Consolidated Bioprocessing for Ethanol Production

CHAPTER

Consolidated Bioprocessing for Ethanol Production

7

Zhiliang Fan* Department of Biological and Agricultural Engineering, University of California, Davis, California, USA *Corresponding author: [email protected]

7.1  INTRODUCTION Given rising energy needs, emerging environmental issues, and mounting international security concerns, there is growing interest in the production of sustainable fuel alternatives from renewable resources. Ethanol is one of the most actively pursued renewable biofuels due to its technological maturity and its favorable features as a transportation fuel, including its high octane rating, low cetane number, and higher vapor pressure [1]. In the United States, most ethanol is produced from sugar- or starch-based materials, which are in limited supply and are also used as food sources. The US government has set a mandate that by 2022 the total annual production of renewable fuels should reach 36 billion gallons. Meeting such an objective requires the development of new technologies for the efficient and sustainable production of biofuels from cellulosic biomass, which is an abundant, low-cost, alternative source that does not compete with the food supply. The United States has the capacity to grow enough biomass to displace more than 30% of US petroleum consumption [2]. The production of biofuels from biomass can play a significant role in meeting the nation’s mobility needs, alleviating dependence on foreign oil, and reducing greenhouse gas emission [3,4]. In addition, processing cellulosic biomass can increase the demand for agricultural products, improve rural economy, create local green jobs, and move our society toward sustainability [4,5]. The term lignocellulosic biomass refers to plant materials that have cellulose, hemicellulose, and lignin as their major components [6]. Common examples of lignocellulosic biomass include energy crops, waste from agricultural and forest products, industrial waste, and municipal waste [6,7]. On a dry-weight basis, unprocessed lignocellulosic biomass typically contains 40-55% cellulose, 25-50% hemicellulose, 10-40% lignin, and about 5% extractives and ash. The relative amounts of each component vary with the source of the biomass [6,7]. Biorefineries Copyright © 2014 Elsevier B.V. All rights reserved.

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The major component of lignocellulosic biomass is cellulose, which is a waterinsoluble polymer composed of anhydroglucose units linked by β-1, 4-glycosidic bonds. The crystalline structure of cellulose and the fact that cellulose fibers are embedded within hemicellulose and lignin make cellulose very resistant to hydrolysis [6,8]. In contrast, hemicellulose is a branched polymer. The hydrolyzate of hemicellulose contains xylose, arabinose, mannose, and galactose in addition to glucose. Hemicellulose can be much more easily hydrolyzed than cellulose [6,8]. Lignin is a hydrophobic aromatic macromolecule that is covalently linked to hemicellulose, conferring mechanical strength to the plant cell wall [6,8]. Lignocellulosic biomass is too recalcitrant for direct conversion to biofuels or chemicals. Therefore, additional steps such as thermochemical or biochemical methods are needed to convert it to reactive intermediates. In thermochemical methods, cellulosic biomass is converted to synthetic gas or pyrolysis bio-oil as the reactive intermediates [4,5]. In biochemical methods, the hydrolysis of both cellulose and hemicellulose by cellulases and other hydrolases produce sugar or sugar-like molecules as the reactive intermediates, which can be then converted to ethanol by fermentation [5].

7.2  BIOCHEMICAL PROCESSES FOR ETHANOL PRODUCTION FROM CELLULOSIC BIOMASS When the production of ethanol from lignocellulosic biomass features sugars as reactive intermediates and involves enzymatic hydrolysis, it consists of five steps: pretreatment, cellulase production, enzymatic hydrolysis, microbial fermentation, and product recovery.

7.2.1 PRETREATMENT Cellulose has a highly crystalline structure with strong intramolecular and extramolecular hydrogen bonds, and it is interwoven with hemicellulose and surrounded by a lignin seal [8]. As a result, it is difficult to enzymatically hydrolyze lignocellulosic biomass directly. Hence, pretreatment is needed to make cellulose more assessable for enzymatic hydrolysis, which is achieved by removing the lignin seal, solubilizing the hemicellulose, and decreasing the degree of crystallization of the cellulose. Over the years, many different physical, thermal, chemical, and biological techniques have been applied to achieve the above goals [9,10]. To date, chemical pretreatment of cellulosic biomass at elevated temperatures has provided the highest yields and lowest costs for enhancing the release of sugars from cellulosic biomass in subsequent enzymatic hydrolysis [11]. The most commonly used chemicals include dilute sulfuric acid, sulfur dioxide, ammonia, and lime [11].

7.2.2  CELLULASE PRODUCTION Achieving complete hydrolysis of cellulose into monomer sugars requires the synergetic effects of three major types of enzymes: endo-β-1,4-glucanases (EC

7.2  Biochemical processes for ethanol production

3.2.1.4), cellobiohydrolases (EC 3.2.1.91), and β-glucosidase (EC 3.2.1.21) [12]. Endoglucanases (EGs) attack the more amorphous parts of the cellulose chains and cut them in the middle, releasing more free ends. Cellobiohydrolases (CBHs) depolymerize and hydrolyze the highly crystalline regions in cellulose from the ends of the chain, releasing soluble cellooligosaccharide with cellobiose as the major product. Finally, β-glucosidases (BGLs) hydrolyze cellooligosaccharides into monomer glucose [12]. Another very important general feature of cellulases is their modular structure. Cellulases often include both catalytic modules and carbohydrate-binding modules (CBMs). The CBMs effect binding to the cellulose surface and bring the catalytic domain in close proximity to the substrate. The type of CBMs is particularly important for the initiation and processivity of exoglucanases [13]. In nature, two distinct cellulase systems are produced by aerobic and anaerobic cellulolytic microorganisms. Aerobic cellulolytic microorganisms, including both fungi and bacteria, produce noncomplexed cellulase systems. They produce substantial amounts of extracellular cellulase enzymes in the culture supernatants [4,14]. Each cellulase component may contain its own catalytic domain and cellulosebinding module. The CBM facilitates the binding of the individual cellulase components to cellulose. Synergy can be found among and between different cellulase components [15]. These cellulolytic microorganisms tend to produce relatively high titers of cellulases and high cell yields as a result of aerobic growth and a plentiful ATP supply [4]. Most of the anaerobic cellulolytic bacteria produce complexed cellulase systems. Clostridium thermocellum is one of the most well characterized examples in this category [16,17]. The cellulosome produced by C. thermocellum contains a scaffoldin unit, which organizes the various cellulolytic subunits (e.g., the enzymes) into the complex. Within a cellulosome, multiple catalytic domains with different enzyme functions are assembled into the scaffoldin unit through the specific interaction between the dockerin module contained by each enzymatic subunit and the cohesion module on the scaffoldin [16,17]. The cellulosome is attached to the cell surface, with very few cellulases released to the liquid broth [18]. C. thermocellum grows optimally on cellulose when attached to the substrate [4,19]. The cellulosome also contains one or a few CBM, which anchor the cellulosome on the surface of the cellulose [18]. Together, the cellulose, cellulosome, and cell form a tertiary cellulose-enzyme-microbe system (CEM) [17]. The cellulase titers produced by these anaerobic cellulolytic microorganisms in the complexed cellulase system are low; however, the specific activity is very high as compared to the activity produced in the noncomplexed system by aerobic cellulolytic microorganisms [4,16]. Although anaerobic cellulolytic microorganisms produce cellulase of very high specific activity, slow growth rates and low cellulase titers limit their industrial application. In industry, aerobic cellulolytic fungi are the main workhorses for cellulase production. Trichoderma reesei, the leading microorganism in industrial cellulase production, can produce cellulase in high titers (up to 0.33 g of protein/g of utilized carbohydrate) [20]. Two major components of the cellulases produced by T. reesei are CBH1 and CBHII, which constitute up to 60% and 20%, respectively, of the

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total mass of cellulase protein produced [21]. T. reesei produced at least five different endoglucanases [21]. The BGLs produced by T. reesei are at low levels and are more subject to glucose inhibition than other fungi such as Aspergillus species [4,22]. In the cellulose saccharification processes used on an industrial scale, the cellulase preparations from T. reesei are often supplemented with β-glucosidase from Aspergillus species [22,23].

7.2.3  ENZYMATIC HYDROLYSIS Enzymatic hydrolysis is the process in which cellulases are added to hydrolyze pretreated lignocellulosic biomass into fermentable sugars. The process involves several key steps: (1) transfer of enzymes from the bulk aqueous phase to the surface of the cellulose, (2) adsorption of the enzymes and formation of enzyme-substrate complexes, (3) hydrolysis of the cellulose, (4) transfer of the hydrolysis products from the surface of the cellulosic particles to the bulk aqueous phase, and (5) hydrolysis of cellodextrins and cellobiose to glucose in the aqueous phase [24–27]. The overall rate of the process is influenced by the structural features of lignocellulosic biomass and the composition and source of the cellulases [4,24,28]. The features of cellulose that most affect its hydrolysis rate are its crystallinity and the amount of accessible surface area [25,29,30]. A cellulose fiber is composed of both crystalline and amorphous regions. Cellulose crystallinity affects enzyme adsorption, synergism among cellulase components, and the processivity of cellulases [31–34]. When cellulase binds to crystalline cellulose, hydrolysis rates and yields are more than 100 times lower than when bound to amorphous cellulose [31–34]. Accessible surfaces of a cellulose fiber include both external and internal surfaces [4,24,28]. The external surface area is affected by the shape and size of the cellulosic particles, while the internal surface area depends on the capillary structure of the cellulose fibers, the anatomical structure of the plant cell wall, and the method of pretreatment [4,24,28]. The rate of enzymatic hydrolysis is also influenced by enzyme sources and the proportion of different enzyme components [24,28]. A synergetic effect was observed among different components of cellulases [35–37], as well as among various glycosyl hydrolases (e.g., cellulases, hemicellulases, and ligninases) [37–39]. Cellulase enzymes from different sources have different resistances to product inhibition, which, in turn, affect enzymatic hydrolysis rates [24,40–43].

7.2.4  MICROBIAL FERMENTATION Ethanol production from sugars has been practiced for thousands of years, and the production of ethanol fuel from sugar-based or starch-based feedstock is a well-developed technology. However, ethanol production from cellulosic biomass imposes additional demands on the potential microorganism. Unlike corn hydrolyzate, in which glucose is the only monomer released, lignocellulosic biomass hydrolyzate contains both pentoses and hexoses [44]. It is essential that fermenting microorganisms convert all the sugars in the hydrolyzate to achieve favorable economics. Given that many inhibitors

7.2  Biochemical processes for ethanol production

are formed during the pretreatment process, the fermenting microorganisms must also be able to tolerate those inhibitors and exhibit high process robustness [44]. Two strategies are used to construct microorganisms that are able to convert all the sugars contained in the cellulosic biomass hydrolyzate to ethanol at high yields and at high concentrations [4,5,44]. The native cellulolytic strategy involves identifying a microorganism that has a wide substrate range and then converting it into an efficient ethanologen. The recombinant cellulolytic strategy involves starting with an excellent native ethanologen and broadening its substrate range. Both strategies must be implemented using metabolic engineering [4,5,44]. Saccharomyces cerevisiae and Zymomonas mobilis are attractive starting microorganisms for constructing ethanologens for biomass conversion through the recombinant strategy. They have been widely used in industry for ethanol production using starch-based or sugar-based feedstocks [44,45]. They have many favorable features, including high ethanol yield and productivity, high tolerance to ethanol, process hardiness, and generally recognized as safe (GRAS) status [44,45]. However, both of these organisms can use only hexose monomers in the cellulosic biomass hydrolyzate, and thus, neither can utilize pentose sugars, cellobiose, or xylobiose [44,45]. Efforts have been made to broaden the substrate range for S. cerevisiae, including attempts to enable S. cerevisiae to ferment xylose and arabionose [46,47]. Recent progress includes the production of a functional xylose isomerase from Piromyces sp. in S. cerevisiae, which circumvented the redox imbalance problem created by the previous effort using a xylose reductase (XYL1) and xylitol dehydrogenase (XYL2) system [48]. Subsequently, this strain was improved to utilize xylose much more quickly under anaerobic conditions and to coferment arabionose with xylose [49]. Native S. cerevisiae is not able to utilize cellobiose. Engineering S. cerevisiae to produce a recombinantly secreted BGL from Saccharomycopsis fibuligera enabled the recombinant strain to grow on cellobiose at approximately the same rate as on glucose under anaerobic conditions [50]. Recently, a cellobiose-utilizing yeast was constructed by expressing a cellulodextrin transporter and an intracellular BGL from Neurospora crassa in S. cerevisiae. The recombinant strain efficiently transports cellobiose and hydrolyzes cellobiose into glucose intracellularly [51]. The intracellular hydrolysis of cellobiose leads to very low glucose concentrations in the fermentation broth, which minimizes the glucose repression of xylose fermentation [51,52]. A recombinant strain, which also expresses the xylose utilization pathway, can simultaneously consume cellobiose and xylose [52]. Progress on broadening the substrate range of Z. mobilis has also been made, including enabling Z. mobilis to ferment xylose [53], coferment xylose and arabionose [54–56], and tolerate higher concentrations of acetate and other inhibitors in the cellulosic biomass hydrolyzate [57–59]. Because enteric bacteria such as Escherichia coli and Klebsiella oxytoca have a wide substrate range and can utilize both hexose and pentose sugars in cellulosic biomass hydrolyzate, they are very attractive starting microorganisms in the native strategy for constructing microorganisms that ferment sugars in cellulosic biomass [5,44,60]. In addition to having the ability to transport and utilize all the monomer sugars present in the lignocellulosic biomass, K. oxytoca can also transport and utilize

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their oligomers, such as cellobiose, xylobiose, and xylostriose [61]. Wild-type E. coli was converted to an efficient ethanologen by expressing the Z. mobilis homoethanol pathway (pyruvate decarboxylase and alcohol dehydrogenase). More than 90% of the theoretical yield of ethanol was achieved from glucose by the recombinant strain in a rich medium [62]. Subsequent work has been conducted to improve the ethanol yield [63], to enable better growth in a lean medium [64], and to coferment glucose and xylose, which accelerated the metabolism of a five-sugar mixture (mannose, glucose, arabinose, xylose, and galactose) to ethanol [65]. The Z. mobilis homoethanol pathway was also successfully introduced into K. oxytoca, resulting in K. oxytoca P2, which can convert various sugars into ethanol at more than 90% of the theoretical yield [61]. Subsequently, work has been conducted to improve the ethanol yield from sugars [66]. However, both E. coli and K. oxytoca are less ethanol tolerant than native ethanologens such as Z. mobilis or S. cerevisiae [60].

7.2.5  PRODUCT RECOVERY Distillation is the leading method for ethanol recovery from fermentation broth. Typically, a distillation or a stripping process is used to produce ethanol at a concentration of 90% or more in a so-called beer column. To produce anhydrous ethanol, other processes such as gas-phase molecular sieve adsorption, further distillation in the presence of an entrainer that is subsequently recovered, pervaporation, and other membrane-based operations are used following distillation [67,68]. The concentration of ethanol in the fermentation broth produced from cellulosic biomass is much lower than that produced from corn-based feedstocks due to both biological and processing constraints [68]. A solid content value greater than 15% in the broth is difficult to handle in a typical simultaneous saccharification and fermentation (SSF) process [68]. At carbohydrate contents that are typical for cellulosic feedstocks, the ethanol concentration in the broth (with a 15% solid content) is about 5%. Moreover, ethanol at a concentration greater than 60 g/L is inhibitory to both the cellulase and the fermenting microorganism [69–71]. The cost of product recovery increases with decreasing concentrations of ethanol in the fermentation broth, and ethanol concentrations must exceed 40 g/L to ensure economic ethanol recovery [72]. Economic analysis reveals that pretreatment is the most expensive unit operation in the whole process. The next highest costs are those for the enzymatic hydrolysis of the pretreated cellulose and those for producing enzymes to carry out this task [5,73,74]. Lowering the processing costs of these three steps is essential if cellulosic biorefineries are to become economically viable.

7.3  DEVELOPMENT OF BIOMASS PROCESSING CONFIGURATIONS One of the strategies for lowering overall process costs is process consolidation. Figure 7.1 represents the development of biomass processing configurations featuring

7.4  Aspects of consolidated bioprocessing

Feedstocks

Cellulase production

Pretreatment

Enzymatic Hydrolysis

sugars

Fermentation

SSF (SSCF)

Product recovery

Ethanol

CBP

FIGURE 7.1 The process for ethanol production from lignocellulosic biomass.

enzymatic hydrolysis over the past forty years [5].. If pretreatment, cellulose hydrolysis, fermentation, and product recovery take place in different reactors, the process is called separate hydrolysis and fermentation (SHF). In the SHF configuration, cellulases from the enzyme production step are added to the pretreated material to form glucose from the cellulose fraction. Upon completion of hydrolysis, the fermentative microorganism is added to convert sugars to ethanol. SSF consolidates enzymatic cellulose hydrolysis and hexose fermentation in one reactor [75]. Cellulases are added to the pretreated materials to hydrolyze the cellulose fraction to glucose, while the fermentative microorganism converts glucose into biofuels in the same reactor. Given that glucose, an inhibitor of cellulase, is converted by the fermenting microorganism into ethanol, SSF can efficiently remove or reduce the inhibitory effect of glucose on cellulases, thus achieving faster biomass hydrolysis rates and higher ethanol yields as compared to SHF [73]. The simultaneous saccharification and cofermentation (SSCF) process consolidates enzymatic hydrolysis, and hexose and pentose fermentation into one step due to advanced engineering techniques that enable the fermentative microorganism to coferment pentoses and hexoses [46,53,60]. Higher product yields and greater process efficiency can be achieved by SSCF as compared to SSF. Finally, consolidated bioprocessing (CBP), which is discussed next, features the combination of cellulase production, enzymatic cellulose hydrolysis, and fermentation in a single process step.

7.4  ASPECTS OF CONSOLIDATED BIOPROCESSING 7.4.1  ECONOMIC BENEFITS OF CBP CBP features a high level of process consolidation and lacks a dedicated step for cellulase production. A model developed at the National Renewable Energy Laboratory

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(NREL) to analyze advances in biomass processing systems indicated that increasing levels of process consolidation can result in greater cost reductions, and CBP offers the largest cost reduction of any process improvement considered to date [5,67,68]. The cost savings realized by CBP result from the reduction of capital costs and operational costs that occur when the dedicated cellulase production step is eliminated. In addition, CBP can potentially achieve higher process efficiency, which can also lead to lower processing costs [76]. High cellulase costs are the major impediments to low-cost biomass processing. In the past ten years, much effort has been dedicated to reducing the cost of cellulase production for biomass conversion, including efforts to improve both the enzymes and the economics of the enzyme production process. The cost of cellulase production in a biomass conversion process depends on a variety of factors, including the feedstock used, the source of cellulases, and the conversion technology chosen [77]. However, even in the most optimistic case, the cost of cellulase is still too high for the production of a commodity product such as ethanol [77]. The cost savings that could be achieved with a dedicated cellulase production process are substantial [4,76]. In addition, the CBP configuration can potentially lead to more efficient biomass hydrolysis. Two features of CBP, microbe-enzyme synergy and the use of thermophiles, may lead to savings in both capital costs and operational costs by yielding higher biomass hydrolysis rates, which, in turn, lead to reduced reactor volumes and shorter processing cycles [4,76,77].

7.4.1.1  The effects of microbe-enzyme synergy in CBP In the SSF system, cellulose hydrolysis is accomplished using a cellulose-enzyme (CE) system. In a CBP configuration using anaerobic bacteria with a complex cellulase system, cellulose hydrolysis is accomplished by a tertiary cellulose-enzymemicrobe system (CEM) in addition to a CE system. According to a qualitative analysis conducted by Lu et al. [78], the cellulose hydrolysis rates mediated mainly by the CEM complex plus CE in a CBP setup could be 2.7–4.7 times greater than the rates achieved by a CE complex in SSF. This microbe-enzyme synergy requires the presence of metabolically active adherent cellulolytic microorganisms and is believed to result from the microbe’s preferred access to hydrolysis products, local high cellulase concentrations, and the removal of hydrolysis products from the fermentation broth [78]. These increased hydrolysis rates suggest that a CEM system may be an efficient way to decrease enzymatic hydrolysis costs.

7.4.1.2  The use of thermophiles in CBP Some of the cellulolytic microorganisms of potential use in CBP have optimal growth temperatures in the range of thermophiles. For example, C. thermocellum has an optimal growth temperature of 65°C [79], and Caldicellulosiruptor sp. has an optimal growth temperature of 80°C, temperatures at which substantial solubilization of lignocelluloses can be achieved without pretreatment [80]. The advantages of using a high-temperature system include higher reaction rates, reduced sensitivity to contaminants, lower cooling requirements, and the possibility of recovering ethanol

7.5  Approaches to developing CBP-enabling microorganisms

directly from the fermentation broth, thus reducing the inhibition effect and downstream separation costs [81]. All these features can result in process cost savings.

7.5  APPROACHES TO DEVELOPING CBP-ENABLING MICROORGANISMS The CBP system for ethanol production requires a microorganism or a microorganism consortium that is capable of efficiently producing cellulases and efficiently fermenting the resulting sugars into ethanol. CBP-enabling microorganisms or microorganism consortia do not exist naturally in nature and thus must be constructed through genetic engineering [4,5]. Such microorganisms can be developed via two strategies: the native cellulolytic strategy, which involves engineering naturally occurring cellulolytic microorganisms with the ability to ferment sugars into ethanol at high yields and at high concentrations, and the recombinant cellulolytic strategy, which involves engineering noncellulolytic organisms that can produce ethanol at high yields and have the ability to directly utilize cellulose as the substrate [5,76].

7.5.1  THE NATIVE STRATEGY FOR DEVELOPING CBP-ENABLING MICROORGANISMS Using the native strategy for developing CBP-enabling microorganisms that produce ethanol begins with proper cellulolytic microorganisms and then converts them into efficient ethanologens. The production of CBP-enabling microorganisms can theoretically start with any cellulase producer, and cellulolytic fungi are potentially viable CBP hosts [82,83]. Yet, most of the work on developing CBP-enabling microorganisms through the native cellulolytic strategy has focused on anaerobic cellulolytic bacteria. Native cellulolytic anaerobes have branched fermentation pathways that yield a wide variety of fermentation products in addition to ethanol, which lowers the ethanol yield from sugar fermentation. Thus, the major goal of the native strategy is to improve ethanol yield from sugars by deleting the branched pathways, and other goals include increasing the ethanol tolerance of CBP-enabling microorganisms and enabling them to utilize both hexose and pentose sugars [82]. Most of the cellulolytic bacteria that are potential CBP hosts are Grampositive obligate anaerobes. They include cellulolytic bacteria that produce free cellulases, such as Clostridium phytofermentans, Thermoanaerobacter sp., and Thermanaerobacterium sp., as well as cellulolytic microorganisms that produce cellulosomes, such as C. thermocellum and C. cellulolyticum [4,77]. Progress in engineering these microorganisms has been hampered by the paucity of sufficiently advanced genetic tools. The major obstacles include the lack of an efficient gene transfer system for introducing foreign DNA into the cell, the resistance of the host to foreign DNA, and insufficiently developed genetic markers for facilitating selection [84]. If the host is a thermophile, additional challenges in developing a genetic system include the stability of antibiotics at elevated temperatures, the selection of

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genes coding for markers, and the stability of gene products at elevated temperatures [85,86]. However, in the past ten years, substantial progress has been achieved in developing genetic systems for these microorganisms, including the design of new methods for transformation [87,88], new markers [89–91], and the application of group II intron technology to inactive gene targets in some Clostridium species [92–95]. Advances in the development of genetic tools enable progress in CBP microorganism development through the native strategy. For example, researchers recently blocked the production of acetic acid and lactic acid in the anaerobe Thermoanaerbacterium saccharolyticum, a strain that can efficiently utilize xylan and all other biomass sugars [88,96–98]. The engineered strain can produce ethanol at a yield of 0.46 grams of ethanol per gram of xylose or other sugars. In another study, three genes encoding competing pathways were deleted from a C. thermocellum strain, which was subsequently improved by evolution, and the strain produced 14 g/L of ethanol from 40 g/L of Avicel in a 72-h fermentation. This level of production corresponds to 60% of the maximal theoretical yield from cellulose and a cellulose utilization rate of 13.3 g/day/L [99]. Using a coculture composed of this strain and another engineered T. saccharolyticum strain that is deficient in organic acid production, 38 g/L of ethanol was produced from 92 g/L of Avicel in 146 hours, resulting in 80% of the maximal theoretical yield from cellulose. The reasons for coculturing with T. saccharolyticum that can improve the ethanol yield from cellulose are unknown. It was speculated that some metabolite produced by C. thermocellum was metabolized by T. saccharolyticum to produce ethanol [99]. Ethanol tolerance is another very important parameter affecting the feasibility of CBP. The higher an ethanol titer the microorganism can produce, the more economical the recovery cost will be. However, as an end-product of fermentation, ethanol inhibits the fermentative microorganisms by inhibiting glycolytic enzymes and harming the cell membrane [100,101]. Naturally occurring cellulolytic microorganisms were unable to produce high concentrations of ethanol. In addition, there seems to be a gap between the maximum concentration of ethanol produced by cellulolytic microorganisms and the maximum concentration of ethanol that the organisms can tolerate [76]. For example, the maximum concentration of ethanol that C. thermocellum could produce did not exceed 30 g/L, and an ethanol concentration as low as 10 g/L inhibited the growth of this microorganism. However, C. thermocellum could tolerate ethanol at a concentration of 64 g/L when the ethanol was exogenously added to the culture [102]. More recent studies have been conducted to reveal the mechanisms of ethanol tolerance in ethanol-adapted C. thermocellum [103,104]. Genome sequencing of an ethanol tolerant strain revealed more than 400 mutations, including changes in genes responsible for cellulose degradation, ethanol metabolism, and membrane protein production [104]. Specifically, a mutated alcohol dehydrogenase gene alone (adhE) was found to confer most of the ethanol-tolerant phenotype of the ethanoladapted strain. The specificity of the mutated ADH seemed to cause a shift toward

7.5  Approaches to developing CBP-enabling microorganisms

using NADPH as the cofactor, suggesting that the ethanol tolerance of this mutant may be related to central metabolism and redox cofactor balance[104].

7.5.2  THE RECOMBINANT STRATEGY FOR DEVELOPING CBP-ENABLING MICROORGANISMS The use of recombinant technology to develop of CBP-enabling microorganisms for ethanol production involves identifying an ethanologen that can produce ethanol at high yields and high titers and then conferring that microorganism with cellulolytic ability through the the heterologous expression of cellulases [5,76,77]. In order to enable growth on crystalline cellulose, the total cellulase activity produced by the recombinant microorganisms must be sufficiently high [77]. The total cellulase activity of a microorganism is a function of both the amount of the enzyme it can produce and the specific activity of the cellulase system. In turn, the specific activity of the cellulase system depends on the composition, the specific activities, and the spatial arrangement of each component [77]. The recombinant expression of various cellulase components has been attempted in a variety of genetically tractable hosts, including native ethanologens (e.g., S. cerevisiae and Z. mobilis) and recombinant ethanologens (e.g., E. coli and K. oxytoca). The major difficulties and challenges for the recombinant strategy include the adverse effects of the coexpression of multiple heterologous genes on cell performance, the modulation of the expression of different genes at appropriate levels, and the improper protein folding that can prevent secretion [77,82]. In general, the heterologous expression of endogluconases (EGs) in recombinant hosts is far more successful than that of CBHs [77,105]. One reason for the contrasting degrees of expression is that the specific activity of EGs is observed to be two to three times higher than that of CBH on amorphous cellulose, and much fewer EGs are needed to achieve successful activity [105]. In addition, recombinant EGs seem to be able to retain more activity, especially when hyperglycosylated by a host such as S. cerevisiae, than can be retained by CBHs [106]. The functional and high-level expression of CBHs remains a bottleneck in the development of CBP-enabling microorganisms using the recombinant strategy [76,77,105]. Multiple attempts have been made to express endoglucanases in recombinant E. coli and K. oxytoca in order to confer them with cellulolytic ability. One successful example using E. coli involved the expression of the enzymes Cel5Z and Cel8Y from Erwinia chrysanthemi [107]. Initial attempts to express these two enzymes resulted in a recombinant strain in which most of the resulting Cel5Z accumulated and formed inclusion bodies in the periplasmic space, and most of the Cel8Y was secreted [107]. Zhou and coinvestigators further improved the system by reconstructing the type II secretion system coded by the out gene from E. chrysanthemi in E. coli, which facilitated secretion of 50% of the recombinant Cel5Z [108]. The recombinant E. coli strain produced a total of 13,000 IU of active enzyme per liter of culture, of which 7,800 IU were in the supernatant. The total active endoglucanase produced was estimated to reach 4–6% of total cellular protein [108].

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Endoglucanase expression has been similarly pursued in K. oxytoca, and Zhou and Ingram also expressed Cel5Z and Cel8Y from E. chrysanthemi in the ethanologen K. oxytoca P2, while introducing the out gene. Both Cel5Z and Cel8Y were secreted into the supernatant. The strain produced about 24,000 IU/L of endoglucanase activity, which enabled the strain to grow on amorphous cellulose without the addition of exogenous cellulase [109,110]. Several cellulase-encoding genes, such as the endoglucanase gene eglX from Pseudomonas fluorescens and celZ from E. chrysanthemi, were introduced into Z. mobilis. In those cases, however, either no activity was detected due to poor expression or enzyme activity was detected in the periplasmic space only [111,112]. The yeast Saccharomyces cerevisiae has been used with varying degrees of success as the host for expressing one or more cellulase components from both bacteria and fungi [113–118]. Various groups have reported that the coexpression of a β-glucosidase and an endoglucanase enabled the one-step conversion of phosphoric acid-swollen cellulose into ethanol using high-density cell cultures. [119,120]. However, high-level expression of CBHs remains a bottleneck to achieving the conversion of crystalline cellulose into ethanol using recombinant strains. CBHI and CBHII from various sources have been expressed in S. cerevisiae, and the levels of CBHI and CBHII expressed vary significantly with the source of gene, codon usage, choice of promoters, cellular stress reaction to a specific protein, and secretion level by the host [105]. Additionally, CBHs that are recombinantly expressed often lose some activity as compared to native CBHs, mainly due to overglycosylation by the host and faulty folding [105,118,121]. However, significant progress has recently been made in functionally expressing and secreting CBHs. Ilmen et al. reported the successful coexpression of CBHI (Cel6A) and CBHII (Cel7a) in S. cerevisiae; for the first time, the enzyme activity was high enough to enable fermentation on crystalline cellulose [121]. The strain coexpressing CBHI and CBHII was able to grow on Avicel with the exogenous addition of BGL. About 3 g/L of ethanol was produced from 20 g/L of cellulose, achieving 30% of the theoretical maximum yield. The cellulose conversion was 23% [121]. Another advance in the development of CBP-enabling microorganisms involves mimicking the complex cellulase architecture by developing an artificial recombinant cellulosome to enhance cellulolytic activity. Wen et al. expressed the scaffoldin with three C. thermocellum cohesions, the C. thermocellum cellulose-binding domain, and three enzyme components (CBHII and EGII from T. reesei, and BGL from A. aculeatus) fused with C. thermocellum dockerin domains in S. cerevisiae [122]. An artificial minicellulosome was successfully assembled on the cell surface. The recombinant strain displaying this minicellulosome produced 3.5 g/L of ethanol from phosphoric acid-swollen cellulose (PASC). Ethanol production by this minicellulosome was three times greater than that in a culture with the same amounts of purified individual components added [122]. Further progress was made by Tsai et al. (2010), who expressed the four individual components separately using different synthetic yeasts [123]. The synthetic yeast consortium displaying the minicellulosome achieved 60% solubilization of 10 g/L of PASC in 50 hours. However, in either

7.6 Perspectives

case, the cellulolytic activities displayed by the minicellulosome were still too weak to enable direct growth on PASC [123].

7.6  PERSPECTIVES CBP has great potential to provide low-cost cellulosic bioprocessing for ethanol production. To realize the potential of CBP, microorganisms that are able to utilize cellulose at high rates and produce ethanol at high yields and high titers are needed. The development of CBP microorganisms via the native and recombinant strategies has achieved substantial progress. However, strains that are suitable for commercial applications are not yet available. Additional progress using both strategies will result from further development of genetic engineering tools and greater understanding of the principles of microbial cellulose utilization.

REFERENCES [1] Wyman C. Ethanol production from lignocellulosic biomass: overview. Washington DC: Taylor and Francis; 1996. [2] Perlack RD, Stokes BJ. U.S. billion-ton update: biomass supply for a bioenergy and bioproducts industry. In: U.S. Department of Energy, editor. Oak Ridge, TN: Oak Ridge National Laboratory; 2011. [3] Urbanchuk JM. Contribution of the ethanol industry to the economy of the United States. Renewable Fuels Association; 2011. [4] Lynd LR, Weimer PJ, van Zyl WH, Pretorius IS. Microbial cellulose utilization: fundamentals and biotechnology. Microbiol Mol Biol Rev 2002;66:506–77, table of contents. [5] Lynd LR, Wyman CE, Gerngross TU. Biocommodity engineering. Biotechnol Prog 1999;15:777–93. [6] Wiselogel A, Tyson J, Johnsson D. Biomass feedstock: resources and composition. In: Wyman CE, editor. Handbook on bioethanol: production and utilization. Washington, DC: Taylor and Francis; 1996. p. 105–18. [7] Sun Y, Cheng J. Hydrolysis of lignocellulosic materials for ethanol production: a review. Bioresour Technol 2002;83:1–11. [8] Hatfield RD, Ralph J, Grabber JH. Cell wall structural foundations: Molecular basis for improving forage digestibilities. Crop Sci 1999;39:27–37. [9] Hsu TA. Pretreatment of biomass. In: Wyman CE, editor. Handbook on bioethanol, production and utilization. Washington, DC: Taylor & Francis; 1996. p. 179–212. [10] McMillan JD. Pretreatment of lignocellulosic biomass. In: Himmel ME, Baker JO, Overend RP, editors. Enzymatic conversion of biomass for fuels production. Washington: Amer Chemical Soc; 1994. p. 292–324. [11] Mosier N, Wyman C, Dale B, Elander R, Lee YY, Holtzapple M, et al. Features of promising technologies for pretreatment of lignocellulosic biomass. Bioresour Technol 2005;96:673–86. [12] Teeri TT. Crystalline cellulose degradation: New insight into the function of cellobiohydrolases. Trends Biotechnol 1997;15:160–7.

153

154

CHAPTER 7  Ethanol Production

[13] Teeri TT, Koivula A, Linder M, Wohlfahrt G, Divne C, Jones TA. Trichoderma reesei cellobiohydrolases: why so efficient on crystalline cellulose? Biochem Soc Trans 1998;26:173–8. [14] Eriksson KE, Blanchette RA, Ander P. Microbial and enzymatic degradation of wood and wood components. New York, N. Y.: Springer-Verlag; 1990. [15] Din N, Damude HG, Gilkes NR, Miller RC, Warren RAJ, Kilburn DG. C1-Cx revisited— intramolecular synergism in a cellulase. Proc Natl Acad Sci U S A 1994;91:11383–7. [16] Schwarz WH. The cellulosome and cellulose degradation by anaerobic bacteria. Appl Microbiol Biotechnol 2001;56:634–49. [17] Bayer EA, Morag E, Lamed R. The cellulosome–a treasure-trove for biotechnology. Trends Biotechnol 1994;12:379–86. [18] Bayer EA, Kenig R, Lamed R. Adherence of Clostridium thermocellum to cellulose. J Bacteriol 1983;156:818–27. [19] Wiegel J, Dykstra M. Clostridium-Thermocellum—Adhesion and Sporulation While Adhered to Cellulose and Hemicellulose. Appl Microbiol Biotechnol 1984;20:59–65. [20] Esterbauer H, Steiner W, Labudova I, Hermann A, Hayn M. Production of Trichoderma Cellulase in Laboratory and Pilot Scale. Bioresour Technol 1991;36:51–65. [21] Wood TM. Fungal cellulases. Biochem Soc Trans 1992;20:46–53. [22] Reczey K, Brumbauer A, Bollok M, Szengyel Z, Zacchi G. Use of hemicellulose hydrolyzate for beta-glucosidase fermentation. Appl Biochem Biotechnol 1998;70–2:225–35. [23] Sternberg D, Vijayakumar P, Reese ET. Beta-glucosidase: microbial-production and effect on enzymatic-hydrolysis of cellulose. Can J Microbiol 1977;23:139–47. [24] Walker LP, Wilson DB. Enzymatic hydrolysis of cellulose—an overview. Bioresour Technol 1991;36:3–14. [25] Lee SB, Shin HS, Ryu DDY, Mandels M. Adsorption of cellulase on cellulose: effect of physicochemical properties of cellulose on adsorption and rate of hydrolysis. Biotechnol Bioeng 1982;24:2137–53. [26] Ladisch MR, Gong CS, Tsao GT. Cellobiose hydrolysis by endoglucanase (glucan glucanohydrolase) from Trichoderma-reesei - kinetics and mechanism. Biotechnol Bioeng 1980;22:1107–26. [27] Woodward J, Hayes MK, Lee NE. Hydrolysis of cellulose by saturating and nonsaturating concentrations of cellulase - implications for synergism. Bio/Technology 1988;6:301–4. [28] Yang B, Dai Z, Ding SY, Wyman CE. Enzymatic hydrolysis of cellulosic biomass. Biofuels 2011;2:421–50. [29] Cowling EB, Kirk TK. Properties of cellulose and lignocellulosic materials as substrates for enzymatic conversion processes. Biotechnol Bioeng 1976;6:95–123. [30] Cowling EB. Physical and chemical constraints in hydrolysis of cellulose and lignocellulosic materials. Biotechnol Bioeng 1975;5:163–81. [31] Ooshima H, Burns DS, Converse AO. Adsorption of cellulase from trichoderma-reesei on cellulose and lignacious residue in wood pretreated by dilute sulfuric acid with explosive decompression. Biotechnol Bioeng 1990;36:446–52. [32] Ooshima H, Sakata M, Harano Y. Adsorption of cellulase from Trichoderma viride on cellulose. Biotechnol Bioeng 1983;25:3103–14. [33] Sinitsyn AP, Gusakov AV, Vlasenko EY. Effect of structural and physicochemical features of cellulosic substrates on the efficiency of enzymatic hydrolysis. Appl Biochem Biotechnol 1991;30:43–59.

7.6 Perspectives

[34] Jeoh T, Ishizawa CI, Davis MF, Himmel ME, Adney WS, Johnson DK. Cellulase digestibility of pretreated biomass is limited by cellulose accessibility. Biotechnol Bioeng 2007;98:112–22. [35] Henrissat B, Driguez H, Viet C, Schulein M. Synergism of cellulases from Trichoderma reesei in the degradation of cellulose. Bio/Technology 1985;3:722–6. [36] Kanda T, Wakabayashi K, Nisizawa K. Modes of action of exo-cellulases and endocellulases in the degradation of cellulose-I and cellulose-II. J Biochem 1980;87:1635–9. [37] Woodward J. Synergism in cellulase systems. Bioresour Technol 1991;36:67–75. [38] Coughlan MP, Moloney AP, McCrae SI, Wood TM. Cross-synergistic interactions between components of the cellulase systems of Talaromyces emersonii, Fusarium solani, Penicillium funiculosum and Trichoderma koningii. Biochem Soc Trans 1987;15:263–4. [39] Wood TM, Wilson CA, McCrae SI. Synergism between components of the cellulase system of the anaerobic rumen fungus Neocallimastix frontalis and those of the aerobic fungi Penicillium pinophilum and Trichoderma koningii in degrading crystalline cellulose. Appl Microbiol Biotechnol 1994;41:257–61. [40] Perlack RD, Wright LL, Turhollow AF, Graham RL, Stokes BJ, Erbach DC. Biomass as feedstock for a bioenergy and bioproducts industry: the technical feasibility of a billionton annual supply. In: Oak Ridge National Laboratory, editor. Oak Ridge, TN: Oak Ridge National Laboratory; 2005. [41] Gong CS, Ladisch MR, Tsao GT. Cellobiase from trichoderma-viride - purification, properties, kinetics, and mechanism. Biotechnol Bioeng 1977;19:959–81. [42] Kruus K, Andreacchi A, Wang WK, Wu JHD. Product inhibition of the recombinant CelS, an exoglucanase component of the Clostridium thermocellum cellulosome. Appl Microbiol Biotechnol 1995;44:399–404. [43] Kruus K, Wang WK, Ching JT, Wu JHD. Exoglucanase activities of the recombinant clostridium-thermocellum cels, a major cellulosome component. J Bacteriol 1995;177:1641–4. [44] Zaldivar J, Nielsen J, Olsson L. Fuel ethanol production from lignocellulose: a challenge for metabolic engineering and process integration. Appl Microbiol Biotechnol 2001;56:17–34. [45] Dien BS, Cotta MA, Jeffries TW. Bacteria engineered for fuel ethanol production: current status. Appl Microbiol Biotechnol 2003;63:258–66. [46] Ho NWY, Chen ZD, Brainard AP. Genetically engineered Sacccharomyces yeast capable of effective cofermentation of glucose and xylose. Appl Environ Microbiol 1998;64:1852–9. [47] Hahn-Hagerdal B, Karhumaa K, Fonseca C, Spencer-Martins I, Gorwa-Grauslund MF. Towards industrial pentose-fermenting yeast strains. Appl Microbiol Biotechnol 2007;74:937–53. [48] Kuyper M, Harhangi HR, Stave AK, Winkler AA, Jetten MS, de Laat WT, et al. Highlevel functional expression of a fungal xylose isomerase: the key to efficient ethanolic fermentation of xylose by Saccharomyces cerevisiae? FEMS Yeast Res 2003;4:69–78. [49] Karhumaa K, Wiedemann B, Hahn-Hagerdal B, Boles E, Gorwa-Grauslund MF. Coutilization of L-arabinose and D-xylose by laboratory and industrial Saccharomyces cerevisiae strains. Microb Cell Fact 2006;5:18. [50] van Rooyen R, Hahn-Hagerdal B, La Grange DC, van Zyl WH. Construction of cellobiose-growing and fermenting Saccharomyces cerevisiae strains. J Biotechnol 2005;120:284–95.

155

156

CHAPTER 7  Ethanol Production

[51] Galazka JM, Tian CG, Beeson WT, Martinez B, Glass NL, Cate JHD. Cellodextrin Transport in Yeast for Improved Biofuel Production. Science 2010;330:84–6. [52] Ha SJ, Galazka JM, Kim SR, Choi JH, Yang X, Seo JH, et al. Engineered Saccharomyces cerevisiae capable of simultaneous cellobiose and xylose fermentation. Proc Natl Acad Sci U S A 2011;108:504–9. [53] Zhang M, Eddy C, Deanda K, Finkestein M, Picataggio S. Metabolic engineer ing of a pentose metabolism pathway in ethanologenic Zymomonas mobilis. Science 1995;267:240–3. [54] Deanda K, Zhang M, Eddy C, Picataggio S. Development of an arabinose-fermenting Zymomonas mobilis strain by metabolic pathway engineering. Appl Environ Microbiol 1996;62:4465–70. [55] Mohagheghi A, Evans K, Chou YC, Zhang M. Cofermentation of glucose, xylose, and arabinose by genomic DNA-integrated xylose/arabinose fermenting strain of Zymomonas mobilis AX101. Appl Biochem Biotechnol 2002;98–100:885–98. [56] Mohagheghi A, Evans K, Finkelstein M, Zhang M. Cofermentation of glucose, xylose, and arabinose by mixed cultures of two genetically engineered Zymomonas mobilis strains. Appl Biochem Biotechnol 1998;70–72:285–99. [57] Jeon YJ, Svenson CJ, Joachimsthal EL, Rogers PL. Kinetic analysis of ethanol production by an acetate-resistant strain of recombinant Zymomonas mobilis. Biotechnol Lett 2002;24:819–24. [58] Joachimstahl E, Haggett KD, Jang JH, Rogers PL. A mutant of Zymomonas mobilis ZM4 capable of ethanol production from glucose in the presence of high acetate concentrations. Biotechnol Lett 1998;20:137–42. [59] Ranatunga TD, Jervis J, Helm RF, McMillan JD, Hatzis C. Identification of inhibitory components toxic toward Zymomonas mobilis CP4(pZB5) xylose fermentation. Appl Biochem Biotechnol 1997;67:185–98. [60] Ingram LO, Aldrich HC, Borges AC, Causey TB, Martinez A, Morales F, et al. Enteric bacterial catalysts for fuel ethanol production. Biotechnol Prog 1999;15:855–66. [61] Wood BE, Ingram LO. Ethanol-Production from Cellobiose, Amorphous Cellulose, and Crystalline Cellulose by Recombinant Klebsiella-Oxytoca Containing Chromosomally Integrated Zymomonas-Mobilis Genes for Ethanol-Production and Plasmids Expressing Thermostable Cellulase Genes from Clostridium-Thermocellum. Appl Environ Microbiol 1992;58:2103–10. [62] Ingram LO, Conway T, Clark DP, Sewell GW, Preston JF. Genetic engineering of ethanol production in Escherichia coli. Appl Environ Microbiol 1987;53:2420–5. [63] Ohta K, Beall DS, Mejia JP, Shanmugam KT, Ingram LO. Genetic improvement of Escherichia coli for ethanol production: chromosomal integration of Zymomonas mobilis genes encoding pyruvate decarboxylase and alcohol dehydrogenase II. Appl Environ Microbiol 1991;57:893–900. [64] Yomano LP, York SW, Zhou S, Shanmugam KT, Ingram LO. Re-engineering Escherichia coli for ethanol production. Biotechnol Lett 2008;30:2097–103. [65] Yomano LP, York SW, Shanmugam KT, Ingram LO. Deletion of methylglyoxal synthase gene (mgsA) increased sugar co-metabolism in ethanol-producing Escherichia coli. Biotechnol Lett 2009;31:1389–98. [66] Wood BE, Yomano LP, York SW, Ingram LO. Development of industrial-mediumrequired elimination of the 2,3-butanediol fermentation pathway to maintain ethanol yield in an ethanologenic strain of Klebsiella oxytoca. Biotechnol Prog 2005;21:1366–72.

7.6 Perspectives

[67] Lynd LR, Elander RT, Wyman CE. Likely features and costs of mature biomass ethanol technology. Appl Biochem Biotechnol 1996;57–8:741–61. [68] Lynd LR. Overview and evaluation of fuel ethanol from cellulosic biomass: Technology, economics, the environment, and policy. Annu Rev Energy Environ 1996;21:403–65. [69] Wu Z, Lee YY. Inhibition of the enzymatic hydrolysis of cellulose by ethanol. Biotechnol Lett 1997;19:977–9. [70] Holtzapple M, Cognata M, Shu Y, Hendrickson C. Inhibition of Trichoderma reesei cellulase by sugars and solvents. Biotechnol Bioeng 1990;36:275–87. [71] Ghose TK, Tyagi RD. Rapid ethanol fermentation of cellulose hydrolysate, 2. product and substrate-inhibition and optimization of fermentor design. Biotechnol Bioeng 1979;21:1401–20. [72] Zacchi G, Axelsson A. Economic-evaluation of preconcentration in production of ethanol from dilute sugar solutions. Biotechnol Bioeng 1989;34:223–33. [73] Wyman CE. What is (and is not) vital to advancing cellulosic ethanol. Trends Biotechnol 2007;25:153–7. [74] Aden, A., Ruth, M., Ibsen, K., Jechura, J., Neeves, K., Sheehan, J., Wallace, B., Montague, L., Slayton, A., and Lukas, J. (2002) Lignocellulosic biomass to ethanol process design and economics utilizing co-current dilute acid prehydrolysis and enzymatic hydrolysis for corn stover. NREL/TP-510–32438. [75] Wright JD. Ethanol from biomass by enzymatic hydrolysis. Chem Eng Prog 1988;84:62–74. [76] Lynd LR, van Zyl WH, McBride JE, Laser M. Consolidated bioprocessing of cellulosic biomass: an update. Curr Opin Biotechnol 2005;16:577–83. [77] Olson DG, McBride JE, Shaw AJ, Lynd LR. Recent progress in consolidated bioprocessing. Curr Opin Biotechnol 2012;23:396–405. [78] Lu YP, Zhang YHP, Lynd LR. Enzyme-microbe synergy during cellulose hydrolysis by Clostridium thermocellum. Proc Natl Acad Sci U S A 2006;103:16165–9. [79] McBee RH. The characteristics of Clostridium thermocellum. J Bacteriol 1954;67:505–6. [80] Yang SJ, Kataeva I, Hamilton-Brehm SD, Engle NL, Tschaplinski TJ, Doeppke C, et al. Efficient Degradation of Lignocellulosic Plant Biomass, without Pretreatment, by the Thermophilic Anaerobe “Anaerocellum thermophilum” DSM 6725. Appl Environ Microbiol 2009;75:4762–9. [81] Sonnleitner B, Fiechter A. Advantages of using thermophiles in biotechnological processes: expectations and reality. Trends Biotechnol 1983;1. [82] Xu Q, Singh A, Himmel ME. Perspectives and new directions for the production of bioethanol using consolidated bioprocessing of lignocellulose. Curr Opin Biotechnol 2009;20:364–71. [83] Amore A, Faraco V. Potential of fungi as category I Consolidated BioProcessing organisms for cellulosic ethanol production. Renew Sustain Energy Rev 2012;16:3286–301. [84] Demain AL, Newcomb M, Wu JHD. Cellulase, clostridia, and ethanol. Microbiol Mol Biol Rev 2005;69:124. [85] Mai V, Lorenz WW, Wiegel J. Transformation of Thermoanaerobacterium sp. strain JW/ SL-YS485 with plasmid pIKM1 conferring kanamycin resistance. FEMS Microbiol Lett 1997;148:163–7. [86] Mai V, Wiegel J. Recombinant DNA applications in thermophiles. In: Demian AL, Davis JE, editors. Manual of industrial microbiology and biotechnology. 2nd ed. Washington, D. C: ASM Press; 1999. p. 511–9.

157

158

CHAPTER 7  Ethanol Production

[87] Tyurin MV, Lynd LR, Wiegel J. Gene transfer systems for obligately anaerobic thermophilic bacteria. In: Rainey FA, Oren A, editors. Extremophiles. London: Academic Press Ltd-Elsevier Science Ltd; 2006. p. 309–30. [88] Tyurin MV, Desai SG, Lynd LR. Electrotransformation of Clostridium thermocellum. Appl Environ Microbiol 2004;70:883–90. [89] Tripathi SA, Olson DG, Argyros DA, Miller BB, Barrett TF, Murphy DM, et al. Development of pyrF-based genetic system for targeted gene deletion in Clostridium thermocellum and creation of a pta mutant. Appl Environ Microbiol 2010;76:6591–9. [90] Shaw AJ, Covalla SF, Hogsett DA, Herring CD. Marker removal system for Thermoanaerobacterium saccharolyticum and development of a markerless ethanologen. Appl Environ Microbiol 2011;77:2534–6. [91] Shaw AJ, Hogsett DA, Lynd LR. Natural competence in Thermoanaerobacter and Thermoanaerobacterium species. Appl Environ Microbiol 2010;76:4713–9. [92] Tolonen AC, Chilaka AC, Church GM. Targeted gene inactivation in Clostridium phytofermentans shows that cellulose degradation requires the family 9 hydrolase Cphy3367. Mol Microbiol 2009;74:1300–13. [93] Heap JT, Cartman ST, Kuehne SA, Cooksley C, Minton NP. ClosTron-targeted mutagenesis. Methods Mol Biol 2010;646:165–82. [94] Heap JT, Kuehne SA, Ehsaan M, Cartman ST, Cooksley CM, Scott JC, et al. The ClosTron: Mutagenesis in Clostridium refined and streamlined. J Microbiol Methods 2010;80:49–55. [95] Heap JT, Pennington OJ, Cartman ST, Carter GP, Minton NP. The ClosTron: a universal gene knock-out system for the genus Clostridium. J Microbiol Methods 2007;70:452–64. [96] Shaw AJ, Jenney FE, Adams MWW, Lynd LR. End-product pathways in the xylose fermenting bacterium, Thermoanaerobacterium saccharolyticum. Enzym Microb Technol 2008;42:453–8. [97] Desai SG, Guerinot ML, Lynd LR. Cloning of L-lactate dehydrogenase and elimination of lactic acid production via gene knockout in Thermoanaerobacterium saccharolyticum JW/SL-YS485. Appl Microbiol Biotechnol 2004;65:600–5. [98] Shaw AJ, Podkaminer KK, Desai SG, Bardsley JS, Rogers SR, Thorne PG, et al. Metabolic engineering of a thermophilic bacterium to produce ethanol at high yield. Proc Natl Acad Sci U S A 2008;105:13769–74. [99] Argyros DA, Tripathi SA, Barrett TF, Rogers SR, Feinberg LF, Olson DG, et al. High Ethanol Titers from Cellulose by Using Metabolically Engineered Thermophilic, Anaerobic Microbes. Appl Environ Microbiol 2011;77:8288–94. [100] Jones RP. Biological principles for the effects of ethanol. Enzym Microb Technol 1989;11:130–53. [101] Ingram LO. Ethanol tolerance in bacteria. Crit Rev Biotechnol 1990;9:305–19. [102] Rani KS, Seenayya G. High ethanol tolerance of new isolates of Clostridium thermocellum strains SS21 and SS22. World J Microbiol Biotechnol 1999;15:173–8. [103] Shao X, Raman B, Zhu M, Mielenz JR, Brown SD, Guss AM, et al. Mutant selection and phenotypic and genetic characterization of ethanol-tolerant strains of Clostridium thermocellum. Appl Microbiol Biotechnol 2011;92:641–52. [104] Brown SD, Guss AM, Karpinets TV, Parks JM, Smolin N, Yang S, et al. Mutant alcohol dehydrogenase leads to improved ethanol tolerance in Clostridium thermocellum. Proc Natl Acad Sci U S A 2011;108:13752–7.

7.6 Perspectives

[105] van Zyl WH, Lynd LR, den Haan R, McBride JE. Consolidated bioprocessing for bioethanol production using Saccharomyces cerevisiae. Adv Biochem Eng Biotechnol 2007;108:205–35. [106] Vanarsdell JN, Kwok S, Schweickart VL, Ladner MB, Gelfand DH, Innis MA. Cloning, characterization, and expression in Saccharomyces cerevisiae of endoglucanase-I from Trichoderma reesei. Bio/Technology 1987;5:60–4. [107] Wood BE, Beall DS, Ingram LO. Production of recombinant bacterial endoglucanase as a co-product with ethanol during fermentation using derivatives of Escherichia coli KO11. Biotechnol Bioeng 1997;55:547–55. [108] Zhou SD, Yomano LP, Saleh AZ, Davis FC, Aldrich HC, Ingram LO. Enhancement of expression and apparent secretion of Erwinia chrysanthemi endoglucanase (encoded by celZ) in Escherichia coli B. Appl Environ Microbiol 1999;65:2439–45. [109] Zhou SD, Ingram LO. Simultaneous saccharification and fermentation of amorphous cellulose to ethanol by recombinant Klebsiella oxytoca SZ21 without supplemental cellulase. Biotechnol Lett 2001;23:1455–62. [110] Zhou SD, Davis FC, Ingram LO. Gene integration and expression and extracellular secretion of Erwinia chrysanthemi endoglucanase CelY (celY) and CelZ (celZ) in ethanologenic Klebsiella oxytoca P2. Appl Environ Microbiol 2001;67:6–14. [111] Su P, Liu CQ, Lucas RJ, Delaney SF, Dunn NW. Simultaneous Expression of Genes Encoding Endoglucanase and Beta-Glucosidase in Zymomonas-Mobilis. Biotechnol Lett 1993;15:979–84. [112] Lejeune A, Eveleigh DE, Colson C. Expression of an Endoglucanase Gene of Pseudomonas-Fluorescens Var Cellulosa in Zymomonas-Mobilis. FEMS Microbiol Lett 1988;49:363–6. [113] Katahira S, Mizuike A, Fukuda H, Kondo A. Ethanol fermentation from lignocellulosic hydrolysate by a recombinant xylose- and cellooligosaccharide-assimilating yeast strain. Appl Microbiol Biotechnol 2006;72:1136–43. [114] Fujita Y, Ito J, Ueda M, Fukuda H, Kondo A. Synergistic saccharification, and direct fermentation to ethanol, of amorphous cellulose by use of an engineered yeast strain codisplaying three types of cellulolytic enzyme. Appl Environ Microbiol 2004;70:1207–12. [115] Fujita Y, Takahashi S, Ueda M, Tanaka A, Okada H, Morikawa Y, et al. Direct and efficient production of ethanol from cellulosic material with a yeast strain displaying cellulolytic enzymes. Appl Environ Microbiol 2002;68:5136–41. [116] Cho KM, Yoo YJ, Kang HS. delta-Integration of endo/exo-glucanase and betaglucosidase genes into the yeast chromosomes for direct conversion of cellulose to ethanol. Enzym Microb Technol 1999;25:23–30. [117] Van Rensburg P, Van Zyl WH, Pretorius IS. Engineering yeast for efficient cellulose degradation. Yeast 1998;14:67–76. [118] Den Haan R, McBride JE, La Grange DC, Lynd LR, Van Zyl WH. Functional expression of cellobiohydrolases in Saccharomyces cerevisiae towards one-step conversion of cellulose to ethanol. Enzym Microb Technol 2007;40:1291–9. [119] Den Haan R, Rose SH, Lynd LR, van Zyl WH. Hydrolysis and fermentation of amorphous cellulose by recombinant Saccharomyces cerevisiae. Metab Eng 2007;9:87–94. [120] Jeon E, Hyeon JE, Suh DJ, Suh YW, Kim SW, Song KH, et al. Production of cellulosic ethanol in Saccharomyces cerevisiae heterologous expressing Clostridium thermocellum endoglucanase and Saccharomycopsis fibuligera beta-glucosidase genes. Mol Cells 2009;28:369–73.

159

160

CHAPTER 7  Ethanol Production

[121] Ilmen M, den Haan R, Brevnova E, McBride J, Wiswall E, Froehlich A, et al. High level secretion of cellobiohydrolases by Saccharomyces cerevisiae. Biotechnol Biofuels 2011;4:30. [122] Wen F, Sun J, Zhao HM. Yeast Surface Display of Trifunctional Minicellulosomes for Simultaneous Saccharification and Fermentation of Cellulose to Ethanol. Appl Environ Microbiol 2010;76:1251–60. [123] Tsai SL, Goyal G, Chen W. Surface Display of a Functional Minicellulosome by Intracellular Complementation Using a Synthetic Yeast Consortium and Its Application to Cellulose Hydrolysis and Ethanol Production. Appl Environ Microbiol 2010;76:7514–20.