Mats Galbe1 • Per Sassner1 • Anders Wingren2 • Guido Zacchi1 (И)

department of Chemical Engineering, Lund University, P. O. Box 124, 221 00 Lund, Sweden

Guido. Zacchi@chemeng. lth. se

2SEKAB E-Technology, P. O. Box 286, 891 26 Ornskoldsvik, Sweden

1 Introduction……………………………………………………………………………………………… 304

2 Flowsheeting……………………………………………………………………………………………… 309

2.1 Simulation of Ethanol Production from Lignocellulosic Materials…………………. 310

3 Process Economics…………………………………………………………………………………….. 311

3.1 Effect of Various Parameters on the Energy Demand and Production Cost 318

3.2 Lignocellulose versus Starch—a Comparison……………………………….. 322

3.3 Co-location with other Plants…………………………………………………………………….. 325

4 Conclusions………………………………………………………………………………………………. 325

References……………………………………………………………………………………………………. 326

Abstract This work presents a review of studies on the process economics of ethanol production from lignocellulosic materials published since 1996. Our objective was to identify the most costly process steps and the impact of various parameters on the fi­nal production cost, e. g. plant capacity, raw material cost, and overall product yield, as well as process configuration. The variation in estimated ethanol production cost is considerable, ranging from about 0.13 to 0.81 US$ per liter ethanol. This can be ex­plained to a large extent by actual process differences and variations in the assumptions underlying the techno-economic evaluations. The most important parameters for the economic outcome are the feedstock cost, which varied between 30 and 90 US$ per metric ton in the papers studied, and the plant capacity, which influences the capi­tal cost. To reduce the ethanol production cost it is necessary to reach high ethanol yields, as well as a high ethanol concentration during fermentation, to be able to de­crease the energy required for distillation and other downstream process steps. Improved pretreatment methods, enhanced enzymatic hydrolysis with cheaper and more effective enzymes, as well as improved fermentation systems present major research challenges if we are to make lignocellulose-based ethanol production competitive with sugar — and starch-based ethanol. Process integration, either internally or externally with other types of plants, e. g. heat and power plants, also offers a way of reducing the final ethanol production cost.

Keywords Bioethanol production • Biomass • Flowsheeting • Process economics

1

Introduction

There is no single process design offering the most cost-efficient way to pro­duce ethanol from biomass. Many factors that affect the desired product have to be taken into consideration. Regarding ethanol production, some of the most important parameters are the capital cost of the plant, the type and cost of raw material, the utilization efficiency of the materials involved in the pro­cess and the energy demand. The design of the plant, as well as its individual process steps, must be based on accurate and reliable data. These comprise both physical and chemical data and cost estimation data. It is naturally best to use data gathered from the same or a similar type of plant as the intended one. Most of the data required are available, or can be adapted and used for a new plant design. This is not the situation when lignocellulosic materials are considered as feedstock for ethanol production.

Ethanol has traditionally been produced from sugar cane and sugar beet juice [1] or from various starch-containing materials, e. g. corn or wheat [2-4]. Figure 1 shows a simplified flowsheet of an ethanol produc­tion process based on starch-containing materials. Liquefaction of the starch fraction is accomplished by adding hydrolytic enzymes (a-amylases) at tem­peratures of around 90 ° C. After the liquefaction step the starch molecules are further hydrolyzed by the addition of glucoamylases. This produces sug­ars, which are readily fermented by yeast, e. g. Saccharomyces cerevisiae, to ethanol. The main co-product is usually animal feed, consisting of the re­maining fraction of the raw material, mainly proteins and fiber, which is sometimes referred to as DDGS—distillers dried grains with solubles [5]. There is considerable experience in starch-based ethanol production, and the technology can be considered mature. The design and cost estimates of new plants are, therefore, rather accurate.

The availability of agricultural land for non-food crops and the limited market for animal feed places a limit on the amount of ethanol that can be produced from starch-based materials in a cost competitive way [6]. Ethanol production from lignocellulosic raw materials, on the other hand, reduces the potential conflict between land use for food (and feed) production and energy feedstock production. The raw material is less expensive than conventional agricultural feedstock and can be produced with lower inputs of fertilizers, pesticides, and energy. Lignocellulosic materials contain about 50-60% car­bohydrates in the form of cellulose (made up of glucose) and hemicellulose (consisting of various pentose and hexose sugars), which may be fermented to ethanol, and 20-35% lignin. The latter is the main co-product, which could be used for the production of heat and electricity or, in the longer perspective, for the production of specialty chemicals. There is thus no co-product limita­tion on the use of lignocellulosic materials for ethanol production. The only limitation is the availability of the raw material and, of course, the production

cost. During recent years, there has been a considerable increase in interest in research on and the development of the conversion to ethanol of lignocellu — losic materials, such as agricultural and forest residues, as well as dedicated energy crops.

However, in contrast to starch-containing materials, cellulose-containing raw materials, such as forest residues and straw, have not yet been commer­cialized in the ethanol industry. The reasons for this are several. For instance, there are physical barriers such as:

• the complex structure of lignocellulosic materials, making them recalci­trant to hydrolysis;

• the presence of various hexose and pentose sugars in hemicellulose, mak­ing fermentation more difficult; and thirdly,

• the presence of various compounds that inhibit the fermenting organism. These compounds either originate from the raw material itself, e. g. ex­tractives, or are formed during the early process steps, e. g. degradation products of sugars and lignin. This makes it difficult to reach high ethanol concentrations during fermentation, which in turn results in a high energy demand and thus high production cost.

There is a big risk involved in being the first to invest in commercialization of a lignocellulose to ethanol plant and this is the main reason why there is no full-size plant in operation today.

Interest in lignocellulose-based ethanol production has recently brought about action on high political levels. For example, in the USA, the Energy Pol­icy Act of 2005 requires blending of 7.5 billion gallons (^ 28.4 million m3) of alternative fuels by 2012 [7] and recently, in his State of the Union Address (Jan 31, 2006), the US President announced the goal of replacing more than 75% of imported oil with alternative fuels by the year 2025 [8]. The major part of this alternative fuel will probably consist of ethanol, and to be able to meet these demands this will have to be largely produced from lignocellulosic materials. In Europe the European Commission plans to progressively replace 20% of conventional fossil fuels with alternative fuels in the transport sector by 2020, with an intermediate goal of 5.75% in 2010 [9]. Bioethanol is also expected to be one of the main means of achieving this goal.

Experience in the production of ethanol from lignocellulosic materials is limited, at least using modern technology. Full-scale plants have only been run occasionally during times of war. Examples are the Bergius process (con­centrated HCl) operated in Germany during World War II, and the Scholler process (dilute H2SO4), which was used in the former Soviet Union, Japan and Brazil [10]. Thus, design and cost estimation for lignocellulosic-based processes cannot be based on reliable operational experience, but data gath­ered on lab scale, or at best on pilot scale, must be used. It is true that some of the process steps are of the same type as in a starch-based process, but there are several major differences. For example, the by-products from the various processes are not the same. Some of these are considered valuable co­products, which will contribute to the profit from the process, while others are waste materials that must be dealt with in wastewater treatment plants, or disposed of by other means.

During the past 20 years or so, a great deal of effort has been devoted to research on various areas, such as the pretreatment of raw material, enzy­matic hydrolysis of cellulose, including the production of more cost-effective enzymes, and the development of new microorganisms and fermentation techniques to ferment all the sugars available in lignocellulosic materials. An enormous amount of data has been generated (see the work by Galbe, Vikarii, Cherry, Hahn Hagerdal, and Ingram, all in this volume), which today forms the basis for techno-economic calculations. However, although the re­sults may be accurate, there is still a huge scale-up problem involved in going from batch pretreatment reactors on the liter scale, to continuous reactors of several cubic meters, and from 1- to 100-liter fermentors to vessels with a vol­ume of 1000 cubic meters or more. Issues such as material corrosion, rapid heat evolution, excessive foaming, and precipitation of solids and incrusta­tion, which may not even be considered on the lab scale may become serious problems in a full-scale process.

Pilot-scale trials have been run in several places during the past decade. The National Renewable Energy Laboratory (NREL) (Golden, Colorado, USA) has constructed a pilot fermentation facility to test bioprocessing technolo­gies for the production of ethanol and other fuels or chemicals from cellulosic biomass [11]. The Process Development Unit (PDU) of the Bioethanol Pilot Plant was set up to investigate biomass fuel and chemical production pro­cesses from start to finish on a scale of about 900 kg day-1 of dry feedstock. The plant is, however, not a fully integrated unit that can run continuously.

A 1000 kg day-1 plant, using spruce as the raw material, has been in op­eration in Ornskoldsvik in Sweden since the middle of 2004 [12]. Abengoa Bioenergy Corp. has constructed a pilot plant in York, Nebraska, USA [13] and is now constructing a demonstration scale plant in Salamanca, Spain, with an annual production capacity of 5000 m3 ethanol. This will be brought into operation at the beginning of 2007 [14]. This demonstration plant, which will be co-located with a 195 000 m3 y-1 starch-based plant, will utilize the straw from wheat and thus contribute to the overall production capacity. Furthermore, Iogen Corp. is operating a pre-commercial demonstration fa­cility, located in Ottawa, Canada, where ethanol is made from agricultural residues [15]. The plant is able to handle up to 40 metric tons of feedstock daily, consisting of wheat, oat, and barley straw, and is designed to produce up to 3 million liters of ethanol annually.

Data from these types of plants will increase the reliability of cost estimates significantly. They can also be used to identify process problems associated with continuous processing, such as the accumulation of toxic substances in various process streams, and fouling of heat exchanger surfaces. However, in most cases this will be proprietary information not available in the scientific literature.

Two process concepts have been investigated more than others regarding ethanol production from lignocellulosic materials. The main difference be­tween the two is the way in which the cellulose chain is broken apart; either dilute sulfuric acid or cellulolytic enzymes are used to hydrolyze the cellulose molecules. Figure 2 shows the main features of a dilute acid hydrolysis pro­cess. The raw material is treated with 0.1-3% (w/w) H2SO4 at temperatures normally ranging from 160 to 200 °C. It may be advantageous to perform dilute-acid hydrolysis in two steps since the hemicellulose fraction is more easily degraded than is the cellulose fraction. A disadvantage of the dilute acid process is the somewhat low ethanol yield and the necessity of using ex­pensive construction materials that are resistant to corrosion by acid at high temperatures. The acid must also be neutralized, which leads to the forma­tion of large amounts of gypsum, CaSO4, or other compounds that have to be disposed of.

An alternative to acid hydrolysis is enzymatic hydrolysis (Fig. 3). Cellu­lolytic enzymes are produced by microorganisms and have the ability to cleave off short sugar units from the cellulose chain, as described in detail

by Vikarii 2007 and Merino 2007 (this volume). The enzymatic process is op­erated at much milder conditions than the dilute acid process, which is of great importance for several reasons. The yield can be expected to be higher, the construction materials will be less costly and the formation of toxic by­products will also be reduced. However, the enzyme action suffers from being slow if the raw material is not pretreated prior to enzymatic hydrolysis. Pre­treatment can be performed in a number a ways. Depending on the type of raw material (hardwood, softwood or agricultural residue) a certain pre-

treatment method can be more or less successful. Pretreatment is described in more detail by Galbe 2007 (this volume). Fermentation can be performed either in a separate fermentor tank, a process configuration normally re­ferred to as separate hydrolysis and fermentation (SHF), or simultaneously with the hydrolysis of the cellulose chains, so-called simultaneous sacchar­ification and fermentation (SSF). If the pentose sugars are also fermented, the process is sometimes referred to as simultaneous saccharification and co­fermentation (SSCF). The downstream processing section is similar for the dilute acid hydrolysis and the enzymatic processes, or at least includes the same process steps (Figs. 2 and 3).

Simulation of processes with the aid of flowsheeting programs is an in­valuable tool in studying how changes in process design affect the overall performance of a plant. Plants operating 24/7 cannot be experimented on, since the profit loss may be considerable if an ill-planned test causes standstill for a day or two. By performing “experiments” on a plant using computers the outcome of a design change can be evaluated beforehand, which will make a change in the process less risky.

This work will focus on the process economic aspects of ethanol pro­duction from lignocellulosic materials and provide targets for where process improvements should be investigated. The enzymatic process will be consid­ered in detail, as most research over the years has been concentrated on this type of process. However, as mentioned earlier, the process suffers from the fact that process data from large production plants are very scarce. Neverthe­less, the data gathered so far on lab and bench scales can be used as input data in flowsheeting programs for comparison of various process alternatives and to help identify bottlenecks in a process. A summary of various pub­lished reports and papers will be made. Unfortunately, this is an area that has clearly been neglected by many researchers, since the number of publications is small.

2

It soon became evident that the mere introduction ofpentose utilization path­ways in S. cerevisiae was not enough to render the recombinant strains traits for efficient ethanol fermentation [43] (strain RWB202, Tables 3 and 4), [47, 48,78,79] (strain TMB3001, Tables 1-3). A number of metabolic engin­eering strategies to enhance ethanolic xylose (and arabinose) fermentation in S. cerevisiae have been explored, the most important of which will be discussed below. The initial xylose utilization pathway, the cellular redox metabolism, and the flux of central carbon metabolism have been the main targets of these engineering strategies. Figure 3 highlights the metabolic re­actions that have been engineered to improve ethanolic xylose fermentation by S. cerevisiae.

4.1

1 Introduction……………………………………………………………………………………………… 206

2 Baker’s Yeast (S. cerevisiae) as a CBP Host…………………………………………………….. 208

3 Engineering S. cerevisiae for Sugar Fermentation……………………………………………… 210

4 Expression of Cellulases in S. cerevisiae…………………………………………………………. 211

5 Expression of Hemicellulases in S. cerevisiae…………………………………………………… 218

6 Selection for the Development of Superior CBP Yeasts………………………………….. 224

7 Integration of Different Enzymatic Activities

into a Single CBP Yeast and Transfer to Industrial Strains………………………. 228

References…………………………………………………………………………………………………….. 230

Abstract Consolidated bioprocessing (CBP) of lignocellulose to bioethanol refers to the combining of the four biological events required for this conversion process (production of saccharolytic enzymes, hydrolysis of the polysaccharides present in pretreated biomass, fermentation of hexose sugars, and fermentation of pentose sugars) in one reactor. CBP is gaining increasing recognition as a potential breakthrough for low-cost biomass pro­cessing. Although no natural microorganism exhibits all the features desired for CBP, a number of microorganisms, both bacteria and fungi, possess some of the desirable properties. This review focuses on progress made toward the development of baker’s yeast (Saccharomyces cerevisiae) for CBP. The current status of saccharolytic enzyme (cellulases and hemicellulases) expression in S. cerevisiae to complement its natural fermentative ability is highlighted. Attention is also devoted to the challenges ahead to integrate all required enzymatic activities in an industrial S. cerevisiae strain(s) and the need for molecular and selection strategies pursuant to developing a yeast capable of CBP.

Keywords Consolidated bioprocessing • Cellulolytic yeast •

One-step bioethanol production • Saccharomyces cerevisiae

1

Introduction

Biomass is the only foreseeable renewable feedstock for sustainable produc­tion of biofuels. The main technological impediment to more widespread utilization of this resource is the lack of low-cost technologies to overcome the recalcitrance of the cellulosic structure [1]. Four biological events occur during conversion of lignocellulose to ethanol via processes featuring enzy­matic hydrolysis: production of saccharolytic enzyme (cellulases and hemi — cellulases), hydrolysis of the polysaccharides present in pretreated biomass, fermentation of hexose sugars, and fermentation of pentose sugars [2]. The hydrolysis and fermentation steps have been combined in simultaneous sac­charification and fermentation (SSF) of hexoses and simultaneous saccharifi­cation and cofermentation (SSCF) of both hexoses and pentoses schemes. The ultimate objective would be a one-step “consolidated” bioprocessing (CBP) of lignocellulose to bioethanol, where all four of these steps occur in one re­actor and are mediated by a single microorganism or microbial consortium able to ferment pretreated biomass without added saccharolytic enzymes (Fig. 1).

CBP is gaining increasing recognition as a potential breakthrough for low — cost biomass processing. A fourfold reduction in the cost of biological pro­cessing and a twofold reduction in the cost of processing overall is projected when a mature CBP process is substituted for an advanced SSCF process fea­turing cellulase costing US $0.10 per gallon ethanol [3]. The US Department of Energy (DOE) Biomass Program multiyear technical plan states: “Mak­ing the leap from technology that can compete in niche or marginal markets for fuels and products also requires expanding the array of possible concepts and strategies for processing biomass. Concepts such as consolidated bio­processing… offer new possibilities for leapfrog improvements in yield and cost.” [4]. The detailed analysis of mature biomass conversion processes by Greene et al. [5] finds CBP to be responsible for the largest cost reduction of all R&D-driven improvements incorporated into mature technology scenar­ios featuring projected ethanol selling prices of less than US $0.70 per gallon. Finally, a recent report entitled Breaking the Biological Barriers to Cellulosic Ethanol states: “CBP is widely considered to be the ultimate low-cost configu­ration for cellulose hydrolysis and fermentation.” [6].

In addition to being desirable, recent studies of naturally occurring cel­lulolytic microorganisms provide increasing indications that CBP is feas­ible. Lu et al. [7] showed that cellulase-specific cellulose hydrolysis rates exhibited by growing cultures of Clostridium thermocellum exceed specific rates exhibited by the Trichoderma reesei cellulase system by approximately 20-fold, with a substantial part of this difference resulting from “enzyme — microbe synergy” involving enhanced effectiveness of cellulases acting as part of cellulose-enzyme-microbe complexes. Whereas cellulase synthesis

was thought to be a substantial metabolic burden for anaerobic microbes fermenting cellulose without added saccharolytic enzymes, C. thermocellum realizes cellulose-specific bioenergetic benefits that exceed the bioenergetic cost of cellulase synthesis [8]. These and other observations provide guid­ance with respect to features that may be beneficial in the course of creating recombinant cellulolytic microbes, and also underscore the point that micro­bial cellulose utilization is differentiable from enzymatic hydrolysis from both fundamental and applied perspectives [1,3].

Although no natural microorganism exhibits all the features desired for CBP, a number of microorganisms, both bacteria and fungi, possess some of the desirable properties. These microorganisms can broadly be divided into two groups: (1) native cellulolytic microorganisms that possess supe­rior saccharolytic capabilities, but not necessarily product formation, and

(2) recombinant cellulolytic microorganisms that naturally give high prod­uct yields, but into which saccharolytic systems need to be engineered [1, 9]. Examples of native cellulolytic microorganisms under consideration in­clude anaerobic bacteria with highly efficient complexed saccharolytic sys­tems, such as mesophilic and thermophilic Clostridium species [9,10], and fungi that naturally produce a large repertoire of saccharolytic enzymes, such as Fusarium oxysporum [11] and a Trichoderma species [12]. However, the anaerobic bacteria produce a variety of fermentation products, limiting the ethanol yield, whereas the filamentous fungi are slow cellulose degraders and give low yields of ethanol [13]. Candidates considered as potential re­combinant cellulolytic microorganisms into which saccharolytic systems have been engineered include the bacteria Zymomonas mobilis [14,15], Escherichia coli [16,17] and Klebsiella oxytoca [18,19], and the yeast Saccharomyces cere — visiae and xylose-fermenting yeasts Pachysolen tannophilus [20], Pichia stipi — tis, and Candida shehatae [21].

While both native and recombinant cellulolytic microorganisms merit in­vestigation, this review will focus on the well-known ethanol producing yeast S. cerevisiae, which has a long commercial history as microorganism of choice for beer, wine, baker’s yeast, and commercial ethanol production. In particu­lar, we address recent progress in heterologous cellulase expression pursuant to development of recombinant cellulose-fermenting yeast strains [22-25].

2

The application of 13C and 31P Nuclear Magnetic Resonance (NMR) spec­troscopy can provide information on both metabolic and energy status dur­ing cell growth through determination of the levels of various phosphorylated intermediates and energy rich compounds as shown in earlier studies on wild-type strains of Z. mobilis [48,52-55].

More recent research with 31P NMR has identified a less energized state of ZM4 (pZB5) when grown on xylose media [56,57]. 31P NMR studies have established that levels of nucleoside tri-phosphates (mostly ATP) and sugar phosphates were lower for growth on xylose compared to that on glu­cose, with this energy limitation resulting in a potential growth restriction. The presence of by-products identified as xylitol, acetate, lactate, acetoin and dihydroxyacetone by 13C NMR spectroscopy and high-performance li­quid chromatography may also result in some inhibition of growth. Further 31P NMR studies [58] have shown that the addition of inhibitory concentra­tions of sodium acetate caused decreased levels of nucleotide tri-phosphates and sugar phosphates, together with increased cytoplasm acidification.

2.4

Other major biofuel producers include China, which has grown its bioethanol production sector rapidly since 2000 to become the third-largest single bioethanol producer after the USA. Total capacity from four plants in 2005 was about 1.3 billion L, but continued high prices for international oil has led the National Development and Reform Commission to announce that biofuel production will increase dramatically, providing China with the ability to re­place about 2 million t of crude oil by 2010, and 10 million t by 2020 [63]. The Commission also announced that China would begin shifting to non-grain feedstocks, including sweet sorghum, for bioethanol production [63]. Jilin Fuel Alcohol remains the world’s largest corn-based bioethanol plant with a current capacity in excess of 350 million Lyear-1 [64]. The biofuel industry in China has been subsidized, mostly in terms of funds to construct biofuel plants. Some Chinese provinces have announced biofuel mandates, although the national government has not yet made any decision about legislating bio­fuel use [63].

A country poised to be a major biofuel producer is Canada, which cur­rently produces about 250 million L annually [15]. Much of the funding being made available to fund research and development in biofuels in Canada has depended upon the federal government’s environment strategy. This strat­egy has evolved significantly with the ascension of a Conservative minority federal government in 2005, who made a campaign promise to introduce a 5% biofuels mandate. An agreement with provincial governments on the 5% mandate was reached in May, 2006, which will see this mandate take full effect by 2010 [65]. Recently, the federal government announced the proposed Clean Air Act, which was tabled on 19 October 2006 [66]. Unfortunately, the pro­posed Act does nothing to codify the government’s biofuels target, and does not provide concrete policy incentives for additional biofuel use. To help spur some biofuel development, Agriculture and Agri-Food Canada is providing CAD $ 10 million (approximately US $ 8.7 million) in the fiscal year 2006/2007 through the Biofuels Opportunities for Producers Initiative (BOPI). The ob­jective of the Initiative is to help agricultural producers develop business plans for new biofuels projects [67].

Previous governments have provided more substantial support to biofu­els, including a cumulative investment of CAD $ 2.7 billion (US $ 2.34 billion) into the implementation of the former Climate Change Plan for Canada [68], which included incentives for the development and use of environmentally — friendly technologies including bioethanol. The federal Canadian govern­ment provided direct funding for the industry through the Ethanol Expansion Program, which in 2004 and 2005 provided a total of CAD $ 118 million (US$ 102 million) in direct funding for 11 projects, six of which are cur­rently in active development [69]. The federal government provides an excise tax exemption for biofuels, as do the provinces of Manitoba, On­tario, and Alberta [70]. Most recently, the Alberta government has an­nounced a commitment of CAD $ 239 million (US $ 207 million) to expand the province’s bioenergy sector by encouraging products including biofuel development [71]. Other nations with biofuel-friendly policies include Aus­tralia, where a bioethanol production subsidy is in place that replaces excise tax exemptions at a rate of approximately US$ 0.21 L-1 produced. Capital subsidies have been provided for two bioethanol production plants [64]. In Thailand, excise taxes are waived for bioethanol. In Latin America, produc­tion schemes in Peru and Columbia have been linked to urban renewable fuel standards in Columbia [64]. In a move designed to utilize surplus pro­duction, the sugar industry in India has successfully lobbied the government for state-level E5 fuel mandates, which were passed in September 2002 and which apply to nine states and four territories. In order to support these man­dates, an excise tax exemption was granted and bioethanol prices have been fixed by a Tariff Commission [72]. Production from other nations will become more important as capacity comes on-line and the international market for bioethanol continues to develop.

3

Random methods such as mutagenesis, adaptation, hybridization, and evo­lutionary engineering [130] have been employed to obtain improved xylose­utilizing [5,42,110,131] (strains TMB3400, C1, C5, BH42, RWB218, RWB202- AFX, H2490-4, Tables 1, 3, and 4) and arabinose-utilizing [71] S. cerevisiae strains. Some of the resultant strains have been analyzed in order to identify molecular traits related to the improved ethanolic fermentation of pentose sugars. High-throughput technologies, such as transcription analysis [71,91, 109,132], enzyme and metabolite analysis [110], and proteome analysis [57], have been used. In many cases, the mutations and alterations observed in mutant strains are the same as have been earlier rationally engineered, con­firming previous knowledge and hypotheses about control and regulation of pentose metabolism. So far, no report exists where completely novel infor­mation would have been obtained from high-throughput molecular analyses. Thus, the investigations have mainly served to confirm and demonstrate the validity of the technologies.

5

Because dependence on nutritional supplementation increases the process cost, the ideal biocatalyst should produce high amounts of ethanol in sim­ple mineral salts growth medium. While KO11 and LY01 both attained high ethanol yields and titers in rich media, these microbial biocatalysts perform poorly in minimal media. With nutritional supplementation, KO11 produced 45 g L-1 ethanol from 100 g L-1 glucose in 72 h; in minimal medium less than 30 gL-1 were produced in 96 h [31]. Results for LY01 were similarly disap­pointing: the final cell mass and ethanol titer attained in minimal medium were tenfold lower than in rich medium [32]. Considering that these strains were selected in rich media, this is not a surprising result. The low ethanol production by KO11 in minimal media has been attributed to suboptimal partitioning of pyruvate for biosynthesis [33,34]. Low acetyl CoA and high NADH levels result in inhibition of citrate synthase, limiting the availability of 2-oxoglutarate for biosynthesis. 2-Oxoglutarate is required for the biosynthe­sis of many amino acids and is an important compound for osmotic tolerance. This proposed inhibition of citrate synthase was supported by the finding that expression of a NADH-insensitive citrate synthase from Bacillus increased the growth and ethanol production of KO11 by about 75% [33].

The ability of microbial biocatalysts to retain ethanologenicity over time without dependence on antibiotics is important for minimizing production costs. While instability of KO11 has been reported [35,36], other reports have demonstrated maintenance of KO11 ethanologenicity for up to 27 days in continuous stirred tank and fluidized beds reactors [37].

In addition to the production of 48 gL-1 ethanol in rich media, KO11 also produced up to 192 mgL-1 of the undesirable co-product ethyl acetate. An es­terase with ethyl acetate hydroylase activity (estZ) from Pseudomonas putida was introduced into KO11 and the presence of this enzyme reduced the ethyl acetate level to less than 20 mg L-1, a level comparable to that of yeast fermen­tation [38].

2.2

Birgitte K. Ahring (И) • Peter Westermann

Bioscience and Technology, BioCentrum-DTU, Technical University of Denmark,

Building 227, 2800 Lyngby, Denmark

bka@biocentrum. dtu. dk

1 Introduction……………………………………………………………………………………………… 290

2 Hydrogen Production………………………………………………………………………………… 291

3 Methane Production…………………………………………………………………………………. 291

4 Production of Biofuels Using the Maxifuel Concept……………………………………… 292

4.1 Pretreatment…………………………………………………………………………………………….. 295

4.2 Hydrolysis………………………………………………………………………………………………… 296

4.3 Separation………………………………………………………………………………………………… 296

4.4 Fermentation…………………………………………………………………………………………….. 296

4.5 Waste Water Treatment…………………………………………………………………………….. 297

4.6 Bio/Catalytic Refineries……………………………………………………………………………… 299

4.7 Integrating Conventional and Bio/Catalytic Refineries………………………………….. 299

5 Conclusion……………………………………………………………………………………………….. 301

References……………………………………………………………………………………………………. 301

Abstract Large scale transformation of biomass to more versatile energy carriers has most commonly been focused on one product such as ethanol or methane. Due to the nature of the biomass and thermodynamic and biological constraints, this approach is not optimal if the energy content of the biomass is supposed to be exploited maximally. In natural ecosystems, biomass is degraded to numerous intermediary compounds, and we suggest that this principle is utilized in biorefinery concepts, which could provide dif­ferent fuels with different end use possibilities. In this chapter we describe one of the first pilot-scale biorefineries for multiple fuel production and also discuss perspectives for further enhancement of biofuel yields from biomass. The major fuels produced in this refinery are ethanol, hydrogen, and methane.

We also discuss the applicability of our biorefinery concept as a bolt-on plant on conventional corn — or grain-based bioethanol plants, and suggest that petroleum-base re­fineries and biorefineries appropriately can be coupled during the transition period from a fossil fuel to a renewable fuel economy.

Keywords Biorefinery • Fuel cells • Hydrogen • Methane • Reforming

1

Introduction

Traditionally, the development of biological processes to transform biomass to more versatile energy carriers has focused on the production of one en­ergy carrier, either hydrogen, methane, or ethanol. Among these products, only methane is released from the conversion of organic matter in nature; both hydrogen and ethanol are intermediates during anaerobic degradation and are further metabolized to methane in nature [1]. The production of these two energy carriers, therefore, demands a physical separation of indi­vidual processes in the anaerobic degradation chain, or the use of defined microbial cultures under controlled conditions. This can be carried out in a biorefinery, which is a facility that integrates biomass conversion processes and equipment to produce fuels, power, and chemicals from biomass [2,3]. The biorefinery concept is analogous to today’s petroleum refineries, which produce multiple fuels and products from petroleum.

Instead of concentrating on the biological production of only one en­ergy carrier, the simultaneous production of hydrogen, methane, and ethanol leaves the possibility to optimize the exploitation of the specific energy car­riers to suit specific needs, corresponding to the current use of specific fossil fuels for specific purposes. Hydrogen can for instance be used in fuel cells for urban transportation. Ethanol can be used in fuel cells in rural areas, and me­thane can be used in fuel cells for local electricity and heat production in fuel cells or micro-turbines [4]. Although the fuel cell technology was developed initially for molecular hydrogen, this technology is in rapid progression, and fuel cell systems dealing with more complex compounds such as ethanol are currently being developed [5,6].

Despite the obvious advantages of combining the production of different energy carriers, only a few concepts have been published. Common to the known concepts is a much better exploitation of the biomass by suiting spe­cific microbiological processes to the conversion of different fractions of the substrates to different fuels. The different processes are thereby exploited in an additive sequential fermentation, transforming most of the energy avail­able in the substrate to usable energy carriers. Furthermore, biorefineries might be considered as more environmentally friendly processes since pro­cess water and nutrients from the different processes can be recirculated, and waste production can be kept minimal [4].

By producing multiple products, a biorefinery can also take advantage of the differences in biomass components and intermediates and maximize the value derived from the biomass feedstock. A biorefinery might, for ex­ample, produce one or several low-volume, but high-value, chemical products and a low-value, but high-volume liquid transportation fuel, while generat­ing electricity and process heat for its own use and perhaps enough for sale of electricity. The high-value products enhance profitability, the high-volume fuel helps meet national energy needs, and the power production reduces costs and avoids greenhouse-gas emissions.

2

Mixtures of selected thermostable enzymes (Table 2) were first evaluated for their hydrolytic efficiency by measuring the FPU activities at different tem­peratures (Fig. 3). The temperature optima of the new thermostable mixtures in the FPU activity assay were 5-10 °C higher than those of the commercial enzyme mixtures when a relatively short reaction time (60 min) in this assay was used. The relative FPU activity was set at the value of 100 at the refer­ence point at 50 °C. The maximum FPU activity of the novel enzyme mixture was about 25% higher at the optimum temperature at 65 °C as compared with the highest activity of the commercial reference enzyme at 60 °C. As could be expected, at lower temperature (35 °C), corresponding to the fermentation temperature of traditional yeasts in a simultaneous saccharification and fer­mentation process (SSF), the FPU activities of the thermostable preparations were slightly lower than those of the commercial T. reesei enzymes.

The thermostable enzyme mixture without added xylanase activity (TM 1) was evaluated on pure cellulose (Avicel) and compared with the commercial enzyme preparations (Celluclast supplemented with в-glucosidase) at 45 °C, 55 °C and 60 °C in a 48 h hydrolysis (Fig. 4). On pure cellulose, the mixture of thermostable enzymes gave nearly similar hydrolysis results at 60 °C as the T. reesei enzymes at 45 °C, i. e. thus enabling an increase of temperature of about 15 °C. At 60 °C, the hydrolysis yield of Avicel was about three — to four­fold better with the thermostable enzymes than with the commercial fungal enzymes. The highest hydrolysis yield was about 90% of the theoretical.

On the spruce substrate, the thermostable enzyme mixture resulted in an even more significant improvement in the performance at higher hydrolysis temperature as compared with the commercial enzymes. Thus, the hydrolysis yield was about threefold better at 55 °C and about fivefold better at 60 °C
using the thermostable enzyme mixture (Fig. 5). The hydrolysis was, however, also decreased with the thermoenzyme mixture at 60 °C. When comparing the hydrolytic performance of the commercial enzymes by increasing the temperature from 45 °C to 60 °C on Avicel and on spruce, it can be observed that the increased hydrolysis temperature decreased the performance on the natural lignocellulose substrate significantly more: from 70-10% on spruce, as compared with 90-30% on Avicel within 48 h. Obviously, the spruce sub­strate, even washed, contained compounds that, with increasing temperature, inhibited or inactivated not only the T. reesei enzymes, but also the ther­mostable enzymes.

High temperature enzyme mixtures suitable for hemicellulose-containing raw materials were evaluated in the hydrolysis of steam pretreated corn stover substrate (Fig. 6). With this raw material, the hydrolysis by the thermostable enzyme mixture at 45 °C was better than with the commercial preparation. The hydrolysis was still efficient at 55 °C and only slightly decreased at 60 °C with the thermostable enzyme mixture. The relative decrease of the hy­drolytic performance of both enzyme preparations was less pronounced on the corn stover substrate than with the spruce substrate at elevated tempera­tures. Based on HPLC analysis (Table 4) of the corn stover hydrolysates, the yield of glucose was around 90-95% of the theoretical after 72 h. The cor­responding yield of xylose was about 80-90% at temperatures up to 60 °C. The hydrolysis yields of the minor monosaccharide sugars, arabinose and galactose, were not significantly improved by the thermophilic enzyme mix­tures, indicating the absence of corresponding thermostable enzymes, i. e. arabinosidases and galactanases in the mixtures. In the hydrolysis of the

Table 4 Sugars released from steam pretreated spruce and corn stover (% of the initial sugar component in the substrate), analysed by HPLC

Enzymes Hydrolysis Sugars released Sugars released from corn stover

temp. from spruce

% of theoretical % of theoretical

(°C) Glucose Glucose Xylose Arabinose Galactose

Enzyme dosage was for reference enzymes: Celluclast 5 FPU g-1 substrate supplemented with 100 nkatg-1 Novozym 188; and for thermostable enzyme (TM 3) 5 FPU g-1 substrate. Hydrolysis time 72 h at pH 5, triplicates with mixing. Release of xylose, mannose and ara — binose from spruce substrate was below the reliable detection limit (less than 0.1% of the substrate) spruce substrate (Fig. 5, Table 4), only glucose was released. The individual sugar analyses corresponded well with the measured values of the reducing sugars.

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Xylose isomerase (XI, D-xylose ketol isomerase, EC 5.3.1.5) catalyses the re­versible isomerisation of D-xylose to D-xylulose. This enzyme has been the subject of much applied research because it also catalyses the isomerisation of D-glucose and D-fructose. In this role of “glucose isomerase”, xylose iso — merase is applied on a huge scale for the production of high-fructose corn syrup and continues to be one of the most abundantly applied industrial en­zymes. The high-fructose syrup application has led to intensive screening and protein engineering studies, with increased activity and stability of XIs at el­evated temperature as a priority target [11,23]. For excellent reviews on the molecular and industrial aspects of XI, the reader is referred to a number of specialised reviews [4,11,12].

In the context of the present paper, several characteristics of XIs are note­worthy. First and foremost, and in contrast to the xylose reductase/xylitol dehydrogenase pathway, the XI reaction does not involve pyridine nucleotide cofactors. As this will entirely circumvent the cofactor regeneration chal­lenges associated with the xylose reductase/xylose dehydrogenase pathway, functional expression of a XI in S. cerevisiae has long been regarded the most promising approach to engineering S. cerevisiae for alcoholic fermentation of D-xylose [14].

XIs generally require divalent cations, but the specificity of the metal re­quirement is strongly dependent on the source of the enzyme, with many enzymes requiring Co2+, but others Mn2+ or Mg2+ [11]. Although S. cere­visiae has been demonstrated to accumulate cobalt intracellularly [18], it is not clear whether this metal is available in the cytosol or sequestered in, for example, the vacuole. Other aspects with potential relevance for yeast metabolic engineering include the high temperature optimum (60-80 °C) and the relatively high pH optimum (7.0-9.0) of many of the XIs that have been characterised [11]. As S. cerevisiae is a mesophilic micro-organism with a cytosolic pH slightly below 7, intracellular expression of heterologous struc­tural genes for XIs may not always lead to optimal activity.

Even in the pre-genomics era, it was clear that XIs are widespread among prokaryotic micro-organisms, and also occur in several plants [11]. Figure 3 shows a phylogenetic tree of XI gene sequences based on an October 2006 GenBank database search. This phylogenetic tree gives a good indication of the diversity of XI genes and the phylogenetic relationships between se­quences from related organisms. With respect to eukaryotes, the tree contains four sequenced XI sequences from the plants Hordeum vulgare, Arabidop — sis thaliana, Oryza sativa and Medicago truncatula, which cluster together (Fig. 3). The phylogenetic tree contains only one other eukaryotic XI se­quence, namely that of the anaerobic fungus Piromyces sp. E2 [28]. Inter­estingly, this eukaryotic XI sequence clusters with those of the prokaryotic phylum Bacteroidetes, which has led to the suggestion that the fungus may have acquired XI via horizontal gene transfer [28], as previously suggested for other enzymes in anaerobic fungi [20].

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