Lignocellulolytic microorganisms produce diverse enzymes to degrade cellulose, matrix polysaccharides (i. e., hemicellulose) and even lignin, into soluble carbons to support cellular metabolism (Lynd et al. 2002; Doi 2008). Extensive examination of these degraders and active enzymes has uncovered a wide variety of biological mechanisms in lignocellulose hydrolysis. By definition, biochemical deconstruction relies on biological catalysts. In the first three conversion platforms described above, i. e., SHF, SSF, and SSCF, the enzymes are heterologously expressed, purified and added to pretreated, neutralized biomass. A long-standing goal has been to reduce costs and increase efficiency by exploiting multifunctional hydrolase complexes, or as for CBP, using organisms whose suite of enzymes provide them with hyper-degrading abilities. To give an overview of how lignocellulose is decomposed for conversion to biofuels, here we discuss the diversity and discovery, classification and action, and engineering strategies of lignocellulolytic enzymes, with a focus on cellulases, xylanases, and ligninases, and complexes of these enzymes. Throughout, barriers to efficient deconstruction will be discussed along with potential strategies to overcome them.

Decomposition of most lignocellulose biomass requires the cleavage of O-glycosidic bonds, which link sugar units to form large polysaccharides. Glycosyl hydrolases (GHs) acting on these bonds are roughly classified into endo-acting and exo-acting enzymes (Naumoff 2011). Endo-acting glycosidases cleave the internal glycosidic linkages of polymers; Exo-acting ones act on the bond between the sugar residue at the end of the chain and the rest of the polymer. GHs have versatile enzymatic properties, in terms of substrate specificity, product diversity and catalytic efficiency. Table 3 summarizes the known enzyme families that function in cellulose, xlyan and lignin hydrolysis. In addition to possessing a single hydrolase catalytic

Table 3. Classification of lignocellulolytic enzymes. Class Enzyme EC number GH families Mode of action References Cellulase Endoglucanase (endo-l,4-p-glucanase or 1,4-p-D — glucan 4-glucanohydrolase EC 3.2.1.4 GH5-10, GH12, GH18, GH19, GH26, GH44, GH45, GH48, GH51, GH61, GH74, and GH124 Hydrolyzes interior 1,4-p-D — glucosidic linkages (Lynd et al. 2002; Cantarel et al. 2009) Exo-l,4-p-glucosidase (1, 4-p-D — glucan glucanohydrolase) EC 3.2.1.74 GH1, GH3, GH5 and GH9 Hydrolyzes terminal 1,4-p — linkages to release one glucose (Lynd et al. 2002; Cantarel et al. 2009) Cellobiohydrolase (exoglucanase or 1,4-p-D-glucan cellobiohydrolase) EC 3.2.1.91 GH5-7 and GH9 Hydrolyzes 1,4-p-D-glucosidic linkages to release cellobiose (Lynd et al. 2002; Cantarel et al. 2009) p-glucosidase (cellobiase or p-D-glucoside glucohydrolase) EC 3.2.1.21 GH1, GH3, GH5, GH9, GH30 and GHU6 Hydrolyzes terminal p-D — glucose residues to yield one glucose (Lynd et al. 2002; Cantarel et al. 2009) Cellobiose phosphorylase EC 2.4.1.20 GH94 Phosphorylates cellobiose to yield glucose and glucose 1-phosphate (Cantarel et al. 2009) Cellobiose dehydrogenase EC 1.1.99.18 Oxidizes cellobiose into cellobiono-lactone (Henriksson et al. 2000; Cantarel et al. 2009; Phillips et al. 2011) Hemicellulase Endo-xylanase (1,4-p-D-xylan xylanohydrolase) EC 3.2.1.8 GH5, GH7-12, GH16, GH26, GH30, GH43, GH44, GH51 and GH62 Hydrolyzes mainly interior p-1,4-xylose linkages of the xylan backbone (Gilbert et al. 2008; Cantarel et al. 2009)

Exo-xylanase (Exo-l,4-p-D-xylanase) EC 3.2.1.37 GH1, GH3, GH30, GH39, GH43, GH51, GH52, GH54, GH116 and GH120 Hydrolyzes p-1,4-xylose linkages to release xylobioses (Gilbert et al. 2008; Cantarel et al. 2009) p-xylosidase (1,4-p-D-xylan xylohydrolase) EC 3.2.1.37 GH1, GH3, GH30, GH39, GH43, GH51, GH52, GH54, GH116 and GH120 Releases xylose from xylobiose and short chain xylooligosaccharides (Gilbert et al. 2008; Cantarel et al. 2009) a-arabinofuranosidase EC3.2.1.55 GH3, GH10, GH43, GH51, GH54 and GH62 Hydrolyzes terminal nonreducing a-arabinofuranose from arabinoxylans (Gilbert et al. 2008; Cantarel et al. 2009) a-glucuronidase EC 3.2.1.139 GH4 and GH67 Releases glucuronic acid from glucuronoxylans (Cantarel et al. 2009) Acetylxylan esterase EC 3.1.1.72 GH5 and GH11 Hydrolyzes acetylester bonds in acetyl xylans (Cantarel et al. 2009) Feruloyl-/p-coumaroyl — esterase EC 3.1.1.73 GH10 and GH78 Hydrolyzes feruloyl or p-coumaroyl ester bonds in xylans (Cantarel et al. 2009) Endomananase (p-mannanase) EC 3.2.1.78 GH5, GH9, GH26, GH44 and GH113 Hydrolyzes interior mannoglycosidic bonds in mannan-based polysaccharides (Gilbert et al. 2008; Cantarel et al. 2009) p-mannosidase (p-l,4-D-mannoside mannohydrolase) EC 3.2.1.25 GH1, GH2and GH5 Hydrolyzes and releases mannose units from the non­reducing end of mannosides (Gilbert et al. 2008; Cantarel et al. 2009; Lundell et al. 2010) Table 3. contd….

Table 3. contd. Class Enzyme EC number GH families Mode of action References Ligninase Lignin peroxidase (LiP, ligninase) EC 1.11.1.14 Catalyzes the one-electron oxidation of various aromatic compounds (Koua et al. 2009; Lundell et al. 2010) Manganese peroxidase (MnP, Mn-dependent peroxidase) EC 1.11.1.13 Catalyzes the oxidation of Mn(II) to Mn(III), which in turn can oxidize several phenolic substrates (Koua et al. 2009; Lundell et al. 2010) Laccase (benzenedioLoxygen oxidoreductase) EC 1.10.3.2 Catalyzes the oxidation of phenols, polyphenols and anilines by one-electron abstraction (Wong 2009; Arora et al. 2010) Versatile peroxidase (VP) EC 1.11.1.16 — Catalyzes the oxidation of various aromatic compounds (Koua et al. 2009; Lundell et al. 2010) Oxidases (e. g., glucose oxidase, aryl alcohol oxidase and dehydrogenase) EC 1.1.3.4 EC 1.1.3.7 EC 1.1 91 Generates H202 or conducts aldehyde-alcohol transformation (Leonowicz et al. 2001)

domain, many GHs are linked by flexible amino acid chains to an additional catalytic domain, a carbohydrate-binding module (CBM), and/or, a type I dockerin domain (Fontes et al. 2010; Naumoff 2011). CBMs bring catalytic domains close to specific substrates and increase catalytic processivity. Type I dockerin domains mediate binding to a complex of cellulose degradation proteins, called a cellulosome, which will be further discussed below (Fontes et al. 2010). Figure 6 summarizes the various enzymes that participate in cellulose and matrix polysaccharide digestion.

• Glucose •« Cellobiose Celloderxtrin Endoglucanase (2) Cellobiohydrolase m Exo-l,4-|3-glucosidase CT 3-glucosidase & Cellobiose phosphorylase a Carbohydrate-binding module (CBM)

■—a Cohesin

®l¥ Endoglucanase with dockerin

Cellobiohydrolase with dockerin 0* Exo-l,4-|3-glucosidase with dockerin Гр Other hydrolases Q/b Other dockerin-carrying components

eg. Hydrolases, protease and protease inhibitor

Hydrolase Discovery and Diversity

Extensive studies on lignocellulosic degraders and their hydrolytic mechanisms have uncovered a vast diversity of hydrolases from isolated microbes and microbial communities. In the most recent update of the CAZy database, GHs are classified into 131 families and 14 different clans, A to N, based on amino acid sequence similarities and structural folds, respectively (Cantarel et al. 2009; Naumoff 2011). Cellulases are spread across at least 12 different GH families, seven of which can be distributed into four different clans; xylanases are classified into 12 GH families (King et al. 2011). Some GH families contain both cellulases and xylanases (e. g., GH5) while others contain cellulases but no xylanases (e. g., GH7) or vice versa (e. g., GH11). Additionally, a single clan, the GH-As, contains a GH5 endoglucanase, GH26 mannanase and a GH53 endo-p-1,4-galactanase (Gilbert 2010). These observations suggest that GHs show a large diversity in structure and enzymatic activity as the result of convergent and divergent evolution.

High-throughput techniques for DNA sequencing, activity measurements, and proteomics accelerate opportunities to understand the diversity of degradation mechanisms of widely distributed lignocellulose-
degrading microorganisms. Cow rumen microbes specialize in degrading ligoncellulosic biomass, but most members of this complex community resist cultivation. To characterize biomass-degrading genes and genomes, Hess et al. (2011) sequenced metagenomic DNA from microbes adhering to plant fiber incubated in the rumen of a cow. These researchers identified 27,755 putative carbohydrate-active genes and expressed 90 candidate proteins, of which 57% were enzymatically active against cellulosic substrates (Hess et al. 2011). They also assembled 15 uncultured microbial genomes (Hess et al.

2011) . The metagenomics approach also has been used to isolate cellulolytic and xylanolytic genes from such sources as rice straw compost (Yeh et al.

2012) and the hindgut paunch of a wood-feeding ‘higher’ termite species (Warnecke et al. 2007). Metagenomic approaches have also successfully mined for genes encoding enzymes responsible for cellular tolerance to the biomass inhibitors, syringaldehyde and 2-furoic acid (Sommer et al.

2010) . Similarly, Beloqui et al. (2006) identified a novel polyphenol oxidase through activity screening of a metagenome expression library from bovine rumen microflora. Proteome-wide systems analysis of a cellulosic biofuel — producing microbes has also been conducted. For example, quantitative mass spectrometry integrated with physiological characterization revealed proteome-wide expression changes and more than 100 CAZy family proteins expressed in C. phytofermentans (Tolonen et al. 2011).