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

The assessment of corn-derived ethanol was the most extensive of the reports (60% of the total printed pages in the final collection of papers) and formed a notional blueprint for a facility sited in Illinois with a projected working life of 20 years and operating costs of approximately 950/gallon of hydrous ethanol (table 5.1). The final factory gate selling price was computed to be $1.05/gallon (1978 prices) in the base case of the 50-million-gallon/year capacity including the results of a 15% dis­counted cash flow/interest rate of return analysis; the selling price was a little lower (980/gallon) with twice the annual capacity but considerably higher ($1.55/gallon) at only 10 million gallons/year. The quoted comparative price for refinery gasoline was 400/gallon; after allowing for the lower energy content of ethanol (70% of that of gasoline — chapter 1, section 1.3), the “real” cost of corn-derived ethanol would have been $1.50/gallon for the 50-million-gallon facility, that is, 3.75-fold higher than gasoline at that time.

Various options were explored in the study to define the sensitivity of the required selling price for ethanol:

1. The DOE required the analyses to define a selling price that would cover not only the annual operating expenses but also yield a return on equity, the base case being a 15% discounted cash flow/interest rate of return; increas­ing this factor to 20% resulted in a higher selling price ($1.16/gallon for the base-case scenario).

2. Lengthening the depreciation schedule from 10 to 20 years increased the selling price by 20/gallon.

3. Increasing the working capital to 20% of the total production cost increased the selling price by 30/gallon.

4. A higher investment tax credit (50%) would reduce the selling price by 20/gallon.

5. Financing only 80% of the plant investment could reduce the selling price by 100/gallon.

6. For every 10% rise in the price of corn, the selling price would increase by 80/gallon (after allowing for a triggered rise in the selling price of the solid animal feed coproduct).

7. For every 10% rise in the price of the animal feed coproduct, the selling price would decrease by 40/gallon.

8. Replacing local coal by corn stover as the fuel for steam generation would increase the selling price by 40/gallon — although a lower total investment (by approximately $1 million) would have been an advantage resulting from the removal of the need for flue gas desulfurization.

All of these changes are comparatively minor, and other quantified changes to the over­all process were likely to have been equally small: ammonium sulfate (a coproduct aris­ing from flue gas desulfurization) was only generated in small amounts, approximately 3 tons/day, and no allowance was made for capturing and selling the CO2 generated in the fermentation step. No denaturant was included in the final cost breakdown.

Alternative feedstocks were also explored. Milo (grain sorghum) offered a slight reduction in the selling price of ethanol (down to $1.02/gallon) but was considered a

small-acreage crop at that time. Both wheat and sweet sorghum were likely to increase the final factory gate selling price to $1.31/gallon and $1.40/gallon, respectively. Although wheat and milo grain could be processed in essentially the same equipment used for corn, sweet sorghum required a higher investment in plant facilities.

The best taxonomically and physiologically characterized examples of H2 pro­ducers are clostridia, but other genera (including bacilli), as well as a microbial flora from anoxic marsh sediments and other environments are known that are capable of the H2 production and either the ABE “solvent” fermentation (chap­ter 6, section 6.3.3), the accumulation of one or more of the ABE trio, carbox­ylic acids (acetic, butyric, etc.) and/or other products (acetoin, 2,3-butanediol, etc.).26

A wide spectrum of carbon sources supports H2 production at rates up to 1,000 ml/hr/g cells at a maximum yield of 4 mol of H2/mol glucose with the stoichiometry:26

C6H12O6 + 2H2O ^ 2CH3COOH + 2CO2 + 4H2

This reaction is sufficiently exothermic (to support microbial growth). The yield of H2 is, however, subject to feedback inhibition by H2, requiring that the partial pres­sure of the gas be kept low to avoid problems with growth rate or a shift to acid production.

Another H2-forming fermentation has butyric acid as its major acidic product: C6H12O6 ^ C3H7COOH + 2CO2 + 2H2

although the molar production of H2 is only half that of acetate-accumulating strains.

The key enzyme in heterotrophic H2 producers is hydrogenase, an enzyme that catalyzes the reoxidation of reduced ferredoxin (Fd), an iron-containing protein reduced by ferredoxin-NAD and pyruvate-ferredoxin oxidoreductases, with the lib­eration of molecular hydrogen (figure 7.5):27

2Fd2+ + 2H+ ^ 2Fd3+ + H2

A summary of hydrogenase-containing bacteria is given in table 7.2.

Hydrogenases are a diverse group of enzymes and are often cataloged on the basis of the metal ion they contain as an essential component of the active site.28 The fastest H2-evolving species under laboratory conditions, Clostridium acetobutylicum, produces two different hydrogenases:29-31

• An iron-containing enzyme, whose gene is located on the chromosome

• A dual-metal (nickel, iron) enzyme whose gene is located on a large plasmid

Glucose

I Butyric Acid

Fructose-1, 6-bisphosphate

1

up to 0.59 ?

up to 0.9

?

up to 1.5

The iron-dependent hydrogenase from C. acetobutylicum has a specific activ­ity eightfold higher than similar enzymes from green algae even when all three enzymes are expressed in and purified from the clostridial host.32 The active sites of iron-dependent hydrogenases may be the simplest such structures yet studied at the molecular level. Of enormous potential importance for the industrial development of hydrogenases is the finding that even simple complexes of iron sulfide and CO mimic hydrogenase action.33 The crucial structure involves two iron atoms with different valency states at different stages of the reaction mechanism (figure 7.6). These findings raise the possibility of rational design of improved hydrogenases by the binding of novel metal complexes with existing protein scaffolds from known enzymes.

In contrast, nickel-iron bimetallic hydrogenases possess complex organometal — lic structures with CO and cyanide (CN-) as additional components, the metal ions bound to the protein via multiple thiol groups of cysteine resides, and an impor­tant coupling between the active site and iron-sulfur clusters.34,35 Multigene arrays are required for the biosynthesis of mature enzyme.36 Nevertheless, progress has been impressive in synthesizing chemical mimics of the organometallic centers that contain elements of the stereochemistry and atomic properties of the active site.37 The enzyme kinetics of nickel-iron hydrogenases remain challenging, and it is pos­sible that more than one type of catalytic activation step is necessary for efficient functioning in vivo.38 39

Such advances in basic understanding will, however, open the door to replacing expensive metal catalysts (e. g., platinum) in hydrogen fuel cells by iron — or iron/ nickel-based biocatalysts — the sensitivity of many hydrogenases to inhibition by O2

OC CO

Fe1 — Fe1———— Fe0 — Fe1

H’H H- ■

I I

Fe11—Fe1 ———— Fe11—Fe1

H+

FIGURE 7.6 Simple organometallic complexes as biochemical mimics of hydrogenase enzymes. (Modified from Darensbourg et al.33)

is a serious drawback but, of the many organism known to produce hydrogenases, some contain forms with no apparent sensitivity to O2 and can function under ambient levels of the gas.40

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.

Constructing a large-scale bioethanol industry also implies a major change in the industrial landscape: whereas oil refineries are predominately coastal, biorefiner­ies would be situated in agricultural areas or (with the development of a mature industry) close to forests and other biomass reserves. Gasoline distribution to retail outlets is without doubt a mature industry — in the United States, shipments of 6.4 billion liters of petroleum and petroleum products are made each day, 66% by pipe­line (in 320,000 miles of pipeline), but only 4% by truck and 2% by rail; from the Gulf Coast to New York, shipping costs for gasoline amount to only 0.8 0/l.76 It was the early development of a national distribution system for gasoline that decided the use of this fuel rather than ethanol for the emerging automobile industry before 1920.77

To support nationwide consumption of E10, cellulosic ethanol would be 61% of the total, the remainder being corn-derived; assuming that switchgrass will be a major contributor to the feedstock mix, ethanol production would be centered in a wide swathe of states from North Dakota to Georgia, whereas demand would have geographic maxima from west to east (figure 5.7). Ethanol shipping would be predominantly by truck or rail until the industry evolved to take over existing petroleum pipelines or to justify the construction of new ones; linear optimization showed that shipping by truck would entail a cost of $0.13/l ($0.49/gallon), whereas rail transport would entail lower costs, $0.05/l ($0.19/gallon). In contrast, gasoline transportation to retail outlets only incurs costs of $0.003/l ($0.01/gallon).76 The same study concluded that national solutions, although they would spur innovation and eventually lead to economies of scale, would increase shipping distances and add to total truck movements; an investment of $25 billion would be required for a dedicated ethanol pipeline system, “just to make petroleum pipelines obsolete in the long-term.”

Tax incentives and subsidies are, therefore, highly likely to be features of policy making relevant to the adoption of biofuels in OECD economies generally. Fund­ing the ethanol supply chain will be crucial; minimizing shipping costs implies the construction of as many production sites as possible, based on the use of raw materi­als from multiple geographical areas (forest, agricultural wastes, dedicated energy crops, municipal solid waste, etc.), ideally to match the likely distribution of major urban demand centers for ethanol blends (figure 5.7).

Xylitol was a significant biochemical feature of the metabolic routes for the xylose presented to ethanologenic cultures in hydrolysates of the hemicelluloses from ligno — cellulosic biomass (chapter 2, section 2.3, and chapter 3, section 3.2). With an organ­oleptic sweetness to human taste approximately equivalent to sucrose, however, it is a fine chemical product in its own right as a low-calorie sweetener — xylose sugars
are not metabolized by the human consumers of xylitol-containing chewing gums (figure 3.2). Other envisaged uses include15

• Production of anhydrosugars (as chemical intermediates) and unsaturated polyester resins

• Manufacture of propylene and ethylene glycols as antifreeze agents and unsaturated polyester resins

• Oxidation to xylonic and xylaric acids to produce novel polymers (polyes­ters and nylon-type structures)

The production of xylitol for use as a building block for derivatives essentially requires no technical development, and if the xylose feedstock is inexpensive (as a product of biomass processing), then the production of xylitol could be done for very low cost.

The accumulation of xylitol during ethanologenesis from lignocellulosic sub­strates is, of course, unwanted and quite undesirable — for process as well as for eco­nomic reason (chapter 3, section 3.2). Viewed as an economically valuable product, xylitol formation and production acquire a different biotechnological perspective, and patenting activity has recently been intense (table 8.2). Biochemical efforts also continue to locate and exploit enzymes for bioprocessing hemicelluloses and hemi — cellulosic waste streams, for example:

•

An L-xylulose reductase identified from the genome sequence of the fil­amentous mold Neurospora crassa has been heterologously produced in E. coli for the production of xylitol.49

• A P-xylosidase from Taloromyces emersonii has been shown to be superior to the enzyme from the industrial fungus Hypocrea jecorina in releasing xylose from vinasse, the solid-waste material from ethanol fermentations.50

• Xylan residues in hemicelluloses can be variably esterified with acetyl, fer — uloyl, and p-coumaryl residues; hemicellulose deacetylating esterases have been characterized from fungal species and shown to only be highly effec­tive when mixed in multiplexes of enzymes capable of using all the possible structures as substrates for enzyme action.51

• Feruloyl esterases have recently been designed as novel chimeric forms with cellulose and hemicellulose binding proteins to improve their efficien­cies with plant polymeric substrates.52,53

• Ferulic acid has a number of potential commercial applications as an anti­oxidant, food preservative, anti-inflammatory agent, photoprotectant, and food flavor precursor; major sources — brewer’s spent grain, wheat bran, sugarbeet pulp, and corn cobs — make up 1-2% of the daily output of the global food industry, and ferulic acid can be released from brewer’s grain and wheat bran by feruloyl esterases from the thermophilic fungus Humi — cola insolens.54

Once acquired, a hemicellulose hydrolysate contains a variety of hexoses and pentoses; yeast highly suitable for the production of xylitol from the xylose present in the mix may, however, preferentially utilize glucose. One solution to this problem is to remove the rapidly utilized hexoses before removing the cells and replacing them with a purposefully xylose-grown cell batch, as was demonstrated with dilute acid hydrolysates of corn fiber.55 With Saccharomyces cerevisiae expressing the Pichia stipitis gene for xylose reductase, the presence of glucose inhibited xylose uptake and biased the culture toward ethanol production; controlling the glucose concentra­tion by feeding the fermentation to maintain a high xylose:glucose ratio resulted in a near-quantitative conversion of xylose to xylitol, reaching a titer of 105 g/l xylitol.56 With a double-recombinant strain of S. cerevisiae carrying the xylose reductase genes from both P. stipitis and Candida shehatae, quite different microbial biochem­istry occurred in a more process-friendly formation (at close to theoretical levels) of xylitol from a mixture of xylose (the major carbon source) and glucose, galactose, or mannose as the cosubstrate — indeed, the presence of the cosubstrate was manda­tory for continued metabolism of the pentose sugar.57 In a gene-disrupted mutant of C. tropicalis, with no measurable xylitol dehydrogenase activity, glycerol proved to be the best cosubstrate, allowing cofactor regeneration and redox balancing, with a xylitol yield that was 98% of the maximum possible.58

Bioprocess engineering for xylitol production appears straightforward; aerated cultures with pH regulation and operated at 30°C appear common. With Brazilian sugarcane bagasse as the source of the hemicellulosic sugars, xylitol from the har­vested culture broth was recovered at up to 94% purity crystallization from a clari­fied and concentrated broth.59

Xylitol dehydrogenase enzymes catalyze reversible reactions. The catabolism of xylitol proceeds via the formation of D-xylulose (figure 3.2). Fungal pathways of L-arabinose utilization can include L-xylulose as an intermediate, a much less common pentulose sugar.60-63 Such “rare” sugars are of increasing interest to metabolic bio­chemists because it has long been appreciated that many naturally occurring antibi­otics and other bioactives contain highly unusual sugar residues, sometimes highly modified hexoses derived by lengthy biosynthetic pathways (e. g., erythromycins); many secondary metabolites elaborated by microbes may only exhibit weak anti­biotic activity but are or can easily be converted into chemicals with a wide range of biologically important effects: antitumor, antiviral, immunosuppressive, anti- cholesterolemic, cytotoxic, insecticidal, or herbicidal.64 To the synthetic chemist, therefore, the availability of such novel carbohydrates in large quantities offers new horizons in development novel therapeutic agents: for example, L-xylulose offers a promising route to inhibiting the glycosylation of proteins, including those of viruses.65

Equally, such unconventional chemicals may have very easily exploited proper­ties as novel fine chemicals. The hexose D-tagatose is an isomeric form of the com­monly occurring sugar D-galactose (a hexose present in hemicelluloses, figure 1.23) and has attracted interest for commercial development.66 Because of its very rare occurrence in the natural world, a structure such as d-tagatose presents a “tooth — friendly” metabolically intractable sugar to human biochemistry. Fortuitously, the enzyme L-arabinose isomerase includes among its spectrum of possible substrates d-galactose; the enzyme can be found in common bacteria with advanced molecular genetics and biotechnologies, including E. coli, Bacillus subtilis, and Salmonella typhimurium and, when expressed in suitable hosts, can convert the hexose into d — tagatose with a 95% yield.67 Even more promising for industrial use is the efficient bioconversion of d-galactose to d-tagatose using the immobilized enzyme, more active than free L-arabinose isomerase and stable for at least 7 days.68 Both the enzyme and recombinant L-arabinose isomerase-expressing cells can be used in packed-bed bioreactors, the cells being particular adaptable to long production cycles.69,70

Synthetic chemistry offers only expensive and low-yielding routes to the rare sugars, but uncommon tetroses, pentoses, and hexoses can all be manufactured with whole cells or extracted enzymes acting on cheap and plentiful carbon sources, including hemicellulose sugars; bioproduction strategies use an expanding toolkit of enzymes, including D-tagatose 4-epimerase, aldose isomerase, and aldose reduc­tase.71-73 Together, the rare sugars offer new markets for sugars and sugar derivatives of at least the same magnitude as that for high-fructose syrups manufactured with xylose (glucose) isomerase.

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

Iogen’s choice of a wheat straw feedstock was made on practical and commercial grounds from a limited choice of agricultural and other biomass resources in Canada available on a sufficiently large scale to support a bioethanol industry (section 4.1). Wheat, as a monoculture, is inevitably subject to crop losses arising from pathogen infestation. Modern biotechnology and genetic manipulation offer novel solutions to the development of resistance mechanisms as well as yield improvements through increased efficiency of nutrient usage and tolerance to drought, and other seasonal and unpredictable stresses. But the deliberate release of any such genetically modi­fied (GM) species is contentious, highly so in Europe where environmental cam­paigners are still skeptical that GM technologies offer any advantage over traditional plant breeding and are without the associated risks of monopoly positions adopted by international seed companies, the acquisition of desirable traits by “weed” species, and the horizontal transfer of antibiotic resistance genes to microbes.282 The positive aspects of plant biotechnology have, in contrast, been succinctly expressed:

Genetic transformation has offered new opportunities compared with traditional breed­ing practices since it allows the integration into a host genome of specific sequences leading to a strong reduction of the casualness of gene transfer.283

Because large numbers of insertional mutants have been collected in a highly manip — ulable “model” plant species (Arabidopsis thaliana), it has been possible for some years to inactivate any plant gene with a high degree of accuracy and certainty.

In comparison with natural gas-based FT syntheses, biomass requires more inten­sive engineering, and gas-cleaning technology has been slow to evolve for indus­trial purposes — although for the successful use of biomass, it is essential because of the sensitivity of FT catalysts to contaminants. In year 2000 U. S. doller terms, investment costs of $200-340 million would be required for an industrial facility, offering conversion efficiencies of 33-40% for atmospheric gasification systems and 42-50% for pressured systems, but the estimated production costs for FT die­sel were high, more than 10 times those of conventional diesel.89 Two years later,

Biomass Air, O2

Syngas

the same research group from the Netherlands was predicting the same produc­tion costs and concluded that, unless the environmental benefits of FT diesel were valued in economic terms, the technology would only become viable if crude oil prices rose substantially.90 This did, in the event, occur (figure 5.1), and the cost differential has undoubtedly narrowed — although with no signs of a surge in investor confidence.

If it could be produced economically, using an energy crop such as switchgrass as the substrate, FT diesel rates better than E85 (from corn-derived ethanol) as a biofuel in assessments performed by the Argonne National Laboratory (figure 6.9).9192 FT diesel greatly outperformed E85 for total fossil fuels savings and also exhibited much reduced emissions of total particulates, sulfur oxides, and nitric oxides — although it fared worse than E85 using the criterion of total CO. Compared with conventional diesel fuel, FT diesel had higher total emissions of volatile organic carbon, CO, and nitric oxides (figure 6.9).

An experimental biofuel conversion technology, presently explored only in the Netherlands, is that for HydroThermalUpgrading diesel.93 At high temperature (300-350°C) and pressure, wet biomass feedstocks such as beet pulp, sludge, and bagasse can be converted to a hydrocarbon-containing liquid that, after suitable refining, can be blended with conventional diesel in any proportion without engine adjustments. A pilot plant remains the focus for further process optimization and development.

3.4.1 "Traditional" Microbial Ethanologens

Given the present indeterminacy regarding “best” choices for producing organism and biomass substrate, it is timely to summarize recent advances in the metabolic and genetic sciences relevant to yeast and bacterial ethanologens, and from them to project forward key areas for their application to industrial ethanol production after 2007. Four themes can readily be identified:

1. A better understanding of stress responses

2. Relating transport phenomena to sugar utilization and ethanol productivity

3. Defining the roles of particular genes in the metabolism of ethanologens

4. Applying whole-genome methodologies and in silico simulations to opti­mize the balance of carbon flows along synergistic or competing pathways

Yeast ethanologens show dose-dependent inhibition by furfural and HMF and are generally more sensitive to furfural; a S. cerevisiae strain converted HMF to 2,5-bis-HMF, a previously postulated alcohol formed from the parent aldehyde, and this suggests that an enzymic pathway for detoxifying these components of lignocel- lulosic hydrolysates should be attainable by gene cloning and/or mutation to improve the activities and substrate preferences of the enzymes.244 That such a reduction reaction as that to form 2,5-bis-HMF might utilize NADPH as cofactor was shown by screening a gene disruption library from S. cerevisiae: one type of mutation was in zwfl, the gene encoding glucose 6-phosphate dehydrogenase, the NADPH-generating entry point to the oxidative pentose pathway; overexpression of zwfl allowed growth in furfural concentrations normally toxic, whereas similar effects with non-redox- associated gene products could be explained as a result of inhibiting the overall activ­ity of the pathway.245 The ADH6 gene of S. cerevisiae encodes a reductase enzyme that reduces HMF and NADPH supports a specific activity of the enzyme some 150­fold higher than with NADH; yeast strains overexpressing this gene had a higher in vivo conversion rate of HMF in both aerobic and anaerobic cultures.246

A key trait for biofuels research is to enable growth in the presence of very high concentrations of both glucose and ethanol, the former in the batch medium (discussed in chapter 4, section 4.3), the latter accumulating as the fermentation proceeds. In yeasts, analysis of the effects of ethanol concentration and tempera­ture point to nonspecific actions on the cell membranes as leading to the variety of cellular responses, including reduced fermentation rate and viability.247 Adapting cells to ethanol causes changes in the cellular membrane composition, in particu­lar the degree of unsaturation of the fatty acids and the amount of sterols present, both indicative of alterations in the fluidity characteristics of the lipid-based mem­branes.248 The low tolerance of Pichia stipitis to ethanol can be correlated with a disruption of protein transport across the cell membrane, an energy-requiring pro­cess involving a membrane-localized ATPase; the system is activated by glucose, but ethanol uncouples ATP hydrolysis from protein transport, thus wasting energy for no purpose, and structural alteration of the ATPase to affect its sensitivity to ethanol could be a quick-win option for yeast ethanologens in general.249

In Z. mobilis, there is evidence for a glucose-sensing system to confer (or increase) the cells’ tolerance to the osmotic stress imposed by high sugar concentrations; the exact functions of the products of the four genes simultaneously transcribed in a glu­cose-regulated operon are unknown, but manipulating this genetic locus would aid adaptation to high concentrations (>100 g/l) of glucose in industrial media.250 The announcement of the complete sequencing of the genome of Z. mobilis by scientists in Seoul, South Korea, in 2005 provided rationalizations for features of the idio­syncratic carbohydrate metabolism in the absence of three key genes for the EMP/ tricarboxylic acid cycle paradigm, that is, 6-phosphofructokinase, 2-oxoglutarate dehydrogenase, and malate dehydrogenase.251 The nonoxidative pentose phosphate pathway is mostly missing, although the genes are present for the biosynthesis of phosphorylated ribose and thence histidine and nucleotides for both DNA and RNA (figure 3.12). The 2-Mbp circular chromosome encodes for 1,998 predicted func­tional genes; of these, nearly 20% showed no similarities to known genes, suggesting a high probability of coding sequences of industrial significance for ethanologens. In comparison to a strain of Z. mobilis with lower tolerance to ethanol and rates of ethanol production, glucose uptake, and specific growth rate, strain ZM4 contains 54 additional genes, including four transport proteins and two oxidoreductases, all potentially mediating the higher ethanol productivity of the strain. Moreover, two genes coding for capsular carbohydrate synthesis may be involved in generating an altered morphology more resistant to osmotic stress. Intriguingly, 25 of the “new” genes showed similarities to bacteriophage genes, indicating a horizontal transfer of genetic material via phages.251

Fructose

6-Phosphogluconate

2-Oxo-3-deoxy-6-phosphogluconate

Glyceraldehyde 3-phosphate

Ribulose 5-phosphate

1,3-Diphosphoglycerate

2-Phosphoglycerate

5-Phosphoribosyl pyrophosphate

Phosphoc^o/pyruvate

Oxaloacetate

Succinate

FIGURE 3.12 Fragmentary carbohydrate metabolism and interconversions in Z. mobilis as confirmed by whole-genome sequencing. (From Seo et al.251)

Analysis of mutants of the yeast Yarrowia lipolytica hypersensitive to acetic acid and ethanol identified a single gene that could direct multiple phenotypic changes, including colony morphology, the morphology of intracellular membranes, and the appearance of vacuoles and mitochondria, and induce early death on glucose even with­out the presence of acetate or ethanol; this GPR1 gene may be a target for manipulation
and mutation to increase solute tolerance.252 A more direct approach in S. cerevisiae involved multiple mutagenesis of a transcription factor and selection of dominant muta­tions that conferred increased ethanol tolerance and more efficient glucose conversion to ethanol: 20% for growth in the presence of an initial glucose concentration of 100 g/l, 69% for space-time yield (grams of ethanol per liter per hour), 41% for specific pro­ductivity, and 15% for ethanol yield from glucose.253 It is likely that the expression of an ensemble of hundreds of genes was increased by the mutations; of the 12 most over­expressed genes, deletions mutations abolished the increased glucose tolerance in all but one gene, and selective overexpression of each of the top three up-regulated genes could replicate the tolerance of the best mutant. Because this work was, by necessity, carried out on a laboratory haploid strain, it is possible that similar changes could have occurred gradually during many generations in industrial strains. The effect on gene expression was, however, greatly diminished at 120 g/l of glucose.

Comparable analyses of gene expression and function are now being introduced into commercial potable alcohol research. A common driver between breweries and industrial bioethanol plants is (or will be) the use of concentrated, osmotically stressful fermentation media. In breweries, the capability to use high-gravity brewing in which the wort is more concentrated than in traditional practice saves energy, time, and space; the ethanol-containing spent wort can be diluted either with lower alcohol-content wort or with water. Variants of brewer’s yeast (S. pastorianus) were obtained subjecting UV- mutagenized cells to consecutive rounds of fermentation in very-high-gravity wort; variants showing faster fermentation times and/or more complete sugar utilization were identified for investigations of gene expression with microarray technologies.254 Of the 13 genes either overexpressed or underexpressed in comparison with a control strain, a hexokinase gene was potentially significant as signaling a reduced carbon catabolite repression of maltose and maltotriose uptake; more speculatively, two ampli­fied genes in amino acid biosynthesis pathways could point to increased requirements for amino acid production for growth in the highly concentrated medium. Although deletion mutants are usually difficult to isolate with polyploid industrial strains, the dose-variable amplification of target genes is more feasible.

Strains of S. cerevisiae genetically engineered for xylose utilization depend, unlike naturally xylose-consuming yeasts, on xylose uptake via the endogenous hexose transporters; with a high XR activity, xylose uptake by this ad hoc arrange­ment limits the pentose utilization rate at low xylose concentrations, suggesting that expression of a yeast xylose/pentose transporter in S. cerevisiae would be beneficial for the kinetics and extent of xylose consumption from hemicellulose hydrolysates.255 A high-affinity xylose transporter is present in the cell membranes of the efficient xylose fermenting yeast Candida succiphila when grown on xylose and in Kluyvero — myces marxianus when grown on glucose under fully aerobic conditions.256

In ethanologenic bacteria, there are also grounds for postulating that radically chang­ing the mechanism of xylose uptake would improve both anaerobic growth and xylose utilization. Mutants of E. coli lacking PFL or acetate kinase activity fail to grow anaero­bically on xylose, that is, the cells are energy-limited without the means to generate ATP from acetyl phosphate; in contrast, the mutants can grow anaerobically on arabinose — and the difference resides in the sugar transport mechanisms: energy-expending active transport for xylose but an energy-conserving arabinose.257 Reconfiguring xylose uptake

is, therefore, an attractive option for removing energetic constraints on xylose metabo­lism. In yeast also, devising means of conserving energy during xylose metabolism would reduce the bioenergetic problems posed by xylose catabolism — gene expres­sion profiles during xylose consumption resemble those of the starvation response, and xylose appears to induce metabolic behavioral responses that mix features of those elic­ited by genuinely fermentable and respirative carbon sources.258

Looking beyond purely monomeric sugar substrates, the soil bacterium Geobacil­lus stearothermophilus is highly efficient at degrading hemicelluloses, possessing a 30- gene array for this purpose, organized into nine transcriptional units; in the presence of a xylan, endo-1,4-P-xylanase is secreted, and the resulting xylooligosaccharides enter the cell by a specialized transport system, now known to be a three-gene unit encod­ing an ATP-binding cassette transport system, whose transcription is repressed by xylose but activated by a response regulator.259 This regulation allows the cell to rapidly amplify the expression of the transport system when substrates are available. Inside the cell, the oligosaccharides are hydrolyzed to monomeric sugars by the actions of a fam­ily of xylanases, P-xylosidase, xylan acetylesterases, arabinofuranosidases, and gluc­uronidases. Such a complete biochemical toolkit for hemicellulose degradation should be considered for heterologous expression in bacterial bioethanol producers.

Control of the gene encoding the major PDC in wild-type S. cerevisiae was shown by analysis of gene transcription to be the result of ethanol repression rather than glucose induction; an ethanol-repression ERA sequence was identified in the yeast DNA; the Era protein binds to the PDC1 promoter sequence under all growth conditions, and an autoregulatory (A) factor stimulates PDC1 transcription but also binds to the pdc1 enzyme — the more enzyme is formed, the more A factor is bound, thus making the transcription of the FDC1 gene self-regulating.260 This may be a useful target for strain improvement; industrial strains may, however, have already been “blindly” selected at this locus.

Focusing on single-gene traits, a research group at Tianjin University (China) has recently demonstrated three approaches to reduce glycerol, acetate, and pyruvate accumulation during ethanol production by S. cerevisiae:

1. Simply deleting the gene encoding glycerol formation from glycolysis, that is, glycerol 3-phosphate dehydrogenase reduced the growth rate compared with the wild type but reduced acetate and pyruvate accumulation. Simul­taneously overexpressing the GLTl gene for glutamate synthase (catalyzing the reductive formation of glutamate from glutamine and 2-oxo-glutarate) restored the growth characteristics while reducing glycerol accumulation and increasing ethanol formation.261

2. Deleting the gene (FSP1) encoding the transporter protein for glycerol also improved ethanol production while decreasing the accumulation of glycerol, acetate, and pyruvate—presumably increasing the intracellular concentration of glycerol (by blocking its export) accelerates its metabolism by a reversal of the glycerol kinase and glycerol 3-phosphate dehydrogenase reactions.262

3. Improved production of ethanol was also achieved by deleting FPS1 and overexpressing GLTl, accompanied by reduced accumulation of glycerol, acetate, and pyruvate.263

A more subtle metabolic conundrum may limit ethanol accumulation in Z. mobilis: although the cells are internally “unstructured” prokaryotes, lacking membrane — limited intracellular organelles, different redox microenvironments may occur in which two forms of ADH (rather than one) could simultaneously catalyze ethanol synthesis and oxidation, setting up a futile cycle; this has only been demonstrated under aerobic growth conditions, but the functions of the two ADH enzymes (one Zn-dependent, the other Fe-dependent) inside putative redox microcompartments within the cell warrant further investigation.264

S. cerevisiae, along with E. coli, B. subtilis, and a few other organisms, formed the initial “Rosetta Stone” for genome research, providing a well-characterized group of genes for comparative sequence analysis; coding sequences of other organisms lacking detailed knowledge of their biochemistry could be assigned protein func­tionalities based on their degree of homology to known genes. Genome-scale meta­bolic networks can now be reconstructed and their properties examined in computer simulations of biotechnological processes. Such models can certainly generate good fits to experimental data for the rate of growth, glucose utilization, and (under micro — aerobiosis) ethanol formation but have limited predictive value.265 One limitation is that the biosynthetic genes comprise too conservative a functional array, lacking the sensing mechanisms and signaling pathways that sense and respond to nutri­tional status and other environmental factors.266 These elements function to estab­lish developmental and morphological programs in the wild-type ecology and have unpredictable effects on growth patterns in the fluctuating nutritional topologies of large imperfectly mixed industrial fermentors. If computer-aided design for metabo­lism is to be feasible for industrially relevant microbes, added sophistications must be incorporated — equally, however, radical restructuring is necessary to accommo­date the gross differences in metabolic patterns between bacteria such as E. coli and, for example, the Entner-Doudoroff pathway organisms or when P. stipitis metabo­lism is compared in detail with the far better characterized pathway dynamics in S. cerevisiae.267,268 Comparing metabolic profiles in 14 yeast species revealed many important similarities with S. cerevisiae, but Pichia augusta may possess the ability to radically rebalance the growth demand for NADPH from the pentose phosphate pathway (figure 3.2); this suggests a transhydrogenase activity that could be used to rebalance NADH/NADPH redox cofactors for xylose metabolism (section 3.2.1).269

In the immediate and short-term future, therefore, applied genomics can suc­cessfully define the impacts of more focused changes in quantitative gene expression. Developing a kinetic model of the pentose phosphate and Entner-Doudoroff pathways in Z. mobilis, and comparing the predictions with experimental results, showed that the activities of xylose isomerase and transaldolase were of the greatest significance for ethanol production but that overexpressing the enzymic link between the two pathways (phosphoglucose isomerase) was unnecessary.270 Such systematic analysis can help not only in maximizing fermentation efficiencies but also in minimizing the degree of expression of heterologous enzymes to reduce “metabolic burdens” in recombinant strains. Although in silico predictions failed to account for contin­ued xylitol accumulation by a recombinant, xylose-utilizing S. cerevisiae strain on xylose (suggesting continued problems in balancing cofactor requirement in reduc — tase/dehydrogenase reactions and/or still unidentified limitations to later catabolic steps in xylose consumption), computer simulations did accurately highlight glycerol formation as a key target for process improvement: expressing a nonphosphorylating NADPH-dependent glyceraldehyde 3-phosphate dehydrogenase increased carbon flow to pyruvate (rather than being diverted to glycerol formation), reduced glycerol accumulation, and improved ethanol production by up to 25%.271272