Chapter 10

Enzyme Production Systems forBiomass ConversionJohn A. Howard, Zivko Nikolov, and Elizabeth E. Hood

10.1 Introduction

Plant biomass is a complex matrix of polymers comprising the polysaccharides cellulose andhemicellulose, as well as the phenolic polymer lignin, as the major structural components.Cellulose, the most abundant biopolymer on earth, is a simple, linear, β-1,4-linked polymerof glucose. However, its semi-crystalline structure is notoriously resistant to hydrolysis byboth enzymatic and chemical means. The cellulose in secondary cell walls is embeddedin a matrix of hemicellulose and lignin with lesser amounts of pectin and protein. Thesesecondary walls are the source of biomass targeted for conversion because of their abundantdry weight at maturity when compared to herbaceous plant materials. A strategy designedto use lignocellulose for fuel or bio-products must include the ability to efficiently convertthe polysaccharide and lignin components of plant cell walls to simple sugars and phenolicmonomers, respectively. Deconstruction of lignocellulose can be accomplished by heat andchemical means (Taherzadeh and Karimi, 2007) but the preferred environmentally friendlymethod is to use enzymes.

10.2 The Challenge: Volume and Cost of Enzymes Required

With the current state of technology for biomass conversion, the overwhelming enzyme re-quirement is for cellulases: endo-cellulase, exo-cellulase, and glucosidase (Merino and Cherry,2007). The specific activity of most cellulases is quite low (Jorgensen et al., 2007; Sticklen,2008b) and much effort has focused on increasing their activity levels. However, even withimproved enzymes and improved methods of production, the amount of cellulase required to

Plant Biomass Conversion, First Edition. Edited by Elizabeth E. Hood, Peter Nelson and Randall Powell.C© 2011 John Wiley & Sons Inc.

227

228 Plant Biomass Conversion

Table 10.1. Estimates of enzyme amounts necessary to digest fixed amounts of biomass.

Tons of BiomassBillions GallonsEthanol Possible

Enzymes PerGallon (g)

Tons ofEnzymes (MMa)

Cost of Enzymeper Gallonb ($)

425 million 36 100 3.6 0.1730 1.1 0.12

Assumptions: 85 gallons ethanol per ton; tons of enzymes required to produce that amount per year.Enzymes required per gallon will decrease with improvements. Thirty-six billion gallons will be producedby 2022.aMM, million.bNREL estimate; David Hsu.

deconstruct the volumes of biomass necessary for 30% replacement of gasoline are in themillions of tons. For the purpose of this chapter, we are using a base case of enzyme amountsand cost as shown in Table 10.1. At a loading level of 100 g enzyme per gallon of ethanolproduced, and a production goal of 36 billion gallons from the Bush mandate, 3.6 millionmetric tons of cellulase will be required. This is the amount per year! It is assumed for thisdiscussion that commercial scale microbial fermenters would hold 500,000 L and yield 50 gcellulase/L of culture broth per week (i.e., per batch), conditions that would allow productionof 1,250 metric tons of enzyme per year per fermenter, assuming 100% recovery. Under theseconditions, a minimum of 2,900 fermenters would be required to produce enough enzymeto saccharify 425 million tons of biomass. Moreover, if each fully equipped and installedfermenter costs about $10 million, the capital investment for fermenters alone would amountto $30 billion.

This is an unprecedented challenge in terms of the amount of enzymes and the extremely lowcost that is required for competitively priced ethanol. In addition, the amount of upfront capitalrequired for fermenter capacity is also problematic. If the enzyme load was 100 g/gallon at acost of $0.17/gallon, a total capital investment of over $57 billion dollars would be required toreturn gross annual revenues of ∼$6 billion. This situation has led many groups to investigateways to reduce this cost burden. There is every reason to believe that the cost will go down.While we have used this previously published model as the base case, continual reportsappear of improvements from many different groups on many different enzyme cost reductiontechnologies. This chapter will discuss the different approaches that are being considered tolower the overall cost and the impact that these approaches may have on reducing the costburden.

10.3 Theoretical Ways to Address the Challenge of Quantity ofEnzyme and Cost Requirements

There are four steps to the conversion of biomass to ethanol: (1) pretreatment to remove ligninand hemicellulose, (2) hydrolysis to allow saccharification of cellulose, (3) fermentation forethanol production, and (4) purification of ethanol and other products. The hydrolysis step usingcellulase enzymes accounts for 40% of current cost input (Miyamoto, 1997). Pretreatment isanother energy and cost hurdle, which can be addressed by an enzymatic or thermochemicalprocess. The pretreatment step has an impact on the amount and type of enzymes that are

10 Enzyme Production Systems for Biomass Conversion 229

required for saccharification (see Chapter 9, this volume). The focus of this chapter is on whatcan be done to each of the four steps that will have an impact on the enzyme cost.

10.3.1 Increase Susceptibility for Biomass Deconstruction

The plant cell wall is a complex network of cellulose fibrils coated and linked by hydrogenbonds to hemicellulose. Hemicelluloses are a heterogeneous group of linear and branchedpolysaccharides primarily composed of pentoses, predominantly xylose (Shallom and Shoham,2003). Hemicelluloses vary in composition between different plant sources, but often containside chains which can sterically block the accessibility of cellulose to cellulase and therebylimit digestion. The hemicellulose polymers are further covalently linked to lignin molecules,forming an insoluble and robust structure. The amount of lignin can vary with different typesof tissues as well (Campbell and Sederoff, 1996).

Saccharification of cellulose microfibrils prior to fermentation requires that the shieldinghemicelluloses and lignin be removed to increase cellulase access. Therefore, any treatmentthat reduces steric hindrance and allows greater access to cellulose can theoretically reducethe amount of enzymes required for saccharification.

Pretreatment of Biomass

A plethora of pretreatment paradigms have been reported (Mosier et al., 2004; Sun and Cheng,2002) and compared (Wyman et al., 2005) (see also Chapter 9 in this volume). Physicaland chemical methods including milling, high-temperature treatment with steam, acid, oralkali, as well as steam or alkali explosion, and biological pretreatment have been usedsuccessfully. Protocols include a thermochemical pretreatment that uses weak acid hydrolysisat an elevated temperature to remove hemicelluloses and expose cellulose fibrils to enzymaticdeconstruction. This treatment allows for limited sugar degradation and the potential forrecovery of components such as lignin, polylactic acid, and hemicellulose sugars for otherapplications (Ragauskas et al., 2006; Zhang et al., 2007).

Biologically based pretreatment protocols utilize white rot fungi of several genera (Sunand Cheng, 2002). Lignin-degrading enzymes are secreted extracellularly by many white rotfungi and fall into three groups: lignin peroxidases (EC 1.11.1.7, LiP), manganese perox-idases (EC 1.11.1.7; MnP), and laccase (EC 1.10.3.2). LiP is synthesized by a family ofabout ten closely related genes, lipA through lipJ, whose expression is affected by cultureconditions. MnP expression is regulated by the presence of Mn in culture. Both MnP and LiPhave been expressed in heterologous systems, and their activity tested following site-directedmutagenesis. Laccase genes are less well characterized, and their presence is not thought tobe essential for effective lignin degradation. All three enzymes act through oxidative, ratherthan hydrolytic, mechanisms and have acidic optima that make them useful for biomass pre-treatment under acidic conditions (Cullen, 2002). Laccases and ligninases are used in the pulpand paper industry for lignin removal. Delignification is a central process to the utilizationof lignocellulose, and therefore the genome of the model lignin-degrading white rot fungusPhanerochaete chrysosporium has been sequenced and methods for its genetic manipulationby homologous recombination and transformation by auxotrophic complementation have beendeveloped (Cullen, 2002). A strain of Sporotrichum pulverulentum lacking cellulase has beenproduced to prevent hydrolysis during delignification (Ander and Eriksson, 1977). Other delig-nifying enzymes are produced by white rot fungi that might speed up the process, but overall

230 Plant Biomass Conversion

the process remains slow. It is, however, energetically and environmentally less demandingthan thermochemical pretreatment.

The enzymes that degrade hemicellulose are as heterogeneous as the substrate, but areusually modular enzymes with a carbohydrate-binding module (CBM) and a catalytic domain.Many also have dockerin domains to bind the enzyme to the microbial cell surface or tothe conserved cohesin domains on cellulosomes (Bayer et al., 2007; Gilbert, 2007). Thebiotechnological applications of hemicellulases include biobleaching and biopulping in thepulp and paper industry, improvement of animal feed quality, as well as the processing offlour in the bakery industry (Shallom and Shoham, 2003). Currently, hemicellulases are notused for biomass degradation, but studies are being conducted on extension of their use toeconomic pretreatment (Galbe and Zacchi, 2002; Mielenz, 2001; Sticklen, 2008b). Further,hemicelluloses could be converted into constituent pentoses to be fermented into ethanol,which would increase overall ethanol yield. However, this requires different organisms thatferment pentoses (See Chapter 8 in this Volume).

The removal of lignin and hemicellulose from corn stover has been compared in batch andflow-through systems (Yang and Wyman, 2004). The effectiveness of flow-through systems atremoving lignin and hemicelluloses was generally higher than batch systems. Lignin removalgenerally increased cellulose digestibility, but its removal was not essential in batch systems.

In summary, the more effective the pretreatment is in exposing the cellulose and removinglignin and hemicelluloses, the lower the enzyme requirement for saccharification. The mosteffective pretreatment today involves thermochemical processing, which requires large capitalinvestments and produces environmental waste. At the other extreme is enzymatic pretreat-ment, which is environmentally friendly, but not as efficient with current technology. Thedilemma is that enzymatic pretreatment represents additional cost contraints on an alreadycost-prohibitive amount of enzymes used in saccharification. Therefore, while the long-termhope is to have a completely enzymatic process, in the short term, it appears that thermaland chemical pretreatments will need to be optimized to be compatible with the enzymaticsaccharification process (see Chapter 9).

Genetic Alteration of Feedstock

Plants show varying susceptibility to deconstruction, related to the levels of obstructive ligninand hemicelluloses. A survey of the composition of cell walls across plants (Sun and Cheng,2002) allows comparison of the best pretreatment and hydrolysis procedures to follow. Suscep-tibility of cell walls to degradation may vary because lignin content may vary even within thesame species, e.g., in trees (Campbell and Sederoff, 1996). Experiments have indicated thatrice hulls require a ten times higher concentration of a mixture of enzymes to effect breakdownas hardwood requires (Jimenez-Flores et al., 2010). This difference may reflect the presenceof materials such as silica that interfere with cellulose pretreatment and hydrolysis (Gressel,2007; Sainz, 2009; Zhu et al., 2009). Assessing digestibility based on composition will helpwith choosing the best substrates for hydrolysis as well as tailoring pretreatment and choosingthe cocktail of hydrolytic enzymes for optimal activity.

Selecting the most amenable source for enzymatic deconstruction is not always practical,especially when using agricultural by-products such as corn stover and rice hulls or woodby-products. Therefore, the potential to increase the digestibility of these biomass sourcesdirectly is desirable. Plant lines that have competitive agronomic characteristics but alsopossess increased digestibility of their by-products can be obtained. The two targeted areas

10 Enzyme Production Systems for Biomass Conversion 231

where this approach would have a major impact are in reducing the costs of pretreatment andsaccharification.

Plants can either be selected or engineered to be more susceptible to digestion (Campbelland Sederoff, 1996; Chen and Dixon, 2007). Attempts have been made to increase the amountof lignin-degrading enzymes in vegetative tissue in order to reduce the pretreatment burden.The challenge is to obtain plants with greater susceptibility to enzyme digestion but which stillperform well in the field. MnP expression in alfalfa led to stunted development and reducedyield of dry matter, although the plants did produce seed (Austin et al., 1995). In maize, theexpression of MnP correlated to tissue-specific effects. MnP expressed preferentially in seedshowed no signs of deleterious effects on plant health. In contrast, cytoplasmic targeting witha constitutive promoter caused lesions in older leaves, although germination and floweringwere not affected (Clough et al., 2006).

A decrease in lignin content was achieved in aspen by down-regulation of the coumarate:coenzyme A ligase (Pt4CL1), which led to a 45% decrease in lignin and a compensatory15% increase in cellulose content of the modified plant. This altered ratio favors bioethanolproduction (Li et al., 2003). Modifying lignin biosynthetic enzymes to lower lignin in cellwalls is an obvious way to reduce biomass recalcitrance, and sugar yields from modifiedalfalfa lines with lower lignin-forming enzymes were nearly double over wild type (Chen andDixon, 2007). Technologies have been developed to modulate levels of lignin biosyntheticenzymes in situ using antisense constructs in alfalfa (Reddy et al., 2005) and RNAi in maize(Sticklen, 2008a).

Biomass composition and enzymatic digestibility of plants can be altered by reducing thelevels of synthetic enzymes and increasing the levels of degradative enzymes. Reducingthe expression of poplar glycosyltransferase using RNA interference led to a reduction inthe glucuronoxylan content of poplar and consequently increased its digestibility by cellulase(Lee et al., 2009). Arabidopsis plants expressing a repressor derived from a secondary cellwall thickening-promoting factor (NST1) were twice as susceptible to enzymatic hydrolysis ascontrol plants (Iwase et al., 2009). By contrast, increasing expression in Arabidopsis of poplarcellulase caused modification of cellulose–hemicellulose crosslinks, leading to increased plantsize and improved cellulose hydrolysis (Park et al., 2003). Hartati et al. (2008) expressedpoplar cellulase in the leguminous plant, sengon. Transgenic plants showed increased growthand leaf size, attributed to paracrystalline cellulose disruption and less xyloglucan contentin the wall. The expression of several hydrolytic enzymes in poplar made the wood moresusceptible to enzymatic hydrolysis (Kaida et al., 2009). However, one must bear in mind thatthermochemical pretreatment of transgenic plants containing enzymes may lead to inactivationof the endogenous enzyme, with the potential need to add exogenous cellulases followingpretreatment (Oraby et al., 2007).

10.3.2 Decrease Exogenous Enzyme Load

The deconstruction of microcrystalline cellulose is far from a simple chemical reaction. Theprocess requires the activity of several different enzymes, thereby complicating a simpleone-solution method for improvement. However, this complexity also provides the potentialfor improvements in many different ways, each of which are discussed separately below.Ultimately, many of these approaches can be combined to provide additive or even synergisticeffects to increase the overall activity of cellulase action.

A minimum of three categories of enzymes in the glycosyl hydrolase superfamily arerequired for deconstruction of cellulose after the biomass has undergone pretreatment. They

232 Plant Biomass Conversion

are endoglucanase (EC 3.2.1.4), exoglucanase (also called cellobiohydrolase; EC 3.2.1.91),and β-glucosidase (EC 3.2.1.21). Endoglucanases hydrolyze internal β-1,4 glycosidic bondswithin a cellulose microfibril. The fibrils are then exposed to the action of exoglucanasesthat processively trim cellobiose units from exposed ends, again by hydrolysis of the β-1,4glycosidic bond. Finally, β-glucosidase converts cellobiose and cellotriose units to glucose. β-Glucosidase shows severe end-product inhibition and therefore optimal activities are achievedupon glucose removal either by physical methods or by fermentation to ethanol (Andric et al.,2010). These enzymes work in synergy, that is, the combined effect on cellulose deconstructionis greater than the individual effects added together. Therefore, any consideration of economicalconversion will benefit from optimizing this synergistic activity. This process has been testedwidely, and many studies testify to the value of using purified, reconstituted cellulase mixturesover crude extracts from a single organism (Gusakov et al., 2007; Rosgaard et al., 2007) aswell as tailoring cellulase mixtures to biomass source (reviewed further below).

Cellulase enzymes are modular, with domains linked by a flexible, glycosylated, hingeregion, whose length is important to optimal cellulase activity. The catalytic domain variesin shape between endoglucanases and exoglucanases. Endoglucanases have an open catalyticsite, and exoglucanases have a tubular catalytic groove to fit the ends of the cellulose fibril(Henrissat, 1994). All the enzymes share a carbohydrate binding domain (CBD) on the N- or C-terminus of the catalytic domain that allows the enzymes to adsorb to insoluble cellulose. Site-directed mutagenesis has shown that surface aromatic amino acids flanked by polar residueswithin the binding cleft are important for tight binding (Kormos et al., 2000). Tryptophaninstead of tyrosine increases affinity of cellobiohydrolase from the four families of CBDs sofar identified. These differing affinities for substrate could be manipulated using site-directedmutagenesis to optimize binding properties for different substrates, i.e., different crystallineforms of cellulose as it goes through deconstruction. Some cellulases have two CBDs and thisorganization synergistically increases affinity of the enzyme for cellulose, and CDBs can bemutagenized to modulate broader or altered enzyme specificity (Linder and Teeri, 1997). Thus,cellulase can be modified to optimize specificity by promoting synergism between enzymes, byvarying the length of the linker region, modifying amino acids in the CBD to increase substratespecificity and increasing binding strength by increasing number or affinity of CBDs.

Cellulolytic enzymes are synthesized and secreted outside the cell by fungi such as Tri-choderma reesei or into membrane-bound complexes called cellulosomes in bacteria such asClostridium thermocellum (Bayer et al., 1998, 2004). Enzymes from different sources vary intheir pH and temperature optima, inducibility, stability, and other parameters (Cantarel et al.,2009) (see also www.cazy.org). These factors need to be accounted for when using multipleenzymes to degrade cellulose. In some organisms, cellulolytic activity may depend on thepresence of a single gene (Tolonen et al., 2009). Disruption of the cphy3367 gene for its onlyfamily 9 glycosyl hydrolase in Clostridium phytofermentans abolished cellulolytic activity de-spite the presence and even elevated levels of other CAZy glycosyl hydrolases, indicating thatthere was an obligate requirement for Cphy3367 for cellulose metabolism to even occur. Thus,it is not enough to have the best single enzyme of the class but rather the best complement ofenzymes that can work together, a condition that must be established empirically.

Tailor Enzyme Cocktails to Biomass Source

The variety of possible enzymes for use in deconstruction protocols is not uniform acrosssources of biomass, suggesting that enzyme cocktails should be varied to fit the substrate

10 Enzyme Production Systems for Biomass Conversion 233

(Zhang and Lynd, 2004). Attention also should be paid to pretreatment and optimal celluloseadsorption by enzymes to maximize deconstruction based on the composition of the specificsubstrate. In other words, one can combine the optimal mixture of enzymes with the specificsubstrate to decrease enzyme load. For example, the composition of cellulose, hemicellulose,and lignin fractions from different sources varies significantly (Sun and Chen, 2002) and wouldbe deconstructed with different kinetics even if the same pretreatment and enzyme mixtureswere used.

In addition to using different enzyme cocktails for different biomass sources, each sourceshould be separately evaluated as to the optimal amounts of each enzyme to be used in itsdeconstruction. This concept has been assessed using SO2-impregnated spruce and aspenwood (Breuil et al., 1990). It was shown that although aspen wood was completely hydrolyzedby any of five commercial cellulase preparations, spruce wood contained inhibitory agentsthat prevented complete hydrolysis. Enzymatic pretreatment of spruce might increase thehydrolysis profile by removing some inhibitory agents, such as lignin that might competitivelyadsorb cellulases (Zhang and Lynd, 2004). The same lab showed previously that removal ofend products of hydrolysis was important to achieving optimal substrate hydrolysis (Chanet al., 1989).

On a related issue, the rate of hydrolysis decreases with time, even when taking into accountthe effect of end-product inhibition and enzyme deactivation (Eriksson et al., 2002). Both thesurface area and cellulose adsorptive capacity have been shown to decrease with hydrolysisover time. Hydrolysis rates may be maintained by periodic addition of new substrate (Zhangand Lynd, 2004) and by varying CBD sites on enzymes (Carrard et al., 2000). Enzymesfrom Chrysosporium lucknowense were used to demonstrate improved cellulose hydrolysisfrom reconstituted purified enzymes over crude extracts (Gusakov et al., 2007). These authorsalso demonstrated that modifying the mixture to contain enzymes with poor CBM overcamethe pseudo-inactivation of cellulases during hydrolysis, presumably due to greater ability tomigrate over the surface, thus clearing obstacles on the cellulose.

Zhou et al. (2009) compared a crude cellulase extract from a Trichoderma viride T 100-14mutant strain with purified enzymes from the same strain in various combinations to identifythe optimal amounts of each enzyme for glucose production from steam exploded corn stover.The optimal enzyme combination produced 2.1 times more glucose than the crude extract.This type of manipulation to obtain maximum conversion will be useful in an industrial settingto maximize efficiency and lower cost.

Obtain New Sources of Enzyme-Encoding Genes

In spite of the progress in optimization of hydrolysis, cellulases constitute a relatively high costfor the bioconversion process and account for up to 40% of the total biomass conversion cost(Miyamoto, 1997; Zhang and Lynd, 2004). Since the 1970s, screening has led to the discoveryof many sources of cellulase from fungi, termites, aerobic and anaerobic bacteria, and evensymbiotic microorganisms from pinfish transitioning from a carnivorous to herbivorous diet(Luczkovich and Stellwag, 1993). Novel screening methods include isolation of novel cellulaseenzymes from soil metagenome libraries (Rondon et al., 2000). This process makes it feasibleto isolate enzymes from organisms that are refractory to cultivation.

Degenerate PCR primers were used for screening DNA of fungal communities in corn stoverfor the presence of glycosyl hydrolase family genes (Jacobsen et al., 2005). This rationale ispowerful, given that only organisms that are able to efficiently use lignocellulosics as a carbon

234 Plant Biomass Conversion

source would grow on it. This strategy was also applied to leaf litter and soil from oak-dominated and sugar maple-dominated hardwood forests (Edwards et al., 2008) to screen fororganisms containing sequences similar to the exoglucanase gene, cbhI. Novel sequences wererecovered, indicating that this strategy holds promise for identifying new cellulolytic genesfrom natural communities. Another natural cellulase digesting community is the rumen ofherbivores (Coleman, 1985; Teather and Wood, 1982), and several cellulase genes have beencloned from these sources (Cappa et al., 1997; Ekinci et al., 2002).

Increase the Specific Activity of Cellulolytic Enzymes

An approach that has worked exceptionally well for other enzymes is to structurally engineerthem to increase catalytic activity, thereby reducing the amount of the enzymes required andeffectively lowering the overall cost. Approaches to increasing the specific activity of cellulasehave focused on understanding the catalytic process. The CAZY database (www.cazy.org)stores data regarding structural features and carbohydrate binding domains of a number ofcellulases (Cantarel et al., 2009). Several enzymes have been crystallized and the active sitesexamined following mutagenesis of conserved residues (Zhang et al., 2006). In addition tomodification of the catalytic site, specific activity can be modified by increasing the affinityof the enzyme for the substrate through altering residues in the carbohydrate binding domain(CBD) or increasing the number of the enzyme’s CBDs (Linder and Teeri, 1997). Two specificapproaches, directed evolution and rational design, are considered below.

Directed Evolution

DNA shuffling using PCR is a powerful way to randomly modify the structure of enzymes,which are later screened for activity (Rabinovich et al., 2002). Some authors (Kim et al.,2000) have utilized the process of DNA shuffling to create a library of cellulase genes thatare expressed on the surface of bacteria as fusions to the ice nucleation protein for screeningon carboxymethylcellulose (CMC) as substrate. Improvements of up to 5-fold in CMCaseactivity were obtained by this method, and sequencing indicated that amino acid changes inthe catalytic site were responsible for increased specific activity. This approach was used (Kaperet al., 2002) to shuffle the nonhomologous sequences of two different hyperthermostable β-glucosidases with temperature optima of 105◦C and 95◦C to obtain a more active enzymewith a more moderate optimum of 70◦C. Another approach (Heinzelman et al., 2009) usingstructure-guided recombination created chimeras of three class II cellobiohydrolases. Theresultant chimeras showed higher thermostability than the most stable parent, indicating thatthis technique could be used to generate a large combinatorial library of enzymes with desiredproperties. Techniques such as these may help to manipulate pH and temperature optima forenzymes for industrial applications. Another group (Toyama et al., 2003) used the alternativeprocess of nuclear shuffling in swollen T. reesei conidia to achieve autopolyploidy usingcolchicine treatment. Successive treatment with benomyl and ethylmethanesulfonate producedhyperproducers, but these were slow growing compared to the original strain (Toyama andToyama, 2001), which may be a problem for commercial production.

Rational Design

In contrast to directed evolution where a variety of products are generated by DNA shufflingand then screened for activity, rational design uses targeted approaches to modifying enzymes.

10 Enzyme Production Systems for Biomass Conversion 235

Table 10.2. Improvement in enzyme specific activity of cellulolytic enzymes.

Enzyme Improvement Method Improvement Reference

Cel9A endo-exoglucanase fromThermobifida fusca

Rational design/computer modeling

40% activity Escovar-Kousenet al., 2004

Endo-β -1,4-glucanasefrom B. subtilis BSE616

DNA shuffling 5-fold activity offusion; 2.2-foldactivity of free enzyme

Kim et al., 2000

β-D-glucosidase fromThermotoga neapolitana

Directed evolution 31% activity McCarthy et al., 2004

Cellobiohydrolase fromTalaromyces emersonii inS. cerevisiae

Rational design/structure-guidedengineering of S–Sbonds

Thermostabilityincreased by 9◦C;improved activity atRT

Voutilainen et al., 2009

Cel B from Pyrococcusfuriosus

DNA shuffling 1.5- to 3.5-foldthermostability at70◦C

Kaper et al., 2002

Lac S from Sulfolobussolfataricus

DNA shuffling 3.5- to 8.6-foldincreased lactosehydrolysis

Kaper et al., 2002

Cellobiohydrolase Cel7BMelanocarpusalbomyces in S.cerevisiae

Random mutagenesis 2-fold increasedactivity at 70◦C

Voutilainen et al., 2007

B. subtilisendo-β-1,4-glucanase(Cel5A)

Directed evolution(error-prone PCR)

2.03- to 2.68-fold Lin et al., 2009

Availability of crystallographic or site-directed mutagenesis data allows understanding of thestructure of the catalytic site of cellulases (Zhang et al., 2006) and provides the basis for arational design approach to optimize the interaction of the enzyme with the substrate. Thecrystal structure of the catalytic domains of several, predominantly bacterial, hemicellulasesand cellulases has been achieved (Davies and Henrissat, 1995). Molecular modeling can then beused to compute and analyze modifications in structure that might improve binding or catalysis.Such an approach was utilized by Kormos et al. (2000) to identify and modify amino acids inthe CBM binding groove and identified tyrosines 19 and 85 in the groove as essential for tightbinding of CBDN1 of endoglucanase C from Cellulomonas fimi. Amino acids in the catalyticsite of endoglucanase CelC of Clostridium thermocellum that contributed to catalytic activitywere identified (Escovar-Kousen et al., 2004; Navas and Beguin, 1992). Computer modelingwas also used to identify conserved amino acids in active sites of an endo/exoglucanase thatwere then mutated to obtain 40% improvement in activity (Escovar-Kousen et al., 2004).

Table 10.2 shows some of the enzymes whose activity was modified by directed evolutionor rational design. However, the maximum increase in activity is still a relatively modest2- to 5-fold despite the intense effort. Specific activity may be slightly more improved byoptimizing the length of linker and the number of CBM domains, but if commercial productionis the target, other approaches must be considered to improve the economics of biomasshydrolysis.

236 Plant Biomass Conversion

Synergistic Proteins That Increase Cellulase Activity

Increasing the efficiency of cellulases by the action of other proteins is another way by whichconversion of cellulose to fermentable sugars can be increased. Expansins are a group ofproteins that “loosen” cell walls during turgor-driven wall expansion, fertilization, and growth(Cosgrove, 1999, 2005) probably by weakening noncovalent tethering of cellulose fibrilsthrough xyloglucan bridges. Cell wall loosening immediately suggests a procedure to increasecellulase access in biomass, but the mechanism of action of expansins remains unclear. Thetwo categories of expansins, A and B, share domain structure, have acidic pH optima, and havea conserved putative CBD and an endoglucanase-like (EG-45) domain, although no hydrolyticactivity of expansins has been detected.

Polysaccharide hydrolases are another category of enzymes that may weaken the matrixof the plant cell wall, exposing the cellulose fibrils for deconstruction. It appears that mostpolysaccharide hydrolases act on the hemicellulose crosslinks of the wall, and specifically onthe xylan and the mixed linkage 1,3:1,4-β-glucan seen primarily in grasses, which comprise alarge portion of agricultural biomass (Cosgrove, 2005, 2000, 1999). These have an acidic pHoptimum and could conceivably be used in conjunction with acid pretreatment.

Other reactions that may be beneficial include utilization of xyloglucan endotransglycosy-lases (XETs) and ascorbate. XETs are plant enzymes that catalyze a rearrangement reactionof the xyloglucan network in the plant cell wall and are a proposed part of cell wall loos-ening (Cosgrove, 1999). Hydroxyl radicals produced by treatment with ascorbate reduce theviscosity of xyloglucan solutions and may expose the cellulose fibrils in cell walls by makingthe hemicellulose matrix less rigid (Cosgrove, 1999). This reaction may be carried out inconjunction with the xylan hydrolase reaction as part of an acid pretreatment protocol.

Protease pretreatment may loosen the cell wall matrix by digesting the protein components.Large quantities of the protease enzyme trypsin are procurable at relatively low cost byextracting and purifying either from the native source or from recombinant proteins madein plants (Horn et al., 2004; Woodard et al., 2003,), but care must be taken to inactivate theenzyme to avoid digestion of cellulolytic enzymes in subsequent steps.

In summary, the addition of several different types of enzymes can aid in lowering theamount of cellulase required for the conversion of biomass. The utility of this approach willdepend on the cost of the additional proteins compared to the cost of the cellulase. Severalcommercially important enzymes have been produced in plants (Hood and Howard, 2008)and indicate a path towards expression of several cell-wall-modifying enzymes in plants forbiomass utility.

10.3.3 Increase Accumulation of Enzymes in Production Host

There is an inverse correlation between the expression level of enzymes in their host andthe cost of production. Therefore, a key strategy is to increase the overall accumulation ofthe enzymes in the host tissue to decrease the overall cost. The National Renewable EnergyLaboratory has a target to reduce the cost of ethanol production to $1.07/gallon (in 2002dollars) by the year 2012—a target that is only one year away. A second target is to produce 60billion gallons of bioethanol/year by the year 2030 (NREL/BR-510-40742). These aggressivetargets demand aggressive approaches to reducing enzyme production costs. While all of themethods discussed above are likely to be helpful in achieving the goal, the overall cost ofenzyme production will likely be a key factor for any solution.

10 Enzyme Production Systems for Biomass Conversion 237

Microbial Production Systems

Both prokaryotic bacteria and eukaryotic fungi produce cellulases, and the enzymes showconsiderable overlap in structure and function (Gilbert and Hazlewood, 1993). A primary dif-ference exists between aerobic producers and anaerobic producers, however. Most anaerobesproduce tightly associated and aggregated enzymes called cellulosomes, which are associatedwith membranes through dockerin molecules, whereas aerobes produce soluble, secreted en-zymes. Temperature optima vary based on source: many thermophilic bacteria produce highlyefficient cellulases, but their temperature optima make them uneconomical for hydrolysis.

Optimization of conditions for high cellulase enzyme synthesis has been studied usingvarious substrates, inducers, fermentation conditions, and microbial systems. One approachused Trichoderma reesei with Solid Substrate Fermentation on wheat bran and manipulatedtemperature and moisture conditions to obtain a 6.2-fold increase in activity over controls(Singhania et al., 2007). Other parameters that have been tested are batch vs. continuousfermentation, substrate concentration, fungal vs. bacterial systems, and shake vs. stationarycultures (Bailey and Tahtiharju, 2003; Bailey et al., 1972; Gomes et al., 1992; Jana et al.,1994; Kadam and Keutzer, 1995; Nikolov et al., 2000; Schaffner and Toledo, 1991; Singhaniaet al., 2007; Spear et al., 1993; Tholudur et al., 1999; Xia and Shen, 2004; Zhou et al., 2009;Zhuang et al., 2007). The enormous literature on manipulation of individual parameters makesit clear that conditions have to be established separately for each organism, and either crudeor reconstituted purified mixtures of cellulases tested for activity. The production optimum isnot necessarily the optimum for activity.

Screening strategies for high-producing strains rely on quantitative assays based on productaccumulation, substrate reduction, or change in physical properties of substrates (reviewed inZhang et al., 2006). These may be conveniently carried out on solid media and enzyme activityinferred by substrate digestion producing haloes. Chromogenic substrates for evaluation ofenzyme activity, high throughput and high-density microtiter plates, and PCR cyclers withhot lids to prevent evaporation from samples are among the facilitated screening tools thatallow rapid screening of large numbers of samples (reviewed in Zhang et al., 2006). Selectionstrategies to identify the highest producers are slower and use methods such as cross-feedingwhere soluble enzymes diffuse around colonies growing in resource-limited areas exhibitingcompetition to identify high-producing strains (Zhang et al., 2006). A colorimetric microplateassay to screen virulent plant pathogens for cellulolytic activity allows high throughput, quanti-tative screening of fungal extracts, allowing more efficient bio-prospecting for enzymes (Kinget al., 2009). Organisms identified by such screening strategies can then be evaluated furtherby other methods.

Plants to Provide Supply of Enzymes

Producing exogenous enzymes in microbes is efficient and is the main method used todayfor the production of most industrial enzymes. Microbial fermentation for the recovery ofcommercial amounts of cellulase requires high capital costs for setup and operational costs forrunning. Thus, investigation into the use of plants as an alternate production system for trans-genic proteins for biomass conversion has developed. Plants need little infrastructure beyondensuring that transgenic plants are contained appropriately, even possibly within dedicatedethanol-producing areas (Howard and Hood, 2007).

There are several theoretical reasons why plants may be the best source for the long-termsupply of exogenous enzymes, including (1) plants represent the least expensive method to

238 Plant Biomass Conversion

produce protein in general, (2) plants do not require the large amount of capital for productioncompared to microbial fermenters, (3) plant production can be scaled-up or -down withoutmajor changes in infrastructure, (4) almost any plant can function as a production system,(5) proteins can be targeted to specific compartments, allowing for increased accumulationin the desired tissue with little interference in other tissues (reduces potential toxicity to thecell), (6) plants have convenient storage, transport, and processing of component materials,and (7) plants have the potential to have lines containing different enzymes combined throughcrossing (Howard and Hood, 2005; Sainz, 2009; Twyman et al., 2003). Sainz (2009) providesa table of cellulases that have been expressed so far in plants. The choice of plant species andtissue for accumulation of foreign proteins depends on the specific transgene to be expressed,production system employed, transportation needs, and processing and storage requirements.Transformation, expression, and purification technologies also affect the cost of the productsproduced in plants (Howard and Hood, 2005, 2007).

Subcellular targeting is critical for accumulation and protein integrity of cellulase or otherindustrial enzymes (Clough et al., 2006; Hood et al., 2007). Other factors that affect proteinaccumulation include (1) the use of the catalytic domain alone, which supports a higher levelof expression than the whole enzyme (Sticklen, 2006); (2) dual targeting to two compartmentsis more efficient in some cases than one (Hyunjong et al., 2006); and (3) optimization of codonusage (Xue et al., 2003). Although the catalytic domain alone was optimal for protein accumu-lation (Sticklen, 2006), for functionality, the catalytic and binding domains are both required.Targeting enzymes to subcellular compartments may overcome toxicity, permit correct fold-ing, and enable glycosylation. The chloroplast is another desired location because of ease ofcontainment. Using downstream box fusions of Thermobifida fusca cellulase in chloroplasts oftransgenic tobacco caused increased expression up to 10.7% TSP. This is significantly higherthan the 1% accumulation seen with nuclear-targeted fusions (Gray et al., 2009). However,homoplastic transformation protocols are still elusive for most plants (Grevich and Daniell,2005; Sticklen, 2008b).

Proteins produced in plants can accumulate at significant levels, and in addition, the lig-nocellulosic by-products can themselves be used for production of ethanol. For example, aheterologous endo-1,4 β-glucanase E1 gene from Acidothermus cellulolyticus under controlof a cauliflower mosaic virus 35S promoter in maize was used to obtain active enzyme at1.13% TSP. Total soluble protein from transgenic maize allowed AFEX-pretreated corn stoverto be directly processed into glucose (Ransom et al., 2007) .

Plants with underground storage tubers can be induced to accumulate cellulase in the leafby chloroplast targeting, while the tubers can be used as food. An Acidothermus cellulolyticusendoglucanase E1 gene was placed under the control of an RbcS-3C promoter, alfalfa mosaicvirus 5’-untranslated leader, and RbcS-2A signal peptide (Dai et al., 2000) and showed proteinaccumulation up to 2.6% TSP in the leaf.

In addition to simply using plants as a production vehicle in the way microbes are used,other more integrated approaches have been developed. One attractive strategy is to have theenzymes produced from by-products or from tissue normally discarded from existing cropplants (Howard and Hood, 2007). The germ tissue from maize, a by-product of many grainethanol facilities, was used to demonstrate high accumulation levels of E1 (16% TSP) cellulaseas well as cellobiohydrolase (18% TSP) (Hood et al., 2007). Endoglucanase expression from anapoplast-directed protein, expressed from a constitutive promoter, was obtained at high levelsin rice without apparent deleterious effects. Transgenic rice extracts were active in convertingpretreated rice and maize biomass into ethanol at 22% and 30%, respectively, compared to62% and 95% conversion using purified commercial enzyme (Oraby et al., 2007). These last

10 Enzyme Production Systems for Biomass Conversion 239

several examples show great promise in that it allows the crop to be harvested for food andthe rest of the plant to be used for production of cellulase enzymes after harvest.

In addition to cellulases, plants have been used to produce other potentially useful proteinsfor biomass conversion such as ligninases (Clough et al., 2006; Hood et al., 2003) and xylanase(Borisjuk et al., 1999). Transgenic production of trypsin as a zymogen has been successfullyachieved in maize tissue, a system that provides cost effective transport, storage, and use(Woodard et al., 2003). As mentioned above, in addition to being cost-effective sources forprotein production, transgenic plant lines can be crossed to other transgenic lines to allow theproduction of several enzymes in the same plant.

Another option is to express degradative enzymes in vegetative tissue to decrease the burdenof adding exogenous cellulase. This would increase the effectiveness of cellulase activity byworking from the inside of cell wall tissue, while the outer tissue is exposed to exogenous cel-lulase. Cellulose hydrolytic enzymes have been engineered into potato, tobacco, Arabidopsis,rice, barley, and maize. Many of these proteins were targeted for expression to specific tissuesor sub-cellular compartments to overcome potential toxicity (Sticklen, 2006; Torney et al.,2007). In order to be of use for saccharification, these enzymes must be used either with anenzymatic pretreatment, be extracted prior to thermochemical pretreatment, or be stable underthermochemical pretreatment conditions. The required changes to integrate this approach withdownstream processing limit the rapid acceptance for incremental improvements, but this maychange if progress continues in the future toward less harsh pretreatment conditions, in whichmore of an enzymatic approach is integrated.

Supply Live Microbes

Currently, enzymes from fungi (Ramamurthy et al., 1992) or bacteria (Gilbert and Hazlewood,1993) are used in crude or purified, reconstituted mixtures for the aerobic saccharification step.Rather than add the enzymes directly, it is possible to add live cellulase-producing organisms forthe initial aerobic saccharification followed by addition of yeasts for the anaerobic fermentationprocesses (Mamma et al., 1996). As microbes can be finicky about growing on differentsubstrates, this approach will require more attention than the simple addition of enzymes.It will also require that the products of the aerobic saccharification reaction be compatiblewith the organism used for fermentation. This approach, however, does offer a way to greatlyreduce the costs of making the enzymes in separate fermentation facilities if these factors canbe adequately controlled.

Simultaneous Saccharification and Fermentation

The process of deriving ethanol from lignocellulosic feedstock requires that carbohydratepolymers be deconstructed and fermented to ethanol. This process can be carried out underseveral paradigms. In Separate Hydrolysis and Fermentation (SHF), the hydrolysis step iscarried out aerobically and the products separately fermented by anaerobes. This processis inefficient because of the separate chambers required for each process and also becauseproducts of hydrolysis that accumulate can be inhibitory to the hydrolytic enzymes (Taherzadehand Karimi, 2007).

Simultaneous Saccharification and Fermentation (SSF) refers to cellulose hydrolysis carriedout in the presence of a fermentative organism to relieve end-product inhibition by removalof glucose through fermentation (Sun and Cheng, 2002; Taherzadeh and Karimi, 2007).

240 Plant Biomass Conversion

The theoretical advantages of SSF over SHF are many: increase in hydrolysis rate due toremoval of inhibitory sugars by fermentation, shorter process time, and less capital cost due tosingle reactor usage. In addition, enzyme and sterility requirements can also be reduced (Sunand Cheng, 2002). Simultaneous Saccharification and Co-Fermentation (SSCF) increasesthe theoretical ethanol yield by incorporating organisms that ferment pentose to ethanol.Mixed cultures of hydrolytic and fermentative strains can be enhanced by modifying hexosefermenting strains (see Chapter 8 in this volume). This has been achieved in Zymomonasmobilis and Saccharomyces cerevisiae to also ferment pentose (Hahn-Hagerdal et al., 2007;Olofsson et al., 2008; Rudolf et al., 2008; Zhang et al., 1995), thus increasing ethanol yieldfrom biomass.

More recently, the concept of Consolidated Bioprocessing (CBP), reviewed in Lynd et al.(2005), has been garnering a lot of interest because of the potential for drastically reducingproduction cost. CBP incorporates biological production of cellulase, biomass hydrolysis, andsugar fermentation, all in a single step. Therefore, the basic requirement is the development ofCBP-enabling microorganisms with high levels of protein expression such as S. cerevisiae orE. coli, or at least a compatible mixed culture, and engineering in cellulose utilization at highefficiencies. Many of the suggested organisms either naturally or through modification fermentsugars. Lynd et al. (2005) proposed the utilization of Cellulose–Enzyme–Microbe complexesas opposed to Cellulose-Enzyme (CE) complexes, citing the higher rates of hydrolysis withthe former, but in principle, this procedure could be adapted to both, depending on the kineticsobserved. In addition, using biomass feedstocks that are engineered for low recalcitrance willprovide extra benefits.

SSF, SSCF, and CBP (Table 10.3) can be carried out in batch, fed-batch (semi-continuous),or continuous cycles. The advantages of fed-batch and continuous cycles are the relieffrom buildup of sugars, which severely inhibit cellulase enzymes. However, since hydrol-ysis is kinetically slower than fermentation, alcohol-tolerance of strains used is also aconsideration.

10.4 Cost of Producing Exogenous Enzymes

There are many types of enzyme production systems in use today, including microbial cultures,animal cell cultures, submerged plant cultures, row crops, Baculovirus and many more. Eachof these can play an important role for protein production and each has inherent limitations.To reach the cost targets for cellulase, only row crops and microbes are inherently at a lowenough cost to be viable for cellulase production.

In addition to the inherent cost of the host, the cost of producing a specific protein isinversely proportional to the ability of the host to accumulate that protein. Therefore, a hostthat can accumulate large amounts of the desired protein will be able to produce the protein fora lower cost. The production practices of both microbes and row crops have been optimizedfor decades, and while they continue to enjoy incremental improvements, it is unlikely thatthere will be a quantum change in their cost structure. Improvements therefore must come ineither their ability to accumulate the desired protein or other efficiencies related to the overallintegration of the process.

In the case of microbial production systems, the lowest cost of cellulase enzymes is byextraction from fungal cultures. Years of experience using these cultures have allowed opti-mization for cellulase production. In the near term, we should expect that these fungal systems

10 Enzyme Production Systems for Biomass Conversion 241

Table 10.3. Fermentation procedures.

Type of Treatment Advantages Disadvantages Comments References

Simulataneoussaccharificationand fermentation(SSF)

Single reactor;improved ratesand yields

Sub-optimalconditions;ethanol-inhibitedfermentation;difficulty recyclinghydrolytic enzymesand fermentativeorganisms;separate C5 and C6fermentation

Conversion ratestypically 30%higher in SSF thanSHF using; fedbatch usage limitsproblems withinhibitors, mass,and heat transfer

Drissen et al.,2008; Olofssonet al., 2008b

Simultaneoussaccharificationandco-fermentation(SSCF)

Single reactor,higher yieldxylose, andglucoseconversion

Suboptimalconditions; xyloseuptake needs lowglucoseconcentrations foruptake; lowtemperatures forfermentation do notfavor hydrolyticenzymes fromthermophiles;needs heat-tolerantyeast strains

78% of theoreticalyield of (35 g/L)ethanol obtainedwith 7% solidloading infed-batch using S.cerevisiaeTMB340 strain

Olofsson et al.,2008a, 2008b

Consolidated bioprocessing (CBP)

Lower costs,higher efficiency

Requiresengineeredhydrolytic strains tobe ethanol tolerant;possible nontargetmetaboliteproduction

Lynd et al., 2005

will continue to provide the lowest cost cellulase enzymes. While there are continual im-provements being made as to strains that accumulate higher levels of enzymes, it is unlikelythat huge improvements will be seen from the already very high current levels. Likewise, theoptimization of the production practices are likely to provide incremental improvements butunlikely to have a major effect in driving the cost down because of the maturity of the system.It is more likely that microbial production systems will rely on synergistic proteins or increasesin specific activity to help drive cost down by decreasing the protein load for bioconversion.

In contrast to microbial production, relatively little experience has been gained with accu-mulating cellulase in plants and no commercial production of cellulase using plants that existstoday. Plant extracts as an inexpensive enzyme formulation will require a well-conceptualizedand developed bioprocess. In the near term, the most realistic way to produce and formulatebiomass hydrolytic enzymes will be from crude aqueous plant extracts of leaves, seed, stalks,or their fractions (Sainz, 2009). Changes in downstream biomass processing would not berequired and would be compatible with the current system of using microbial extracts.

Direct addition of dry fractionated plant tissue may be the most desirable approach be-cause of simple processing, storage stability, and low cost associated with water additionand subsequent drying of residual spent transgenic biomass. The extent of processing of

242 Plant Biomass Conversion

plant material containing biomass-degrading enzymes will depend on whether or not the planttissue is amenable to dry or wet fractionation (pressing fresh juice is also wet fractionation)without undergoing aqueous extraction.

Direct application of fractionated transgenic plant tissue in saccharification of pretreatedbiomass hinges on enzyme concentration in the transgenic plant fraction. For example, atenzyme loading of 8 g/kg biomass (20 g/kg cellulose), the amount of associated plant materialshould not exceed more than 20% of pretreated biomass concentration because the increaseof total solids in the saccharification reaction mix could adversely affect cellulose hydrolysis,viscosity, nonproductive binding of enzymes, inhibition, etc. In this case, the amount ofhydrolytic enzyme(s) should be at least 4% of dry wt. (40 g/kg) of fractionated transgenicplant tissue. Note that not all plant tissues are equally practical for direct application. Forexample, soybeans would be expensive for direct application as soy protein (meal) is the majorrevenue and feed source. The same is true for whole corn seed, unless extremely high enzymeaccumulation levels are reached. Thus, seed crops must be extracted to recover the enzymefrom by-products, allowing the seed to be sold for its originally intended use to capture thecredits. To eliminate drying and transportation cost of extracted enzyme, corn mills, corn-toethanol plants, and soy crushers should be an integral part of biomass conversion, which maynot be a trivial stipulation.

The conclusion from this discussion is that expression of enzymes in tissue or residue withlittle or no by-product value would be the ideal candidates for direct delivery. Examples ofthese include rice straw, corn stover, corn germ, and switchgrass. Of course, the key is toachieve accumulation levels of at least 4% dry wt.

10.4.1 Cost Analysis

Taking into account what we know about production cost, we have made some generalizationsabout the enzyme cost and potential cost in the future. We have used models for productioncost described below, knowing that there are continual improvements in enzyme activity asmentioned in the sections above that will affect the absolute cost. Any increase in specificactivity or synergism with cellulase should apply equally to both microbes and plants, thereforethe hope is these numbers will show the relative merits of the two systems and be a guide tothe absolute costs.

To determine the amount of enzyme per gallon ethanol and unit enzyme cost ($/kg protein)required to achieve our target cost of $0.17/gal ethanol (Table 10.1), we assumed enzymeloading of 12 FPU/g cellulose. Using enzyme specific activity of 600 FPU/g, this enzymeloading is equivalent to 20 g enzyme/kg cellulose or 8 g enzyme/kg biomass, assuming anaverage of 40% cellulose in biomass (Mosier et al., 2004). At 85 gal ethanol/ton biomass, thenecessary amount of enzyme per gallon ethanol produced is 94 g/gal (100 g/gal in Table 10.1).Now, to achieve the near-term target of $0.17, one has to be able to produce cellulases for lessthan $2.0/kg enzyme.

Cost of Cellulases Produced by Microbial Fermentation

The best cost models for production are undoubtedly in private companies that keep thisinformation confidential. We have used a variety of public information sources to arriveat our cost for production models, knowing these numbers will continue to change with

10 Enzyme Production Systems for Biomass Conversion 243

Table 10.4. Cost estimate of enzyme produced by microbial fermentation.a

Fermentation capacity (cu.m.) 2,600Fermentation titer (g/L) 50Batch cycle (days) 7Recovery yield (%) 85Product per week (ton) 111Number of batches/year 50Plant output per year 5,525Fermentation (variable and fixed) cost ($/kg) 6.0Recovery (variable and fixed) cost ($/kg) 4.2Total enzyme production cost ($/kg) 10.2

aHepner and Assoc., 1995

improvements. The hope is that these models can be used as a guide to compare relative trendsin the industry.

A 2002 NREL report (NREL/TP-510-32438) showed that 2000 tons/day would be anoptimal size of a lignocellulosic biomass conversion facility; the facility would utilize 10% ofavailable corn acres and haul stover from within a 50 miles radius. At 8 kg enzyme loading perton biomass, an enzyme production facility capable of supplying 16 tons of enzyme per day(5,600 tons enzyme per year) has to be built on site to eliminate transportation and minimizeenzyme formulation and stabilization cost. If enzyme is produced by a fungal fermentationprocess that lasts 7 days and yields 50 g cellulase/L, the required fermentation capacity wouldbe 2.2 MM L per week. A fermentation facility with 5 fermenters of 500,000 L each would costabout $50 MM. The enzyme facility (centrifugation, concentration, and polishing) to process2.2 MM L of fermentation broth per week would probably require an additional investment ofup to $40 MM. The 2022 mandate of producing 36 billion gallons of ethanol translates to (36× 100/gal) a total of 3.6 MM metric tons of enzyme (Table 10.1). With 5,600 tons of enzymeper facility, 640 fermentation facilities have to be built; thus, the total capital investment forenzyme production would amount to approximately $57 billion (640 × 50 MM = 32 MM forfermentation + 640 × 40 MM = 25.6 MM for processing). A model of the cost breakdownof enzyme produced by fermentation is given above (Table 10.4). The total cost based on thismodel shows microbial production to be at approximately $10/kg with the assumptions ofprotein loading from above.

Cost of Cellulase Produced in Transgenic Plants

The bioprocessing challenge for transgenic tissue with enzymes is to deliver formulatedenzyme preparations to a saccharification facility at the lowest possible cost. Protein andtissue stability, tissue fractionation, protein extraction and formulation, as well as storage andtransportation all add to this cost. For example, if transportation costs of $14/ metric ton(NREL/TP-510-32438) were assumed, then transporting an enzyme formulation that contains5% w/w enzyme would already cost $0.28/kg enzyme. In addition, any wet fractionation suchas fresh tissue pressing, aqueous extraction of dry plant tissue followed by residue separationwould add at least $1/kg to the bioprocessing cost. Therefore, in order to get the enzyme costof less than $2/kg, production of transgenic plant material should not cost more than $1.00/kgenzyme, which translates to raw material cost of $100/dry metric ton for material containing10% enzyme.

244 Plant Biomass Conversion

Table 10.5. Cost estimate of enzyme produced from defatted corn germ.

Defatted corn germ price ($/kg germ) 0.1Enzyme expression level (% dry base) 1.5Extraction and recovery yield (%) 85Enzyme output per year (ton) 5,600Cost of enzyme produced in corn germ ($/kg enzyme) 7.8Recovery (variable and fixed) cost ($/kg enzyme) 7.0Byproduct creditsResidual germ cake at $55/ton ($/kg enzyme) 4.0Total enzyme production cost ($/kg) 10.8

This calculation clearly exemplifies the cost and processing constraints imposed on en-zyme production from transgenic plants. Thus, to achieve production cost targets for biomassenzymes, the following considerations must be employed: (1) eliminate transportation costsby integrating enzyme processing into biomass conversion facility, (2) minimize fractiona-tion/extraction cost of transgenic material, and (3) reduce the contribution of transgenic plantmaterial to enzyme production cost by capturing plant biomass value through by-productcredits or cellulose.

One of the most crucial assumptions today for plants is how well can the tissue accumulatethe cellulase enzymes. Little experience is yet achieved with plants as compared to microbialcultures to know what the uppermost accumulation levels can be. Therefore, the model usedfor plants is based on the level of accumulation required for plants to be at par with the costof microbial production and then estimating the necessary accumulation level. Table 10.5provides an example of cellulase produced in corn germ and extracted at comparable costwith microbes. In this case, the level of protein accumulation required is 1.5 % of germ dryweight. While this seems intuitively reasonable, it has also been achieved (Hood and Howard,unpublished results), making this a realistic case for maize germ as well as potentially forother row crops.

There is every reason to believe that expression levels will continue to improve in plantsas it is a relatively young science. For this reason, we developed a future case where thecellulase represents 7.5 % of germ dry weight or 25% of the total soluble protein. On the basisof accumulation of proteins in other systems, this seems realistic and especially feasible forseed proteins where the vast majority of these serve only to provide amino acid reservoirs forgermination. At this level of accumulation, the cost of extracted enzyme drops to $2.50/kg.

If instead of extracting the germ we can use it directly, the model predicts that cellulaseaccumulation levels of 1.5% dry weight are needed to match the comparable microbiallyproduced cellulase. In addition, there is the benefit of a much lower capital investment. Asdiscussed above, however, this may require addition of too many solids, and therefore eventhough the cost is comparable, it may not be practical and the level may need to be as high as4% to become practical.

If the level of cellulase reaches 7.5% dry weight, then it is well below the limitations foradditional solids. The cost drops to $1.50/kg, making this the most economical scenario forbiomass conversion and below the cost target. This level of expression has been reached in thelaboratory and improvements in expression technology continue in this relatively new field.

The other practical consideration is the acreage required to grow crops to produce thisamount of enzyme. We modeled this concept previously, considering the proximity limitationsof the lignocellulosic biomass to the ethanol facility to avoid large transportation costs. The

10 Enzyme Production Systems for Biomass Conversion 245

Table 10.6. Comparison of production systems for 5,600 metric tons of enzyme per year.

System Source

ManufacturingCost Estimates($/kg) Capital Investment Comments

Microbial Microbialfermentation50 g/L fungus

$10 ∼$100 MM All aspects feasibletoday. Needimprovements tolower protein loadingto meet cost targets

Extraction fromseed w/by-product(current)

Extraction fromcorn germ@ 1.5% of drywt

$10 ∼$40 MM Expression levelsreached but notscaled up orcommercialized

Extraction fromseed w/by-product(future)

Extraction fromcorn germ@7.5%of dry wt

$2.50 <$20 MM Can approach costtargets without majorimprovementslowering proteinloading

Seedby-product(current)

Direct use ofcorn germ@1.5% of dry wt

$7.8 <$2 MM Can meet cost targetswithout improvementsin lowering proteinloading but may notbe practical becauseof high solidpercentage

Seedby-product(future)

Direct use ofcorn germ@ 7.5% dry wt

$1.50 <$2 MM The lowest cost ifexpression levels canbe raised to theirpotential. No otherpractical limitations

Extraction fromvegetative tissue

Extraction fromrice leaves@ 0.1% dry wt

10 ∼$40 MM Must be extractedprior to pretreatmentto keep activity (Orabyet al., 2007)

study demonstrated that if cellulase levels were 0.1% of the dry weight of the seed (1% ofgerm dry weight), it was more than sufficient to keep the acreage of the cellulase crop lessthan the acreage needed to supply the lignocellulosic biomass (Howard and Hood, 2007).

Table 10.6 shows the cost factors above along with that of a previously discussed rice modelto illustrate cellulase coming from the vegetative crop (Oraby et al., 2007). In summary, plantproduction systems that accumulate cellulase in the normally unused or low value portionof the plants are now approaching competitive cost structure with microbial systems. Asexpression technology improves and cellulase reaches 4% of the dry weight, direct delivery ofplant tissue can be the system of choice and will easily reach the current cost targets.

10.5 Summary and Future Prospects

The current production of bioethanol from starch produces the ethical dilemma of “foodor fuel,” but the use of agricultural residues (Chapter 2 in this volume) and other

246 Plant Biomass Conversion

lignocellulosic biomass (Chapters 3–5 in this volume) can potentially resolve this issue. How-ever, lignocellulosic biomass is considerably more recalcitrant to digestion than starch andrequires pretreatment and large amounts of enzymes for its deconstruction, which are expen-sive using current production methods. The complex problem of converting biomass to fuel isstill in its infancy. The near-term will clearly rely on microbial production systems. However,none of the current systems can meet the target enzyme cost today using the assumptions weemployed in our models. In order to meet the cost target, improved facility design, process,organisms, enzymes, titer, quantity, recovery, and efficiency must be addressed simultane-ously. Because of the magnitude of the problem, the most likely outcome will not be a singlebreakthrough but rather combining the outcomes of several different lines of research. As anexample, increased susceptibility of feedstocks along with increased synergy of enzymes willdecrease enzyme loading, as will increases in specific activity. Companies such as Novozymesand Genencor reportedly have reduced the cost of enzymes necessary for producing a gallon ofethanol from $5.40 in 2000 to $0.20 in 2005 (Moreira, 2005). Despite this reported improve-ment, enzymes still represent a key stumbling block to bioconversion and a practical limitationto achieving this theoretical cost because of the large capital investment needed. Microbialproduction of enzymes will need to rely more on decreased protein loading than on increasein accumulation to reach lower costs. In this regard, the use of plants selected to be moresusceptible to digestion can also help lower the cost requirements for exogenous enzymes.

In the long term, plants may also provide the lowest cost of exogenous enzymes comingeither from seed by-products or from vegetative tissue. While plants can also benefit from anydecreases in protein loading similarly to microbes, plants can also lower cost by increasingaccumulation of the enzymes and by better integration of the overall process. One can imaginea fully integrated and self-contained system where a crop such as maize provides the stoverfor the cellulose biomass designed to be susceptible to digestion and the enzymes required fordigestion contained in both the stover and the germ by-product. The endosperm (starch fraction)can continue to be used as it is today to produce grain ethanol. Plants have the potential tobecome enzyme production systems, and the ultimate one-stop shop for the efficient productionof cellulase enzymes as well as the substrate for deconstruction to bioethanol.

References

Ander, P. & Eriksson, K. 1977. Selective degradation of wood components by white-rot fungi.Physiologia Plantarum, 41, 239–248.

Andric, P., Meyer, A., Jensen, P. & Dam-Johansen, K. 2010. Reactor design for minimizingproduct inhibition during enzymatic lignocellulose hydrolysis: I. Significance and mecha-nism of cellobiose and glucose inhibition on cellulolytic enzymes. Biotechnology Advances,28, 308–324.

Austin, S., Bingham, E., Mathews, D., Shahan, M., Will, J. & Burgess, R. 1995. Productionand field performance of transgenic alfalfa (Medicago sativa L.) expressing alpha-amylaseand manganese-dependent lignin peroxidase. Euphytica, 85, 381–393.

Bailey, M. J. & Tahtiharju, J. 2003. Efficient cellulase production by Trichoderma reeseiin continuous cultivation on lactose medium with a computer-controlled feeding strategy.Applied Microbiology and Biotechnology, 62, 156–162.

Bailey, P. J., Keilich, G. & Husemann, E. 1972. Isolation and characterisation of cellulase(beta-1,4-glucan 4-glucanohydrolase) from Chaetomium globosum Kunze. Material undOrganismen, 7, 29–43.

10 Enzyme Production Systems for Biomass Conversion 247

Bayer, E., Belaich, J., Shoham, Y. & Lamed, R. 2004. The cellulosomes: Multienzyme ma-chines for degradation of plant cell wall polysaccharides. Annual Review of Microbiology,58, 521–554.

Bayer, E. A., Chanzy, H., Lamed, R. & Shoham, Y. 1998. Cellulose, cellulases and cellulo-somes. Current Opinion in Structural Biology, 8, 548–557.

Borisjuk, N., Borisjuk, L., Logendra, S., Petersen, F., Gleba, Y. & Raskin, I. 1999. Pro-duction of recombinant proteins in plant root exudates. Nature Biotechnology, 17, 466–469.

Breuil, C., Chan, M. & Saddler, J. N. 1990. Comparison of the hydrolytic activity of commercialcellulase preparations. Applied Microbiology and Biotechnology, 34, 31–35.

Campbell, M. & Sederoff, R. 1996. Variation in lignin content and composition (mechanismsof control and implications for the genetic improvement of plants). Plant Physiology,110, 3.

Cantarel, B., Coutinho, P., Rancurel, C., Bernard, T., Lombard, V. & Henrissat, B. 2009. Thecarbohydrate-active enZymes database (CAZy): An expert resource for glycogenomics.Nucleic Acids Research, 37, D233.

Cappa, F., Riboli, B., Rossi, F., Callegari, M. L. & Cocconcelli, P. S. 1997. Construction of novelRuminococcus albus, strains with improved cellulase activity by cloning of Streptomycesrochei endoglucanase gene. Biotechnology Letters, 19, 1151–1155.

Carrard, G., Koivula, A., Soderlund, H. & Beguin, P. 2000. Cellulose-binding domains promotehydrolysis of different sites on crystalline cellulose. Proceedings of the National Academyof Sciences of the United States of America, 97, 10342–10347.

Chan, M., Breuil, C., Schwald, W. & Saddler, J. N. 1989. Comparison of methods for quantify-ing the hydrolytic potential of cellulase enzymes. Applied Microbiology and Biotechnology,31, 413–418.

Chen, F. & Dixon, R. 2007. Lignin modification improves fermentable sugar yields for biofuelproduction. Nature Biotechnology, 25, 759–762.

Clough, R. C., Pappu, K., Thompson, K., Beifuss, K., Lane, J., Delaney, D. E., Harkey, R.,Drees, C., Howard, J. A. & Hood, E. E. 2006. Manganese peroxidase from the white-rotfungus Phanerochaete chrysosporium is enzymatically active and accumulates to high levelsin transgenic maize seed. Plant Biotechnology Journal, 4, 53–62.

Coleman, G. S. 1985. The cellulase content of 15 species of entodiniomorphid protozoa,mixed bacteria and plant debris isolated from the ovine rumen. The Journal of AgriculturalScience, 104, 349–360.

Cosgrove, D. J. 1999. Enzymes and other agents that enhance cell wall extensibility. AnnualReview of Plant Physiology Plant Molecular Biology, 50, 391–417.

Cosgrove, D. J. 2000. Expansive growth of plant cell walls. Plant Physiology and Biochemistry,38, 109–124.

Cosgrove, D. J. 2005. Growth of the plant cell wall. Nature Reviews Molecular Cell Biology,6, 850–861.

Cullen, D. 2002. Molecular genetics of lignin-degrading fungi and their applications inorganopollutant degradation. In: Kempken, F. V. E. (ed.) The Mycota XI Agricultural Appli-cations. 1st edn. Berlin, Heidelberg, New York: Springer-Verlag.

Dai, Z., Hooker, B., Anderson, D. & Thomas, S. 2000. Improved plant-based production ofE1 endoglucanase using potato: Expression optimization and tissue targeting. MolecularBreeding, 6, 277–285.

Davies, G. & Henrissat, B. 1995. Structures and mechanisms of glycosyl hydrolases. Structure,3, 853–859.

248 Plant Biomass Conversion

Drissen, R., Maas, R., Tramper, J. & Beeftink, H. 2008. Modelling ethanol production fromcellulose: separate hydrolysis and fermentation versus simultaneous saccharification andfermentation. Biocatalysis and Biotransformation, 27, 27–35.

Edwards, I., Upchurch, R. & Zak, D. 2008. Isolation of fungal cellobiohydrolase I genes fromsporocarps and forest soils by PCR. Applied and Environmental Microbiology, 74, 3481.

Ekinci, M. S., Martin, J. C. & Flint, H. J. 2002. Expression of a cellulase gene, celA, fromthe rumen fungus Neocallimastix patriciarum in Streptococcus bovis by means of promoterfusions. Biotechnology Letters, 24, 735–741.

Eriksson, T., Karlsson, J. & Tjerneld, F. 2002. A model explaining declining rate in hydrolysisof lignocellulose substrates with cellobiohydrolase I (Cel7A) and endoglucanase I (Cel7B)of Trichoderma reesei. Applied Biochemistry and Biotechnology, 101, 41–60.

Escovar-Kousen, J. M., Wilson, D. & Irwin, D. 2004. Integration of computer modeling andinitial studies of site-directed mutagenesis to improve cellulase activity on Cel9A fromThermobifida fusca. Applied Biochemistry and Biotechnology, 113, 287–297.

Galbe, M. & Zacchi, G. 2002. A review of the production of ethanol from softwood. AppliedMicrobiology and Biotechnology, 59, 618–628.

Gilbert, H. & Hazlewood, G. 1993. Bacterial cellulases and xylanases. Microbiology, 139,187–194.

Gilbert, H. J. 2007. Cellulosomes: Microbial nanomachines that display plasticity in quaternarystructure. Molecular Microbiology, 63, 1568–1576.

Gomes, I., Gomes, J., Steiner, W. & Esterbauer, H. 1992. Production of cellulase and xylanaseby a wild strain of Trichoderma viride. Applied Microbiology and Biotechnology, 36,701–707.

Gray, B. N., Ahner, B. A. & Hanson, M. R. 2009. High-level bacterial cellulase accumulationin chloroplast-transformed tobacco mediated by downstream box fusions. Biotechnologyand Bioengineering, 102, 1045–1054.

Gressel, J. 2007. Transgenics are imperative for biofuel crops. Plant Science, 174, 246–263.

Grevich, J. & Daniell, H. 2005. Chloroplast genetic engineering: Recent advances and futureperspectives. Critical Reviews in Plant Sciences, 24, 83–107.

Gusakov, A., Salanovich, T., Antonov, A., Ustinov, B., Okunev, O., Burlingame, R., Emalfarb,M., Baez, M. & Sinitsyn, A. 2007. Design of highly efficient cellulase mixtures for enzymatichydrolysis of cellulose. Biotechnology and Bioengineering, 97, 1028–1038.

Hahn-Hagerdal, B., Karhumaa, K., Fonseca, C., Spencer-Martins, I. & Gorwa-Grauslund,M. 2007. Towards industrial pentose-fermenting yeast strains. Applied Microbiology andBiotechnology, 74, 937–953.

Hartati, S., Sudarmonowati, E., Park, Y. W., Kaku, T., Kaida, R., Baba, K. I. & Hayashi, T.2008. Overexpression of Poplar Cellulase Accelerates Growth and Disturbs the ClosingMovements of Leaves in Sengon. Plant Physiology, 147, 552–561.

Heinzelman, P., Snow, C., Wu, I., Nguyen, C., Villalobos, A., Govindarajan, S., Minshull, J.& Arnold, F. 2009. A family of thermostable fungal cellulases created by structure-guidedrecombination. Proceedings of the National Academy of Sciences, 106, 5610.

Henrissat, B. 1994. Cellulases and their interaction with cellulose. Cellulose, 1, 169–196.Hood, E.E. & Howard, J.A. 2008. Over-expression of novel proteins in maize. In: Kriz, A.

& Larkins, B. (eds.) Molecular Genetic Approaches to Maize Improvement. Heidelberg:Springer-Berlin; pp. 91–105.

Hood, E. E., Bailey, M. R., Beifuss, K., Magallanes-Lundback, M., Horn, M. E., Callaway,E., Drees, C., Delaney, D. E., Clough, R. & Howard, J. A. 2003. Criteria for high-level

10 Enzyme Production Systems for Biomass Conversion 249

expression of a fungal laccase gene in transgenic maize. Plant Biotechnology Journal, 1,129–140.

Hood, E. E., Love, R., Lane, J., Bray, J., Clough, R., Pappu, K., Drees, C., Hood, K. R., Yoon,S., Ahmad, A. & Howard, J. A. 2007. Subcellular targeting is a key condition for high-levelaccumulation of cellulase protein in transgenic maize seed. Plant Biotechnology Journal,5, 709–719.

Horn, M., Woodard, S. & Howard, J. 2004. Plant molecular farming: Systems and products.Plant Cell Reports, 22, 711–720.

Howard, J. A. & Hood, E. 2005. Bioindustrial and biopharmaceutical products produced inplants. Advances in Agronomy, 85, 91–124.

Howard, J. A. & Hood, E. E. 2007. Methods for growing nonfood products in transgenicplants. Crop Science, 47, 1255–1262.

Hyunjong, B., Lee, D. & Hwang, I. 2006. Dual targeting of xylanase to chloroplasts and perox-isomes as a means to increase protein accumulation in plant cells. Journal of ExperimentalBotany, 57, 161.

Iwase, A., Hideno, A., Watanabe, K., Mitsuda, N. & Ohme-Takagi, M. 2009. A chimeric NSTrepressor has the potential to improve glucose productivity from plant cell walls. Journal ofBiotechnology, 142, 279–284.

Jacobsen, J., Lydolph, M. & Lange, L. 2005. Culture independent PCR: An alternative enzymediscovery strategy. Journal of Microbiological Methods, 60, 63–71.

Jana, S. K., Ghosh, V. K. & Singh, A. 1994. Production and hydrolytic potential of cellulaseenzymes from a mutant strain of Trichoderma reesei. Biotechnology and Applied Biochem-istry, 20, 233–239.

Jorgensen, H., Kristensen, J. B. & Felby, C. 2007. Enzymatic conversion of lignocellulose intofermentable sugars: Challenges and opportunities. Biofuels, Bioproducts and Biorefining,1, 119–134.

Kadam, K. L. & Keutzer, W. J. 1995. Enhancement in cellulase production by Trichodermareesei Rut-C30 due to citric acid. Biotechnology Letters, 17, 1111–1114.

Kaida, R., Kaku, T., Baba, K., Oyadomari, M., Watanabe, T., Nishida, K., Kanaya, T., Shani,Z., Shoseyov, O. & Hayashi, T. 2009. Loosening xyloglucan accelerates the enzymaticdegradation of cellulose in wood. Molecular Plant, 2, 904–909.

Kaper, T., Brouns, S., Geerling, A., De Vos, W. & Van Der Oost, J. 2002. DNA family shufflingof hyperthermostable beta-glycosidases. Biochemical Journal, 368, 461.

Kim, Y., Jung, H. & Pan, J. 2000. Bacterial cell surface display of an enzyme library for selectivescreening of improved cellulase variants. Applied and Environmental Microbiology, 66,788.

King, B., Donnelly, M., Bergstrom, G., Walker, L. & Gibson, D. 2009. An optimized microplateassay system for quantitative evaluation of plant cell wall-degrading enzyme activity offungal culture extracts. Biotechnology Bioengineering, 102, 1033–1044.

Kormos, J., Johnson, P., Brun, E., Tomme, P., Mcintosh, L., Haynes, C. & Kilburn, D. 2000.Binding site analysis of cellulose binding domain CBD (N1) from endoglucanse C ofCellulomonas fimi by site-directed mutagenesis. Biochemistry, 39, 8844–8852.

Lee, C., Teng, Q., Huang, W., Zhong, R. & Ye, Z. 2009. Down-regulation of PoGT47Cexpression in poplar results in a reduced glucuronoxylan content and an increased wooddigestibility by cellulase. Plant and Cell Physiology, 50, 1075.

Li, L., Zhou, Y., Cheng, X., Sun, J., Marita, J., Ralph, J. & Chiang, V. 2003. Combinatorial mod-ification of multiple lignin traits in trees through multigene cotransformation. Proceedingsof the National Academy of Sciences of the United States of America, 100, 4939.

250 Plant Biomass Conversion

Lin, L., Meng, X., Liu, P., Hong, Y., Wu, G., Huang, X., Li, C., Dong, J., Xiao, L. & Liu, Z.2009. Improved catalytic efficiency of Endo- -1, 4-glucanase from Bacillus subtilis BME-15by directed evolution. Applied Microbiology and Biotechnology, 82, 671–679.

Linder, M. & Teeri, T. 1997. The roles and function of cellulose-binding domains. Journal ofBiotechnology, 57, 15–28.

Luczkovich, J. & Stellwag, E. 1993. Isolation of cellulolytic microbes from the intestinal tractof the pinfish, Lagodon rhomboides: Size-related changes in diet and microbial abundance.Marine Biology, 116, 381–388.

Lynd, L., Zyl, W., McBride, J. & Laser, M. 2005. Consolidated bioprocessing of cellulosicbiomass: An update. Current Opinion in Biotechnology, 16, 577–583.

Mamma, D., Koullas, D., Fountoukidis, G., Kekos, D., Macris, B. & Koukios, E.1996. Bioethanol from sweet sorghum: Simultaneous saccharification and fermenta-tion of carbohydrates by a mixed microbial culture. Process Biochemistry, 31, 377–381.

McCarthy, J., Uzelac, A., Davis, D. & Eveleigh, D. 2004. Improved Catalytic Efficiency andActive Site Modification of 1, 4- -d-Glucan Glucohydrolase A from Thermotoga neapolitanaby Directed Evolution. Journal of Biological Chemistry, 279, 11495.

Merino, S. T. & Cherry, J. 2007. Progress and challenges in enzyme development for biomassutilization. Advances in Biochemical Engineering and Biotechnology, 108, 95–120.

Mielenz, J. R. 2001. Ethanol production from biomass: Technology and commercializationstatus. Current Opinion in Microbiology, 4, 324–329.

Miyamoto, K. 1997. Renewable biological systems for alternative sustainable energy produc-tion (FAO Agricultural Services Bulletin-128). Molecular biology, 4, 3.

Moreira, N. 2005. Growing Expectations. Science News, 168, 218–220.Mosier, N. S., Wilker, J. J. & Ladisch, M. R. 2004. Rapid chromatography for evaluating

adsorption characteristics of cellulase-binding domain mimetics. Biotechnology and Bio-engineering, 86, 756–764.

Navas, J. & Beguin, P. 1992. Site-directed mutagenesis of conserved residues of Clostridiumthermocellum endoglucanase CelC. Biochemical and Biophysical Research Communica-tions(Print), 189, 807–812.

Nikolov, T., Bakalova, N., Petrova, S., Benadova, R., Spasov, S. & Kolev, D. 2000. An effectivemethod for bioconversion of delignified waste-cellulose fibers from the paper industry witha cellulase complex. Bioresource Technology, 71, 1–4.

Olofsson, K., Bertilsson, M. & Liden, G. 2008. A short review on SSF–an interesting processoption for ethanol production from lignocellulosic feedstocks. Biotechnology for Biofuels,1, 7.

Olofsson, K., Rudolf, A. & Liden, G. 2008. Designing simultaneous saccharification andfermentation for improved xylose conversion by a recombinant strain of Saccharomycescerevisiae. Journal of Biotechnology, 134, 112–120.

Oraby, H., Venkatesh, B., Dale, B., Ahmad, R., Ransom, C., Oehmke, J. & Sticklen, M.2007. Enhanced conversion of plant biomass into glucose using transgenic rice-producedendoglucanase for cellulosic ethanol. Transgenic Research, 16, 739–749.

Park, Y., Tominaga, R., Sugiyama, J., Furuta, Y., Tanimoto, E., Samejima, M., Sakai, F. &Hayashi, T. 2003. Enhancement of growth by expression of poplar cellulase in Arabidopsisthaliana. Plant Journal, 33, 1099–1106.

Rabinovich, M. L., Mel’nik, M. S. & Bolobova, A. V. 2002. [Cellulases from microorganisms].Prikl Biokhim Mikrobiol, 38, 355–373.

10 Enzyme Production Systems for Biomass Conversion 251

Ragauskas, A., Williams, C., Davison, B., Britovsek, G., Cairney, J., Eckert, C., Frederick Jr,W., Hallett, J., Leak, D. & Liotta, C. 2006. The path forward for biofuels and biomaterials.Science, 311, 484.

Ramamurthy, V., Thacker, S. P. & Kothari, R. M. 1992. Optimized protocol for the pilot-scalepreparation of fungal cellulase. Journal of Industrial Microbiology, 9, 121–125.

Ransom, C., Balan, V., Biswas, G., Dale, B., Crockett, E., Sticklen, M. 2007. Heterologousacidothermus cellulolyticus 1, 4-beta-endoglucanase E1 produced within the corn biomassconverts corn stover into glucose. Applied Biochemistry and Biotechnology, 137, 207–219.

Reddy, M., Chen, F., Shadle, G., Jackson, L., Aljoe, H. & Dixon, R. 2005. Targeted down-regulation of cytochrome P450 enzymes for forage quality improvement in alfalfa (Medicagosativa L.). Proceedings of the National Academy of Sciences, 102, 16573.

Rondon, M., August, P., Bettermann, A., Brady, S., Grossman, T., Liles, M., Loiacono, K.,Lynch, B., Macneil, I. & Minor, C. 2000. Cloning the soil metagenome: A strategy foraccessing the genetic and functional diversity of uncultured microorganisms. Applied andEnvironmental Microbiology, 66, 2541.

Rosgaard, L., Pedersen, S., Langston, J., Akerhielm, D., Cherry, J. & Meyer, A. 2007. Eval-uation of minimal Trichoderma reesei cellulase mixtures on differently pretreated barleystraw substrates. Biotechnology Progress, 23, 1270–1276.

Rudolf, A., Baudel, H., Zacchi, G., Hahn-Hagerdal, B. & Liden, G. 2008. Simultaneous sac-charification and fermentation of steam-pretreated bagasse using Saccharomyces cerevisiaeTMB3400 and Pichia stipitis CBS6054. Biotechnology and Bioengineering, 99, 783–790.

Sainz, M. 2009. Commercial cellulosic ethanol: The role of plant-expressed enzymes. In VitroCellular & Developmental Biology-Plant, 45, 314–329.

Schaffner, D. W. & Toledo, R. T. 1991. Cellulase production by Trichoderma reesei whencultured on xylose-based media supplemented with sorbose. Biotechnology and Bioengi-neering, 37, 12–16.

Shallom, D. & Shoham, Y. 2003. Microbial hemicellulases. Current Opinion in Microbiology,6, 219–228.

Singhania, R. R., Sukumaran, R. K. & Pandey, A. 2007. Improved cellulase production by Tri-choderma reesei RUT C30 under SSF through process optimization. Applied Biochemistryand Biotechnology, 142, 60–70.

Spear, L., Gallagher, J., McHale, L. & McHale, A. P. 1993. Production of cellulase andbeta-glucosidase activities following growth of Streptomyces hygroscopicus on cellulosecontaining media. Biotechnology Letters, 15, 1265–1268.

Sticklen, M. 2006. Plant genetic engineering to improve biomass characteristics for biofuels.Current Opinion in Biotechnology, 17, 315–319.

Sticklen, M. 2008a. Altering regulation of maize lignin biosynthesis enzymes via RNAitechnology. US Patent App. 20,080/213,871, 2007.

Sticklen, M. B. 2008b. Plant genetic engineering for biofuel production: Towards affordablecellulosic ethanol. Nature Reviews Genetics, 9, 433–443.

Sun, Y. & Cheng, J. 2002. Hydrolysis of lignocellulosic materials for ethanol production: Areview. Bioresource Technology, 83, 1–11.

Taherzadeh, M. & Karimi, K. 2007. Enzyme-based hydrolysis processes for ethanol fromlignocellulosic materials: A review. BioResources, 2, 707–738.

Teather, R. & Wood, P. 1982. Use of Congo red–polysaccharide interactions in enumeration andcharacterization of cellulolytic bacteria from the bovine rumen. Applied and EnvironmentalMicrobiology, 43, 777–780.

252 Plant Biomass Conversion

Tholudur, A., Ramirez, W. F. & Mcmillan, J. D. 1999. Mathematical modeling and optimiza-tion of cellulase protein production using Trichoderma reesei RL-P37. Biotechnology andBioengineering, 66, 1–16.

Tolonen, A., Chilaka, A., Church, G. & Church, G. 2009. Targeted gene inactivation inClostridium phytofermentans shows that cellulose degradation requires the family 9 hydro-lase Cphy3367. Molecular Microbiology, 74, 1300–1313.

Torney, F., Moeller, L., Scarpa, A. & Wang, K. 2007. Genetic engineering approaches toimprove bioethanol production from maize. Current Opinion in Biotechnology, 18, 193–199.

Toyama, H. & Toyama, N. 2001. The effect of additional autopolyploidization in a slow growingcellulase hyperproducer of Trichoderma. Applied Biochemistry and Biotechnology, 91/93,787–790.

Toyama, H., Yano, M., Gisushi, A., Hotta, T. & Toyama, N. 2003. Active nuclear shufflingsystem using a swollen conidium of Trichoderma reesei. Applied Biochemistry and Biotech-nology, 108, 821–824.

Twyman, R. M., Stoger, E., Schillberg, S., Christou, P. & Fischer, R. 2003. Molecular farmingin plants: Host systems and expression technology. Trends in Biotechnology, 21, 570–578.

Voutilainen, S., Murray, P., Tuohy, M. & Koivula, A. 2010. Expression of Talaromyces emer-sonii cellobiohydrolase Cel7A in Saccharomyces cerevisiae and rational mutagenesis toimprove its thermostability and activity. Protein Engineering Design and Selection, 23,69–79.

Woodard, S., Mayor, J., Bailey, M., Barker, D., Love, R., Lane, J., Delaney, D., McComas-Wagner, J., Mallubhotla, H. & Hood, E. 2003. Maize (Zea mays)-derived bovine trypsin:Characterization of the first large-scale, commercial protein product from transgenic plants.Biotechnology and Applied Biochemistry, 38, 123–130.

Wyman, C. E., Dale, B. E., Elander, R. T., Holtzapple, M., Ladisch, M. R. & Lee, Y. Y. 2005.Comparative sugar recovery data from laboratory scale application of leading pretreatmenttechnologies to corn stover. Bioresource Technology, 96, 2026–2032.

Xia, L. M. & Shen, X. L. 2004. High-yield cellulase production by Trichoderma reesei ZU-02on corn cob residue. Bioresource Technology, 91, 259–262.

Xue, G., Patel, M., Johnson, J., Smyth, D. & Vickers, C. 2003. Selectable marker-free transgenicbarley producing a high level of cellulase (1, 4-ß-glucanase) in developing grains. PlantCell Reports, 21, 1088–1094.

Yang, B. & Wyman, C. 2004. Effect of xylan and lignin removal by batch and flowthroughpretreatment on the enzymatic digestibility of corn stover cellulose. Biotechnology andBioengineering, 86, 88–98.

Zhang, M., Eddy, C., Deanda, K., Finkelstein, M. & Picataggio, S. 1995. Metabolic engineeringof a pentose metabolism pathway in ethanologenic Zymomonas mobilis. Science, 267, 240.

Zhang, P., Himmel, M. & Mielenz, J. 2006. Outlook for cellulase improvement: Screening andselection strategies. Biotechnology Advances, 24, 452–481.

Zhang, Y.-H. P., Ding, S.-Y., Mielenz, J. R., Cui, J.-B., Elander, R. T., Laser, M., Himmel, M.E., Mcmillan, J. R. & Lynd, L. R. 2007. Fractionating recalcitrant lignocellulose at modestreaction conditions. Biotechnology and Bioengineering, 97, 214–223.

Zhang, Y. H. P. & Lynd, L. R. 2004. Toward an aggregated understanding of enzymatic hy-drolysis of cellulose: Noncomplexed cellulase systems. Biotechnology and Bioengineering,88, 797–824.

Zhou, J., Wang, Y.-H., Chu, J., Luo, L.-Z., Zhuang, Y.-P. & Zhang, S.-L. 2009. Optimizationof cellulase mixture for efficient hydrolysis of steam-exploded corn stover by statisticallydesigned experiments. Bioresource Technology, 100, 819–825.

10 Enzyme Production Systems for Biomass Conversion 253

Zhu, J., Wang, G., Pan, X. & Gleisner, R. 2009. The status of and key barriers in lignocellulosicethanol production: A technological perspective. International Conference on BiomassEnergy Technologies, Guangzhou, China.

Zhuang, J., Marchant, M. A., Nokes, S. E. & Strobel, H. J. 2007. Economic analysis ofcellulase production methods for bio-ethanol. Applied Engineering in Agriculture, 23, 679–687.