We have combined biological production of ethanol, hydrogen, and methane in the Maxifuel concept (Fig. 2). The concept is designed to address the major barriers for bioethanol production from lignocellulosic materials. The overall process outline has been defined to yield the maximum amount of biofuels per unit of raw material and to increase the process benefit by utilization of the residues for further energy conversion and by-product refining. The main product is bioethanol for use as transportation fuel and emphasis has been on optimizing ethanol production. The supply and efficient conversion of the raw material is the major economic burden of bioethanol production and full and optimized use of the raw material is a key to success. Production of other bio­fuels such as methane and hydrogen, and other valuable by-products such as
a solid fuel from the parts of the biomass not suitable for ethanol production, adds full value to the overall process. The concept exploits an environmen­tally friendly way of producing bioethanol where recirculation and reuse of all streams produced in the process are integrated into the process. This is in contrast to most other bioethanol process schemes where water has to be added continuously and toxic waste water is left after the process. The basic ideas of producing biogas along with bioethanol and then to recycle the pro­cess water, or part of the process water, within the process are patented [14]. A combination of these innovative ideas along with the best available tech­nologies has ensured an economic feasibility with a competitive advantage over other concepts. The development of the optimized process of bioethanol production from lignocellulosic biomass can be further integrated into a con­ventional bioethanol production where corn or grain fibers will be a residue of low value. Conversion of this fraction into ethanol can increase the produc­tivity by up to 20% along with an improvement of the protein feed produced in the process (Fig. 3).

The Maxifuel concept is patented and consists of the following process steps (Fig. 2):

• Pretreatment

• Hydrolysis

• Fermentation of C6 sugars

• Separation

• Fermentation of C5 sugars

• Anaerobic digestion of process water and recirculation

All fermentable carbohydrates in the raw material are converted into ethanol and hydrogen, while much of the unused fraction such as residues from

tion can be recirculated to the pretreatment unit and used together with fresh raw material.

4.1

Barbel Hahn-Hagerdal1 (И) • Kaisa Karhumaa1 • Marie Jeppsson1,2 •

Marie F. Gorwa-Grauslund1

Applied Microbiology, Lund University, P. O. Box 124, 221 00 Lund, Sweden Barbel. Hahn-Hagerdal@tmb. lth. se

2 Present address:

GS Development AB, Jagershillgatan 15, 213 75 Malmo, Sweden

1 Introduction……………………………………………………………………………………………… 148

2 Xylose……………………………………………………………………………………………………… 149

2.1 Xylose Utilization Pathways………………………………………………………………………. 158

2.2 Expression of XI in S. cerevisiae……………………………………………………………………. 158

2.3 Expression of XR and XDH in S. cerevisiae……………………………………………………. 159

3 Arabinose…………………………………………………………………………………………………. 160

3.1 Arabinose Utilization Pathways…………………………………………………………………. 160

3.2 Engineering Arabinose Utilization in S. cerevisiae…………………………………………… 161

4 Improving Ethanolic Fermentation by Pentose-Utilizing S. cerevisiae. . . 162

4.1 Sugar Transport………………………………………………………………………………………… 163

4.2 Improving the Conversion of Xylose to Xylulose…………………………………………… 164

4.2.1 Cofactor Dependence……………………………………………………………………………….. 164

4.2.2 Activity of Initial Pentose Pathway Enzymes……………………………………………… 164

4.2.3 GRE3 Deletion…………………………………………………………………………………………. 165

4.3 Xylulokinase……………………………………………………………………………………………. 165

4.4 Pentose Phosphate Pathway……………………………………………………………………… 166

4.5 Engineering the Redox Metabolism of the Cell…………………………………………….. 167

4.5.1 Oxidative PPP…………………………………………………………………………………………. 167

4.5.2 Transhydrogenase and Redox Enzymes……………………………………………………… 168

4.6 Glycolytic Flux…………………………………………………………………………………………. 169

4.7 Other Modifications…………………………………………………………………………………. 169

4.8 Random Methods…………………………………………………………………………………….. 170

5 Industrial Pentose-Fermenting Strains…………………………………………………………. 170

5.1 Inhibitor Tolerance…………………………………………………………………………………….. 171

5.2 Strain Stability…………………………………………………………………………………………… 171

5.3 Fermentation of Hydrolysates……………………………………………………………………. 172

6 Conclusion and Future Outlook………………………………………………………………….. 172

References……………………………………………………………………………………………………. 173

Abstract The introduction of pentose utilization pathways in baker’s yeast Saccharomyces cerevisiae is summarized together with metabolic engineering strategies to improve

ethanolic pentose fermentation. Bacterial and fungal xylose and arabinose pathways have been expressed in S. cerevisiae but do not generally convey significant ethanolic fermen­tation traits to this yeast. A large number of rational metabolic engineering strategies directed among others toward sugar transport, initial pentose conversion, the pentose phosphate pathway, and the cellular redox metabolism have been exploited. The directed metabolic engineering approach has often been combined with random approaches in­cluding adaptation, mutagenesis, and hybridization. The knowledge gained about pentose fermentation in S. cerevisiae is primarily limited to genetically and physiologically well — characterized laboratory strains. The translation of this knowledge to strains performing in an industrial context is discussed.

Keywords Arabinose • Ethanol • Fermentation • Lignocellulose • Xylose • Yeast

Abbreviations

G6PDH Glucose-6-phosphate dehydrogenase

GAPDH Glyceraldehyde-3-phosphate dehydrogenase

mRNA Messenger RNA

PPP Pentose phosphate pathway

RKI Ribose-5-phosphate ketol-isomerase

RPE Ribulose-5-phosphate 3-epimerase

TAL Transaldolase

TKL Transketolase

XDH Xylitol dehydrogenase

XI Xylose isomerase

XK Xylulokinase

XR Xylose reductase

1

Introduction

When in the late 1970s it was discovered independently in two laborato­ries in North America [1,2] that baker’s yeast Saccharomyces cerevisiae could ferment the pentose sugar xylulose to ethanol, it was proclaimed that the development of recombinant xylose-fermenting strains of S. cerevisiae was a task that would be efficiently solved within a couple of years. Still, more than 25 years later, only a limited number of industrial S. cerevisiae strains that ferment pentose sugars have been generated [3-9]. Furthermore, there are relatively few studies on the performance of these strains under industrial conditions in lignocellulosic hydrolysates [6,10-14]. The difficulty in devel­oping efficient pentose-fermenting S. cerevisiae strains is no doubt that the regulation of metabolism in the eukaryotic yeasts is much less understood than that of, for example, the prokaryotic bacterium Escherichia coli. Conse­quently, the research on pentose-fermenting strains of S. cerevisiae has had the spin-off effect of generating more knowledge on the metabolism of this species, not least in relation to other yeasts.

The rationale for developing pentose-utilizing S. cerevisiae strains relies on the fact that this yeast has been used for the industrial production of ethanol and carbon dioxide as long as human history has been recorded. Presently,

S. cerevisiae forms the basis for the world’s largest fermentation industry pro­ducing beer, wine, potable and industrial ethanol, and baker’s yeast. In add­ition, this organism serves as a eukaryotic model organism with an intensely studied cell biology and arrays of genetic engineering tools [15]. However, the most important reason for developing pentose-fermenting S. cerevisiae is the fact that such strains can be integrated into existing ethanol plants al­ready using this yeast. Two independent investigations have estimated that integrated approaches to the production of lignocellulosic ethanol will reduce the production cost by nearly 20% [16,17].

This chapter summarizes the metabolic engineering approaches taken to develop pentose-fermenting strains of S. cerevisiae. Different engineer­ing strategies and their physiological context are described below, and the respective fermentation results from each study are chronologically sum­marized in Tables 1-4. Metabolic engineering for arabinose utilization is reported separately, since engineering L-arabinose utilization in S. cerevisiae has only recently been addressed. As will be detailed below, the fermenta­tion of pentose sugars is governed by carbon catabolite repression and by reoxidation of reduced cofactors. Fermentation results of recombinant S. cere­visiae strains have therefore been summarized in relation to batch (Tables 1 and 2) and continuous culture (Tables 3 and 4), and in relation to anaerobic (Tables 1 and 3), oxygen-limited (Table 2), and aerobic conditions (Table 4). Moreover, the data have been organized in relation to the respective con­trol strain to highlight the relative improvement of a particular engineering strategy. Studies that do not use the four aforementioned experimental con­ditions, or for which information on fermentation parameters is insufficient, have been omitted.

2

Metabolic engineering is defined as the improvement of cellular activities by manipulation of enzymic, transport and regulatory functions of the cell with the use of recombinant DNA technology [6]. After the successful expression of a XI in S. cerevisiae [42], reactions downstream of D-xylulose and the, presumably Gre3-dependent, formation of xylitol were identified as priority targets (see previous section).

As it is unlikely that the high capacity of glycolysis in S. cerevisiae would limit D-xylose fermentation rates; limitations in D-xylose fermentation are likely to reside either in the reaction catalysed by xylulokinase or in one of the four reactions of the non-oxidative pentose phosphate pathway. Modulat­ing the flux through a certain pathway by up-modulation of single enzymes often has little effect, as can be shown by metabolic control analysis [50]. Hence, it was decided to simultaneously increase the levels of all five enzymes. To this end, the S. cerevisiae structural genes encoding xylulokinase (XKS1), ribulose-5-phosphate epimerase (RPE1), transketolase (TKL1), transaldolase (TAL1) and ribulose-5-phosphate isomerase (RPI1) were over-expressed to­gether with the Piromyces sp. E2 XylA gene [43]. Since the non-specific aldose reductase encoded by GRE3 had previously been implicated in xyli — tol formation by S. cerevisiae, this gene was also deleted in the engineered strain [45,66].

Research on pentose metabolism in S. cerevisiae is increasingly impeded by the fact that key biochemical intermediates can no longer be purchased commercially [35,43]. While this precluded enzyme-activity assays for several of the over-expressed genes, mRNA analysis indicated that over-expression, either from strong constitutive promoters inserted in front of chromosomal genes or from plasmid-borne expression cassettes, was successful.

Remarkably, the S. cerevisiae strain (RWB 217) harbouring the six over­expressions and single deletion was directly capable of anaerobic growth on D-xylose as the sole carbon source at a growth rate of 0.09 h-1 [43]. Start­ing with a low-density inoculum, this strain consumed 20 g L-1 of D-xylose within 40 h, with an ethanol yield on D-xylose of 0.43 gg-1. This ethanol yield, which is lower than the theoretical yield of 0.51 gg-1 due to the for­mation of biomass and glycerol, was virtually identical to the ethanol yield found on glucose in exponentially growing, anaerobic S. cerevisiae cultures. Deletion of GRE3 reduced xylitol production to trace amounts (0.4 mM from 20 g L-1 D-xylose), indicating that alternative D-xylose — or D-xylulose reduc­ing enzymes were active at very low rates in this S. cerevisiae background. In the engineered strain, D-xylulose no longer accumulated in the broth, indicating that limitations downstream of D-xylulose had been successfully eliminated.

In an independent study, Karhumaa et al. (2005) expressed the XI gene from T. thermophilus together with the same combination of pentose phos­phate pathway enzymes [35]. In these strains the specific activity of XI was 0.008-0.017 |xmol (mg protein)-1 min-1 at 30 0C. In contrast to the efficient anaerobic growth of the above-described S. cerevisiae expressing the Piromyces sp. E2 XI, D-xylose consumption by the T. thermophylus XI-containing strain (TMB 3045) was not observed under aerobic conditions. After additional se-

lection, a strain capable of aerobic growth on D-xylose at a maximum specific growth rate of 0.045 h-1 was isolated (TBM 3050). Confusingly, although the abstract claims anaerobic production of ethanol, the experimental description and results section describe the production of 0.29 g ethanol (g D-xylose)-1 at a rate of 2.4 mg (gbiomass)-1 h-1 under oxygen-limited conditions [35]. The ethanol production rates, are more than 400-fold lower than observed in the Piromyces XylA-based strain [35,42]. This observation, combined with the interesting observation that TMB 3045 and TMB 3050 display almost identi­cal specific growth rates on D-xylulose, indicates the importance of high-level functional expression of XI for efficient D-xylose fermentation.

In lignocellulosic hydrolysates, D-xylose is generally the second most abundant sugar, with glucose accounting for the majority of the fermentable sugar [24,46,69]. Rapid consumption of glucose-xylose mixtures — either sequential or simultaneous — is therefore crucial for successful industrial im­plementation. When the metabolically engineered strain RWB 217 (described above) was grown in anaerobic batch cultures on mixtures of 20 g L-1 glucose and 20 g L-1 D-xylose (Fig. 5), sequential utilisation was observed. Although both sugars were consumed within 40 h, D-xylose consumption only com­menced when the glucose concentration dropped below 4 gL-1. Instead of increasing exponentially, as anticipated based on the kinetics of D-xylose con­sumption in D-xylose-only cultures, the specific rate of D-xylose consumption decreased over time. Clearly, the kinetics of D-xylose consumption by cells grown in the presence of glucose were sub-optimal. This challenge was ad­dressed by evolutionary engineering.

6

Xylitol has recently been recognized as one of the top 12 value-added chem­icals from biomass by the DOE [139]. This pentahydroxy sugar alcohol is commonly used to replace sucrose in food products and in toothpastes as a natural, non-nutritive sweetener that inhibits dental caries [140]. In add­ition, xylitol can serve as a valuable synthetic building block for deriva­tives intended for new polymer opportunities [139]. Production of xylitol, which typically involves hydrogenation of xylose derived from hemicellulose — xylan hydrolysates with an active catalyst such as nickel, ruthenium, or rhodium [139], is currently very limited. Numerous yeast strains have been developed that are capable of producing xylitol in complex medium [141 — 144]. Xylitol production (up to 237 gL-1) by Candidata tropicalis has been optimized by growth in complex media containing urea and numerous ex­pensive vitamin supplements [145]. More recently, strain PC09 was derived from E. coli W3110, which is capable of fermenting a broad range of sug­ars in mineral medium. PC09 can process glucose and xylose blends into xylitol by using an NAD(P)H-dependent xylose reductase from Candida boi — dinii (CbXR) to reduce xylose to xylitol, whereas glucose serves as the cell growth substrate and to regenerate the reducing equivalents [146]. Resting cells and controlled fermentations of PC09 produced 71 and 250 mM xylitol while consuming 15 and 150 mM glucose, respectively. In the controlled fer­mentations, approximately 25 mM xylulose was formed as co-product [146].

Because glucose was used to regenerate reducing equivalents and was not converted to xylitol, the xylitol yield was quantified in terms of a molar yield of reduced product formed per glucose consumed. In the case of zero growth, a maximum molar yield of 10-12 is expected; resting cells and controlled fermentations of PC09 had molar yields of 4.7 and 1.7, respectively. While the molar yield is relatively low compared to the theoretical maximum, this process could prove to be more economical after further optimization and metabolic engineering.

5.4

Process simulation of ethanol production from spruce using a process con­cept based on SO2-catalyzed steam pretreatment followed by SSF, as shown in Fig. 3 ([20], Wingren et al. 2007 (submitted)), has been used to illustrate the effect of various process parameters on the energy demand and on the ethanol production cost. The general conclusions are, however, also valid for most of the process configurations described in Table 1. The model input was based on experimental data obtained from a process development unit. SSF was performed at 10% WIS with 2 gL-1 yeast. In the model, the overall ethanol yield was 296 liters per metric dry ton, corresponding to 69.4% of the theoretical based on the hexosan content in the raw material. Pentose fer­mentation was not included. Regarding production cost data, the proposed ethanol plant is assumed to be located in Sweden, with a capacity of 200 000 dry tons of raw material annually.

The ethanol yield affects both the raw material and capital costs and is the single most important parameter in reducing the cost of ethanol pro­duction, as was already stated in 1988 [39]. High energy efficiency is also of great importance for the process to be economically feasible. In most techno­economic evaluations, live steam for the process is generated in a steam boiler by burning part of the solid residue. From the excess solids it is possible to generate heat and electricity or pellets that can be sold to improve the pro­cess economics. Thus, the energy demand of the process affects the amount of solid residue that may add to the income as a solid fuel co-product and, therefore, it is very important for the process to be energy-efficient.

The heat duty of the process depends to a large extent on the process con­figuration. For the process alternative described above, the heat duty of the energy-demanding process steps is shown in Fig. 5. The white bars represent the primary steam demand while the gray bars represent the amount of sec­ondary steam that is generated in each process step. The overall process heat duty, i. e. the total energy demand in the form of boiler-generated steam, is the sum of the black bars. Distillation (including preheating of the SSF broth) and evaporation account for the major part of the process energy demand. The contributions from pretreatment and drying, with the latter assumed to work as a steam dryer, are comparatively small, due to the generation of secondary steam in these process steps.

The energy demand of the distillation step, in which the ethanol in the mash from fermentation is concentrated, is highly dependent on the ethanol

Ethanol feed concentration (% [w/w])

Fig. 6 Energy demand in the distillation step, where ethanol is concentrated to 94 wt %, as a function of the ethanol feed concentration. The step was assumed to consist of two stripper columns (25 trays each) and a rectification column (35 trays) heat integrated by operating at different pressures. The inlet feed temperature was increased from 80 °C to the boiling temperature before entering each stripper column feed concentration, as shown in Fig. 6. The distillation step normally consists of a stripper column, in which the ethanol is separated from all solid and non-volatile compounds, and a rectification column, in which the ethanol is concentrated close to the azeotropic point. The implementation of heat inte­gration, for instance by using the overhead vapor from the stripper as the heat
source in the reboiler of the rectification column, significantly reduces the en­ergy demand. Nevertheless, it is of great importance to obtain a high ethanol concentration in the distillation feed. In a starch-based process the ethanol concentration in the stream entering the distillation step is normally above 8% (w/w). In a lignocellulose-based process, however, the aim has been to reach at least 4-5% (w/w) ethanol. In addition, a high ethanol concentration results in a high concentration of non-volatile compounds, which also leads to a decrease in energy demand in the evaporation step.

Recirculation of process streams is one way of reducing the overall energy demand, which results in a decrease in overall production cost, as shown by Wingren et al. [38]. Recirculation of part of the stream after distillation back to the fermentation step would result in an increased concentration of non­volatiles and thus a reduction in the energy demand in the evaporation step. Recirculation of part of the stream before distillation would also result in an increase in the ethanol concentration and thus a reduction in the energy de­mand in both the distillation and evaporation steps. This is true for both the SSF and SHF configurations. However, in the same study it was shown that it is even more beneficial to increase the substrate concentration in the SSF step. This would affect not only the costs related to distillation and evaporation, but also the cost of SSF. On the basis of this fact, one of the main objectives of several experimental studies performed during recent years has been to increase the substrate concentration in SSF [40-43]. This results in reduced water consumption, which greatly reduces the energy demand for distillation and evaporation, provided that the ethanol yield is maintained at a high level. In Fig. 7, the process heat duty (in MJ L-1) and the overall production cost

(in US$ L-1) are presented as functions of the WIS concentration in SSF. The ethanol yield and the amount of yeast (NB: not the yeast concentration), were the same as in the 10-% WIS case when varying the WIS concentration. The reduction in production cost is due to an increase in co-product credit and a reduction in the fixed capital cost.

Process simulations clearly demonstrate the potential reductions in pro­duction cost and energy demand that can be obtained by running SSF at higher substrate concentrations. However, given the large number of com­pounds involved, and due to the fact that they may act synergistically, it is impossible to predict the impact of increased concentrations on the perform­ance of the yeast and enzymes using process models. Effects on parameters such as productivity (yield, residence time), yeast and enzyme dosages have to be determined experimentally, preferably on pilot scale.

Savings in energy demand can also be accomplished by changes in the process design. Evaporation is the traditional, but energy-demanding, way to concentrate the water-soluble, non-volatile components in the stillage stream. To reduce the energy requirements for evaporation, multiple evaporation ef­fects are used. This has a significant effect on the overall process heat duty, as shown in Fig. 8. (In the simulation results presented in Figs. 6 and 7, evapo­ration was carried out with five effects.) The energy savings have, of course, to be weighed against the increase in capital cost. Also shown in Fig. 8 is a case where the use of mechanical vapor recompression (MVR) has been implemented in the evaporation unit. In a traditional multiple-effect evap­orator system, a large proportion of the energy supplied ends up as latent heat in the vapor phase leaving the last effect in the evaporator. This vapor is

§

normally condensed using cooling water. Another option is to compress the vapor, thereby raising the temperature to a level at which the latent heat can be utilized. The vapor can then be used as a heating medium to replace most of the primary steam. When compression is carried out by aid of a mechanical compressor the process is referred to as MVR. An electrical motor or a steam turbine provides power to the compressor. The overall process heat duty was reduced from 15.1 (base case configuration) to 10.3 MJ L-1 when MVR was applied to the evaporation step (Fig. 8), while the overall electric power re­quirement was estimated to increase from 2.2 (base case configuration) to

2.8 MJ L-1 (data not shown).

It has also been proposed that the entire evaporation step be replaced by an anaerobic digestion step, in which most of the organic material (unfermented sugars, acids, yeast, etc) is converted to biogas mainly consisting of methane and carbon dioxide. This was estimated to reduce the production cost by about 7%. The performance of such a system is dependent on a number of parameters such as the composition of the feed, residence time, temperature, etc. A crucial question is also how to handle the sludge from the anaerobic digestion. Further investigation is required since very limited data regarding the performance of this kind of system have been published.

3.2

Natural S. cerevisiae strains reduce xylose to xylitol with an endogenous xylose (aldose) reductase encoded by the GRE3 gene [28]. Xylitol strongly inhibits the activity of XI [94], and therefore deleting the GRE3 gene im­proves efficient xylose utilization in XI-expressing S. cerevisiae strains [95]. Improved ethanol yields at the expense of reduced xylitol yields were in­deed observed for XI-carrying strains [96,97] (strain RWB217, Table 1). Furthermore, the GRE3 deletion decreased xylitol formation also in strains carrying XR and XDH, albeit only under continuous fermentation [98] (strain TMB3120, Tables 2 and 3). However, the aldose reductase endoded by GRE3 belongs to a group of generally stress-induced proteins [28] and the deletion of it reduces the growth by 30% [96]. This limits the usefulness of GRE3 deletion in strains aimed at industrial applications.

4.3

Hemicellulose refers to a number of heterogeneous structures, such as (ara — bino)xylan, galacto(gluco)mannan, and xyloglucan [107,108]. These chem-

REF SFI EG1 CEL5 CEL5

PASC

ically diverse polymers are linked together through covalent and hydrogen bonds, as well as being intertwined. Although many pretreatments remove variable amounts of hemicellulosics, it remains imperative from an economic perspective that sugars contained in the hemicellulose fraction of lignocellu- lose are also converted to ethanol [9,36]. The hydrolysis of xylans, the second most abundant sugar polymer in nature, and utilization of xylose, its main constituent, are therefore crucial in a viable CBP configuration.

The cross-linked and partially crystalline nature of the matrix offer great resistance to enzymatic hydrolysis. Furthermore, as the structure of xylan

is variable, involving not only linear P-1,4-linked chains of xylose, but also branched heteropolysaccharides; its degradation requires the synergistic ac­tion of a range of different enzymes [109-111]. To date, hydrolytic enzymes for the cleavage of almost all chemical bonds found in plant structures have been identified in microbial sources (Fig. 2). Hemicellulases, such as в- xylanases and в-mannanases, have drawn attention as they can help facilitate industrial processes such as bleaching in the pulp and paper industry.

There have been several reports of the successful expression of hemi — cellulases in S. cerevisiae (Table 3). Xylan hydrolyzing enzymes such as в- xylanase, в-xylosidase, and auxiliary enzymes such as a-glucuronidase and a-arabinofuranosidase have all been produced successfully in yeast [114,120, 121,128]. The heterologous production of mannanase and a-xylosidase, ac­tive against mannan and xyloglucan, respectively, was also reported [107,124, 127].

Degradation of the e-1,4-xylan backbone requires the action of endo-в — 1,4-xylanases (e-1,4-D-xylan xylanohydrolase EC 3.2.1.8) and в-xylosidases (e-1,4-D-xylan xylohydrolase EC 3.2.1.37) (Fig. 2a). cDNA copies of в- xylanase encoding genes cloned from the yeasts Cryptococcus albidus and Aureobasidium pullulans and the filamentous fungi A. niger, A. kawachii, and

T. reesei have been expressed in S. cerevisiae under transcriptional control of glycolytic promoters [112-114,116,129]. Secreted active enzyme could be as­sayed in all cases. The cDNA copy of the T. reesei в-xylanase II (xyn2) was expressed in S. cerevisiae under the transcriptional control of the PGK1 and ADH2 promoters [114]. Efficient secretion of the heterologous в-xylanase was achieved by the native T. reesei xyn2 secretion signal and the recombinant в-xylanase was 27 kDa in size. The molecular mass of the mature protein in T. reesei was found to be 21 kDa, with virtually no glycosylation. The extra molecular weight of the heterologous Xyn2 protein secreted by S. cerevisiae was shown to be the result of hyperglycosylation of the protein; however, the extra sugar moieties did not influence the activity of the enzyme.

The в-xylosidase encoding gene of Bacillus pumilus (xynB) was cloned from a genomic DNA library and expressed in S. cerevisiae under tran­scriptional control of the S. cerevisiae ADH2 promoter [118]. To promote secretion of the enzyme, the gene was fused in reading frame with the S. cere­visiae mating pheromone a-factor (MFa1) secretion signal. Biologically ac­tive в-xylosidase was obtained, but remained mostly cell associated. When this fusion gene and T. reesei xyn2 were coexpressed in S. cerevisiae under transcriptional control of the S. cerevisiae ADH2 promoter, a 25% increase in the amount of reducing sugars released from birchwood xylan was obtained, compared to strains expressing в-xylanase alone. However, no xylose was produced from birchwood xylan, presumably due to very low в-xylosidase activity. A cDNA copy of the A. niger в-xylosidase encoding gene was sub­sequently cloned [115]. The mature protein encoding region was fused in reading frame with the S. cerevisiae MFa1 secretion signal to ensure secretion

U = micromole substrate released/min, DCW = dry cell weight, NR = not reported; sub­strate used for activity determination is given in parentheses; italics indicate calculation based on assumptions (0.45 g DCW/g glucose, 0.45 g protein/g DCW, 1.3 x 107 cells/g DCW, 1 OD(600) = 0.57 g DCW/L).

BG-xylan = birchwood glucuronoxylan, PNP-|3-X = p-nitrophenyl-|3-D-xylopyranoside, AZCL-XG = azurine-dyed cross-linked xyloglucan, ABIU = aldobiouronic acid, ATRU = aldotriouronic acid, ATEU = aldotetraouronic acid, PNPA = p-nitrophenyl-a-L-arabino — furanoside, LBG = locust bean gum, INM = ivory nut mannan, PGGM = pinewood galactoglucomannan, PNPaGal = p-nitrophenyl-a-D-galactopyranoside from S. cerevisiae and secreted в-xylosidase activity was obtained. When this fusion gene and T. reesei xyn2 were coexpressed in S. cerevisiae, high levels of в-xylanase and в-xylosidase activity were obtained in autoselective strains grown in rich medium. Coproduction of these two enzymes allowed this re­combinant S. cerevisiae strain to convert up to 46% of birchwood xylan to xylose [115].

Using a cell surface engineering system based on a-agglutinin, S. cere­visiae strains displaying the в-xylanase II separately or in combination with the в-xylosidase from Aspergillus oryzae on the cell surface were con­structed [119,130]. When xylan was incubated with high cell concentrations of these strains, HPLC analysis showed that xylose was the main product of the yeast strain codisplaying the в-xylanase and в-xylosidase, while xylobiose and xylotriose were detected as the main products of the yeast strain dis­playing the в-xylanase. Subsequently, a xylan utilizing S. cerevisiae strain was constructed by introducing genes for xylose utilization, specifically, those en­coding xylose reductase and xylitol dehydrogenase from Pichia stipitis and xylulokinase from S. cerevisiae into the strain codisplaying the P-xylanase and в-xylosidase. Ethanol was directly produced from birchwood xylan, and the yield in terms of grams of ethanol per gram of carbohydrate consumed was 0.30 g/g. This strain, though not able to completely degrade xylan and still suffering from the redox imbalance problem during xylose utilization, supports the potential of using S. cerevisiae in a CBP configuration for con­verting xylan to ethanol.

In order to achieve complete degradation of complex substituted xylans, a series of accessory or debranching enzymes are also needed, namely a-D — glucuronidases (EC 3.2.1), a-L-arabinofuranosidases (a-L-arabinofuranoside arabinofuranosidase EC 3.2.1.55), and acetylesterases or acetyl xylan es­terases (EC 3.1.1.6) [131]. Successful expression of an a-glucuronidase in S. cerevisiae was recently reported [120]. The secreted enzyme was active on aldouronic acids from aldobiuronic to aldopentauronic acid. The T. ree — sei a-arabinofuranosidase encoding gene, abfl, was expressed in S. cere­visiae and the resulting enzyme released L-arabinose from p-nitrophenyl-a — L-arabinofuranoside and arabinoxylans [117]. Successful expression of the gene encoding a-L-arabinofuranosidase B (abfB) from A. niger was also shown [121,122].

The major hemicelluloses in softwoods are acetylated galactoglucoman — nans [107]. These consist of a backbone of P-1,4-linked mannose and glucose residues substituted with a-1,6-linked galactosyl side groups. Mannanase (endo-1,4-P-mannanase; mannan endo-1,4-|3-mannosidase; EC 3.2.1.78) ran­domly hydrolyzes the 1,4-^-mannosidic bonds of the main chain of gluco — mannan and galactomannan (Fig. 2c). Endomannanases of T. reesei (man1), Aspergillus aculeatus (man1), and Orpinomyces PC-2 (manA) have all been expressed and secreted in S. cerevisiae [107,108,124]. The secreted en­zymes showed activity toward locust bean gum and ivory nut mannan with the A. aculeatus enzyme exhibiting the highest titer and activity. The en­zyme a-galactosidase (a-D-galactoside galactohydrolase) catalyzes hydrolysis of a-1,6-linked galactosyl residues from galacto(gluco)mannans and simple oligosaccharides such as raffinose and is required for the complete hydro­lysis of galactomannan [125]. Three a-galactosidases, agl1, agl2, and agl3, from T. reesei were cloned and expressed in S. cerevisiae. The recombinant enzymes were able to hydrolyze raffinose, melibiose, and p-nitrophenyl-a-D — galactopyranoside and release galactose from galacto(gluco)mannan.

Xyloglucan is the main hemicellulosic polysaccharide present in the pri­mary cell walls of dicotyledonous plants [127]. It consists of a linear 1,4- в-linked D-glucan backbone that carries a-D-xylosyl, P-D-galactosyl-1,2-a — D-xylosyl, and a-L-fucosyl-1,2-P-D-galactosyl-1,2-a-D-xylosyl side chains at­tached to the OH-6 of в-glucosyl residues. a-Xylosidase releases the unsubsti­tuted side chain xylosyl residue attached to the backbone glucosyl residue sit­uated farthest from the reducing end of the molecule (Fig. 2d). When a gene encoding the Arabidopsis thaliana a-xylosidase (AtXYLl) was expressed in S. cerevisiae, activity could be detected inside the cell [127]. A xyloglucan — specific endo-P-1,4-glucanase from Aspergillus aculeatus was isolated and ex­pressed in yeast [126]. The recombinant enzyme was active in yeast, showing clearing zone formation on azurine-dyed cross-linked xyloglucan containing plates.

As the technologies of pentose sugar utilization and hemicellulase produc­tion in S. cerevisiae mature, integration of these processes and subsequent single-step processing of biomass hemicellulose to commodity products such as ethanol becomes ever more easily envisioned.

6

Several studies on ethanol production by wild-type strains of Z. mobilis on industrial starch-based raw materials have been reported. Bringer et al. [69] investigated an industrial-scale process and Poosaran et al. [70] evaluated a cassava-derived starch hydrolysate. In the latter case in a batch culture at controlled T = 30 °C and pH = 5.0, fermentation using Z. mobilis ZM4 gave an ethanol yield of 95% theoretical, a productivity of 6 gL-1 h-1 and a final ethanol concentration of 114 gL-1. Under the same conditions, a strain of Sac — charomyces uvarum gave an ethanol yield of 90% theoretical, a productivity of 4gL-1 h-1 and a final ethanol concentration of 106gL-1 for a cassava starch suspension (23% glucose equivalent). A comparative batch and continuous culture study with starch hydrolysate using yeast and Z. mobilis 29191 has also been reported by Beavan et al. [71].

Extensive studies with various strains of Z. mobilis have been reported by using sugar cane syrup and molasses [72-76] and for sugar beet mo­lasses [77,78] with evidence of yield reductions on sucrose based media due to production of the fructose polymer levan as by-product [3,6] and rate reductions due to high salt concentrations in the molasses. Improved produc­tivities were reported following membrane desalting of high salt-containing sugar cane molasses [72].

Most recently, Davis et al. [79] studied the fermentation of a hydrolyzed waste starch stream from flour wet milling using both Z. mobilis ZM4 and an industrial ethanol-producing strain of S. cerevisiae. With glucose concen­trations in the range 80-110gL-1, Z. mobilis ZM4 demonstrated superior fermentation characteristics. In a repeated batch process (five cycles), rapid concentration of the cells and increased productivities were achieved by cell settling between batches using the flocculent strain Z. mobilis ZM401 (ATCC 31822) as characterized by Skotnicki et al. [80]—see Fig. 6.

Similar flocculent mutants of wild-type Z. mobilis strains CP4 and ATCC 29191 have been isolated by Lawford et al. [16] and Fein et al. [81] using a spe­cially designed chemostat. These strains were deposited with the ATCC as strains 35 000 and 35 001, respectively. The use of such flocculent cultures was demonstrated to increase volumetric productivity by as much as ten-fold [82]

and may have considerable potential in future large-scale processes for more stable fermentations.

Recombinant strains of Z. mobilis developed for xylose utilization have been evaluated on various agricultural residues including oat hull hydrolysate produced by the Iogen process [40]. Oat hull hydrolysate contains glucose, xylose and arabinose in a mass ratio of 8 : 3 : 0.5. Synthetic hydrolysate (6% w/v glucose; 3% w/v xylose; 0.75% w/v acetic acid) at initial pH 5.75 was mixed with either 2 ml L-1 corn steep liquor (CSL) or 1.2 g L-1 di-ammonium phosphate as N source and used for evaluation of ethanol production. From the results it was concluded that the highest productivity was achieved with Z. mobilis ZM4 (pZB5). In this and other studies, CSL was also found to be an effective nutrient source to replace yeast extract in the fermentation media for Z. mobilis [83-85].

Further studies were reported by Mohagheghi et al. [41] with an integrant strain (designated Z. mobilis Fig. 8b) derived from ZM4 (pZB5) using over­limed corn stover hydrolysate. The hydrolysate contained 16 gL-1 glucose, 69gL-1 xylose and 11 gL-1 acetic acid at pH = 5.0. This medium was sup­plemented with 100 gL-1 glucose and diluted to various concentrations prior to fermentation. The authors found that up to 50 g L-1 ethanol was produced by the integrant strain with diluted 80% corn stover hydrolysate. Yields of 83-87% theoretical (based on sugars utilized) were reported.

One of the potential issues for large-scale Z. mobilis fermentations is whether or not contamination control is needed particularly in the presence of ethanol-tolerant strains of Lactobacilli. Such contamination constitutes a problem in many yeast-based processes and can reduce yields by an es­timated 2-5%. However, its impact is reduced as pH decreases to 3.0-3.5 towards the end of batch fermentation (in the absence of pH control). “Acid washing” of the residual yeast at this pH or lower is often used to minimize contamination in yeast subsequently used in a repeated batch process. Z. mo­bilis is more sensitive to low pH than S. cerevisiae and contamination was identified as a problem by Bringer et al. [69] in their study on an industrial — scale process for conversion of starch to ethanol using Z. mobilis although Lawford and Rousseau [63] demonstrated that lactic acid in such circum­stances is not likely to be inhibitory to Z. mobilis. Interestingly, although rarely observed in Z. mobilis fermentations due to the usual high metabolic flux rates in the ED pathway, conditions have been reported which can pro­mote lactic acid synthesis in Z. mobilis [37,85].

The issue of contamination control was addressed directly by Grote et al. [86] in which a continuous culture of Z. mobilis ZM4 was directly contaminated with a 10% (v/v) inoculum of Lactobacillus sp. isolated as an ethanol-tolerant contaminant from an industrial plant (Grain Processing Corporation, Muscatine, Iowa). It was found at D = 0.1 h-1 under condi­tions of glucose limitation, pH control at 5.0 and ethanol concentrations of 60-65 gL-1, that the addition of the contaminant caused only a temporary disturbance in the process. Steady state conditions with no evidence of sus­tained contamination were regained within five to six generations. These results suggest that contamination is not likely to be a significant problem once an active culture of Z. mobilis is established providing that the pH is maintained above 3.5-4.0. A similar conclusion was reached in a recent study [87] using an acid-tolerant strain of Z. mobilis under non-sterilized feed and operating conditions.

3

Reports on xylose fermentation by recombinant strains in industrial sub­strates are relatively few [6,10,12-14]. Laboratory strains are usually not viable in toxic lignocellulose hydrolysates, and strains with industrial back­ground must be used. In general, xylose fermentation in hydrolysates occurs more slowly than in laboratory media even by industrial strains. For example, the rate of xylose fermentation of strain TMB3400 in dilute-acid spruce hy­drolysate was an order of magnitude lower than that in mineral medium [54]. Another general observation is that very little xylitol is produced when xy­lose in lignocellulose hydrolysates is fermented by XR — and XDH-carrying industrial strains. This is most likely due to the presence of external electron acceptors in industrial media [74-76,144], paradoxically removing the prob­lem of xylitol formation, which has been considered the main drawback of the XR — and XDH-based metabolic engineering strategy.

6

Recombinant expression of the Z. mobilis homoethanol pathway has been the cornerstone of E. coli ethanologenesis. However, recent progress has en­abled ethanol production by a mutant E. coli strain lacking foreign genes [46]. Due to the inability to regenerate NAD+ and maintain redox balance, wild — type E. coli is unable to grow anaerobically in the absence of both IdhA and pflB [47]. Chemical mutagenesis was used to isolate AldhA/ApflB deriva­tives capable of anaerobic growth. The resulting strain SE2378 fermented glucose and xylose to ethanol with 82% yield. Further analysis of SE2378 revealed an essential mutation within the pyruvate dehydrogenase (PDH) operon. In native strains, pyruvate formate-lyase is primarily responsible for production of acetyl-CoA during anaerobic growth; PDH is reportedly inactive [48] or weakly active [49] under these conditions. The essential mu­tation in the pdh operon restored function during anaerobic growth and produced an additional NADH for each pyruvate. This additional NADH al­lowed the balanced production of 2 moles of ethanol per mole of glucose by a novel pathway not previously known in nature. The anaerobic spe­cific growth rate of SE2378 was reduced approximately 50% relative to the parental strain in rich media and no growth was observed in glucose mini­mal media without acetate, glutamate, or corn steep liquor supplementation. Despite growth challenges, the maximum specific productivity of SE2378, 2.24 g ethanol h-1 gcells-1, is comparable to KO11 and ethanol was pro­duced from 50 g L-1 glucose and xylose at greater than 80% of the theoretical yield.

2.5