Patent Application
20090061502
The present invention relates to a Saccharomyces cerevisiae strain being able to grow and produce ethanol in the presence of inhibiting compounds and substances, in particular furfural and derivatives thereof while reducing such compounds, and in particular it relates to a strain that is able to produce ethanol in a fed-batch or continuous production system.
Because of its low net contribution to the production of carbon dioxide, ethanol produced from renewable resources, such as lignocellulose, is considered an attractive alternative for partly replacing fossil fuels (1). Many sources of lignocellulosic materials (e.g. wood, forest residues and agricultural residues) can potentially be used for ethanol production (2). Prior to fermentation, however, the cellulose and the hemicellulose in the lignocellulose must be converted to monomeric sugars by a combination of physical (e.g. grinding, steam explosion), chemical (e.g. dilute acid) and perhaps also enzymatic treatments (2). In addition to monomeric sugars also a number of other compounds are formed during these processes, several of which are potent inhibitors. Examples of such compounds are carboxylic acids, furans and phenolic compounds (3, 4, 5, 6, 7). The microorganism used for fermentation of hydrolyzates should consequently exhibit three characteristics: a) it should have high ethanol tolerance, b) it should be resistant to inhibitors found in the hydrolyzate and c) it should have a broad substrate utilization range, since the hydrolyzate contains several different sugars. The quantitively most important sugars in hydrolyzate from spruce are glucose, mannose and xylose (6).
Due to its high ethanol yield, high specific productivity and high ethanol tolerance, Saccharomyces cerevisiae is the preferred microorganism for conversion of hydrolyzate to ethanol. It has also been shown that this yeast species is more tolerant to inhibitors such as acetic acid, furfural and 5-hydroxy-methyl furfural (HMF) than several other potential production microorganisms (8). The tolerance to, in particular, many of the aldehyde compounds can most likely be explained by a bioconversion of these compounds by the yeast to, in general, the less inhibitory corresponding alcohols. It is for instance known that S. cerevisiae converts furfural into the less inhibiting compound furfuryl alcohol (9, 10). With respect to sugar utilization, S. cerevisiae efficiently converts both glucose and mannose into ethanol, but is unable to convert xylose into ethanol. Other yeast species, e.g. Pichia stipitis and Candida shehatae are able to convert xylose into ethanol. However, these yeasts have a relatively low ethanol and inhibitor tolerance, and, furthermore, require microaerobic conditions in order to give a high productivity (8, 11). Work has consequently been made to genetically engineer S. cerevisiae in order to obtain xylose-fermenting capacity. In the xylose-metabolizing yeasts, xylose is channeled into the pentose phosphate pathway (PPP) in a three-step process. Xylose is first converted to xylitol by a xylose reductase (XP). Xylitol is then oxidized to xylulose by xylitol dehydrogenase (XDH), and finally, xylulose is phosphorylated to xylulose 5-phosphate by xylulose kinase (XK) (12). The first two enzymes are lacking in S. cerevisiae. Furthermore, the activity of XK in S. cerevisiae has been shown to be low (13), which has been suggested to limit the consumption rate in S. cerevisiae strains expressing XR and XDH (14). However, strain background appears to be important for the effect of XK (15, 16). In the present work, a genetically modified xylose-utilizing strain of S. cerevisiae was studied: TMB3006 (17). This strain express the heterologous genes XYL1 and XYL2 (encoding the enzymes XR and XDH, respectively) from P. stipitis, and overexpress the native gene XKS1 (encoding XK).
It has previously been shown that strongly inhibiting dilute-acid hydrolyzates, not fermentable in a batch process, can be fermented by S. cerevisiae without prior detoxification in a fed-batch operation. This, however, requires a carefully controlled hydrolyzate feed-rate (18, 19, 20). The most likely explanation for the success of fed-batch operation is that inhibitors are maintained at low levels because of their conversion to less toxic compounds. In the development of a closed-loop control strategy for fed-batch fermentation, the S. cerevisiae strain CBS 8066 (21) was used. It is known that different strains of S. cerevisiae show significant differences in fermentative capacity and inhibitor tolerance in batch cultivation. In the work by Carlos et. al. (22) significant differences in the ethanol productivity of several strains were found in batch cultivations with different levels of an inhibitor cocktail in a synthetic media. Only three out of 13 tested strains produced ethanol in batch fermentation with the highest level of the inhibitor cocktail. Not only the performance in batch culture, but even more so the performance in fed-batch culture is important for selection of a suitable strain. Besides the possibility to control the level of several potential inhibitors, the fed-batch operation offers another advantage in comparison to batch operation, which is the possibility of a parallel uptake of several sugars. The reason is that the concentration of sugars can be maintained at a low level by controlling the feed rate. In this way saturation of uptake systems (or saturation of the glycolytic flux) can be avoided, making co-fermentation of different sugars possible. In S. cerevisiae, xylose is believed to be transported by the same uptake system as glucose, however, with a much lower affinity (23). It is also known that a concomitant uptake of glucose increases the xylose consumption rate (24). One may therefore anticipate that a higher specific conversion rate xylose in hydrolyzates can be obtained in fed-batch cultivations.
Larroy, C., M. R. Fernandez, G. E, X. Pares, and J. A. Biosca. 2002. Characterization of the Saccharomyces cerevisiae YMR318C (ADH6) gene product as a broad specificity NADPH-dependant alcohol dehydrogenase: relevance in aldehyde reduction. Biochem J. 361:163-172 (38) have characterised the enzyme ADHVI and tested its kinetics for several substrates, primarily aliphatic and aromatic aldehydes. In their work the authors have primarily concentrated on the aromatic aldehydes (cinnamaldehyde and veratraldehyde). The authors suggest that ADHVI may give the yeast the opportunity to survive in ligninolytic environments where products derived from lignin biodegradation may be available. However, no tests have been made concerning the ability of ADHVI to use HMF as a substrate, HMF being a carbohydrate derived product. Furthermore, the paper does not discuss, or experimentally investigate, the potential role of ADHVI in protection against or conversion of inhibitors resulting from breakdown of the carbohydrates such as cellulose and/or hemicellulose.
Dickinson, J. R., L. Eshantha, J. Salgado, and M. J. E. Hewlins. 2003. The catabolism of amino acids to long chain complex alcohols in Saccharomyces cerevisiae. The Journal of Biological Chemistry 278:8028-8034. have studied the final step in the formation of long chain or complex alcohols in S. cerevisiae. They conclude that any of one of the six alcohol dehydrogenases (encoded by ADH1, ADH2, ADH3, ADH4, ADH5 or SFA1) is sufficient for the final stage of long chain or complex alcohol formation. Mutant strains were grown on single amino acids and fusel alcohol formation was measured. No measurements of enzyme activities in lysates nor any assessment of co-factor requirements were made. Importantly, the gene ADH6 was not at all studied, since it was regarded unlikely by the authors that an NADPH-dependent enzyme would be involved in fusel alcohol formation. The paper by Dickinson et al is therefore completely unrelated to the conversion of HMF and furfural, and is completely unrelated to any conversion catalyzed by the gene product of ADH6.
Martín, C., and L. J. Jönsson. 2003. Comparison of the resistance of industrial and laboratory strains of Saccharomyces and Zygosaccharomyces to lignocellulose-derived fermentation inhibitors. Enzyme and Microbial Technology 32:386-395, (3) have performed a comparison between 13 different yeast strains with respect to their resistance to lignocellulose-derived fermentation inhibitors. The strains are exposed to the following inhibitors: formic acid, acetic acid, furfural, HMF, cinnamic acid and coniferyl aldehyde. It is concluded that there is a big difference between the strains ability to tolerate and convert the inhibitors. However, no mechanistic investigations on the enzymes responsible for the conversion are presented or discussed in the paper. Specifically, there is no referral to either the gene product of ADH6 nor co-factor dependence in the reduction, made in the paper.
In the present work, different strains of S. cerevisiae were characterized in both batch and fed-batch fermentations of dilute-acid hydrolyzates. A total of four different strains were studied: CBS 8066, commercial bakers yeast, TMB3000, and TMB3006. The specific ethanol productivity, specific growth rate, consumption rates of monosaccharides, cell viability and furan reduction activities were determined. The results suggest that the furan reducing capacity is a key factor behind tolerance to lignocellulosic hydrolyzates.
The four strains of Saccharomyces cerevisiae used are given in Table 1. The strains were maintained on agar plates with the following composition: 10 g/l yeast extract, 20 g/l soy peptone, 20 g/l agar and 20 g/l glucose. Inoculum cultures were grown in 300 ml cotton plugged E-flasks with 100 ml of synthetic media according to Taherzadeh et. al. (25) with 15 g/l glucose as carbon and energy source. The inoculum cultures were grown for 24 h at 30° C. and with a shaker speed of 150 rpm before 20 ml was added to the fermentor to start the cultivation.
The hydrolyzate used was produced from forest residue, originating mainly from spruce, in a two-stage dilute-acid hydrolysis process using sulphuric acid as the catalyst (19). The hydrolyzates obtained from the two stages were mixed and stored at 8° C. until used. The composition of the hydrolyzate is given in Table 2.
Fermentation experiments were performed in a 3.3 l BioFlo III bioreactor (New Brunswick Scientific, Edison, N3, USA). The stirring rate was 400 rpm and the fermentor was continuously sparged with 600 or 1000 ml/min nitrogen gas (oxygen content guaranteed to be less than 5 ppm, ADR class2, 1(a), AGA, Sweden). The pH was maintained at 5.0 with 2.0 M NaOH. All experiments started with an initial batch phase in 1 l of synthetic media according to Taherzadeh et. al. (25) with 50 g glucose as carbon and energy source. However, the concentrations of media components other than glucose were tripled to compensate for the dilution during fed-batch operation. Hydrolyzate feeding was started at the depletion of the glucose, when the carbon evolution rate had decreased to less than 1 mmol/h.
Two types of fermentation experiment were made for each strain. In the first type of experiment, 1.5 liter of the hydrolyzate was added to the reactor using the maximum feed rate of the medium pump (approximately 2 liters/h) after the initial batch cultivation. This is referred to as “batch” fermentation. The second type of experiment was a fed-batch experiment, in which the hydrolyzate feed rate was controlled using a step-response method developed by Nilsson et. al. (19). In short, the feed-rate was changed in a step-wise manner, in which the step increase was proportional to the derivative of the measured carbon dioxide evolution rate from the previous step. Feed rate control was obtained by controlling the frequency of a peristaltic pump (Watson-Marlow Alitea AB, Sweden). Also in these experiments, a total of 1.5 l of hydrolyzate was added.
Additional fed-batch experiments were made with the xylose-fermenting yeast (TMB3006) using low feed-rates (12.5 and 25 ml/h). The purpose of these experiments was to obtain low medium concentrations of glucose and mannose, expected to give an increased xylose uptake rate.
A gas monitor (model 1311, Brüel and Kjaer, Denmark) (described by Christensen et. al.) was used to measure the carbon dioxide evolution rate (CER). The gas analyzer had three channels for measurement of carbon dioxide, oxygen and ethanol in the off-gas from the reactor. The ethanol signal was calibrated against ethanol concentrations measured in the broth by HPLC, since it was assumed that the ethanol in the gas phase was in equilibrium with the ethanol in the broth. Calibration for oxygen and carbon dioxide was done using a gas containing 20.0% oxygen and 5.0% carbon dioxide.
A flow-injection-analysis (FIA) system (26) was used to measure biomass concentration in the reactor. This was done by measuring the optical density at 610 nm, at a frequency of 1 h−1. After every fermentation the FIA-signal was calibrated against measured dry-weight. Duplicate 10 ml samples of the fermentation broth were centrifuged at 3000 rpm for 3 min in pre-weighted tubes. The cells were washed with distilled water, centrifuged again and dried over night at 105° C. before they were weighted again. The dry weight was measured three times during each fermentation.
Cell viability was measured as the ratio between colony forming units (CFU) and counted cell numbers three times during each fermentation. Samples were withdrawn from the fermentation broth and diluted to give a concentration of around 1000 cells/ml, and CFUs were determined from triplicate agar plates onto which 0.1 ml samples of diluted broth were spread. Cell numbers were calculated under microscope using a Bürker chamber. Prior to the calculation the samples were diluted 100 times.
Samples for analysis of metabolite concentrations were taken regularly from the reactor. The samples were centrifuged and filtered trough 0.2 μm filters. The concentrations of glucose, mannose, xylose, galactose and arabinose were measured on an Aminex HPX-87P column (Bio-Rad, USA) at 80° C. The concentrations of ethanol, HMF, furfural, glycerol and acetic acid were measured on an Aminex HPX-87H column (Bio-Rad, USA) at 65° C. All compounds were detected with a refractive index detector, except for HMF and furfural, which were detected with a UV-detector (210 nm).
To compensate for evaporated ethanol during the fermentations, the Mole fraction of ethanol in the gas phase was assumed to be proportional to the mole fraction of ethanol in the liquid phase. The amount of evaporated ethanol could thereby be estimated by integration of the gas flow leaving the reaction multiplied with the mole fraction of ethanol in the gas, as described by Nilsson et. al., (18).
Cell extracts were prepared for measurements of enzyme activities in the strains TMB3000 and CBS 8066. Crude extracts were made using Y-PER reagent (Pierce, Rockford, Ill., USA). The cell extracts were kept in an ultra freezer (−80° C.) until used. The protein content: in the cell free preparation was determined by Coomassie Protein Assay Reagent using bovine serum albumin as a standard (Pierce, Rockford, Ill., USA).
Furfural and HMF reducing activity was measured according to Wahlbom et. al. (27). 20 μl of the cell free extract (diluted ten times in 100 mM phosphate buffer) was diluted in 2.0 ml of 100 mM phosphate buffer (50 mM KH2PO4 and 50 mM K2HPO4) and furfural was added to a concentration of 10 mM. The samples were heated to 30° C. and thereafter the reaction was started by addition of NADH to a concentration of 100 μM. The oxidation of NADH was followed as the change in absorbance at 340 nm. The same procedure was repeated with NADPH as the co-factor, but the sample amount was increased to 200 μl due to the lower activity. The total volume was still 2.0 ml and the concentrations of furfural and NADH 10 mM and 100 μM respectively.
The same procedure was repeated for measurement of HMF reduction capacity, 200 μl of diluted cell extract was used except for the measurement for strain TMB3000 with NADH as the co-factor where 20 μl sample was used due to the higher activity. The concentration of HMF was 10 mM. Activities were measured with both NADH and NADPH.
Measurement of ADH activity
ADH activity was measured according to Bruinenberg et. al. (28). The cell free extract was diluted 10 times and 20 μl of this dilution was added to 2.0 ml of 100 mM phosphate buffer. Ethanol was added to give a concentration of 100 mM. After heating to 30° the reaction was started by addition of NAD+ to a concentration of 100 μM. The reduction of NAD+was followed as the change in absorbance at 340 nm.
To analyze the mRNA content in strain CBS 8066 and TMB 3000 continuous cultures were run. The synthetic media was according to (25), but 33% more concentrated and the glucose concentration was 20 g/l. The liquid volume in the reactor (Belach BR 0.5 bioreactor, Belach Bioteknik AB, Solna, Sweden) was 500 ml and after the glucose in the batch had been consumed the feed was started at a dilution rate of 0.1 h−1. The reactor was sparged with 300 ml of nitrogen/min. pH was maintained at 5.0 with 0.75 M NaOH and the temperature at 30° C. The stirrer speed was set to 500 rpm. To investigate which genes were induced by HMF, cell samples for mRNA analysis were taken both after feeding the reactor with media without HMF and with media including 0.5 g HMF I−1. To get a steady state in the reactor the samples were taken 5 resident times after start of feed or change in feed media.
mRNA Preparation
Samples from the reactor were spinned in ice at 3000 rpm for 1 min and thereafter frozen in liquid nitrogen and stored at −80 C until mRNA was isolated from the samples. The mRNA was isolated using Fast RNA kit (Q-biogene, USA). The mRNA was then purified, cDNA synthesized, in-vitro transcribed and fragmented as described by Affymetrix. Hybridization, washing, staining and scanning of microarray-chips (Yeast Genome S98 Arrays) were made with a Affymetrix Gene Chip Oven 640, a Fluidics Station 400 and a GeneArray Scanner (Affymetrix).
Selected strains (over expressing LAT1, ALD6, ADH5, ADH6, GDH3, OYE3, IDP3, ADH7, AAD15, ERG27, HMG1, LYS5, SPS19, SGE1) from the ExClone collection (Resgen, Invitrogen Corporation (UK)) were grown in 300 ml shake flasks (with carbon dioxide traps) containing 100 ml SD-Ura emission media and 40 g/l glucose as described by the supplier. However, 80 μM of Cu2+ was added when the shake flasks were inoculated and a 100 mM phosphate buffer were used. Samples for enzyme activity measurements were taken after 16 hours of growth at 30° C. and 150 rpm.
Batch and fed-batch fermentations were performed with four yeast strains. After an initial batch growth phase on synthetic media, 1.5 liters of hydrolyzate was added to the reactor. In the batch fermentations hydrolyzate was added with the maximal rate (approximately 2000 ml/h), whereas in the fed-batch experiments the feed-rate was controlled using a closed-loop control algorithm. In short, the feed-rate was changed in a step-wise manner, in which the step increase was proportional to the derivative of the measured carbon dioxide evolution rate from the previous step. Feed rate control was obtained by controlling the frequency of a peristaltic pump (Watson-Marlow Alitea AB, Sweden). (see Materials and Methods).
There were significant differences between the strains, in particular with respect to fermentation rates, as reflected by the carbon dioxide evolution rate (
The ethanol productivity was higher in fed-batch compared to batch fermentation for all strains tested (Table 3). For CBS 8066, the average ethanol productivity increased by 131%. However, a gradual decrease in CER could not be avoided, and there was no cell growth, although a high viability was maintained. In fact, the viability was well maintained for all strains during fed-batch operation.
Apart from CBS 8066, the other strains grew in fed-batch fermentation (
The strain carrying genes coding for XR, XDH and XK chromosomally integrated, TMB3006, consumed 6% of the xylose in the hydrolyzate. Xylose was assumed to be converted to ethanol, since no xylitol was detected (
Furfural and HMF reduction capacity was measured on cell extract sampled during fed-batch experiments. The enzyme activities were measured with both NADH and NADPH as co-factors (
Already before addition of any hydrolyzate, there was a clear difference between the activities in the two strains (
For TMB3000 the ADH activity was on average 40% higher than for CBS 8066 (
To investigate which enzyme(s) was responsible for the high conversion rates of in particular HMF and, in particular, with NADH as the cofactor, continuous cultivations where run with TMB 3000 and CBS 8066 with and without HMF present in the synthetic media. We searched for known reductase and hydrogenase genes that were upregulated at least twice in TMB 3000 in comparison with the strain CBS8066, both with or without the presence of HMF. —Maybe you should include the list here again—As seen in
Strains from the ExClone collection in which the genes identified from the mRNA analysis described above were upregulated, were grown in shake flasks cultivations, and the obtained activities for reduction of furans were measured (
The present experiments clearly demonstrate that the ability of S. cerevisiae to ferment dilute-acid hydrolyzates of cellulosic material is highly strain specific. Importantly, and in accordance with previous work on the strain CBS 8066 (29, 20, 19, 18), higher productivities were obtained in fed-batch operation for all strains tested. The specific ethanol productivities for the most inhibitor tolerant strains (TMB3000 and TMB3006) increased by 69% in comparison to batch operation. Growth in batch cultivation was negligible for all strains, but the specific ethanol productivity varied significantly between strains also in batch fermentation. In contrast, all strains—with the exception of the strain CBS 8066- to some extent grew in anaerobic fed-batch cultivations. The lower degree of inhibition in fed-batch cultivation is most likely attributed to the in-situ conversion of one or more inhibitors—including furan compounds and other aldehydes (30, 31, 22).
The physiological effects of furfural on S. cerevisiae have been previously studied extensively in synthetic model media. It has been shown in furfural-containing chemostat cultivation (both anaerobic and aerobic), that growth is inhibited if the specific furfural conversion rate exceeds a maximum critical conversion rate, During anaerobic conditions, the determining rate is the rate of reduction to furfuryl alcohol, whereas for aerobic conditions the critical rate is instead the oxidation rate to form furoic acid. At a too high furfural feed load, the furfural concentration increases in the medium, which presumably leads to inhibition of a number of key enzymes, including PDH and AIDH and washout occurs. For strain CBS 8066 the critical specific conversion rate of furfural was found to be between 0.10 and 0.15 g/g h during anaerobic conditions. In the present work, the concentration of furfural was very low (<0.04 g/l) in all fed-batch cultivations, and it appears that the critical conversion rate of furfural was not exceeded. However, there were larger differences with respect to the HMF concentrations. In the cultivations with the best growing strains, TMB3000 and TMB3006, the HMF concentration was maintained at a relatively low level (<0.23 g/l) whereas in the fed-batch fermentation with CBS 8066 only little conversion of HMF took place. For the two strains CBS 8066 and TMB3000, fed-batch fermentations were repeated and the activities of furfural and HMF reduction were measured (
In an industrial medium the furfural concentration may 1 g/l up to 3 g/l depending on its origin.
Normally the furfuryl alcohol will be measured as the fermentation is anaerobic, and the product is then almost exclusively furfuryl alcohol.
The transformation capacity, conversion rate (determined by the enzymatic activity) determines how fast it is possible to add the furans. If they should be added too fast, furans will be accumulated in the medium, which will lead to an inhibition of central functions by means of interactions between furans and a number of enzymes such as PDH, PDC and others. This in turn leads to an inhibitor growth and down-regulation of the fermentation rate.
The average enzyme activities for furfural and HMF conversion in CBS 8066 was similar to those found for strain TMB3001, a strain derived from CEN.PK PK113-7A. The average activities for furfural conversion in CBS 8066 were 353 mU/mg protein with NADH as co-factor and 22.8 mU/mg protein with NADPH as co-factor, compared to 490 and 22 mU/mg protein, respectively, found in TMB3001. For HMF conversion, the average activities for CBS 8066 were 1.8 mU/mg protein (NADH) and 12.4 mU/mg protein (NADPH), compared with 2.2 and 22 mU/mg protein, respectively, for TMB3001. The Enzyme activities obtained for the strain TMB3000 were very different. The average furfural reduction activity was several times higher than for CBS 8066 and TMB3001, although the co-factor preference was similar. The most striking difference was, however, the high activity for HMF reduction with NADH as co-factor (
The furfural and HMF conversion activities provide an explanation for the advantage of TMB3000 over CBS 8066 in lignocellulose fermentation. High activities ensure high conversion rates of furfural and HMF, and possibly other inhibitory aldehydes (32), so that the concentration of these inhibitors is kept low in the fermentation. For strain CBS 8066 the measured in vitro activity for furfural reduction would correspond to an in vivo reduction rate of 0.69 g/g h. This, in fact agrees well with the maximum conversion rate reported in synthetic media for the same strain (0.6 g/g h). The corresponding predicted specific conversion rate of HMF would be 0.03 g/g h, which is somewhat lower than the value reported in synthetic media of 0.14 g/g h. For strain TMB3000 measured in vitro reduction activities for furfural and HMF were 2.26 g/g h and 0.98 g/g h respectively and this would correspond to feeding rates of about 3 l/h, at the cell density and volumes used which is much higher than those applied in the present work (cf.
There was a considerable furan reduction activity in the cell extract already before the cells had been exposed to the inhibitors of the hydrolyzate, and furthermore, activity measurements showed that the furan reduction activity did not increase significantly with time during exposure to hydrolyzate, indicating that the responsible enzyme(s) were not induced. The ability to reduce furfural has previously been attributed to the enzyme alcohol dehydrogenase (ADH) (32, 33), although this has been questioned (35). The ratio between measured ADH activities for CBS 8066 and TMB3000 (
Below it is further shown that the enzyme encoded by the gene ADH6 in Saccharomyces cerevisiae is able to convert HMF using the co-factor NADPH. Yeast strains that over-express this gene have a substantially higher conversion rate of HMF in both aerobic and anaerobic cultures. Importantly, we have furthermore shown that strains over-expressing ADH6 has a substantially higher ethanol productivity and are less effected by inhibition during fermentation of a dilute-acid lignocellulose hydrolyzate. Strains genetically modified to give a high expression of ADH6 will therefore be advantageous for the conversion of lignocellulosic hydrolyzates.
The alcohol dehydrogenase VI (ADH6) gene from Saccharomyces cerevisiae TMB3000 and CEN.PK 113-5D were amplified using the primers ADH6-FOR and ADH6-REV (Table 4). The 5′ region of the primers ADH6-FOR and ADH6-REV contain 34 and 33 nucleotides corresponding to the sequence of the HXT promoter and PGK1 terminator, respectively. After PCR amplification, the PCR products were analyzed by electrophoresis in agarose gels and purified using QIAquick PCR Purification kit (QIAGEN). The vector pYEplacHXT was linearized using the restriction endonuclease Bam HI. A mix containing the linear vector to (6.2 Kb), the ADH6 product from TMB 3000 (T-ADH6) or CEN.PK 113-5D (C-ADH6) was used to transform S. cerevisiae CEN.PK 113-5D by lithium acetate method (38). Yeast cells were grown overnight, in 5 mL YPD, at 30° C. In the morning, a 50 mL YPD solution was inoculated using 3 mL of the pre-culture. Growth was followed until OD600=1.2, when the yeast cells were centrifuged and finally suspended in 10 mL of sterile water. One milliliter of cells were pipetted in micro-centrifuge tubes and centrifuged for approximately 20 seconds in top speed. After supernatant removal, the cells were resuspended in 1 mL of 100 mM lithium acetate (LiAc) and incubated at 30° C. for 10-15 minutes. The suspension was centrifuged at top speed for 30 seconds, the supernatant removed and the transformation mix (240 μl PEG 50% w/v, 36 μl 1.0 M lithium acetate, 52 μg 2 mg/mL ssDNA, 28 μl sterile water, 1.0 μl 40 ng/μl pYEplacHXT vector and 3 μl 40 ng/μl PCR product) was added to the pellet. After subsequent incubations at 30° C. for 30 min and 42° C. for 20 min, the mix was centrifuged at top speed for 30 seconds and the transformation mix removed. The yeast cells were re-suspended in 150 μl of YNB and left at room temperature for approximately 2 hours. After incubation the mix containing cells was plated on YNB-plates, which were incubated at 30° C. for 3-4 days. A yeast control strain was constructed by transformation with the empty pYEplacHXT vector. Transformant yeast strains were selected by colony PCR using ADH6 primers and ethanol oxidation capacity. Plasmids from two transformants (C-ADH6-2 and T-ADH6-2) were recovered, amplified in E. coli DH5α and submitted to automatic sequencing.
Growth experiments were carried out in 300 ml unbaffled shake-flasks. The volume of synthetic media was 200 ml with the composition given in (25) and contained 13 g glucose. The pH was adjusted to 5.5 with 2 M NaOH at the start of the cultivations. The shaker speed was 170 rpm and the temperature was 30° C. The anaerobic shake flasks were equipped with glycerol traps, whereas the aerobic shake flasks were sparged with air. When OD620 reached 3.0 the pH was readjusted to 5.5 and HMF was added to a concentration of 1.5 g/l.
Batch fermentations were made with the strain CEN.PK 113-5D and T/ADH6-2. The reactor (Belach BR 0.5 bioreactor, Belach Bioteknik AB, Solna, Sweden) was initially filled with 300 ml synthetic media according to (25), which contained 30 g. glucose. pH was maintained at 5.0 with 0.75 M NaOH and the temperature at 30° C. The reactor was sparged with 300 ml/min of nitrogen and the stirrer speed was set to 500 rpm. When the carbon dioxide evolution rate had reached a maximum, 300 ml hydrolyzate was added.
The hydrolyzate used was produced from forest residue, originating mainly from spruce, in a two-stage dilute-acid hydrolysis process using sulphuric acid as the catalyst (19). The hydrolyzates obtained from the two stages were mixed and stored at 8° C. until used. The composition of the hydrolyzate is given in Table 5.
Cell extracts of strains over-expressing ADH6 were prepared for measurements of enzyme activities. Crude extracts were made using Y-PER reagent (Pierce, Rockford, Ill., USA), The protein content in the cell free preparation was determined using Micro BCA Protein Assay Kit (Pierce).
Enzyme activities for theoxidation of ethanol and the reduction of furfural, 5-hydroxymethyl-furfural (HMF) and dihydroxyacetone phosphate (DHAP) were measured on cell extract samples. The rate of ethanol oxidation was determined by monitoring the reduction of NAD+ photometrically at a wavelength of 340 nm. The enzyme assay, based on (37), contained 5.0 mM NAD+ and 1.7 M of ethanol in 100 mM glycine buffer at pH 9.0 in 1.0 cm path length cuvettes. The samples were incubated at 30° C. and the reaction was started by addition of ethanol. HMF and furfural reducing activities were measured according to (27). 5-10 μL of cell free extract (using different dilutions) was diluted in 1 mL of 100 mM phosphate buffer (50 mM KH2PO4 and 50 mM K2HPO4) and NADH was added to a concentration of 100 μM. The samples were incubated at 30° C. and thereafter the reaction was started by addition of HMF or furfural to a concentration of 10 mM. The oxidation of NADPH was followed as the change in absorbance at 340 nm. The procedure was repeated with NADH as the co-factor, but the sample amount was increased due to the lower activity. The total volume was still 1.0 mL. The same procedure was carried out when using DHAP, except that only 0.7 mM of this substrate was used. The molar absorption coefficient (s) used for NADH and NADPH was ε340=6.22 mM−1cm−1.
ADH6 gene was PCR amplified from CEN.PK or TMB3000 genomic DNA and cloned into the yeast vector pYEplac-HXT, generating pYEplacHXT-C/ADH6 and pYEplacHXT-T/ADH6 vectors respectively. The plasmids were used for the transformation of CEN.PK113-5D strain. Colony PCR was used to select yeast strains that carried a pYEplacHXT-ADH6 vector. Clones having the ADH6 gene from CEN.PK and TMB3000 were called C/ADH6-m (m=1, 2 etc) and ADH6-n (n=1, 2, etc), respectively). Genes with increased expression of ADH6 gene were selected amongst transformants for their increased ethanol oxidation capacity compared to the control strain CEN.PK113-5D carrying the empty vector YEplac-HXT (
HMF and furfural conversion capacity of ADH6 over-expressing strains was analyzed using NADH and NADPH as cofactors in enzymatic assays (
In vivo HMF conversion was analyzed in minimal medium using aerobic and anaerobic conditions in shake-flasks. The ADH6 over-expressing strains showed higher specific HMF uptake (3.5-3.9 fold) in aerobic as well as in anaerobic conditions (Tables 6 and 7). The specific uptake of HMF appeared correlated with an increase in glycerol production (Tables 6 and 7). In order to analyze a possible direct activity of ADH6 gene product in the glycerol metabolic pathway, the C-ADH6-2 and T-ADH6-2 DHAP reduction capacity was analyzed by enzymatic assays. Enzyme activity measurements did not shown any increase in DHAP reduction (
The control strain and a strain over-expressing ADH6 from TMB3000 (T/ADH6-2) were used in anaerobic batch fermentations with a dilute-acid hydrolyzate (
Conclusions drawn from this latter experiment series are:
1Calculated as the average specific ethanol productivity during the fermentation of hydrolyzate, until CER decreased to less than 5 mmol/h.
2CFU value taken a 2-8 hours after start of the feeding of hydrolyzate (%)
| Number | Date | Country | Kind |
|---|---|---|---|
| 0401303-3 | May 2004 | SE | national |
| Number | Date | Country | |
|---|---|---|---|
| Parent | 11560490 | Nov 2006 | US |
| Child | 12201292 | US |
| Number | Date | Country | |
|---|---|---|---|
| Parent | PCT/SE05/00738 | May 2005 | US |
| Child | 11560490 | US |
