In view of the intrinsic redox restrictions associated with the combined in­troduction of xylose reductase and xylitol dehydrogenase into S. cerevisiae, it is relevant to explore alternative metabolic engineering strategies. As will be discussed below, expression of heterologous genes for xylose isomerase (an enzyme that does not naturally occur in S. cerevisiae) offers such an alterna­tive [14]. In the following sections, we will briefly discuss the properties and taxonomic distribution of xylose isomerases. This will be followed by a brief overview of previous attempts at functional expression of xylose isomerases in S. cerevisiae. We will then discuss how, in the past few years, fast progress has been made due to the discovery of a new, fungal xylose isomerase gene. Finally, we will discuss the status of the xylose isomerase strategy with regard to full-scale industrial application.

2

Following the release and annotation of a genome, the next logical step is to evaluate the messenger RNA expression level on a whole genome scale, referred to as transcriptome analysis. Targeted metabolic engineering relies heavily on the assumption that a genetic perturbation — gene deletion, con­stitutive overexpression, regulated induction, or modulation — will confer a metabolic flux response. This stems from the central dogma of biology: DNA is transcribed to RNA and subsequently translated to polypeptides that give rise to phenotype. Prior to transcriptome analysis, genes were assumed to be expressed followed by post-translational regulation, with little under­standing of interactions across gene loci [81]. In fact, transcriptome profiling of reference strains has provided a first approximation as to which pathways are active and, equally important, inactive, assuming that up-regulated gene expression leads to up-regulated pathway activity. It has since been shown that this is not always true — elevated mRNA levels do not always translate to elevated protein levels or activity. It has also provided significant insight into alternative modes of regulation, such as transcription factor-mediated as opposed to post-translational regulation. This has permitted narrowing of the experimental space that metabolic engineers need to consider, and made available new strategies to consider. Additionally, transcriptome pro­filing provides a quantitative in vivo assessment of several key metrics fol­lowing a genetic perturbation relative to a reference case: (1) what is the net change in mRNA expression levels of the targeted gene(s), (2) what is the net change in mRNA expression levels of non-targeted gene(s), and (3) what is the net change in mRNA expression levels of either reference or constructed strains under specific environmental conditions. These questions aim to iso­late which genes and pathways may serve as targets and/or explanations for observed or induced phenotypes. Measurement of the transcriptome, via readily available microarray technology, has evolved into a routinely meas­ured data set for many industrially relevant organisms, including E. coli and S. cerevisiae, and is playing a central role in both forward and reverse metabolic engineering [63,82,83].

Among the first applications of transcriptome measurements with in­dustrial relevance to bioethanol production was establishing the baseline response of S. cerevisiae to diverse carbon substrates and medium com­positions — essential for optimizing strains to given feedstocks and vice versa. Steady-state chemostat cultures were used to measure transcriptome responses under glucose, ethanol, ammonium, phosphate, and sulfate lim­itations [84]. Results suggested that genes related to high-affinity glucose uptake, the TCA cycle, and oxidative phosphorylation were up-regulated in glucose-limiting conditions, while genes involved in gluconeogenesis and ni­trogen catabolite repression were up-regulated in ethanol-grown cells [84]. In a similar but earlier study, transcriptome measurements were performed of S. cerevisiae grown using glucose-limited chemostats coupled with nitro­gen, phosphorus, and sulfur limitations [85]. In total, 1881 transcripts (31% of the total 6084 different open reading frames probed) were significantly up — or down-regulated between at least two conditions, and a total of 51 genes demonstrated a >tenfold higher or lower expression within a given condi­tion [85]. The transcriptome profiles under each condition have provided genetic motifs that may be recognized and regulated by transcription factors. These may be used in metabolic engineering strategies that could cater to a specific growth medium composition.

With the experimental mechanics of collecting transcriptome data becom­ing common place, attention and focus is now placed on data analysis methods and integration with other x-ome data sets. It has become abundantly clear that transcriptome data alone, unless used for the purposes of environmen­tal screening or quality control (i. e., confirming that an engineered genotype is producing the corresponding transcription profile), provides limited bi­ological insight. Several efforts have emerged coupling transcriptome with metabolome and fluxome data [86-89]. For example, elementary flux modes for three carbon substrates (glucose, ethanol, and galactose) were deter­mined using the catabolic reactions from the genome-scale metabolic model of S. cerevisiae, and then used for gene deletion phenotype analysis. Control — effective fluxes were used to predict transcript ratios of metabolic genes for growth under each substrate, resulting in a high correlation between the theor­etical and experimental expression levels of 38 genes when ethanol and glucose media were considered [90]. This example demonstrates that incorporating transcriptional functionality and regulation into metabolic networks for in silico predictions provides both more biologically representative models and a means of bridging transcriptome and fluxome data.

In another example, the topology of the genome-scale metabolic model constructed for S. cerevisiae is examined by correlating transcriptional data with metabolism. Specifically, an algorithm was developed enabling the iden­tification of metabolites around which the most significant transcriptional changes occur (referred to as reporter metabolites) [91]. Due to the highly connected and integrated nature of metabolism, genetic or environmental per­turbations introduced at a given genetic locus will affect specific metabolites and then propagate throughout the metabolic network. Using transcriptome experimental data, a priori predictions of which metabolites are likely to be affected can be made, and serve as rational targets for additional inspection and metabolic engineering [91]. This algorithm has been recently extended to include reporter reactions, whereby transcriptional data is correlated with the metabolic reactions of the reconstructed S. cerevisiae genome-scale metabolic network model to identify those reactions around which a genetic or environ­mental perturbation conferring transcriptional changes cluster [92].

As more genomes continue to become available, and microarray technol­ogy continues to become more accessible with cost-effective customizable DNA microarrays now available, transcriptome data will continue to increase. Bioinformatics for data handling, integration of transcriptome with other x-ome data, and the development of various network models that rely on tran- scriptome data for biological interpretation will continue to develop. From an industrial biotechnology perspective transcriptome measurements and analysis have played a large role in reverse metabolic engineering; transcrip­tional surveying of a strain constructed either via random mutagenesis or directed evolution [63,82,83,93]. For example, lysine production via C. glu — tamicum has undergone transcriptome and fluxome measurements to elu­cidate greater than 50 years of traditional metabolic engineering (random mutagenesis), providing new targets for improved strategies [94-96]. This ef­fort, applied to other industrial biotechnology processes, is likely to intensify.

3.3

The key to developing an economically viable biorefinery is to employ a holis­tic approach that integrates the unit steps, maximizing the yield at each, while minimizing both capital and operating costs. At each step of the process, from pretreatment to fermentation, effort must be made to minimize any loss in potential ethanol production. In the example in Fig. 2, the production of degraded sugars during pretreatment, incomplete cellulose or hemicellulose

conversion to fermentable sugars during hydrolysis, and fermentation losses due to sugar consumption by the yeast all contribute to lost value in the con­version. If biomass feedstock such as corn stover, purchased at $5/ton, could be converted with perfect efficiency to its theoretical potential of 113 gal­lons of ethanol per ton of stover with an ethanol selling price of $2/gallon, the value of the ethanol would be ~$225/ton, creating an “operating cost window” for depreciation of capital, operation, and profit of ~$220/ton [5]. Losses in any unit step that reduces the overall yield will reduce the value per ton, whether the losses result from reduced enzyme hydrolysis, poor fer — mentability of the hydrolyzate sugars, or reduced fermentation yield. It is also important to note that maximizing the conversion of the two most abundant sugars, glucose and xylose, is important to viable economics. If only cellulose is utilized with no conversion of hemicellulose, the theoretical yield drops 39% to 69 gallons/ton, reducing the cost window to ~ $135/ton. Unless the xylose is utilized to produce something of equal or higher value, it is un­likely that such a process could be viable. Similarly, a pretreatment selected on the basis of a reduced capital cost for installed equipment, but increasing the required enzyme dosage, may reduce the operating cost window significantly.

3

It soon became evident that the mere introduction ofpentose utilization path­ways in S. cerevisiae was not enough to render the recombinant strains traits for efficient ethanol fermentation [43] (strain RWB202, Tables 3 and 4), [47, 48,78,79] (strain TMB3001, Tables 1-3). A number of metabolic engin­eering strategies to enhance ethanolic xylose (and arabinose) fermentation in S. cerevisiae have been explored, the most important of which will be discussed below. The initial xylose utilization pathway, the cellular redox metabolism, and the flux of central carbon metabolism have been the main targets of these engineering strategies. Figure 3 highlights the metabolic re­actions that have been engineered to improve ethanolic xylose fermentation by S. cerevisiae.

4.1

1 Introduction……………………………………………………………………………………………… 206

2 Baker’s Yeast (S. cerevisiae) as a CBP Host…………………………………………………….. 208

3 Engineering S. cerevisiae for Sugar Fermentation……………………………………………… 210

4 Expression of Cellulases in S. cerevisiae…………………………………………………………. 211

5 Expression of Hemicellulases in S. cerevisiae…………………………………………………… 218

6 Selection for the Development of Superior CBP Yeasts………………………………….. 224

7 Integration of Different Enzymatic Activities

into a Single CBP Yeast and Transfer to Industrial Strains………………………. 228

References…………………………………………………………………………………………………….. 230

Abstract Consolidated bioprocessing (CBP) of lignocellulose to bioethanol refers to the combining of the four biological events required for this conversion process (production of saccharolytic enzymes, hydrolysis of the polysaccharides present in pretreated biomass, fermentation of hexose sugars, and fermentation of pentose sugars) in one reactor. CBP is gaining increasing recognition as a potential breakthrough for low-cost biomass pro­cessing. Although no natural microorganism exhibits all the features desired for CBP, a number of microorganisms, both bacteria and fungi, possess some of the desirable properties. This review focuses on progress made toward the development of baker’s yeast (Saccharomyces cerevisiae) for CBP. The current status of saccharolytic enzyme (cellulases and hemicellulases) expression in S. cerevisiae to complement its natural fermentative ability is highlighted. Attention is also devoted to the challenges ahead to integrate all required enzymatic activities in an industrial S. cerevisiae strain(s) and the need for molecular and selection strategies pursuant to developing a yeast capable of CBP.

Keywords Consolidated bioprocessing • Cellulolytic yeast •

One-step bioethanol production • Saccharomyces cerevisiae

1

Introduction

Biomass is the only foreseeable renewable feedstock for sustainable produc­tion of biofuels. The main technological impediment to more widespread utilization of this resource is the lack of low-cost technologies to overcome the recalcitrance of the cellulosic structure [1]. Four biological events occur during conversion of lignocellulose to ethanol via processes featuring enzy­matic hydrolysis: production of saccharolytic enzyme (cellulases and hemi — cellulases), hydrolysis of the polysaccharides present in pretreated biomass, fermentation of hexose sugars, and fermentation of pentose sugars [2]. The hydrolysis and fermentation steps have been combined in simultaneous sac­charification and fermentation (SSF) of hexoses and simultaneous saccharifi­cation and cofermentation (SSCF) of both hexoses and pentoses schemes. The ultimate objective would be a one-step “consolidated” bioprocessing (CBP) of lignocellulose to bioethanol, where all four of these steps occur in one re­actor and are mediated by a single microorganism or microbial consortium able to ferment pretreated biomass without added saccharolytic enzymes (Fig. 1).

CBP is gaining increasing recognition as a potential breakthrough for low — cost biomass processing. A fourfold reduction in the cost of biological pro­cessing and a twofold reduction in the cost of processing overall is projected when a mature CBP process is substituted for an advanced SSCF process fea­turing cellulase costing US $0.10 per gallon ethanol [3]. The US Department of Energy (DOE) Biomass Program multiyear technical plan states: “Mak­ing the leap from technology that can compete in niche or marginal markets for fuels and products also requires expanding the array of possible concepts and strategies for processing biomass. Concepts such as consolidated bio­processing… offer new possibilities for leapfrog improvements in yield and cost.” [4]. The detailed analysis of mature biomass conversion processes by Greene et al. [5] finds CBP to be responsible for the largest cost reduction of all R&D-driven improvements incorporated into mature technology scenar­ios featuring projected ethanol selling prices of less than US $0.70 per gallon. Finally, a recent report entitled Breaking the Biological Barriers to Cellulosic Ethanol states: “CBP is widely considered to be the ultimate low-cost configu­ration for cellulose hydrolysis and fermentation.” [6].

In addition to being desirable, recent studies of naturally occurring cel­lulolytic microorganisms provide increasing indications that CBP is feas­ible. Lu et al. [7] showed that cellulase-specific cellulose hydrolysis rates exhibited by growing cultures of Clostridium thermocellum exceed specific rates exhibited by the Trichoderma reesei cellulase system by approximately 20-fold, with a substantial part of this difference resulting from “enzyme — microbe synergy” involving enhanced effectiveness of cellulases acting as part of cellulose-enzyme-microbe complexes. Whereas cellulase synthesis

was thought to be a substantial metabolic burden for anaerobic microbes fermenting cellulose without added saccharolytic enzymes, C. thermocellum realizes cellulose-specific bioenergetic benefits that exceed the bioenergetic cost of cellulase synthesis [8]. These and other observations provide guid­ance with respect to features that may be beneficial in the course of creating recombinant cellulolytic microbes, and also underscore the point that micro­bial cellulose utilization is differentiable from enzymatic hydrolysis from both fundamental and applied perspectives [1,3].

Although no natural microorganism exhibits all the features desired for CBP, a number of microorganisms, both bacteria and fungi, possess some of the desirable properties. These microorganisms can broadly be divided into two groups: (1) native cellulolytic microorganisms that possess supe­rior saccharolytic capabilities, but not necessarily product formation, and

(2) recombinant cellulolytic microorganisms that naturally give high prod­uct yields, but into which saccharolytic systems need to be engineered [1, 9]. Examples of native cellulolytic microorganisms under consideration in­clude anaerobic bacteria with highly efficient complexed saccharolytic sys­tems, such as mesophilic and thermophilic Clostridium species [9,10], and fungi that naturally produce a large repertoire of saccharolytic enzymes, such as Fusarium oxysporum [11] and a Trichoderma species [12]. However, the anaerobic bacteria produce a variety of fermentation products, limiting the ethanol yield, whereas the filamentous fungi are slow cellulose degraders and give low yields of ethanol [13]. Candidates considered as potential re­combinant cellulolytic microorganisms into which saccharolytic systems have been engineered include the bacteria Zymomonas mobilis [14,15], Escherichia coli [16,17] and Klebsiella oxytoca [18,19], and the yeast Saccharomyces cere — visiae and xylose-fermenting yeasts Pachysolen tannophilus [20], Pichia stipi — tis, and Candida shehatae [21].

While both native and recombinant cellulolytic microorganisms merit in­vestigation, this review will focus on the well-known ethanol producing yeast S. cerevisiae, which has a long commercial history as microorganism of choice for beer, wine, baker’s yeast, and commercial ethanol production. In particu­lar, we address recent progress in heterologous cellulase expression pursuant to development of recombinant cellulose-fermenting yeast strains [22-25].

2

This category includes methods in between, or a mixture of, purely physical and chemical methods. Steam pretreatment is one of the most widely used methods for pretreatment of lignocellulosics. This pretreatment method used to be called steam explosion, since it was believed that an “explosive” action on the fibres was necessary to render a material suitable for hydrolysis. It has been shown that it is more likely that the effect of steam pretreatment is due to acid hydrolysis of the hemicellulose, which is the reason why some cellulosic materials are easier than others to break down [30,31]. In particu­lar, agricultural residues and some types of hardwood contain organic acids, which act as catalysts for the hemicellulose hydrolysis. Using steam pretreat­ment the raw material is usually treated with high-pressure saturated steam at a temperature typically between 160 and 240 °C (corresponding to a pres­sure between 6 and 34 bar), which is maintained for several seconds to a few minutes, after which the pressure is released. During pretreatment some of the raw material, predominantly hemicellulose, is solubilized and found in the liquid phase as oligomeric and monomeric sugars. The cellulose in the solid phase then becomes more accessible to the enzymes. It is in some cases dif­ficult to find conditions that result in high yields of both hexose and pentose sugars, and at the same time also create a cellulose fraction which is easy to hydrolyse to glucose. This may call for steam pretreatment using two steps, where hemicellulose sugars are recovered at lower severity, while the cellulose fraction is subjected to pretreatment at higher severity.

Steam pretreatment can be improved by using an acid catalyst, such as H2SO4 or SO2. The acid increases the recovery of hemicellulosic sugars, and also improves the enzymatic hydrolysis of the solid residue. The use of an acid catalyst in steam pretreatment results in an action similar to dilute acid hydro­lysis but with less liquid involved. It is especially important to use an acid catalyst for softwood, since softwood in general is more difficult to degrade.

Steam pretreatment with addition of a catalyst is the pretreatment method for hydrolysis and enzymatic digestibility improvement that is closest to commercialization. It has been widely tested in pilot-scale equipment, for example, in the NREL pilot plant in Golden, CO (USA) [32] and in the SEKAB pilot plant in Ornskoldsvik (Sweden) [33], and is also used in a demonstration-scale ethanol plant at Iogen in Ottawa (Canada) [34].

Hydrothermolysis, or liquid hot-water (LHW) treatment, involves treat­ment in water at high temperature. This method is similar to steam pre­treatment, but lower temperatures and lower dry matter (DM) content are used, and thus more poly — and oligosaccharides are recovered [35,36]. A cat­alyst, such as an acid, can be added, making the method similar to dilute acid pretreatment. Since the water content is much higher than in steam pre­treatment, the resulting sugar solution is more diluted and thus causes the downstream processes to be more energy demanding. In the range 1-10 wt % DM virtually no difference in ethanol yield was found when bagasse was treated at 220 °C, after which SSF was performed using S. cerevisiae [37].

Wet oxidation pretreatment involves the treatment of the biomass with water and air, or oxygen, at temperatures above 120 °C, sometimes with the addition of an alkali catalyst. This method is suited to materials with low lignin content, since the yield has been shown to decrease with increased lignin content, and since a large fraction of the lignin is oxidized and solubi­lized [38]. As with many other delignification methods, the lignin cannot be used as a solid fuel, which considerably reduces the income from by-products in large-scale production. As discussed in the “Process Economics” chapter, it is extremely important to recover as much as possible of the lignin fraction (Sassner et al., in this volume).

Ammonia fibre explosion (AFEX) is also an alkaline method which, sim­ilarly to the steam pretreatment process, operates at high pressures. The biomass is treated with liquid ammonia for about 10-60 min at moderate temperatures (below 100 °C) and high pressure (above 3 MPa) [39,40]. Up to 2 kg of ammonia is used per kg of dry biomass. The ammonia is recycled after pretreatment by reducing the pressure, as ammonia is very volatile at atmospheric pressure. During pretreatment only a small amount of the solid material is solubilized, i. e. almost no hemicellulose or lignin is removed. The hemicellulose is degraded to oligomer sugars and deacetylated [41], which is a probable reason for the hemicellulose not becoming soluble. However, the structure of the material is changed, resulting in increased water holding cap­acity and higher digestibility. Like the other alkaline pretreatment methods AFEX performs best on agricultural waste, but has not proven to be efficient on wood due to its higher lignin content [42,43]. According to Sun et al. the AFEX process does not produce inhibitors that may affect downstream bio­logical processes [44].

Another type of process utilizing ammonia is the ammonia recycle perco­lation (ARP) method [45,46]. In the process aqueous ammonia (10-15 wt %) passes through biomass at elevated temperatures (150-170 °C), after which the ammonia is recovered. ARP is an efficient delignification method for hardwood and agricultural residues, but is somewhat less effective for soft­wood.

3.4

There is arguably no other industrial enzyme application that poses a greater challenge to the enzyme producer than supplying cost-effective enzymes for biomass utilization. The high enzyme loading required, combined with the low value of the final product, in the form of ethanol, requires not only that the enzymes be as efficient as possible, but that the cost of producing them be as low as possible. To this end, significant effort has been expended over the past 6 years to increase the productivity of the fungal strains used to pro­duce the enzymes, to reduce the cost of the enzyme fermentation process by reducing the cost of carbon and nitrogen sources for the fermentations, and to reduce the complexity of enzyme recovery and formulation.

Improving the host by classical mutagenesis is one way of developing a host strain with improved total protein production and improved activ­ities. This approach has a long and successful history. Montenecourt and Eveleigh [32] isolated RutC30, one of the best existing Trichoderma cellulase mutants, using a combination of ultraviolet irradiation and nitrosomethyl guanidine (NTG). Recently, Toyama, et al. [45] demonstrated a method to screen for increased cellulase production using growth through an overlay of cellulose substrate (Avicel) in Petri plates. In an effort to increase total cellulase productivity, a combination of these methods were utilized on the T. reesei strain currently used to produce Celluclast 1.5 L. Chemical muta­genesis was used to generate mutants that were screened using the method of Toyama [45] with minor changes. Briefly, mutagenized spores were sus­pended in an agar medium, poured into a plate and allowed to harden. The spore-containing layer was then covered with a top layer of agar contain­ing washed, acid pretreated corn stover (PCS) as the sole carbon source. Colonies growing through the PCS layer fastest were isolated and used in a secondary screening. In this, spores from selected fast-growing colonies were inoculated into shake flasks containing cellulase-inducing media. After 5 days of growth, broth samples were tested by robotic assay for produc­tion of reducing sugars from hydrolysis of PCS. Total protein assays were then conducted, and mutants expressing elevated cellulase and/or total pro­tein were then re-grown in 2-L fermentors. Broth from the fermentors was then analyzed again in PCS hydrolysis assays and for total protein. Some mu­tants were identified as having improved PCS hydrolysis and increased total protein secretion compared with the control. Top strains isolated in this man­ner showed significant increases in protein production and secreted cellulase activity.

Another method of improving a cellulase productivity is through increas­ing the expression of target proteins using genetic engineering. In many cases the total cost of supplying a heterologous mix of enzymes can be reduced by creating a single expression host expressing not only the native cellulases and hemicellulases, but expressing additional components, such as the BG and GH61 proteins, without negatively impacting the expression of the na­tive proteins. The introduction of multiple genes into a single host is no easy feat. A significant amount of work was done to identify strong promoters, to identify a number of selectable markers, and to develop a successful trans­formation technique that allows for co-transformation of multiple transgenes. These technological improvements have allowed us to rapidly and efficiently investigate the effect of introducing various enzymes into the T. reesei cellu — lase mix.

In addition to controlling gene expression transcriptionally, by utilizing promoters of different strengths, we have focused on enhancing individual protein yield by optimizing protein secretion. One example is the replacement of the A. oryzae BG signal sequence with a signal peptide from H. insolens Cel45A EG, which improved the BG secretion in T. reesei by two- to threefold relative to the unfused gene (Fig. 10).

As previously mentioned, several GH61 proteins result in a “boost” in PCS hydrolysis when supplemented to Celluclast 1.5 L. In addition, our stud­ies show that increased levels of в-glucosidase are required in our Tricho — derma host. Therefore, numerous co-transformations of T. reesei with various GH61s, A. oryzae в-glucosidase, and other genes of interest were carried out. Those transformants expressing both a GH61 and the в-glucosidase were then screened in PCS hydrolysis assays in order to identify the top strains in true performance assays. Those strains demonstrating the best perform-

ance in PCS hydrolysis were then fermented in 2-L bioreactors and retested in PCS hydrolysis assays. Eventually, a single strain was identified exhibiting im­proved hydrolysis from our original strains and high total protein production (Fig. 11).

5.1

Random methods such as mutagenesis, adaptation, hybridization, and evo­lutionary engineering [130] have been employed to obtain improved xylose­utilizing [5,42,110,131] (strains TMB3400, C1, C5, BH42, RWB218, RWB202- AFX, H2490-4, Tables 1, 3, and 4) and arabinose-utilizing [71] S. cerevisiae strains. Some of the resultant strains have been analyzed in order to identify molecular traits related to the improved ethanolic fermentation of pentose sugars. High-throughput technologies, such as transcription analysis [71,91, 109,132], enzyme and metabolite analysis [110], and proteome analysis [57], have been used. In many cases, the mutations and alterations observed in mutant strains are the same as have been earlier rationally engineered, con­firming previous knowledge and hypotheses about control and regulation of pentose metabolism. So far, no report exists where completely novel infor­mation would have been obtained from high-throughput molecular analyses. Thus, the investigations have mainly served to confirm and demonstrate the validity of the technologies.

5

Because dependence on nutritional supplementation increases the process cost, the ideal biocatalyst should produce high amounts of ethanol in sim­ple mineral salts growth medium. While KO11 and LY01 both attained high ethanol yields and titers in rich media, these microbial biocatalysts perform poorly in minimal media. With nutritional supplementation, KO11 produced 45 g L-1 ethanol from 100 g L-1 glucose in 72 h; in minimal medium less than 30 gL-1 were produced in 96 h [31]. Results for LY01 were similarly disap­pointing: the final cell mass and ethanol titer attained in minimal medium were tenfold lower than in rich medium [32]. Considering that these strains were selected in rich media, this is not a surprising result. The low ethanol production by KO11 in minimal media has been attributed to suboptimal partitioning of pyruvate for biosynthesis [33,34]. Low acetyl CoA and high NADH levels result in inhibition of citrate synthase, limiting the availability of 2-oxoglutarate for biosynthesis. 2-Oxoglutarate is required for the biosynthe­sis of many amino acids and is an important compound for osmotic tolerance. This proposed inhibition of citrate synthase was supported by the finding that expression of a NADH-insensitive citrate synthase from Bacillus increased the growth and ethanol production of KO11 by about 75% [33].

The ability of microbial biocatalysts to retain ethanologenicity over time without dependence on antibiotics is important for minimizing production costs. While instability of KO11 has been reported [35,36], other reports have demonstrated maintenance of KO11 ethanologenicity for up to 27 days in continuous stirred tank and fluidized beds reactors [37].

In addition to the production of 48 gL-1 ethanol in rich media, KO11 also produced up to 192 mgL-1 of the undesirable co-product ethyl acetate. An es­terase with ethyl acetate hydroylase activity (estZ) from Pseudomonas putida was introduced into KO11 and the presence of this enzyme reduced the ethyl acetate level to less than 20 mg L-1, a level comparable to that of yeast fermen­tation [38].

2.2

Pretreatment methods such as solvent extraction (organosolv pulping) [57] or ammonia fiber explosion treatment (AFEX) [58] either modify or remove lignin, while a large proportion of lignin remains intact in the solid phase after SO2-catalyzed steam pretreatment [59] or dilute acid pretreatment [60]. Since lignin is intertwined amongst cellulose and hemicellulose, varying the pretreatment method or conditions employed to improve cellulose or hemi — cellulose yields will undoubtedly affect the lignin content [57,59]. For ex­ample, it has been shown that the amount of lignin in the solid fraction of the substrate increases as the severity of SP is increased. Based on 13 severity fac­tors used during SO2-catalyzed SP of corn fiber, it was shown that the amount of lignin in the solid phase increased as the severity of the pretreatment was raised [59]. The lignin was most likely concentrated in the solid fraction due to the solubilization and degradation of the carbohydrates as the severity was raised [59]. It should be noted that the Klason method that is commonly used to estimate the lignin content of the pretreated substrate can result in arti­ficially high values for lignin, as sugar degradation products and entrapped low molecular weight phenolics can also be measured as “lignin”. For ex­ample, it was shown that the Klason lignin contents of steam-pretreated aspen at a series of severities ranged from 6-30%. However, the supposedly corres­ponding methoxyl groups in the samples were only in the 0.8-7.7% range. This possible elevation of lignin values should be taken into consideration when determining Klason lignin content of a steam-pretreated substrate [61]. Realizing that it is difficult to reduce the amount of substrate lignin by fine — tuning SP process conditions, various researchers have explored methods of “post-treatment” to reduce the lignin content of steam-pretreated substrates.

In the past [62-64], we have tried to enhance the removal of lignin from steam-pretreated substrates, and consequentially increase the rate and ex­tent of hydrolysis by cellulases, by applying several chemical post-treatments (Table 1). Oxygen-alkali and hydrogen peroxide post-treatments removed similar amounts of lignin and thus improved hydrolysis. However, of note, we also showed [64] that the removal of only 7% of the lignin from a steam — pretreated Douglas-fir substrate using a cold NaOH treatment resulted in a 30% improvement in hydrolysis yields, indicating that, in addition to the amount of lignin, the location of lignin is also an important factor affect­ing hydrolysis. Palonen et al. [65] have applied an alternative delignification method employing laccase enzymes in combination with mediators to steam — pretreated softwood, resulting in a slight release of aromatics into the system (delignification was not measured) and a corresponding 21% increase in hydrolysis yield. These researchers also showed that the oxidation of lignin surfaces by the application of laccases in the absence of mediators, as shown by others with pulp fibers [66,67], also resulted in a 13% improvement in hydrolysis yield. These results suggest that the modification of lignin surfaces may also play a role in reducing its inhibitory effect on hydrolysis, perhaps affecting non-productive binding of cellulases to lignin. Since most of the studies have been concerned with altering lignin content by affecting the pretreatment conditions or applying post-treatments, there have only been

limited studies linking the various chemical structures in lignin to changes in hydrolytic activity of cellulases.

3.2