4.3.1

Sorbitol/Gluconate Production

The production of sorbitol by Z. mobilis when grown on sucrose or a mixture of glucose and fructose has been reported earlier by several groups [94,95]. In subsequent studies on the mechanism of sorbitol production, an enzyme complex was identified by Leigh et al. [96] which was capable of oxidizing glucose to gluconic acid concomitant with the reduction of fructose to sor­bitol. This enzyme was described as a glucose-fructose oxidoreductase with a tightly coupled (non-dialyzable) co-factor identified as NADP [97]. The mechanism for sorbitol/gluconic acid production and the associated enzymes are shown in Fig. 8 with the pathway from gluconate to ethanol not being functional if cells of Z. mobilis are fully permeabilized. As shown in Fig. 8, the possibility exists also of producing a mixture of sorbitol and gluconolactone if gluconolactonase activity is deleted.

Kinetic studies have been reported for a 60% sugar solution (300 gL-1 glucose and 300 g L-1 fructose) using toluene-treated permeabilized cells of Z. mobilis in which a sorbitol concentration of 290 g L-1 and a gluconic acid concentration of 283 gL-1 were achieved after 15 h in a batch process [98]. A continuous process with immobilized cells was developed with only a small loss of enzyme activity (less than 5%) evident after 120 h. With a strongly basic anion exchange resin and a buffer system at pH = 9.0, good separation of sorbitol and gluconic acid was achieved. Subsequent studies using immo-

4.3.2

The performance of the thermostable enzyme mixtures was studied in hydro­lysis experiment in test tubes (5 mL). Enzyme mixtures were dosed on the basis of FPU activity to the substrates (10 gL-1 dry matter) suspended in 50 mM sodium acetate, pH 5. The standard enzyme dosage was 10 FPUg-1 cellulose. Triplicate samples were incubated with mixing at 35 °C, 45 °C, 55 °C or 60 °C for 24 h, 48 h or 72 h. Reference samples with inactivated enzymes and corresponding substrates were also prepared.

Chemical Analysis

The release of hydrolysis products was measured as reducing sugars assayed by the DNS method using glucose as standard [10]. The results were cor­rected by taking into account the blank samples containing corresponding amounts of inactivated enzymes and substrate. The mono — and oligosac­charides formed were also analysed by high-performance anion-exchange chromatography on a Dionex 4500i series chromatograph with pulsed amper — ometric detection (HPAEC-PAD), as described earlier [75].

6

In contrast to S. cerevisiae, many yeast species are capable of utilising xy­lose as the sole carbon and energy source for respiratory growth. However, only few of these yeasts are capable of fermenting xylose to ethanol under oxygen-limited conditions, such as for instance Pichia stipitis and Pachysolen tannophilus [65].

Maybe not surprisingly, xylose-metabolising yeasts have predominantly been isolated from wood-related environments. The pathway for D-xylose metabolism used by these yeasts to convert D-xylose to D-xylulose was first described in 1955 [25] and involves a two-step conversion that involves two oxidoreductases (Fig. 1): xylose reductase (EC 1.1.1.21) and xylitol dehydro­genase (EC 1.1.1.9). The xylose reductase has a strong preference for NADPH, whereas the subsequent oxidation of xylitol via xylitol dehydrogenase pro­duces NADH (Table 1).

Clearly, this difference in cofactor specificity can result in redox imbalance. To generate the NADPH for the xylose reductase reaction, part of the D-xylose carbon must be directed through the oxidative pentose phosphate pathway (involving the glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase reactions). While this results in a loss of some carbon as CO2,

which goes at the expense of the ethanol yield on D-xylose, it enables the efficient regeneration ofNADPH [16,32,45,69].

However, the cells have to take additional measures to reoxidise the ex­cess NADH generated in the xylitol dehydrogenase reaction. In the presence of oxygen, this excess NADH can be reoxidised by respiration. This will re­quire accurate dosage of oxygen to prevent full respiration of D-xylose. Such accurate control is difficult to envisage in large-scale processes for ethanol production, which should preferably involve a minimum of aeration to reduce costs.

Under anaerobic conditions, reoxidation of excess NADH can be ac­complished via the production of compounds that are more reduced than D-xylose, such as xylitol and/or glycerol. The production of xylitol occurs via xylose reductases, which have a dual co-enzyme specificity and thereby can also use NADH, or alternatively via other aspecific reductases. As this mech­anism involves the consumption of one D-xylose for each NADH generated, it has a tremendously negative impact on the ethanol yield from D-xylose [45]. Glycerol production is a well-known redox sink during hexose fermenta­tion and especially under anaerobic conditions, but requires both carbon and ATP [67].

The preference of xylose reductase for NADPH is not only species — but also strain-dependent (Table 1). The in vivo ratio ofNADPH over NADH utilisa­tion by xylose reductase and the redox balance requirements determine the

requirement for NADH sinks such as xylitol and glycerol (Fig. 2) in anaerobic cultures [14,69]. When this NADPH/NADH ratio equals zero, xylose reduc­tase only uses NADH and thereby consumes all NADH produced in the xylitol dehydrogenase reaction. Since in addition no regeneration of NADPH is re­quired for the xylose reductase reaction, redox-balanced xylose metabolism will occur according to Eq. 1:

Ratio = 0: 6 xylose ^ 10 ethanol + 10 CO2 + 10 ATP. (1)

At a ratio of one (Eq. 2), one out of every two D-xylose molecules can be further metabolised to ethanol, whereas the other is reduced to xylitol to maintain NADH balance. In addition, some carbon has to be redirected for the generation of NADPH, resulting in the formation of only 9 mol of ethanol from 12 mol of D-xylose (45% of the theoretical yield). Following these redox-balance considerations, catabolism via a xylose reductase with a NADPH/NADH-utilisation ratio of one will follow:

Ratio = 1: 12 xylose ^ 9 ethanol + 12 CO2 + 9 ATP + 6 xylitol. (2)

At ratios above one, NADH-dependent xylitol formation cannot compensate for the production of NADH in the xylitol dehydrogenase reaction and glycerol formation becomes essential for redox balancing [32,45,69]. When the xylose

reductase solely uses NADPH (an infinite NADPH/NADH ratio) this would result in the formation of only 0.5 mol ethanol per mol of xylose fermented.

Ratio = to : 6 xylose + 3 ATP ^ 3 ethanol + 6 glycerol + 6 CO2 . (3)

Despite these inherent redox restrictions and ensuing loss of ethanol yield on D-xylose, the expression of xylose reductase and xylitol dehydrogenase has long been the most successful strategy to enable D-xylose consumption by S. cerevisiae (elsewhere in this volume, and [29,32,33,39,63]). Although attempts have been made to change the cofactor specificity of xylose reduc­tase, fermentation properties of a S. cerevisiae strain containing this gene are not available [55]. Similarly, expression of a transhydrogenase in S. cere­visiae, with the aim of converting excess NADH into NADPH, did not result in reduced byproduct formation [51]. The latter result is perhaps not alto­gether surprising as, with NADPH/NADP+ ratios generally being higher than NADH/NAD+ ratios [51], reduction of NADP+ with NADH is thermodynam­ically unfavourable.

Despite the inherent redox constraints of S. cerevisiae strains based on the xylose reductase/xylitol dehydrogenase strategy, this strategy has resulted in many important insights into the kinetics of D-xylose metabolism by en­gineered S. cerevisiae strains. These findings include the benefits of over­expression of xylulokinase [29, 56], the side role of the S. cerevisiae aldose reductase (Gre3) (besides the heterologous dual specificity xylose reduc­tases) in xylitol formation [66], the role of the enzymes of the non-oxidative part of the pentose phosphate pathway [34,43], characterisation of D-xylose transport [27,62] and many studies on the inhibitor tolerance/sensitivity of D-xylose-consuming strains [54]. The latter will be especially crucial for suc­cessful application of D-xylose-consuming S. cerevisiae strains for ethanol production from lignocellulosic hydrolysates (see Sect. 7).

1.3

In order to attain the desired high product titers, biocatalysts must be sup­plied with high levels of sugars. These high sugar levels in turn create osmotic stress, which is compounded by the desire to use simple mineral salts media. Osmolytes such as trehalose, betaine, proline, and glutamate help bacteria maintain appropriate cell turgor and volume despite changes in extracellu­lar osmolality; osmolyte uptake and synthesis are reviewed in [80]. Increased activity of the native trehalose synthesis pathway elevated the growth rate of E. coli W3110 in the presence of various osmotic stress agents [81], and betaine supplementation increased the production of D-lactic acid by E. coli SZ132 in NBS mineral salts media [42]. A combination of betaine supple­mentation and elevated trehalose synthesis increased the tolerance of W3110 to xylose, glucose, sodium lactate, and sodium chloride more than betaine supplementation or elevated trehalose synthesis alone [82].

As described above, the poor performance of ethanologenic E. coli strain KO11 in minimal media has been attributed to NADH-mediated inhibition of citrate synthase, limiting the availability of glutamate, a protective os — molyte [33]. Additionally, the increased performance of LY01 relative to KO11 can be partly attributed to increased osmolyte production and uptake [83]. NMR analysis confirmed that intracellular pools of glutamate, trehalose, and betaine are very low in KO11 during anaerobic growth relative to aerobic growth in the same medium [84]. Growth and ethanol production of KO11 was increased by supplementation with various osmolytes, demonstrating that the glutamate limitation is related to osmotic stress, not to a specific metabolic demand for glutamate [84].

4.3

A further development of biorefineries is the use of hybrid techniques com­bining biological conversion with catalytic downstream processing (Wester — mann P, J0rgensen B, Lange L, Ahring BK, Christensen CH (2007) Int J Hydro­gen Energy, accepted for publication). For instance, highly efficient autother­mal reformers capable of converting 1 mol ethanol to 5 mol hydrogen have recently been demonstrated [27]. Since 2 mol of ethanol can be achieved for each sugar molecule, the hydrogen yield of this two-step process is 83% of the theoretical maximum, compared to the 10-20% achieved by direct hydrogen fermentation. Hydrogen produced in the thermophilic ethanol fermentation process described above would add to this yield, approaching the theoretical maximum yield of 12 mol hydrogen/mol monosaccharide.

Hydrogen has been suggested as a future energy carrier to succeed the fossil fuel era [28]. The introduction of downstream catalytic conversion of biofuels leaves the possibility of combining a less complex fuel handling technology (ethanol instead of hydrogen) for transportation purposes with all the benefits of the fuel cell technology. In the transition period before a hydrogen-based energy economy has been realized, a gradual change to the use of renewable energy can be facilitated by the use of catalytically con­verted biofuels in existing internal combustion engines. Although ethanol in even high ethanol:gasoline mixtures can be used for ground transportation with few modifications of the engines, biogasoline produced by catalytic con­version of methane and bioethanol will have potential use as a high energy alternative for aviation and air transport. If these transportation means are sustained in the future, the availability of safe liquid fuels with high energy content storable under ambient conditions is a prerequisite.

4.7

Lignocellulosic raw materials contain much less L-arabinose than D-xylose, and solving the problem of xylose fermentation has been prioritized. The relative amounts of the sugars strongly depend on the raw material. For example, corn stover contains of 19% xylan and 3% arabinan, whereas wheat bran contains 19% xylan and 15% arabinan [58]. As a consequence, L-arabinose-utilizing strains of S. cerevisiae have been developed only re­cently. Furthermore, the conversion of L-arabinose into intermediates of the PPP requires more enzymatic reactions than the conversion of xy­lose (Fig. 2). In many bacteria, such as E. coli, L-arabinose is utilized via L-arabinose isomerase (AraA), L-ribulokinase (AraB), and L-ribulose-5- phosphate 4-epimerase (AraD) [59]. Xylulose-5-phosphate is then further metabolized via the PPP. Enzymatic activities of alternative bacterial arabi — nose and xylose utilization pathways have also been described [60-62].

The fungal arabinose utilization pathway consists of four alternating reduction-oxidation reactions (Fig. 2), where L-arabinose is converted to

D-xylitol via L-arabi(ni)tol and L-xylulose [23,63-65]. D-Xylitol is then further metabolized by XDH and XK, resulting in the PPP intermediate D-xylulose-5-phosphate. The first two complete fungal arabinose utilization pathways were recently kinetically characterized for Candida arabinofermen — tans PYCC 5603r and Pichia guilliermondii PYCC 3012 [65]. The fungal xylose and arabinose utilization pathways share the enzymes XR, XDH, and XK, since XR also reduces L-arabinose [65-68]. Indeed, all arabinose-utilizing yeast and fungi also utilize xylose, whereas not all xylose-growing yeasts uti­lize arabinose [66,69]. Similar to the fungal xylose pathway, the cofactors of the enzymes in the fungal arabinose pathway cannot be regenerated within the pathway but require oxygen or an external electron acceptor for regener­ation (Fig. 2).

3.2

Corn stover is another potentially interesting feedstock for ethanol produc­tion, especially in the USA. The fermentation characteristics of S. cerevisiae RWB 218 on corn stover hydrolysate were tested under industrially relevant fed-batch conditions (W. de Laat, unpublished data). The corn stover pulp obtained after steam explosion (190 ° C, 5 min, ENEA, Italy) was diluted with water to 150gL-1 dry matter and subsequently hydrolysed with 10g cellu — lase protein (kg hydrolysate dry matter)-1 (GC220, Genencor, 96 h at 50 °C). After filtration, the resulting sugar solution contained 40 g L-1 glucose, 9 g L-1 D-xylose and 4 g L-1 acetic acid.

Fermentation experiments were initiated by a 32 h batch phase on mo­lasses medium (containing 100gL-1 sucrose, pH 4.8, 32°C) in a volume of 200 mL. Subsequently, 455 mL of corn stover hydrolysate was added during a 16 h fed-batch phase. During the fed-batch phase, glucose was almost com­pletely consumed. However, only 40% of the D-xylose fed to the culture was consumed during this phase (Fig. 9). After 16 h, the fed-batch phase was terminated and the culture was allowed to consume accumulated sugars. Con­version was complete after 35 h. At a biomass concentration of 1.0—1.5 gL-1, this corresponded to a D-xylose fermentation rate of 0.5 mmolg-1 h-1 dur­ing this latter phase. The overall ethanol yield on total sugars was 0.46 gg-1, which corresponds to 90% of the theoretical maximum yield on glucose and D-xylose. Consistent with the wheat straw hydrolysate fermentations, xylitol formation was not observed.

8

2.1

Increased Substrate Range Through Expression of a Single Heterologous Gene

One of the possible disadvantages of Z. mobilis is that it has a limited carbon substrate range as it can only use the simple C6 sugars glucose, fructose and sucrose. As a result early studies on its genetic manipulation focused on ex­tending its substrate range for ethanol production. Skotnicki et al. [18] first reported high frequency conjugal transfer of plasmids from Escherichia coli and Pseudomonas aeruginosa, and this was followed by expression of the lac Z gene and production of ^-galactosidase in strains of Z. mobilis [21,22]. How­ever, the strain ZM6100 (RP1:Tn 951) derived from this work was shown to progressively lose all plasmid markers in batch culture under non-selective conditions. Subsequently a new strain, ZM6306, was developed in contin­uous culture which showed 100% stability for all plasmid markers when grown without selection pressure. Synthesis of ^-galactosidase was induced in continuous culture by addition of lactose resulting in increased ethanol production and unutilized galactose [23].

Further studies to extend the substrate range were reported which involved the cloning and expression of a в-glucosidase gene from Xanthomonas al — bineans [24] and a-glucosidase gene from a Bacillus sp [44], however enzyme expression levels were low.

2.2

The second series of data illustrated in Fig. 1 shows that development of the bioethanol industry in the USA began in the 1980s. The drivers for the in­dustry were in part the rapid surges in global oil prices experienced in the 1970s and 1980s, which led to rising prices of fuel. There was also the presence of a strong agricultural lobby which was (and is) interested in creating ad­ditional revenue streams for farmers. The US bioethanol industry uses corn, and to a lesser extent wheat, as a feedstock for wet — and dry-milling processes. A number of different policy options have been employed to help build the industry. Both federal and state governments have offered the industry dir­ect funding in the form of public-private partnerships and research funds, as well as tax incentives and state-level renewable fuel mandates, i. e., legislated amounts of renewable fuels contained in fuel sales within the state, defined by blending level or by renewable fuels [22,23]. A more focused discussion of state-level funding and tax incentives, and the effectiveness of these options, may be found in Sects. 3 and 4, respectively.

In the USA, most bioethanol production capacity is concentrated in the Midwest, where corn is found in abundance, and where state and federal government incentives have combined to make an attractive en­vironment for investment in the infrastructure required for bioethanol production. Over half of US production capacity is found in just three states, each of which have supplied significant capital resources to the bioethanol industry. The US states with the highest bioethanol capacities include Illinois (annual bioethanol production capacity, 5.1 billion L), Iowa (3.7 billion L), South Dakota (2.2 billion L), Minnesota (1.9 billion L), and Ne­braska (1.8billionL) [18]. These states are notable in that they have provided direct funding incentives in addition to federal funding, as discussed in Sect. 3.

The total financial commitment that the USA has made to biofuels dwarfs the investment that other countries have made. By 2006, total cumulative US funding through national or state programs applicable to bioethanol has ex­ceeded US $ 2.5 billion [23]. The largest amount of funding has been offered by the federal government. Annual program spending by all government agencies, primarily the US Department of Agriculture and the US Depart­ment of Energy, on alternative fuels exceeded US$ 253 million in 1998 and has risen since to more than US$ 300 million [18,24]. This has resulted in improving the technology that is utilized by the industry, and has broadened the potential number of coproducts that can be generated from the bioethanol production process. The remainder of federal funds supports a number of in­centive programs, including the Alcohol Fuel Credit (a corporate tax credit designated for industry producing bioethanol), deductions for both clean — fuel vehicles and refueling properties, and the Renewable Energy Systems and Energy Efficiency Improvements Program. The latter program is designed to aid in the construction of new facilities, and will cover up to 25% of con­struction costs. Maximum grants for a single project under this program are US $ 500 000, and the fund generally pays out between US $ 3-5 million in any given year [22,23]. Finally, it should be pointed out that significant funding in the USA has been directed towards developing cost-effective coproducts from the biofuel production process, allowing the creation of “biorefineries” with improved economic and environmental performance. Pilot facilities are already operating under some of these funding programs [23].

Most recent policy developments in the USA stem from the Energy Pol­icy Act of 2005, H. R. 6, which was signed into law by President G. W. Bush on 8 August 2005 [25]. This act created a nationwide renewable fuels standard (RFS) that will raise the use of biofuels (mostly bioethanol and biodiesel) to 28.4 billion L year-1 by 2012, which is effectively 5% of the total fuel sales. The Act also introduced credits for the purchase or lease of flex-fuel vehicles by taxpayers, although these credits diminish as the sales of flex-fuel vehicles progress by manufacturer through the fiscal year [25]. The 2005 Energy Policy Act has had some unintended consequences as related to biofuels, however. Section 701 of the Act requires flex-fuel vehicles in the US federal fleet to op­erate on alternative fuels 100% of the time. By Executive Order 13149, federal flex-fuel vehicles were previously required to operate on alternative fuels the majority of the time (i. e., 51% or more) [26]. Thus, Section 701 has effectively doubled E85 use by the federal fleet, and the increased demand has raised prices and decreased the practical availability of E85 fuels. The long-term impact of this policy on the market has yet to be seen.

The recently-announced “20/20” vision for biofuels (introduced as a Sen­ate Bill on 29 July 2005) defines a future biofuel production goal for the USA as 20 billion gal (approximately 75.7 billion L) by 2020 [27]. As the US starch — based bioethanol capacity is already quite high, it is unlikely that continued growth could achieve this goal. Accordingly, in his State of the Union Address for 2006, the President outlined the Advanced Energy Initiative, which seeks to reduce US dependence on imported oil by accelerating the development of new, renewable alternatives to gasoline and diesel fuels [28]. These alter­natives include bioethanol and other future biofuels derived from cellulosic biomass. Cellulosic biomass is an attractive energy feedstock because it is an abundant, domestic, renewable source that can be converted to liquid trans­portation fuels including bioethanol, which can be used readily by current — generation vehicles and distributed through the existing transportation-fuel infrastructure. To determine feedstock availability for cellulosic bioethanol processes, the US Department of Agriculture commissioned a report that explored the technical feasibility of a billion-tonne annual supply. This re­port found that approximately 1.24 billion t of dry cellulosic biomass can be sustainably produced each year, with about 910 million t coming from agri­culture and an additional 330 million t from the forest sector [29]. Using the efficiency of conversion technologies observed in the literature to date [6], this would translate to between 110 and 250 million L year-1, compared to current US gasoline use of approximately 500 million L year-1.

US production of biofuels is significant, but today only comprises about 2.6% of liquid fuel consumption. In order to become a more significant component of the transportation fuel sector, biofuel production must grow tremendously, which will require access to cellulosic biomass. The Advanced Energy Initiative includes the Biorefinery Initiative, which sets a goal of mak­ing cellulosic bioethanol cost-competitive by 2012 and which provides signifi­cant funding to achieve this goal (US $ 91 million in 2006, US $ 150 million in 2007) [30]. Biorefining pilot facilities are already operating with starch — based feedstocks, and these processes have the potential to be applied to cellulose-based biofuel production facilities, which will contribute to the eco­nomic viability of these operations. If these measures are successful, cellu — losic bioethanol production could easily become the dominant biofuel within the USA.

2.3

The problem of cofactor regeneration has also been addressed by engineering reactions distant from the xylose utilization pathway, as demonstrated by dif­ferent approaches to introduce a transhydrogenase function in S. cerevisiae. Heterologous expression of a bacterial transhydrogenase [118] in S. cerevisiae carrying XR and XDH reduced xylitol formation, but also increased glycerol, rather than ethanol formation [116] (strain TMB3254, Table 2), indicating that the transhydrogenase reaction did not proceed in the direction favorable for ethanolic xylose fermentation [116,118].

Intracellular cofactor concentrations have also been altered, introduc­ing in S. cerevisiae the NAD(P)+-dependent glyceraldehyde-3-phosphate dehydrogenase (GAPDH) from Kluyveromyces lactis [119] (strain H2673, Table 1). When the ZWF1 gene was simultaneously deleted, the expression of GAPDH improved ethanol formation [115] (strain H2684, Table 1). Similarly, when a NAD(P)+-dependent nonphosphorylating GAPDH from Streptococ­cus mutans was overexpressed in an XR-XDH-XK-carrying strain, increased ethanol formation was observed [120] (strain CPBCB4, Table 3). The result suggested that less carbon was lost as carbon dioxide when NADPH was formed outside the oxidative PPP and that NAD+ consumption in the lower glycolysis was simultaneously reduced.

Engineering the ammonium assimilation pathway [121] has also been used to modify the intracellular cofactor concentrations. Based on the as­sumption that NADH would be used for ammonium assimilation to generate NAD+ for the XDH reaction, the NADPH-dependent glutamate dehydroge­nase gene GDH1 was deleted, and an NADH-dependent isoenzyme (GDH2) was overexpressed. Reduced xylitol formation and higher ethanol forma­tion were observed [121] (strain CPB. CR4, Tables 1 and 3). Alternatively, the GS-GOGAT complex coded by the genes GLT1 and GLN1 was overex­pressed, which only affected xylose fermentation in a continuous fermenta­tion setup [121] (strain CPB. CR5, Tables 1 and 3).

4.6