Ethanol production from lignocellulosic biomass has to include a pretreat­ment more intensive than those used in processing sugar and starch-rich biomass in order to release the sugar compounds contained in the biomass. Agricultural residues like wheat straw or other types of biomass derived from plant material contain lignin, which is constructed to resist microbial attack and to add strength to the plant. Pretreatments are used to open the biomass by degrading the lignocellulosic structure and by partially hydrolysing the substrate. Current pretreatment methods, however, contribute to 30-40% of the total costs of bioethanol production from lignocellulosic biomass. The National Renewable Energy Laboratories (NREL) estimates that in an Nth generation plant (mature technology), feedstock handling and pretreat­ment would account for approximately 20% of the total ethanol production costs [15].

Several pretreatment methods have been developed [16] (see also Zac — chi, this volume). However, in all methods the biomass concentrations need to be higher than 20% dry weight to ensure a suitable ethanol concen­tration for the subsequent distillation process. A new patented pretreat­ment process, Wet-ox explosion (WE), has been developed in our labora­tory combining steam-explosion and wet oxidation using small amounts of oxygen [17]. The optimal combination of process parameters such as tem­perature (170-200 °C), pressure (12-30 bar), amount of oxygen addition, and residence time (2-15 min) has been tested. Depending on the biomass mate­rial used, the method will yield variable sugar yields but overall the results show that the method will be efficient and cost-effective for opening of most major biomass materials such as straw, corn stover, bagasse, and woody materials. Table 1 shows the apparent advantages and disadvantages of this pretreatment method.

Table 1 Advantages and disadvantages of Wet-Ox-Explosion

Disadvantages

Fast and efficient Requires water supply

No emission products Advanced technology

Low heat consumption No standard equipment

No detoxification Only tested on pilot scale

Easily convertible substrates No waste products

4.2

Since S. cerevisiae cannot utilize xylose, but does utilize and ferment its iso­mer D-xylulose [1,2], the obvious first step to allow xylose metabolism is to introduce a heterologous pathway converting xylose to xylulose. Over the years, several approaches have been explored to express a pentose utilization pathway from naturally pentose-utilizing bacteria and fungi in S. cerevisiae. Figure 1 summarizes the initial pathways for D-xylose utilization in bacteria and fungi.

Ul

О

NR: not reported

a Batch culture by pulsing a chemostat culture b Calculated from reference c Ethanol yield on xylose d Calculated after glucose depletion

NR: not reported a Calculated from reference

NR: not reported

a Measured and corrected by closing the DR balance b Calculated from reference

2.1

6.1

Evolutionary Engineering of D-Xylose-Consuming S. cerevisiae for Improved Mixed Substrate Utilisation

The sub-optimal kinetics of mixed-substrate utilisation by the genetically engineered XylA-expressing strain RWB 217 [43] suggested a low affinity (qmax/Ks) for D-xylose. Soon after the invention of the chemostat it was al­ready established that prolonged cultivation in nutrient-limited chemostats leads to selection of spontaneous mutants with an improved affinity for the growth-limiting nutrient [52,53]. This principle, which has since been demonstrated for many micro-organisms and nutrients [40,58,72,73] was applied to improve the affinity of S. cerevisiae RWB 217 for D-xylose [44].

Indeed, during prolonged anaerobic D-xylose-limited chemostat cultivation at a dilution rate of 0.06 h-1, the residual D-xylose concentration decreased threefold, indicating that cells with improved affinity for D-xylose were se­lected for [44]. After 1000 h (85 generations) of this directed evolution in chemostat cultures, single-colony isolates were tested for batch growth on a mixture of glucose and D-xylose. Although the fermentation kinetics of some of these single-cell lines, as evaluated by carbon dioxide production profiles, were already drastically improved relative to the parental strain (Fig. 6), the D-xylose phase remained slower than anticipated based on batch cultivation on D-xylose alone. A further 85 generations of chemostat cultivation resulted in only marginal improvement of the D-xylose consumption characteristics.

To select for further improvement of D-xylose fermentation kinetics, an additional evolutionary engineering strategy was applied, which involved sequential anaerobic batch cultivation on glucose-xylose mixtures [44]. To maximise the number of generations that the cells grow on D-xylose, the D-xylose concentration in the cultures was raised to 90 gL-1, with a glucose concentration of 20gL-1. After 20 cycles, the evolved culture was capable of complete anaerobic conversion of a mixture of 20 g L-1 glucose and 20 g L-1 D-xylose in about 20 h, with an inoculum size of 5% (v/v) [44].

Characterisation of the resulting strain RWB 218 (derived from single colony isolate) showed that D-xylose consumption followed the consump­tion of glucose rapidly (Fig. 7). The D-xylose consumption rate observed in these cultures was 0.9 g D-xylose (gdryweight)-1 h-1. This evolved Xl-based strain, in contrast to strains based on xylose reductase and xylitol dehydroge­nase, produced only 0.45 mM of xylitol, indicating that redox imbalance does

not occur during alcoholic fermentation of D-xylose. The ethanol yield on total sugar in batch cultures co-fermenting glucose and D-xylose was typic­ally 0.40 g g-1, which is identical to the ethanol yield that would be achieved in glucose-grown cultures in a similar set-up. Even when tested in more concen­trated sugar mixtures (100 g L-1 glucose and 25 g L-1 D-xylose), resembling an industrial situation, this strain consumed both sugars within 24 h, starting with 1.1 gL-1 yeast dry weight as the inoculum [44].

With evolutionary engineering as a proven tool for obtaining (yeast) strains with improved properties, a full understanding of the underlying molecular changes becomes the next challenge. In an attempt to unravel the changes between the original metabolically engineered and the subsequently evolved Piromyces XI-based strains, anaerobic chemostat cultivations on D-xylose as the sole carbon source were used as the basis for transcriptome analysis with Affymetrix DNA arrays (J. T. Pronk, unpublished data). The most striking observation amongst the genes with a changed transcript level was the repre­sentation of various members of the hexose transport family, including HXT1, HXT2 and HXT4. Interestingly, HXT1 and HXT4 have been associated with D-xylose transport in previous studies [27,62]. To investigate whether the improved fermentation characteristics were indeed due to changes in sugar transport, zero trans-influx assays were performed using both the strain that was only metabolically engineered and the subsequently evolved strain [44]. The D-xylose uptake kinetics obtained for the metabolically engineered strain (Km 132 mM, Vmax 15.8 mmol (gdryweight)-1 h-1) were in agreement with other studies [22,39]. Strikingly, the D-xylose uptake kinetics of the evolved strain had changed drastically, with a 25% reduction in the Km (to 99 mM) and a twofold increase of Vmax to 32 mmol (g dryweight)-1 h-1.

6.2

Succinate, a natural E. coli fermentation product, can serve as substrate for the production of many compounds currently derived from petroleum [112]. Although there have been numerous reports of succinate production by E. coli and other biocatalysts (for example [147,148]), these processes often involve undesirable nutritional supplementation, multiple steps and low product titers.

Due to our success, described above, in using a combination of directed engineering and metabolic evolution to design ethanol and lactic acid mi­crobial biocatalysts, we have used a similar approach to develop a succinate — producing microbial biocatalyst that attains high product titers in simple mineral salts media [149]. Directed engineering consisted of elimination of the lactate, acetate, and ethanol-forming pathways (IdhA, ackA, adhE), leaving succinate production as the primary route of NADH oxidation. The poor growth and fermentation of the resulting strain in mineral salts media were improved by metabolic evolution. Further directed engineering (focA, pflB, mgsA) reduced co-product formation. The resulting microbial biocat­alysts, KJ060 and KJ073 (poxB), produced nearly 700 mM succinate from glucose with a molar yield of 1.2-1.6; the maximum theoretical molar yield is 1.71 (Jantama et al., unpublished results). KJ060 and KJ073 produced 250 and 183 mM acetate, 39 and 118 mM malate, 0 and 5 mM pyruvate, and 2 and 0 mM lactate as co-products.

5.5

Production of ethanol from starch-based crops such as wheat and corn is a well-known technology. Such processes have been optimized over a long time and are reaching a level of maturity where further cost reductions, based on improvements in conversion technology, are becoming more difficult. In contrast, processes using lignocellulosic raw materials are still under develop­ment and significant reductions in ethanol production cost can be expected.

With some modifications it is possible to make a basic evaluation of a starch-based process (Fig. 1) and compare it with a process based on ligno­cellulosic material (Fig. 3). This was done by Wingren [17]. The purpose of the evaluation was not to determine the absolute ethanol production cost, but to compare the processes using the same fundamental cost basis and the same assumptions in the investment analysis. A comparison of this kind provides valuable information on the major differences between a commercial process and a process under development.

Both plants were designed for an annual ethanol production of 55 000 m3, which is a rather small plant. This value is on a pure ethanol basis, although the actual distillate was assumed to be 94% (w/w), i. e. no dehydration step was included as this would have been the same in both cases. Also, no off­sites, e. g. production of heat and electricity, were included, only the pure ethanol production facility. In the evaluation no credit was given for carbon dioxide. The cost of the enzymes in a starch-based plant is lower than in a lig- nocellulosic plant. In the study it was assumed to be 0.014 US$ L-1 ethanol, which is slightly higher than the cost reported for the enzymes in a corn — based plant located in the USA [18].

The raw material flow is higher in the lignocellulosic process, 200 000, compared to 126 500, dry metric tons y-1 for the starch-based plant, due to the lower overall ethanol yield and the somewhat lower amount of fermentable sugars in the raw material. The overall energy demand in the lignocellulosic process was estimated to be 16 MJ L-1 ethanol compared to 10 MJ L-1 for the wheat-based process. The fixed capital investment was estimated to be 99 and 53 million US$ for the lignocellulosic and the starch-based processes, respec­tively. A breakdown of costs is presented in Table 3. The estimated ethanol production cost was 0.60 and 0.58 US$ L-1 for the lignocellulosic and starch — based processes, respectively. Major differences were found in the cost of raw material, enzymes, capital, steam as well as income from the co-products. It should be noted that, although significantly higher than in the starch-based process, the enzyme cost in the lignocellulosic process was based on a pro­jected future cost. In the starch-based process the cost of the raw material constitutes as much as 65% of the total production cost. This is typically the case for well-established, mature processes. Thus, the economics of a starch — based process is very dependent on the cost of feedstock.

The lignocellulosic process is more dependent on the income from the co-products. However, the potential price of the syrup is uncertain since its fuel properties are unknown. At 12.9 US$ MWh-1 the income from this co­product was estimated to be 0.03 US$ L-1. In a scenario where the co-product instead has to be disposed of and cannot be utilized as a fuel, the ethanol production cost for the lignocellulosic process would be 0.63 US$ L-1. The

income from the pellets in the lignocellulosic plant reduces the ethanol pro­duction cost by 0.15 US$ L-1 at 20 US$ MWh-1. As in the case of the syrup, the true price of this co-product will be dependent on its fuel properties.

The results of this comparative study led to some important conclusions regarding potential cost reductions in the lignocellulosic process, compared with the starch-based process. The overall ethanol yield in the lignocellu­losic process evaluated is 68% of the theoretical based on the available glucan and mannan in the raw material, a figure that can probably be increased. In addition, a pentose — and galactose-fermenting organism could increase the ethanol production per unit raw material without increasing the capital cost. This is especially important if the raw material is rich in pentoses, e. g. as in straw or hardwood. A reduction in enzyme loading would also be rewarding provided that the ethanol yield could be maintained. Figure 9 shows a break­down of capital costs together with energy costs for the two processes. The largest difference in costs is seen in the conversion steps and in the evapo­ration step. The pretreatment step in the lignocellulosic process represents around 0.093 US$ L-1 ethanol. This cost is attributed to both a high energy demand and to the high cost of the reactor system. This shows the need to improve pretreatment and/or enzymatic hydrolysis so that less severe pre­treatment is required. The higher cost of the SSF step compared with the fermentation step in the starch-based process is due to the longer residence

time and the lower substrate concentration in the lignocellulosic process. An increase in substrate load and productivity in the lignocellulosic process would reduce this difference. The difference in cost between the starch-based process and the lignocellulosic process in the downstream processing steps (evaporation and distillation) would also be reduced if the ethanol concentra­tion in the SSF step could be increased.

3.3

The S. cerevisiae genome contains the gene XKS1 coding for XK [26,27], but the XK activity in wild-type S. cerevisiae is too low to support ethanolic xylose fermentation in strains engineered with a xylose pathway [26,99,100]. It is only when additional copies of XKS1 are expressed that recombinant xylose­utilizing S. cerevisiae produce ethanol from xylose [79] (strain TMB3001, Tables 1-4), [100] (strain H1691, Table 1), [101] (strain 1400 (pLNH32), Table 2; strain H2490, Tables 3 and 4). However, nonphysiological or unreg­ulated kinase activity may cause a metabolic disorder [102]. It was indeed experimentally demonstrated that only fine-tuned overexpression of XKS1 in S. cerevisiae led to improved xylose fermentation to ethanol [103,104]. Sim­ilarly, it was shown that arabinose-utilizing recombinant S. cerevisiae strains expressed a mutated L-ribulokinase gene with lower specific activity, indicat­ing that a low kinase activity had been selected as advantageous for arabinose utilization [71].

4.4

Den Haan et al. [49] calculated that a 20- to 120-fold improvement in CBH ex­pression, as well as simultaneous high-level expression of other cellulase com­ponents, will be necessary for slow growth on crystalline cellulose. This calcu­lation assumes a strain that can grow at 0.02 h-1 has a typical anaerobic yield of 0.1 gbiomass/g substrate or an aerobic yield of 0.45 gbiomass/gsubstrate, that the expressed cellulase has a specific activity which is the same as that of crude T. reesei cellulase on avicel (0.6 U/mg), and that CBH1 would make up the same fraction of total cellulase protein as in the T. reesei system [9]. While techniques for rational design of cellulases for improvement in ex­pression level and potentially specific activity will be important to achieving this goal, techniques involving random natural and/or induced mutation will also play an important role. The well-established success of directed evolu­tion techniques for enzymes and enzyme systems (e. g., see reviews in [132, 133]) can be transferred to engineering organisms for CBP, although this application does present unique challenges due to the lack of a good high — throughput screening technique for activity on insoluble cellulosic substrates. On the other hand, the natural connection between cellulase expression and growth on cellulose for CBP organisms makes whole cell selection-based strategies for improving cellulase production a powerful way to screen very large libraries of candidate cells, mimicking the evolutionary process found in nature.

An assumption for any selection-based improvement for CBP organisms is that mutations can result in increased cellulase activity expression. For total cellulase activity such mutations would increase either the per cell expression level (g cellulase/g cell) or the cellulase specific activity (U/mg cellulase). Mu­tagenesis and screening techniques have allowed researchers to isolate strains of S. cerevisiae with “super-secreting” phenotypes [134-136], and similar techniques for the expression of individual cellulase components have been successful [137]. Also, random mutation has been used to change the prop­erties of cellulase enzymes (e. g., [138-140]; a further review can be found in [141]), although to our knowledge enhanced overall specific activity of cellulase on insoluble substrates has not been demonstrated via directed evo­lution. However, the specific activity of a mixture of cellulases also depends on the relative amounts of cellulase components to achieve the highest degree of enzyme-enzyme synergy [142], as well as other parameters (as yet not elu­cidated) that determine enzyme-microbe synergy [6]. These features could be impacted by mutation and therefore lead to enhanced specific activity of cellulase systems expressed by recombinant cellulolytic CBP organisms.

Earlier in this review (Sect. 3) the relationship between cellulase activity and growth rate was examined from a whole-population perspective, using parameters that are averages for many cells. The relationship between growth rate and cellulase expression for an individual cell, especially a cell harbor­ing mutations affecting cellulase expression, as compared to other cells in the population depends on the diffusion of the soluble reaction products from the point they are created at the cellulose surface to the point they are taken up by a particular cell. When a connection between growth rate and en­zyme production can be established, selection in liquid culture—particularly continuous culture—has the potential to screen many cells. For example, if a continuous reactor had a cell concentration of 1010 cells/L and was operat­ing at a dilution rate of 0.02 h-1, then 108 cells/(L*h) would be screened, and a 100-h continuous culture would screen 1010 cells.

The power of this system has been recognized previously (see [143-145] for reviews) and demonstrated in many examples where the enzyme of in­terest is located intracellularly [146-153], including some cases where the limiting enzyme made up 25% of the total cellular protein after selection, an approximately fourfold increase in expression in both cases [154,155]. In a very recent study, the authors were able to create a strain of S. cerevisiae capable of utilizing xylose as the sole carbon source with a 6-h doubling time without using recombinant genetic techniques—only using selection on xylose minimal media from a strain that could grow only very poorly ini­tially [156]. For secreted enzymes (both cell-associated or extracellularly), far fewer studies have shown improvements via selection in liquid culture. Francis and Hansche [157] were able to isolate a mutant of S. cerevisiae in a well-mixed chemostat with 1.7-fold improvement in acid phosphatase activ­ity, and Naki et al. [158] were able to isolate mutants of Bacillus subtilis with about fivefold increased secretion of protease by growing the cells in a hollow fiber apparatus, which physically separated cells, with bovine serum albumin as the limiting nitrogen source. Therefore, understanding the physical char­acteristics of the cell/enzyme/substrate system and the resulting magnitude of differences in growth rate between mutants is critical to applying selection to this system.

When cells are grown on solid media, with significant space between ini­tial cell colonies, those cells that produce more or better cellulase will retain the products of their hydrolytic reactions, and will form larger colonies. This technique—selection by people judging the size of colonies—has the ad­vantage of maintaining separation between cells. It has the disadvantage of limiting the number of cells that can be screened. It is hard to imagine how more than 109 cells (103 colonies/plate* 106 plates) can be screened in a rea­sonable amount of time, even utilizing high-throughput approaches.

When cells are grown in well-mixed liquid culture, the situation is much different because the products of hydrolysis are free to diffuse. A schematic representation of some of the liquid culture cases relevant to recombinant cellulolytic CBP organisms is presented in Fig. 4. In case A, where cellulase enzymes are secreted away from the cell, cellulases with cellulose binding do­mains will diffuse to cellulose, bind to it, and release soluble hydrolysis prod­ucts. In the final step of the overall hydrolysis reaction secreted в-glucosidase converts soluble glucose oligomers into glucose (an overview of fungal cellu­lase systems can be found in [1]). Lelieveld [159] predicted that in cases such as this, the limiting enzyme will form a gradient in the diffusion boundary layer around the cell, creating a gradient of the limiting nutrient as well. Such a gradient would provide a link between mutations conferring increased en­zymatic activity and supply of the limiting nutrient to the cell. With respect to selecting CBP organisms, when a cell secretes a growth limiting cellulase that binds cellulose, it will not necessarily take up the products of the reaction preferentially compared to another cell in the population. Thus, increased activity of that cellulase cannot be selected for. The remaining question is whether the postulated gradient of в-glucosidase exists, and if so what is the effect of the glucose gradient (ДА) on a mutant’s growth rate compared to a parent strain producing less в-glucosidase. Fan et al. [160] used a 2-D reac — tion/diffusion model to predict that the differences in growth rates between mutants and parents in this case are too small to allow the mutant to outgrow the parent in a reasonable length of time.

Recombinant xylanases and cellulases can also be expressed as tethered enzymes [59,119,130] (Fig. 4, cases B and C). In the case where a cell does not bind to the cellulose substrate (case B) (e. g., cellulases with cellulose bind­ing domains are not tethered to the cell surface), the limiting enzyme reaction is once again в-glucosidase conversion of cello-oligomers to glucose. The в — glucosidase enzyme is concentrated at the cell surface, setting up a larger gradient (ДВ) than in case A. However, in this case Fan et al. [160] found that unless the Monod constant (kS) for the substrate was very low, this gradi­ent would not be large enough to allow mutants to outgrow parents in liquid culture.

Case C presents the situation when cellulases are tethered to the surface and the cell binds to a substrate particle. In this case, the particle acts to trap the hydrolysis products, creating a substantial difference between the glucose

concentration in this gap and the substrate and the bulk fluid. When this cell/enzyme/substrate relationship is operative, Fan et al. [160] predict that differences in enzyme expression level will lead to differences in growth rates between mutant and parent cells, and that this will allow selection-based pop­ulation changes to occur in a reasonable amount of time. To date, the promise of selection for improving cellulase production by recombinant cellulolytic microorganisms has not been realized, and knowledge of the local concen­tration of glucose around such cells is limited to prediction. However, it is known that cellulose hydrolysis by naturally occurring cellulolytic microor­ganisms occurs much faster when mediated by cells adhering to the substrate as compared to nonadherent mutants [161]. It has been suggested that adher­ence confers a competitive advantage associated with first access to hydrolysis products.

7

As discussed earlier, the complete genome sequence of Z. mobilis ZM4 has been reported recently [12] following earlier related studies by Korean scien­tists [88-90]. It was found that the genome consists of 2 056416 base pairs forming a circular chromosome with 1998 open reading frames (ORFs) and three ribosomal RNA transcription units. As reported by the authors, “the genome lacks recognizable genes for 6-phosphofructokinase, an essential en­zyme in the Embden-Meyerhof-Parnas pathway, and for two enzymes in the tricarboxylic acid (TCA) pathway, the 2-oxoglutarate complex and malate dehydrogenase. Glucose can be metabolized therefore only by the Entner — Doudoroff pathway”.

Comparison of whole genome microarray data for Z. mobilis ZM1 (ATCC10988) and ZM4 (ATCC 31821) revealed that the 54 ORFs present in ZM4 were absent for ZM1. Four of these ORFs that encode trans­port proteins or permeases, and two that encoded for specific enzymes— NAD(P)H:quinone oxidoreductase and an oxidoreductase related to short — chain alcohol dehydrogenases, were found to be highly expressed in Z. mobilis ZM4. The authors suggested that it is possible these genes relate to the higher specific rates of sugar uptake and ethanol production for ZM4 when compared to ZM1. They also reported that two genes encoding capsular carbohydrate synthesis enzymes were only actively expressed in ZM4 and may contribute to its relatively high resistance to increased osmotic pressure found in high sugar solutions (e. g. in 250-300 g L-1 glucose media).

4

The enzymatic hydrolysis of the pretreated raw material and the fermenta­tion of the hydrolysed sugars can be performed separately or simultaneously, commonly referred to as SHF (separate hydrolysis and fermentation) or as SSF (simultaneous saccharification and fermentation). The SSF process con­figuration has been generally considered more favourable for reducing the ethanol production costs [72,81]. The hydrolysis rate in the separate hydro­lysis is strongly inhibited by the accumulation of the end products, cellobiose and glucose [60]. In the simultaneous hydrolysis and fermentation, the end product inhibition is alleviated by the continuous removal of glucose by the fermenting organism. In the separate hydrolysis and fermentation the most severe end product inhibition caused by cellobiose has been overcome by adding an adequately high amount of в-glucosidase. For the same reason, the enzyme dosage needed is obviously lower in the SSF. Other claimed ad­vantages of the SSF are the lower risk of contamination and reduction of investment costs by combined reactors. The low concentration of free glucose and the presence of ethanol make it more difficult for contaminating micro­organisms to take over the fermentation and decrease the ethanol yield. The drawback of the SSF is that the conditions, i. e. the pH and temperature of the hydrolysis and fermentation, are suboptimal in a combined process. The optimal temperature for the enzymatic hydrolysis is clearly higher than that of the presently used fermenting organisms. Another drawback of the SSF is the difficulty in optimising the fermentation of techniques, i. e. by running continuous fermentation or recirculating and reusing the yeast due to the presence of the solid residues from the hydrolysis.

To improve the overall process economics and to achieve a faster hydrolysis rate by using thermostable enzymes, various modifications of the present process configurations can be considered (Fig. 1). After the pretreatment, the temperature of the substrate is high, and is reduced to achieve the operat­ing temperature in the following process stages. In the traditional SSF, the temperature is about 35 °C. In a separate hydrolysis and fermentation pro­cess, the first total hydrolysis stage is carried out at about 45-50 °C with the present commercial enzymes, or above 60 °C with novel thermostable

enzymes. Other options include a partial prehydrolysis at higher tempera­tures, denoted as liquefaction, where the viscosity of the substrate is de­creased using a chosen composition of thermostable cellulases based on one or several enzymes. The liquefaction stage, i. e. an enzymatic treatment improving the rheological properties (improved flowability, reduced viscos­ity) of the slurry, can significantly improve the mixing properties of the substrate slurry [83]. This partial hydrolysis can be carried out even with the limited number of thermostable cellulolytic and hemicellulolytic en­zymes available. Using a set of thermoactive enzymes in the prehydrolysis, it was possible to reduce the viscosity and increase the sugar formation [83]. The high viscosity is a consequence of a high initial substrate consistency, needed to achieve a high final sugar and ethanol concentration and to de­crease the distillation costs [69]. With a theoretical ethanol yield of 25-30% of the raw material, the raw material consistency should be at least 15% (d. w.) to reach an ethanol concentration of 4-5%. Some of the technical obstacles related to high consistency can thus be overcome by a rapid de­crease of viscosity. After a liquefying partial hydrolysis, the saccharification stage using a complete or complementary set of hydrolytic enzymes, ei­ther simultaneously or separately from the fermentation (SSF or SHF), can be carried out. A separate hydrolysis stage (SHF) can be carried out at el­evated temperatures with the complete set of hydrolytic thermostable en­zymes needed for a chosen substrate. Finally, thermostable enzymes could be supplemented to bacterial fermentations using anaerobic, ethanol pro­ducing strains, such as Clostridia, to improve their conversion rate of cellu — losic substrates into sugars (SSF or consolidated bioprocessing). Thus, new thermostable enzymes would allow the design of more flexible process con-

figurations, based on the availability of novel thermostable lignocellulolytic enzymes.

The performance of chosen thermostable cellulolytic enzymes with present commercial fungal enzymes was compared in this paper. The refer­ence enzyme preparations contain the whole set of cellulolytic enzymes, i. e. cellobiohydrolases and endoglucanases, as well as several hemicellulolytic ac­tivities and в-glucosidases. These enzymes work at temperatures up to about 45 °C in long-term hydrolysis conditions and up to 50 °C in short-term con­ditions. New enzyme compositions were designed and tested in the hydrolysis of various steam pretreated raw materials.

5

In conclusion, the domestication of S. cerevisiae for carbon dioxide and ethanol formation from hexose sugars has led to the fact that the metabolism of hexose and pentose sugars in this yeast are fundamentally different. As evidenced by genome-scale transcriptome and proteome analyses of numer­ous recombinant pentose-utilizing S. cerevisiae strains, the difference is not only limited to the initial sugar conversion pathways, but also comprises the central metabolism and the glycolytic pathway. The major future challenge remains to translate the knowledge acquired from laboratory strains to indus­trial production strains.

Acknowledgements The authors acknowledge the financial support from the Swedish En­ergy Agency.