Cellulase Improvement

Biotechnology Advances 24 (2006) 452 481 www.elsevier.com/locate/biotechadv

Research review paper

Outlook for cellulase improvement: Screening and selection strategiesY.-H. Percival Zhang a,, Michael E. Himmel b , Jonathan R. Mielenz ca

Biological Systems Engineering Department, Virginia Polytechnic Institute and State University, Blacksburg, VA 24061, USA b National Bioenergy Center, National Renewable Energy Laboratory, Golden, CO 80401, USA c Life Science Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA Received 31 January 2006; received in revised form 6 March 2006; accepted 11 March 2006 Available online 27 March 2006

Abstract Cellulose is the most abundant renewable natural biological resource, and the production of biobased products and bioenergy from less costly renewable lignocellulosic materials is important for the sustainable development of human beings. A reduction in cellulase production cost, an improvement in cellulase performance, and an increase in sugar yields are all vital to reduce the processing costs of biorefineries. Improvements in specific cellulase activities for non-complexed cellulase mixtures can be implemented through cellulase engineering based on rational design or directed evolution for each cellulase component enzyme, as well as on the reconstitution of cellulase components. Here, we review quantitative cellulase activity assays using soluble and insoluble substrates, and focus on their advantages and limitations. Because there are no clear relationships between cellulase activities on soluble substrates and those on insoluble substrates, soluble substrates should not be used to screen or select improved cellulases for processing relevant solid substrates, such as plant cell walls. Cellulase improvement strategies based on directed evolution using screening on soluble substrates have been only moderately successful, and have primarily targeted improvement in thermal tolerance. Heterogeneity of insoluble cellulose, unclear dynamic interactions between insoluble substrate and cellulase components, and the complex competitive and/or synergic relationship among cellulase components limit rational design and/or strategies, depending on activity screening approaches. Herein, we hypothesize that continuous culture using insoluble cellulosic substrates could be a powerful selection tool for enriching beneficial cellulase mutants from the large library displayed on the cell surface. 2006 Elsevier Inc. All rights reserved.Keywords: Cellulase activity assay; Cellulose; Cellulosome; Continuous culture; Enzymatic cellulose hydrolysis; High throughput screening; Selection; Sugar assay

Abbreviations: AFEX, ammonia fiber explosion; BC, bacterial cellulose; BCA, 2,2-bicinchroninate; BMCC, bacterial microcrystalline cellulose; CMC, carboxymethyl cellulose; CBM, cellulose-binding module; CBP, consolidated bioprocessing; CrI, crystallinity index; DMAc, N,Ndimethylacetamide; DNS, dinitrosalicyclic acid; DP, degree of polymerization of cellulose; DS, degree of substitution; DTT, dithiothreitol; Fa, fraction of -glucosidic bond accessible to cellulase; FPA, filter paper activity; FRE, fraction of the reducing end to all anhydroglucose units of cellulose, 1/DP; HEC, hydroxyethyl cellulose; PASC, phosphoric acid swollen cellulose; RAC, regenerated amorphous cellulose; PAHBAH, 4hydroxybenzoylhydrazine; RS, selection ratio; TNP-CMC, trinitrophenyl-carboxymethyl cellulose. Corresponding author. Tel.: +1 540 231 7414; fax: +1 540 231 3199. E-mail address: [email protected] (Y.-H. Percival Zhang). 0734-9750/$ - see front matter 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.biotechadv.2006.03.003

Y.-H. Percival Zhang et al. / Biotechnology Advances 24 (2006) 452481

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Contents Introduction . . . . . . . . . . . . . . . . . . Cellulose hydrolysis mechanisms . . . . . . . Substrates for cellulase activity assays . . . . . 3.1. Soluble substrates . . . . . . . . . . . . 3.2. Insoluble substrates . . . . . . . . . . . 4. Quantitative assays . . . . . . . . . . . . . . . 4.1. Hydrolysis products . . . . . . . . . . . 4.2. Cellulase activity assays. . . . . . . . . 4.2.1. Endoglucanases . . . . . . . . 4.2.2. Exoglucanases . . . . . . . . . 4.2.3. -D-glucosidases . . . . . . . . 4.2.4. Total cellulase . . . . . . . . . 5. Cellulase improvement and screening/selection 5.1. Rational design . . . . . . . . . . . . . 5.2. Directed evolution. . . . . . . . . . . . 5.3. Screening . . . . . . . . . . . . . . . . 5.4. Selection . . . . . . . . . . . . . . . . 6. Summary . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . 1. 2. 3. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453 455 457 458 458 460 461 462 462 463 464 464 465 465 467 469 470 472 472 472

1. Introduction Cellulose is the primary product of photosynthesis in terrestrial environments, and the most abundant renewable bioresource produced in the biosphere ( 100 billion dry tons/year) (Holtzapple, 1993; Jarvis, 2003; Zhang and Lynd, 2004b). Cellulose biodegradation by cellulases and cellulosomes, produced by numerous microorganisms, represents a major carbon flow from fixed carbon sinks to atmospheric CO2 (Berner, 2003; Falkowski et al., 2000; Melillo et al., 2002), is very important in several agricultural and waste treatment processes (Angenent et al., 2004; Das and Singh, 2004; Haight, 2005; Hamer, 2003; Humphrey et al., 1977; Russell and Rychlik, 2001; Schloss et al., 2005; van Wyk, 2001), and could be widely used to produce sustainable biobased products and bioenergy to replace depleting fossil fuels (Angenent et al., 2004; Demain et al., 2005; Galbe and Zacchi, 2002; Hall et al., 1993; Hoffert et al., 2002; Kamm and Kamm, 2004; Lynd, 1996; Lynd et al., 1991, 2002, 1999; Mielenz, 2001; Mohanty et al., 2000; Moreira, 2005; Reddy and Yang, 2005; Wyman, 1994, 1999, 2003). Additionally, studies have shown that the use of biobased products and bioenergy can achieve zero net carbon dioxide emission (Demain, 2004; Demain et al., 2005; Hoffert et al., 2002; Lynd et al., 1991, 1999). Development of technologies for effectively converting less costly agricultural and forestry residues to fermentable sugars

offers outstanding potential to benefit the national interest through: (1) improved strategic security, (2) decreased trade deficits, (3) healthier rural economies, (4) improved environmental quality, (5) technology exports, and (6) a sustainable energy resource supply (Angenent et al., 2004; Caldeira et al., 2003; Demain et al., 2005; Hoffert et al., 1998, 2002; Kamm and Kamm, 2004; Lynd, 1996; Lynd et al., 1991, 1999, 2002; Moreira, 2005; Wirth et al., 2003; Wyman, 1999). Effective conversion of recalcitrant lignocellulose to fermentable sugars requires three sequential steps: (1) size reduction, (2) pretreatment/fractionation, and (3) enzymatic hydrolysis (Wyman, 1999; Zhang and Lynd, 2004b). One of the most important and difficult technological challenges is to overcome the recalcitrance of natural lignocellulosic materials, which must be enzymatically hydrolyzed to produce fermentable sugars (Chang et al., 1981; Demain et al., 2005; Fan et al., 1982; Grethlein, 1984; Hsu, 1996; Lin et al., 1981; McMillian, 1994; Millett et al., 1976; Moreira, 2005; Mosier et al., 2005; Saddler et al., 1993; Weil et al., 1994; Wyman, 1999; Wyman et al., 2005a). Cellulases are relatively costly enzymes, and a significant reduction in cost will be important for their commercial use in biorefineries. Cellulase-based strategies that will make the biorefinery processing more economical include: increasing commercial enzyme volumetric productivity, producing enzymes using

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cheaper substrates, producing enzyme preparations with greater stability for specific processes, and producing cellulases with higher specific activity on solid substrates. Recently, the biotechnology companies Genencor International and Novozymes Biotech have reported the development of technology that has reduced the cellulase cost for the cellulose-to-ethanol process from US$5.40 per gallon of ethanol to approximately 20 cents per gallon of ethanol (Moreira, 2005), in which the two main strategies were (1) an economical improvement in production of cellulase to reduce US$ per gram of enzyme by process and strain enhancement, e.g., cheaper medium from lactose to glucose and alternative inducer system and (2) an improvement in the cellulase enzyme performance to reduce grams of enzyme for achieving equivalent hydrolysis by cocktails and component improvement (Knauf and Moniruzzaman, 2004). But this claim has not yet been widely accepted because the cellulase mixture was tested only for the specific pretreated lignocellulosic substrate and cannot be applied to other pretreated lignocelluloses. Currently, most commercial cellulases (including glucosidase) are produced by Trichoderma species and Aspergillus species (Cherry and Fidantsef, 2003; Esterbauer et al., 1991; Kirk et al., 2002). Cellulases are used in the textile industry for cotton softening and denim finishing; in the detergent market for color care, cleaning, and anti-deposition; in the food industry for mashing; and in the pulp and paper industries for de-

inking, drainage improvement, and fiber modification (Cherry and Fidantsef, 2003; Kirk et al., 2002). The cellulase market is expected to expand dramatically when cellulases are used to hydrolyze pretreated cellulosic materials to sugars, which can be fermented to commodities such as bioethanol and biobased products on a large scale (Cherry and Fidantsef, 2003; Himmel et al., 1999; van Beilen and Li, 2002). For example, the potential cellulase market has been estimated to be as high as US$400 million per year if cellulases are used for hydrolyzing the available corn stover in the midwestern United States (van Beilen and Li, 2002). This market scenario represents an increase of 33% in the total US industrial enzyme market (Wolfson, 2005). The large market potential and the important role that cellulases play in the emerging bioenergy and bio-based products industries provide a great motivation to develop better cellulase preparations for plant cell wall cellulose hydrolysis. These improved cellulases must also have characteristics necessary for biorefineries, such as higher catalytic efficiency on insoluble cellulosic substrates, increased stability at elevated temperature and at a certain pH, and higher tolerance to end-product inhibition. Fig. 1 shows that cellulase engineering for noncomplexed cellulase systems contains three major research directions: (1) rational design for each cellulase, based on knowledge of the cellulase structure and the catalytic mechanism (Schulein, 2000; Wilson, 2004; Wither, 2001); (2) directed evolution for each

Wild type Cellulase Components Reconstitute cellulase cocktail endos exosR expsNR -Gase Improved cellulase components

Rational Design

Directed Evolution

Screen or select on solid substrate

Fig. 1. Scheme of cellulase engineering for non-complexed cellulases. Endos, endoglucanases; exosR, exoglucanases acting on reducing ends; exosNR, exoglucanases acting on non-reducing ends; -Gase, -glucosidase.


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cellulase, in which the improved enzymes or ones with new properties were selected or screened after random mutagenesis and/or molecular recombination (Arnold, 2001; Cherry and Fidantsef, 2003; Hibbert et al., 2005; Schmidt-Dannert and Arnold, 1999; Shoemaker et al., 2003; Tao and Cornish, 2002); and (3) the reconstitution of cellulase mixtures (cocktails) active on insoluble cellulosic substrates, yielding an improved hydrolysis rate or higher cellulose digestibility (Baker et al., 1998; Boisset et al., 2001; Himmel et al., 1999; Irwin et al., 1993; Kim et al., 1998; Sheehan and Himmel, 1999; Walker et al., 1993; Wilson and Walker, 1991; Zhang and Lynd, 2004b). With respect to engineering complexed cellulase systems (cellulosomes), the idea of chimeric constructs of cellulosomal domains/components was proposed by Bayer et al. (1994), and the reconstruction of cellulosome components is becoming another hot research area (Fierobe et al., 2001, 2002, 2005; Mingardon et al., 2005; Sabathe and Soucaille, 2003), which we do not review here. The cornerstone of enzyme engineering is to achieve a direct correlation between the enzyme assays or screening approaches and the changes in enzyme functions in the desired application. Development of a useful, predictive cellulase assay or screening is particularly difficult because of the nature of solid heterogeneous substrates, such as plant cell walls. Available quantitative cellulase assays and screenings have been analyzed and compared herein, including their advantages and limitations. Also, successful cellulase examples using directed evolution are examined, and a possible strategy of combinatorial molecular breeding and continuous culture with solid cellulosic materials to select a cellulase with higher activity is discussed. 2. Cellulose hydrolysis mechanisms Cellulose is a linear condensation polymer consisting of D-anhydroglucopyranose joined together by -1,4glycosidic bonds with a degree of polymerization (DP) from 100 to 20,000 (Krassig, 1993; O'Sullivan, 1997; Tomme et al., 1995; Zhang and Lynd, 2004b). Anhydrocellobiose is the repeating unit of cellulose. Coupling of adjacent cellulose chains and sheets of cellulose by hydrogen bonds and van der Waal's forces results in a parallel alignment and a crystalline structure with straight, stable supra-molecular fibers of great tensile strength and low accessibility (Demain et al., 2005; Krassig, 1993; Nishiyama et al., 2003; Notley et al., 2004; Zhang and Lynd, 2004b; Zhbankov, 1992). The cellulose molecule is very stable, with a half life of

58 million years for -glucosidic bond cleavage at 25 C (Wolfenden and Snider, 2001), while the much faster enzyme-driven cellulose biodegradation process is vital to return the carbon in sediments to the atmosphere (Berner, 2003; Cox et al., 2000; Falkowski et al., 2000; Schlamadinger and Marland, 1996). The widely accepted mechanism for enzymatic cellulose hydrolysis involves synergistic actions by endoglucanase (EC 3.2.1.4), exoglucanase or cellobiohydrolase (EC 3.2.1.91), and -glucosidase (EC 3.2.1.21) (Henrissat, 1994; Knowles et al., 1987; Lynd et al., 2002; Teeri, 1997; Wood and GaricaCampayo, 1990; Zhang and Lynd, 2004b). Endoglucanases hydrolyze accessible intramolecular -1,4glucosidic bonds of cellulose chains randomly to produce new chain ends; exoglucanases processively cleave cellulose chains at the ends to release soluble cellobiose or glucose; and -glucosidases hydrolyze cellobiose to glucose in order to eliminate cellobiose inhibition. These three hydrolysis processes occur simultaneously as shown in Fig. 2. Primary hydrolysis that occurs on the surface of solid substrates releases soluble sugars with a degree of polymerization (DP) up to 6 into the liquid phase upon hydrolysis by endoglucanases and exoglucanases. The enzymatic depolymerization step performed by endoglucanases and exoglucanases is the rate-limiting step for the whole cellulose hydrolysis process. Secondary hydrolysis that occurs in the liquid phase involves primarily the hydrolysis of cellobiose to glucose by -glucosidases, although some -glucosidases also hydrolyze longer cellodextrins (Zhang and Lynd, 2004b). During cellulose hydrolysis, the solid substrate characteristics vary, including (1) changes in the cellulose chain end number resulting from generation by endoglucanases and consumption by exoglucanases (Kleman-Leyer et al., 1992, 1994, 1996; Kongruang et al., 2004; Srisodsuk et al., 1998; Zhang and Lynd, 2005b) and (2) changes in cellulose accessibility resulting from substrate consumption and cellulose fragmentation (Banka et al., 1998; Boisset et al., 2000; Chanzy et al., 1983; Din et al., 1991, 1994; Halliwell and Riaz, 1970; Lee et al., 1996, 2000; Saloheimo et al., 2002; Walker et al., 1990, 1992; Wang et al., 2003; Woodward et al., 1992). The combined actions of endoglucanases and exoglucanases modify the cellulose surface characteristics (topography) over time, resulting in rapid changes in hydrolysis rates. The complicated interactions among endoglucanases, exoglucanases, and the changing substrate characteristics during hydrolysis have been simulated by a new functionally based mathematical model

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Solid Phase

(primary hydrolysis)

Liquid Phase (secondary hydrolysis)

endos exosR exosNR -Gase

Fig. 2. Mechanistic scheme of enzymatic cellulose hydrolysis by Trichoderma non-complexed cellulase system.

(Zhang and Lynd, in press), applying a set of enzymatic parameters for endoglucanase I, cellobiohydrolases I and II to a variety of substrates with two important substrate properties: the fraction of -glucosidic bond accessible to cellulase (Fa) (Zhang and Lynd, 2004b) and the degree of polymerization (DP) (Okazaki and Moo-Young, 1978; Zhang and Lynd, 2004b) (see Table 1). In this way, disparate information from the

literature was framed in a coherent way to facilitate an understanding of enzymatic cellulose hydrolysis. For example, the reaction rates simulated by the model were consistent with a substantial number of observations reported in the literature, including the effects of substrate characteristics on exoglucanase and endoglucanase activities; the effects of substrate characteristics and experimental conditions on the degree of

Table 1 Summary of typical values of model celluloses for crystallinity index (CrI), the fraction of -glucosidic bond accessible to cellulase (Fa), which is estimated by maximum cellulase adsorption capacity (Zhang and Lynd, 2004b), the number average of degree of polymerization (DPN), the fraction of reducing ends (FRE), and relative ratio of FRE/Fa Substrates CrI Fa (%) DPN FRE (%) Low 26 1002000 16.67 0.05 High 50 1 FRE/Fa Low 0.167 0.0005 High 0.5 0.01

Soluble Cellodextrins and their derivatives CMC Insoluble Cotton Whatman No. 1 filter paper Bacterial cellulose Microcrytalline cellulose (Avicel) PASC Pulp (Solka Floc) Pretreated cellulosic substrates

N.A. N.A.

100 100

0.80.95 0.45 0.80.95 0.50.6 0 0.40.7 0.40.7

0.2 1.8 6 0.6 12 1.8 0.6

10003000 7502800 6002000 150500 1001000 7501500 4001000

0.033 0.036 0.05 0.2 0.1 0.067 0.1

0.1 0.133 0.167 0.667 1 0.133 0.25

0.167 0.0198 0.00833 0.333 0.00833 0.0370 0.167

0.5 0.0741 0.0278 1.11 0.0833 0.0741 0.417


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endo-exo synergy; the effects of endoglucanase partition coefficient on the hydrolysis rates; and the effects of enzyme loading on relative reaction rates for different substrates. The model also suggests that it is nearly impossible to predict hydrolysis performance of cellulase mixtures from one solid substrate to another solid substrate, because of large variations in total cellulase concentration, ratio of endo/exocellulases, reaction time, and substrate characteristics. Therefore, enzyme reconstitution may have be conducted so as to achieve better performance for a specific substrate (Knauf and Moniruzzaman, 2004). Unlike non-complexed fungal cellulase, anaerobic microorganisms possess complexed cellulase systems, called cellulosomes (Bayer et al., 1994, 1998, 2004; Beguin and Alzari, 1998; Demain et al., 2005; Doi and Kosugi, 2004; Doi et al., 1998; Doi and Tamaru, 2001; Leschine, 1995; Schwarz, 2001). Leschine (1995) estimated that anaerobic cellulose degradation could account for only 510% of total cellulose biodegradation, but it could be underestimated because anaerobic cellulose hydrolysis is responsible for considerable carbon recycling in the anoxic zones of ponds, lakes, oceans, and intestines of ruminants and guts of termites (P.J. Weimer, personal communication). Furthermore, an understanding anaerobic cellulase systems are of significant importance to basic

sciences, such as the evolution of cellulase genes, the structures of cellulases, and the formation and hydrolysis of reacting biofilms on cellulose surfaces (Lynd et al., in press). Anaerobic cellulose fermentation has both current and future applications, such as agricultural processes anaerobic waste treatment, and consolidated bioprocessing (CBP), respectively (Lynd, 1996; Lynd et al., 1999, 2002, 2005). Recently, a microbial cellulose hydrolysis mechanism has been reported for the anaerobic cellulolytic bacterium Clostridium thermocellum that assimilates longer soluble hydrolysis products with an average degree of polymerization of 4 rather than glucose and cellobiose. The improved bioenergetics resulting from longer chain sugar assimilation supports the biological feasibility of anaerobic fermentation without added saccharolytic enzymes (Zhang and Lynd, 2005c). More information about the cellulosomebased microbial cellulose hydrolysis research is available elsewhere (Lynd, 1996; Lynd et al., 2002, 1999, 2005; Zhang and Lynd, 2003a, 2004a, 2005a). 3. Substrates for cellulase activity assays Substrates for cellulase activity assays can be divided into two categories, based on their solubility in water (Table 2).

Table 2 Substrates containing -1,4-glucosidic bonds hydrolyzed by cellulases and their detections Substrate Soluble Short chain (low DP) Cellodextrins Radio-labeled cellodextrins Cellodextrin derivatives -methylumbelliferyl-oligosaccharides p-nitrophenol-oligosaccharides Long chain cellulose derivatives Carboxymethyl cellulose (CMC), hydroxyethyl cellulose (HEC) Dyed CMC Insoluble Crystalline celluloseCotton, microcrystalline cellulose (Avicel), Valonia cellulose, bacterial cellulose Amorphous cellulose - PASC, alkali-swollen cellulose RAC Dyed cellulose Fluorescent cellulose Chromogenic and fluorephoric derivatives Trinitrophenyl-carboxymethylcellulose (TNP-CMC) Fluram-cellulose Practical cellulose-containing substrates -cellulose, pretreated lignocellulosic biomassa

Detection a

Enzymes

RS, HPLC; TLC TLC plus liquid scintillation Fluorophore liberation, TLC Chromophore liberation, TLC RS; viscosity Dye liberation

Endo, Exo, BG Endo, Exo, BG Endo, Exo, BG Endo, Exo, BG Endo Endo

RS, TSS, HPLC RS, TSS, HPLC RS, TSS, HPLC, TLC Dye liberation Fluorophore liberation Chromophore liberation Fluorophore liberation HPLC, RS

Total, Endo, Exo Total, Endo, Exo Total, Endo Total Endo Endo, Total Total

RS, reducing sugars; TSS, total soluble sugars.

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3.1. Soluble substrates Soluble substrates include low DP cellodextrins from 2 to 6 sugar units and their derivatives, as well as long DP cellulose derivatives (ca. several hundreds of sugar units). They are often used for measuring individual cellulase component activity (Table 2). Cellodextrins are soluble for DP 6, and very slightly soluble for 6 < DP < 12 (Miller, 1960; Miller, 1963; Pereira et al., 1988; Zhang and Lynd, 2003b, 2005b). Their solubility decreases drastically with increasing DP because of strong intermolecular hydrogen bonds and system entropic effects. Cellodextrins are often prepared through cellulose hydrolysis by fuming HCl (Miller, 1960, 1963), sulfuric acid (Voloch et al., 1984), acetylation (Schmid et al., 1988; Wolfram and Dacons, 1952), or mixed acids (HCl and H2SO4) (Zhang and Lynd, 2003b). Cellodextrins are also prepared through biosynthesis using C. thermocellum cellobiose and cellodextrin phosphorylases (Ng and Zeikus, 1986; Strobel et al., 1995; Zhang and Lynd, 2005c, 2006), or T. reesei -glucosidase (Chirico and Brown, 1987). Cellodextrin mixtures can be separated into single components using chromatographic methods such as charcoal-celite (Miller, 1963), thin layer (Chirico and Brown, 1985; Zhang and Lynd, 2006), cation exchange (Huebner et al., 1978; Voloch et al., 1984), or sizeexclusion (Schmid et al., 1988; Shintate et al., 2003; Zhang and Lynd, 2003b, 2006). Chromogenic p-nitrophenyl glycosides and fluorogenic methylumbelliferyl-D-glycosides derived from soluble cellodextrins are very useful for the study of initial cellulase kinetics (Tuohy et al., 2002; Wolfgang and Wilson, 1999), reaction specificity (Bhat et al., 1990; Claeyssens and Aerts, 1992; Tomme et al., 1996; van Tilbeurgh and Claeyssens, 1985; van Tilbeurgh et al., 1982, 1985; Zverlov et al., 2002b), and binding site thermodynamics (Barr and Holewinski, 2002). They are also used to determine the inhibition constants of cellulase in the presence of added cellobiose and glucose (Tuohy et al., 2002), because chromophores released from substituted glycosides can be easily measured independently of sugars. Long DP cellulose derivatives can be dissolved in water because of their chemical substitutions. Ionicsubstituted carboxymethyl cellulose (CMC) is often used for determining endoglucanase activity, called CMCase, because endoglucanases cleave intramolecular -1,4-glucosidic bonds randomly, resulting in a dramatic reduction in the DP (i.e., specific viscosity) of CMC. CMC has two very important physical parameters the degree of substitution (DS) and DP. The solubility

of CMC is closely associated with the DS that has a maximum stoichiometric value of 3. CMC is soluble in water when DS > 0.30.7 (Karlsson et al., 2001; Klemm et al., 1998a; Wood and Bhat, 1988). Commercial CMCs usually have a DS < 1.5. It is strongly recommended that a reducing sugar assay or viscosity assay using CMC as a substrate should be limited to the first 2% hydrolysis of substrate when DS = 0.7 (Wood and Bhat, 1988). This is important because only nonsubstituted glucose units are accessible to cellulase, and hydrolysis action requires at least two or three contiguous non-substituted residues. The DP of CMC is not important for the reducing sugar assay, but it is very important for determining viscosity reduction. CMC dissolution in water should be done by gentle swirling to avoid DP reduction (Sharrock, 1988). Also, the viscosity of ionic CMC is influenced by pH, ionic strength, and polyvalent cation concentration. Therefore, it is recommended to use nonionic substituted celluloses, such as hydroxyethyl cellulose (HEC), for determining endoglucanase activity (Wood and Bhat, 1988). Dyed soluble CMC is made by mixing CMC with dyesRemazol Brilliant Blue R (Flp and Ponyi, 1997; McCleary, 1980; Wirth and Wolf, 1992) or Ruthenium Red (Rescigno et al., 1994). The colors released from soluble cellulose derivatives in the supernatant can be measured after precipitation of the soluble substrates. A dyed CMC is also used as a substrate on solid agar plates, sometimes called zymograms (Bera-Maillet et al., 2000; Eriksson and Petterson, 1973; Holt and Hartman, 1994). After hydrolysis, staining, and washing, halo zones can be observed in the colored background. 3.2. Insoluble substrates Insoluble cellulose-containing substrates for cellulase activity assays include nearly pure celluloses (cotton linter, Whatman No. 1 filter paper, bacterial cellulose, microcrystalline cellulose, and amorphous cellulose) and impure cellulose-containing substrates (dyed cellulose, -cellulose, and pretreated lignocellulose). Native cellulose, referred to as cellulose I, has two distinct crystallite forms, I which is dominant in bacterial and algal cellulose, and I, which is dominant in higher plants (Atalla and Vanderhart, 1984). Native cellulose (cellulose I) can be converted to other crystalline forms (IIIV) by various treatments (Klein and Snodgrass, 1993; Krassig, 1993; O'Sullivan, 1997). Table 1 shows several key physical valuescrystallinity index, degree of polymerization, and cellulose


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accessibility to cellulase that can be estimated based on maximum cellulase adsorption (Zhang and Lynd, 2004b). The crystallinity index (CrI) of cellulose, quantitatively measured from its wide range X-ray diffraction pattern (Krassig, 1993; Ramos et al., 2005; Zhang and Lynd, 2004b), is not strongly associated with hydrolysis rates (Converse, 1993; Mansfield et al., 1999; Zhang and Lynd, 2004b). Nevertheless, it is still a convenient indicator representing the change in cellulose characteristics for one material before and after treatment. Cotton, bacterial cellulose, and the Valonia ventricosa algal cellulose are examples of highly crystalline cellulose (Boisset et al., 1999; Fierobe et al., 2002), whereas amorphous cellulose is at the other extreme. Microcrystalline cellulose, filter paper, -cellulose, and pretreated cellulosic substrates have modest CrI values, and can be regarded as a combination of crystalline fraction and amorphous fraction, but there is no clear borderline between two fractions. Cotton fiber is made from natural cotton after impurities, such as wax, pectin, and colored matter, have been removed (Wood, 1988). Whatman No.1 filter paper is made from long fiber cotton pulp with a low CrI = 45% (Dong et al., 1998; Henrissat et al., 1985). Microcrystalline cellulose, called hydrocellulose or avicel (the commercial name), can be purchased from several companies, such as FMC, Merck, and Sigma. It is made through the following steps: hydrolysis of wood pulp by dilute hydrochloric acid to remove the amorphous cellulose fraction, formation of colloidal dispersions by high shear fields, followed by spray drying of the washed pulp slurry (Fleming et al., 2001; Zhang and Lynd, 2004b). However, microcrystalline cellulose still contains a significant fraction of amorphous cellulose. Avicel is a good substrate for exoglucanase activity assay, because it has a low DP and relatively low accessibility (i.e., the highest ratio of FRE/Fa) (Table 1). Therefore, some researchers feel that avicelase activity is equivalent to exoglucanase activity (Wood and Bhat, 1988). However, some endoglucanases can release considerable reducing sugars from avicel (Zhang and Lynd, 2004b). Bacterial cellulose (BC) is prepared from the pellicle produced by Acetobacter xylinum (ATCC 23769) (Hestrin, 1963) or from Nata de Coco (Daiwa Fine Produces, Singapore) (Boisset et al., 2000). Bacterial microcrystalline cellulose (BMCC) can be prepared from BC by partial acid hydrolysis to remove the amorphous cellulose fraction, resulting in a reduction in DP (Valjamae et al., 1999).

Amorphous cellulose is prepared by converting the crystalline fraction of cellulose to the amorphous form by mechanical or chemical methods. These celluloses include mechanically made amorphous cellulose, alkaliswollen cellulose, and phosphorous acid swollen cellulose (PASC, Walseth cellulose). Mechanically made amorphous cellulose is often prepared by ball milling or severe blending (Fan et al., 1980; Ghose, 1969; Henrissat et al., 1985; Wood, 1988). Alkaliswollen amorphous cellulose is made by swelling cellulose power in a high concentration of NaOH (e.g., 16% wt/wt) producing the cellulose type II from type I (O'Sullivan, 1997; Wood, 1988). Phosphoric acid swollen cellulose (PASC) is most commonly made by swelling dry cellulose powder by adding 85% ophosphoric acid (Walseth, 1952; Wood, 1988). High concentration phosphoric acid treatment could result in some degree of conversion of type II cellulose from type I (Weimer et al., 1990). The properties of amorphous cellulose made by ball milling, NaOH and H3PO4, vary greatly, depending on cellulose origins, reaction temperature and time, as well as reagent types and concentrations. Therefore, it is nearly impossible to compare hydrolysis rates on various types of amorphous cellulose from different laboratories or even different batches of amorphous cellulose preparations from the same laboratory. Amorphous cellulose should be kept in hydrated condition; simple air-drying dehydration results in a loss of substrate reactivity (Zhang and Lynd, 2004b). The loss of substrate reactivity during dehydration can be minimized through freeze drying or drying after solvent exchange (Fan et al., 1981; Lee et al., 1980). Regenerated cellulose is often made by converting insoluble cellulose to soluble form using cellulose solvents, such as nitric acid, sulfuric acid, ammoniacal cupric hydroxide (Cu(NH3)4(OH)2), N,N-dimethylacetamide (DMAc)/LiCl (Striegel, 1997), and 1-butyl-3methylimidazolium Cl (Swatloski et al., 2002), followed by restoration to physically insoluble form. The major commercial regenerated cellulose is viscose rayon, which is not pure amorphous cellulose due to some recrystallization. Regenerated amorphous cellulose (RAC) can be made by using cold 85% H3PO4 to dissolve cellulose slurry, followed by precipitation with cold water. RAC is a very good homogeneous substrate for cellulase activity assays (Zhang et al., 2006), and is different from Walseth cellulose, prepared from heterogeneous swollen cellulose (Walseth, 1952). RAC has a consistent quality from batch to batch, and is an ideal insoluble nonsubstitutation cellulose substrate for measuring extremely low cellulase activity.

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-Cellulose contains major cellulose and a small amount of hemicellulose. The commercial Sigma cellulose is often used as a reference cellulosic material to evaluate the hydrolysis ability of total cellulase (Kim et al., 2003). Holocellulose is a solid residue of wood (lignocellulose) after removal of lignin; -cellulose is a solid residue of holocellulose after removal of major hemicellulose by alkali extraction (Green, 1963); after the neutralization of soluble alkali extract materials from holocellulose, the insoluble fraction and the soluble fraction are -cellulose and -cellulose, respectively (Corbett, 1963a; Corbett, 1963b). Lignocellulose pretreatment breaks up the recalcitrant structure of lignocellulose so that cellulase can hydrolyze pretreated lignocellulose faster and more efficiently. Current leading lignocellulose pretreatment technologies, including dilute acid, hot water, flow through, ammonia fiber explosion (AFEX), ammonia recycle percolation, and lime, have been recently reviewed elsewhere (Mosier et al., 2005; Wyman et al., 2005a,b). In addition, two other pretreatments steam explosion and organosolvhave been intensively investigated (Arato et al., 2005; Bura et al., 2002, 2003; Galbe and Zacchi, 2002; Ohgren et al., 2005; Pan et al., 2005a,b; Pye and Lora, 1991; Sassner et al., 2005 ; Soderstrom et al., 2003; Wingren et al., 2003). The substrate characteristics (e.g., cellulose accessibility, DP, hemicellulose content, and lignin content) of pretreated lignocelluloses vary greatly, strongly depending on pretreatment methods and severity, and on lignocellulose origins. For example, the goal of AFEX is to break up the linkages among lignin, hemicellulose, and cellulose, but not to remove any main component. Therefore, the addition of hemicellulase into the cellulase mixture would be important for improving overall hydrolysis performance for AFEX-treated feedstock (Teymouri et al., 2005). Dilute acid pretreatment not only to breaks the linkage among lignin, hemicellulose, and cellulose, but also removes major hemicellulose. Therefore, the addition of hemicellulase is not necessary for an improvement in cellulase mixture performance; while the addition of non-hydrolysis proteins (e.g. bovine serum albumin) into the cellulase mixture could reduce the use of cellulase because of minimization of non-hydrolysis adsorption of cellulase to lignin (Pan et al., 2005b; Wyman CE, personal communication). Organosolv pretreatment significantly removes both hemicellulose and lignin (Arato et al., 2005; Pan et al., 2005a; Pye and Lora, 1991). Therefore, neither hemicellulase nor other protein blockers need to be added. A novel cellulose-solvent-based lignocellulose fractionation is

under development by our laboratory; the hydrolysis rates of residual cellulose samples containing little hemicellulose and lignin cannot be improved by the addition of either hemicellulase or non-hydrolysis protein (Zhang et al., unpublished). In a word, improvements in the overall performance of cellulase mixture by cocktailing are strongly dependent on residual lignocellulose properties, and remains in the trial-and-test stage. Dyed cellulose is prepared by mixing cellulose with a variety of dyes, such as Remazol Brilliant Blue (Holtzapple et al., 1984; Wood, 1988), Reactive Orange (Gusakov et al., 1985), Reactive Blue 19 (Yamada et al., 2005), and fluorescent dye 5-(4,6dichlorotriazinyl) aminofluresceinsm (Helbert et al., 2003). Because of large variations in the surface areas of cellulose and the binding conditions, the quantitative relationship between released dye and reducing sugars must be established for each batch of dyed cellulose. Insoluble cellulose derivatives, such as slightly substituted CMC, can be mixed with a variety of dyes, including Cibacron Blue 3GA and Reactive Orange 14 to produce insoluble dyed-CMC (Ten et al., 2004). Insoluble cellulose derivatives can also be chemically substituted with trinitrophenyl groups to produce chromogenic trinitrophenyl-carboxymethyl cellulose (TNP-CMC) and fluorophoric Fluram cellulose (Huang and Tang, 1976). The TNP-CMC has a 25fold greater sensitivity for endoglucanase activity than does the reducing sugar dinitrosalicyclic acid method, and Fluram cellulose gives another 10-fold increase in sensitivity over TNP-CMC (Huang and Tang, 1976). However, an increased substitution of TNP-CMC reduces substrate solubility and impairs cellulase action along -linked chains (Wood and Bhat, 1988). Sometimes, TNP-CMC is a useful substrate for enzyme solutions containing reducing agents when the reducing sugar assay cannot be conducted (Shinmyo et al., 1979). For example, the cellulosome from the anaerobic bacterium C. thermocellum requires the presence of reducing agents such as DTT or cysteine for activity (Johnson et al., 1982a; Morag et al., 1992; Zhang and Lynd, 2003a). 4. Quantitative assays All existing cellulase activity assays can be divided into three types: (1) the accumulation of products after hydrolysis, (2) the reduction in substrate quantity, and (3) the change in the physical properties of substrates.


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4.1. Hydrolysis products The majority of assays involve the accumulation of hydrolysis products, including reducing sugars, total sugars, and chromophores. The most common reducing sugar assays include the dinitrosalicyclic acid (DNS) method (Ghose, 1987; Miller, 1959), the NelsonSomogyi method (Nelson, 1944; Somogyi, 1952), the 2,2-bicinchroninate (BCA) method (Waffenschmidt and Janeicke, 1987; Zhang and Lynd, 2005b), the 4hydroxybenzoylhydrazine (PAHBAH) method (Lever, 1972; Lever et al., 1973), and the ferricyanide methods (Kidby and Davidson, 1973; Park and Johnson, 1949) in Table 3. Total soluble sugars, regardless of their chain lengths, can be measured directly by the phenol-H2SO4 method (Dubois et al., 1956; Zhang and Lynd, 2005b) or the anthrone-H2SO4 method (Roe, 1955; Viles and Silverman, 1949). Glucose can be measured by an enzymatic glucose kit using coupled hexokinase and glucose-6-phosphate dehydrogenase (Zhang and Lynd, 2004a), or HPLC after post-hydrolysis conversion to glucose. Detection ranges of many sugar assays can be modified using two strategies: (1) a further dilution after the color reaction and (2) varying sugar volume per sample prior to the reaction. For example, the DNS method was originally designed for 20600 g reducing sugar per sample (Miller, 1959), but its detection range

can be expanded to samples of 1002500 g, followed by water dilution (Ghose, 1987). The same is true for the Nelson-Somogyi method. The Sigma enzymatic glucose assay kit was designed to measure sugar concentrations from 200 to 5000 g/L using a reaction mixture consisting of a 10-L sample plus a 1000-L enzyme solution. However, its detection limits can be lowered to 4100 g/L using a reaction mixture of 500-L sample plus 500-L 2-fold concentrated enzyme solution (Zhang and Lynd, 2004a). Major reducing sugar assays depend on the reduction of inorganic oxidants such as cupric ions (Cu2+) or ferricyanide, which accepts electrons from the donating aldehyde groups of reducing cellulose chain ends. Their detection ranges vary from less than 1 g per sample to> 2500 g per sample (Table 3). The DNS and NelsonSomogyi methods are two of the most common assays for measuring reducing sugars for cellulase activity assays because of their relatively high sugar detection range (i.e., no sample dilution required) and low interference from cellulase (i.e., no protein removal required). However, the primary drawback for this method is the poor stoichiometric relationship between cellodextrins and the glucose standard (Coward-Kelly et al., 2003; Ghose, 1987; Kongruang et al., 2004; Wood and Bhat, 1988; Zhang and Lynd, 2005b). For example, the results may suffer from an underestimation of cellulase activity when glucose is used as the standard

Table 3 The common colorimetric sugar assays Method Reducing Sugar Assay DNS DNS Nelson-Somogyi Nelson-Somogyi Nelson Ferricyanide-1 Ferricyanide-2 PAHBAH PAHBAH BCA Modified BCA Total Sugar Assay Phenol-H2SO4 Anthrone-H2SO4 Enzymatic Glucose Assay Glucose-HK/PGHD kit Glucose-HK/PGHD kit Sample (mL) Micro Macro Micro Macro Semi-Micro 13 0.5 15 2 2 13 1 0.5 0.01 0.5 1 Reagent (mL) 3 3 2+2 2+2 2 1+5 0.25 1.5 3 0.5 1 G amount (g/sample) 20600 1002500 110 10600 5100 19 0.181.8 0.55 550 0.24.5 0.49 G concn. (mg/L) 6.7600 2005000 0.210 5300 2.550 0.39 0.181.8 110 5005000 0.49 0.49 Ref.

Micro Macro

Miller, 1959 Ghose, 1987 Somogyi, 1952 Somogyi, 1952 Nelson, 1944 Park and Johnson, 1949 Kidby and Davidson, 1973 Lever, 1972 Lever, 1972 Waffenschmidt and Janeicke, 1987 Zhang and Lynd, 2005b

1 1

1+5 1+5

5100 5100

10100 10100

Dubois et al., 1956; Zhang and Lynd, 2005b Roe, 1955; Viles and Silverman, 1949

0.01 0.5

1 0.5

250 250

2005000 4100

Sigma Kit Zhang and Lynd, 2004a

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and -glucosidase is not in excess (Breuil and Saddler, 1985a,b; Schwarz et al., 1988). The ferricyanide, PAHBAH, and BCA methods, having higher sensitivity to reducing sugar, can detect as little as several micrograms per sample, but suffer from non-specific interference from protein. Total carbohydrate assays, including the phenolH2SO4 method and the anthrone-H2SO4 method, offer two obvious advantages as compared with reducing sugar assays: a strict stoichemetic relationship between cellodextrins (glucose equivalent) and the glucose standard, and little or no interference from protein. But they are limited for application to pure celluloses, because any carbohydrates and their derivatives can have strong interference readings. Using an enzymatic glucose assay kit or HPLC can overcome nonspecific readings from other sugars, but this requires an extra stepconversion of longer cellodextrins to glucose. Total loss of substrate can be measured by several means, such as gravimetry and chemical methods. These methods are not as popular as those involving product accumulation because they involve tedious procedures, such as sample centrifugation or filtration followed by drying. Gravimetry should be employed with care, because the standard deviation of this method is strongly associated with sample weight. For example, two samples of 1mg and 100mg weighed by an analytical balance with accuracy of 0.1mg have 10% and 0.1% standard deviation, respectively. Chemical methods for determining substrate loss include the phenol-H2SO4 (Dubois et al., 1956), the anthrone-H2SO4 (Viles and Silverman, 1949), and the K2Cr2O7H2SO4 methods (Wood, 1988) for residual cellulose, and quantitative saccharification for different carbohydrate components (Ruiz and Ehrman, 1996). Measurable physical cellulose properties representing cellulase activity include swollen factor, fiber strength, structure collapse, turbidity, and viscosity. Earlier assays, involving measurement of the physical changes of the residual solid cellulose, are reviewed here for historical interest. Examples of these assays include the swelling factor (measured by alkali uptake) and the reduction in tensile strength of thread and pulp (Oksanen et al., 2000; Wood, 1975). Typically, the lack of sensitivity limits the use of these assays, except on special occasions (Oksanen et al., 2000; Pere et al., 2001; Wong et al., 2000). For example, Toyama et al. measured total cellulase activity based on the time needed to disintegrate a 1 1 cm filter paper square (Wood, 1988). The turbidometric assay measures a reduction in the absorbance of particle suspension during the hydrolysis process (Enari and Niku-Paavola,

1988; Johnson et al., 1982a,b; Nummi et al., 1981), which monitors the overall hydrolysis rate over a long time but does not measure well the initial hydrolysis rate for individual enzymes. Amorphous cellulose is recommended for turbidometric assays (Enari and NikuPaavola, 1988) because crystalline cellulose hydrolysis could lead to an initial absorbance increase (Zhang, unpublished). Viscosimetric determinations have been used as an assay for the initial hydrolysis rate for endoglucanases using soluble cellulose derivatives (Demeester et al., 1976; Hulme, 1988; Manning, 1981; Miller et al., 1960). Application of this method relies on the assumption that the ratio of viscosity-average molecular weight to number-average molecular weight should remain constant during the period of the assay, which may be true only for a short time (Hulme, 1988). This method is also experimentally cumbersome and difficult to automate. 4.2. Cellulase activity assays The two basic approaches to measuring cellulase activity are (1) measuring the individual cellulase (endoglucanases, exoglucanases, and -glucosidases) activities, and (2) measuring the total cellulase activity. In general, hydrolase enzyme activities are expressed in the form of the initial hydrolysis rate for the individual enzyme component within a short time, or the end-point hydrolysis for the total enzyme mixture to achieve a fixed hydrolysis degree within a given time. For cellulase activity assays, there is always a gap between initial cellulase activity assays and final hydrolysis measurement (Sheehan and Himmel, 1999). To be most meaningful, individual cellulase component assays must also be based on a reliable estimation of the amount of individual enzyme component present in the assay. This information permits the calculation of specific activity, i.e., bonds broken per milligram enzyme per unit time. 4.2.1. Endoglucanases Endoglucanases cleave intramolecular -1,4-glucosidic linkages randomly, and their activities are often measured on a soluble high DP cellulose derivative, such as CMC with the lowest ratio of FRN/Fa (Table 1). The modes of actions of endoglucanases and exoglucanases differ in that endoglucanases decrease the specific viscosity of CMC significantly with little hydrolysis due to intramolecular cleavages, whereas exoglucanases hydrolyze long chains from the ends in a processive process (Irwin et al., 1993; Teeri, 1997; Zhang and Lynd, 2004b). Endoglucanase activities can


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be measured based on a reduction in substrate viscosity and/or an increase in reducing ends determined by a reducing sugar assay. Because exoglucanases also increase the number of reducing ends, it is strongly recommended that endoglucanase activities be measured by both methods (viscosity and reducing ends). Because the carboxymethyl substitutions on CMC make some glucosidic bonds less susceptible to enzyme action, a linear relationship between initial hydrolysis rates and serially diluted enzyme solutions requires (1) dilute enzyme preparation, (2) a short incubation period (e.g., 24 min) or a very low enzyme loading, (3) a low DS CMC, and (4) a sensitive reducing sugar assay. Many workers agree that the BCA method for reducing sugar assay is superior to the DNS method (Carcia et al., 1993). For example, the modified BCA method, which is conducted at 75 C to avoid -glucosidic bond cleavage during the assay, delivers a strict stoichiometry for the reducing ends of cellodextrins regardless of sugar chain lengths (Zhang and Lynd, 2005b) and offers a much higher sensitivity as shown in Table 3 (Zhang and Lynd, 2005b). Soluble oligosaccharides and their chromophoresubstituted substrates, such as p-nitrophenyl glucosides and methylumbelliferyl--D-glucosides, are also used to measure endoglucanase activities based on the release of chromophores or the formation of shorter oligosaccharide fragments, which are measured by HPLC or TLC (Bhat et al., 1990; Claeyssens and Aerts, 1992; van Tilbeurgh and Claeyssens, 1985; Zverlov et al., 2002a, 2002b, 2003, 2005). Endoglucanase activities can also be easily detected on agar plates by staining residual polysaccharides (CMC, cellulose) with various dyes because these dyes are adsorbed only by long chains of polysaccharides (Flp and Ponyi, 1997; Hagerman et al., 1985; Jang et al., 2003; Jung et al., 1998; Kim et al., 2000; Murashima et al., 2002a; Piontek et al., 1998; Rescigno et al., 1994; Ten et al., 2004). These methods are semi-quantitative, and are well suited to monitoring large numbers of samples. Precision is limited because of the relationship between the cleared zone diameters and the logarithm of enzyme activities. For example, differences in enzyme activity levels less than 2-fold are difficult to detect by eye (Sharrock, 1988). Unfortunately, most exoglucanase activities are not detected by these methods, since the processive action of exoglucanases is blocked by carboxymethyl substitutions, which prohibits cellulose chain from shortening. The lack of efficient exoglucanase plate screening method explains some of the difficulty in detecting exoglucanase genes cloned from C. thermocellum (Demain et al., 2005).

4.2.2. Exoglucanases Exoglucanases cleave the accessible ends of cellulose molecules to liberate glucose and cellobiose. T. reesei cellobiohydrolase (CBH) I and II act on the reducing and non-reducing cellulose chain ends, respectively (Teeri, 1997; Teeri et al., 1998; Zhang and Lynd, 2004b). Avicel has been used for measuring exoglucanase activity because it has the highest ratio of FNR/Fa among insoluble cellulosic substrates (Table 1). During chromatographic fractionation of cellulase mixtures, enzymes with little activity on soluble CMC, but showing relatively high activity on avicel, are usually identified as exoglucanases. Unfortunately, amorphous cellulose and soluble cellodextrins are substrates for both purified exoglucanases and endoglucanases. Therefore, unlike endoglucanases and glucosidases, there are no substrates specific for exoglucanases within the cellulase mixtures (Sharrock, 1988; Wood and Bhat, 1988). Claeyssens and his coworkers (van Tilbeurgh et al., 1982) found that 4-methylumbelliferyl--D-lactoside was an effective substrate for T. reesei CBH I, yielding lactose and phenol as reaction products, but it was not a substrate for T. reesei CBH II (van Tilbeurgh and Claeyssens, 1985) and some endoglucanases (van Tilbeurgh et al., 1982). T. reesei EG I, structurally homologous to CBH I, also cleaves 4-methylumbelliferyl--D-lactoside, yet these enzymes can be differentiated by adding cellobiose, an inhibitor that strongly suppresses cellobiohydrolase activity (Claeyssens and Aerts, 1992). T. reesei CBH II does not hydrolyze 4methylumbelliferyl--D-aglycones of either glucose or cellobiose units, but does cleave 4-methylumbelliferyl-D-glycosides with longer glucose chains (van Tilbeurgh et al., 1985). Deshpande et al. (1984) reported a selective assay for exoglucanases in the presence of endoglucanases and glucosidases. This assay is based on the following: (1) exoglucanase specifically hydrolyzes the aglyconic bond of p-nitrophenyl--D-cellobioside to yield cellobiose and p-nitrophenol, (2) -glucosidase activity is inhibited by D-glucono-1,5--lactone (Holtzapple et al., 1990), and (3) the influence of exoglucanase hydrolysis activities must be quantified in the assay procedure in the presence of added purified endoglucanases. However, this technique has its own limitations: (1) CBH II activity cannot be measured using p-nitrophenyl--Dcellobioside, (2) the specific activity of the available purified endoglucanases may not be representative of all existing endoglucanases in the mixture, and (3) the product ratio from endoglucanase actions may be influenced by the presence of exoglucanases.

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4.2.3. -D-glucosidases -D-glucosidases hydrolyze soluble cellobiose and other cellodextrins with a DP up to 6 to produce glucose in the aqueous phase. The hydrolysis rates decrease markedly as the substrate DPs increase (Zhang and Lynd, 2004b). The term cellobiase is often misleading due to this key enzyme's broad substrate specificity beyond a DP of 2. -D-glucosidases are very amenable to a wide range of simple sensitive assay methods, based on colored or fluorescent products released from pnitrophenyl -D-1,4-glucopyranoside (Deshpande et al., 1984; Strobel and Russell, 1987), -naphthyl--Dglucopyranoside, 6-bromo-2-naphthyl--D-glucopyranoside (Polacheck et al., 1987), and 4-methylumbelliferyl--D-glucopyranoside (Setlow et al., 2004). Also, -D-glucosidase activities can be measured using cellobiose, which is not hydrolyzed by endoglucanases and exoglucanases, and using longer cellodextrins, which are hydrolyzed by endoglucanases and exoglucanases (Ghose, 1987; Gong et al., 1977; McCarthy et al., 2004; Zhang and Lynd, 2004b). 4.2.4. Total cellulase The total cellulase system consists of endoglucanases, exoglucanases, and -D-glucosidases, all of which hydrolyze crystalline cellulose synergically. Total cellulase activity assays are always measured using insoluble substrates, including pure cellulosic substrates such as Whatman No. 1 filter paper, cotton linter, microcrystalline cellulose, bacterial cellulose, algal cellulose; and cellulose-containing substrates such as dyed cellulose, -cellulose, and pretreated lignocellulose. The heterogeneity of insoluble cellulose and the complexity of the cellulase system cause formidable problems in measuring total cellulase activity. Experimental results show that the heterogeneous structure of cellulose (filter paper and bacterial cellulose) gives rise to a rapid decrease in the hydrolysis rate within a short time (less than an hour), even when the effects of cellulase deactivation and product inhibition are taken into account (Valjamae et al., 1998; Zhang et al., 1999). In an attempt to clarify this situation, a functionally based model has been developed to demonstrate that the degree of synergism between endoglucanase and exoglucanase is influenced by substrate characteristics, experimental conditions, and enzyme loading/composition ratio (Zhang and Lynd, in press). This model clearly suggests the complexity of total cellulase activity assays and infers that it is nearly impossible to apply the results of the total cellulase activity assay measured on one solid substrate to a different solid substrate. This is one of the reasons that the U.S. DOE-sponsored cellulase

development projects, conducted by Genencor International and Novozymes Biotech, tailored cellulase mixture performance based only on an identical sampledilute acid pretreated corn stover substrate that was prepared in the pilot plant of the National Renewable Energy Laboratory (Golden, CO) (Knauf and Moniruzzaman, 2004). The most common total cellulase activity assay is the filter paper assay (FPA) using Whatman No. 1 filter paper as the substrate, which was established and published by the International Union of Pure and Applied Chemistry (IUPAC) (Ghose, 1987). This assay requires a fixed amount (2mg) of glucose released from a 50-mg sample of filter paper (i.e., 3.6% hydrolysis of the substrate), which ensures that both amorphous and crystalline fractions of the substrate are hydrolyzed. A series of enzyme dilution solutions is required to achieve the fixed degree of hydrolysis. The strong points of this assay are (1) it is based on a widely available substrate, (2) it uses a substrate that is moderately susceptible to cellulases, and (3) it is based on a simple procedure (the removal of residual substrate is not necessary prior to the addition of the DNS reagent). However, the FPA is reproduced in most laboratories with some considerable effort and it has long been recognized for its complexity and susceptibility to operators' errors (Coward-Kelly et al., 2003; Decker et al., 2003). Reliability of results could be influenced by (1) the -D-glucosidase level present in the cellulase mixture (Breuil and Saddler, 1985a,b; Schwarz et al., 1988; Sharrock, 1988), because the DNS readings are strongly influenced by the reducing end ratio of glucose, cellobiose, and longer cellodextrins (Ghose, 1987; Kongruang et al., 2004; Wood and Bhat, 1988; Zhang and Lynd, 2005b); (2) the freshness of the DNS reagent, which is often ignored (Miller, 1959); (3) the DNS reaction conditions, such as boiling severity, heat transfer, and reaction time (Coward-Kelly et al., 2003); (4) the variations in substrate weight based on the area size (1 6 cm a strip), because this method does not require substrate excess (i.e., substrate amounts strongly influence enzyme activity) (Griffin, 1973); and (5) filter paper cutting methods, because the different papercutting methods such as paper punching, razoring, or scissoring could lead to different accessible reducing ends of the substrate (Zhang and Lynd, 2005b). Dyed celluloses are widely used for determining sugar inhibition for total cellulase because they avoid the high background interference from added sugars (Gusakov et al., 1985c; Holtzapple et al., 1984; Wood, 1988). Fluorescent-dyed cellulose is also used for the same purpose, and the higher signal per molecule of


465

fluorescent dye permits detection of lower cellulase activities. Researchers should consider the following: (1) the calibration curve between dye release and reducing sugar accumulation should be established for each batch of substrate, because dye adsorption depends on cellulosic substrate properties and preparation conditions; (2) the calibration curve works only for a small hydrolysis conversion range, because dye molecules cannot enter into the internal cellulose structure; and (3) the different hydrolysis modes of endoglucanases and exoglucanases have different dye release preferences (Helbert et al., 2003). Using dyed cellulose, Holtzapple et al. (1984) showed that glucose and cellobiose were noncompetitive inhibitors to the T. reesei cellulase. On the contrary, the T. longibrachiatum cellulase was competitively inhibited by cellobiose and glucose (Gusakov et al., 1985c). Some feel that the different inhibition patterns may be attributed to large variations in characteristics of dyed celluloses (Gruno et al., 2004). Cotton fiber, microcrystalline cellulose, bacterial cellulose, and algal cellulose are several other common pure cellulosic substrates. Powder microcrystalline cellulose could become a preferred substrate to replace filter paper because (1) it can be rapidly dispensed volumetrically as a slurry and thus permits robotics methods; (2) it can be easily pelleted by centrifugation, and the total sugars released are measured more exactly by the phenol-H2SO4 method than by the DNS assay; (3) it is a more recalcitrant substrate, yielding a more stringent substrate for total cellulase activity than does filter paper; and (4) activities measured on microcrystalline cellulose could more accurately represent hydrolysis ability on pretreated lignocellulose, because its characteristics are closer to those of pretreated lignocelluloses, based on cellulose accessibility to cellulase and the degree of polymerization (Zhang and Lynd, 2004b). Sigmacell-20, a readily available microcrystalline cellulose powder, could also be a good alternative substrate for a total cellulase activity assay, replacing Whatman No. 1 filter paper. Keep that in mind, some of the pretreated lignocellulose still contains significant amounts of hemicellulose and lignin, while microcrystalline cellulose does not contain hemicellulose and lignin. -Cellulose and pretreated lignocellulose are often used to evaluate the digestibility of commercial cellulase or of a reconstituted cellulase mixture for a prolonged reaction. The primary difference, as compared to cellulase activity assays using model cellulosic substrates, is the time required for assays, which ranges from several minutes to hours for model substrates

(initial hydrolysis rate) to several days for pretreated lignocellulose to obtain the final digestibility (cellulose conversion). Clearly, the presence of hemicellulose and even lignin results in more complexity. Again, the desired outcome of the experiment must indicate the substrate chosen, especially in the case of total cellulase performance. In conclusion, the measurement of isolated individual cellulase activity is relatively easy, but it is still challenging to measure T. reesei CBH I and CBH II activities specifically in the presence of endoglucanases. There is no clear relationship between the hydrolysis rates obtained on soluble substrates and those on insoluble substrates, mainly because of huge differences in substrate accessibility and DP. For insoluble cellulose, it is highly unlikely that any substantial solubilization of crystalline or semicrystalline cellulose will proceed linearly with time, due to varying -glucosidicbond accessibilities and chain end availability for different regions of fibers. Researchers must state clearly all parameters of their assay conditions, and resist temptation to compare their results to those of other researchers using different substrates, assay methods, etc. For example, the specific activity of Thermobifida fusca YX endoglucanase is reported to be at least ten-fold higher than that of T. reesei endoglucanase on soluble CMC (Himmel et al., 1993); however, this activity ratio is not maintained if the assays are performed with insoluble cellulose (Himmel et al., 1999). 5. Cellulase improvement and screening/selection Two strategies are available for improving the properties of individual cellulase components: (1) rational design and (2) directed evolution. 5.1. Rational design Rational design is the earliest approach to protein engineering, was introduced after the development of recombinant DNA methods and site-directed mutagenesis more than 20 years ago, and is still widely used. This strategy requires detailed knowledge of the protein structure, of the structural causes of biological catalysis or structure-based molecular modeling, and of the ideally structurefunction relationship. As shown in Fig. 3, the process of rational design involves (1) choice of a suitable enzyme, (2) identification of the amino acid sites to be changed, based usually on a high resolution crystallographic structure, and (3) characterization of the mutants. The availability of data on the protein

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Protein structure

Structure-based molecular modeling

Site-directed mutagenesis

Repeat(optional)

Transformation and Expression

Characterization of mutants

Fig. 3. Scheme of rational protein design.

structure of an enzyme or of homologous proteins typically governs the choice of a suitable enzyme for modification. The identification of the region of the protein to be modified generally requires the knowledge of not only the existing function of the region but also the desired modified or new function. The modification of amino acid sequence can be achieved through sitedirected mutagenesis, exchange of elements of secondary structure, and even exchange of whole domains and/ or generation of fusion proteins. The faith in the power of rational design relies on the belief that our current scientific knowledge is sufficient to predict function from structure. But such information of structures and mechanisms is not available for the vast majority of enzymes. Even if the structure and catalysis mechanism of the target enzyme are well characterized, the molecular mutation basis for the desired function may not be achieved (Arnold, 2001). Rational design appears to be a logical method for researchers to examine possible amino acid sites near to the active site or the binding pocket in a 3-dimensional structure (Bornscheuer and Pohl, 2001). But many important enzymatic properties are not localized in a small number of catalytic residues a priori. Indeed, many residues distributed over large parts of the protein often confer important properties. Even when large functional changes can be obtained with a few amino

acid substitutions, it will often be difficult or impossible to discern the specific mutations responsible. For example, a significant increase (10 6 -fold) in the specificity constant (kcat/KM) of aspartate aminotransferase favoring valine requires 17 amino-acid changes, only one of which occurs within the active-site (Benkovic and Mames-Schiffer, 2003). Recently, a successful computational design to convert non-active ribose binding protein to triose phosphate isomerase was based on 1822 mutations and exhibited a 105106 fold activity enhancement (Dwyer et al., 2004). Unfortunately the success of computational models is often limited to well-understood reactions and enzymes. Different from most enzymes catalyzing soluble substrates in the aqueous phase, cellulase acting on insoluble heterogeneous cellulose is a more complex process, involving: (1) the changes in heterogeneous cellulose characteristics during hydrolysis (Banka et al., 1998; Boisset et al., 2000; Chanzy et al., 1983; Din et al., 1991, 1994; Halliwell and Riaz, 1970; Lee et al., 1996, 2000; Saloheimo et al., 2002; Walker et al., 1990, 1992; Wang et al., 2003; Woodward et al., 1992; Zhang and Lynd, 2004b); (2) cellulase diffusion, adsorption, and catalysis on the surface of cellulose, i.e., decreases from a 3-dimension diffusion (in liquid phase) to a 2dimension diffusion (on solid surfaces) (Henis et al., 1988; Katchalski-Katzir et al., 1985) and even 1-


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dimension processivity along cellulose chains for cellobiohydrolases (Teeri, 1997); (3) the non-productive cellulase binding on the cellulose surface (Beldman et al., 1987; Sheehan and Himmel, 1999); and (4) the yet unexplained dynamic interactions among the cellulosebinding module (CBM), the catalytic domain, and a single glucan chain end lifted from the cellulose surface (Skopec et al., 2003). Several excellent reviews summarize numerous studies using site-directed mutagenesis for investigating cellulase mechanisms and improving enzyme properties (Schulein, 2000; Wilson, 2004; Wither, 2001). Not surprisingly, few researchers using site-directed mutagenesis have reported successful examples of significantly higher activity cellulase mutants on insoluble substrates (Escovar-Kousen et al., 2004; Sakon et al., 1996; Zhang et al., 2000a,b; Zhang and Wilson, 1997). One clear example, however, is the report by Baker and coworkers of a 20% improvement in the activity on microcrystalline cellulose of a modified endoglucanase Cel5A from Acidothermus cellulolyticus (Baker et al., 2005). The Cel5A endoglucanase, whose high-resolution crystallographic structure has been available (Sakon et al., 1996), was subjected to a series of mutations designed to alter the chemistry of the product-leaving side of the active site cleft. Using structural information and following a thesis that end product inhibition could be relieved by a substitution of non-aromatic residue at site 245, a mutant (Y245G) was shown to increase KI of cellobiose by 15-fold. However, today there are no general rules for sitedirected mutagenesis strategies for improving cellulase activity on solid cellulase substrates and it remains in a trial-and-test process.

5.2. Directed evolution Our still limited knowledge about the characteristics of insoluble cellulose substrates, the dynamic interactions between cellulases and insoluble substrates, and the complex synergetic and/or competitive relationships among cellulase components, significantly limits rational design for improving cellulase properties, despite increasing understanding of cellulase structures and hydrolysis mechanisms, characterization of cellulose properties, and cellulase adsorption (Bothwell et al., 1997; Bothwell and Walker, 1995; Bourne and Henrissat, 2001; Lynd et al., 2002; Wither, 2001; Zhang and Lynd, 2004b). In 1999, Michael Himmel (Sheehan and Himmel, 1999) wrote: non-informational approaches to protein engineering should be used to complement existing efforts based on informational or rational design strategies in order to ensure success of the DOE cellulase improvement program. One approach to non-informational mutant identification is irrational design using directed evolution. The greatest advantage of directed evolution is that it is independent of knowledge of enzyme structure and of the interactions between enzyme and substrate. The greatest challenge of this method is developing tools to correctly evaluate the performance of mutants generated by recombinant DNA techniques. The success of a directed evolution experiment depends greatly on the method chosen for finding the best mutant enzyme, often stated as you get what you screen for (Hibbert et al., 2005; Schmidt-Dannert, 2001; Schmidt-Dannert and Arnold, 1999) (see Fig. 4). Table 4 lists the published examples of the cellulases with properties altered using directed evolution. Four

Fig. 4. Scheme of directed protein evolution.

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Table 4 List of cellulases and relevant enzymes whose properties have been changed using directed evolution techniques Enzyme Endoglucanase Endoglucanase Endoglucanase Endoglucanase -D-glucosidase -D -glucosidase -D -glucosidase -D -glucosidase -glycosidase Mutated -glucosidase (glycosynthase) Mutated endoglucanase (glycosynthase) Altered property Thermal stability Activity Alkali pH Cold adoption Thermal stability Thermal stability Activity Activity Activity Activity DNA technique Family shuffling DNA shuffling epPCR Family shuffling DNA shuffling epPCR epPCR + Family shuffling epPCR Family shuffling epPCR cassette mutegenesis Screening/Selection Facilitated screening-Congo red + CMC agar Facilitated screening-Congo red + CMC agar Facilitated screening-Congo red + CMC agar Facilitated screening-Congo red + CMC agar Random Screening-chromogenic substrate Random Screening-chromogenic substrate Random Screening-chromogenic substrate Random Screening-coupled to color reaction Random Screening-chromogenic substrate Facilitated Screening-fluorogenic substrate Chemical complementation Ref. Murashima et al., 2002b Kim et al., 2000 Wang et al., 2005 Catcheside et al., 2003 Lebbink et al., 2000 Gonzalez-Blasco et al., 2000 Arrizubieta and Polaina, 2000 McCarthy et al., 2004 Kaper et al., 2002 Kim et al., 2004 Lin et al., 2004

directed evolution examples have been reported for endoglucanases, all of which are identified by facilitated screening on solid plates containing CMC, followed by Congo Red staining (Catcheside et al., 2003; Kim et al., 2000; Murashima et al., 2002a; Wang et al., 2005). Kim et al. (2000) reported that a 5-fold higher specific activity Bacillus subtilis endoglucanase mutant was found by screening cellulase mutants, generated by DNA shuffling and displayed on the surface of E. coli by fusion of the Pseudomonas syringae ice-nucleation protein. Doi et al. (Murashima et al., 2002b) enhanced the thermostability of an endoglucanase by seven-fold using the family shuffling technique based on the parental Clostridium cellulosomal endoglucanasesEngB and EngD. Gao et al. (Wang et al., 2005) found that a T. reesei EG III mutant generated using the errorprone PCR technique and expressed in Saccharyomyces cerevisiae was found to have an optimal pH of 5.4, corresponding to a basic pH shift of 0.6. Another example identified hybrid mutants using the family shuffling technique for T. reesei cel12A and Hypocrea schweinitzii cel12A genes (Catcheside et al., 2003). -D-glucosidase mutants have been reported to be screened blindly using 96-microplate wells because of lack of facilitated screening tools (Arrizubieta and Polaina, 2000; Gonzalez-Blasco et al., 2000; Lebbink et al., 2000; McCarthy et al., 2004). Improvements in the low temperature catalysis (3-fold) for the hyperthermostable Pyrococcus furiosus -D-glucosidase CelB (Lebbink et al., 2000) and the thermostabilities and catalytic efficiencies for the Paenibacillus polymyxa BgblA and BglA were obtained using the chromogenic substrate, p-nitrophenyl--D-glucopyranoside (Arrizu-

bieta and Polaina, 2000; Gonzalez-Blasco et al., 2000). The hydrolysis rate of the Thermotoga neapolitana 1,4-D-glucan -glucohydrolase (GghA) (EC 3.2.1.74) mutant is increased by 31% after error-prone PCR mutagenesis, in which blind screening was based on glucose released from a non-chromogenic substrate (cellobiose) and measured by the coupled reactions of thermostable glucokinase and glucose-6-phosphate dehydrogenase (McCarthy et al., 2004). In another recent example, after DNA family shuffling, a glycosidase mutant was found to display lactose hydrolysis rates 3.5-fold and 8.6-fold higher than the parental P. furiosus CelB and Sulfolobus solfataricus LacS, respectively, where glucose released from lactose was measured using a coupled glucose oxidase and phenol 4-aminophenazone peroxidase reaction (Kaper et al., 2002). In some cases, glycosyl hydrolases, e.g., Agrobacterium sp. -D-glucosidase, can be converted to glycosynthases by site-directed mutagenesis (Mackenzie et al., 1998). There is no intrinsic way to screen or select for glycosynthase activities today. The specific activity of glycosynthase from Agrobacterium sp. -D-glucosidase was improved (Kim et al., 2004) using a novel coupledenzyme assay and screening on solid plates because another endoglucanase releases fluorophores from the fluorogenic product synthesized by glycosynthase (Mayer et al., 2001). Another selection method for a glycosynthase mutant library is the chemical complementation method (Lin et al., 2004), based on the principle that the glycosynthase activity is linked to the transcription of a LEU2 reporter gene, resulting in cell growth dependant on glycosynthase activity. A 5-fold


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higher activity of glycosynthase is obtained using this approach (Lin et al., 2004). Although a number of successful examples using directed evolution for desired cellulases have been published, the largest limitation of all current selection and screening methods is based on soluble substrates. It is still very challenging to design a method to screen or select cellulase mutants using solid cellulosic substrates. 5.3. Screening The screening strategy is a critical step for finding the desired mutants from a large mutant library. Screening can be divided into two categories: (1) facilitated screening, which distinguishes mutants on the basis of distinct phenotypes, such as chromospheres released or halos formed, and (2) random screening, which picks mutants blindly (Taylor et al., 2001). A typical facilitated screening, carried out on solid agar, relies on product solubilization followed by an enzymatic reaction that gives rise to a zone of identity, such as chromophores released from chromogenic substrates. The assays may be coupled to a second enzyme whose product can in turn be easily monitored, as demonstrated by a successful coupling for cytochrome P450 to horseradish peroxidase (Joo et al., 1999a,b). With the help of microscopic plate images, it is feasible to screen a much larger number of clones on solid plates (e.g., several hundreds per cm2) (Delagrave et al., 2001; Joo et al., 1999b; Youvan et al., 1995). Recently, an ultra-high throughput facilitated screening method, based on solid microbeads, has been developed in which single cells containing mutant genes are immobilized on solid beads. After a chromogenic substrate is applied, stronger colored beads containing desired mutants are identified under the microscope (Freeman et al., 2004). Another facilitated screening method, conducted in the liquid phase, applies a flow cytometer for detecting chromospheres released from chromogenic substrates, which are catalyzed by the celldisplayed enzyme. Numerous reviews pertaining to cell surface displayed enzyme library screening by flow cytometers are available elsewhere (Aharoni et al., 2005; Becker et al., 2004; Cohen et al., 2001; Goddard and Reymond, 2004; Lin and Cornish, 2002; Wahler and Reymond, 2001; Wittrup, 2001). Endoglucanase activities are detected easily by examination of halos on solid agar plates using CMC as the substrate, followed by Congo Red staining and washing. Higher hydrolysis rates of mutants usually result in larger halos (in Section 4.2.1). It is not

surprising that all reported endoglucanase examples using directed evolution have been screened using the CMC/Congro Red method (in Section 5.2). It may be operative to screen exoglucanase mutants on solid plates using soluble chromogenic substrates, such as nitrophenol-cellobioside. However, it is worth noting that the best screening methods for endoglucanases and exoglucanases, capable of hydrolyzing insoluble cellulose, must be implemented on insoluble cellulose rather than on soluble cellulose derivatives. Random screening is another choice, if facilitated screening is not available. It is often implemented using 96-well microtiter plates, although some researchers are moving towards 384-well and higher density plate formats with the help of accurate, low-volume dispensing instruments (Sundberg, 2000). For example, Diversa has developed an ultra-throughput screening platform, the Gigamatrix, having 400,000 wells containing only 50 nL of liquid substrate per well (Wolfson, 2005). But the reformatting of 96-well plates into higher density requires high assay sensitivity and high evaporation control. Additional product measurement can be achieved using HPLC, mass spectrometry, capillary electrophoresis, or IR-thermography (Wahler and Reymond, 2001). A number of improved -glycosidase mutants after random mutagenesis are found using 96-well microplates, as reported in Section 5.2. Recently, in order to measure total cellulase activity, the FPA has been miniaturized from a 1.5-ml enzyme solution to 60 L, which is implemented in a 96-microplate well (Xiao et al., 2004). Water evaporation from samples is prevented using a PCR thermocycler having a built-in 105 C hot lid. Also, Decker and coworkers (2003) have developed a high throughput cellulase assay system using 96microplates equipped in a Cyberlabs C400 robotics deck with the substrates such as Whatman No. 1 filter paper disks (0.25in. diameter), Solka-Floc, SigmaCell-20, Avicel PH101 (FMC, Philadelphia, PA), and cotton linters (Fluka/Sigma Aldrich). This custom system has a maximum output of 84 samples per day and produces values that correlate to the traditional FPA. However, no application of these systems to the screening of higher activity cellulases has been reported. Considering the inherent limitations of the FPA (see Section 4.2.4), this automated approach could be benefited by replacing the DNS method with the phenol-H2SO4 method because the latter (1) has a higher sugar sensitivity (Table 3), (2) is independent of oxygen presence (unlike the DNS reagent) (Miller, 1959), especially for miniaturization that has a very high surface/volume ratio, (3) yields a strict stoichiometric relationship between color

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formation and total soluble sugars released, and (4) is thus independent of -D-glucosidase levels. Different from FPA, the recommended method requires centrifugation for soluble sugars and solid cellulose residue prior to the phenol-sulfuric acid assay. 5.4. Selection Selection is always preferred over screening because it has several-order-of-magnitude higher efficiency than screening (Griffithsa et al., 2004; Olsen et al., 2000; Otten and Quax, 2005). However, selection requires a phenotypic functional link between the target gene and its encoding product that confers selective advantage to its producer. This method is often implemented based on the principles of resistance to cytotoxic agents (e.g., antibiotics) (Stemmer, 1994a,b) or of complementation of auxotroph (Griffithsa et al., 2004; Jurgens et al., 2000; Smiley and Benkovic, 1994). Today, selection on solid media in petri dishes (200300 colonies per dish) is commonly used because a large number of mutants can be identified conveniently by visual inspection of growth or zones around the colonies as a consequence of a diffusing product. Recently, more and more attention has been paid to the traditional liquid selection (or enrichment) technique (Sauer et al., 2000). An exceedingly higher cell concentration of 1012 individual cells per liter and a longer cultivation time (generations) allow continuous culture to become a powerful selection system for the ultra-large size of the mutant library even when selective advantages are very small. Experimental selection strategy is associated with (1) the selection ratio, (2) the location of the targeted enzyme, and (3) the solubility of the substrate for the

targeted enzyme. Fig. 5 presents possible selection strategies and real examples. The selection ratio (RS) is defined as l RS mutant lparent in which mutant and parent are the specific growth rates of the beneficial mutant strain and the parental control, respectively, in terms of time (h- 1). RS values could vary from 1 to infinity. A value of one implies no selective advantage for the mutants in growth rates, and a value of infinity implies an essentially infinite selective advantage, e.g., no growth for wild strains in the presence of antibiotic. When RS > 1.2, a selection experiment can be implemented preferentially on solid media because of easy operation. When RS < 1.2, liquid continuous culture could be the only choice, because a limited number of generations (e.g., 25) cannot generate the significant difference detected, but this limitation can be overcome by employing the continuous culture chemostat. The location of the targeted enzyme is another important concern for selection. Selection for intracellular enzymes is relatively easy, because such benefit is confined to the same organism that produces the protein. A typical example is kanamycin selection, in which only bacteria containing the aminoglycoside phosphototransferase (kan) gene expressed intracellularly can grow in the presence of kanamycin. Other examples of carbohydrate-metabolizing enzymes are intracellular -galactosidase (Horiuchi et al., 1962) and cell-membraneassociated lactose permease (Tsen et al., 1996), when lactose is the controlling growth carbon source. A Saccharomyces cerevisiae mutant containing several recombinant key intracellular genes for xylose

Enzyme LocationIntracellular High RS: Yes. Solid Media Extracellular - tethered Extracellular - secreted High RS: Yes. Solid Media e.g. -lactamase gene in the presence of ampicillin Low RS: Yes. Microcolonization e.g. subiltis gene on BSA

Substrate Solubility

Soluble Insoluble

e.g. kan gene in the presence of kanamycin Low RS: Yes. Chemostat e.g. -galactosidase gene on lactose

High RS: No.

High RS: Yes. Solid Media. e.g. Isolation of cellulolytic microorganisms in the presence of cellulose

Low RS: No.

Low RS: Possible. No example

Low RS: No.

Fig. 5. Selection strategies depending on selection ratio (RS), enzyme location, and substrate solubility.


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catabolism has been selected to utilize xylose efficiently under anaerobic conditions after 460 generations (Sonderegger and Sauer, 2003). When a key enzyme is secreted extracellularly, the benefit from this enzyme is shared with others, resulting in cross feeding. A typical example is the growth of plasmid-deficient stains in liquid fermentations when fermentations are prolonged or the formation of micro-colonies on petri plates because the secreted -lactamase degrades ampicillin with time. A novel method, called microcolonization, has been invented to solve cross feeding challenges for a secreted enzyme (Naki et al., 1998). To employ this method, cells are compartmented into hollow fibers and each single colony grows in its own segment. When soluble substrate is fed to each single cell, diffusion of the secreted enzymes and hydrolytic products (cross feeding) is limited spatially, resulting in effective selective pressure. A 5-fold greater protease-producing mutant was obtained using bovine serum albumin as a sole nitrogen source and by this selection technique (Naki et al., 1998). The examples pertaining to the influence of enzyme location on evolution of specific cellulase activity could exist in nature. The genes encoding the secretory T. reesei cellobiohydrolases are thought to have no selection pressure to increase their catalytic efficiency (Divne et al., 1994; Konstantinidis et al., 1993; Sinnott, 1998). The fact that the cell surface associated cellulase from C. thermocellum ( 2.4 IU/mg, measured at substrate-excess conditions) (Zhang and Lynd, 2003a) has several-fold higher specific activity than the secretory T. reesei cellulase ( 0.6 IU/mg, measured at substrate-excess) (Lynd et al., 2002) on crystalline cellulose could be attributed to the enzyme location. (Note, here we do not use the old data that the specific hydrolysis rate of C. thermocellum cellulase on crystalline cellulose (substrate-excess) was 50-fold higher than the T. reesei cellulase hydrolysis (substrate-limited) (Johnson et al., 1982a,b). Another comparative example (cellulase vs. amylase) also suggests the influence of different enzymes' locations on their evolution rates. The different main biological roles (cellulose as a major component of the plant cell wall and starch as an energy reserve) determine that the majority of cellulases and amylases are located extracellularly and intracellularly, respectively. Therefore, it is not surprising that amylases could have experienced higher evolution rates for achieving higher catalytic efficiency. The difference in catalytic efficiencies for two enzymes on soluble substrates is verified by the results from the Sinot laboratory (Sinnott, 1998). Furthermore, if amylases already have

a little higher specific activity than cellulases, less expression of amylase for utilizing substrates could result in a higher evolution rate than cellulases because of different expression levels (Drummond et al., 2005). Therefore, the d

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