When designing novel thermostable enzyme systems, the structural features of the substrates determine the number of enzymes needed for total hydro­lysis. The crystallinity of cellulose, the available surface area and the dis­tribution of lignin and hemicellulose are the major substrate-related factors limiting the hydrolysis rate of cellulose. An efficient pretreatment is the most straightforward solution for improving the hydrolysis rate and decreasing the amount of enzymes needed. Using various pretreatment techniques, either most of the hemicellulose or lignin is removed. It has been observed that the removal of hemicellulose has a direct correlation with the efficiency of the hydrolysis [56]. Even low amounts of residual xylan can limit the extent and rate of the hydrolysis. This can be overcome by addition of suitable hemicel — lulases, especially xylanases, to substrates with high original xylan content. Usually, the xylanase activity in commercial T. reesei preparations has been adequately high to overcome this limitation on xylan-containing substrates. Lignin content and distribution has also been proposed to be a substrate — related factor that affects the efficiency of enzymatic hydrolysis [49]. The close association of lignin and cellulose may prevent swelling of the fibrous substrate and result in limited accessibility of enzymes. The role of lignin or lignin-derived compounds in destabilising or deactivating enzymes is obvi­ously also crucial. High temperature and pressure during the pretreatment result in a variety of soluble inhibitors for the enzymes and the yeast. In this work, it was observed that the inhibition of cellulases on lignin-containing substrates was increased at higher temperatures (Figs. 4 and 5).

The optimal cellulase composition varies depending on the substrate used but usually, the major cellulases comprise two cellobiohydrolases (about 60-70% of total protein), and two major and several minor endoglucanases (about 25% of the total protein). Various models and mechanisms for the syn­ergistic action of cellulases have been proposed. These studies have focused on the T. reesei exo-exo synergism [31,53,77,80] or on the endo-exo syn­ergism [39,46,52,80]. The key role of в-glucosidase in a separate hydrolysis process has been clearly demonstrated, and is due to the end product inhi­bition of especially cellobiohydrolases caused by cellobiose [20,54]. In T. ree — sei this activity is partly mycelium-bound and obviously limits the enzyme performance in commercial T. reesei preparations. Therefore, P-glucosidase is usually supplemented, generally originating from Aspergillus niger. In­terestingly, in this work it was shown that just three major thermostable cellulases, i. e. one cellobiohydrolase and one endoglucanase supplemented by в-glucosidase, used in a preliminarily optimised ratio were able to pro­duce a hydrolysis yield comparable with that obtained with the whole set of cellulolytic and accessory enzymes present in the commercial T. reesei prep­arations. Further research would be necessary to clarify the detailed mech­anisms of these enzymes. Although the endo-exo synergism was obviously efficient enough to result in a high sugar yield, it could be further improved by optimising the thermostable cellulase components. The optimal ratio of the major enzymes was shown to be close to that of T. reesei. In this work, the individual thermostable cellulases were preliminarily screened based on their activity profiles and not based on their synergistic action. Therefore, the hydrolysis result can be considered extremely promising. Previously, thermo­stable enzymes from different organisms have not been combined to form new efficient mixtures. Expectedly, further optimisation, as well as supple­mentation of other synergistically acting enzymes would further improve the hydrolytic efficiency. In the present work, only thermostable xylanase was added to the mixture of the three cellulases (endoglucanase, cellobiohydrolase and P-glucosidase).

Previously, thermostable enzymes have only been studied as individually added proteins to improve the performance of the cellulases from T. ree — sei [62]. The T. reesei cellulase system is rapidly inactivated at temperatures above 45 °C, and the optimal temperature is generally considered to be be­low 45 °C on substrates requiring longer hydrolysis times, e. g. due to higher substrate consistency. Crude culture filtrates from various moderately ther­mophilic fungi (C. thermophilum, T. terrestris, T. aurantiacus, C. thermo — philus, M. thermophila) were added on the protein basis to a commercial T. reesei preparation. Obviously, due to the relatively high proportion of T. reesei enzymes in the mixture, and the consequent inactivation of these en­zymes at elevated temperatures, no improvement of the hydrolysis at higher temperatures could be observed. The main advantage was expected to be due to more active endoglucanases or due to a improved improved ratio of endoglucanase and cellobiohydrolase in the crude fermentation broth. In addition, unidentified enzyme activities in the preparations may also have caused some effects.

In this work, the individual cloned thermostable enzymes were produced with a T. reesei strain where the four genes encoding the major cellulases, i. e. Cel7A, Cel6A, Cel7B and Cel5A, had been deleted. Thus, only the mi­nor endoglucanases Cel12A, Cel61A and Cel45A, as well as xylanases and other accessory enzymes, were present in the T. reesei background. In add­ition, most of these activities were inactivated in a thermal treatment. Only the Cel45A was somewhat more resistant to thermal inactivation and re­tained most activity at higher temperatures. Thus, the hydrolysis results were non-disputably obtained due the cloned thermostable enzymes, and the back­ground activities were negligible. This was also clear from the hydrolysis experiments with the commercial T. reesei enzymes, showing clearly a de­creased performance at temperatures of 50 °C or above.

In addition to improved performance in the hydrolysis of lignocellulosic substrates, thermophilic enzymes allow the design of more flexible process configurations. Traditionally, T. reesei enzymes are used either in a separate hydrolysis and fermentation process (SHF) or in a simultaneous saccharifi­cation and fermentation (SSF) process. It is commonly stated that the major advantage of the SHF is that both process steps (hydrolysis and fermenta­tion) can be run under optimal conditions. Typically, hydrolysis of the SHF is carried out at around 45-50 °C at pH 5, and the fermentation at 35 °C at a lower pH. The SSF, on the other hand is usually carried out at 35 °C at pH 4.5-5. A more efficient hydrolysis is expected to take place at higher tem­peratures. In this work, using thermostable enzymes, it was indeed possible to obtain about 10 °C higher operation temperature than with the present commercial T. reesei enzyme preparations. The applicable hydrolysis tem­perature could be 60-65 °C for the hydrolysis of corn stover substrate and about 55 °C for spruce substrate. The hydrolysis rates at 55 °C were higher than those of the commercial enzymes at 45 °C. The enzymatic hydrolysis at higher temperatures would potentially reduce the reaction time and the enzyme loading.

It can be concluded that the cloned thermostable enzymes in preliminarily optimised preparations clearly demonstrate that the hydrolysis of lignocel — lulosic raw materials can be further improved, leading to potential savings in the hydrolysis costs. Previously, it has been shown that the costs of cel — lulases can be radically decreased, e. g. by improving the specific activity, by omitting the downstream processing of enzyme production or by improving the production process by other means. A mixture of only four thermostable enzymes was shown to be superior to the present commercial T. reesei prep­arations, which are comprised of at least ten enzymes acting synergistically on cellulose and on other components of the lignocellulosic substrates under optimal conditions. Further supplementation of other cellulases or accessory enzymes would expectedly further improve the hydrolysis result and the over­all economy of the process.

Acknowledgements The EU Commission is gratefully acknowledged for the financial sup­port (project number: NNE5-2001-00447, TIME).

Expression of the Piromyces sp. E2 XylA gene under control of a strong, con­stitutive TPI1 promoter on a 2^-based plasmid (pAKX002) in the haploid laboratory strain S. cerevisiae CEN. PK resulted in XI activities ranging from 0.33 to 1.1 ^mol (mgprotein)-1 min-1 in cell extracts [42]. These activities are similar to those of key enzymes of alcoholic fermentation in glucose — fermenting cultures [68]. Apparently, conditions in the cytosol of S. cerevisiae do not preclude accurate folding of the fungal XI, as has previously been re­ported for the Streptomyces rubiginosus XI [21]. In addition, in contrast to the previously expressed XI from T. thermophilus, the Piromyces XI yielded the above-mentioned activities at a temperature of 30 °C.

Although the high XI activities found in XylA-expressing S. cerevisiae strains provided an excellent starting point for further strain development, they did not as such enable a high specific rate of D-xylose fermentation. In fact, the specific growth rate in aerobic cultures on 20 g L-1 D-xylose as the sole carbon source was only 0.005 h-1 (Fig. 4). A similar very low spe­cific growth rate was found in earlier engineered S. cerevisiae strains that expressed the P. stipitis xylose reductase and xylitol dehydrogenase genes [38, 39]. The low rate of D-xylose conversion in strains with a high XI activity sug­gested that D-xylose consumption was either controlled by D-xylose transport or by reactions downstream from D-xylulose.

Since the low specific growth rates of the Piromyces XylA-expressing strains on D-xylose complicated studies in batch cultures, initial studies on D-xylose consumption kinetics and product formation were performed in anaerobic chemostat cultures grown on glucose-xylose mixtures. Anaero­bic chemostat cultivations on glucose alone demonstrated that expression of a heterologous XI did not interfere with product formation during growth on glucose [42]. However, when D-xylose was also included in the medium of the anaerobic glucose-limited chemostat cultures, a significant effect of XylA expression was observed. With 20% of the added D-xylose being consumed, a significant increase of the ethanol yield on consumed glucose was observed (from 0.40 g g-1 to 0.44 g g-1). Although no labeling studies were performed, it stands to reason that this ethanol was produced from the consumed D-xylose.

Interestingly, these anaerobic chemostat cultures of the XylA-expressing strains excreted significant amounts of D-xylulose. At a specific D-xylose con­sumption rate of 0.73 mmol (gbiomass)-1 h-1 this yeast excreted D-xylulose at a rate of 0.20 mmol (gbiomass)-1 h-1 (corresponding to 30% of consumed D-xylose), which suggested that reactions downstream of D-xylulose were rate-controlling. Moreover, small amounts of xylitol were produced in these cultivations, suggesting involvement of a non-specific aldose reductase such as encoded by GRE3 [66]. This information on D-xylulose and xylitol produc­tion was used in subsequent metabolic engineering attempts to improve the D-xylose consumption rate and to minimise xylitol formation.

5

During oxidative growth, roughly half of the sugar carbons can be diverted into cell mass and CO2 [129,130]. For this reason, bacterial production of commodity chemicals has traditionally focused on generating reduced end — products using anaerobic conditions, in order to minimize the loss of carbon as cell material or CO2. Current biological production of acetate involves complex growth conditions consisting of two separate organisms: an ini­tial fermentation of sugars to ethanol by Saccharomyces and subsequent oxidation to acetate by Acetobacter under aerobic conditions [131-133]. Strain TC36, an E. coli W3110 derivative, was engineered to merge aspects of both fermentative and oxidative metabolism for the production of acetate via a single microbial biocatalyst [77]. TC36 contains multiple chromosomal gene deletions to eliminate production of formate focA-pflB), succinate frdBC), lactate (ldhA), and ethanol (adhE), to disrupt the tricarboxylic acid cycle (sucA) and, most notably, to inactivate oxidative phosphorylation (atpFH) in order to direct the flow of carbons from sugar to acetate with minimal car­bon loss to other fermentation products, CO2, and cell mass. A maximum of 878 mM acetate was produced by TC36 in mineral salts medium. Though this is a lower titer than that achieved during ethanol oxidation by Aceto — bacter, TC36 has a twofold higher production rate, can metabolize a wide range of sugars, and requires a simple, single step process in mineral salts medium.

Pyruvate is used as a food additive, nutriceutical, weight control supple­ment, and starting material for the production of amino acids and acetalde­hyde [15,134]. Pyruvate can be produced by either chemical or biological processes. Chemical synthesis from tartrate entails the use of toxic solvents, requires a great deal of energy, and is very costly [135]. Biological produc­tion involves two auxotrophic microorganisms that require costly nutritional supplements and strict regulation of media composition [134,136], or an E. coli strain that produces pyruvate from glucose and acetate in complex medium [137]. More recently, a microbial biocatalyst has been developed for the efficient synthesis of pyruvate from sugar requiring only inexpen­sive mineral salts medium [138]. Strain TC36, an E. coli W3110 derivative described above, was used as a platform to generate the pyruvate pro­ducer TC44. This strain encompasses two additional chromosomal deletions, ackA and poxB, to allow pyruvate accumulation and eliminate acetate pro­duction. TC44 yields (0.75 g pyruvate per gram of glucose), titer (749 mM maximum) and production rate (1.2 g of pyruvate L-1 h-1) in mineral salts medium were comparable to or better than the previously described bio­catalysts, which required costly nutritional supplements and complex me­dia [138]. This strain improves the cost of pyruvate production by reduc­ing the costs of materials, process controls, product purification, and waste disposal.

5.3

The number of studies on economic aspects of ethanol production from biomass is rather limited. This depends to a large extent on the fact that the ethanol production from biomass has not yet been demonstrated on commercial scale. The ethanol production cost varies between the studies performed from about 0.13 to 0.81 US$ per liter ethanol, see Table 1. Dur­ing 2006 the selling price of bioethanol, produced from starch or sugar-based materials, has fluctuated around 0.65 US$ per liter of ethanol with a peak at 1.12 US$/liter [21]. The future selling price will be dependent on demand and availability, which is influenced by political decisions, such as the EU direc­tive mentioned before. Also tax exemptions, e. g. exemption of CO2 tax and energy tax adopted in Sweden, and protective duty, as that applied in the EU, will impact the pricing for customers.

The large differences in ethanol production costs in Table 1 can be ex­plained by variations in the process design and in the assumptions underly­ing the techno-economic evaluations. Process variations are due to different conversion technologies, e. g. an enzymatic process with SHF, SSF or SSCF or the use of various types of raw materials. Thus, for meaningful compari­son the actual differences have to be identified. The discrepancies that arise due to various assumptions, in many cases, overshadow the actual differences.

Typical examples are the raw material cost (even if the same raw material is used), plant capacity and investment parameters, e. g. pay-off time and in­terest on capital. Also, the country in which the proposed plant is assumed to be located is of importance. One of the main influences on the produc­tion cost originates from the assumed annual capacities of the ethanol plants, which varied from 196 000 to 3 110 000 tons of raw material. This has a con­siderable influence on the total production cost (Fig. 4). Another difference is found in the overall ethanol yields assumed, e. g. if pentoses are converted to ethanol or not. Also, changes in process configurations, or a change in the equipment included in the ethanol production process also influences the overall cost, e. g. whether utilities such as process steam and electricity are in­cluded. Therefore, care must be taken when comparing ethanol production costs from different studies. However, this does not mean that the economic studies are without value. They give important information about which parts of the process are most costly, and where bottlenecks, which need to be ad­dressed by further research, can be expected.

Most economic studies performed on the enzymatic bioethanol process during the past ten years have been for a configuration using some kind of

steam pretreatment employing an acid catalyst. NREL has for many years been conducting detailed techno-economic evaluations of ethanol produc­tion from lignocellulosic materials. In the 1999 report [16] hardwood (poplar) was considered as the raw material and the proposed annual capacity was 700 000 dry metric tons. The following process configuration was assumed. The raw material is pretreated with dilute sulfuric acid at 190 ° C for 10 min­utes. The liquid hydrolyzate is detoxified by ion exchange and overlimed, after which an SSF step is employed. Some of the slurry following pretreatment is used for enzyme production. In the SSF step the remaining cellulose is con­verted to glucose and both hexoses and pentoses are considered fermentable. The ethanol is removed from the mash through stripping and the stillage is dewatered by means of centrifugation. The solids, together with the con­centrated liquid from the evaporation step, are transferred to a boiler for steam and electricity production. The estimated ethanol production cost was 0.38 US$ L-1. Especially the database of physical properties [22], but also sev­eral of the unit operation models from this study, has been used by many other investigators.

Lynd et al. [23] evaluated a process based on dilute acid hydrolysis, pentose fermentation and SSF using hardwood as the raw material, assuming the fol­lowing procedure. Distillation bottoms are centrifuged and the solid residue, together with methane and sludge from the anaerobic digester, is sent to the boiler where process steam and electricity are generated. The plant capacity was assumed to be 658 000 dry tons per year. The ethanol production cost was estimated to be 0.31 US$ L-1. A techno-economic evaluation based on the same process concept and the same raw material as the above, with an annual capacity of 640 000 tons, was conducted by Stone et al. [24]. This re­sulted in an estimated production cost of 0.34 US$ L-1. In the study presented by Lynd et al. [23] a more advanced scenario was also evaluated where fu­ture improvements in conversion technology were included. These include, but are not limited to, a higher overall ethanol yield, the use of a microorgan­ism capable of not only fermenting sugars to ethanol but also of hydrolyzing the cellulose (direct microbial conversion), and shorter residence times in the process steps. With these improvements, together with an increased cap­acity (2 738 000 dry tons per year) the projected ethanol production cost was 0.13 US$ L-1.

A comparative study of an SSF-based process and a process using dilute acid hydrolysis was performed by So and Brown [25]. The SSF process used the same conversion technology as the process evaluated by Lynd et al. [23], while the dilute sulfuric acid step was assumed to be carried out at 180 °C with an acid concentration of 5 gL-1. Estimated production costs at a plant capacity of 25 million gallons of ethanol per year (equivalent to around 260 000 tons of dry raw material per year) were 0.34 US$ L-1 for the SSF-based plant and 0.36 US$ L-1 for the process employing a dilute sulfuric acid process.

In the NREL report of 2002 [26] the raw material was changed to corn stover. Several changes were also made to the model from 1999. Instead of running SSF, an SHF configuration (including pentose fermentation) was employed, where the saccharification was carried out separately prior to fer­mentation. The reason for this was to be able to carry out saccharification at a higher temperature than in the fermentation step. The enzyme produc­tion step was also removed and it was assumed that the enzymes had to be purchased from an enzyme-producing company at an estimated cost of 0.10 US$ per gallon of ethanol. This represents a projected future cost rather than the present cost. Changes were also made to the separation of solids from the stillage stream. Instead of centrifuges, as suggested in the 1999 report, a horizontal belt filter (of the type manufactured by Pneumapress Filter Corporation, CA, USA) was employed and the concentration of wa­ter insoluble solids (WIS) in the filter cake was assumed to be about 50%. The overall ethanol yield increased significantly compared to the 1999 report (from 73.6% to 85.5% concerning cellulose, while 85% yield was assumed from all hemicellulose sugars). The estimated production cost was reduced to 0.28 US$ L-1.

According to a techno-economic evaluation of ethanol production from biomass performed by “SRI Consulting’s Process Economic Program” (PEP) [27], the capital investment required for a plant producing ethanol from 2000 metric tons of straw per day would be around 260 million US$. The plant is assumed to produce 190 million liters of ethanol per year, which gives an investment cost of around 1.37 US$ per liter ethanol and 370 US$ per ton raw material. This is about 2.5 times higher than the investment cost of a corn-based ethanol plant with the same feed capacity. On the basis of the ethanol produced, the ratio would increase to above 3 as the yield of ethanol per ton raw material is higher for corn than for lignocellulosic materials.

The highest contribution to the capital cost, 45% of the total, was equip­ment for the production of heat and electricity for the process and for sale to the grid, wastewater treatment and other utilities. This is not a direct cost of the ethanol production equipment, and in our opinion is often underesti­mated in most studies on ethanol production cost. Another cost that differs widely between studies is the cost of raw material. This depends on both dif­ferences in the type of raw material (agricultural residues, forest residues or energy crops) and on the location of the raw material. According to the Road Map for Agricultural Biomass Feedstock Supply in the US presented by the DOE [28] the goal is to reach a feedstock cost of 30 US$ per dry ton. On the basis of this figure the net raw material cost, i. e. after by-product credit, in the PEP study would be about 0.07 US$ per liter of ethanol, which corres­ponds to 20% of the total production cost of 0.34 US$ L-1. This production cost is, however, without any profit. The cost of biomass in Sweden, and other European countries is much higher, exceeding 90 US$ per metric ton of dry matter [29], which results in a net raw material cost of about 60 US$ per met­ric ton. This would have increased the total production cost in the PEP study to about 0.44 US$ L-1. However, according to the PEP study, the main reason for the higher investment cost for biomass-produced ethanol is due to the cost of conditioning and pretreating the biomass to make the cellulose accessible to enzymatic hydrolysis, which was estimated to represent 27% of the total fixed capital.

Eggeman et al. [30] investigated the pretreatment cost in ethanol produc­tion from corn stover for five different pretreatment methods: dilute acid, hot water, ammonia fiber explosion (AFEX), ammonia recycle percolation (ARP) and lime. The pretreatment design was based on experimental data from various research groups [31] and was implemented in the Aspen Plus model for a full-scale bioethanol plant previously developed by NREL [26]. The model was based on a corn stover feed rate of 2000 dry metric tons per day. The process configuration was based on pretreatment, SSF, ethanol re­covery and internal production of heat and electricity from the syrup and solid residue from the process. The process configuration was identical for all processes except for the pretreatment step. The dilute acid pretreatment process resulted in the lowest ethanol production cost, 0.26 US$ L-1 for the base case alternative where oligomers released in the pretreatment and hydro­lysis steps were not considered for ethanol production. The production cost includes depreciation, but no income tax or return on capital, to make it com­parable to the other costs presented in this review. The total investment cost was estimated to be 185.8 million US$, of which the pretreatment step consti­tuted 25 million US$, i. e. 13.5% of the total. The largest investment cost was for steam and power, 41.8 million US$, which represents 22.5% of the total investment cost.

Two-step pretreatment has been suggested to improve the overall sugar yield in several studies [32-34]. The first step is performed at low severity to release hemicellulose sugars, which are then removed, followed by the sec­ond step at more severe conditions to make the cellulose more accessible to enzymatic attack. Wingren et al. [35] compared the ethanol production cost of two-step steam pretreatment of SO2-impregnated spruce with that for one — step pretreatment, based on experimental data for pretreatment at optimal conditions. The production plants considered were designed for a yearly cap­acity of 200 000 tons raw material, and the only difference between the two plants was the pretreatment step. The process was based on SSF of the pre­treated material. The two-step process resulted in a higher ethanol yield (see Table 2) and a lower requirement for enzymes. However, due to the higher energy demand and higher capital cost the estimated ethanol production cost was the same, 0.55 US$ L-1. In the most optimistic scenario, where the material from the first step was dewatered to 50% dry matter (DM) with­out reducing the pressure, and the overall ethanol yield was assumed to be the highest achieved in the experimental work, 77%, the production cost de­creased by 5.6% to 0.52 US$ L-1. This shows the potential of the two-step pretreatment process, which, however, remains to be verified in pilot trials.

Hamelinck et al. [36] investigated the effect of expected future improve­ments in the conversion of biomass to ethanol, with poplar as a model raw material. The paper contains detailed information on the technical and eco­nomic data used in the analysis. Three scenarios were investigated: short­term (5 y), middle-term (10-15 y) and long-term (> 20 y). Improvements were mainly expected to be in enhanced pretreatment and bioconversion steps, changing from SSF to SSCF of hexose and pentose sugars and finally Consolidated BioProcessing (CBP), resulting in higher ethanol yield and re­duced capital costs. This is based on improvements of both enzyme efficiency and fermentation microorganisms. For CBP a completely new microorgan-

ism, not yet known, was assumed. The production capacity is also assumed to increase, from a biomass input (in metric tons per year) of 620 000 to 1550 000 and 3 110 000 for the short, medium and long term, respectively. Also, the cost of the raw material was assumed to fall from about 58 US$ per metric ton for the short term to 48 and 39 US$ for the middle and long term.

The total investment cost for the short-term scenario was about 395 mil­lion US$, i. e. 475 US$ per ton raw material yearly capacity. The ethanol production cost was determined to be 0.51 US$ L-1. Forty-five percent was due to capital cost and about 35% to the net raw material cost, i. e. after credit for co-produced electricity. For the middle-term scenario the total invest­ment cost increased to around 465 million US$, corresponding to 300 US$ per ton raw material yearly capacity. The ethanol production cost decreased to 0.31 US$ L-1, of which about 44% arose from capital cost and about 31% due to net raw material cost. For the long-term scenario the total invest­ment cost increased to around 820 million US$, corresponding to 263 US$ per ton raw material yearly capacity. The ethanol production cost decreased to 0.20 US$ L-1, of which about 50% was due to capital cost and about 42% to the net raw material cost. The long-term ethanol production cost was still considerably higher than that predicted by Lynd et al. [23] who estimated the future ethanol production cost for a plant based on CBP to be between 0.1 and 0.16 US$ L-1. The main reasons for their lower cost are a higher conversion yield and lower capital costs. It must be pointed out that these are scenarios based on future projected improvements, which are very uncertain.

The number of studies on softwood is more limited. Fransson et al. [27] studied the potential for a two-stage dilute acid process using softwood as raw material. The plant is assumed to be co-located with a heat and power plant from which steam is purchased and co-products (solid residue) are sold. The first hydrolysis step is assumed to be run in co-current fashion, whereas the second reactor is working in counter-current mode to maximize sugar yield and reduce sugar degradation. A sulfuric-acid concentration of 5 g/L is used in both steps. The sugar stream is detoxified in an overliming step and then fermented to ethanol. The dilute ethanol is concentrated in a distillation step and the stillage is then evaporated. The plant is designed to process around 300 000 tons of raw material annually. The estimated ethanol production cost was 0.53 US$ L-1.

In another study by Wingren et al. [38], the production of fuel ethanol from spruce using the enzymatic process was investigated. The softwood was steam pretreated after impregnation with SO2. Two configurations, one based on SSF and the other on SHF, were evaluated and compared. The process conditions selected were based mainly on laboratory data and the processes were simulated using Aspen Plus, while the capital costs were estimated using Icarus Process Evaluator. The ethanol production cost was estimated to be 0.69 and 0.76 US$ L-1 for the SSF and SHF cases, respectively, based on a raw material cost of 63 US$ per dry ton. The main reason for SSF being less ex­pensive was due to the capital cost being lower and the overall ethanol yield higher. Improvements in the SSF process, by running SSF at 8% WIS rather than at 5%, and by recycling of process streams, were shown to result in a de­crease in production cost to 0.51 US$ L-1.

3.1

In the first generation of recombinant xylose-utilizing S. cerevisiae strains the activity of the enzyme(s) converting xylose to xylulose has been insuffi­cient to support ethanolic fermentation of xylose [42,43,54] (strain RWB202, Tables 3 and 4). For example, overexpression of the nonoxidative PPP im­proved xylose fermentation only when the XR and XDH activities were enhanced [54,90] (strain TMB3057, Table 1, vs strain TMB3026, Table 3), indicating that when the flux through central metabolism was high, the control of xylose metabolism was in the steps converting xylose to xylu­lose [42] (strain TMB3050, Table 2). Invariably, increased XR and XDH ac­tivities have been observed in mutant S. cerevisiae strains with improved xylose utilization [31,57,91]. Similarly, high activity of Piromyces XI allowed higher xylose fermentation rates than the lower bacterial XI activity [42,92] (strain RWB202-AFX, Table 1; strain TMB3050, Table 2). Also, in recombi­nant arabinose-utilizing S. cerevisiae strains, enhanced levels of the arabinose isomerase significantly improved arabinose fermentation [8].

The fact that not only the cofactor specificities, but also the relative activ­ities of XR and XDH affect xylitol formation suggests that the redox model for xylitol formation [32] may have to be reevaluated. Not only the cofac­tor preferences of the enzymes involved but also, equally importantly, the levels of the XR and XDH activities affect xylitol formation during xylose fermentation [54]. A plain increase in XR and/or XDH activity, allowing an increased flux through the initial pathway, significantly reduces xylitol excretion [53,54,93] (strain TMB3062, Table 1; strain TMB3061, Table 2).

4.2.3

The major requirement for S. cerevisiae as CBP yeast would be sufficient ex­pression and production of extracellular saccharolytic enzymes [1]. In the context of creating such a CBP yeast, the first question researchers would like to answer is, “How much saccharolytic enzyme, particularly cellulase expres­sion, is enough to enable CBP conversion of plant material to ethanol, and is that amount feasible in S. cerevisiae?” The obvious follow-up question is, “How do we accomplish those levels of expression?” Recent analyses [9,44] have approached the first question from a kinetic standpoint, balancing the demand for soluble products of cellulose hydrolysis (glucose) by cells with the production of those products by cellulase systems. Demand is a simple func­tion of the growth rate and the cell yield: ^/YX/S = g glucose/gcells/h, while supply is just the cell-specific cellulase activity: g glucose/g cells/h. These re­lationships can be used to calculate a number of useful quantities, including the percentage of total cell protein that needs to be cellulase to achieve a par­ticular growth rate on a cellulosic substrate. The relative levels of individual cellulase component expression can be calculated based on the known ratios of those components in native systems.

In the last two decades there have been several reports on the expression of cellulases in S. cerevisiae. Table 2 summarizes some of the results found to date. Most reports regarding the expression of cellulases and hemicellulases in yeast employed strong glycolytic (or other constitutively expressed) pro­moters to drive expression of the heterologous gene(s). Although the choice of promoter and leader sequences will undoubtedly have a great influence on expression levels attained, there are not enough data in the literature to suggest any general trends as to what are the best promoter and leader sequences to use when expressing cellulases and hemicellulases. Several re­searchers have sought to produce cellulases in an organism that would not yield interfering activities so as to gain insight into the mechanism of the original cellulolytic enzyme [99], whereas others have sought to enable the yeast to hydrolyze nonnative cellulolytic substrates [43,59,78,102]. Although most of the cellulases that have been successfully produced in S. cerevisiae were of fungal origin, there are reports of successful bacterial cellulase pro­duction [76,82].

Full enzymatic hydrolysis of crystalline cellulose requires three major types of enzymatic activity: (1) endoglucanases (1,4-P-D-glucan 4-glucano — hydrolases; EC 3.2.1.4); (2) exoglucanases, including D-cellodextrinases (1,4-P-D-glucan glucanohydrolases; EC 3.2.1.74) and cellobiohydrolases (1,4-P-D-glucan cellobiohydrolases; EC 3.2.1.91); and (3) P-glucosidases (в-glucoside glucohydrolases; EC 3.2.1.21) (Fig. 2a). Cellobiohydrolase (CBH) enzymes are key components for fungal cellulase systems, and their func­tional secretion is critical for allowing CBP. For example, CBHs make up ~ 80% of the total mass for the T. reesei system, and CBH1 plays a particu­larly important role, making up 60% of the total mass [103]. CBHs have been successfully produced and secreted by S. cerevisiae and were tested for ac­tivity on a variety of substrates ranging from small synthetic molecules to amorphous and crystalline forms of cellulose (Table 1). Some reports have shown decreased specific activity on certain substrates, presumably as a re­sult of hyperglycosylation [47, 48]. However, in a recent study it was shown that the specific activity of a glycosylated heterologous CBH1 did not dif­fer significantly from that of the native enzyme produced by T. reesei [49].

U = micromole substrate released/min, NR = not reported; italics indicate calculation based on assumptions (0.45 g DCW/g glucose, 0.45 g protein/g DCW, 1.3 x 107 cells/mg DCW, 1 OD(600) = 0.57 g DCW/L).

CBH = cellobiohydrolase, EG = endoglucanase, BGL = beta-glucosidase, AC = amorph­ous cellulose, BMCC = bacterial microcrystalline cellulose, BBG = barley beta-glucan, CC = crystalline cellulose, IOSC = insoluble cellooligosaccharides, C2 = cellobiose, C3 = cel — lotriose, C4 = cellotetraose, PNPC = p-nitrophenol cellobioside, PNPL = p-nitrophenol lactoside, MUC = methylumbelliferyl cellobioside, MUL = methylumbelliferyl lactoside, Xglu = 5-bromo-4-chloro-3-indolyl-|3-D-glucopyranoside

Reports of CBH production in yeast have also shown that a relatively low titer of secreted cellulase is found, although the range of reported values is quite large—0.002 to 1.5% of total cell protein. Coupled with the low spe­cific activity of CBHs, CBH expression has been identified as a limiting factor for CBP using yeast [9]. However, in a recent report the amount of CBH1 required to enable growth on crystalline cellulose was determined and was found to be, in terms of total cellular protein, within the capacity of heterol­ogous protein production in S. cerevisiae, i. e., between 1 and 10% of total cell protein [49,104-106].

Fig.2 Illustration of the complexity of cellulose and hemicellulose and the enzymes in — ► volved in their degradation. Cellulose (a) and hemicellulose structures for arabinoxylan (b), galactomannan (c) , and xyloglucan (d) depicting the different side chains present. Hexoses are distinguished from pentoses by the presence of a protruding line from the cyclic hexagon (pyranose ring), depicting the CH2OH group. Hydrolase enzymes and the bonds targeted for cleavage in the four polysaccharide structures are indicated by arrows [100,101]

Fungal and bacterial endoglucanase (EG) production in S. cerevisiae have been by and large more successful than CBH production (Table 2). This is not surprising considering that EG enzymes usually have specific activities 2 to 3 orders of magnitude higher on synthetic and amorphous cellulose substrates, such as phosphoric acid swollen cellulose (PASC) and carboxymethyl cellu­lose (CMC), in comparison to CBHs. It is thus easier to measure the presence of even small amounts of heterologous EG compared to CBHs. Although se­creted heterologous EGs were usually reported to be hyperglycosylated, this did not necessarily influence their specific activity negatively [61]. Sufficiency analysis shows that, assuming that a T. reesei system is reconstructed, even if all of the non-CBH cellulase system components were EG, it would still only need to make up ~ 0.3% of cell protein, well within the range of possibility for a S. cerevisiae secretion system. The successful expression of P-glucosidases in S. cerevisiae at sufficient levels to sustain growth on cellobiose as sole car­bon source at a rate comparable to glucose suggests that BGL expression will not be a limiting step in cellulase system reconstruction [43,44].

A number of studies have expressed multiple cellulase enzymes in at­tempts to recreate a fully cellulolytic, fermentative system [45,59,64,78,102]. Van Rensburg et al. [51] constructed a yeast capable of hydrolyzing numer­ous cellulosic substrates and growing on cellobiose, while Cho et al. [78] showed that decreased loadings of cellulase could be used for SSF experi­ments with their strain expressing a BGL enzyme and an enzyme with dual exo/endocellulase activity. Fujita et al. [59,64] reported coexpression and sur­face display of cellulases in S. cerevisiae, and a recombinant strain displaying the T. reesei endoglucanase II, cellobiohydrolase II, and the Aspergillus ac — uleatus P-glucosidase 1 was built. High cell density suspensions of this strain were able to directly convert PASC to ethanol with a yield of approximately 3 g L-1 from 10 g L-1 within 40 h [59]. Den Haan et al. [102] reported growth on and direct conversion of PASC to ethanol by a S. cerevisiae strain coex­pressing the T. reesei EG1 and the Saccharomycopsis fibuligera BGL1 (Fig. 3). Anaerobic growth (0.03 h-1) up to 0.27 gL-1 dry cell weight was observed with this strain on medium containing 10gL-1 PASC as sole carbohydrate source with concomitant ethanol production of up to 1.0 gL-1. As an ex- ocellulase activity such as CBH is required for the successful hydrolysis of crystalline cellulose, it is postulated that the addition of successful, high-level expression of CBH to this strain will enable CBP of crystalline cellulose to ethanol.

5

A number of components in lignocellulosic hydrolysates can inhibit the growth and ethanol production of bacteria and yeasts, and acetic acid has been identified as a major potential inhibitor of Z. mobilis in such acid — produced hydrolysates [61-65]. Lawford and Rousseau [32] examined the role of glucose feeding as a means of improving fermentation perform­ance in acetate-containing media. Another approach to solving this problem has been to use a hydrolysate-fed chemostat to produce adapted or mutant strains [33,34]. Following chemical mutagenesis, Joachimsthal et al. [66] iso­lated a mutant strain, designated ZM4/AcR with a higher acetate resistance than the parent strain. This strain was then transformed by Jeon et al. [67] to the mutant recombinant ZM4/AcR (pZB5). Compared to ZM4 (pZB5), this strain showed enhanced kinetics in batch culture in the presence of 12 gL-1 sodium acetate (8.8 g L-1 acetic acid) at pH = 5.0 in batch culture on 40 g L-1 glucose, 40 g L-1 xylose medium. In continuous culture there was evidence of increased maintenance energy requirements/uncoupling of metabolism in the presence of acetate.

In more recent studies Saez-Miranda et al. [68] have determined ATP levels for growth on glucose/xylose media in the presence of different concentra­tions of acetic acid. From their results they have found that ATP production and accumulation rates are most sensitive to acetic acid at lower pH values— a result consistent with the earlier NMR studies by Kim et al. [57] which demonstrated increasing de-energization of the cells as the inhibitory effects of acetic acid increased. The greater toxicity of acetic acid at lower pH is related to its pKa value as only unprotonated acid can be transported into the cells.

The effects of a range of inhibitory compounds at levels reported previ­ously for a pre-treated hardwood hydrolysate [65] on specific rates of xylose utilization and ethanol production for ZM4 (pZB5) have been analyzed by Kim et al. [57]. From the results, sodium acetate was found to have the greatest inhibitory effect at the concentration tested (10.9 gL-1 at pH = 6.0), followed by vanillin (0.04 gL-1), syringaldehyde (0.13 gL-1) hydroxymethyl- furfural (0.9gL-1) and furfural (0.3 gL-1). Vanillic acid (0.08 gL-1) did not show any inhibitory effects at this experimental concentration. At the levels tested, these inhibitory compounds did not affect ethanol yields on xylose. Volumetric rates of xylose utilization and ethanol produc­tion were reduced by up to 20% by addition of the individual inhibitory components.

2.7

The ability of biofuels to contribute positively to the environmental and eco­nomic performance of a country, and to improve energy security in the long term, makes the nascent industry a tool that policymakers can employ to meet national priorities in these areas. A review of the priorities that gov­ernments are pursuing when designing biofuel-related policy illustrates some issues that the emerging bioethanol industry might consider. These issues may have particular relevance to the commercialization of the lignocellulosic — based component of the industry.

In the USA, the primary political drivers that support research and de­velopment into bioethanol for fuel are related to the economy and to energy security. Two agencies have become the primary implementing bodies for US policies related to bioethanol. The Department of Agriculture (USDA) has a mandate to increase rural employment, diversify agricultural economies, and stimulate rural development by harnessing crops and crop residues and identifying new uses for this material. The Department of Energy (DOE) has a mandate to diversify the energy supply, expand the availability of renew­able energy sources, and develop new technologies to exploit renewables in all forms.

From an economic perspective, bioethanol policy in the USA has been highly successful. Since 1976, bioethanol production capacity has grown sig­nificantly. Almost a decade ago, the US industry passed 5 billion L in an­nual production and was credited with the creation of an estimated 200 000 new jobs and US$ 500 million in annual tax receipts [4]. Today, there are 94 bioethanol plants in the USA, producing about 18.5 billion L year-1, with an additional 16 plants and 2.5 billion L of capacity under construc­tion [13]. Urbanchuk [85] estimated that expansion to this level would require US $ 5.3 billion investment in new facilities and would increase demand for crops by 1.6 billion bushels per year. In that report, the author anticipates that a bioethanol industry of this size could reduce the US trade deficit by US$ 34 billion year-1, create 214 000 new jobs within the USA, and gener­ate US$ 51.7 billion in new US household income. It should be noted that the success of the US industry is in part due to the presence of import tar­iffs on bioethanol (duty of 2.5% market value, plus US$ 0.143 L-1) [23]. While some regions (notably the Caribbean) may export duty-free bioethanol within a quota, the maximum amount of duty-free bioethanol entering the USA is currently 7% per year. This means that it is not cost-effective to import large supplies of bioethanol from other producers, such as Brazil.

From a security perspective, bioethanol policy has been less success­ful. American demand for petroleum continues to outpace domestic sup­ply, resulting in growing petroleum imports, anticipated to be nearly 70% by 2020 [18]. Only about 3% of US energy requirements are supplied by biomass [56], and only about 2.6% of American total transportation fuel consumption is derived from biofuels [18]. Five individual US states (South Dakota, Nebraska, Minnesota, Iowa, and Illinois) now produce enough bioethanol to provide an E10 option to their entire local population. From the perspective of energy security, the USA could benefit from continued ex­pansion of the bioethanol industry and increased utilization of the industry’s potential.

Globally, Germany has the best capacity to substitute biofuels for fossil — based fuels, with current capacity of about 3.75% total demand, followed by the USA (2.6%), Sweden (2.2%), France (1.2%), Austria (1.1%), and Spain (0.44%) [9,35,36,43,48,60].

The issue of climate change has become a major, global concern, but the sectors most closely linked to bioethanol production — including energy pro­ducers, farmers, and foresters — will feel the impact of this issue more closely. Climate change is the driver behind many new policies that influence the ac­tions taken by these sectors. Perhaps the best-known of these is the Kyoto Protocol, which has been ratified by Russia, by the members of the EU, and by Canada in North America. The Clean Skies Initiative in the USA is another example of these policies. Because the use of bioethanol has the potential to significantly reduce net greenhouse gas emissions compared to petroleum products, an expansion of bioethanol production may become a significant part of national climate change strategies. It must be noted, however, that sig­nificant amounts of bioethanol must be substituted for petroleum products in order for these reductions to make a significant impact on total greenhouse gas emissions.

6

Conclusions

Successful policy options to support biofuel production may take a number of forms, including targets and mandates, exemption of biofuels from national excise taxation schemes, direct government funding of capital projects to in­crease capacity or upgrade distribution networks, or consumption mandates for government or corporate vehicle fleets. As discussed in this review, these policies can be differentiated by their relative emphasis on government, in­dustry, or consumer actions. In most biofuel-producing countries examined here, a number of policies have been enacted in order to develop industrial capacity and encourage consumption. It is very difficult to measure the indi­vidual success of these policies because of the synergistic effects that multiple policies may have.

In the USA, an analysis of state-level excise tax exemptions shows no correlation with bioethanol industry capacity, which suggests that these ex­emptions are not a crucial factor in the creation of industrial facilities. Direct funding and support was found to play a much more positive role in the creation of production capacity. It was noted that strong funding for es­tablishment of facilities, including all aspects of research, development, and deployment, was present in each of the states where significant bioethanol production was present. In a comparison of production capacity between 2003 and 2005, it was observed that the correlation between direct funding opportunities and bioethanol production capacity has dropped somewhat. This indicates that other factors, including feedstock supply, the presence or absence of interested industrial players, and other market forces play a signifi­cant role in the establishment of the industry.

In advising governments on the creation of bioethanol-friendly policy, the US experience offers some valuable lessons to consider. The US goals behind policies supporting the bioethanol industry are dominated by (1) economic and social issues, and (2) security-based concerns. Of these priorities, the bioethanol industry has been more successful in meeting social criteria such as rural employment. The starch-based segment of the bioethanol industry has enjoyed particular success in the USA, particularly in Minnesota, Illi­nois, and Iowa. In the past, these jurisdictions have utilized a number of schemes, including direct payments, grants, corporate tax breaks, and ex­cise tax exemptions, as incentives to lure the industry and build bioethanol capacity.

The ability of the industry to increase energy security in the USA, on the other hand, has been limited by the relatively small capacity of their production facilities at the current time. This should serve as a cautionary measure for governments in both Canada and the EU, who have invested biofuel-related policy with more emphasis on the environment and on energy security than they have upon social or economic concerns. Improved en­ergy security through biofuel production can only be achieved when enough capacity is brought on-line. Thus, security-related policy geared to the short­term cannot succeed to any great extent. Policymakers must realize that, in the immediate future, the goals of most successful policies will be related to the economy, and perhaps to the environment. The implication here is that security-related policy, such as mandated renewable fuel use, is likely to take the form of long-term programs that have very little immediate reward.

One important finding was that a balance between research funding and funding for the creation of facilities might be more conducive to support­ing the industry. It was noted that the USA has devoted a significant amount of funds to research as well as to supporting facility creation. A commit­ment to advancing the technology and improving efficiencies may serve to increase the industry’s comfort level in committing resources to this sec­tor. The US example may have important lessons for other countries, where an effective balance between research and commercialization has not been reached. For instance, the total French commitment to biofuels in 2002 was just under US$ 200 million, of which about US$ 180 million is devoted to investment subsidies for biofuels, and a further US $ 11 million was put to­wards wood energy programs. Only about US $ 9 million was earmarked for research and development into renewables, including biofuels research [32]. Although the incentives that the French government offer are dramatic, the research focus of this country has been in other areas, notably nuclear power. This may in part explain the relatively low level of bioethanol production in France, which is currently at about 140 million L year-1 (or 629 million L when bio-ETBE production is considered) [35]. In Spain, the total investment is much lower at approximately US $ 30 million per year, but over half of this amount (US $ 17 million) is available for research and development into var­ious renewables, while the other half may be used for commercial facilities or demonstration plants [32]. Perhaps because of this, Spanish production of bioethanol is at about 521 million Lyear-1 [43]. The balance between research and production incentives that is present in both Spain and the USA, and the resultant human capital, may in part account for the success that these nations have had in nurturing the bioethanol industry.

The experiences gained in developing bioethanol capacity, using both sugar — and starch-based processes, contain many lessons for other biofuels, including biodiesel and the lignocellulose-based bioethanol industry. These fuels can be seen as a response to a variety of domestic issues, including the need to diversify local economies, increased concerns over environmen­tal damage associated with fossil fuel use, and a growing security rationale for a shift to domestic fuel sources. The emerging industry, including the lignocellulosic-based sector, may in turn find opportunities for strategic link­ages and partnerships that capitalize upon these political issues.

Our findings indicate that successful policy interventions can take many forms, but that success measured as biofuel production capacity is equally dependent upon external factors, which include feedstock availability, an ac­tive industry, and competitive energy prices. It is important that policies be crafted that reflect “realistic” use scenarios for bioethanol and other biofuels over future time-frames.

Acknowledgements The authors would like to thank the International Energy Agency (IEA) Bioenergy Task 39 for providing some of the funding required to support this work. The authors also recognize the assistance of Jack Saddler, John Neeft, and other colleagues within Task 39, as well as the Forest Products Biotechnology Group at the University of British Columbia.

[1] The unit bbl is an abbreviation for barrel, a common unit of measurement for petroleum, equiva­lent to 42 US gallons or approximately 159 L.

In addition to tolerance and robustness, strain stability is a prerequisite when designing yeast strains for industrial use. Strains carrying mul­ticopy plasmids are generally not applicable in industry due to their instability [123,145]. Multicopy plasmids require auxotrophic or antibiotic resistance markers to be retained in the cell, both of which are not ap­plicable in industrial media containing complex nutrients and being used in large volumes. Thus, chromosomal integration is necessary for any genes to be introduced in industrially applied yeast strains. Ideally this re­quires sufficient specific activity of the introduced heterologous enzymes, so that single-copy integration supplies enough activity for metabolic func­tion. Multiple chromosomal integration has also been utilized to gener­ate stable pentose-fermenting strains with high activity of the enzymes introduced [6-8].

Metabolic engineering strategies applied on industrial strains have been limited to the introduction of the initial xylose and arabinose utilization pathways [4,5,8,101]. Only the XR-XDH pathway has been developed in in­dustrial S. cerevisiae strains [4,5,101] (strains A4 and A6, Table 1; strain F, Tables 1 and 4; strain TMB3400, Tables 1, 3 and 4; strain 1400(pLNH32), Table 2). No chromosomally integrated XI constructs have been reported. XI expression in S. cerevisiae seems to require a multicopy expression system to provide sufficient enzyme activity for xylose growth and fermentation [43]. Due to the difficulty of applying complex metabolic engineering strategies in industrial strains, procedures for random strain improvement have been relied upon to improve xylose utilization.

5.3

The same PET operon used in engineering of KO11 has also been used to construct a series of ethanologenic K12-derivatives, designated FBR for the Fermentation Biochemistry Research Unit. These strains were engineered with the goal of maximizing strain stability [43,44]. The most recent strain in this line, FBR5, produced ethanol from a variety of substrates at 86-92% of the theoretical yield [44]. Long-term stability of this strain was demonstrated by the maintenance of ethanol yields over 26 days of continuous culture on glucose or xylose [45]. However, the final ethanol concentration and yield from FBR5 in LB xylose are lower than LY168 in minimal medium (Table 2); additionally, these strains have the disadvantage of rich media dependence and contain plasmids.

2.4