintained; i.e., the concentration was less than the detection level, but the HMF concentrations increased somewhat compared to those in cycloheximide-free medium, particularly for CBS 8066 (Fig. (Fig.2B).2B). Some biomass formation was measured during the first hour of the fed-batch fermentations (Fig. (Fig.2B),2B), which probably resulted from increased cell size rather than increased cell number, since cycloheximide (10 mg/liter) inhibits protein synthesis in S. cerevisiae within a few minutes (5).

The average furfural-reducing activity with NADH as the cofactor for TMB3000 was three times that for CBS 8066, and with NADPH as the cofactor it was five times higher for fed-batch grown cells (Table (Table1).1). The largest difference was observed for HMF-reducing activity with NADH as the cofactor. This activity was very low in CBS 8066 but 200-fold higher in TMB3000 (Table (Table1).1). With NADPH as the cofactor, the activities differed by a factor of three. Induction of the reducing activity caused by the presence of the inhibitors in the hydrolysate was assessed by comparing the enzyme activities during feeding of hydrolysate and the activities in stationary cells. The inductions with the clearest significance were found for NADPH-coupled reduction of both furfural and HMF for TMB3000 (Table (Table1).1). To further investigate the differences between furan-reducing activities, the strains were grown in continuous cultures with synthetic media. The furan-reducing activities were determined at steady states after feeding of media with and without added HMF (0.5 g/liter). The difference in furan-reducing activities between the strains measured during fed-batch fermentation of hydrolysate remained in the continuous fermentation (Table (Table1).1). The furan-reducing activities were 4 to 14 times higher for TMB3000 than for CBS 8066; the only exception was reduction of HMF with NADH as the cofactor, in which the activity was more than 300 times higher, which is in good qualitative agreement with the fed-batch results. The presence of HMF in the medium did not cause a significant increase in any of the activities measured.

TABLE 1.
Furfural and HMF reduction activities in crude cell extracts from CBS 8066 and TMB3000 when different cofactors were used

A maximum reduction capacity could be calculated from the in vitro measurements of furan reduction, and this value could be compared to the measured in vivo conversion rates during the fermentations (Table (Table2).2). The calculated maximum reduction rates (based on the sum of NADH and NADPH coupled activities) were found to be 0.67 g furfural g biomass−1 h−1 for CBS 8066 and 2.0 g g−1 h−1 for TMB3000. The in vitro activity for furfural reduction by strain CBS 8066 agrees well with the maximum conversion rate in synthetic media reported for CBS 8066 (0.6 g g−1 h−1) (20). These values are much higher than the fed-batch conversion rates that we measured. The concentration of furfural also remained below the detection limit during the fermentation, further strengthening the conclusion that the conversion of furfural did not limit the feed rate. The measured in vitro enzyme activity for HMF conversion in CBS 8066 (0.033 g g biomass−1 h−1) was very close to the in vivo conversion rate (0.034 g g biomass−1 h−1) in hydrolysate (Table (Table2).2). The HMF conversion rate was somewhat lower than that previously reported for synthetic media (0.14 g g biomass−1 h−1) (19). For TMB3000 the in vitro enzyme activity was 0.92 g g biomass−1 h−1.

TABLE 2.
Measured average in vivo rates of specific conversion of furfural and HMF and in vitro enzyme activitiesa

Furfural reduction has been attributed to the enzyme ADH (4, 12). The average ADH activities during fed-batch fermentation were 1,000 mU/mg for TMB3000 and 620 mU/mg for CBS 8066. The difference is smaller than that observed for either furfural conversion or HMF conversion in the strains (Table (Table1).1). However, at the end of the fed-batch phase the ADH activity was three times higher for TMB3000 than for CBS 8066.

DISCUSSION

One characteristic presumed to be responsible for inhibitor tolerance in yeasts is the ability to reduce toxic aldehydes, such as the furans furfural and HMF. We found that an inhibitor-tolerant yeast strain (TMB3000) had a higher capacity for reduction of furan compounds than a less tolerant strain (CBS 8066) and that the cofactor specificities for the conversion of HMF were different. The combination of higher conversion activity and an NADH-coupled HMF reduction suffices to explain the higher ethanol productivity of TMB3000 in fed-batch fermentations of lignocellulosic hydrolysates. Higher reducing activity enables higher rates of conversion of furfural and HMF and possibly also other inhibitory aldehydes (10), which ensures that the concentrations of these inhibitors in the fermentations remain low.

We hypothesize that there are two ways to inhibit the growth of S. cerevisiae during fermentation of undetoxified lignocellulose hydrolysate. First, growth may be inhibited if the cells are exposed to high levels of inhibitors (for example, in batch fermentation). This type of inhibition occurred for both CBS 8066 and TMB3000 during batch fermentation of the lignocellulose hydrolysate, when neither strain could grow. Growth also may be inhibited if the reduction of the inhibitors competes for cofactors needed for growth. We found that HMF reduction in CBS 8066 occurs with NADPH as a cofactor. An NADPH-coupled reduction diverts NADPH from anabolic reactions, whereas an NADH-coupled reduction, like that found in TMB3000, does not directly interfere with the NADPH balance.

The measured enzyme activities for furan conversion by CBS 8066 in the present study were similar to those previously reported for strain TMB3001, which was derived from CEN.PK113-7A (24). The average activities for furfural conversion in CBS 8066 were 370 mU/mg protein with NADH as the cofactor and 20 mU/mg protein with NADPH as the cofactor, compared to 490 and 22 mU/mg protein, respectively, for TMB3001. For HMF conversion, the average activities for CBS 8066 were 1.4 mU/mg protein (NADH) and 12 mU/mg protein (NADPH), compared with 2.2 and 22 mU/mg protein, respectively, for TMB3001 (24).

The furan reduction activities were substantially higher in strain TMB3000. There was considerable furan reduction activity in the cell extracts of both strains before the cells were exposed to the hydrolysate (i.e., after the growth phase on synthetic batch medium). The NADPH-coupled reduction activities for both furfural and HMF were slightly increased during prolonged exposure to hydrolysate for TMB3000, even though the enzymes catalyzing the NADH-coupled conversion of HMF in TMB3000 were expressed during growth on glucose, based on the chemostat experiments with synthetic medium.

During fermentation of a toxic hydrolysate it is essential to maintain at least some cell growth. Ethanol productivity decreases during fermentation if growth is completely inhibited, as shown in cycloheximide addition experiments. For growth to continue, the concentration of inhibitors must be kept low, which can be achieved by in situ detoxification during fed-batch fermentation. The strain used also must have a high inhibitor conversion activity (e.g., reduction of furans) to enable high productivity. An NADPH-coupled reduction is likely to have a negative effect on growth and on ethanol productivity. Thus, the NADH-coupled reduction of HMF that we identified is potentially an important key to increasing ethanol productivity by fermentation of acid-hydrolyzed lignocellulose.

Acknowledgments

This work was financially supported by the Swedish Energy Agency.

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