If ethanol production in Brazil exemplified the extrapolation of a mature technol­ogy for sugar-based fermentation and subsequent distillation, the development of the second major ethanol fuel market — from corn in the United States — adopted a different approach to alcohol production, adapting and developing that employing starchy seeds in the production of malt and grain spirits (bourbon, rye, whiskey, whisky, etc.). The biological difference from sugar-based ethanol fermentations lies in the carbon substrate, that is, starch glucan polymers (figure 1.13). Historically, seeds and grains have been partially germinated by brewers to generate the enzymes capable of depolymerizing “storage” polysaccharides. With whisky, for example, barley (Hordeum vulgare L.) seeds are germinated and specialized cells in the seed produce hydrolytic enzymes for the degradation of polysaccharides, cell walls, and proteins; the “malted” barley can be used as a source of enzyme activities to break down the components of starch in cooked cereals (e. g., maize [Zea mays L.]) sol­ubilized in sequential hot-water extractions (which are combined before the yeast cells are added) but not sterilized so as to maintain the enzyme activities into the fermentation stage.3435 Starch is usually a mixture of linear (amylose) and branched (amylopectin) polyglucans. For starch hydrolysis, the key enzyme is a-amylase, active on a-1,4 but not a-1,6 linkages (in amylopectin); consequently, amylose is bro­ken down to maltose and maltotriose and (on prolonged incubation) to free glucose and maltose, but amylopectin is only reduced to a mixture of maltose, glucose, and

oligosaccharides containing a-1,6-linked glucose residues, thus limiting the amount of fermentable sugars liberated (figure 1.14). Cereal-based ethanol production plants use the same biochemical operations but replace malted grains with a-amylase and other polysaccharide-degrading enzymes added as purified products.

For much of the twentieth century, ethanol production as a feedstock in the formation of a large number of chemical intermediates and products was dominated in the United States by synthetic routes from ethylene as a product of the petrochemical industry, reaching 8.8 x 105 tonnes/year in 1970.36 The oil price shocks of the early 1970s certainly focused attention on ethanol as an “extender” to gasoline, but a mix of legislation and economic initiatives starting in the 1970s was required to engender a large-scale bioprocessing industry; in particular, three federal environmental regulations were important:3738 [6]

Although ethanol was always a good oxygenate candidate for gasoline, the compound first approved by the Environmental Protection Agency was methyl tertiary butyl ether (MTBE), a petrochemical industry product. Use of MTBE increased until 1999, but reports then appeared of environmental pollution incidents caused by MTBE spillage; state bans on MTBE came into force during 2002,39 and its consumption began to decline (figure 1.15). In the Midwest, ethanol was by then established as a corn-derived, value-added product; when the tide turned against MTBE use, ethanol production increased rapidly after showing little sustained growth for most of the 1990s (figure 1.16). California, New York, and Connecticut switched from MTBE to ethanol in 2004; after 2006, with many refiners discon­tinuing MTBE use, U. S. ethanol demand was expected to expand considerably.40 In the seven years after January 1999, the number of ethanol refineries in the United States nearly doubled, and production capacity increased by 2.5-fold (figure 1.17). In 2005, the United States became the largest ethanol producer nation; Brazil and the United States accounted for 70% of global production, and apart from China, India,

France, and Russia, no other nation accounted for more than 1% of the total etha­nol produced. To further underline the perceived contribution of renewable fuels to national energy use, the 2005 Energy Policy Act created a Renewable Fuels Standard that envisioned renewable fuel use increasing from 4 billion gallons/year in 2006 to

7.5 billion gallons/year in 2012. This implies a further expansion of ethanol produc­tion because of the dominant position of E85 (85% ethanol, 15% gasoline) vehicles in the AFV and hybrid-fuel marketplace (figure 1.18).

Fuel ethanol production in the United States has been almost exclusively from corn, although sorghum (Sorghum bicolor L.), barley, wheat, cheese whey, and brewery waste have made small contributions. A detailed study of sugar sources for ethanol production concluded that only sugarcane molasses offered competitive feedstock and processing costs to established corn-based technologies (figure 1.19), although annual capital cost investments could be comparable for corn, sugarcane, and sugarbeet molasses and juice as rival feedstocks.41 Corn ethanol production developed from wet milling of corn; data compiled in the mid-1990s indicates that more than 70% of the large ethanol facilities then used wet milling.38 Wet milling, schematized in figure 1.20, produces four important liquid or solid by-products:

• Corn steep liquor (a lactic acid bacterial fermentation product, starting from ca. 5% of the total dry weight of the grain extracted with warm water, with uses in the fermentation industry as a nitrogen source)42

• Corn oil (with industrial and domestic markets)

• Corn gluten feed (a low-value animal feed)

• Corn gluten meal (a higher-value, high-protein animal feed)

Together with the possibility of collecting CO2 from the fermentation step as a saleable commodity, this multiplicity of products gave wet milling flexibility in times of variable input and output prices, although requiring a higher initial capital invest — ment.38 Other sources of flexibility and variation in the wet milling procedure arise at the starch processing stage; while a-amylase is used to liquefy the starch, saccharifi­cation (using glucoamylase) can be differently controlled, at one extreme producing

— Electric

Hydrogen

Liquefied natural gas Liquefied petroleum gas Diesel-electric hybrid

FIGURE 1.18 Alternative — and hybrid-fuel vehicles. (Data from U. S. Department of Energy, Energy Information Administration.)

a high-glucose, low-solids substrate for fermentation and at the other producing a low glucose concentration but which is continually replenished during the fermentation by the ongoing activity of the glucoamylase in the broth.

In contrast to wet milling, dry milling produces only CO2 and distillers dried grains with solubles (DDGS) as by-products but has become the favored approach for corn ethanol production because of lower start-up costs.43 Dry milling should conserve more of the nutrients for yeast growth in the fermentation step — in particular, nitrogenous inputs (free amino acids, peptides, and protein), inorganic

and organic phosphates, and some other inorganic ions (potassium, sodium, mag­nesium, etc.) — but this has little, if any, impact on overall process economics (table 1.2). A detailed account of a dry milling process was published by Alltech in 2004.44 The scheme in figure 1.21 is a simplified derivative of the information provided then as a representative example of the complete bioprocess for ethanol and DDGS.

Unlike Brazilian sucrose-based ethanol, corn-derived ethanol has been technology-driven, especially in the field of enzymes and improved yeast strains with high ethanol tolerance and may be (or become) capable of yielding up to 23% by volume of ethanol in batch fermentations within 60 hours.44 45 Typical commercially available enzymes used liberate the sugars present in starches. Their properties are summarized in table 1.3. Innovations in biocatalysts and fermenta­tion engineering for corn ethanol facilities are covered at greater length in chapter 3. The availability of enzyme preparations with increasingly high activities for starch degradation to maltooligosaccharides and glucose has been complemented by the use of proteases that can degrade corn kernel proteins to liberate amino acids and peptides to accelerate the early growth of yeast cells in the fermentor; protein digestion also aids the access of amylases to difficult-to-digest starch resi­dues, thus enhancing overall process efficiency and starch to ethanol conversion. table 1.4 contains indicative patents and patent applications awarded or filed since 2003 for corn ethanol technologies.

As the multiplicity of U. S. corn ethanol producers has increased, the relative contributions of large and small facilities have shifted: in 1996, Archer Daniels Midland accounted for more than 70% of the total ethanol production, but by late

2006, this had fallen to just 21%.3846 Although the largest 4 producers still account for 42%, 8 smaller companies each claim 1 to 2% of the total capacity (figure 1.22). The mix of producers includes local initiatives and farmer-owned facilities, and production is heavily concentrated in the Midwest (to minimize transportation costs for raw materials) but with existing and planned expansion in states from Georgia to Oregon. In September 2006, ethanol production capacity in the United States amounted to 5 billion gallons/year, with a further 3 billion gallons/year under construction.46

Presently, ethanol blends commercially available are the 10% (E10) and 85% (E85) versions. The 2004 Volumetric Ethanol Excise Tax Credit made E85 eligible for a 51 cent/gallon tax break; various states (including Pennsylvania, Maine, Min­nesota, and Kansas) levy lower taxes on E85 to compensate for the lower mileage with this fuel. In Hawaii, the tax rate positively discriminates in favor of E85.47 The 2005 Energy Policy Act established tax credits for the installation of a clean-fuel infrastructure, and state income tax credits for installing E85 fueling equipment have been introduced. FFVs capable of using standard gasoline or E85 began to appear in 1995-1998 (Ford), and since then, Daimler Chrysler, General Motors, Isuzu, Lincoln, Mazda, Mercedes Benz, Mercury, and Nissan have introduced

models as FFVs.47 Usage of ethanol blends is highest in California — 46% of total U. S. consumption.46

Outside North America, construction of the first bioethanol facility in Europe to utilize corn as the feedstock commenced in May 2006 in France; AB Bioenergy France aims to begin production in 2007. The parent company Abengoa Bioenergy (www. abengoa. com) operates three facilities in Spain, producing 5,550 million liters of ethanol a year from wheat and barley grain. A plant in Norrkoping, Sweden, began producing 50 million liters of ethanol annually from wheat in 2001; the product is blended with conventional imported gasoline at up to 5% by volume. These and other representative bioethanol facilities in Europe and Asia are listed in table 1.5. Similar industrial plants, to use a variety of agricultural feedstocks, are presently planned or under construction in Turkey, Bulgaria, Romania, El Salvador, Colombia, and elsewhere.

By 1983, in experimental laboratory programs, selected Bacillus strains had achieved ethanol formation to 20 g/l (from 50 g/l sucrose as the carbon substrate) at 60°C, with ethanol as the major fermentation product; acetic and formic acids remained serious by-products, however, and evidence from laboratory studies suggested that ethanol accumulation followed (and depended on) the formation of those growth- inhibiting acids.46 The ability to run ethanol fermentations at 70-80°C with ther­mophilic microbes remains both a fascination and a conscious attempt to accelerate bioprocesses, despite the low ethanol tolerance and poor hexose-converting abilities of anaerobic thermophilic bacteria. In 2004, exploratory work at the Technical Uni­versity of Denmark tested isolates from novel sources (hot springs, paper pulp mills, and brewery wastewater), using three main criteria for suitable organisms:236

1. The ability to ferment D-xylose to ethanol

2. High viability and ethanol productivity with pretreated wheat straw

3. Tolerance to high sugar concentrations

Five good (but unidentified) strains were identified by this screening program, all from hot springs in Iceland,[29] the best isolate could grow in xylose solutions of up to 60 g/l.

Thermophilic and mesophilic clostridia also have their advocates, especially with reference to the direct fermentation of cellulosic polymers by the cellulosome multien­zyme complexes, as discussed in chapter 2, section 2.4.1). Bypassing the cellulosome is possible if cellulose degradation products (rather than polymeric celluloses) are used as carbon sources — this equates to using bacteria with cellulase-treated materi­als, including agricultural residues and paper recyclates. Laboratory studies with C. cellulolyticum tested cellobiose in this fashion but with chemostat culture so as to more closely control growth rates and metabolism.237 The results demonstrated that a more efficient partitioning of carbon flow to ethanol was possible than with cellulose as the substrate but that the fermentation remained complex, with acids being the major products (figure 3.11). Nevertheless, clostridia are open to metabolic engineering to reduce the waste of carbohydrates as acids and polymeric products or as vehicles for “consolidated bioprocessing” where cellulase production, cellulose hydrolysis, and fer­mentation all occur in one step — this is covered in chapter 4 (section 4.5).

Historically, the use of dilute acid hydrolysis predated enzymic hydrolysis as a methodology used for cellulose processing beyond the laboratory stage of develop­ment (table 2.3). In the former Soviet Union, large-scale processes for single-cell protein[13] as animal feeds using acid-hydrolyzed woody materials were developed.49 More recently, highly engineered reactors have been devised and investigated for the efficient hydrolysis of lignocellulosic biomass with dilute sulfuric acid, including50

• Batch reactors operating at temperatures up to 220°C

• Plug-flow reactors, that is, flow-through reactors in which liquid and solid phases travel at the same velocity and reduce the residence time at high temperature (up to 230°C)

• Percolation reactors, including two-stage reverse-flow and countercur­rent geometries

Hydrolysis efficiencies can now rival those in enzymic (cellulase) hydrolyses with the advantages that none of the feedstock need be dedicated to support enzyme production and very low acid concentrations used at high temperatures may be economically competitive with enzyme-based approaches.

Two-stage processes employ mild hydrolysis conditions (e. g., 0.7% sulfuric acid, 190°C) to recover pentose sugars efficiently, whereas the more acid-resistant cellulose requires a second stage at higher temperature (e. g., 215°C); sugars are recovered from both stages for subsequent fermentation steps.51 Concentrated (30-70%) sulfuric acid hydrolysis can be performed at moderate temperature (40°C) and result in more than 90% recovery of glucose but the procedure is lengthy (2-6 hr) and requires efficient recovery of the acid posttreatment for economic feasibility.52

The major drawback remains that of the degradation of hexoses and pentoses to growth-inhibitory products: hydroxymethylfurfural (HMF) from glucose, furfural from xylose, together with acetic acid (figure 2.6). HMF is also known to break down in the presence of water to produce formic acid and other inhibitors of ethanol-producing organisms.53 In addition, all thermochemical methods of pretreatments suffer, to varying extents, from this problem; even total inhibition of ethanol production in a fermentation step subsequent to biomass presteaming has been observed (figure 2.7).

FIGURE 2.6 Chemical degradation of hemicellulose, xylose, and glucose during acid — catalyzed hydrolysis.

Two contrasting views have become apparent for dealing with this: either the growth — inhibitory aldehydes are removed by adsorption or they can be considered to be an additional coproduct stream capable of purification and resale.5455

Bioprocess engineering indicates that simply feeding a cellulosic hydrolysate with high concentrations of furfurals and acetic acid to yeast cells, rather than present­ing the full “load” of inhibitors in the batched medium, conditions the microorgan­ism to detoxify and/or metabolize the inhibitory products of sugar degradation.5657 A more proactive strategy is to remove the inhibitors by microbiological means, and a U. S. patent details a fungus (Coniochaeta lignaria) that can metabolize and detoxify furfural and HMF in agricultural biomass hydrolysates before their sac­charification and subsequent use for bioethanol production.58

In addition to isolating grains for processing, cereal-milling plants also generate fiber — rich fractions as a coproduct stream. In the wet milling of corn, the fiber fraction has traditionally been added into a feed product (figure 1.20). The University of Illinois developed a modified dry milling procedure to recover fiber fractions before fermenta­tion: this quick fiber contained 65% by weight of total carbohydrate and 32% by weight of glucans, and dilute acid pretreatment was used before fermentation of the substrate to ethanol by either Escherichia coli or S. cervisiae.102 Destarched, cellulose-rich, and arabinoxylan-rich fractions of the corn fiber support the growth of strains of Hypocrea jecorina and their secretion of hydrolases for plant polysaccharides; these enzymes act synergistically with commercial cellulases on corn fiber hydrolysate and represent a valuable source of on-site enzymes for corn fiber product utilization.103 Similarly, large quantities of wheat bran are produced worldwide as a coproduct of wheat milling; residual starch in the bran material can be hydrolyzed to glucose and oligoglucans by amylolytic enzymes, and acid hydrolysis pretreatment followed by cellulase treatment gives a sugar (pentose and hexose) yield of 80% of the theoretical: 135, 228, and 167 g/kg of starch-free bran for arabinose, xylose, and glucose, respectively.104

Rice husks are approximately 36% by weight cellulose and 12% hemicellulose; as such, this agricultural by-product could be a major low-cost feedstock for ethanol production; 60% of the total sugars could be released by acid hydrolysis and treatment with a mixture of enzymes ф-glucosidase, xylanase, and esterase) with no formation of furfuraldehyde sugar degradation products.105 Recombinant E. coli could ferment the released sugars to ethanol; high-pH treatment of the hydrolysate reduced the time required for maximal production of ethanol substantially, from 64 to 39 hours. In a study from India, rice straw was pretreated with and without exogenous acid, and the released hemicellulose sugars fermented by a strain of C. shehatae; ethanol production was also demonstrated by yeast cells immobilized in calcium alginate beads — an example of an advanced fermentation technology discussed in more detail in the next section.106

Fast-growing willow trees are a major focus of research interest as a bioenergy crop in Scandinavia; high sugar recoveries were achieved from lignocellulosic mate­rial by steaming sulfuric acid-impregnated material for a brief period (4-8 minutes) at 200°C, and then digesting the cellulose enzymically, liberating glucose with 92% efficiency and xylose with 86% efficiency. The pretreated substrate could also be used for SSF with a S. cerevisiae strain.107

Many “exotic” plant materials have been included in surveys of potential bio­mass and bioenergy sources; example of these are considered in chapter 5, section 5.5.2, when sustainability issues are covered at the interfaces among agronomy, the cultivation of bioenergy crops, land use, and food production. As a lignocellulosic, straw from the grass species Paja brava, a Bolivian high-plains resident species, can be considered here. Steamed, acid-impregnated material gave hemicellulose frac­tions at 190°C that could be fermented by three pentose-utilizing yeasts, P. stipitis, C. shehatae, and Pachysolen tannophilus, while a higher temperature (230°C) was necessary for cellulose hydrolysis.108 Much more widely available worldwide is the mixed solid waste of lumber, paper, tree pruning, and others; this is a highly digest­ible resource for cellulase, the sugars being readily fermented by S. cerevisiae and the residual solids potentially usable for combustion in heat and power generation.109

Bacteria are traditionally unwelcome to wine producers and merchants because they are causative spoiling agents; for fuel ethanol production, they are frequent contami­nants in nonsterile mashes where they produce lactic and acetic acids, which, in high concentration, inhibit growth and ethanol production by yeasts.3435 In a pilot plant constructed and operated to demonstrate ethanol production from corn fiber-derived sugars, for example, Lactobacilli were contaminants that could utilize arabinose, accumulating acids that impaired the performance of the ethanologenic yeast; the unwanted bacteria could be controlled with expensive antibiotics, but this experience shows the importance of constructing ethanologens to consume all the major carbon

sources in lignocellulosic substrates so as to maximize the competitive advantage of being the dominant microbial life form at the outset of the fementation.36

Bacteria are much less widely known as ethanol producers than are yeasts but Escherichia, Klebsiella, Erwinia, and Zymomonas species have all received serious and detailed consideration for industrial use and have all been the hosts for recom­binant DNA technologies within the last 25 years (table 3.3).37-43 With time, and perhaps partly as a result of the renewed interest in their fermentative capabilities, some bacteria considered to be strictly aerobic have been reassessed; for example, the common and much-studied soil bacterium Bacillus subtilis changed profoundly in its acknowledged ability to live anaerobically between the 1993 and 2002 editions of the American Society for Microbiology’s monograph on the species and its rela­tives; B. subtilis can indeed ferment glucose to ethanol, 2,3-butanediol, and lactic acid, and its sequenced genome contains two ADH genes.44 The ability of bacteria to grow at much higher temperatures than is possible with most yeast ethanologens led to proposals early in the history of the application of modern technology to fuel etha­nol production that being able to run high-yielding alcohol fermentations at 70°C or above (to accelerate the process and reduce the economic cost of ethanol recovery) could have far-reaching industrial implications.45,46

Bacteria can mostly accept pentose sugars and a variety of other carbon sub­strates as inputs for ethanol production (table 3.3). Unusually, Zymomonas mobilis can only use glucose, fructose, and sucrose but can be easily engineered to utilize pentoses by gene transfer from other organisms.47 This lack of pentose use by the

wild-type organism probably restricted its early commercialization because other­wise Z. mobilis has extremely desirable features as an ethanologen:

• It is a GRAS organism.

• It accumulates ethanol in high concentration as the major fermentation product with a 5-10% higher ethanol yield per unit of glucose used and with a 2.5-fold higher specific productivity than S. cerevisiae.48

• The major pathway for glucose is the Entner-Doudoroff pathway (figure 3.4); the inferior bioenergetics of this pathway in comparison with glycolysis means that more glucose is channeled to ethanol production than to growth, and the enzymes required comprise up to 50% of the total cellular protein.48

• No Pasteur effect on glucose consumption rate is detectable, although inter­actions between energy and growth are important.49

Escherichia coli and other bacteria are, as discussed in chapter 2 (section 2.2), prone to incompletely metabolizing glucose and accumulating large amounts of carboxylic acids, notably acetic acid; with some authors, this has been included under the heading of the “Crabtree effect.”50,51 For E. coli as a vehicle for the production of recombinant proteins, acetate accumulation is an acknowledged inhibitory factor; in ethanol production, it is simply a metabolic waste of glucose carbon. Other than this (avoidable) diversion of resources, enteric and other simple bacteria are easily genetically manipulated, grow well in both complex and defined media, can use a wide variety of nitrogen sources for growth, and have been the subjects of decades of experience and expertise for industrial-scale fermentations — Z. mobilis also was developed for ethanol production more than 20 years ago, including its pilot-scale use in a high-productivity continuous process using hollow fiber membranes for cell retention and recycling.52

Sucrose functions in plants as a highly water-soluble and readily transported product of carbon fixation in leaves, although it can also accumulate in storage organs (e. g., sugarbeets). Starch is mainly a storage polymer in, for example, cereal grains. Cellulose, on the other hand, is essentially a structural polymer in plants (figure 1.23), highly insoluble, organized into crystalline macroscopic fibers, mixed with other polysaccharides (e. g., hemicelluloses), and protected from enzymic attack in native woods by the physical presence of lignin (figure 1.24). Lignins are polyphenolic polymers generated by enzyme-catalyzed free radi­cal reactions from phenylpropanoid alcohols. Unlike nucleic acids and proteins, they have no informational content but are neither inert to enzyme-catalyzed degradation nor incapable of being converted (by hydrogenolysis or oxidative breakdown) to useful chemical intermediates, for example, in the manufacture of synthetic resins, perfume, dyes, and pharmaceuticals.48 As wood chemicals,

Patent Applications and Patents Awarded for Corn Ethanol Technologies

extracted celluloses are, however, far more widely known, especially in the paper industry and also (as acetylated, nitrated, and other derivatives) find applications as varied as components of explosives, cigarette filters, cosmetics, and medical products such as gauze and bandages. Among tree species, hardwoods and soft­woods differ in their compositions, hardwoods having (perhaps, paradoxically) less lignin (figure 1.25). Detailed data for more than 120 tree species lists cel­lulose contents as high as 57% (and as low as 38%), with lignin in the 17 to 37% range (by weight).48 This plasticity of biomass chemical composition suggests that plant breeding programs and genetic technologies can accelerate the evolution of “bioenergy” plants as novel cultivars.

Hemicelluloses are a diverse group of polysaccharides, with different plant species elaborating structures with two or three types of sugar (sometimes further modified by O-methylation or O-acetylation) and a sugar acid; the major sugar monomers are the pentoses xylose and arabinose and the hexoses glucose, galac­tose, and mannose (figure 1.23). Most plant species contain xylans (1,4-linked poly­mers of xylose); in addition, hardwood and softwood trees contain copolymers of glucose and mannose (glucomannans) — larchwoods are unusual in having a core polymer of galactose. Table 1.6 summarizes compositional data based on sugar

FIGURE 1.23 Chemical structures of cellulose and sugar components of hemicelluloses.

type, taken from analyses presented by the National Renewable Energy Laboratory (Golden, Colorado) for a range of tree species, paper products for recycling, cere­als, and grasses.49

In principle, a bioprocess for producing ethanol from a lignocellulosic substrate could be modeled on those developed for cornstarch (figures 1.20 and 1.21). If only cellulosic glucose is considered as a substrate, the essential stages are

• Milling/grinding of the plant material to reduce particle size

• Chemical and/or physical pretreatment of the plant material to increase the exposure of the cellulose to enzyme (cellulase) attack

• Separation of soluble sugars and oligosaccharides

• Addition of either cellulase or a microorganism capable of secreting active cellulase and utilizing the released sugars for ethanol production by fermentation (simultaneous saccharification and fermentation) or direct microbial conversion

If the hemicellulosic sugars are also to be utilized, then either hemicellulases need to be added or a mixture of organisms used in cofermentations or sequential fermenta­tions. As more of the total potential substrate is included in the fermentation step, the biology inevitably becomes more complex — more so if variable feedstocks are to be used during the year or growing season — as the total available carbohydrate input to the biological fermentation step will alter significantly (table 1.6).

In 1996, several years’ experience with pilot plants worldwide using either enzyme conversion or acid-catalyzed hydrolysis of candidate cellulosic feedstocks inspired the prediction that technologies for the conversion of lignocellulosic bio­mass to ethanol would be rapidly commercialized.50 A decade on, “generic” tech­nologies have failed to emerge on large-scale production sites; in April 2004, Iogen

w о

FIGURE 1.24 Outline of lignin biosynthesis.

Compositional Analyses of Tree Species, Paper Recyclates, Cereal Wastes, and Grasses (% Dry Weight Basis)

(Ottawa, Ontario, Canada) opened a demonstration facility capable of processing 40 tonnes of feedstock/day and producing 3 million liters of ethanol annually from wheat, oat, and barley straw, corn cobs, and corn stalk.51 Iogen was founded in 1974 and has received CAN$91.8 million in research funding from the government of Canada, Petro-Canada, and Shell Global Solutions International BV. Iogen and its

European partners are studying the feasibility of producing cellulosic ethanol in Germany and, in May 2006, attracted CAN$30 million from Wall Street investors Goldman Sachs.52-54 The unique status of Iogen indicated that the technical hurdles to be overcome for cellulosic materials were considerably higher than for corn­starch, and it is highly likely that major advances in biotechnology will be required to tailor enzymes for the fledgling industry as well as providing novel biocatalysts for fermentative steps and optimizing plant species as “energy crops.”55 Signifi­cantly, Iogen is also an industrial producer of enzymes used in textiles, pulp and paper, and animal feed.[7]

Since 2006, a pilot facility in Jennings, Louisiana, has operated for the production of cellulosic ethanol; in February 2007, construction commenced on an adjacent demonstration facility designed to utilize regionally available feedstocks, including sugarcane bagasse, and the technology has also been licensed to Japanese companies to develop a project facility at Osaka to produce 1.3 million liters of ethanol annu­ally from demolition wood waste (www. verenium. com). After developing facilities to produce more than a million tons of ethanol from corn and wheat grain by 1995, Chinese scientists at Shandong University devised a bioethanol production process from corn cob, and a 50,000 ton/year plant for ethanol and xylose-derived products is planned to be constructed at Yucheng.56

The “promise” of lignocellulosic bioethanol remains quantitatively persuasive. Estimates of land area available for biomass energy crops and of the utilization of wood industry, agricultural, and municipal solid waste total 1.3-2.3 billion tons of cellulosic biomass as potential annual inputs to bioethanol production, poten­tially equivalent to a biofuel supply matching 30-50% of current U. S. gasoline consumption.3757 In stark contrast, even if all U. S. corn production were to be dedi­cated to ethanol, only 12% of the gasoline demand would be met.58 Data from Can­ada show similar scenarios, i. e., the total 2004 demand for fuel ethanol was met from 2,025 million liters of wheat, barley, corn, and potatoes, but the available nonfood crop supplies then already amounted to nearly 11,500 million liters as corn stover, straw, wood residues, and forest residues.59

The 2005 Energy Policy Act (http://www. ferc. gov) continued the influential role of legislation on renewable energy sources with initiatives to

• Increase cellulosic ethanol production to 250 million gallons/year

• Establish loan guarantees for new facilities

• Create an advanced biofuels technologies program

Continued interest in novel biotechnological solutions to the problems of lignocellulosic bioethanol are highly likely to be maintained over the next decade. The scientific aspects of present developments and future requirements are discussed in chapters 2 and 3.

Definitive comparisons of recombinant ethanologenic bacteria in tightly controlled, side-by-side comparisons have not been made public. From data compiled in 2003 from various sources with E. coli, K. oxytoca, and Z. mobilis strains fermenting

Geysers from Yellowstone National Park also yielded one promising isolate.

mixtures of glucose, xylose, and arabinose, conflicting trends are evident.162 For maximum ethanol concentration, the ranking order was

Z. mobilis AX101 >> K. oxytoca P2 = E. coli FBR5,

but for ethanol yield (percentage of maximum possible conversion), E. coli was superior:

E. coli FBR5 > K. oxytoca P2 = Z. mobilis AX101,

and the rankings of ethanol production rate (grams per liter per hour) were again different:

E. coli FBR5 >> Z. mobilis AX101 >> K. oxytoca P2.

All three strains can utilize arabinose, xylose, and glucose, but Z. mobilis AX101 can­not utilize the hemicellulose component hexose sugars galactose or mannose. In tests of E. coli, K. oxytoca, and Erwinia chrysanthemi strains, only E. coli KO11 was able to convert enzyme-degraded polygalacturonic acid (a pectin polymer) to ethanol.221

Enteric bacteria (including E. coli and Klebsiella sp.) have the additional hurdle to overcome of being perceived as potentially injurious to health, and K. oxytoca has been implicated in cases of infectious, hospital-acquired, and antibiotic-associated diseases.238-240 K. oxytoca is well known as a producer of a broad-spectrum P — lactamase, an enzyme capable of inactivating penicillins and other p-lactam antibiotics.241 Immunosuppression of patients under medical supervision or as a result of pathogenesis has led to the identification of infections by hitherto unknown yeast species or by those not considered previously to be pathogenic, including Kluyveromyces marxianus, five species of Candida, and three species of Pichia.17 Biosafety issues and assessments will, therefore, be important where planning (zon­ing) permissions are required to construct bacterial bioethanol facilities.

Because commercially relevant biomass plants for lignocellulosic ethanol only began operating in 2004 (chapter 2, section 2.7), future planned sites in divers parts of the world will inevitably make choices of producing organism that will enor­mously influence the continued development and selection of candidate strains.242 Different substrates and/or producing regions may arrive at different choices for optimized ethanol producer, especially if local enzyme producers influence the choice, if licensing agreements cannot be made on the basis of exclusivity, or if national interests encourage (or dictate) seamless transfer of technologies from labo­ratories to commercial facilities. More than a decade ago, a publication from the National Renewable Energy Laboratory ranked Z. mobilis ahead of (in descending order of suitability): recombinant Saccharomyces, homofermentative Lactobacillus, heterofermentative Lactobacillus, recombinant E. coli, xylose-assimilating yeasts, and clostridia.202 Their list of essential traits included •

• Low fermentation pH (to discourage contaminants)

• High fermentation selectivity

• Broad substrate utilization range

• GRAS status

The secondary list of 19 “desirable” traits included being Crabtree-positive (see above, section 3.1.1), high growth rates, tolerance to high salts, high shear, and elevated tem­perature. No commercialization of the Z. mobilis biocatalyst is yet established, but if bacterial ethanologens remain serious candidates for commercial bioethanol produc­tion, clear evidence for this should appear in the next decade as increasing numbers of scale-up bioethanol facilities are constructed for a variety of biomass feedstocks.

The increasing list of patents issued to companies and institutions for ethanologens testifies to the endeavors in this field of research and development (table 3.6). More problematic is that the field has been gene-led rather than genome — or (more usefully still) metabolome-led, that is, with full cognizance and appreciation of the flexibility and surprises implicit in the biochemical pathway matrices. The “stockpiling” of useful strains, vectors, and genetic manipulation techniques has built a process platform for the commercialization of ethanol production from lignocellulose biomass. The scien­tific community can find grounds for optimism in the new insights in metabolic engi­neering described in this chapter; what is less clear is the precise timescale — 10, 15, or more years — required to translate strain potential into industrial-scale production.243

2.4.1 Enzymology of Cellulose Degradation by Cellulases

Enzymic saccharification of pretreated biomass has gradually supplanted acid hydrolysis in pilot plant developments for ethanol production from lignocellulosic substrates (table 2.3). “Cellulase” is a deceptively complex concept, a convenient shorthand term for four enzyme activities and molecular entities, each with their Enzyme Commission (EC) identifying numbers, required for the complete hydrolytic breakdown of macromolecular cellulose to glucose:5960

1. Endoglucanases (1,4-P-D-glucan-4-glucanohydrolases, EC 3.2.1.4) decrease the degree of polymerization of macromolecular cellulose by attacking accessible sites and breaking the linear cellulose chain.

2. Cellodextrinases (1,4-P-D-glucan glucanohydrolases, EC 3.2.1.74) attack the chain ends of the cellulose polymers, liberating glucose.

3. Cellobiohydrolases (1,4-P-D-glucan cellobiohydrolases, EC 3.2.1.91) attack the chain ends of the cellulose polymers, liberating the disaccharide cellobiose, the repeating unit of the linear 1,4-linked polyglucan chain (figure 1.23).

4. Finally, P-glucosidases (EC 3.2.1.21) hydrolyze soluble cellodextrins (1,4- P-D-glucans) and cellobiose to glucose.

Plurals have been used for each discrete enzyme activity because cellulolytic organisms often possess multiple genes and separable enzymically active proteins; for example, the fungus Hypocrea jecorina[14] contains two cellobiohydrolases, five endoglucanases, and two P-glucosidases.61-63

Cellulases are widely distributed throughout the global biosphere because not only is cellulose the single most abundant polymer, but many organisms have also evolved in widely different habitats to feed on this most abundant of resources. Bacteria and fungi produce cellulases in natural environments and while contained in the diges­tive systems of ruminant animals and wood-decomposing insects (e. g., termites), but insects themselves may also produce cellulases, and other higher life forms — plants and plant pathogenic nematodes — certainly do.6064 Higher plants need to reversibly “soften” or irreversibly destroy cell wall structures in defined circumstances as part of normal developmental processes, including plant cell growth, leaf and flower abscis­sion, and fruit ripening; these are highly regulated events in cellular morphology.

The online Swiss-Prot database (consulted in December 2006) listed no fewer than 120 endoglucanases, 22 cellobiohydrolases, 4 cellodextrinases, and 27 P-glucosidases (cellobiases) in its nearly 250,000 entries of proteins fully sequenced at the amino acid level; most of these cellulase components are from bacteria and fungi, but higher plants and the blue mussel (Mytilus edulis) are also represented in the collection.65 The enormous taxonomic diversity of cellulase producers has aroused much speculation; it is likely, for example, that once the ability to produce cellulose had evolved with algae and land plants, cellulase producers arose on separate occasions in different ecological niches; moreover, gene transfer between widely different organisms is thought to occur easily in such densely populated microbial environments as the rumen.66

From the biotechnological perspective, fungal and bacterial cellulase producers have been foci of attention as potential industrial sources. More than 60 cellulolytic fungi have been reported, including soft-rot, brown-rot, and white-rot species — the last group includes members that can degrade both cellulose and lignin in wood samples.67 The penetration of fungal hyphae through the solid growth substrate represented by intact wood results in an enormous surface area of contact between the microbial population and lignocellulosic structures; the release of soluble enzymes then results in an efficient hydrolysis of accessible cellulose as different exo — and endoglucanases attack macroscopic cellulose individually at separate sites, a process often referred to as “synergy” — it is also highly significant that even different mem­bers of the same exo — or endoglucanase “family” have different substrate selectivities (table 2.4), thus ensuring a microdiversity among the catalytic population and a maxi­mized capacity to hydrolyze bonds in cellulosic glucans that may have different poly­mer-polymer interactions.59 Aerobic bacteria have a similar strategy in that physical adherence to cellulose microfibers is not a prerequisite for cellulose degradation, and a multiplicity of “cellulases” is secreted for maximal cellulose degradation — pres­ently, the most extreme example is a marine bacterium whose extraordinary meta­bolic versatility is coded by 180 enzymes for polysaccharide hydrolysis, including 13 exo — and endoglucanases, two cellodextrinases, and three cellobiases.68

Anaerobic bacteria, however, contain many examples of a quite different biochemical approach: the construction of multienzyme complexes (cellulosomes) on the outer surface of the bacterial cell wall; anaerobic cellulolytics grow optimally when attached to the cellulose substrate, and for some species, this contact is obligatory.60 The ability of such anaerobic organisms to break down cellulose and to ferment the resulting sugars to a variety of products including ethanol has prompted several investigators to promote them as ideal candidates for ethanol production from lignocellulosic biomass.69

The drive to commercialize cellulases — in applications as diverse as the stonewashing of denims, household laundry detergent manufacture, animal feed

TABLE 2.4

Substrate Selectivities of Trichoderma Cellulase Components

Endoglucanase

Cellobiohydrolase Cellobiohydrolase

3

2

5

1

3

1

1

1

Note: Numbers represent relative activity: 0 inactive, 5 maximum activity production, textile “biopolishing,” paper deinking, baking, and fruitjuice and beverage processing70 — has ensured that the biochemistry of the exo — and endoglucanases that attack macromolecular cellulose has been extensively researched. Most of these enzymes share a fundamental molecular architecture comprising two “domains” or “modules”: a cellulose-binding region (CBD or CBM) and a catalytic module or core.71 As more cellulase enzymes have been sequenced at the levels of either amino acids or (almost invariably now) genes, families of conserved polypeptide structures for CBD/CBM have been recognized; they form part of the 34 presently recognized carbohydrate-binding modules collated in a continuously updated data — base.72 All proteins in three families (CBM1, CBM5, and CBM10) bind to crystalline cellulose, whereas proteins in the CBM4 and CBM6 families bind to cellulose as well as to xylans and other polysaccharides using different polysaccharide binding sites.73 Removal of the portion of the cellulase responsible for binding to cellulose reduces cellulase activity with cellulose as the substrate but not with cellodextrins; conversely, the isolated binding domains retain their affinity for cellulose but lack catalytic action.74 The contribution of cellulose binding to overall cellulase activ­ity has more recently been elegantly demonstrated in a more positive manner: the endoglucanase II from Hypocrea jecorina has five amino acid residues in a topo­graphically distinct planar surface in the CBM; selectively altering the amino acids at two positions increased and decreased cellulose binding affinity and cellulose hydrolysis rate in synchrony (table 2.5) and demonstrated that the native enzyme could be catalytically improved by combinatorial mutagenesis.75

• The Cip units are anchored to the cell wall via other cohesin domains.

• Large and stable complexes are formed with molecular weights in the range of 2-16 MDa, and polycellulosomes occur with molecular weights up to 100 MDa.

The elaboration of such complex structures is thought to aid the energy-deficient anaerobes by maximizing the uptake of cellodextrins, cellobiose, and glucose by the spatially adjacent bacteria and ensuring a greatly increased binding affinity to the

cellulose.59,60

The cellulases of cellulolytic anaerobes may also be more catalytically effi­cient than those of typical aerobes and, in particular, the soluble cellulases secreted by fungi — but this is controversial because enzyme kinetics of the cellulose/cel — lulase system are problematic: the equations used for soluble enzyme and low — molecular-weight substrates are inadequate to describe the molecular interactions for cellulases “dwarfed” and physically binding to macroscopic and insoluble cel­luloses.59 Nevertheless, some results can be interpreted as showing that clostridial cellulases are up to 15-fold more catalytically efficient (based on specific activity measurements, i. e., units of enzyme activity per unit enzyme protein) than are fun­gal cellulases.60 Similarly, a comparison of fungal (Hypocrea, Humicola, and Irpex [Polyporus]) and aerobic bacterial (Bacillus, Pseudomonas, and Thermomonospora) cellulases noted a higher specific activity of more than two orders of magnitude with the bacterial enzymes.67

After acid hydrolysis (or some other pretreatment) and cellulase digestion, the product of the process is a mixed carbon source for fermentation by an ethanologenic microbe. Few details have been made public by Iogen about their development of nutritional balances, nitrogen sources, or media recipes for the production stage fer — mentation.1 This is not surprising because, for most industrial processes, medium optimization is a category of “trade secret,” unless patenting and publication priori­ties deem otherwise. Few industrial fermentations (for products such as enzymes, antibiotics, acids, and vitamins) rival the conversion efficiency obtained in ethanol production; one notable exception — about which a vast literature is available — is that of citric acid manufacture using yeasts and fungi.110 Many of the main features of citric acid fermentations have echoes in ethanol processes, in particular the use of suboptimal media for growth to generate “biological factories” of cell populations supplied with very high concentrations of glucose that cannot be used for further growth or the accumulation of complex products but can be readily fluxed to the simple intermediate of glucose catabolism — the biochemistry of Aspergillus niger strains used for the production of citric acid at concentrations higher than 100 g/l is as limited (from the viewpoints of biological ingenuity and bioenergetics) as ethanol production by an organism such as Z. mobilis (see figure 3.4).

Nevertheless, interest in media development for ethanol production has been intense for many years in the potable alcohol industry, and some innovations and developments in that industrial field have been successfully translated to that of fuel ethanol production.

3.2.1 Increased Pentose Utilization by Ethanologenic Yeasts by Genetic

Manipulation with Yeast Genes for Xylose Metabolism via Xylitol

It has been known for many years that S. cerevisiae cells can take up xylose from nutrient media; the transport system is one shared by at least 25 sugars (both natural and synthetic), and only major alterations to the pyranose ring structure of hexoses (e. g., in 2-deoxyglucose, a compound that lacks one of the hydroxyl groups of glu­cose) reduce the affinity of the transport system for a carbohydrate.53 Moreover, both D-xylose and L-arabinose can be reduced by S. cerevisiae, the products being the sugar alcohols, xylitol, and L-arabinitol, respectively; three separate genes encode enzymes with overlapping selectivities for xylose and arabinose as substrates.54

Indeed, the wild-type S. cerevisiae genome does contain genes for both xylose reductase and xylitol dehydrogenase, thus being able to isomerize xylulose from xylose, and the resulting xylulose (after phosphorylation catalyzed by a specific xylulokinase) can enter the pentose phosphate pathway (figure 3.2).55,56 Overex­pressing the endogenous yeast genes for xylose catabolism renders the organism capable of growth on xylose in the presence of glucose as cosubstrate under

conditions, although no ethanol is formed; S. cerevisiae may, therefore, have evolved originally to utilize xylose and other pentoses, but this has been muted, possibly because its natural ecological niche altered or the organism changed its range of favored environments.55

Ethanol production from xylose is a rare phenomenon; of 200 species of yeasts tested under controlled conditions in the laboratory, only six accumulated ethanol to more than 1 g/l (0.1% by volume): Pichia stipitis, P. segobiensis, Candida shehatae, C. tenuis, Brettanomyces naardenensis, and Pachysolen tannophilus.56 Even rarer is the ability among yeasts to hydrolyze xylans, only P. stipitis and C. shehatae[24] hav­ing xylanase activity; P. stipitis could, moreover, convert xylan into ethanol at 60% of the theoretical yield as computed from the xylose content of the polymer.58 Three naturally xylose-fermenting yeasts have been used as donors for genes encoding enzymes of xylose utilization for transfer to S. cerevisiae: P. stipitis, C. shehatae, and C. parapsilosis.59-61 These organisms all metabolize xylose by the enzymes of the same low-activity pathway known in S. cerevisiae (figure 3.2), and the relevant enzymes appear to include arabinose as a possible substrate — at least, when the enzymes are assayed in the laboratory.

P. stipitis has been the most widely used donor, probably because it shows relatively little accumulation of xylitol when growing on and fermenting xylose, thus wasting less sugar as xylitol.62 This advantageous property of the yeast does not appear to reside in the enzymes for xylose catabolism but in the occurrence of an alternative respiration pathway (a cyanide-insensitive route widely distributed among yeasts of industrial importance); inhibiting this alternative pathway renders P. stipitis quite capable of accumulating the sugar alcohols xylitol, arabinitol, and ribitol.63 Respiration in P. stipitis is repressed by neither high concentrations of fer­mentable sugars nor by O2 limitation (i. e., the yeast is Crabtree-negative), and as an ethanologen, P. stipitis suffers from the reduction of fermentative ability by aerobic conditions.64

Transferring genetic information from P. stipitis in intact nuclei to S. cerevi­siae produced karyoductants, that is, diploid cells where nuclei from one species have been introduced into protoplasts of another, the two nuclei subsequently fusing, with the ability to grow on both xylose and arabinose; the hybrid organism was, however, inferior to P. stipitis in ethanol production and secreted far more xylitol to the medium than did the donor; its ethanol tolerance was, on the other hand, almost exactly midway between the tolerance ranges of S. cerevisiae and P. stipitis.65 For direct genetic manipulation of S. cerevisiae, however, the most favored strategy (starting in the early 1990s) has been to insert the two genes (xyl1 and xyl2) from P. stipitis coding for xylose reductase (XR) and xylitol dehydrogenase (XDH), respectively.66 Differing ratios of expression of the two foreign genes resulted in smaller or higher amounts of xylitol, glycerol, and acetic acid, and the optimum XR:XDH ratio of 0.06:1 can yield no xylitol, less glycerol and acetic acid, and more ethanol than with other engineered S. cerevisiae strains.67

The first patented Saccharomyces strain to coferment xylose and glucose to ethanol (not S. cerevisiae but a fusion between S. diastaticus and S. ovarum able to produce ethanol at 40°C) was constructed by the Laboratory of Renewable

Resources Engineering at Purdue University, with four specific traits tailored for industrial use:68-70

1. To effectively direct carbon flow from xylose to ethanol production rather than to xylitol and other by-products

2. To effectively coferment mixtures of glucose and xylose

3. To easily convert industrial strains of S. cerevisiae to coferment xylose and glucose using plasmids with readily identifiable antibiotic resistance markers controlling gene expression under the direction of promoters of S. cerevisiae glycolytic genes

4. To support rapid bioprocesses with growth on nutritionally rich media

In addition to XR and XDH, the yeast’s own xylulose-phosphorylating xylulokinase (XK, figure 3.2) was also overexpressed via high-copy-number yeast-£. coli shuttle plasmids.68 This extra gene manipulation was crucial because both earlier and contem­porary attempts to transform S. cerevisiae with only genes for XR and XDH produced transformants with slow xylose utilization and poor ethanol production.68 The synthesis of the xylose-metabolizing enzymes not only did not require the presence of xylose, but glucose was incapable of repressing their formation. It was known that S. cerevisiae could consume xylulose anaerobically but only at less than 5% of the rate of glucose utilization; XK activity was very low in unengineered cells, and it was reasoned that providing much higher levels of the enzyme was necessary to metabolize xylose via xylulose because the P. stipitis XDH catalyzed a reversible reaction between xylitol and xylulose, with the equilibrium heavily on the side of xylitol.70-72 Such strains were quickly shown to ferment corn fiber sugars to ethanol and later utilized by the Iogen Corporation in their demonstration process for producing ethanol from wheat straw.7374

The vital importance of increased XK activity in tandem with the XR/XDH pathway for xylose consumption in yeast was demonstrated in S. cerevisiae: not only was xylose consumption increased but xylose as the sole carbon source could be con­verted to ethanol under both aerobic and anaerobic conditions, although ethanol pro­duction was at its most efficient in microaerobiosis (2% O2).75 Large increases in the intracellular concentrations of the xylose-derived metabolites (xylulose 5-phosphate and ribulose 5-phosphate) were demonstrated in the XK-overexpressing strains, but a major drawback was that xylitol formation greatly exceeded ethanol production when O2 levels in the fermentation decreased.

In the intervening years (and subsequently), considerable efforts have been dedicated to achieving higher ethanol productivity with the triple XR/XDH/XK constructs. Apart from continuing attempts to more fully understand the metabolism of xylose by uncon­ventional or little-studied yeast species, two main centers of attention have been evident:

1. Strategies for harmonizing the different cofactor requirements in the path­way, that is, NAPDH-dependent (or NAPDH-preferring) XR and NAD- requiring XDH, and thus reducing xylitol formation

2. Overexpressing a wider array of other pentose-metabolizing enzymes to maximize the rate of xylose use or (broadening the metabolic scope) increas­ing kinetic factors in the central pathways of carbohydrate metabolism

Because the early attempts to overexpress P. stipitis genes for XR and XDH in S. cerevisiae often resulted in high rates of xylitol formation, if NADPH formed in the oxidative pentose phosphate pathway (figure 3.2) equilibrated with intracellular NAD to form NADH, the reduced availability of NAD could restrict the rate of the XDH reaction in the direction of xylitol oxidation to xylulose; adding external oxidants capable of being reduced by NADH improved ethanol formation and reduced xylitol formation. Two of these were furfural and 5-hydroxyfurfural, known to be sugar deg­radation products present in lignocellulose acid hydrolysates — see chapter 2, sec­tion 2.3.4.78 The adventitious removal of toxic impurities by these reactions probably explained why xylitol accumulation was very low when lignocellulose acid hydroly­sates were used as carbon sources for XR — and XDH-transformed S. cervisiae 7 This line of reasoning does not accord entirely with the high activities of XR measurable in vitro with NADH (63% of the rate with NADPH), but site-specific mutagenesis on the cloned P. stipitis XR gene could alter the activity with NADH to 90% of that with NADPH as cofactor and concomitantly greatly reduce xylitol accumulation although with only a marginally increased xylose utilization rate.69 Further optimization of the xylitol pathway for xylose assimilating was, therefore, entirely possible. Simply coalescing the XR and XDH enzymes into a single fusion protein, with the two active units separated by short peptide linkers, and expressing the chimeric gene in S. cere — visiae resulted in the formation of a bifunctional enzyme; the total activities of XR and XDH were similar to the activities when monomeric enzymes were produced, but the molar yield of xylitol from xylose was reduced, the ethanol yield was higher, and the formation of glycerol was lower, suggesting that the artificially evolved enzyme complex was more selective for NADH in its XR domain as a consequence of the two active sites generating and utilizing NADH being near each other.80

There have also been four direct methodologies tested for altering the preference of XR to use the NADPH cofactor:

1. A mutated gene for a P. stipitis XR with a lower affinity for NADPH replaced the wild-type XR gene and increased the yield of ethanol on xylose while decreasing the xylitol yield but also increasing the acetate and glycerol yields in batch fermentation.81

2. The ammonia-assimilating enzyme glutamate dehydrogenase in S. cerevisiae (and other yeasts) can be either NADPH or NADH specific, setting an artificial transhydrogenase cycle by simultaneously expressing genes for both forms of the enzyme improved xylose utilization rates and ethanol productivity.82

3. Deleting the gene for the NADPH-specific glutamate dehydrogenase aimed to increase the intracellular NADH concentration and the competition between NADH and NADPH for XR but greatly reduced growth rate, ethanol yield, and xylitol yield on a mixture of glucose and xylose; overexpressing the gene for the NADH-specific enzyme in the absence of the NADPH-requir — ing form, however, restored much of the loss in specific growth area and increased both xylose consumption rate when glucose had been exhausted and the ethanol yield while maintaining a low xylitol yield.83

4. Redox (NADPH) regeneration for the XR reaction was approached from a different angle by expressing in a xylose-utilizing S. cerevisiae strain the gene for an NADP-dependent D-glyceraldehyde 3-phosphate dehydrogenase, an enzyme providing precursors for ethanol from either glucose or xylose; the resulting strain fermented xylose to ethanol at a faster rate and with a higher yield.84

Two recent discoveries offer novel biochemical and molecular opportunities: first, the selectivity of P. stipitis XDH has been changed from NAD to NADP by multiple — site-directed mutagenesis of the gene, thereby harmonizing the redox balance with XR; second, an NADH-preferring XR has been demonstrated in the yeast Candida parapsilopsis as a source for a new round of genetic and metabolic engineering.6185

Beyond the initial conversions of xylose and xylitol, pentose metabolism becomes relatively uniform across kingdoms and genera. Most microbial species — and plants, animals, and mammals (including Homo sapiens) — can interconvert some pentose structures via the nonoxidative pentose phosphate pathway (figure 3.2). These reactions are readily reversible, but extended and reorganized, the pathway can function to fully oxidize glucose via the glucose 6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase reactions (both forming NADPH), although the pathway is far more important for the provision of essential biosynthetic intermedi­ates for nucleic acids, amino acids, and cell wall polymers; the reactions can even (when required) run “backward” to generate pentose sugars from triose intermedi­ates of glycolysis.86 Increasing the rate of entry of xylulose into the pentose phosphate pathway by overexpressing endogenous XK activity has been shown to be effec­tive for increasing xylose metabolism to ethanol (while reducing xylitol formation) with XR/XDH transformants of S. cerevisiae.81 In contrast, disrupting the oxidative pentose phosphate pathway genes for either glucose 6-phosphate or 6-phosphoglu — conate oxidation increased ethanol production and decreased xylitol accumulation from xylose by greatly reducing (or eliminating) the main supply route for NADPH; this genetic change also increased the formation of the side products acetic acid and glycerol, and a further deleterious result was a marked decrease in the xylose consumption rate — again, a predictable consequence of low NADPH inside the cells as a coenzyme in the XR reaction.88 Deleting the gene for glucose 6-phosphate dehydrogenase in addition to introducing one for NADP-dependent D-glyceraldehyde 3-phosphate dehydrogenase was an effective means of converting a strain ferment­ing xylose mostly to xylitol and CO2 to an ethanologenic phenotype.84

The first report of overexpression of selected enzymes of the main nonoxidative pentose phosphate pathway (transketolase and transaldolase) in S. cerevisiae harbor­ing the P. stipitis genes for XR and XDH concluded that transaldolase levels found naturally in the yeast were insufficient for efficient metabolism of xylose via the pathway: although xylose could support growth, no ethanol could be produced, and a reduced O2 supply merely impaired growth and increased xylitol accumulation.89 In the most ambitious exercise in metabolic engineering of pentose metabolism by

S. cerevisiae reported to date, high activities of XR and XDH were combined with overexpression of endogenous XK and four enzymes of the nonoxidative pentose phosphate pathway (transketolase, transaldolase, ribulose 5-phosphate epimerase, and ribose 5-phosphate ketolisomerase) and deletion of the endogenous, nonspecific NADPH-dependent aldose reductase (AR), catalyzing the formation of xylitol from xylose.90 In comparison with a strain with lower XR and XDH activities and no other genetic modification other than XK overexpression, fermentation performance on a mixture of glucose (20 g/l) and xylose (50 g/l) was improved, with higher ethanol production, much lower xylitol formation, and faster utilization of xylose; deleting the nonspecific AR had no effect when XR and XDH activities were high, but glyc­erol accumulation was higher (figure 3.5).

To devise strains more suitable for use with industrially relevant mixtures of carbohydrates, the ability of strains to use oligosaccharides remaining undegraded to free hexose and pentose sugars in hydrolysates of cellulose and hemicelluloses is essential. Research groups in South Africa and Japan have explored combinations of heterologous xylanases and P-xylosidases:

• A fusion protein consisting of the xynB P-xylosidase gene from Bacillus pumilus and the S. cerevisiae Mfa1 signal peptide (to ensure the correct posttranslational processing) and the XYN2 P-xylanase gene from Hypo — crea jecorina were separately coexpressed in S. cerevisiae under the con­trol of the glucose-derepressible ADH2 ADH promoter and terminator; coproduction of these xylan-degrading enzymes hydrolyzed birch wood xylan, but no free xylose resulted, probably because of the low affinity of the P-xylosidase for its xylobiose disaccharide substrate.91

• A similar fusion strategy with the xlnD P-xylosidase gene from Aspergillus niger and the XYN2 P-xylanase gene from H. jecorina enabled the yeast to hydrolyze birch wood xylan to free xylose.92

• A xylan-utilizing S. cerevisiae was constructed using cell surface engi­neering based on в-agglutinin (a cell surface glycoprotein involved in cell-cell interactions) to display xylanase II from Hypocrea jecorina and a в-xylosidase from Aspergillus oryzae; with P. stipitis XR and XDH and overexpressed endogenous XK, the strain could generate ethanol directly from birch wood xylan with a conversion efficiency of 0.3 g/g carbohydrate used.93

High XR, XDH, and XK activities combined with the expression of a gene from Aspergillus acleatus for displaying в-glucosidase on the cell surface enabled S. cere­visiae to utilize xylose — and cellulose-derived oligosaccharides from an acid hydro­lysate of wood chips, accumulating 30 g/l of ethanol from a total of 73 g/l of hexose and pentose sugars in 36 hr.94