Cellulose degradation by polysaccharide monooxygenases.

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Cellulose Degradation by Polysaccharide Monooxygenases William T. Beeson,1 Van V. Vu,2 Elise A. Span,2 Christopher M. Phillips,3 and Michael A. Marletta2 1

Department of Chemistry, University of California, Berkeley, California 94720

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Department of Chemistry, The Scripps Research Institute, La Jolla, California 92037; email: [email protected]

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BP Biofuels Advanced Technology Inc., San Diego, California 92121

Annu. Rev. Biochem. 2015. 84:30.1–30.24

Keywords

The Annual Review of Biochemistry is online at biochem.annualreviews.org

copper enzymes, biofuels, redox enzymes, fungi, oxygen activation

This article’s doi: 10.1146/annurev-biochem-060614-034439

Abstract

c 2015 by Annual Reviews. Copyright All rights reserved

Polysaccharide monooxygenases (PMOs), also known as lytic PMOs (LPMOs), enhance the depolymerization of recalcitrant polysaccharides by hydrolytic enzymes and are found in the majority of cellulolytic fungi and actinomycete bacteria. For more than a decade, PMOs were incorrectly annotated as family 61 glycoside hydrolases (GH61s) or family 33 carbohydrate-binding modules (CBM33s). PMOs have an unusual surface-exposed active site with a tightly bound Cu(II) ion that catalyzes the regioselective hydroxylation of crystalline cellulose, leading to glycosidic bond cleavage. The genomes of some cellulolytic fungi contain more than 20 genes encoding cellulose-active PMOs, suggesting a diversity of biological activities. PMOs show great promise in reducing the cost of conversion of lignocellulosic biomass to fermentable sugars; however, many questions remain about their reaction mechanism and biological function. This review addresses, in depth, the structural and mechanistic aspects of oxidative depolymerization of cellulose by PMOs and considers their biological function and phylogenetic diversity.

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Contents


INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.2 THE DISCOVERY OF POLYSACCHARIDE MONOOXYGENASES . . . . . . . . . . . . 30.3 POLYSACCHARIDE MONOOXYGENASE STRUCTURE . . . . . . . . . . . . . . . . . . . . . . 30.5 Tertiary Structure and the Active Site . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.5 Electron Transfer During Catalysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.8 Substrate Binding and Regioselectivity of Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.9 THE POLYSACCHARIDE MONOOXYGENASE REACTION . . . . . . . . . . . . . . . . . .30.11 Copper-Dependent Monooxygenase Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .30.11 Single Turnover Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .30.12 DIVERSITY AND BIOLOGICAL FUNCTIONS OF POLYSACCHARIDE MONOOXYGENASES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .30.15 PERSPECTIVES AND APPLICATIONS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .30.17 Polysaccharide Monooxygenases in Industrial Biofuel Production . . . . . . . . . . . . . . . . .30.18

INTRODUCTION Plant biomass is an abundant renewable resource; in 2011, a study was released by the US Department of Energy stating that more than 1.3 billion tons of plant biomass could be sustainably produced for biofuel production, enough to provide 30% of US transportation fuels (1, 2). There are also significant opportunities to use biomass as a source of sugar for production of renewable bioproducts and chemicals with higher margins than fuels (3). The primary technological bottleneck associated with development of lignocellulosic fuels or chemicals lies in the development of an inexpensive process for the conversion of plant cell wall polysaccharides to fermentable sugars. The expense is related to the fundamental recalcitrance of lignocellulose when compared with other agricultural feedstocks, such as sucrose or starch (4, 5). Lignocellulosic biomass comprises primarily the polysaccharides cellulose and hemicellulose, plus lignin, a heterogeneous phenolic polymer (6). Cellulose, the major structural polysaccharide of the plant cell wall, is a homopolymer of β-1,4-glucose and is the most abundant biopolymer on earth. The intricate hydrogen-bonding (H-bonding) network between and within glucan chains, coupled with degrees of polymerization in excess of thousands of glucose monomers, severely limits the accessibility of hydrolytic enzymes to the glycosidic linkages. This lack of accessibility dramatically slows the hydrolytic depolymerization of cellulose (7). In nature, filamentous fungi and bacteria are the predominant degraders of plant biomass, and this process accounts for a significant fraction of the global carbon cycle (8). The accepted model for enzymatic depolymerization of cellulose is centered on hydrolytic cellulase enzymes classified as exoglucanases (or cellobiohydrolases) and endoglucanases (9–11). Cellulases contain an active-site cleft or tunnel lined with aromatic residues that facilitate the separation of a glucan chain from cellulose (12, 13). Hydrolysis of the glycosidic bond then occurs via carboxylate residue general acid/base catalysis. It has been hypothesized that the limiting step for depolymerization of crystalline cellulose by cellulases is separation of the glucan chain from the strong H-bonding network of cellulose into the active-site cleft. Recently, the conventional hydrolytic model of cellulose degradation has been challenged by several reports of the occurrence and functional importance of a novel class of copper-dependent

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polysaccharide monooxygenases (PMOs) found in both fungi and bacteria (14–16). These enzymes depolymerize cellulose through an oxidative mechanism involving hydroxylation of cellulose at the C1 or C4 carbon, leading to subsequent cleavage of the glycosidic bond. Following cleavage, an oxidized chain end is produced containing an aldonolactone or a 4-ketoaldose. The reaction requires both molecular oxygen and an extracellular electron source that can be derived from cellobiose dehydrogenase (CDH) or small-molecule reductants present in lignocellulosic biomass. This mechanism of chain cleavage is fundamentally different from that used by cellulases, presumably circumventing the energetically difficult separation of a glucan chain from highly crystalline cellulose and thereby directly creating new sites for exoglucanase action. In this review, we focus on the current understanding of copper-dependent PMO cellulose depolymerization by examining PMO discovery, structure, predicted mechanism, biological role, and potential application in biofuels or renewable chemicals production. Although our primary focus is on cellulose-active fungal PMOs, where relevant we also discuss bacterial PMOs and related enzymes that work on alternative polysaccharides. Much of what is known about the fungal cellulolytic PMOs is likely applicable across the PMO superfamily.

THE DISCOVERY OF POLYSACCHARIDE MONOOXYGENASES In the early 1990s, the first fungal PMOs were identified from complementary DNA libraries as secreted enzymes with the potential to be involved in cellulose degradation (17, 18). These enzymes were originally reported to be hydrolases and were thus annotated as family 61 glycoside hydrolase (GH61) enzymes until late 2011, when they were termed PMOs, and later LPMOs and auxiliary activity family 9 (AA9) (19). In 2001, TrCel61A, isolated from the industrial cellulase producer Trichoderma reesei, was named endoglucanase 4 due to its reported hydrolytic activity on cellulose (20). The hydrolytic activity demonstrated by TrCel61A was several hundredfold lower than that of other Trichoderma endoglucanases and has since been a matter of debate. More recent studies have suggested that this activity, which is very low on all polysaccharide substrates tested, may be due to contamination. The first clue suggesting that GH61s may not be hydrolases came from the first crystal structure of TrCel61B in 2008 (21). The structure revealed a highly conserved, flat surface, unlike the tunnel or cleft active sites found in cellulases. Notably, this protein lacked the conserved carboxylate residues that catalyze hydrolytic cleavage. The flat surface of TrCel61B has a highly conserved metal-binding site that contains a nickel ion derived from the crystallization buffer. Whereas the TrCel61B sequence showed no significant hits through BLAST (Basic Local Alignment Search Tool ) searches, a DALI (distance alignment matrix method) search revealed weak structural similarity to CBP21, a chitin-binding protein from the bacterium Serratia marcescens. Chitin is composed of β-1,4-linked N-acetylglucosamine and, like cellulose, is a crystalline polysaccharide. CBP21 had been proposed to enhance chitin degradation through a noncatalytic mechanism (22, 23). The advent of next-generation sequencing led to multiple reports on the genomes, transcriptomes, and secretomes of cellulose-degrading fungi, which demonstrated the widespread occurrence of GH61 proteins. Expression and secretion of GH61s in response to cellulose were first observed in T. reesei (24) and later in a number of other fungi (25–28). The number of GH61 genes in some fungal genomes was also unexpected, with many species of cellulolytic fungi having significantly more genes for GH61s than cellulases. More in-depth biochemical characterization of GH61s was reported in 2010, when Harris et al. (29) showed that GH61s can enhance the activity of cellulases acting on acid-pretreated corn stover, but not pure cellulose, through an unknown mechanism. This enhancing activity was

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PMO: polysaccharide monooxygenase CDH: cellobiose dehydrogenase, the biological redox partner of PMOs in fungi Family 61 glycoside hydrolases (GH61s): former classification of cellulose-active fungal PMOs Auxiliary activity family 9 (AA9): refers to the CAZypedia (encyclopedia of carbohydrate-active enzymes) classification of fungal cellulose-active PMOs TrCel61A: the first fungal PMO isolated; from Trichoderma reesei TrCel61B: the first fungal PMO crystallized (PDB 2VTC); from Trichoderma reesei (also known as HjCel61B under the species’ alternate name, Hypocrea jecorina) CBP21: chitin-binding protein 21 from Serratia marcescens; first PMO crystal structure (PDB 2BEM) and first NMR structure (PDB 2LHS)

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Family 33 carbohydratebinding modules (CBM33s): former classification of bacterial PMOs TaGH61A: a PMO3 from Thermoascus aurantiacus; crystal structure available (PDB 2YET) Type 1 PMO (PMO1): member of a subfamily of phylogenetically related PMOs that oxidize C1 in cellulose Type 2 PMO (PMO2): member of a subfamily of phylogenetically related PMOs that oxidize C4 in cellulose

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completely abolished by point mutations in residues involved in metal binding. These results were further supported by a number of related patent filings by a team of scientists at Novozymes, dating back to 2005 (30, 31). Later in 2010, Vaaje-Kolstad et al. (32) reported that the bacterial CBP21 protein is actually an enzyme that catalyzes oxidative depolymerization of chitin in the presence of molecular oxygen and a chemical reductant. The same paper also reported the incorporation of an oxygen atom from molecular oxygen into the oxidized chito-oligosaccharide products; however, this chitinolytic activity was reported to be dependent on Zn(II) and Mg(II), metal ions that cannot generate an oxidant from molecular oxygen. At that time, Vaaje-Kolstad et al. (32) proposed that a similar reaction was likely to occur in the fungal GH61s on cellulose. Key experiments linking the CBP21 oxidative cleavage reaction to fungal enzymes soon occurred through identification of extracellular electron sources capable of reductively activating GH61s (33–35). While small-molecule reductants are present in plant cell walls, most fungi also express CDH. CDH is an extracellular hemoflavoenzyme and member of the glucose–methanol– choline oxidoreductase superfamily that catalyzes the oxidation of cellobiose to cellobionolactone (36, 37). The reaction occurs at the flavin domain with subsequent electron transfer (ET) to an N-terminal heme domain. The biological electron acceptor for CDH was unknown, but had long been proposed to be an extracellular ferric complex thought to participate in Fenton chemistry, generating hydroxyl radicals that would degrade plant cell walls (38). Langston et al. (35) showed that CDH could activate GH61s; shortly thereafter, a study using a genetics approach showed that the deletion of the major CDH isoform in Neurospora crassa results in a twofold decrease in secreted cellulase activity on pure cellulose (34). The catalytically active flavin domain of CDH was unable to restore GH61 activity, suggesting that the heme domain is required for ET to GH61s. Many details about the ET from CDH to GH61s remain to be elucidated, and the role of free CDH–heme domains in many fungi is an area that merits further investigation. Two important advances followed. The first showed that copper is the functional active-site metal, and the second revealed that GH61s generate products at both the reducing and nonreducing ends of the glucan chain. Crystal structures of GH61s and bacterial family 33 carbohydratebinding modules (CBM33s) were first solved with nickel (21), magnesium, and zinc (29) in the metal-binding site. The ability to enhance polysaccharide depolymerization, first shown in 2010, was also reported with many divalent metals. In 2011, Quinlan et al. (33) determined the first crystal structure of a GH61 bound to copper by solving the structure of Thermoascus aurantiacus GH61A (TaGH61A). These authors were also the first to report that the N-terminal histidine in PMOs is methylated at Nε , although electron density suggesting methylation is clearly evident in earlier structures. At about the same time, Phillips et al. (34) reported that natively purified N. crassa GH61s contains copper and that copper is the only metal that enables catalysis by an N. crassa GH61 (NCU01050). Both of these studies also showed evidence for GH61s that oxidized the nonreducing end of sugars. Quinlan et al. proposed C6 oxidation, whereas Phillips et al. proposed C4 oxidation and a mechanism that could explain the oxidative cleavage. Experimental evidence of C4 oxidation soon followed when Beeson et al. (39) unambiguously confirmed reaction at C4. Given these new findings, we suggested changing the name of these enzymes from GH61s and CBM33s to PMOs, reflecting the common reaction catalyzed. Furthermore, we proposed classifying PMOs as type 1 and type 2 PMOs on the basis of their ability to oxidize the reducing and nonreducing ends of the glucan chain. This nomenclature was modeled on the primary fungal cellulases, cellobiohydrolases 1 and 2, which hydrolyze glycosidic linkages from the reducing and nonreducing ends, respectively. In the past few years, research on PMOs has dramatically increased, and many new discoveries related to the structure, function, and diversity of these enzymes have been reported. The remainder of this review discusses the key findings of these efforts. Beeson et al.


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POLYSACCHARIDE MONOOXYGENASE STRUCTURE The earliest indication that GH61 enzymes were not glycoside hydrolases came from the first GH61 crystal structure, TrCel61B, which was reported in 2008 (21). TrCel61B lacked the canonical active-site groove with acid/base residues found in glycoside hydrolases; instead, it presented a flat surface with a putative metal-binding site. This finding strongly suggested that this was a new enzyme family with a different mechanism of glycosidic bond cleavage. Subsequent structural studies of fungal cellulose-active PMOs performed between 2010 and 2013 have shed considerable light on PMO function, specifically with respect to active-site residues, internal ET, substratebinding interactions, and the regioselectivity of oxidation (29, 33, 40, 41). Structural studies of bacterial PMOs, many of which are chitin active, have similarly contributed to these areas of knowledge (22, 42–45). In 2014, a structure was reported to represent a new family of chitinactive fungal PMOs (46), and in the same year structures were solved for bacterial PMOs with activity toward cellulose (47). Most PMO structural studies have utilized X-ray crystallography; however, the CBP21 solution structure was solved by NMR (42), and in some cases computational studies have augmented structural analyses (40, 41).

CBM: carbohydrate-binding module; CBM1 binds cellulose

Tertiary Structure and the Active Site The overall fold of PMOs has been described as immunoglobulin- or fibronectin type III–like and consists of a β-sandwich of typically 8–10 β-strands (Figure 1a). A series of short loops between strands taper to a conical tip at the end opposite the active site; longer loops often contain short helices and form the plane of the active site. These longer loops vary greatly among PMOs; it is thought that they function in substrate recognition and exhibit variation in response to diverse and sometimes polymorphic substrates (40, 48). The fungal PMO catalytic domain also has a putative surface for CDH binding (40). In some PMOs, the catalytic domain is attached via a flexible linker to a carbohydrate-binding module (CBM1 for cellulolytic PMOs). The CBM1 structure (Figure 1b) is well characterized and contains three coplanar aromatic residues thought to be important for cellulose binding (29, 49). PMO family members typically share low sequence identity, but all structures identified to date show a high degree of structural similarity. Specifically, fungal PMO structures may exhibit pairwise root-mean-square deviations as low as ∼0.6 A˚ over ∼70% of all modeled Cα atoms, while sharing only ∼47% sequence identity [e.g., compare Protein Data Bank (PDB) entries 3EII and 4B5Q, or 2VTC and 2YET]. Most conserved residues are at or near the active-site surface, a region of ∼30 A˚ × 40 A˚. All structures solved since 2011 contain the functional active-site copper ion (33, 40, 41). Bacterial PMOs and a chitin-active fungal PMO share the same overall fold with cellulose-active fungal PMOs; bacterial PMO family members exhibit a more helical secondary structure. The copper coordination sphere observed in PMO structures unambiguously includes three nitrogen ligands contributed by two histidine residues, of which one is N-terminal, in a motif termed the histidine brace (33). These ligands form a three-coordinate, T-shaped coordination plane that is tilted ∼30◦ relative to the surface plane (40). Histidine coordination of the copper ion in PMOs is similar to that in particulate methane monooxygenase (pMMO), as well as in the high-pH form of the Cu(II)-binding site of CopC, a bacterial copper homeostasis protein (33, 34, 50–53). All X-ray structures of copper-bound, cellulose-active fungal PMOs determined to date (33, 40, 41) suffer from photoreduction of the copper center by the X-ray beam. Consequently, in most structures there is some ambiguity about the copper oxidation state and coordinating ligands, other than the three nitrogen ligands ∼2 A˚ from the copper ion. Cu(II) prefers coordination numbers www.annualreviews.org • Cellulose Degradation


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of four to six, with five being the most common, and Cu(I) prefers smaller coordination numbers of two to four (54). Cu(I)-bound cellulose-active fungal PMOs appear to be three-coordinate in the histidine brace, with additional ligands in the Cu(II) form. In PMO-Cu(II) X-ray structures, elongated distances between the copper atom and these additional ligands are what indicate partial or complete photoreduction. In particular, the conserved axial tyrosine ligand (Y175 in Figure 1c)

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is observed at distances longer than expected (>2.7 A˚) for formal Cu(II) bonding, even considering Jahn–Teller distortions arising from the d9 electronic configuration of Cu(II) (55). Likewise, an exogenous axial ligand (trans to the tyrosine) and an exogenous equatorial ligand (coplanar with the histidine brace) often appear at unusually long distances from the copper or do not appear at all. Additionally, in some structures the bond length between copper and the N-terminal amino ligand is longer than expected for Cu(II). These findings strongly suggest the presence of Cu(I) or mixed populations of Cu(I) and Cu(II) in the crystals, with averaging of the different coordination states evident in the structures. Exogenous ligands coordinating the metal ion in fungal PMO structures include water (21, 29, 33, 40, 41), the crystallization additives glycerol (41) and polyethylene glycol (33), and dioxygen species (40). The last have suggested insights into the mechanism of O2 activation (see the section titled The Polysaccharide Monooxygenase Reaction, below). NMR studies of PMO active sites are limited due to the paramagnetism of Cu(II). A helical X-ray diffraction method that minimizes the sample radiation dose was recently published, along with the unambiguous Cu(II) and Cu(I) structures of the chitin-active bacterial PMO Enterococcus faecalis CBM33A (EfCBM33A) (Figure 1d ) (45). The trigonal bipyramidal geometry of Cu(II)– EfCBM33A results from copper coordination of two water molecules in addition to the histidine brace. A conserved alanine residue restricts solvent access to the axial position, as in other bacterial PMOs (43, 44, 47, 56). In the reduced form, the coordination assumes a T-shaped geometry within the three-coordinate histidine brace, consistent with the preference of Cu(I) for smaller coordination numbers. Fungal PMOs are expected to have analogous coordination states, with the exception of octahedral geometry for oxidized enzyme, due in part to the active-site tyrosine that contributes an axial oxygen ligand and the unobstructed axial solvent position trans to the tyrosine; these coordination geometries have been confirmed with X-ray absorption spectroscopic studies (57). The active-site tyrosine in fungal PMOs may contribute to the stronger Cu(II) affinity observed in this family relative to that of bacterial PMOs, which have a conserved phenylalanine residue in that position (41). Interestingly, the N-terminal histidine has not been observed to be methylated in any bacterial PMO structure (56). The methylation is expected to be present in all fungal PMOs, although the use of expression hosts Pichia pastoris and Escherichia coli may have precluded its observation, as neither organism is predicted to possess the methyltransferase required for this modification (41, 46). Functions attributed to the methylation include alteration of the shape and/or electronic properties of the active site and a possible catalytic role (56). It also may explain the stronger Cu(II) affinity in fungal PMOs: Calculations predict that Nε -methylation increases the histidine acid dissociation constant by 0.5 units, thereby increasing copper affinity (42).

EfCBM33A: bacterial chitin-active PMO from Enterococcus faecalis; crystal structures are available of Cu(II)- and Cu(I)-bound forms (PDB 4ALC and 4ALT, respectively)

←−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−− Figure 1 (a) Structure of Thermoascus aurantiacus GH61A [TaGH61A; Protein Data Bank (PDB) entry 2YET]. Green color and stick representation indicate residues that are absolutely conserved in cellulose-active fungal polysaccharide monooxygenases (PMOs). (b) Solution structure of carbohydrate-binding molecule 1 (CBM1) (PDB 1CBH) with all planar aromatic residues depicted as sticks. The distance between Y5 and Y32 is 10.6 A˚, and the distance between Y32 and Y31 is 10.5 A˚—roughly the distance between two pyranose rings in the cellulose chain. (c) The active site of TaGH61A depicting mixed Cu(II)/Cu(I) coordination and hydrogen-bonding network. (d ) The active site of a chitin-active bacterial PMO, Enterococcus faecalis CBM33A (EfCBM33A), with (left) Cu(II) (PDB 4ALC) and (right) Cu(I) (PDB 4ALT) bound. (e) Neurospora crassa PMO3 (NcPMO3) (PDB 4EIS) shown with predicted cellobiose dehydrogenase (CDH)-binding site ( yellow), cellulose-binding surface, and putative electron transfer pathways. One pathway is conserved in all cellulose-active fungal PMOs ( green); the other is conserved in type 3 PMOs (magenta). www.annualreviews.org • Cellulose Degradation


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A conserved H-bonding network composed of residues H164, Q173, and Y175 (TaGH61A numbering) and active-site water ligands is found in the cellulose-active fungal PMO family (Figure 1c). Mutation of the glutamine residue to glutamic acid, asparagine, or leucine, which likely disrupts the H-bonding network, resulted in a loss of activity in one study (29). Crystal structures of cellulose-active bacterial PMOs (PDB entries 4OY6 and 4OY7) show that the sidechain positions in the conserved motifs R–X4 –E–X–F and H–X2 –Q–X–Y closely resemble those of H164, Q173, and Y175 in fungal PMOs (47). Two bacterial PMOs containing the H–X2 –Q– X–Y motif oxidize both the C1 and C4 positions of cellulose, whereas two bacterial PMOs with the R–X4 –E–X–F motif exclusively oxidize C1. The H-bonding network is not found in chitinactive bacterial PMOs, but the absolutely conserved E64 (PDB entry 4A02 numbering) might form a weak H-bonding interaction with a coordinated water ligand (Figure 1d ) (45). In the crystal structure of a chitin-active fungal PMO (PDB entry 4MAI) that oxidizes C1 and possibly C4, the conserved residues N136, E138, and Y140 are found in place of H164, Q173, and Y175, respectively (46). In summary, the H-bonding network is present in all cellulose-active fungal and bacterial PMOs, and C4 oxidation has been found only in PMOs with both the H-bonding network and the active-site tyrosine residue.


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Electron Transfer During Catalysis The PMO reaction requires the delivery of two electrons to the copper center. The electron donors can be found in the plant cell wall, as with gallic acid or lignin (16, 58), or can be secreted by the fungus, as with the enzyme CDH (34, 35). Sygmund et al. (59) have measured direct ET from reduced CDH to oxidized PMO. Because the active site is on a flat protein surface, it is possible for the copper center to accept electrons through direct contact with the electron donors. During the reaction, however, the active-site surface of PMOs is likely to be occupied by cellulose, which would prevent the direct contact of the copper center with electron donors, especially with macromolecules such as CDH. Interactions between cellulose and the active-site surface during the reaction may help direct the reactive intermediate(s) toward the target C–H bonds and prevent nonspecific reactions. Therefore, the delivery of the electrons is likely to occur through a protein-based ET pathway. A patch of conserved residues observed in a structure-based sequence alignment of N. crassa PMOs upregulated on cellulose link the conserved histidine of the H-bonding network (see the section titled Tertiary Structure and the Active Site, above) to another side of the protein, which contains a predicted interaction surface with CDH (Figure 1e) (40). When alternate rotamers are taken into account, distances between these conserved residues (Y215, W125, and H160 in Figure 1e) are only 3.5 to 4.5 A˚, suggesting that this network may serve as an ET pathway as found in many other proteins and protein–protein complexes (60–62). In this pathway, electrons are presumably transferred from the histidine to the copper atom via the water molecules involved in the H-bonding network (Figure 1c). These residues are absolutely conserved in all characterized cellulose-active fungal PMOs. Examination of the available crystal structures of PMOs reveals another conserved network of tyrosine, tryptophan, and histidine residues in a conserved subfamily of cellulose-active fungal PMOs (type 3 PMOs; see the section titled Substrate Binding and Regioselectivity of Oxidation, below) that creates a path from the copper center to the predicted CDH-binding site (Figure 1e) (21, 33, 40). The distance between any two consecutive residues of this network ranges from 3 to 4 A˚, suggesting that this network may also serve as an ET pathway. In chitin-active fungal PMOs, a patch of tryptophan residues and a methionine residue may facilitate ET (46); all but one of these residues are absolutely conserved in sequences of chitin-active fungal 30.8

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PMOs. This pathway traverses the larger of the β-sheets; it does not overlap with regions of cellulose-active fungal structures implicated in ET. ET in bacterial PMOs may differ from that in the fungal enzymes. There is no clear evidence to date for a bacterial CDH; however, one recent study has identified two genes in the saphrophytic bacterium Cellvibrio japonicus that contain domains with predicted redox functions and that are required for growth on crystalline cellulose (63). For cellulose-active bacterial PMOs, investigators have suggested that conserved aromatic residues in the β-sandwich play a role in internal ET (47), but structure and greater sequence analyses have not revealed an exact pathway. The structure of chitin-active bacterial PMO EfCBM33A (PDB entry 4A02) displays a patch of three tryptophan residues (W108, W119, and W176) in the core of the protein near the active site that have been attributed to a putative ET pathway in chitin-active bacterial PMOs (43). These residues are less than 5 A˚ apart, positionally conserved in the two other chitin-active bacterial PMO structures (22, 44), and at least 80% conserved in sequences of chitin-active bacterial PMOs. However, electrons in this putative pathway apparently have to transfer to the active-site phenylalanine residue, which would not facilitate ET; thus, how electrons would flow to the copper ion is unclear.

Substrate Binding and Regioselectivity of Oxidation The PMO catalytic domain is generally regarded as a relatively rigid molecule of ∼200 to 250 amino acids with disulfide bonds that contribute to structural stability. Regions thought to exhibit plasticity include loops that make up the plane of the active site, which may be involved in substrate and/or reductant binding; in fungal PMOs, these include loop regions L2, LC, and LS (Figure 1a), which tend to have higher crystallographic B-factors in deposited structures. Loop regions LC and LS make up the predicted CDH-binding groove (40); they are the most mobile substructures in molecular dynamics simulations (41). Loop flexibility at the substrate-binding interface has also been implicated in the solution structure of bacterial CBP21 (42). Structural studies of PMOs bound to cellulose have not been possible due to the insolubility of the substrate. PMOs are assumed largely to use aromatic–carbohydrate π interactions to facilitate cellulose binding. In cellulose-active fungal PMOs, aromatic residues on the putative substrate-binding plane are conserved (Figure 2a). Their arrangement resembles that of the tyrosines in CBM1 (Figure 1b) (29, 40, 41), and as in CBM1, the spacing of the aromatic residues matches the spacing between glucose subunits in crystalline cellulose (Figure 2a) (64). The distance between centers of two pyranose rings in the same cellulose chain is 5.2 A˚, and the distance between tyrosine residues in CBM1 is approximately two times that—10.5 and 10.6 A˚ in the CBM1 solution structure of TrCel7A (Figure 1b). These findings suggest that these three tyrosine residues could bind every other glucose unit over a short stretch of the cellulose polymer. In various PMOs, the distances between aromatic residues tends to be double or triple the intrachain pyranose distance (∼10 A˚ or ∼15 A˚), suggesting that they too could overlap with individual glucose units in the same chain. Alternatively, the spacing of aromatic residues in some PMOs approximately matches the interchain distance in crystalline cellulose (8.2 A˚), suggesting a perpendicular orientation across two or more polymers (Figure 2a) (40). Early bacterial PMO structures, all of chitin-active enzymes, have an abundance of conserved hydrophilic residues in the substrate-binding plane that are thought to H-bond with chitin substrates. These structures still contain one conserved aromatic residue at the active-site surface (22, 42–44). Investigators have proposed that multiple surface aromatic residues are necessary for cellulose binding on the basis of analyses of fungal PMOs, but recent structures of cellulose-active bacterial PMOs (47) have revealed only the single aromatic surface residue (Y79 in Figure 2a) that is positionally conserved in the chitin-active bacterial PMO structures (22, 43, 44). When www.annualreviews.org • Cellulose Degradation


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a

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S83 K85

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Y198

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Figure 2 (a) Phanerochaete chrysosporium GH61D (PcGH61D) structure [Protein Data Bank (PDB) entry 4B5Q], with aromatic residues shown on the active-site surface in teal stick representation. Two oligomers of crystalline cellulose are docked on the substrate-binding plane, based on the cellulose 1β structure (64). Y79 of CelS2 (PDB 4OY7; orange) shows the position of the single bacterial conserved aromatic surface residue. (b) The same surface of PcGH61D [a type 1 polysaccharide monooxygenase (PMO)] with cartoon depictions of subfamily-conserved surface motifs as observed in Neurospora crassa PMO2 (NcPMO2) (PDB 4EIR; green), NcPMO3 (PDB 4EIS; blue), and Myceliophthora thermophila PMO3∗ (MtPMO3∗ ) ( gold ). Select subfamily-conserved residues are depicted as sticks; teal coloring of S83 indicates conservation in both the PMO2 (serine, S) and PMO3 (threonine, T) subfamilies. The copper ion is coordinated by the histidine brace (dark gray sticks).

Type 3 PMO (PMO3): member of a subfamily of phylogenetically related PMOs that can oxidize C1 and C4 in cellulose PMO3∗ : member of a subfamily of phylogenetically related PMOs related to type 3 PMOs that oxidize only C1 in cellulose

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aligned with cellulose-active fungal PMO structures, the bacterially conserved aromatic residue appears at approximately the same distance (∼17 A˚) from the copper center as the nearest aligned fungal conserved aromatic residue (Y28 in Figure 2a), but at an angle ∼25◦ from it in the surface plane. The regioselectivity of oxidation is largely thought to be due to the positioning of a PMO on a specific substrate. Cellulose-active PMOs have been classified into types or subfamilies per the correlation between phylogeny and observed regioselectivity of oxidation. Systematic studies with phylogenetic analyses and activity assays showed that the majority of fungal and bacterial cellulose-active PMOs can be divided into three main groups: (a) type 1 PMOs, which oxidize C1; (b) type 2 PMOs, which oxidize C4; and (c) type 3 PMOs, which can oxidize both C1 and C4 (47, 65). There is also a subset of type 3 PMOs (termed the PMO3∗ subfamily) in fungi that appear to have lost C4 activity and can carry out only C1 oxidation. In addition to the globally conserved residues in all cellulose-active fungal PMOs, there are subfamily-specific residue distributions (40, 65), some of which form conserved secondary structures in the plane of the active site (Figure 2b). These substructures are thought to influence cellulose binding so as to affect the orientation of the PMO catalytic center relative to the glycosidic bond, in effect poising the copper center to act on one C–H bond or the other. For example, a distinctive feature of type 3 fungal PMO sequences is an insert of ∼12 amino acids starting around residue 20. Crystal structures of NcPMO3, TaGH61A, and TrCel61B show that this inserted sequence forms an extended L2 loop with a short helix. Deletion of this sequence from a PMO3 from N. crassa (NCU07760) resulted in a loss of C4 oxidation products (65), and it has been proposed that this extended L2 loop in type 3 PMOs affects substrate positioning and thus active-site orientation at the glycosidic bond. The PMO3∗ subfamily also contains an elongated Beeson et al.



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L2 loop compared with that of type 1 PMOs; however, it is shorter than the PMO3 loop and does not contain any helical secondary structure. Another feature that may be important for C4 oxidation is a conserved surface serine/threonine residue that is immediately before the second histidine ligand to the copper active site. The crystal structures of NcPMO2, NcPMO3, and TaGH61A show that the hydroxyl group of this serine residue is positioned on the active-site surface and forms an H-bond with the second histidine ligand. This serine/threonine residue could influence orientation of the copper center on the substrate. Alternatively, it could modulate the electronic properties of the copper center, thereby altering oxygen activation. Hydrogen-atom abstraction (HAA) from C4 is likely a more difficult thermodynamic task than HAA from C1; the C–H bond dissociation energy is estimated to be several kilocalories per mole higher for C4 (alcohol) than for C1 (aldehyde) (66). In addition, type 2 PMOs have a highly conserved lysine residue immediately following the second histidine ligand and an extra unique sequence of 9 to 14 amino acids a few residues prior to the histidine. The crystal structure of NcPMO2 shows that this extra sequence forms a short helix on the active-site surface; it remains to be determined whether this helix could be analogous to the L2 loop in PMO3 enzymes, interacting with cellulose in such a way as to poise the copper center for C4 oxidation. Regioselectivity preferences have been comprehensively characterized in the family of cellulose-active fungal PMOs (65), and it was originally thought that the ability to oxidize both nonreducing and reducing ends of glucose was unique to this family—only reducing-end oxidation had been observed in other families. This view changed in 2014, when a study showed that some bacterial PMOs can also oxidize at C4 in cellulosic substrates (47), indicating that regioselectivity is a deeply conserved feature of oxidative cellulose degradation. A comparative study between a pair of cellulose-active PMOs from Streptomyces coelicolor, CelS2 (which oxidizes C1) and ScPMO10B (which oxidizes cellulose C1/C4 and chitin C1) showed little difference in their global structures other than slight variations in the region analogous to the L2 loop in fungal PMOs. This loop in bacterial PMO structures forms a much larger protuberance that contains significant α-helical structure and contributes up to 50% of the active-site binding surface (47); small differences in its structure may be significant. Additionally, differences lie in the active site: The CelS2 activesite structure is similar to that of chitin-active bacterial PMOs, whereas the active-site structure of ScPMO10B bears a closer resemblance to that of cellulose-active fungal PMOs. ScPMO10B contains the axial tyrosine ligand and H-bonding network (H–X2 –Q–X–Y motif ), features that appear to be necessary for C4 oxidation (see the section titled Tertiary Structure and the Active Site, above). Although conserved residues and substructures are important for substrate binding and regioselectivity of oxidation, other factors may be involved. For example, varied posttranslational glycosylations and active-site groove depths may play a role in substrate binding and O2 positioning, respectively (40). Glycosylation modifications in CBM1 have been demonstrated to promote cellulose binding via molecular dynamics simulations (67).

NcPMO3: a PMO3 found in Neurospora crassa (NCU07898); crystal structure available (PDB 4EIS) NcPMO2: a PMO2 found in Neurospora crassa (NCU01050); crystal structure available (PDB 4EIR) CelS2: a bacterial PMO (from Streptomyces coelicolor) that oxidizes C1 in cellulose; crystal structure available (PDB 4OY7) ScPMO10B: a bacterial PMO (from Streptomyces coelicolor) that oxidizes C1 and C4 in cellulose and C1 in chitin; crystal structure available (PDB 4OY6)

THE POLYSACCHARIDE MONOOXYGENASE REACTION Copper-Dependent Monooxygenase Activity The oxygen- and reductant-dependent oxidative activity of PMOs was first reported by VaajeKolstad et al. (32) for CBP21 from the chitinolytic bacterium Serratia marcescens. This oxidative activity was initially attributed to a redox inert divalent metal ion such as Mg(II) or Zn(II) in the active site. Prior to this study, others had reported that a divalent metal ion is required for some GH61 proteins to stimulate cellulose degradation; however, the metal was not ascribed to www.annualreviews.org • Cellulose Degradation


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any oxidative activity (29). Despite the discrepancy in the assignment of the native metal cofactor of PMOs, the study by Vaaje-Kolstad et al. (32) provided the first clear indication toward understanding the chemistry of this superfamily. In later studies on fungal cellulose-active PMOs, multiple groups reconstituted apo-PMOs with various metal ions and showed that copper is the native metal cofactor of PMOs. Copper was subsequently found in chitin- and cellulose-active bacterial PMOs (14, 41, 44, 47, 56, 68), as well as in chitin-active fungal PMOs (46). The site of oxidation was initially a challenge to resolve due to the difficulty of analyzing the reaction products. Vaaje-Kolstad et al. (32) used isotope-labeling experiments with either H2 18 O or 18 O2 to show that the C1 position of the N-acetyl-glucosamine unit in chitin is oxidized to the corresponding carboxylic acid with one oxygen atom incorporated from O2 . Oxygenation of the C1 position of cellulose has subsequently been found in many cellulose-active bacterial and fungal PMOs (33, 34, 39, 41, 47, 65, 68–71); some of these have been found to oxidize C4 as well, or to oxidize exclusively C4. On the basis of these studies, investigators proposed a possible mechanism for glycosidic bond cleavage (Figure 3a). In this proposed mechanism, PMOs hydroxylate either the C1 or C4 position of the glycosidic bond in cellulose to form unstable hemiketal intermediates that undergo elimination, resulting in either aldonolactones (C1 oxidation) or 4-ketoaldoses (C4 oxidation). Aldonolactones subsequently undergo hydrolysis, which can be spontaneous or enzyme-catalyzed (72), to form aldonic acid products. 4-Ketoaldoses are the products of C4 oxidation, as reported by Beeson et al. (39) via a combination of mass spectrometry and chemical derivatization and later confirmed in independent research by Isaksen et al. (73) using two-dimensional NMR. Two other studies proposed oxidation at C6 by PMOs solely on the basis of mass spectrometry analysis (33, 69). Vu et al. (65) examined the regioselectivity of phylogenetically diverse fungal cellulose-active PMOs and did not detect C6 oxidation. All these data suggest that C6 oxidation is not taking place.

Single Turnover Mechanism The proposed PMO mechanism has been based primarily on studies of similar oxygen-activating copper enzymes and relevant model complexes, as well as some recent computational research. Phillips et al. (34) and Beeson et al. (39) proposed that PMOs activate O2 to hydroxylate cellulose, with subsequent breakdown of the hemiketal that is generated. Li et al. (40) observed X-ray crystallographic electron density at 3.0 and 3.4 A˚ from the copper center of a type 2 PMO and a type 3 PMO from N. crassa that were assigned as superoxide and hydrogen peroxide, respectively. In the subunit of the type 3 PMO where the hydrogen peroxide was not observed, a nearby tyrosine was hydroxylated. It is likely that the copper center in these PMOs was photoreduced upon exposure to the X-ray beam and that it subsequently reduced or activated O2 . However, the distances from the O–O moieties to the copper center are too long to form bonding interactions, −−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−→ Figure 3 The polysaccharide monooxygenase (PMO) reaction. (a) Regioselective hydroxylation of cellulose by PMOs. (b) Proposed single turnover mechanisms of PMOs. (c–f ) Representative superoxo species. (c) Crystal structure of S [Cu(II)–superoxo intermediate] in peptidylglycine α-hydroxylating monooxygenase (PHM), in which the copper center is coordinated by two histidine residues and a methionine residue [Protein Data Bank (PDB) entry 1SDW]. (d ) Five-coordinate square pyramidal S model for fungal PMOs (86). (e) A putative superoxide moiety is observed 3.0 A˚ from the copper center of the Neurospora crassa PMO NCU01050 (PDB 4EIR); ( f ) Four-coordinate planar S model for fungal PMOs with projected hydrogen-bonding interactions. The filled circles represent oxygen atoms. Abbreviations: HO–R, Cu(III)–OH; HOx, Cu(II)–OH; HP, Cu(II)–hydroperoxo; O–R, Cu(III)–oxyl radical; Ox, Cu(II)–oxyl radical. 30.12

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OH RO

OH

H

H

OH

OH


2 + 2e– + 2H+ – H2

OH RO

b

H

+

RH

HO

PMO–Cu(II)–H

HOx

Ox

O–R

H2

H+,e

Ox

PMO–Cu(I) PM MO –C Cu(I) 2

(Ox1)

R•

HP

H+

e

H+

H 2

PMO–Cu(II)––H

S

PMO–Cu(II)––H H2

Ox

(S2)

RH

•– H+,e PMO–Cu(II)––

(Ox2)

PMO–Cu(II)–•

2H+,e

e

PMO–Cu(II)–•

R• RH

PMO–Cu(II)–•

RH R•

H2 PMO–Cu(II)

RH Tyr170

Tyr170

OH

R–

Tyr170

R•

HO

OH H

RH

Tyr170

+

OR O

H+

HO

PMO–Cu(II)–•

HO

R•

PMO–Cu(III)–H

HO–R

PMO–Cu(III)–•

4-Ketoaldose OH

OH

OH

HO

OH

O

HO

PMO–Cu(II)–H R•

O H 4

RO

O

OH H

OR

OH

OR

O

HO

OH H 1

OH

H

OH

HO O

HO

PMO2 2 + 2e– + 2H+ – H2

OH

HO

O

O

RO

PMO3

O

OH Aldonolactone

PMO1 or PMO3* 2 + 2e– + 2H+ – H2

OR

HO

HO

O

OH

HO

O

RO

OR

O

HO

OH

OH

HO

O

H+,e

HP

(S1)

R•

RH

P PMO–Cu(II)–––

c

d

e H2N

2.1 Å

O

f HN

HO

H

H H N NH H O N O H Cu(II) N N H N

N

N

O

NH

3.0 Å

N N

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and whether the observed oxygen species are relevant to catalysis, or an artifact of photoreduction in the crystal, remains unclear. Oxygen activation by PMOs is likely to start with the binding of O2 to Cu(I)–PMO forming a Cu(II)–superoxo intermediate (S). From S there are several possible reaction paths that could lead to substrate hydroxylation (Figure 3b), all of which require the well-timed delivery of protons and electrons. Paths S1 and S2 rely on HAA by S, whereas paths Ox1 and Ox2 catalyze HAA by a Cu(III)–oxyl radical (O–R) species. Path S1, which Phillips et al. (34) and Beeson et al. (39) proposed for PMOs, is analogous to the generally accepted mechanism of peptidylglycine α-hydroxylating monooxygenase (PHM) and dopamine β-monooxygenase (DβM) (74, 75). In this path, serine abstracts a hydrogen atom from the substrate to form Cu(II)–hydroperoxo (HP) and a substrate radical (74). HP is then further reduced and the O–O bond cleaved to form a Cu(II)-oxyl radical (Ox). Radical rebound then occurs between the substrate radical and Ox to form the product anion, and the enzyme returns to the resting oxidized state. Both S and HP have been observed and characterized in enzymes (Figure 3c) and model complexes, whereas Ox has been implicated in rigorous kinetic studies in model complexes (74, 76–81). In model complexes, S exhibits predominantly Cu(II)– superoxo character but can also vary between Cu(II)–superoxo and Cu(III)–peroxo species (82). An alternative to path S1 is path S2, in which the hydroxyl radical is transferred from HP to the substrate radical to form the hydroxylated product and Ox, as proposed by Chen & Solomon (83) for PHM. In paths Ox1 and Ox2, a proton and an exogenous electron are transferred to S to form HP. A Cu(II)–peroxo species could also be formed prior to HP. In the next step of path Ox1, another proton and electron are transferred to HP to form Ox, which then abstracts a hydrogen atom from the substrate to form a Cu(II)–OH (HO) species. In path Ox2, HP undergoes heterolytic O–O bond cleavage, as found for P450s (84), to form an O–R species. O–R is supposedly highly oxidizing and could be stabilized by the active-site tyrosine residue (Tyr175), which may result in a Cu(II)– oxyl–ligand cation radical analogous to Compound I in P450s (84). Hydrogen-atom transfer from the substrate to O–R leads to the formation of a Cu(III)–OH (HO–R) species that has recently been characterized in a model complex (85). The cation radical may not form in chitin-active bacterial PMOs and the subfamily of cellulose-active bacterial PMOs that have a phenylalanine residue in the place of Tyr175 (see the section titled Polysaccharide Monooxygenase Structure, above). In addition, the nonmethylated N-terminal histidine ligand in bacterial PMOs might not be able to stabilize high-valent copper species as efficiently as the methylated residue in fungal PMOs. Thus, path Ox2 is less likely in bacterial PMOs without the active-site tyrosine residue. Moreover, C4 oxidation, which has been observed only in PMOs with the active-site tyrosine (see the section titled Polysaccharide Monooxygenase Structure, above), might follow path Ox2. Paths Ox1 and S2 were investigated in silico for a PMO from the fungus T. aurantiacus (86). The energy barrier for HAA from the cellulose substrate by S was calculated to be 34.9 kcal/mol. This barrier is significantly higher than the calculated overall 18.8 kcal/mol barrier for the oxyl mechanism (path Ox1). In this computational analysis, path Ox1 is favored over paths S1 and S2. This high energy barrier was unexpected because various experimental results suggested that the S in other metalloenzymes and model complexes is capable of HAA (74, 75, 87–89) from C–H bonds with comparable strengths as those in cellulose. The barrier also differs from the calculation by Chen & Solomon (83) for PHM, in which HAA by S is slightly thermodynamically favorable (G = −2.4 kcal/mol). This discrepancy may result from the novel features of the PMO active site or from the assumptions used in the calculations. The computational study described above proposed a square pyramidal geometry for S, in which the superoxide moiety occupied the axial position trans to Tyr175 while the equatorial


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water remained the same throughout the reaction (Figure 3d ) (86). This model is similar to the putative S structure of NcPMO2 (Figure 3e) (40). In this model, the reactive oxygen species in either S or the resulting Ox might be pointed toward the substrate surface. However, the axial superoxo ligand might not be stably bound due to the Jahn–Teller effect as observed for S in the crystal of NcPMO2. The model is also inconsistent with the fact that the water ligands dissociate from elongated six-coordinate Cu(II) to form a T-shaped three-coordinate Cu(I) center upon reduction of PMOs, as shown by X-ray crystallography for bacterial PMOs and X-ray absorption spectroscopy for fungal PMOs (44, 45, 90). In a separate study, experimentally calibrated density functional theory calculations were used to show that the T-shaped Cu(I) center of fungal PMOs facilitates the thermodynamically difficult one-electron reduction of O2 , resulting in a four-coordinate planar S (Figure 3f ) with very little reorganization energy (90). This calculation is consistent with the rapid inner sphere reoxidation of Cu(I) by bound O2 , and the optimized structure of S was calibrated using relevant model complexes (see the following paragraphs). Four-coordinate planar configuration is a favored geometry of Cu(II) species. The conserved active-site H-bonding network in both cellulose-active fungal and bacterial PMOs (see the section titled Polysaccharide Monooxygenase Structure, above) may play an important role in the activation of oxygen, given that two protons are required for every oxygen molecule utilized. H-bonding networks have been found in the active sites of other metal-dependent hydroxylases, such as the P450s (84, 90). Rigorous studies showed that the H-bonding network of the P450s stabilizes the S and precisely directs proton transfer to the terminal oxygen atom of the end-on dioxygen moiety in the HP intermediate in order to cleave the O–O bond and generate high-valent iron–oxo species capable of C–H bond activation (84, 90). In the four-coordinate planar S model, the conserved catalytically important active-site residues H164 and Q173 may form H bonds with the superoxide ligand (Figure 3d ) and facilitate activation. In the five-coordinate square pyramidal S model (Figure 3b) (86), these residues are not involved in the activation of oxygen. The differences in the active sites between fungal PMOs and bacterial PMOs suggest that these PMOs could use different mechanisms to activate oxygen. A recent study showed that a PMO from N. crassa (NCU02916) uses soluble cellodextrins as a substrate (73), a finding that will enable steady-state and transient kinetic studies. We anticipate that extensive kinetic studies, characterization of reaction intermediates with various spectroscopic and structural techniques, and computational studies based on NCU02916 will take place, which may help unravel the mechanism of cellulose-active fungal PMOs and provide mechanistic information about the PMO superfamily.

DIVERSITY AND BIOLOGICAL FUNCTIONS OF POLYSACCHARIDE MONOOXYGENASES Within the kingdom Fungi, genes encoding cellulose-active PMOs are widely found in the genomes of filamentous basidiomycetes and ascomycetes (91). So far, cellulose-active PMOs have not been reported in the basal fungal lineages and the ascomycetous yeasts. This could be a result of limited sampling of these organisms, in the case of the basal fungi, or of gene loss due to lifestyle changes, in the case of the yeasts. It is likely that the last common ancestor of basidiomycetes and ascomycetes, estimated to have split more than 450 million years ago (92), possessed multiple cellulose-active PMOs. The presence of genes encoding cellulose-active PMOs in basidiomycetes and ascomycetes is strongly correlated with the presence of genes encoding hydrolytic enzymes. In an analysis of the genomes of 31 fungal species, more genes encoding cellulose-active PMOs were www.annualreviews.org • Cellulose Degradation


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identified than the sum of all glycoside hydrolases in families 5, 6, and 7 (91). The authors of this review are unaware of any cellulolytic fungi whose genomes lack putative cellulose-active PMOs, with the exception of the ruminant obligate anaerobic fungus Orpinomyces (93). As an obligate anaerobe, Orpinomyces lives in an environment devoid of molecular oxygen, so cellulose-active PMOs, which require oxygen, would not be active. Taken together, these observations suggest that cellulose-active PMOs may be as integral to effective plant cell wall degradation in fungi as the extensively studied hydrolytic enzymes. Researchers have attempted to understand why cellulose-active PMOs are highly abundant in fungi (65). Type 1 and type 2 PMOs generate unmodified nonreducing and reducing ends, respectively, whereas type 3 PMOs generate both unmodified ends (see the section titled Polysaccharide Monooxygenase Structure, above) (39, 65, 73). Among the 500 fungal PMOs we identified, type 1 and type 3 PMOs are similar in abundance (76% in total), type 2 PMOs account for ∼12%, and the remaining ∼12% of PMOs have not been classified. Analogous studies were also performed for bacterial PMOs (47). Although cellulose-active and chitin-active bacterial PMOs share closely related active-site and tertiary structures, they have very little primary sequence homology. The cellulose-active bacterial PMO clade is further divided into type 1 PMO and type 3 PMO clades. It is unclear whether bacterial and fungal PMOs evolved from a common ancestral PMO, or whether their occurrence in such diverse species is a result of horizontal gene transfer or convergent evolution. Cellulose-active PMOs are tightly regulated at the transcriptional level, similar to hydrolytic enzymes involved in plant cell wall degradation (25, 26, 94–96). The genomes of most highly cellulolytic fungi contain more than 10 genes encoding cellulose-active PMOs; however, generally only a subset of these PMOs are strongly expressed or detected in proteomic analyses of secretomes. For instance, in N. crassa, expression of 11 of 14 cellulose-active PMOs was induced during growth on Miscanthus or pure cellulose, but only four PMOs were reliably detected by proteomic analysis of the secretome (25). Of the four identified PMOs, one was a type 1 PMO, two were type 2 PMOs, and one was a type 3 PMO. The same trend was observed in the secretomes of the related thermophilic fungi Myceliophthora thermophila and Thielavia terrestris, in which a few of the cellulose-active PMOs in the organism’s genome were detected in its secretome, but representatives of both reducing-end and nonreducing-end active PMOs were simultaneously present (94). Biochemical activity confirming the presence of type 2 PMOs in basidomycetes has not yet been reported, but basidiomycete genes predicted to encode these enzymes on the basis of sequence homology have been identified as predicted secreted proteins. In the secretomes of the white-rot basidiomycetes Phanerochaete chrysosporium and Phanerochaete carnosa, PMOs of all three subtypes were simultaneously identified during growth on cellulosic substrates (96, 97). Recently, investigators discovered that the well-studied cellulolytic actinomycete bacterium Thermobifida fusca also simultaneously secretes PMOs that act on both the reducing and nonreducing ends during growth on cellulose (47, 98, 99). This large degree of conservation across ascomycetes, basidiomycetes, and actinomycetes for the simultaneous production of functionally different PMOs implies that both reducing-end and nonreducing-end activities are necessary for efficient depolymerization of cellulose. The discovery of the function of PMOs in N. crassa was enabled by the observation that a deletion of the major CDH in N. crassa strongly decreased the cellulase-enhancing activity of PMOs in the secretome (34). CDH is important for PMO activity because of the sequential transfer of electrons, via the CDH–heme domain, to the copper center on the surface of the PMO used to activate oxygen for C–H bond insertion. The absence of CDH in the industrial cellulase producer Trichoderma reesei, which contains a small number of PMOs, has fueled speculation that CDH may not be critical for PMO function. Several hundred fungal genomes have now been sequenced and


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e Fe2+

O2 Cu

Cu

Cu


Cu

Endoglucanase

CBH1

CBH2

Cellobiose (reducing end on the right)

Type 1 PMO

Type 2 PMO

Type 3 PMO

CDH

Indicates oxidized ends

Figure 4 A model for fungal cellulose degradation. Exoglucanases act on chain ends to generate soluble sugars, which are taken up by the fungus. Endoglucanases and polysaccharide monooxygenases (PMOs) increase the accessibility of cellulose to exoglucanases by creating new chain ends from internal regions of amorphous and crystalline cellulose chains, respectively. Extracellular reductases, such as cellobiose dehydrogenase (CDH), colocalize on the cellulose surface and provide the reducing equivalents necessary for PMO-dependent oxygen activation. Abbreviation: CBH, cellobiohydrolase.

are publicly available online (100). The picture that emerges from analysis of these ∼320 fungal genomes is that Trichoderma is the exception and not the norm. In fungi that contain multiple PMOs and the canonical cellobiohydrolases, CDH is overwhelmingly conserved. Among fungi that contain at least three genes encoding PMOs in their genomes, more than 98% also contain a CDH. In fungi that contain zero PMOs, CDH is always absent. In addition to Trichoderma, there are some brown-rot fungi, such as Postia placenta, that contain a small number of PMOs, no cellobiohydrolases, and no CDH (27). Also, the thermophilic eurotiomycete Thermoascus aurianticus produces an active PMO, but unlike most cellulolytic fungi, it secretes no enzymes with cellulose-binding modules, cellobiohydrolase II, or CDH (101). T. aurianticus grows quite poorly on pure, crystalline cellulose, relative to Neurospora or Myceliophthora. Outside of Trichoderma, how fungi similar to T. aurantiacus and P. placenta depolymerize cellulose is still poorly understood at a mechanistic level. Further studies of these cellulolytic systems may shed light on this topic.

PERSPECTIVES AND APPLICATIONS The discovery of PMOs has fundamentally changed the long-accepted paradigm of fungal cellulose utilization (Figure 4). Much has been learned in the past few years, but key questions remain largely unaddressed. For decades, uncontrolled Fenton chemistry was speculated to be the principal oxidative mechanism employed by fungi for cellulose degradation (37, 38). With the discovery of PMOs, the Fenton chemistry model has been largely refuted, at least with respect to CDH. We initially proposed that oxygen insertion adjacent to the glycosidic bond, catalyzed by PMOs, would be destabilizing and lead to a potentially spontaneous breakdown of a labile intermediate to result in polysaccharide cleavage and production of a carbonyl product (34, 39). The term lytic PMO (LPMO) was later introduced by Eijsink and colleagues (14) because oxygen insertion leads to cleavage of the glycosidic bond. It is now generally accepted that these enzymes act as monooxygenases, but no evidence showing that glycosidic bond cleavage is catalyzed by the PMO has been reported. Until the bond-cleavage step is shown to be enzyme dependent, we feel it is premature to include lytic in the enzyme name. Kinetic studies, which would address these topics experimentally, are still sorely lacking, predominantly because of the analytical challenges www.annualreviews.org • Cellulose Degradation


Lytic polysaccharide monooxygenase (LPMO): alternate name for PMO

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of working with insoluble substrates and unstable products and intermediates. The recent identification of an N. crassa PMO that is active on soluble oligosaccharides should enable the detailed kinetic analyses that are necessary to address some of the remaining mechanistic questions (73). Another key outstanding question in the PMO field is related to the N-terminal histidine methylation. Almost nothing is known about this modification, except that it appears to be absolutely conserved in fungi. Methylation of the histidine ligand changes the electronic properties of the copper active site, and as elaborated on above (see the section titled Polysaccharide Monooxygenase Structure), may have important mechanistic implications for oxygen activation and stabilization of reactive intermediates. The methyltransferase responsible for this posttranslational modification is unknown, but it likely resides in the endoplasmic reticulum. The functional genomic tools available for the genetically tractable fungus N. crassa make it an ideal system to use for the identification of the PMO methyltransferase (102). Similarly, the metabolic pathway responsible for utilizing the C4 oxidized products (4-ketoaldoses) remains to be elucidated, but it is very likely to be present in lignocellulolytic fungi. Additional genetic experiments could also be pursued in N. crassa, in which strains with multiple deletions of PMOs are generated and their growth on crystalline cellulose evaluated. The biological importance of each class of cellulose-active PMOs could then be directly addressed, possibly explaining why so many fungi produce more than 10 PMOs. Perspectives about PMOs are expanding as new biological functions and activities are being discovered. Studies have shown that some bacterial virulence factors, such as Vibrio cholerae colonization factor GbpA and Listeria monocytogenes LMO2467, are PMOs (45, 103–105). GbpA and LMO2467 contain cell-adhesion domains that are also found in putative multimodular PMOs in other pathogenic bacteria such as Bacillus anthracis and Yersinia pestis. Many fungal chitin-active PMOs and cellulose-active PMOs contain extra C-terminal domains with unknown functions, suggesting that these PMOs also have biological functions beyond biomass destruction alone. An N. crassa PMO, NCU02916, which can oxidize both cellulose and soluble cellodextrins (65, 73), has also been shown to oxidize hemicelluloses that contain β(1→4) linkages between glucose or substituted glucose units (106). Moreover, cleavage of polysaccharides with α-glycosidic linkages has been observed in a starch-active PMO family (107, 108). Finally, putative new PMOs with unknown substrates, but conserved PMO motifs, have also been identified (107).


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Polysaccharide Monooxygenases in Industrial Biofuel Production As with most new catalyst discoveries, there will be challenges in realizing the full potential of these enzymes in a commercial application, but initial progress is encouraging. Harris et al. (29) showed data supporting the assertion that the addition of PMOs to commercial cellulase cocktails can reduce the required enzyme dose for conversion of pretreated corn stover as much as twofold. Despite these initially promising results, there may be trade-offs in using oxygen-dependent catalysts for biomass conversion (109). Investigators have devoted much effort to developing a simultaneous saccharification and fermentation (SSF) process for production of cellulosic ethanol. The SSF approach is frequently used in the production of corn ethanol (110). Under the anaerobic or microaerobic conditions necessary for fermentation, oxygen may not be available for the oxidative cleavage reactions catalyzed by PMOs (111). Even if separate hydrolysis and fermentation approaches are ultimately used in the industry, a fraction of the carbohydrates will be oxidized extracellularly by the PMOs and thus may not be available for conventional sugar fermentation. Balancing the energy losses in the feedstock due to oxidation by PMOs with the cost savings from lower enzyme doses or reduced processing time will have to be carefully managed to maximize economic returns. 30.18

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The relatively recent discovery of PMOs is dramatically changing our view of microbial cellulose degradation. Exciting opportunities exist in further elucidating the biological function, fundamental mechanism, and commercial application of these enzymes for enhanced production of renewable chemicals and fuels.

DISCLOSURE STATEMENT


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BI84CH30-Marletta

30.24

Beeson et al.


Single-domain flavoenzymes trigger lytic polysaccharide monooxygenases for oxidative degradation of cellulose.

Lytic polysaccharide monooxygenases disrupt the cellulose fibers structure.

The molecular basis of polysaccharide cleavage by lytic polysaccharide monooxygenases.

A structural overview of GH61 proteins - fungal cellulose degrading polysaccharide monooxygenases.

Structural and functional characterization of a conserved pair of bacterial cellulose-oxidizing lytic polysaccharide monooxygenases.

Fungal cellulose degradation by oxidative enzymes: from dysfunctional GH61 family to powerful lytic polysaccharide monooxygenase family.

Cellulose surface degradation by a lytic polysaccharide monooxygenase and its effect on cellulase hydrolytic efficiency.

Cellulose degradation by oxidative enzymes.

Determinants of regioselective hydroxylation in the fungal polysaccharide monooxygenases.

Classification of fungal and bacterial lytic polysaccharide monooxygenases.

Enhancement of Cellulose Degradation by Cattle Saliva.

Injectable polysaccharide hydrogels reinforced with cellulose nanocrystals: morphology, rheology, degradation, and cytotoxicity.

Evolution of substrate specificity in bacterial AA10 lytic polysaccharide monooxygenases.

Molecular biology of cellulose degradation.

Enzymatic degradation of (ligno)cellulose.

Activation of enzymatic chitin degradation by a lytic polysaccharide monooxygenase.

Evaluating Models of Cellulose Degradation by Fibrobacter succinogenes S85.

Promotion of crystalline cellulose degradation by expansins from Oryza sativa.

Cellulose degradation by Clostridium thermocellum: from manure to molecular biology.

Regulated methionine oxidation by monooxygenases.

Lytic polysaccharide monooxygenases and other oxidative enzymes are abundantly secreted by Aspergillus nidulans grown on different starches.

Spectroscopic and computational insight into the activation of O2 by the mononuclear Cu center in polysaccharide monooxygenases.

Antifouling coating of cellulose acetate thin films with polysaccharide multilayers.

The Contribution of Non-catalytic Carbohydrate Binding Modules to the Activity of Lytic Polysaccharide Monooxygenases.