Like E. coli, Klebsiella oxytoca is able to metabolize a variety of biomass — derived monomeric sugars, but unlike E. coli it also has the native ability to transport and metabolize cellulose subunits cellobiose and cellotriose [50, 51]. The PET operon was expressed in K. oxytoca concurrent with the ori­ginal E. coli work [52] and was later chromosomally integrated, resulting in strain P2 [51]. Ethanol production by K. oxytoca P2 from various substrates has been reported (Table 1) [53-56]. K. oxytoca strain BW21, which was de­rived from strain P2 by elimination of the butanediol pathway, produces over 40 g L-1 ethanol in 48 h in OUM1 medium. OUM1 is a medium designed spe­cifically for K. oxytoca, as described below [57].

When Z. mobilis was selected as the source of the PET operon, Z. mo — bilis and Gram-positive Sarcina ventriculi were the only known bacteria with PDC activity [58-61]. Since that time, PDC activity has been identified in other bacteria, including Gram-negative Acetobacter pasteurianus [62] and Zymobacter palmae [3]. The Z. palmae PDC has a higher specificity and lower pyruvate Km than Z. mobilis, S. ventriculi, and A. pasteurianus [63]. The S. ventriculi PDC is different from the Z. mobilis enzyme but is highly related to the PDC found in fungi; its expression in E. coli requires the presence of accessory tRNA due to differences in codon usage [63]. In A. pasteurianus, PDC seems to have the unusual role of functioning in an aerobic pathway, contributing primarily to the conversion of pyruvate to acetaldehyde [62].

The robustness of Gram-positive organisms is appealing for industrial ap­plications, but initial attempts to express the Z. mobilis homoethanol pathway in Bacillus and lactic acid bacteria had limited success [64-67]. However, the discovery of new PDC forms has enabled renewed engineering attempts. The PDCs from S. ventriculi, A. pasteurianus, and Z. mobilis, as well as S. cerevisiae, were each expressed in Bacillus megaterium, with the S. ven — triculi PDC showing the highest activity. When coupled with ADH from Geobacillus stearothermophilus, the S. ventriculi PDC enabled B. megaterium to convert 13.2 gL-1 pyruvate to 3.3 gL-1 ethanol, a tenfold increase rela­tive to strains lacking PDC [68]. The S. ventriculi PDC was also expressed in Lactobacillus plantarum, with production of up to 6 g L-1 ethanol from 40gL-1 glucose [69]. Recent attempts to express the Z. mobilis pathway in Corynebacterium glutamicum have resulted in production of ethanol from glucose [70].

Considerable effort has been extended to engineering of Z. mobilis for im­proved ethanol production, as covered elsewhere in this volume. However, the ethanol titers attained by ethanologenic E. coli LY168 in minimal medium exceed published values for Z. mobilis in rich medium (Table 2).

3

The hydrolysate released from the pretreatment is typically treated with en­zymes in order to break down cellulose and hemicellulose into hexoses and pentoses that are then further fermented to ethanol. Enzyme costs are, how­ever, generally high, so that the search for new enzymes with high efficiency that can be produced at low costs is the key to overcome the bottleneck of this process step [18] (see also Viikari, this volume). Another possible way to reduce treatment costs is to implement recycle loops in order to feed back­washed enzymes into the vessel of enzymatic hydrolysis [19,20]. Enzymes could further be produced at the plant using a stream of the pretreated ma­terial in an on-site enzyme production.

The hydrolysis step is optimized by performing the enzyme treatment to­gether with yeast fermentation of glucose (simultaneous saccharification and fermentation, SSF). The temperature optimum of the enzymes is, however, of­ten higher than the optimum for yeast. This can depress the advantages of SSF compared to separation of the two processes.

4.3

In naturally xylose-utilizing bacteria, D-xylose is isomerized to D-xylulose [18] by xylose isomerase (XI). Xylulose is then phosphorylated to xylulose 5-phosphate [19], which is an intermediate of the pentose phosphate pathway (PPP). A similar pathway has been found in an anaerobic fungus [20]; how­ever, most naturally xylose-utilizing fungi contain a more complex pathway consisting of reduction-oxidation reactions involving the cofactors NAD(P)H and NAD(P)+ (Fig. 1). Xylose is reduced to xylitol [21-23] by a NAD(P)H — dependent xylose reductase (XR), and xylitol is then oxidized to D-xylulose by a NAD+-dependent xylitol dehydrogenase (XDH) [22,24,25]. As in bacte­ria, xylulose is phosphorylated to D-xylulose 5-phosphate by a xylulokinase (XK) [26,27]. Despite the inability of S. cerevisiae to utilize xylose, the genes encoding the reductive-oxidative xylose pathway enzymes XR, XDH, and XK are present in its genome [26,28,29]; however, they are expressed at too low levels to allow xylose utilization. Even when the genes were overexpressed, no growth on xylose could be detected [30]. Neither was it possible through adaptation protocols to upregulate the expression of these genes to levels high enough to allow significant xylose fermentation [31].

2.2

After the proof of principle of XI expression in S. cerevisiae, not only metabolic engineering, but also evolutionary engineering was applied to im­prove the rate of D-xylose utilisation of a strain solely over-expressing XI [44]. Since improvement of the aerobic consumption rate was initially the target of this selection experiment, serial transfer in a shake flask was chosen as the cultivation condition of this evolution run. Indeed, after 30 serial transfers, the specific growth rate of this culture improved drastically (24-fold) from 0.005 h-1 to 0.12 h-1 (Fig. 8). However, a strain isolated from this selection ex­periment was not yet capable of anaerobic growth. Therefore, an additional ten selection rounds were performed in oxygen-limited batch cultures, fi­nally followed by ten cycles in an anaerobic sequencing batch reactor. From this culture a single colony was isolated (named RWB 202-AFX, for anaero­bic fermentation of D-xylose based on strain RWB 202) and used for further characterisation of the end product of this evolutionary engineering.

It was shown that only the expression of a XI, followed by evolutionary en­gineering for anaerobic growth, can also result in a S. cerevisiae strain that can grow on 2% D-xylose as the sole carbon source, with a growth rate of 0.03 h-1 in anaerobic batch fermentations [45]. However, although this strain displayed a good ethanol yield on D-xylose (0.42 gg-1) and very low pro­duction of xylitol (2.8 mM), the obtained growth rate, and therefore ethanol

production rate, were insufficient to allow economically viable industrial ap­plication. During these batch cultivations, small amounts of D-xylulose (up to 8 mM) were still excreted into the broth, indicating that evolutionary engin­eering alone did not fully overcome the metabolic limitations downstream of this metabolite. This result indicates that although evolutionary engineering is a very powerful tool, it has limitations and, in this case, the combination of knowledge-based metabolic engineering (Sect. 5) combined with evolution­ary engineering (Sect. 6.1) resulted in more desirable attributes and higher ethanol production rates.

7

L-Alanine, used in the pharmaceutical industry [150] and as a food addi­tive [151], is commercially produced by enzymatic decarboxylation of L-aspartic acid with either immobilized cells or cells suspensions [152]. How­ever, recent attention has shifted to fermentative production [150,151].

Given our success in producing microbial biocatalysts by a combination of directed engineering and metabolic evolution, we modified lactic-acid produc­ing E. coli B derivative SZ194 for alanine production. The native IdhA gene was replaced with the ribosome binding site, coding region, and transcriptional terminator of the thermostable alanine dehydrogenase alaD from Geobacillus stearothermophilus XL-65-6 (Zhang et al., unpublished results). While the ini­tial microbial biocatalyst was capable of producing l-alanine as the primary fermentation product, long incubation times were required and the productiv­ity was low. As with other microbial biocatalysts designed in our laboratory, metabolic evolution was used for strain improvement. The strain was further engineered to reduce co-product formation (mgsA) and increase the chiral purity (dadX). The final microbial biocatalyst, XZ132, produced near 1.3 M l — alanine from 12%glucose within 48 h, a yield greater than 95%, in AM1 mineral salts media.

6

One approach to reducing the production cost is integration of ethanol pro­duction with another suitable plant, e. g. a combined heat and power plant, a starch-based ethanol plant or a pulp and paper mill. Significant reduction of the production cost was obtained in a study on co-production of ethanol and electricity from softwood, based on conditions in California, USA [44]. One of the benefits is that the syrup or lignin residue can be used for steam pro­duction without prior drying. Another option is to integrate cellulosic ethanol production with starch-based ethanol production to utilize the whole agri­cultural crop. This will increase the production capacity drastically, and it may also help to boost the ethanol concentration resulting from the ligno — cellulosic process, if the ethanol-containing streams can be distilled in the same distillation units. This will have a beneficial effect on the energy de­mands in the distillation and evaporation steps. It might be a disadvantage if the residue cannot be used for animal feed (DDGS). However, it will still have a fuel value, which will help to improve the economics of the overall process. The biorefinery concept is also an interesting option. Using chem­ical and biological transformations, the raw material is processed to produce ethanol and, e. g., modified lignin, specialty chemicals and maybe anaerobic biogas, adding value to the main product. In this case the income from other products improves the overall process economics [45,46].

4

Conclusions

Flowsheeting, combined with estimates of the production cost, is a valuable tool for the comparison of process alternatives and to determine bottlenecks that require further improvement. It is, however, difficult to compare pro­duction costs from different studies due to the many assumptions made in the simulations, such as ethanol yield, productivity and concentration, as no commercial-scale plants are in existence. Also, differences in capacity and cost of raw material, as well as currency exchange rates, add to the uncer­tainty. This is clearly illustrated by the large variation in the estimated ethanol production cost, from 0.13 to 0.81 US$ L-1 ethanol.

The most important parameters for the economic outcome are the feed­stock cost, which varied between 30 and 90 US$ per metric ton, and the plant capacity, which influences the capital cost. It is thus very important to reach a high overall ethanol yield as this is directly related to feedstock and capital costs for a given production capacity.

One of the major research challenges is to improve the hydrolysis of carbo­hydrates through more efficient and less expensive pretreatment methods, but also by enhanced enzymatic hydrolysis with superior enzymes at a reduced enzyme production cost. The latter is one of the most uncertain costs in most economic analyses.

It is also important to achieve a high ethanol concentration in the fer­mentation or SSF steps to reduce the energy demand. This requires new technology for enzymatic hydrolysis (or SSF) at high solids concentrations and the development of robust fermenting organisms that are more toler­ant to inhibitors. They also have to be able to ferment all sugars in the raw material in concentrated hydrolyzates, while maintaining high ethanol pro­ductivity and a high ethanol concentration.

Finally, process integration within the process and with other types of in­dustrial processes, e. g. a combined heat and power plant or a starch-based ethanol plant, will reduce the production cost further. Regarding the immedi­ate future, we believe that these integrated plant concepts will be used in the first successful industrial-scale production of lignocellulosic fuel ethanol.

Adv Biochem Engin/Biotechnol (2007) 108: 329-357 DOI 10.1007/10_2007_059 © Springer-Verlag Berlin Heidelberg Published online: 11 April 2007

Vancouver, British Columbia V6T 1Z4, Canada warren. mabee@ubc. ca

1 Introduction……………………………………………………………………………………………… 330

2 Biofuel Production…………………………………………………………………………………….. 331

2.1 Brazil……………………………………………………………………………………………………….. 333

2.2 United States…………………………………………………………………………………………….. 334

2.3 European Union………………………………………………………………………………………… 337

2.4 Other Biofuel Producing Nations………………………………………………………………… 341

3 Direct Funding Programs in the USA………………………………………………………….. 343

4 Excise Tax Exemptions in the USA…………………………………………………………….. 347

5 Political Goals and Bioethanol-Related Policy………………………………………………. 350

6 Conclusions………………………………………………………………………………………………. 351

References ……………………………………………………………………………………………………. 354

Abstract Biofuels for use in the transportation sector have been produced on a signifi­cant scale since the 1970s, using a variety of technologies. The biofuels widely available today are predominantly sugar — and starch-based bioethanol, and oilseed — and waste oil — based biodiesel, although new technologies under development may allow the use of lignocellulosic feedstocks. Measures to promote the use of biofuels include renewable fuel mandates, tax incentives, and direct funding for capital projects or fleet upgrades. This paper provides a review of the policies behind the successful establishment of the bio­fuel industry in countries around the world. The impact of direct funding programs and excise tax exemptions are examined using the United States as a case study. It is found that the success of five major bioethanol producing states (Illinois, Iowa, Nebraska, South Dakota, and Minnesota) is closely related to the presence of funding designed to support the industry in its start-up phase, while tax exemptions on bioethanol use do not influ­ence the development of production capacity. The study concludes that successful policy interventions can take many forms, but that success is equally dependent upon external factors, which include biomass availability, an active industry, and competitive energy prices.

Keywords Biofuels • Direct funding • Excise tax exemptions • Policy •

Renewable fuel mandates

1

Introduction

Biofuels derived from sustainable biological sources, including agricultural crops, waste vegetable oils, and woody biomass, is advocated by many in­cluding MacLean et al. [1] and McMillan [2] as a potential substitute for petroleum-derived fuels such as gasoline and diesel. The use of biofuels is generally associated with lower greenhouse gas emissions and improved en­ergy balance compared to petroleum-based fuels [3], which makes them an attractive option for combating climate change and meeting national or in­ternational targets of environmental performance. As the biofuel industry is based on agricultural (or potentially forest) biomass, development of the in­dustry will lead to a diversified rural economy and increased employment, which can support domestic development goals [4-6]. The industry has long been promoted as a means to substitute renewable, sustainable biomass for fossil reserves of oil, which may in turn increase the security of energy sup­plies and reduce dependence upon foreign oil [7]. These attributes make biofuel an attractive option for policymakers, offering solutions to a number of domestic challenges. At the same time, policy is needed in order to increase the competitiveness of bio-based fuels, which are generally more expensive to produce than petroleum-based counterparts [8].

Policy options to support biofuel production may take a number of forms. Some options are “top-down” in form, as they are enacted on a national or regional basis and impact all producers and consumers. One such option is the national target, in which policymakers make a public declaration of their intention to meet a certain level of production (often expressed as a percent­age of overall production) in domestic transportation fuel supply. Top-down policy places the emphasis upon governments, which are then responsible for creating an environment supportive towards industrial expansion. The national target should not be confused with a renewable fuels standard (or obligation), which sets legal standards for the minimum levels at which bio­fuels must be blended into transportation fuels. A renewable fuel standard places the emphasis upon industry, who must then meet the renewable fuel standards with their products in order to be eligible for sale. One commonly observed policy option is exemption of biofuels from national excise taxation schemes, which has the effect of reducing producer costs and thus increasing potential profits. This type of incentive can be identified as a subsidy to indus­try, although lower prices can be and are passed to consumers in competitive markets.

Other policy options act in a “bottom-up” fashion, impacting only par­ticular industrial or consumer participants in the biofuel marketplace. One such option is direct government funding of capital projects to increase cap­acity or upgrade distribution networks. Normally, these types of policies are enacted in a competitive fashion, wherein various industrial producers can compete for projects, which are then carried out in conjunction with govern­ment. Another bottom-up type of policy is targeted at increasing biofuel use in government or corporate vehicle fleets.

In some countries, multiple policies covering the range of options de­scribed above have been enacted to support biofuel development (e. g. [9-11]). The presence of multiple policies within these jurisdictions means that de­termining the effectiveness of individual policies is quite difficult. In this paper, implementation of biofuels in several countries is examined. The abil­ity of two measures to promote domestic biofuel production is compared. The first measure considered is exemptions on fuel excise taxes; the second is funding designed to support projects, infrastructure, or capacity develop­ment for bioethanol production. The industry is then evaluated on its ability to successfully promote broad policy goals of employment, environmental performance, and fuel security. A number of recommendations for the for­mulation of future policies are proposed.

2

In contrast to the energy-conserving function of glycolysis, the main metabolic function of the PPP is to provide anabolic intermediates such as ribulose 5-phosphate, erythrose 4-phosphate, and NADPH for biosynthesis and cell growth. The flux through the nonoxidative PPP in S. cerevisiae was found to be much lower than in other yeasts [105], which was later con­firmed by metabolome analysis [106]. The low PPP activity in S. cerevisiae is sometimes interpreted to be a result of the domestication of S. cerevisiae by prolonged selection for carbon dioxide and ethanol production from hexose sugars. However, PPP activity is a crucial part of pentose metabolism, since it is virtually the only way to introduce xylulose into the central metabolism. It was early pointed out that the PPP activity may limit xylose metabolism in S. cerevisiae [47,107], which was further supported when excretion of PPP intermediates was observed in xylulose — and xylose-metabolizing S. cere­visiae [43,108].

The insufficient flux through the nonoxidative PPP in S. cerevisiae has been indirectly confirmed in several genome-scale and enzymatic analyses of mutant strains with improved xylose metabolism, where invariably ei­ther the transaldolase (TAL1) or the transketolase (TKL1) genes, or both, have been found to be upregulated [57,71,91,109-111]. Directly, the im­portance of the flux through the PPP has been confirmed by the superior pentose utilization and ethanolic fermentation by strains in which the en­zymes of the nonoxidative PPP have been overexpressed. An early attempt to overexpress P. stipitis transketolase in xylose-metabolizing S. cerevisiae was not successful [112], whereas overexpression of the endogenous S. cere­visiae transaldolase (TAL1) resulted in improved growth on xylose [78]. Later, the overexpression of all four nonoxidative PPP genes, including not only TAL1 and TKL1 but also ribulose-5-phosphate 4-epimerase (RPE1) and ribu — lokinase (RKI1), was shown to improve xylulose consumption by S. cere­visiae [90,113] (strain TMB3026, Table 3). Moreover, the improvement result­ing from the overexpression of the four genes was higher than when each gene was overexpressed alone [90]. The simultaneous overexpression of the whole nonoxidative PPP, together with GRE3 deletion, allowed growth on xylose in a strain carrying a bacterial XI [42] (strain TMB3050, Table 2). The usefulness of this combination of modifications was confirmed when it allowed aerobic and anaerobic growth on xylose in a strain carrying the Piromyces XI [97] (strain RWB217, Table 1). PPP overexpression also al­lowed superior xylose fermentation rates in combination with high levels of XR and XDH [42,54] (cf. strains TMB3057, TMB3056, and TMB3062, Table 1 and Fig. 5).

4.5

Significant advances related to recombinant enzyme expression support the potential for S. cerevisiae as a CBP host. However, the challenge of integrating all the different aspects of enzymatic hydrolysis of cellulose and hemicellu — lose and subsequent fermentation of the sugars released to ethanol in a single reactor with a CBP should not be underestimated. A pertinent question of­ten asked by critics is, “Would S. cerevisiae be able to simultaneously express multiple genes, while producing and secreting the different cellulases, hemi- cellulases, and pentose utilizing enzymes required?” Several studies demon­strate coexpression of multiple genes in S. cerevisiae, for example in the case of the expression of tethered cellulolytic and xylanolytic enzymes [59,119], xylose and arabinose utilizing enzymes [40,162], as well as xylose and cel — looligosaccharide utilizing enzymes [45]. The expression and secretion of a variety of cellulases, amylases, and pectinase has also been demonstrated without adversely affecting yeast growth [51, 52].

However, the number of genes expressed is probably not as important a challenge as the need for high-level expression as well as the stress re­sponses that may accompany such high-level expression. Factors that may impose unnecessary stress on the cell are (1) sequestering of transcription factors at highly expressed promoters used for heterologous gene expression,

(2) impact of unfavorable codon bias on the translation of heterologous pro­tein (can be overcome by the use of codon-optimized synthetic genes), and

(3) improper folding of foreign proteins that can evoke the (4) unfolded pro­tein response (UPR) and consequently the endoplasmic reticulum-associated protein degradation (ERAD) response [163]. Some of these effects may be exacerbated by (5) interrupted transport of foreign proteins through the se­cretion pathway, or (6) accumulation of larger proteins at the cell wall due to low permeability [164]. The answer would thus not be simply overexpression of all the required genes to ensure a functional CBP yeast with the desirable enzymatic activities, but much more attention should also be devoted to the careful manipulation of the required enzyme activities and producing them at the right concentration to provide functionality without exerting too much unnecessary stress on the CBP yeast.

Essentially all work carried out thus far involving heterologous expres­sion of saccharolytic enzymes in yeast has involved laboratory strains. Much of this work has to be transferred to industrial strains that provide the fer­mentation capacity and robustness desired for industrial processes. Different strategies have been used for the overexpression of multiple genes in in­dustrial S. cerevisiae strains. High copy-number episomal YEp vectors, often using the two-micron autonomous replicating sequence (ARS), have been very helpful in demonstrating proof of concept in laboratory strains of S. cere­visiae [43,51,102,115]. However, these vectors are usually mitotically unsta­ble and require selection for the episomal plasmid, which often means using a defined medium that is not applicable to industrial uses [164]. The preferred route taken for industrial strains has been the use of integrative YIp vectors that facilitate direct integration of foreign expression cassettes into a target gene on the yeast genome [165,166] or recycling dominant selectable markers for multiple integration [167-170]. Although these methods provide stable expression from the yeast genome and are amendable to industrial strains, the major drawback has been low expression levels and often not delivering high enough quantities of the required gene product.

Different approaches have been pursued in an attempted to combine the advantages of overexpression from multicopy plasmids with the stability of chromosomal integration, which is also applicable to industrial strains when dominant selectable markers are used. These include the use of repetitive chromosomal DNA sequences such as rDNA [171] and 8-sequences [172]. There are approximately 140-200 copies of rDNA existing in the haploid yeast genome; however, rDNA is located in the nucleolus, which may af­fect the accessibility to RNA polymerase II transcription. Also, the size of pMIRY (multiple integration into ribosomal DNA in yeast) vectors could determine the mitotic stability of these multiple integrations [173]. The 8- sequences are the long terminal repeats of S cerevisiae retrotransposon Ty. More than 400 copies of 8-sequences can exist either Ty associated or as sole sites in the haploid yeast genome [174]. 8-Integration thus makes it possible to integrate more copies of a gene of interest into the yeast genome than the conventional integration systems [78,175]. Host strains and integrated gene size can significantly affect the transformation efficiency at 8-sequences; however, the transformation frequency can be 10- to 100-fold those ob­tained when transforming with vectors that target a single gene on the yeast genome [176].

Although the necessary tools exist for multiple and repeated integration of genes of interest into the genome of industrial strains to complement the required features for CBP (Table 1), a more strategic approach would be re­quired to design a yeast that produces the required enzyme activities, yet re­tains the competence to still perform well under industrial conditions for the economic conversion of plant biomass to ethanol. Such an approach will most probably start by building on a platform industrial yeast that cometabolises hexoses and pentoses, and subsequently finding the right combination and level of expression for saccharolytic enzymes. This approach will inevitably use reiterated metabolic engineering and flux analysis, selection and mutage­nesis strategies, and even strain breeding to allow the microorganism itself to overcome rate-limiting hurdles toward developing an efficient CBP yeast. Ex­amples of such approaches in the past have been performed to enhance xylose fermentation in laboratory and industrial strains [33,37,39,177].

4.1

Metabolites and Related Products

The production of a range of byproducts from Z. mobilis is reviewed compre­hensively by Johns et al. [6] and Panesar et al. [8] with the former authors identifying potential commercial opportunities for the following products: fructose (using sucrose and a fructokinase negative mutant), sorbitol and glu­conic acid, levan (a fructose polymer), fructo-oligosaccharides and various enzymes. As pointed out in a review by Scopes [91], Z. mobilis is a rich source (on an enzyme content per g cell basis) of many of the enzymes currently used in diagnostic analysis and research. Interestingly, in other studies by Park et al. [92], it was established that the activities of some of the key ED enzymes (e. g. glucokinase, G-6-phosphate dehydrogenase) were unaffected by the relatively high ethanol concentrations produced during fermentation, while the activity of an enzyme such as transketolase decreased appreciably above ethanol concentrations of 60 gL-1. It has been estimated that for ac­tively growing cells, as much as 30-50% of the cellular protein is comprised of ED enzymes [93]. However, the greatest difficulty for the commercial pro­duction of such enzymes is the low cell yield of Z. mobilis which is typically 0.02-0.03 gg-1 substrate sugar, compared to cell yields close to 0.5 gg-1 for many aerobically grown microorganisms.

4.2

The thermostable enzyme preparations were kindly provided by ROAL, Fin­land. The genes encoding three thermostable enzymes: cellobiohydrolase (CBH/Cel7A) from Thermoascus aurantiacus, fused with the T. reesei CBHI cel­lulose binding domain (CBM), endoglucanase (EG/Cel45A) from Acremonium thermophilum and a xylanase and в-glucosidase (BG/Cel3A) from T. auran­tiacus were inserted under the control of a strong T. reesei cbhl promoter and transformed into a host strain where all the major cellulase genes were deleted (phenotype CBHI/Cel7A — CBHII — Cel6A — EGI/Cel7B — EGII/Cel5A). Fermenter supernatants produced at pilot scale were used to produce vari­ous mixtures of the thermostable enzymes. The background activities in the deletion strains were measured. The composition of the mixture of the three thermostable enzymes was optimised based on the average cellulase compo­sition of T. reesei. The enzyme components were mixed in different ratios and the total cellulase activity of the mixtures was measured at 50 °C(as FPU mL-1) and used as the basis of enzyme dosing. In addition, a family 10 thermostable xylanase from T. aurantiacus, cloned and expressed in the T. reesei deletion strain, was added to some preparations to ensure complete hydrolysis.

Celluclast 1.5 L FG (Novozymes, Denmark) and Econase CE (ABEnzymes, Finland), eventually supplemented with BG from Novozym 188 (Novozymes, Denmark) were used as reference enzymes. The standard enzyme dosage was 10 FPU g-1 cellulose for Celluclast 1.5 L FG, supplemented with additional BG (100-500 nkatg-1 cellulose). Assuming an average 50% cellulose content of the lignocellulose substrates, the enzyme dosage was thus 5 FPU g-1 substrate. For the hydrolysis experiments at elevated temperatures, higher dosage of Celluclast (22 FPU g-1 cellulose) was used.

The total cellulolytic activity used as a basis for dosing of the enzyme mixtures was evaluated using the FPU activity, measured against Whatman no. 1 filter paper [36]. The EG activity was assayed using hydroxyl ethyl cel­lulose as substrate [36]. The CBH activity was determined by using 4-methyl umbelliferyl-b-D-lactoside as substrate, estimating the effect of EGs by carry­ing out the assay with or without 20 mM cellobiose in the reaction [6]. The xy- lanase activity was assayed using birchwood glucuronoxylan as substrate [4] and that of BG using p-nitrophenyl-b-D-glucopyranosidase as substrate [5]. Protein was assayed according Lowry et al. [43]. All the enzyme activity assays were carried out at pH 5.

Substrates

The substrates used were steam pretreated, washed spruce solid fraction kindly provided by Guido Zacchi at the Lund University, Sweden, and steam pretreated corn stover kindly provided by Francesco Zimbardi at ENEA, Italy. The solid fraction of spruce substrate after the pretreatment was sep­arated from the liquid fraction by filtration, washed and used in the hydro­lysis experiments. The composition of the fibre fractions of the substrates is presented in Table 2. In addition to the insoluble fibre fraction, the corn stover substrate contained significant amounts, about 17% (d. w.) of solu­bilised mono — and oligosaccharides, solubilised mainly from hemicelluloses. Based on secondary analytical enzymatic hydrolysis and HPLC analysis, the carbohydrates in the soluble fraction consisted of xylose (74%), arabinose (15%), galactose (5%) and glucose (6%). Comparative hydrolysis experiments were carried out using crystalline cellulose (Avicel, Sigma).