Advances in the fields of genomics and metagenomics have dramatically re­vised our view of microbial biodiversity and the potential for biotechnological applications. In the last decade the revolution in computer processor speeds, memory storage capability, and expanding networks has made possible the large scale sequencing of genomes and management of large integrated databases over the Internet. Since the first microorganism, Haemophilus in­fluenzae, was sequenced in 1995, genome sequencing initiatives have resulted in over 300 sequenced organisms, including 27 archaeal, 337 bacterial, and 41 eukaryotic genomes. As of July 2006, more than 1500 prokaryotic and eukaryotic genome sequencing projects are underway [70,71]. The genome sequences of Escherichia coli and Saccharomyces cerevisiae were not only among the first to be published, but were also the first of wide relevance for the production of industrial biochemicals, including bioethanol. Given that the genome of a particular microorganism, following annotation, provides the theoretical enzyme reaction set, it serves as a preferred starting point for engineering metabolic pathways that will lead to significantly improved titer, yield, productivity, and performance of a microorganism [62].

Annotated genomes certainly compliment experimental designs; however, the design space that can be considered by visual inspection or classical hy­pothesis driven experimentation is limited given the high degree of connec­tivity of the metabolic network. Modifying a given enzyme or metabolite pool is likely to elicit a multilayered regulatory response that not only mitigates the original perturbation, but will shift the equilibrium of other enzymes, metabolite pools, or signalling pathways. To a large extent, this is why ran­dom mutagenesis approaches have been favored over targeted approaches, until recently. The first genome-scale in silico metabolic network model for E. coli was made available in 2000 and was among the first to demonstrate consistency between modeling predictions and in vivo physiology [72,73]. Specifically, the model was used to explore the relationship between acetate, succinate, and oxygen uptake rates when attempting to maximize growth rate, to confirm the hypothesis that E. coli under acetate and succinate car­bon limitations regulates its metabolic network to maximize growth rate. For industrial biotechnology process development, it is often desirable to shift carbon flux from biomass to product formation, thereby maximizing the yield of product on substrate.

The first eukaryotic genome-scale metabolic model was reported in S. cere — visiae in 2003 based on its annotated genome sequence and a thorough examination of online pathway databases, biochemistry textbooks, and jour­nal publications [74]. This genome-scale in silico model, by using a relatively simple synthetic medium, could predict 88% of the growth phenotypes cor­rectly, indicating that this model in many cases can predict cellular behavior. In one step further, Duarte et al. (2004) [74] used the S. cerevisiae genome — scale metabolic network constructed by Forster et al. (2003) [75] to generate a phenotypic phase plane (PhPP) analysis that describes yeast’s metabolic states at various levels of glucose and oxygen availability. Examination of the S. cerevisiae PhPP has led to the identification of two lines of optimal­ity: LOgrowth, which represents optimal biomass production during aerobic, glucose-limited growth, and LOethanol, which corresponds to both maximal ethanol production and optimal growth during microaerobic conditions. The predictions of the S. cerevisiae PhPP and genome scale model were compared to independent experimental data, and the results showed strong agreement between the computed and measured specific growth rates, uptake rates, and secretion rates. Thus, genome-scale in silico models can be used to system­atically reconcile existing data available for S. cerevisiae, particularly now that yeast resources, databases, and tools for global analysis of genomic data have been expanded and made publicly available, such as the Saccharomyces Genome Database [70,71].

Another major challenge of current biotechnology, especially in the lignocellulose-to-ethanol process, is to identify novel biocatalysts and en­zymes for enzymatic hydrolysis from the genomes of organisms and metage­nomic libraries. A large number of protein sequences deduced from the genomes of industrial microorganisms have no assigned function, as they exhibit low (or no) homology to the enzymes and/or proteins functionally characterized thus far [76]. The demand for identification of novel biomass­degrading enzymes as well as for heterologous protein production at higher efficiencies and reduced costs has catalyzed an interest in elucidating the genomic sequence of Trichoderma reesei — the most prolific producer of biomass-degrading enzymes. Diener et al. (2004) [77] has described the creation of a T. reesei strain QM6A large-insert BAC (bacterial artificial chro­mosome) library and its subsequent analysis, which was successfully used to identify both biomass degradation and secretion related genes. These data re­vealed the utility of a BAC library for use in assembly of the T. reesei genome and isolation of genomic sequences of industrial interest.

Even though the above study represents a direct application of sequenc­ing technology for identification of novel biomass-degrading enzymes, it is also often the case to use such high-throughput experimental techniques to elucidate mechanistic understanding of enzymes derived from random, nat­ural selective pressures. The research of Foreman et al. (2003) [78] using

T. reesei RL-P37, a strain that has been selected for improved production of

cellulolytic enzymes [79], is such an example. The mutation(s) that improved cellulase production concurrently improved the inducible expression of ancil­lary genes that do not have a known function in cellulose degradation. These results suggest significant regulatory points of convergence across the spec­trum of cellular processes involved in carbon sensing, signal transduction, and transcriptional regulation. These findings will likely have significant im­plications for the design of industrial processes for commercial production of biomass-degrading enzymes.

In conclusion, the vastly improved computational capability integrated with large-scale miniaturization of biochemical techniques such as BAC, PCR, and microarray chips has delivered significant amounts of genomic data to researchers all over the world [80]. This availability of data has led to an ex­plosion of genome analysis leading to many new discoveries and tools that are not possible in exclusively wet-lab experiments.

It is evident from the above applications of genomics coupled to in silico modeling that industrial biotechnology, and especially bioethanol produc­tion, can benefit from this technology platform both in the identification of metabolic engineering target genes to improve yields, titer, and productivity, and in the discovery of novel enzymatic catalysts. This is further reinforced by the various case studies to be presented in subsequent chapters, including the role genomics has played in the identification of thermostable cellulases, metabolic engineering for pentose and xylose utilization in S. cerevisiae and Pichia stipitis, development of ethanologenic bacteria, and development of Z. mobilis for bioethanol production.

3.2

While possible variations in the process of converting lignocellulosic biomass to ethanol are virtually endless, it can most simply be described as the in­tegration of five unit operations: (1) desizing, (2) thermochemical pretreat­ment, (3) enzymatic hydrolysis, (4) fermentation, and (5) ethanol recovery (Fig. 1). In the first step of the process, the delivered biomass must be made uniform in size to facilitate handling and transport via conveyor or screw drive and to provide a more consistent surface-to-mass ratio for thermo­chemical pretreatment. The pretreatment step is typically a short — (minutes) to long-term (hours) exposure to extremes of temperature (150-200 C), pH (<2 or >10) and pressure (2-5 atm) and may additionally involve a rapid pressure release that facilitates chemical infiltration and fiber explosion. Ide­ally, pretreatment produces a disrupted, hydrated substrate that is accessible to enzymatic attack, but avoids both the production of sugar degradation products and fermentation inhibitors. As discussed below, some pretreat­ments solubilize hemicellulose to oligomeric and/or monomeric sugars com­prised largely of pentoses that can be fermented independently or together with the glucose released from the cellulose fraction. In the next step, the pH is adjusted and enzymes are added to initiate cellulose hydrolysis to fer­mentable sugars. With pretreatments that do not solubilize the hemicellulose fraction, additional enzymes may be required to hydrolyze the hemicellulose

polymer. Hydrolysis typically is performed at pH 5 and 50 °C for 24-120 h, followed by addition of a fermentation organism to begin production of ethanol. In many cases (as described below) fermentation is initiated long be­fore hydrolysis has completed, since both the extent and speed of ethanol pro­duction can often be increased by combining the hydrolysis and fermentation steps. In the final step, ethanol is recovered via distillation, and remaining organic waste is burned for production of heat and/or power.

2.1

The first attempt to introduce an L-arabinose utilization pathway in S. cere­visiae by heterologous expression of the complete E. coli L-arabinose pathway
did not result in appreciable arabinose utilization [70], most likely due to the absence of functional expression of the L-arabinose isomerase. It was only when the E. coli araA gene encoding the L-arabinose isomerase was substi­tuted by the corresponding Bacillus subtilis gene that a functional arabinose pathway was established in S. cerevisiae [71]. Similar to the use of the het­erologous XI pathway, other genetic modifications in addition to the new L-arabinose isomerase were required for the recombinant strain to grow on L-arabinose as sole carbon source [71]: an additional copy of the galactose permease (Gal2), which also transports arabinose [72], and an unspecified adaptation for growth on arabinose [71].

The fungal L-arabinose utilization pathway has also been introduced in S. cerevisiae, combining enzymes from P. stipitis and from the filamentous fungus Trichoderma reesei. The enzymes were actively expressed; however, neither appreciable growth on L-arabinose nor significant ethanolic fermen­tation was observed [73]. The dysfunction of the fungal arabinose pathway with respect to ethanolic fermentation parallels the inability of the naturally arabinose-growing yeasts to ferment L-arabinose to ethanol [50,69]. Instead, these yeasts often produce L-arabitol from L-arabinose (Fig. 2) [65,66,69]. Minute ethanolic fermentation has been observed for six yeast species, C. arabinofermentans, P. guilliermondii, C. auringiensis, C. succiphila, Ambro- siozyma monospora, and Candida sp. YB-2248, but only in rich medium [65, 69]. Rich media may contain other fermentable sugars as well as undefined electron acceptors that serve to regenerate reduced cofactors [32,74-76], which appears necessary for ethanolic arabinose fermentation to occur via the fungal pathway. Also, the presence of low amounts of oxygen aids cofactor regeneration [50,77].

4

Functional expression in S. cerevisiae of a highly active fungal XI has paved the way for metabolic engineering of this yeast towards high-yield, rapid production of ethanol from D-xylose under fully anaerobic conditions. On theoretical grounds, this XI-based approach is superior to the extensively studied xylose reductase/xylitol dehydrogenase strategy. While considerable experimental proof to substantiate this statement has been obtained under “academic” conditions, a next important challenge is to do the same under industrial conditions. While the first experiments in real-life plant biomass hydrolysates are quite promising, there remains plenty of scope for integrat­ing the D-xylose-fermentation genotype with other metabolic and process­engineering strategies for further increased robustness under process condi­tions.

In addition to D-xylose, plant biomass hydrolysates contain several other potentially fermentable substrates that cannot be converted by wild-type S. cerevisiae strains [69]. While these compounds often represent only a few percent of the potentially fermentable carbon, they can have a decisive impact on economical competitiveness and sustainability of high-yield, high-volume processes such as fuel ethanol production. Functional integration of a highly efficient D-xylose fermentation pathway with pathways that are under devel­opment (e. g. arabinose [9,36]) or under consideration (e. g. rhamnose [69]) therefore presents an additional challenge in metabolic engineering for ef­ficient fermentation of plant biomass hydrolysates. We are convinced that creative integration of metabolic engineering, evolutionary engineering and process design can result in rapid breakthroughs in these areas.

The regular and cross-linked cellulose chains form a very efficient barrier against penetration of the enzymes into the fibres. Swelling of the pores can be achieved by alkaline pretreatment through soaking of the material in an alkaline solution, such as NaOH, and then heating it for a certain time. The swelling causes an increase in the internal surface area, and a decrease in the degree of polymerization and crystallinity. Usually a major fraction of the lignin is solubilized together with some of the hemicellulose. A rather large fraction of the hemicellulose sugars are usually recovered as oligomers. Al­kaline pretreatment breaks the bonds between lignin and carbohydrates and disrupts the lignin structure, which makes the carbohydrates more accessible to enzymatic attack. As it acts mainly by delignification, it is more effective on agricultural residues and herbaceous crops than on wood materials, as these materials in general contain less lignin. For softwood species, which contain a large amount of lignin, a small or no effect has been observed. Pretreatment using lime instead of sodium hydroxide is another alkaline method, especially suited for agricultural residues, e. g. corn stover, or hardwood materials, such as poplar [24,25].

Dilute acid pretreatment is performed by soaking the material in dilute acid solution and then heating to temperatures between 140 and 200 °C for a certain time (from several minutes up to an hour). Sulphuric acid, at con­centrations usually below 4 wt %, has been of most interest in such studies as it is inexpensive and effective. The hemicellulose is hydrolysed and the main part is usually obtained as monomer sugars. It has been shown that materials that have been subjected to acid hydrolysis may be harder to ferment because of the presence of toxic substances [26-28].

Another approach is to use an organic or aqueous-organic solvent mixture with addition of an inorganic acid catalyst (H2SO4 or HCl), which is used to break the internal lignin and hemicellulose bonds. These methods are usu­ally referred to as organosolv processes [29]. In these cases the hydrolysed lignin is dissolved and recovered in the organophilic phase. It is important to thoroughly wash the material prior to enzymatic hydrolysis and fermenta­tion, as the solvents may act as inhibitors. Solvents that are used are typically methanol, ethanol, acetone, ethylene glycol, triethylene glycol and phenol. Some of these substances are explosive and highly inflammable and thus dif­ficult to handle.

3.3

Development of improved enzymes for the hydrolysis of the other major carbohydrate polymer present in lignocellulosic biomass is also of commer­cial interest, particularly to those utilizing neutral or alkaline pretreatments that leave much of the hemicellulose intact. To develop these enzymes, an industrial residue of the wheat starch industry was used as a model sub­strate. In Europe, wheat is one of the major substrates for production of fuel ethanol. Processing of wheat starch for glucose results in a by-product stream (vinasse) consisting mainly of the wheat endosperm cell wall ma­terial and leftover yeast cells following the fermentation of the starch. The hemicellulose by-product is approximately 33 wt % carbohydrates of which approximately 66 wt % is arabinoxylan. Arabinoxylans consist of a linear backbone of P-1,4-linked D-xylopyranosyl units that are partially substituted with arabinofuranosyls. The major portion of the arabinoxylan in indus­trial wheat fermentation residues is water-soluble [39], the water-insoluble arabinoxylan is quantitatively more abundant in cell walls isolated directly from unprocessed wheat endosperm [40]. Arabinoxylans are hydrolyzed to monosaccharides by acid treatment or by enzymatic hydrolysis. The enzy­matic hydrolysis is usually preferred because it allows for a more specific and controlled modification and fewer undesirable by-products, making it more suitable for microbial fermentation using organisms that can metabolize both xylose and arabinose [41].

The enzymatic degradation of arabinoxylans requires both side-group cleaving and depolymerizing enzymes. Cleavage of the side chains re­quires the action of several accessory enzyme activities, including a-L — arabinofuranosidases, a-glucuronidases, ferulic acid esterase, and acetyl — esterases [41,42]. Depolymerization requires endo-1,4-|3-xylanases that result in unbranched xylooligosaccharides, including xylotriose and xylobiose, and в-xylosidases that cleave xylobiose and attack the non-reducing ends of short chain xylooligosaccharides to liberate xylose [41].

The hydrolysis of arabinoxylan is critical for improved utilization of wheat hemicellulose in the ethanol industry. Three Novozymes cellulolytic and hemicellulolytic enzyme preparations, Celluclast 1.5 L, Ultraflo L, and Vis — cozyme L were tested in various combinations for their ability to liberate arabinose and xylose from water-soluble wheat arabinoxylan. The substrate was medium viscosity water-soluble wheat arabinoxylan from Megazyme (Bray). The three different enzymes were evaluated individually and also in 50 : 50 combinations to look for possible synergistic effects. Reactions were carried out at pH 5 and 50 °C followed by analysis of arabinose, galactose, glu­cose, xylose, xylobiose, and xylotriose by high-performance anion exchange chromatography (HPAEC) [43]. The molecular weight and distribution of water-soluble wheat arabinoxylan and hydrolyzates were determined by high — performance size exclusion chromatography (HPSEC).

In those reactions containing the individual enzyme preparations, the lev­els of arabinose and xylose increased with increasing enzyme dosage and time. Ultraflo L was superior to Celluclast 1.5 L and Viscozyme L in releasing the arabinose from the water-soluble wheat arabinoxylan, meaning that Ul — traflo L must contain a significant amount of a-L-arabinofuranosidase. Cellu­clast 1.5 L was the best enzyme preparation for liberating xylose, resulting in 26 wt % of the available xylose. Ultraflo L released 16 wt % while Viscozyme L released less than 1.5 wt %. In a mixture of 50 : 50 Celluclast 1.5 L and Ultra — flo L there was no interaction among the arabinose-releasing side activities since the same amount of arabinose was obtained as when the two individual enzyme preparations were used and then the arabinose total was combined. The Viscozyme L preparation exhibited a weak antagonistic effect with Ul — traflo L and Celluclast 1.5 L since the amount of arabinose actually decreased compared to that observed with the individual enzyme preparations. The re­sults indicated that the arabinose-releasing side activities of Viscozyme L had the same activity as those demonstrated by Ultraflo L and Celluclast 1.5 L. Another possibile but less likely explanation is the Viscozyme L contained a — L-arabinofuranosidase inhibitors [43]. The 50 : 50 mixture of Celluclast 1.5 L and Ultraflo L produced an increase in the release of xylose compared with the sum of the individual enzyme preparations (Fig. 8). The mixture released 59 wt % of the available xylose, which was 32 wt % more than the theoret­ical addition of the individual enzyme preparations alone. Combination of Ultraflo L and Viscozyme L showed no such synergism, but incubation of Cel­luclast 1.5 L and Viscozyme L showed a weak synergistic effect in liberating some of the xylose from the wheat arabinoxylan.

To further examine the synergistic affect between Celluclast 1.5 L and Ul — traflo L the amounts of xylobiose and xylotriose released during enzymatic hydrolysis were quantified using HPAEC for both individual and combined enzyme preparations. During the initial stage of incubation, Celluclast 1.5 L

liberated small amounts of both xylobiose and xylotriose, indicating the pres­ence of endo-1,4-^-xylanase activities. As hydrolysis continued, the released xylobiose and xylotriose was hydrolyzed to xylose, indicating the Cellu — clast 1.5 L contained one or more в-xylosidase activities.

Ultraflo L treatment resulted in continual liberation of both xylobiose and xylotriose. Ultraflo L showed a low release of free xylose indicating one or more endo-1,4-^-xylanase activities, but little в-xylosidase activity. The synergistic effect between Celluclast 1.5 L and Ultraflo L in releasing xylose is therefore likely to be a result of the action of a-L-arabinofuranosidase and endo-1,4-^-xylanase activities present in Ultraflo L and the в-xylosidase present in Celluclast 1.5 L [43].

Since a strong synergistic effect was observed with a 50 : 50 combination of Celluclast 1.5 L and Ultraflo L for the breakdown of arabinoxylan, a sec­ond study was conducted to look for similar effects and viscosity reduction in the fermentation residue, vinasse. The effects of enzyme dosage, optimal temperature, and pH were examined in hydrolysis of whole vinasse, vinasse supernatant, and washed vinasse sediment that was provided by Tate & Lyle, Amylum UK (Greenwich, UK). On whole vinasse, the enzyme-catalyzed re­lease of arabinose and xylose by the 50 : 50 combination of Ultraflo L and Cel­luclast 1.5 L decreased as the substrate concentration of the vinasse increased. The monosaccharide release also decreased when the substrate concentration of the vinasse increased. Release of arabinose and xylose from the vinasse sediment was very low. The release of arabinose from the whole vinasse var­ied from 40- 50 g arabinose per kilogram vinasse DM while xylose release was between 75-100 g xylose per kilogram vinasse DM after a 24 h hydrolysis. The

Ultraflo L:Celluclast 1.5 L mixture released 53-75 g arabinose and 75-115 gof xylose per kilogram of vinasse DM after a 24 h hydrolysis [44].

Significant viscosity reduction was obtained by enzyme-catalyzed degra­dation of arabinoxylans present in the fermentation residue stream, vinasse. However, there was limited hydrolysis of the insoluble arabinoxylans in the vinasse sediment. The efficiency of enzymatic degradation of the arabinoxylan in vinasse was dependent on enzyme dosing and substrate dry matter [44].

In an effort to narrow down the specific activities involved in the previous studies, the в-xylosidase from Celluclast 1.5 L was purified and used as a sup­plement to Ultraflo L enzyme preparation. When dosed at 0.25 g в-xylosidase protein per kilogram of arabinoxylan along with Ultraflo L, this enzyme mix released the same or more xylose as the enzyme mix consisting of 50 : 50 Ultraflo L and Celluclast 1.5 L (Fig. 9).

In order to determine the optimal enzyme mix for the hydrolysis of vinasse arabinoxylan, several recombinant enzymes were made and tested in various combinations. Genes were cloned and expressed in the fungal host A. oryzae. Based on our studies the optimal enzyme mix for vinasse hydrolysis consists of a-L-arabinofuranosidase from Meripilus giganteus, a-L — arabinofuranosidase II from Humicola insolens, and T. reesei в-xylosidase. A mixture of 25 : 25 : 50 of a-L-arabinofuranosidase from M. giganteus, a-L- arabinofuranosidase from H. insolens and в-xylosidase from T. reesei was determined to be optimal for maximizing arabinoxylan hydrolysis. The success of this work in identifying and exploiting synergism between hemicellulase component activities is currently being applied to other relevant lignocellulosic substrates that differ significantly in their hemicellulose composition.

5

In addition to the transport flux and the flux through the initial pentose­converting enzymes, the “pulling” effect [55] of the flux through enzymatic reactions downstream of xylitol, as well as through glycolysis, appears to be equally important for ethanolic pentose fermentation. It was early recognized that the presence of glucose during xylose fermentation enhanced the gly­colytic activity [122-124]. Furthermore, it was recently shown that no xylitol was formed in the glucose-xylose coconsumption phase during xylose fer­mentation with recombinant S. cerevisiae in mineral medium [54], nor in lignocellulose hydrolysates which contain hexose sugars [6,12,14].

4.7

Other Modifications

Transcription factors involved in glucose repression have also been modi­fied in order to affect ethanolic xylose fermentation. The gene MIG1, or both MIG1 and MIG2, were deleted in an XR-XDH-XK-carrying strain of S. cere­visiae [125] to generate strains which were constantly glucose de-repressed during glucose-xylose cofermentation. This engineering strategy had little ef­fect on ethanol formation. It rather led to increased xylitol formation [125] (strains CPB. CR2 and CPB. MBH2, Table 3). Similarly, when truncated ver­sions of the MIG1 gene were expressed in xylose-utilizing strains of S. cere — visiae, growth and ethanol formation were only marginally affected [126]. The bacterial phosphoketolase pathway, which converts xylulose-5-phosphate directly to glyceraldehyde-3-phosphate and acetyl-P, has also been introduced in S. cerevisiae to enhance ethanolic xylose fermentation [127,128]. The xyl­itol yield decreased without any increase in the ethanol yield [128] (strain TMB3001c-p6XFP/p4PTA/p5EHADH2, Table 2). In contrast, heterologous ex­pression of a bacterial hemoglobin gene to render the cells a more oxidized state in oxygen-limited conditions was successful [129]. Improved ethanolic xylose fermentation was observed. This strategy is, however, only applicable in oxygenated cultures [129].

4.8

The utility of KO11 for production of ethanol from biomass has been demon­strated with multiple substrates including, but not limited to, rice hulls [19], sugar cane bagasse [20], agricultural residues [20], Pinus sp. hydrolysate [21], corn cobs, hulls and AFEX-pretreated fibers [22,23], orange peel [12], wil­low [24], pectin-rich beet pulp [25], sweet whey [26], brewery waste [27], and cotton gin waste [28]. The final ethanol titers and fermentation times for these substrates are presented in Table 1. Consistent with the robustness of the parental E. coli W, KO11 is relatively robust to changes in temperature and pH [29]. KO11 has also been the subject of an empirical kinetic model [24].

While similar ethanol yields are obtained from glucose and xylose, differ­ences in transport mechanisms result in a lower ATP yield for xylose. Both KO11 and LY01 grow approximately 50% faster and produce three times as much ATP from glucose relative to xylose [30]. As expected, the expression

of xylose metabolic genes is increased during xylose growth relative to glu­cose growth. However, genes contributing to metabolism of other pentose sugars, such as arabinose, ribose and lyxose, also have increased expression during xylose growth, consistent with a relaxation of the cAMP-CRP control system [30].

2.1.3

Lignin is an aromatic network polymer composed of phenylpropane units [53]. It is generally accepted that lignin is the “glue” that binds cellulose and hemicellulose, imparting both rigidity and moisture resistance to the lig — nocellulosic structure. Lignin has also been implicated as an inhibitor of cellulases; therefore, many of the pretreatment methods currently being ex­plored have tried to decrease the lignin content of the solid substrate while minimizing the degradation of carbohydrates [22]. The amount of lignin that must be dealt with by a particular pretreatment and subsequent hydro­lysis depends on the source of biomass. For example, corn fiber has a low lignin content of 7% (w/w) [23] compared to 30% (w/w) in the case of a soft­wood such as Douglas-fir [51]. In addition to the amount of lignin present in a biomass feedstock, the type of lignin differs between agricultural residues, hardwoods and softwoods [54]. Grasses and agricultural residues contain primarily p-hyroxyphenyl units while hardwoods and softwoods contain greater amounts of syringyl and guaiacyl subunits, respectively [54]. Soft­woods lignin also exhibits a greater degree of cross-linking due to an extra linking site provided by the presence of only a single methoxyl group on the guaiacyl aromatic ring [54]. Another factor that must be considered is the ex­istence of lignin carbohydrate complexes (LCCs) that consist of lignin linked to carbohydrates through bonds such as ester, ether or ketal [55]. LCCs have been shown to be particularly problematic for hydrolysis processes, as access to the carbohydrate fraction is restricted by the attached lignin, therefore pre­treatment processes should either expand the pore structure of the substrate or remove the lignin outright [56].

3.1

The activities of commercial reference preparations were first measured at higher temperatures in order to evaluate their general performance and to es­timate the role of the background activities originating from the production strain. The hydrolysis of the pretreated spruce substrate by the commercial preparations (with and without added в-glucosidase, BG) at various tempera­tures from 50 to 70 °C was estimated during the first 24 h of the hydrolysis. The native Trichoderma cellulases and the Aspergillus BG were rapidly inac­tivated during the first 2 h of hydrolysis of the pretreated spruce substrate at temperatures above 60 °C (Fig. 2). The hydrolysis ceased after 24 h at 60 °C and after 48 h at 55 °C (results not shown). As expected, the effect of the added BG on the sugar yield was significant. The relative inactivation of BG was more pronounced even at 60 °C (Fig. 2b). The hydrolytic effect of the rather high loading (about 20 FPU g-1 cellulose) of T. reesei and Aspergillus enzymes was obviously due to the initial stage of hydrolysis during which the enzymes remained active. The hydrolysis yield of sugars from spruce dur­ing the first 2 h was 15% of the theoretical maximum at 70 °C, 22% at 65 °C and 33% at 60 °C. There were indications that the temperature optimum of the commercial T. reesei enzymes in the hydrolysis of the pretreated spruce substrate was about 5 °C lower than on pure cellulose (results not shown).

Fig. 2 Hydrolysis of washed, steam pretreated spruce substrate (cellulose content 18.3 gL-1) with Celluclast 1.5 L FG alone (A) or supplemented with Novozym 188 (B) at various temperatures at pH 5. The dosage of Celluclast was 22 FPU g-1 cellulose and the Novozym 188 P-glucosidase 550 nkatg-1 cellulose. И50 °C, Ш55 °C, ♦бО °C, <>65 °C and • 70 °C

8