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Lytic polysaccharide monooxygenases (LPMOs) in enzymatic processing of lignocellulosic biomass Piotr Chylenski, Bastien Bissaro, Morten Sørlie, Åsmund K. Røhr, Aniko Varnai, Svein J. Horn, and Vincent G.H. Eijsink ACS Catal., Just Accepted Manuscript • DOI: 10.1021/acscatal.9b00246 • Publication Date (Web): 22 Apr 2019 Downloaded from http://pubs.acs.org on April 22, 2019
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ACS Catalysis
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Lytic Polysaccharide Monooxygenases (LPMOs) in Enzymatic
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Processing of Lignocellulosic Biomass
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Piotr Chylenski, Bastien Bissaro, Morten Sørlie, Åsmund K. Røhr, Anikó Várnai, Svein J. Horn
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& Vincent G.H. Eijsink*
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Norwegian University of Life Sciences (NMBU), Faculty of Chemistry, Biotechnology and Food
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Science, P.O. Box 5003, N-1432 Ås, Norway
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*For
correspondence; E-mail:
[email protected] 11 12 13
Abstract: The discovery of lytic polysaccharide monooxygenases (LPMOs) has revolutionized
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enzymatic processing of polysaccharides, in particular recalcitrant insoluble polysaccharides such
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as cellulose. These monocopper enzymes display intriguing and unprecedented catalytic
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chemistry, which make them highly valuable in industrial bioprocessing, but also generate
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considerable challenges in terms of scientific understanding and optimal implementation. One
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issue of particular interest is the fact that both molecular oxygen and hydrogen peroxide can drive
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LPMO reactions. Here we review recent insights into the catalytic mechanism of LPMOs derived
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from structural, spectroscopic and functional studies. We then turn to the question of how one can
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optimally harness the potential of LPMOs in biomass processing, given the current knowledge of
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their catalytic mechanism. Finally, we review recent, more applied studies that have addressed the
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importance of LPMOs in enzymatic conversion of lignocellulosic biomass, and discuss how the
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impact of these powerful enzymes could be improved.
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Keywords: LPMO, cellulose, copper, biomass, hydrogen peroxide, monooxygenase,
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peroxygenase
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1. Introduction
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The discovery of oxidative cleavage of polysaccharides in 2010 by enzymes today referred to as
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Lytic Polysaccharide Monooxygenases (LPMOs; sometimes referred to as PMOs) shed light on
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two intriguing earlier findings.1 In 1974, Eriksson et al. showed that the degradation of cellulose
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by enzymes present in the supernatants of cultures of cellulose-degrading fungi was more efficient
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if the reactions were conducted under aerobic conditions, leading them to suggest that oxidative
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processes played a role.2 In 2005, Vaaje-Kolstad et al. showed that proteins known as “chitin-
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binding proteins” or family 33 carbohydrate-binding modules (CBM33s) boost the efficiency of
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canonical hydrolytic enzymes involved in chitin degradation, namely chitinases.3 Then, in 2010,
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Vaaje-Kolstad showed that these chitin-binding proteins are enzymes that use reducing power and
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O2 to oxidatively cleave glycosidic bonds in chitin.1 In the meantime, it had been shown that
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proteins at the time classified as family 61 glycoside hydrolases (GH61) are structurally similar to
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CBM33s4 and that they boost the activity of cellulases.5,6 Indeed, in 2011 oxidative cleavage of
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cellulose by CBM33s7 and GH61s8-11 was demonstrated.
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Following the initial discovery of chitin- and cellulose-active LPMOs, various LPMOs
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were found to be active on other substrates, namely soluble cello-oligosaccharides,12 various
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hemicelluloses13-15 and starch.16,17 The discovery of the catalytic function of CBM33s and GH61s
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led to their re-classification in the Carbohydrate Active enZymes (CAZy) database as auxiliary
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activities, belonging to family 10 (AA10) and 9 (AA9), respectively.18 The CAZy database, where
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proteins are classified according to sequence similarity,19 currently contains five additional LPMO
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families: AA11, AA13, AA14, AA15 and AA16. It is worth noting that LPMOs belonging to
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different families are not necessarily very different in terms of function. For example, chitin-active
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LPMOs appear in families AA10, AA11 and AA15 and while sequence similarities generally are
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low (hence multiple families in CAZy), most of these chitin-active LPMOs can be identified by a
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single hidden markov model (PF03067).20
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LPMOs are monocopper enzymes (Figure 1) that bind copper in a characteristic “histidine
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brace”,9, 21 similar to that of the particulate methane monooxygenase (pMMO).22,23 Initially, there
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was some confusion about the nature of the catalytic metal,1,6 which is likely due to the ubiquitous
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presence of minor amounts of copper combined with the high affinity of LPMOs for this metal
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ion.9,24 The original reaction scheme for the LPMO reaction entails that each catalytic cycle
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requires two externally delivered electrons and one molecule of O2 (Figure 2A) and several 3 ACS Paragon Plus Environment
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possible catalytic mechanisms have been proposed (see below).8,25,26 In Nature, electrons may be
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delivered by a wide variety of small molecule reductants as well as by flavoproteins such as
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cellobiose dehydrogenase (see below).1,11,27-29
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Importantly, in 2016, Bissaro et al. showed that LPMOs can use H2O2 as co-substrate and
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that controlled supply of H2O2 leads to reaction rates that are orders of magnitude higher than rates
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commonly observed in O2-driven reactions.30 In the H2O2 reaction scheme (Figure 2B), a priming
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reduction of the LPMO, from the Cu(II) to the Cu(I) form is followed by multiple catalytic cycles
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using H2O2 as co-substrate.31 In this reaction scheme, the consumption of reductant is sub-
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stoichiometric relative to the amount of generated products, whereas the O2-driven reaction
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requires stoichiometric amounts of reductant. Bissaro et al. have suggested that O2-driven reactions
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may not occur at all, and that the (low) rates observed for O2-driven reactions reflect the rate of
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H2O2 formation in the reaction mixture,31,32 but this remains controversial.33 Nonetheless, the claim
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that LPMOs can be made to run much faster than previously observed by supply of H2O2 has been
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confirmed by multiple laboratories.32,34-39 Clearly, next to scientific implications, these recent
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findings may have implications for the application of LPMOs in biomass processing, as discussed
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below.
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LPMOs are able to act on the surfaces of insoluble substrates, thus improving the
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accessibility for canonical hydrolases (e.g. chitinases and cellulases) in the most recalcitrant parts
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of the substrate that otherwise would have been degraded much more slowly or not be degraded at
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all.40-43 While the boosting effect of LPMOs on the activity of hydrolytic enzyme cocktails varies
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in magnitude, these enzymes have already found their way to industrial application. LPMOs are
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part of modern commercial cellulose cocktails for biomass processing44,45 and it is well
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documented that they make a considerable contribution to the efficiency of these cocktails.37,46-50
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Since LPMOs are of major scientific and industrial interest it is worthwhile taking a closer
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look at how they work and how they best can be applied.
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2. LPMO Catalysis
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The exact nature of the catalytic mechanism of LPMOs remains a subject of some controversy,
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especially in the light of the recent discovery of the peroxygenase activity of LPMOs. An in-depth
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discussion of structural features and putative mechanistic routes of LPMOs is beyond the scope of 4 ACS Paragon Plus Environment
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the present review, and these aspects have been extensively covered in previous
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reviews.25,26,32,40,51-56 However, it is necessary to mention the key structural and mechanistic
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characteristics pertaining to LPMO catalysis, particularly in the light of recent mechanistic
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discoveries and their implications for biomass processing.
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2.1 Uniqueness of the LPMO structure
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The overall structure of LPMOs makes these enzymes uniquely suited for performing catalysis on
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polysaccharides that are embedded in the crystalline lattice and otherwise unavailable for attack
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by canonical glycoside hydrolases. Despite low sequence identity between LPMOs, both within
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and in between families, all LPMOs share the key feature of having a relatively flat, solvent-
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exposed substrate-binding surface that includes two conserved histidines that coordinate the
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catalytically crucial single copper atom in a structural arrangement known as a “histidine brace”
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(Figure 1C and D).9,21,54,57 A similar T-shaped histidine brace coordinating a single copper atom
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also occurs in the copper transport protein CopC58 and in particulate methane monooxygenases
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(pMMOs, Figure 1E).23 The surroundings of the histidines in the catalytic centers of LPMOs vary
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both among and within LPMO families.53,54 In LPMOs from AA families 9, 11 and 13 the
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relatively buried proximal axial copper coordination position (Figure 3) is occupied by tyrosine,
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whereas in most AA10 LPMOs phenylalanine occupies this position, although tyrosine also
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occurs.59 In fungal LPMOs, the N-terminal histidine is post-translationally methylated at Nε2
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(Figure 1C), and current data suggests that this modification has little effect on catalytic properties
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but may help protecting the enzyme from auto-catalytic oxidative damage.60 The core of the LPMO
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structure has an immunoglobulin- or fibronectin type III-like structure consisting of a distorted β-
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sandwich fold of typically 8–10 β-strands, connected by several helices and loops that generate
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structural diversity among LPMOs and varying topologies of the substrate binding surface and the
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catalytic center.25,53,54,61,62
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Figure 1. Structural features of a fungal and a bacterial LPMO and of pMMO. Panels A and
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B show cartoon representations of TaAA9A (PDB ID code 2YET), whereas panel C-E shows the
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copper binding active sites of TaAA9A, ScAA10C (PDB ID code 4OY7), and Methylocystis sp.
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pMMO (PDB ID code 3RFR). In panels A and B, the side chains of residues that are part of the
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(putative) substrate-binding surface of TaAA9A are shown with cyan carbons, whereas residues
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that are also shown in panel C have green carbons. Copper atoms are shown as orange spheres.
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Close interactions with the copper ion are shown as sticks, with distances in Å, whereas dashed
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magenta lines indicate other relevant (potential) contacts.
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Figure 2. Reaction scheme for LPMO catalytic scenarios. Panel A shows the scheme of a
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monooxygenase reaction originally proposed by Vaaje-Kolstad et al. in 2010.1 Panel B shows the
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scheme of a peroxygenase reaction proposed by Bissaro et al.31 Note that the two scenarios differ
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in terms of the consumption of reductant, which is stoichiometric, relative to the amount of
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products formed, in scheme A and sub-stoichiometric in scheme B.
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LPMOs have varying substrate specificities and vary in their oxidative regioselectivity.
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LPMOs may exclusively oxidize either the C1 or the C4 carbon in the scissile glycosidic bond,
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whereas others produce mixtures of C1- and C4-oxidized products (Figure 4).12,63 Similar
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variations have been found for LPMOs acting on xyloglucan, whereas for the abundantly described
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chitin-active LPMOs only C1-oxidized products have been detected so far. In spite of attempts to
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unravel the determining factors of oxidative regioselectivity59,63,66 and substrate specificity of
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LPMOs,66,67 these factors remain largely unknown. Current data do indicate that productive
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binding of substrate depends on multiple interactions involving a larger part of the substrate-
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binding surface interacting with multiple polysaccharide chains in the substrate.24,39,62 68
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Figure 3. Schematic overview geometries referred to in the text. (A) The typical LPMO
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active site with three nitrogen atoms from the histidine brace coordinating the copper ion. One of
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the axial coordinating positions, the one pointing towards the protein core, also referred to as
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proximal, is usually blocked by a Tyr-OH group or a Phe side chain. Possible interaction sites for
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other molecules at the equatorial (eq) or (distal) axial (ax) position are indicated. (B) Example of
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superoxide that is equatorially bound in a side-on conformation to copper. (C) Example of
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superoxide that is equatorially bound in an end-on conformation to copper.
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Figure 4. Oxidized sugar moieties produced during LPMO-catalyzed degradation of
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cellulose. The primary oxidation products are a lactone or a 4-ketoaldose, which are in equilibrium
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with their hydrated forms, an aldonic acid and a 4-gemdiol, respectively.
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2.2 Catalytic mechanism
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Several possible mechanisms for LPMO catalysis have been proposed and reviewed
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(Figures 2 and 5; see below).25,26,31 The first experimental data shedding light on the key players
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of the reaction was described by Vaaje-Kolstad et al.1 In this study, the chitin-active CBP21
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(SmAA10A) was incubated with -chitin in the presence of an external reductant and molecular
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oxygen (Figure 2A). Analysis of end products of the reaction using mass spectrometry and HPLC
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revealed the formation of aldonic acids with a degree of polymerization (DP) of two and higher.
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Experiments in presence of
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increased mass of 2 Da compared to the standard setup of
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suggested that LPMOs use activated oxygen to perform oxidation of a C1-carbon of an N-
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acetylglucosamine moiety in the chitin chain and that a water molecule takes part in the formation
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of the aldonic acid. Using similar methods, similar conclusions were reached by Beeson et al. for
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a cellulose-active AA9 LPMO.69 Since then, most mechanistic studies have been centered around
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the role of oxygen as co-substrate, and on identification of the key oxidative oxygen species.
18O
16O
2/H2
and
16O
18O
2/H2
both yielded aldonic acids with an 16O
16O.
2/H2
These results clearly
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In 2012, Li et al. described the crystal structure of an LPMO from Neurospora crassa with
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coordination position with an end-on (1) configuration to the copper ion.68 The electron density
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was modeled as a dioxygen species with an unrestrained bond length of 1.16 Å. The authors
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proposed this to be consistent with a superoxide species weakly coordinated to Cu(II), as bond
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lengths for oxygen or superoxide are 1.2-1.3 Å, whereas the bond length in a peroxide would be
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close to 1.49 Å. Since then, the possibility of axial oxygen activation has been explored further by
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Kim et al. (see below),70 but recent work suggests that oxygen activation happens in the equatorial
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plane (Figure 3).57,71-74
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In a 2013 review, Hemsworth et al. suggested that Cu(III)-OH (or possibly Cu-(III)-
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peroxide or “cupryl”) may be the reactive form of oxygen.75 One reason was the isolation of a
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Cu(III)-OH
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diisopropylphenyl)-2,6-pyridinedicarboxamideCuOH]) with a N3 T-shape coordination geometry
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of the copper, akin to the histidine brace.76 The other reason was that the same active species had
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been discussed in relation to C-H bond activation in methane by particulate methane
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monooxygenase (pMMO).77 In relation to this, it is worth noting that it has been suggested that the
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N-terminal amino group can be deprotonated.72 Such deprotonation could possibly facilitate
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formation of a Cu(III) intermediate.
intermediate
of
a
copper
model
compound
([Bu4N][(N,N´-bis(2,6-
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Kim et al. used the structure of an LPMO from Thermoascus aurantiacus (TaAA9A;
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Protein Data Bank, PDB ID code 2YET, Figure 1) to build an active site cluster model and
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performed density functional theory (DFT) calculations to compare two possible reaction paths for
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hydrogen abstraction.70 One tested mechanism employed a η1-superoxo intermediate ([LPMO-
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Cu(II)-OO) which abstracts a substrate hydrogen, while the other alternative entailed formation
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of a copper-oxyl radical (LPMO-Cu(II)-O) that abstracts a hydrogen and subsequently
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hydroxylates the substrate via an oxygen-rebound mechanism (Figure 5). The results predicted
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that oxygen binds end-on (η1) to copper at the axial position, and that a copper-oxyl–mediated,
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oxygen-rebound mechanism is energetically preferred. While axial binding of the oxygen is not
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supported by subsequent studies, the formation of an oxyl intermediate is (Figure 5).
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Figure 5. Schematic summaries of proposed mechanisms for hydrogen atom abstraction by
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an LPMO using O2 (top) or H2O2 (bottom) as co-substrate. Note that alternative scenarios have
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been proposed for both O2- (e.g. refs 25-26, 55) and H2O2-driven (ref 31) catalysis. Also note that,
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while scenarios for H2O2-driven catalysis generally imply that the copper stays reduced in between
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catalytic cycles, this is not the case for all proposed scenarios for O2-driven catalysis, some of
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which entail that the LPMO ends up in the oxidized from at the end of one cycle. The copper-oxyl
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intermediate (LPMO-Cu(II)-O), which is a favoured intermediate state in most modelling studies
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of catalysis, also could be a copper-oxo (LPMO-Cu(III)=O), which bears resemblance to
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Compound I formed by cytochrome P450s, discussed below. See text for more details. These
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schemes are based on the work of Bertini et al.78 and Wang et al.79
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Kjærgård et al. investigated O2 reactivity with the same LPMO, TaAA9A, using EPR and
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stopped-flow spectrophotometry.80 They showed that Cu(I) reoxidation takes place with a
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minimum rate of > 0.15 s-1. Based upon reported redox potentials of ∼275 mV vs. the normal
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hydrogen electrode (NHE)24,61 and the potential of the one-electron reduction of O2 to superoxide
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(−165 mV vs. NHE), the rate of an outer-sphere electron transfer was calculated to be ∼4.5 × 10−4
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s−1, using Marcus theory, which was ∼103 slower than the rate of Cu(I) reoxidation derived from
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the EPR and stopped-flow data. The authors concluded that the single-electron transfer from
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LPMO-Cu(I) to O2 was likely to proceed via an inner-sphere pathway involving rapid formation
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of LPMO-Cu(II)-OO. Further, the coordination distances of the copper N-atom ligands were
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derived from K-edge XANES (X-ray Absorption Near Edge Structure) and EXAFS (Extended X-
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Ray Absorption Fine Structure) experiments, allowing for comparison with the geometry-
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optimized active site models. Addition of an O2 molecule to the optimized structure with a reduced
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copper resulted in a superoxide bound equatorially to the Cu(II) ion in an end-on fashion (Figure
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3). According to the authors, it was not possible to stabilize a side-on bound Cu-O2 structure in the
243
AA9 site.
244
EPR spectroscopy studies by Borisova et al. showed that substrate binding leads to altered
245
g-values and additional superhyperfine coupling patterns, and these authors suggested that the
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observed changes in the spectrum originated from a change in water coordination of the copper
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upon substrate binding.81 This latter hypothesis was strengthened by spectroscopic and
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computational results in a recent study of the chitin-active LPMO SmAA10A.39 Crystal structures
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of LPMO-oligosaccharide complexes described in a seminal paper by Frandsen et al. showed a
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chloride ion, considered to act as a superoxide analogue, bound in the equatorial position (Cu-Cl,
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2.3 Å).57 Frandsen et al.57 observed changes in EPR spectra similar to what was observed by
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Borisova et al.81 and suggested that this was due to the presence of a chloride nucleus leading to a
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significant decrease in the observed gz value of the EPR spectra (from gz = 2.28 in the absence of
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substrate, to 2.27 in the presence of substrate and low chloride concentrations, to 2.23 in the
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presence of substrate and high chloride concentrations). It should be noted, however, that Borisova
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et al. observed the same type of shift at low chloride concentrations.81 Importantly, the structures
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of the complexes described by Frandsen et al. showed that the C6 hydroxymethyl group of the
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glucose unit bound to subsite +1 was close to the copper ion (C-Cu, 3.8 Å) blocking access to the 12 ACS Paragon Plus Environment
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axial copper binding site.57 Frandsen et al. also pointed at the potential interdependence of binding
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of substrate and chloride to the copper ion, which would imply that binding of the oxygen co-
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substrate and a polysaccharide substrate may act in a concerted manner. In this way, production of
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the reactive oxygen species may be controlled by the presence of a substrate.57,82
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In a recent study, Bissaro et al. showed that LPMO reactions can be driven by H2O2.30,31
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By controlling H2O2 supply, stable and fast reaction kinetics were achieved and the LPMOs
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worked in the absence of O2, whereas the reductant was consumed in amounts that were sub-
266
stoichiometric relative to products formed (Figure 2). Experiments with labeled hydrogen peroxide
267
(H218O2) in the presence of a ten-fold molar surplus of O2, showed that the oxygen atom that is
268
introduced into the polysaccharide chain came from H2O2 and not O2 (Figure 6). Introduction of
269
18O
270
and chitin substrates (Figure 6). The authors suggested several possible reaction mechanisms,
271
including mechanisms involving hydroxyl radicals. One plausible mechanism entails the
272
conversion of H2O2 by LPMO-Cu(I) to a water molecule and a Cu(II)-O intermediate, which also
273
can be a Cu(III)=O. From here on, the reaction would proceed via a rebound mechanism, as
274
described above (Figure 5), implying hydrogen abstraction from the substrate, followed by
275
merging of the resulting Cu(II)-associated hydroxide with the substrate radical, leading to
276
hydroxylation of the substrate and regeneration of the Cu(I)-center ready for the next cycle. The
277
most recent studies on the catalytic mechanism of LPMOs have taken the possibility of H2O2-
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driven LPMO reactions into account, albeit to varying extents.
from H218O2 into the final product was shown for both AA9s and AA10s and for both cellulose
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Figure 6. MALDI-ToF analysis of products generated in competition experiments with O2
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and labeled H2O2. (A) MALDI-ToF MS spectrum showing products (DP6 cluster) released upon
283
incubation of Avicel (10 g.L-1) with ScAA10C (0.5 µM) in presence of a low amount of ascorbic
284
acid (AscA; 10 µM), O2 (200-250 M) and varying concentrations of H218O2 (25-100 M). The
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reactions were started by addition of AscA. (B-D) MALDI-ToF MS spectra showing products
286
released from Avicel by ScAA10C (0.5 µM) after 4 min reaction (B), from PASC by PcAA9D (1
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µM) after 15 min reaction (C) and from β-chitin by SmAA10A (0.5 µM) after 60 minutes of 14 ACS Paragon Plus Environment
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reaction (D). All reactions were carried out under normal aerobic conditions, which means that the
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concentration of (non-labeled) 16O2 in solution was in the range of 200-250 µM. Reactions shown
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in panel B were carried out in the presence of 100 µM H216O2 (purple line) or H218O2 (orange line)
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and 1 mM AscA. Reactions shown in panel C were carried out in the presence of 200 µM H216O2
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(purple line) or H218O2 (orange line) and 100 µM AscA. Reactions shown in panel D were carried
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out in the presence of 100 µM H216O2 (purple line) or H218O2 (orange line) and 10 µM AscA. The
294
spectra show that when using H218O2, the characteristic signals for sodium adducts of the aldonic
295
acid form of an oxidized cellohexaose (m/z 1029.7 & 1051.7) or chitohexaose (m/z 1076.0 &
296
1298.0) shift by +2 Da. Abbreviations: DP, degree of polymerization; Nat, native; Lac, oxidized,
297
lactone form; Ald, oxidized, aldonic acid form. Nb. All MS spectra show sodium adducts of the
298
native (Nat), lactone (Lac) and the aldonic acid (Ald) form for the DP6 cluster. Adapted with
299
permission from reference 31. Copyright 2017, Springer Nature.
300 301 302 303
Bertini et al. used DFT calculations on a large active-site model for a fungal AA9 LPMO
304
and with a celloheptaose unit as a substrate mimic to calculate the energies of possible LPMO
305
mechanisms.78 A key finding was that binding of O2 to the T-shaped LPMO-Cu(I) active site
306
resulted in formation of a LPMO-Cu(II)-OO intermediate and a distorted tetrahedral geometry of
307
the Cu atom, a conformation that has not been observed in other computational studies. The
308
presence of the substrate did not change this geometry, but O2 binding was further favored with
309
4.0 kcal/mol. Starting at the LPMO-Cu(II)-OO superoxo intermediate, Bertini et al. then
310
calculated energies for several possible reaction mechanisms.78 The most favored mechanism
311
involved a proton coupled electron transfer to form LPMO-Cu(II)-OOH in the presence of the
312
substrate followed by a second coupled electron transfer and a loss of water to form a LPMO-
313
Cu(II)-O intermediate, that abstracts a hydrogen from the substrate (Figure 5). The final steps of
314
the proposed mechanism are similar to what has been described earlier by Kim et al., i.e. employing
315
an oxygen-rebound mechanism.70 The authors indicated that oxidation of the substrate by H2O2
316
via a Cu(II)-O intermediate would also be possible.
317
Wang et al.79 explicitly addressed H2O2-dependent catalysis by LPMOs, testing reaction
318
pathways suggested by Bissaro et al.31 They combined small model DFT calculations, classical 15 ACS Paragon Plus Environment
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319
molecular dynamics simulations, and quantum mechanical/molecular mechanical calculations to
320
detail the mechanism of a C4 oxidizing LPMO in the presence of H2O2 and cellotriose as model
321
substrate.79 Their key findings were that there is an efficient mechanism to break the O–O bond in
322
H2O2 via a one-electron transfer from the LPMO-Cu(I) to form an HO• radical and a Cu(II)-OH
323
species. This radical is stabilized by hydrogen bonding interactions with the enzyme. Moreover,
324
the calculations showed that the formed radical preferred to abstract a hydrogen atom from the
325
Cu(II)-OH species, to form a Cu(II)-O species, rather than directly abstracting a hydrogen from
326
the substrate. The authors thus concluded that it is the Cu(II)-O species that oxidizes the C4 carbon
327
in the scissile glycosidic bond. It is worth noting that the formation of hydroxyl radicals through
328
the reaction of H2O2 with reduced transition metals is well known from biomass conversion, where
329
the Fenton reaction, usually involving iron, is thought to play a role in the degradation of
330
lignocellulosic material by brown-rot fungi.83 While the Fenton reaction seems rather unspecific,
331
LPMOs may have found a way to harness, control and direct the power of hydroxyl radicals within
332
the confinement of an enzyme-substrate complex.30,31,79 The formation of such powerful oxidative
333
species brings the risk of auto-catalytic damage to the enzymes, as is discussed in section 2.7.
334
Important modelling studies were conducted by Hedegård and Ryde.73,74 In their 2017
335
study, they calculated bond-dissociation energies (BDE) for a number of possible LPMO
336
intermediates to determine if these are sufficiently high to activate the C1-H or C4-H bonds in
337
cellulose, with calculated BDEs of 423 and 434 kJ/mol, respectively.73 The calculated BDEs for
338
Cu(II)-oxyl or a Cu(III)-oxo suggested that the reaction with C1-H or C4-H bonds will result in
339
reaction energies of −25 and −34 kJ/mol, respectively. The reaction with a Cu(III)-hydroxide was
340
47 kJ/mol, but this species was nevertheless considered as a possible candidate for hydrogen atom
341
abstraction. This study also suggested that O–O bond breaking occurs prior to hydrogen
342
abstraction from the substrate.73
343
In their 2018 follow-up study, Hedegård and Ryde employed the same crystal structure of
344
an LPMO-cello-oligosaccharide complex as used in the study by Wang et al., discussed above.74,79
345
They used a QM/MM approach to analyze the catalytic mechanism and identify the species
346
abstracting a hydrogen atom from the polysaccharide substrate. Calculations were performed for
347
both O2 and H2O2 as co-substrates. The calculations showed that when O2 is the co-substrate, a
348
Cu(II)-OO complex is formed upon reaction of Cu(I) with O2 (as shown by Kjærgaard et al.80)
349
and that protonation of this complex by a nearby histidine residue with a concomitant electron 16 ACS Paragon Plus Environment
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350
transfer leads to cleavage of the O–O bond, dissociation of water and formation of an Cu(II)-O
351
intermediate, possibly protonated, that can react with substrate. Analyses with H2O2 as the co-
352
substrate showed that the reaction with LPMO-Cu(I) led to formation of an oxyl or hydroxyl
353
complex, both sufficiently reactive to abstract a hydrogen from the polysaccharide substrate.
354
Comparative calculations showed that the reaction is more favorable with H2O2 as the co-substrate
355
compared to O2. H2O2 was found to interact with the proton of a nearby histidine as well as a with
356
a nearby glutamine residue,74 in accordance with structural data.71
357
All in all, most studies indicate that Cu(II)-oxyl or a closely related species (Cu(III)=O or
358
a protonated oxyl) is the reactive oxygen species and that this species may be generated in both
359
O2- and H2O2-driven mechanisms. For a further discussion of the relevance of these two
360
mechanisms, see section 2.8.
361 362
2.3 Kinetics of LPMO action
363
Despite the increasing number of studies published on LPMOs, detailed kinetic analyses of LPMO
364
action remain scarce (see Table 1 for a summary of kinetic data). Vaaje-Kolstad et al. reported an
365
oxidation rate of 1 min-1 (0.017 s-1) for degradation of -chitin by (bacterial) CBP21 (SmAA10A)
366
in reactions with O2 as co-substrate.1 NcAA9C is a (fungal) AA9 enzyme showing activity on
367
xyloglucan, cellulose and cellodextrins. Again, with O2 as the co-substrate, measured degradation
368
rates for NcAA9C were 0.11 s-1 for xyloglucan and 0.06 s-1 and 0.03 s-1 for an oligomeric
369
xyloglucan and cellulose substrate, respectively.13 Similar values were obtained by Borisova et al.
370
for the same LPMO.81 The pathogen Vibrio cholerae expresses a four-domain AA10-type LPMO
371
(GbpA), which is a virulence factor and not likely involved in biomass processing. Loose et al.
372
showed that the initial reaction rate for GbpA on -chitin nanofibers was 2.7 min-1 (0.045 s-1).84
373
Frandsen et al. analyzed the kinetics of LsAA9A-catalyzed oxidation of a soluble oligomeric
374
substrate analogue of (Glc)4.57 The results were interpreted using a classical Michaelis – Menten
375
approach, to yield a kcat of 0.11 s-1 and a Km of 43 M with respect to the carbohydrate substrate.
376
Importantly, this study was the first to provide an efficiency constant (kcat / Km of 2.6 • 103 M-1 s-
377
1)
378
and a reductant, such as ascorbic acid, typically at a concentration of 1 mM. Generally, the
379
observed reaction rates were slow, varying from considerably below 1 s-1 to below 1 min-1, as
380
recently summarized by Bissaro et al.32
for an LPMO system.57 In all these studies, reactions were typically run in the presence of O2
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381
The first detailed kinetic characterization of H2O2–driven LPMO catalysis was done for
382
degradation of chitin by SmAA10A by Kuusk et al.35 Central to the study was the use of [14C]-
383
labeled chitin, which provided convenient and sensitive detection of released soluble products.
384
Also here, the results were interpreted using a classical Michaelis – Menten approach to yield a
385
kcat of 6.7 s-1, which is two orders of magnitude higher than previously reported (apparent) rate
386
constants for O2-driven reactions. These analyses yielded Km values of 0.58 mg/mL and 2.8 M
387
for chitin and H2O2, respectively. It is worth noting that the kcat/ Km for H2O2 is 2 • 106 M-1 s-1,
388
which is a value commonly seen for peroxygenases.85-86 Of note, the kinetic analyses by Kuusk et
389
al. suggested that LPMO-catalyzed oxidation of chitin in the presence of H2O2 follows a ternary
390
mechanism.35
391
In a subsequent study, Hangasky et al. assessed the kinetics of MtAA9E-catalyzed
392
oxidation of (Glc)6 evaluating both O2 and H2O2 as co-substrates.33 Studies with O2 at a
393
concentration of 208 M and with varying concentrations of (Glc)6, yielded an apparent kcat of
394
10.1 min-1 (0.17 s-1) and a Km of 32 M with respect to (Glc)6, resulting in a kcat/ Km of 5 • 103 s-1
395
M-1. These values are very similar to those obtained by Frandsen et al.57 in their study of LsAA9A-
396
catalyzed oxidation of (Glc)4 (see above). Another series of experiments, with a constant (Glc)6
397
concentration of 1 mM and varying O2 concentrations (0 to 800 M), yielded an apparent kcat of
398
17 min-1 (0.28 s-1) and a Km of 230 M with respect to O2, corresponding to a kcat/ Km of ca. 1 •
399
103 s-1M-1. When H2O2 was used as co-substrate, enzyme rates were much higher and rates
400
corresponding to 50% of added
402
H2O2 had been consumed, the authors calculated observed rate constants (kobs) of 285 – 916 min-1
403
(4.8 - 15 s-1) for concentrations of H2O2 ranging from 12.5 to 100 M. No attempt was made to
404
calculate a Km with respect to H2O2. A plot of kobs vs. H2O2 concentration in a Michaelis – Menten
405
plot using data in Table S9 of this study33 yields a Km of 53 M suggesting a kcat/ Km of 3 • 105 M-1
406
s-1.
407
Of note, currently available kinetic data (Table 1), derived from multiple laboratories, show
408
that H2O2-driven LPMO reactions are orders of magnitude faster and show much better catalytic
409
efficiencies (kcat/ Km), compared to O2-driven LPMO reactions. Bissaro et al. have suggested that
410
the very low rates of LPMO catalysis observed under “standard” conditions (O2 + reductant) reflect
411
the rate of the generation of the “true” LPMO substrate, H2O2, in the reaction mixtures,30-32 18 ACS Paragon Plus Environment
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412
whereas others claim that LPMOs can indeed use O2 directly, and that O2 is the natural co-
413
substrate.33 This current controversy is addressed in some more detail in section 2.8.
414 415 416 Table 1. Kinetics of LPMO reactionsa Enzyme IDb
Reductant
Substratec
O-sourced
kcat (s-1)
KM (M)e
kcat/KM (s-1.M1)f
Apparent oxidative rate (s-1)
Ref.
LsAA9A
AscA (5 mM)
FRET subst. (10-100 µM) Glc6 (0-400 mM) Glc6 (1 mM) Glc6 (1 mM) XG14 (0.2 mM) Glc5 (0.2 mM)
O2 (atm.)
0.11
43 / -
2.6103
-
(57)
0.17
32 / -
5103
-
(33)
0.28
- / 230
1103
-
(33)
3105 g
4.8 - 15
(33)
MtAA9E
NcAA9CCBM1
AscA (2 mM)
AscA (1 mM)
Tamarind XG (5 g·L-1)
O2 (208 µM) O2 (0-800 M) H2O2
O2 (atm.)
PASC (5 g·L-1)
SmAA10Ah
417 418 419 420 421 422 423 424 425 426 427 428 429 430 431
VcAA10BX-YCBM73i
nd
nd
nd
0.06
nd
nd
nd
0.03
nd
nd
nd
0.11
nd
nd
nd
0.11
(13)
Reduced Glutathione (1 mM)
-chitin (0.45 g·L-1)
O2 (atm.)
nd
nd
nd
0.017
(1)
AscA (100 µM)
CNW
H2O2
6.7
0.58 mg.mL-1 / 2.8 M
2106
-
(35)
AscA (1 mM)
-chitin nanofibers (5 g·L-1)
O2 (atm.)
nd
nd
nd
0.045
(84)
a
See reference 32 for a more extensive review of LPMO reaction rates and reaction conditions. Abbreviations: Ls, Lentinus similis; Nc, Neurospora crassa; Sm, Serratia marcescens; Mt, Myceliophthora thermophila (new name Thermothelomyces thermophila); Vc, Vibrio cholerae. c Abbreviatons: FRET subst. = fluorescence-labeled cellotetraose57; CNW, chitin nanowhiskers; PASC, phosphoric acid-swollen cellulose; XG, xyloglucan; XG14, xyloglucan oligomer. d O-source refers to the molecule (i.e. O or H O ) from which the oxygen used in the (per)oxygenase reaction presumably was 2 2 2 derived. Abbreviation: atm., atmospheric. e K values (in µM, unless stated otherwise) for the substrate and the O-source compound, respectively. M f Note that the K may refer to the substrate or the (oxygen containing) co-substrate. M g This value is an estimate made on the basis of data presented in Ref 33; see text for details. h Also known as CBP21 i GbpA is a four-domain protein where X and Y denote unknown domains related to the flagellin protein p5 and pili-binding chaperone FimC, respectively165. nd: not determined b
432 433
2.4 LPMOs among other oxidoreductases
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434
Another family of monooxygenases that can act as peroxygenases to activate C-H bonds are
435
cytochromes P450 (CYP). CYPs are found in all kingdoms of life and have a vast variety of
436
substrates. The active site of CYP contains a heme prosthetic group where the heme-iron is bound
437
to the protein through a cysteine thiolate ligand. In its resting state, the iron has a +3 charge. The
438
catalytic cycle begins with reductive activation of oxygen to form a ferryl–oxo with an porphyrin
439
π-cation radical known as Compound I, which is the hydroxylating species in an oxygen rebound
440
mechanism (Figure 7).87,88 Due to the vast number of CYPs and substrates, there are large
441
variations in observed catalytic rate constants for the various monooxygenase reactions, ranging
442
from 0.1 to 3 s-1.89,90
443
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444 445
Figure 7. A catalytic cycle of oxygen activation by cytochromes 450 and its H2O2-shunt. The
446
upper left state represents the resting state of the enzyme. Note that compound I bears resemblance
447
with the closely related Cu(III)-oxo and Cu(II)-oxyl states that may be formed during LPMO
448
catalysis. Adapted with permission from reference 87. Copyright 2014, Springer.
449 450 21 ACS Paragon Plus Environment
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451
CYPs can short-cut the generation of Compound I by using hydrogen peroxide, which
452
reacts with the CYP in its resting (i.e. non-reduced, or Fe(III)) state in a so-called H2O2-shunt
453
pathway. However, this shunt pathway is generally inefficient. As an example, naphthalene
454
oxidation by H2O2 using CYP3A4, the most abundant P450 enzyme in human liver, has a kcat of
455
1.6 min-1 (0.03 s-1) vs. 43 min-1 (0.7 s-1) for O2 and NADPH supported oxidation.91 A general
456
hypothesis used to explain this low efficiency is that CYPs lack a residue near the active site that
457
can participate in general acid-base catalysis important for the formation of Compound I in H2O2-
458
dependent oxidation by classical heme peroxidases and peroxygenases.87 Moreover, the high
459
hydrophobicity of the heme pocket of CYPs is unfavorable for the formation of Compound I via
460
the H2O2 shunt pathway (as opposed to the “normal” O2-pathway) and the subsequent
461
peroxygenation. A plethora of studies show that the peroxygenase activity of CYP can be enhanced
462
either through site-directed mutagenesis or directed evolution,91,92 but there are not many mutants
463
that show activities that are higher than those obtained in the monooxygenase reaction.87
464
Moreover, peroxide-driven reactions usually require high concentrations of peroxide that promote
465
enzyme inactivation.93-95 The need for such high concentrations is reflected in low catalytic
466
efficiency constants with respect to H2O2. The oxidation of 7-benzyloxyquinoline by CYP3A4
467
with varying concentrations of H2O2 yielded a kcat of 7.6 min-1 (0.13 s-1) and a Km of 61 mM
468
yielding a kcat/Km of 2 M-1s-1.91 In general, CYP have high Km values for H2O2 in the millimolar
469
range (20-250).96-98 Importantly, a few CYPs do seem to be genuine peroxygenases and these
470
enzymes have structural features that place a general acid-base residue near the catalytic center.99
471
These enzymes use H2O2 to catalyze the hydroxylation of long alkyl chain fatty acids with high
472
catalytic activity and substrate specificity.95,99-102 Here, the Km with respect to H2O2 has been
473
determined to be as low as 21 M.103
474
Due to the newly discovered peroxygenase activity of LPMOs, it is tempting to compare
475
this class of enzymes with CYPs. Both enzyme classes are able to activate C-H bonds with both
476
O2 and H2O2 as co-substrates. While LPMOs seem highly substrate specific, CYPs tend to be
477
promiscuous. Similar kcat values are typically observed for the two enzymatic systems when O2 is
478
the co-substrate. This changes rather drastically when H2O2 is the co-substrate, which leads to
479
increased catalytic rates and efficiency constants for LPMOs, and decreased catalytic rates and
480
efficiency constants for CYPs. The most striking characteristic is the clear difference in the Km
481
values, where LPMOs have up to a 1,000-fold higher affinity for H2O2 than CYP. Combined, this 22 ACS Paragon Plus Environment
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482
results in LPMOs having up to a million-fold higher catalytic efficiency constants with respect to
483
H2O2 than CYPs. In contrast to CYPs, the catalytic centers of LPMOs contain residues that may
484
contribute to H2O2-driven catalysis through hydrogen bonding and acid/base functionality (Figure
485
1).39,71,79,104
486
Another clear difference pertains to the redox state of the enzymes: while both CYP and
487
LPMOs activate O2 in their reduced state (i.e. Fe(II) and Cu(I), respectively), activation of H2O2
488
requires the LPMO to be in its reduced Cu(I) form whereas CYP is in its oxidized Fe(III) form
489
when working in shunt mode. A similar shunt pathway in LPMOs (i.e. LPMO-Cu(II) + H2O2)
490
would entail the unlikely reduction of H2O2 by Cu(II), or the formation of a Cu(II)-hydroperoxo
491
intermediate requiring deprotonation of H2O2. While such a mechanism perhaps cannot be
492
excluded, fact is that H2O2-driven catalysis by LPMOs is multiple orders of magnitude faster when
493
the LPMO is first reduced and this situation leads to stable reaction kinetics and high turnover
494
numbers.31,37
495 496 497
2.5 The role and nature of reductants
498
The reduction of LPMO-Cu(II) to LPMO-Cu(I) is generally accepted as a necessary step preceding
499
catalysis. It has been demonstrated that this reduction step can be carried out by plethora of
500
reductants, including small, organic molecules such as ascorbic acid,1 reduced glutathione,1
501
cysteine,17,27 a wide range of plant biomass or fungal phenolic compounds,27,28 lignin and its
502
fractions,6,105-108 and oxidoreductases.8,11,109 The most studied enzymatic electron donor is cellobiose dehydrogenase (CDH).8,11,28,110-
503 504
112
505
terminal heme b-binding AA8 cytochrome domain (CYT) connected by a flexible linker to a C-
506
terminal
507
AA3_1).110,113,114 Some CDHs have an additional C-terminal CBM1 domain binding to
508
cellulose.115 The DH domain is the catalytic part of the enzyme, carrying out a two-electron
509
oxidation of cellobiose and several other oligosaccharides, via the reduction of the FAD cofactor.
510
The reoxidation of the FAD cofactor may be carried out via sequential inter-domain electron
511
transfer (IET) to a CYT domain or via reduction of a two-electron acceptor, including O2.
512
Reduction of O2 leads to the formation of H2O2.116-118 The reduced CYT domain will transfer single
CDHs are bi-modular flavocytochromes, so far only identified in fungi, containing an Nflavin
adenine
dinucleotide
(FAD)-dependent
23 ACS Paragon Plus Environment
dehydrogenase
domain
(DH;
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513
electrons to appropriate acceptors, which includes the LPMOs.110-112 Even though the existence of
514
a putative CDH “docking” site on AA9 LPMOs has been proposed,68 experimental data and
515
computational modelling suggest direct electron transfer at the active site of the LPMO.110,111
516
Recently, a pyrroloquinoline quinone (PQQ)-dependent pyranose dehydrogenase (PDH)
517
from Coprinopsis cinerea (CcPDH) has been shown to activate cellulose-active AA9 LPMOs.109
518
Notably, PDH does not belong to the superfamily of glucose-methanol-choline (GMC)
519
oxidoreductase, which includes CDH. CcPDH has a three-domain structure with an N-terminal
520
heme b-binding AA8 CYT domain, a central PQQ-dependent AA12 DH domain and a C-terminal
521
CBM1.119 The CcPDH shows oxidative activity towards ᴅ-glucosone (2-keto-ᴅ-glucose), L-fucose
522
and some rare pyranoses, and, in contrast to the CDH, does not oxidize cello-oligosaccharides or
523
glucose, which makes this enzyme a promising tool for mechanistic studies of LPMOs and for
524
lignocellulose biorefining.
525
LPMO reactions may also be fueled by photocatalytic systems that provide electrons.120,121
526
In one approach, Bissaro et al. showed that light-driven oxidation of water catalyzed by vanadium-
527
doped titanium dioxide, is capable of providing reducing equivalents to LPMOs and drive LPMO
528
reactions, albeit at low rate.121 In a very important study, Cannella et al. reported that exposure of
529
plant derived pigments (e.g. chlorophyllin) to low intensity light in the presence of ascorbic acid
530
speeds up LPMO catalytic rates by up to 100-fold.120 This effect was attributed by the authors to
531
the delivery of high redox potential electrons to the LPMO from the photo-excited pigment
532
molecule. The nature of the observed enhancement of LPMO catalysis remains controversial. In
533
contrast with claims made in the ground-breaking work by Cannella et al. (see also Möllers et
534
al.122), the authors of this review have proposed that the efficiency of the chlorophyllin-light
535
system may be due to the formation of hydrogen peroxide.30-31 Importantly, the study by Cannella
536
et al. showed that LPMOs could attain much higher catalytic rates than previously thought.120
537
Kracher et al. described the correlation between the reduction potential of the reductant and
538
the catalytic performance of LPMOs.28 Using variation in pH to manipulate redox potentials of
539
reductants, Frommhagen et al. showed a similar correlation.123 Several studies on O2-driven LPMO
540
reactions have shown that higher concentrations of reductant lead to higher catalytic rates and
541
higher consumption of O2.37,112,124 However, high concentrations of reductant also lead to faster
542
inactivation of the LPMO, as discussed below.
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543
Considering the discrepancy between the rate of LPMO reduction (in the milliseconds
544
range; Kracher et al.28) and the reported apparent catalytic rates (in the minute range), it is unlikely
545
that reduction of LPMO-Cu(II) complex is a rate limiting step of an O2-driven LPMO reaction.
546
Thus, in the O2-driven reaction mechanism of LPMOs (Figure 2A), the delivery of the second
547
electron must be the rate-limiting step. In case of the peroxygenase mechanism (Figure 2B),
548
delivery of a second electron is not required. In the latter scenario, the generation of H2O2, either
549
via reactions between the reductant and O2 or via superoxide formation by the reduced LPMO
550
itself,80 may be the rate-limiting step. Of note, Hegnar et al. recently showed a correlation between
551
the (pH-dependent) generation of H2O2 by an LPMO-reductant combination in the absence of
552
substrate, and LPMO activity in the presence of substrate.125
553 554
2.6 The role of the substrate
555
Copper-binding studies indicate that LPMOs bind copper with nanomolar affinities and,
556
importantly, that Cu(I) binds with higher affinity compared to Cu(II).9,24 The catalytic copper site
557
of LPMOs is highly accessible (Figure 1) and it is difficult to envisage how an enzyme with
558
basically no substrate-binding pocket and no confinement around a highly reactive Cu(I) species
559
can control its reactivity and specificity. Although this issue remains partly unresolved, it is clear
560
that the substrate plays a major role,57,126 since binding to substrate, especially binding to an
561
extended substrate surface, secludes the copper from the solution and creates confinement at the
562
active center (Figure 8).39,111
563 564
Figure 8. Binding of LPMO to an extended substrate surface. Panel A shows an experimentally
565
guided model of LPMO SmAA10A bound to crystalline chitin. Panel B shows the active site in
566
the complex and the positioning of substrate hydrogen atoms relative to the copper (green and red 25 ACS Paragon Plus Environment
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567
carbons in the protein show the (almost identical) situations with and without substrate,
568
respectively). The Cu-H1 distance is ~1 Å shorter than the Cu-H4 distance, rationalizing the C1
569
oxidizing activity of this LPMO. Panel C shows the heat mapped LPMO-chitin interaction surface
570
indicating that most interactions occur along carbohydrate residues +3 to -5, all belonging to a
571
single polysaccharide chain. The residues labeled a-h belong to polysaccharide chains adjacent to
572
the central chain. Adapted with permission from reference 39. Copyright 2018, American
573
Chemical Society.
574 575
Insight into substrate-binding to LPMOs comes from NMR studies with both crystalline and
576
soluble substrates,24,111 X-ray crystallographic studies of LPMOs in complex with soluble
577
substrates (see section 2.2),57,126 and modelling.39,62,68 In addition, EPR studies provide insight into
578
how substrate binding changes the electronics of the copper site (see section 2.2).39,57,81 Of note,
579
such electronic effects of substrate binding have been observed for both cellulose and chitin. The
580
structures of enzyme-substrate complexes described in a seminal paper by Frandsen et al.57 and in
581
a follow up paper by Simmons et al.126 show how substrate-binding generates small
582
rearrangements of the copper site. Interestingly, a comparison of the binding of cello-oligomers
583
and xylo-oligomers to the same LPMO showed that the different substrates create quite different
584
environments near the copper.126 While the biological relevance of these differences is unclear, for
585
example because the xylo-oligosaccharides may not be the true substrates, they do underpin that
586
substrate-binding has an effect on the copper environment.
587
In an experimentally guided modeling study, Bissaro et al. generated models of complexes
588
between chitin-active SmAA10A and crystalline chitin (Figure 8).39 In these models the T-shaped
589
Cu-3N active site assumed a position relative to the substrate that was compatible with catalysis.
590
The models further showed that multiple conserved amino acid residues, spread over the putative
591
substrate-binding surface and interacting with multiple polysaccharide chains, are involved in
592
binding and correct positioning of the substrate. It was also shown that small molecules such as
593
water, oxygen and hydrogen peroxide could enter the confined reaction cavity formed in the
594
enzyme-substrate interface through a narrow tunnel, while larger molecules, such as ascorbic acid,
595
may not be able to reach the active site when the enzyme sits on a surface of crystalline substrate.
596
It is important to note that a reduced LPMO that is bound to substrate differs a lot from a
597
reduced LPMO in solution. In the latter situation, the enzyme basically carries an exposed reduced 26 ACS Paragon Plus Environment
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transition metal that can engage in all kinds of (off-pathway) redox reactions (see next section).
599
When bound to its correct substrate, the copper site is secluded from solvent and precisely
600
positioned relative to the to-be-oxidized carbon.39,57,79 It is thus important for a reduced LPMO to
601
bind to its substrate to prevent engagement in off-pathway reactions. Accordingly, it has been
602
shown that reduction of LPMOs increases their affinity for the substrate.36,82,127
603 604
2.7 LPMO inactivation
605
Progress curves for LPMO reactions are often non-linear (Figures 9 and 10). It is now clear
606
that LPMOs, like other metallo-enzymes (see section 2.4) are subject to oxidative damage if the
607
conditions are not right. This may explain the lack of linearity in progress curves, although in some
608
studies the slow disappearance of LPMO activity may also have been due to depletion of another
609
reactant, such as the reductant. Work by Loose et al. studying engineered variants of chitin-active
610
SmAA10A in O2-driven reactions,128 and Bissaro et al. studying cellulose-active ScAA10C in
611
H2O2-driven reactions,31 have shown that LPMO inactivation is accompanied by oxidative damage
612
of residues close to the copper-site.
613
Considering the ongoing discussion on the roles of O2 versus H2O2 in LPMO catalysis (see
614
section 2.8), it is of major importance to note that autocatalytic inactivation of LPMOs happens no
615
matter how the reaction is fueled, as clearly shown by the data presented in Figures 9 and 10.
616 617 618
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619 620
Figure 9. Activity of a bacterial chitin-active LPMO, SmAA10A, in O2-driven reactions.
621
Panel A shows degradation of -chitin under aerobic conditions using a fungal cellobiose
622
dehydrogenase for the supply of reducing equivalents. Higher concentrations of CDH increase the
623
initial activity of the LPMO (higher product levels at 4 h up to a CDH concentration of 3 M), but
624
reduce enzyme stability (reduced increase in product levels between 4h and 24h). Panel B shows
625
degradation of -chitin under aerobic conditions in the presence of varying amounts of ascorbic
626
acid. Higher concentrations of ascorbic acid give higher initial rates but enzyme inactivation
627
becomes noticeable at the highest concentration and the total yield relative to the amount of added
628
reductant decreases. All reactions were carried out at pH 6.0, with 10 mg/ml -chitin and 1 M
629
SmAA10A. The total amount of solubilized oxidized products was determined after enzymatically
630
converting soluble oligomers to a mixture chitobionic acid and N-acetylglucosamine. Adapted
631
with permission from reference 112. Copyright 2016, John Wiley & Sons.
632 633
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634 635
Figure 10. LPMO activity during degradation of Avicel with the LPMO-containing
636
commercial cellulase cocktail Cellic CTec2. Panel A shows the accumulation of C4-oxidized
637
LPMO products over time in reactions containing 5 mM ascorbic acid and with varying oxygen
638
concentrations in the air used to sparge the reaction mixture and present in the head space of the
639
bottle. Higher oxygen concentrations give higher initial yields but also faster LPMO inactivation.
640
Note that the C4-oxidized product monitored here is unstable, which explains why levels gradually
641
go down as production ceases. Panel B shows the accumulation of LPMO products over time in
642
reactions containing 1 mM ascorbic acid that were carried out in fermentors under anaerobic
643
conditions, with feeding of H2O2 at a rate that is indicated in the Figure in M h-1. In the absence
644
of oxygen and H2O2 the LPMO is not active, whereas increasing amounts of added H2O2 lead to
645
faster catalytic rates but also faster enzyme inactivation. All reactions were carried out at pH 5.0,
646
with 100 mg mL-1 Avicel and 4 mg Cellic CTec2 proteins per gram of Avicel. Adapted with
647
permission from reference 37. Copyright 2018, BioMed Central.
648 649 650 651
As briefly alluded to in sections 2.3 & 2.6, reduced LPMOs that are not bound to substrate
652
contain a solvent-exposed Cu(I) ion. Although, LPMOs seem to have evolved to stabilize this
653
Cu(I) in the absence of substrate (since they bind Cu(I) more strongly than Cu(II); Aachmann et
654
al.24), it is plausible that this reduced transition metal engages in reactions with O2 or H2O2 leading 29 ACS Paragon Plus Environment
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655
to the formation of reactive oxygen species, including, possibly, highly damaging species such as
656
hydroxyl radicals. In the absence of substrate, these species will react with something else close to
657
the catalytic center, e.g. the copper coordinating histidines, as has indeed been observed.31 Kinetic
658
studies on H2O2-driven catalysis by chitin-active SmAA10A indicated that the rate for H2O2-driven
659
enzyme inactivation is about 1000-fold lower than the rate for H2O2-driven cleavage of chitin.35 It
660
is likely that the ratio between these two rates will vary between LPMOs and LPMO-substrate
661
combinations.
662
In line with the above considerations, Bissaro et al. showed that the presence of substrate
663
protects LPMOs from oxidative inactivation.31 Similar conclusions can be drawn from data in
664
Hangasky et al.33 Subsequently, studies of the effects of mutating residues on the substrate-binding
665
surface and of removing CBMs have shown a clear correlation between the affinity of the LPMO
666
for its substrate and LPMO stability.66,128,129 Likewise, it has been shown that higher substrate
667
concentrations promote LPMO stability.130 Of note, these studies on the impact of substrate on
668
LPMO stability concern several LPMOs acting on several substrates. There is no doubt that the
669
presence of substrate has a major effect on protecting an LPMO from autocatalytic oxidative
670
damage.
671
Potential enzymatic redox partners of LPMOs such as CDH8,11,110,112 and the
672
pyrroloquinoline quinone-dependent pyranose dehydrogenase (CcPDH),109 described above, often
673
contain a cellulose-binding CBM. Interestingly, Várnai et al. found that removal of the CBM from
674
CcPDH reduced the efficiency of CcPDH-driven LPMO action.109 Although speculative, this
675
suggests that it is beneficial that activation of the LPMO happens in the vicinity of the substrate,
676
i.e. by a substrate-bound rather than a freely moving CcPDH. This would be in line with the notion
677
that substrate binding is crucial to prevent inactivation of activated LPMOs.
678
These stability issues and the role of substrate are crucially important from a practical point
679
of view. During industrial bioprocessing of lignocellulosic biomass, both the amount and the
680
nature of the substrate will change along the reaction, while the LPMO is inactivated. In fact, it is
681
conceivable that in “typical” bioprocessing reactions with LPMO-containing commercial cellulose
682
cocktails, the LPMOs are no longer active during the final phase of the process, where the most
683
recalcitrant part of the substrate remains and the LPMOs may be most needed. These issues are
684
further addressed in section 3 of this review.
685 30 ACS Paragon Plus Environment
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686 687
2.8 The nature of the oxygen co-substrate
688
The discovery, in 2016,30 that LPMOs display peroxygenase activity, and perhaps may not
689
act as monooxygenases at all, has created quite some stir. While the peroxygenase activity and the
690
high catalytic rates that may be achieved by harnessing this activity now have been confirmed in
691
multiple laboratories,31,33,35,38 there remains controversy as to the question whether LPMOs do use
692
O2 directly and thus carry out a monooxygenase reaction. Some, including the authors of this
693
review, argue that H2O2 likely is the only, or, at least, the only kinetically relevant, co-substrate,
694
whereas others claim that O2 not only is relevant but is the natural co-substrate of LPMOs.33
695
Bissaro et al. have recently reviewed existing data on LPMO functionality in light of what is known
696
about other H2O2 producing or consuming redox enzymes that are (potentially) involved in fungal
697
degradation of lignocellulosic biomass.32
698
It is worth noting that a possible mix up of an oxygenase activity and a perxoygenase
699
activity is not unprecedented. In 2013, Wang et al. showed that HppE, a non-heme mono-iron
700
epoxidase involved in the production of fosfomycin, which had been studied for more than a
701
decade,131 is a peroxidase, reducing H2O2, rather than an oxidase, reducing O2.132
702
In their original work on SmAA10A, Vaaje-Kolstad et al. showed that one heavy oxygen
703
atom, provided in the form of 18O2, was incorporated in the oxidized products.1 While this may
704
seem to demonstrate a monooxygenase reaction, advocates of the peroxygenase activity of LPMOs
705
would argue that under the conditions of the assay, O2 may first have been converted to H2O2,
706
which then was used by the LPMO. They would argue that this may happen under all reaction
707
conditions usually used in LPMO research, not in the least because non-substrate bound LPMOs
708
generate H2O2 themselves.12,133 The competition experiments shown in Figure 6 and discussed in
709
section 2.2 may be taken to support this view.
710
In support of the peroxygenase reaction, Bissaro et al. showed that LPMO activity is
711
inhibited in reactions where horseradish peroxidase (HRP) competes for H2O2,31 and similar results
712
have later been described by Hangasky et al. for reactions with insoluble substrates.33 Importantly,
713
Hangasky et al. showed that the inhibitory effect of HRP on LPMO activity becomes less
714
pronounced at higher substrate concentrations and they demonstrated that when using soluble
715
substrate, cellohexaose, the inhibitory effect of HRP is virtually absent.33 These observations were
716
taken to show that, under some conditions, H2O2 does not play a role in LPMO catalysis and that, 31 ACS Paragon Plus Environment
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717
thus, O2 is used directly in a monooxygenase (or “coupled”) reaction. The authors then concluded
718
that LPMOs display both a peroxygenase (or “uncoupled”) reaction and a monooxygenase reaction
719
and that the occurrence of these two mechanisms is substrate-dependent. While we cannot exclude
720
that this conclusion is correct, we would argue that there is a plausible alternative explanation for
721
the observations made by Hangasky et al.33: it is conceivable that at high substrate concentrations,
722
and in particular when using an easily diffusible, soluble substrate, the H2O2–driven LPMO
723
reaction is so fast that HRP cannot compete for the H2O2. For the same reasons, the absence of an
724
effect of (H2O2–consuming) catalase on LPMO catalysis under certain conditions, cannot be taken
725
to indicate that LPMOs do not use H2O2. Kinetics dictate that the LPMO can easily compete with
726
catalase, as recently discussed by Kuusk et al.36
727
In any case, available data show that H2O2-driven LPMO reactions are orders of magnitude
728
faster than O2-driven LPMO reactions, as outlined in section 2.3 (Table 1). The O2 mechanism
729
may exist and be biologically relevant, for example because O2 concentrations will be higher than
730
H2O2 concentrations in many eco-systems. Still, as shown by the competition experiments of
731
Figure 6 and the low KM values for H2O2, low concentrations of H2O2, potentially generated by
732
enzymes secreted specifically for this purpose,32 will lead to H2O2 consumption by LPMOs and
733
speed up LPMO reactions.
734
While this review is not the place to finally settle the issue of the true co-substrate, it is
735
important to point at a few claims that are propagated in the literature and that are clearly incorrect.
736
Firstly, it has been suggested that LPMOs are more prone to inactivation in H2O2-driven reactions
737
compared to O2-driven reactions. This is not true, as illustrated by Figures 9 and 10, above. Of
738
course the degree of inactivation will be affected by a multitude of factors that may vary between
739
experiments, such as the H2O2 concentration in H2O2-driven reactions or the reductant type and
740
concentration in O2-driven reactions, or the substrate used. So, differences in inactivation rates
741
may be observed, but these are most likely due to varying reaction conditions, rather than to
742
fundamentally different catalytic processes.
743
Secondly, it has been claimed, or at least suggested, that LPMOs become less specific if
744
the reaction is fueled by H2O2 rather than O2. For example, when studying H2O2-driven
745
degradation of cellohexaose by a fungal LPMO, Hangasky et al. detected minor amounts of
746
products carrying atypical oxidations, indicating a lack of specificity.33 While these results are
747
indisputable, there is no evidence that the observed lack of specificity is due to the participation of 32 ACS Paragon Plus Environment
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748
H2O2 in the reaction. It should also be noted that cellohexaose is not necessarily the natural or
749
optimal substrate of the LPMO used and its binding may thus not provide the active site
750
confinement that is needed to control and direct the strong oxidative power of the emerging
751
reactive oxygen species (see section 2.3). The elegant structural work by Simmons et al. shows
752
that different oligomeric substrates have different binding modes with clear effects on the spatial
753
arrangement near the copper site.126 Since the substrate is crucial in controlling the orientation of
754
the reactive oxygen species,39,57,79 it is conceivable that aspecific background reactions occur for
755
certain LPMO-substrate combinations. Of note, in many reaction setups, there will be a gradual
756
increase in oxidative damage in the catalytic center of the LPMO,31,128 which could also affect
757
specificity. We have characterized multiple LPMOs (fungal, bacterial), with varying oxidative
758
regioselectivity (C1/C4/C1&C4), acting on multiple substrates (cellulose, cello-oligosaccharides,
759
xyloglucan, chitin) in both O2 and H2O2-driven reactions. In our hands, well controlled O2 and
760
H2O2-driven reactions give the same overall product profiles indicating that there is no basis to
761
claim that LPMOs become less specific if they are fueled with H2O2.
762 763 764
3. Harnessing LPMOs in biomass processing
765 766
3.1 Degradation of lignocellulosic biomass
767
Lignocellulosic biomass is mainly composed of lignin, cellulose and a variety of hemicelluloses.
768
These three polymers are interlinked in a complex matrix where their relative abundance varies
769
depending on the type of biomass. Native lignocellulosic biomass is generally very compact and
770
little accessible to enzymatic attack. Thus, biomass processing is initiated with some sort of
771
pretreatment that rips the fibers apart and makes the biomass accessible for enzymes.134
772
Commercial enzyme cocktails for saccharification of lignocellulosic biomass, such as Cellic
773
CTec® products from Novozymes and Accellerase® products from DuPont, contain enzyme
774
activities that degrade the polysaccharides, leaving lignin as a solid residue. Cellulose is of
775
particular interest due to its abundance in the biomass and due to its simple composition, being a
776
linear polysaccharide of β-1,4 linked D-glucose units.
777
It is generally believed that efficient cellulose degradation requires the presence of three
778
classes of hydrolytic enzymes: endo-1,4-β-glucanases, randomly cleaving internal glucose bonds, 33 ACS Paragon Plus Environment
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779
cellobiohydrolases, which attack the reducing or non-reducing ends of the cellulose polymer
780
releasing cellobiose in a processive manner, and β-glucosidases, converting cellobiose to glucose
781
(Figure 11).5,135 Although the compositions of modern commercial cellulase cocktails are not
782
publicly available, these cocktails likely contain all three enzyme types. In addition, these modern
783
cocktails contain one or more LPMOs,44,45 which cleave internal bonds and increase the overall
784
efficiency of the enzyme blend.37, 44-50 Harnessing the potential of LPMOs in industrial biomass
785
processing is not straightforward and puts new demands on process design. One obvious issue
786
concerns the fact that (controlled) addition of air or hydrogen peroxide is necessary. By acting on
787
the surfaces of insoluble substrates, LPMOs improve the accessibility for canonical cellulases,
788
perhaps particularly in the most recalcitrant parts of the substrate,41,42 and thus improve the overall
789
efficiency of cellulase cocktails.
790
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791
792 793 794
Figure 11. Degradation of cellulose by LPMOs and cellulases. The upper half of the Figure
795
shows LPMO reactions, whereas the lower half illustrates the various cellulases. For clarity,
796
chemical equations for the O2- and H2O2-dependent reaction schemes (indicated by “A” and “B”
797
in the Figure, respectively) that are also shown in Figure 2 appear beneath the figure. Productive
798
LPMO reactions start with step 0, i.e. reduction of the copper, after which an O2- (A) or an H2O2-
799
(B) driven reaction may occur. In H2O2-driven catalysis the copper stays reduced (blue) in between
800
catalytic cycles, whereas the redox state of the copper during and at the end of O2-driven catalytic 35 ACS Paragon Plus Environment
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801
cycles is less clear; see section 2.2 and Figure 5 for more details. The figure also shows multiple
802
redox side reactions: generation (5) and consumption (5’) of H2O2 in reactions involving the
803
reductant, H2O2 generation by the LPMO (3), and inactivation (4) of LPMOs that are not bound to
804
substrate (2). Next to LPMOs, multiple cellulases may contribute to cellulose degradation:
805
endoglucanases (EG), processive cellobiohydrolases moving into both possible directions (6;
806
CBH), and a beta-glucosidase (BG) converting oligomeric products, mostly dimers, to monomeric
807
glucose.
808 809 810
3.2. Breaking down the hemicellulose-cellulose matrix
811
In lignocellulosic biomass, even after an efficient pretreatment, cellulose remains embedded in a
812
hemicellulose-lignin matrix and there are strong indications in the literature that xylan and
813
glucomannan strongly associate with cellulose fibers (Figure 12).136-139 Indeed, accessory
814
hydrolytic enzymes such as xylanases and mannanases are known to have beneficial effects on the
815
saccharification of cellulose by cellulases, likely by increasing cellulose accessibility.49,140-143
816
Importantly, even minor amounts of remaining hemicellulose in a pretreated substrate may inhibit
817
cellulose conversion.142,143
818
While addition of specific hemicellulases to improve cellulose accessibility may be useful
819
in certain situations, additional hemicellulose-removing capacity may come from hemicellulolytic
820
side activities of one or more of the cellulases in standard cellulolytic enzyme cocktails.142,144,145
821
In particular, endoglucanases belonging to the GH7 family, such as Cel7B from Trichoderma
822
reesei, show activities on both xylan and glucomannan that are important in the saccharification
823
of softwood.144
824
Similarly to certain cellulases, substrate promiscuity has been reported for cellulose-active
825
LPMOs belonging to the AA9 family, where known substrates include xyloglucan, glucomannan,
826
mixed-linkage glucan13,146 and xylan.14,126 The ability of cellulose-active LPMOs to cleave
827
hemicelluloses seems independent of their oxidative regioselectivity. For example, xyloglucan is
828
cleaved by C1-oxidizing TtAA9E,120 C4-oxidizing NcAA9C,13 and C1/C4-oxidizing GtAA9A-
829
2.146 Although less well explored up to now, additional industrially relevant functionalities of
830
LPMOs could relate to their ability to remove hemicelluloses that contribute to biomass
831
recalcitrance by binding tightly to cellulose fibrils (Figure 12). Indeed, in a seminal paper, 36 ACS Paragon Plus Environment
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832
Couturier et al. recently described a novel LPMO family (AA14) that specifically acts on those
833
parts of xylan chains that are attached to cellulose fibrils and that promotes cellulose
834
saccharification (Table 2).15 Earlier, Frommhagen et al. had detected LPMO activity on xylan but
835
also in this case only when the xylan was present together with cellulose.14
836 837
838 839 840
Figure 12. Artist impression of the interaction of xylan with cellulose fibrils. The picture,
841
generated by the Dupree group,138,139,147 shows cellulose fibrils (yellow) that are held together by
842
xylan chains that interact with multiple fibrils. Some lignin is also shown. The xylan chains carry
843
multiple decorations, as indicated in the Figure. The boxes, labeled with letters, indicate several
844
substructures in the xylan that differ in their level and type of interaction with cellulose. The
845
recently discovered xylan-active AA14 LPMOs are thought to act on parts of the xylan chains that 37 ACS Paragon Plus Environment
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846
interact with cellulose.15 Reproduced with permission from reference 139. Copyright 2016,
847
Biochemical Society.
848 849 850
3.3 The effects of LPMOs on cellulose conversion
851
The cost of cellulolytic enzyme cocktails has been greatly reduced in the past two decades,5, 44-45
852
but still remains one of the primary bottlenecks for successful commercialization of lignocellulose-
853
derived fuels and chemicals.45,148 Major enzyme producers have been looking into LPMOs and
854
their implementation in enzymatic degradation of biomass for about 15 years.5,44,45
855
Several studies preceding the discovery of the oxidative nature of LPMOs, reported enhanced
856
saccharification of cellulose by utilizing these proteins, which at the time were called GH61s. For
857
example, Merino and Cherry observed that supplementing a Trichoderma reesei secretome, which
858
harbours little GH61 proteins,149,150 with a secretome from Thielavia terrestris increased
859
degradation of pretreated corn stover (PCS), and reduced required enzyme loadings.5 This positive
860
effect was shown to be due the action of GH61 proteins present in the T. terrestris secretome.
861
Similar observations were reported by Harris et al., who showed that addition of a GH61 from
862
Thermoascus aurantiacus to a cellulase cocktail reduced the enzyme loading necessary to reach
863
90% conversion of PCS two-fold.6 Harris et al. also showed a beneficial effect of adding a
864
recombinantly produced GH61 from T. terrestris.6 Interestingly, both Merino and Cherry and
865
Harris et al. noted that none of the tested GH61s exhibited synergistic effects with canonical
866
cellulases during saccharification of “clean” cellulose substrates such as filter paper and Avicel,5,6
867
which in hindsight can be attributed to the lack of lignin and lignin-derived compounds that
868
provide the reducing power that is needed to drive the LPMO reaction.
869
In an early study, Cannella et al. compared the saccharification performance of a mixture of
870
Celluclast, an LPMO-poor149,150 cellulase cocktail, and Novozym 188, a -glucosidase
871
preparation, with the performance of Cellic CTec2, during enzymatic decomposition of
872
hydrothermally pretreated wheat straw.46 They observed increased cellulose saccharification with
873
the CTec2 preparation and detected oxidized glucose and cellobiose in the reaction mixtures.
874
Table 2 provides an overview of studies that have assessed the impact of individual LPMOs
875
on the efficiency of cellulase cocktails in cellulose saccharification. For example, Hu et al. reported
876
that spiking of LPMO-poor Celluclast, with TaAA9A resulted in an 18–63% increase in the 38 ACS Paragon Plus Environment
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ACS Catalysis
877
conversion of cellulose from pretreated corn stover, poplar and loblolly pine, depending on the
878
substrate and type of substrate pretreatment (Table 2).48
879
In a landmark study, Müller et al. analyzed the effect of TaAA9A on the saccharifying
880
efficiency of the Celluclast/Novozym 188 mixture (Table 2), while simultaneously assessing the
881
impacts of reductants and oxygen, and with quantification of LPMO activity through detection of
882
LPMO products.50 Next to showing a beneficial effect of the LPMO, the results showed a clear
883
correlation between the overall saccharification efficiency of the enzyme cocktail and the
884
generation of oxidized sugars. Furthermore, this study showed that LPMO activity and the LPMO
885
effect on saccharification efficiency were absent in reactions run under anaerobic conditions.
886
When working with pure cellulose (e.g. Avicel) the LPMO effect depended on the presence of
887
externally added reductant, whereas such addition was not necessary when working with (lignin-
888
rich) steam-exploded birch.50
889
In a follow-up study, Chylenski et al. showed that optimal use of Cellic CTec3 in the
890
depolymerization of heavily delignified sulfite-pulped Norway spruce, required the addition of
891
oxygen and reductant. A beneficial effect of adding TaAA9A to a Celluclast and β-glucosidase
892
mixture was also found for saccharification of wheat straw.151 Du et al. showed similar boosting
893
effects for an AA9 LPMO from the filamentous fungus Penicillum oxalicum, using alkali-
894
pretreated wheat straw, alkali-pretreated corn stover and delignified corncob residues (Table 2).152
39 ACS Paragon Plus Environment
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896
Table 2. The effect of LPMOs on the efficiency of lignocellulolytic cocktails Added enzyme ID
Base enzyme cocktaila
LPMO (% w/w)a – total protein content (mg/g of biomass)
Reaction conditions (pH/ T, C/ Incubation time, h)
Biomass (DM, % w/v)
T. reesei secretome
20% - 5 mg/g
5.0/50/168
AP-CS (4.7%)
30 (69-89) 63 (43-70) 33 (70-93) 18 (57-67) 23 (65-80) 32 (53-70) 23 (39-48) 33 (64-85) 22 (63-77)
TtAA9E PoxAA9 CgAA9 GtAA9A
Relative increase in yield (%)b
Ref
(6)
Celluclast
5% - 20 mg/g
4.8/50/48
CelluclastN188 Celluclast (80%) + BG (10%) T. reesei secretome
15% - 5 mg/g 15% - 8 mg/g
5.0/50/20 5.0/50/48
OP-CS (2%) OP-P (2%) OP-LP (2%) SE-CS (2%) SE-P (2%) SE-LP (2%) SE-B (10%) SP-NS (5%)
10% - 3 mg/g
5.0/50/96
SE-WS (10%)
44 (39-56)
(151)
20% - 5 mg/g
5.0/50/168
AP-CS (4.7%)
20 (69-83)
(6)
20% - 5 mg/g
4.8/48/72
1 mg/g – nr
5.0/50/48
Alk-WS (2%) Alk-CS (2%) DCCR (2%) Alk-RS (1%) AP-RS (1%) Alk-WS (2%) AP-P (0.5%) AP-Pi (0.5%)
33 (67-89) 30 (63-82) 44 (54-78) 15 (0.83-0.96)* 33 (0.24-0.32)* 30 (4.9-6.4)* 38 (0.34-0.47) 82 (0.22-0.4)
TaAA9A
897 898 899 900 901 902 903 904 905 906 907 908 909 910
Page 40 of 59
Pox cellulase cocktail Celluclast (0.9 FPU/g) Celluclast
(48)
(50) (47)
(152) (153)
10% - 2 mg/g 5.0/50/48 (154) 87% - 7.65 mg/g PcoAA14A CL847 40% - 1.65 T. reesei 5.2/45/24c AP-Pi (0.5%) 25 (0.20-0.25) (15) mg/g secretome AP-P (0.5%) 32 (0.34-0.45) 87% - 7.66 PcoAA14B mg/g AP-Pi (0.5%) 68 (0.22-0.37) a When the total protein load was not reported (nr) the amount of added enzyme cocktail is expressed in filter paper units (FPU) per gram of dry matter (DM) in the column “Base enzyme cocktail”. In this case, the quantity of oxidoreductase is given in mg/g of DM instead of % (w/w) of total protein load in the column “Oxidoreductase-total protein content”. b Increase in biomass hydrolysis yield relative to the reference reaction where the LPMO is either absent or repressed (e.g. anaerobic conditions). The absolute conversion yields (expressed in % of maximum theoretical yield or in g/L of sugars when marked with *), obtained without and with LPMO addition, are given in between brackets in italics. c In this case the LPMO were first incubated with the biomass and AscA for 76 h prior to 24 h further incubation with the cellulase cocktail. Abbreviations: Alk, alkali-pretreated; AP, acid pretreated; BG, -glucosidase; OP, organosolv pretreated; SE, steam-exploded; SP, sulfite pulped; B, birchwood; CS, corn stover; DCCR, delignified corncob residue; LP, lodgepole pine; MCC, microcrystalline cellulose; NS, Norway spruce; P, poplar; Pi, Pine; RS, ricestraw; WS, wheatstraw Cg, Chaetomium globosum; Gt, Gloeophyllum trabeum; Pco, Pycnoporus coccineus; Pox, Penicillium oxalicum JU-A10; Ta, Thermoascus aurantiacus; Tt, Thielavia terrestris.
911 912
The discovery of the peroxygenase activity of LPMOs sheds new light on what may be
913
happening during enzymatic bioprocessing of cellulose. Depending on the substrate and the
914
process setup, access to oxygen, oxygen consumption, and the chemical or enzymatic formation
915
and consumption of hydrogen peroxide will vary. When using “real” biomass, containing lignin
916
and sugars and compounds formed during harsh pretreatments, a lot of redox chemistry may 40 ACS Paragon Plus Environment
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ACS Catalysis
917
happen (e.g. ref 155). In principle, the peroxygenase activity of LPMOs, and the much higher
918
efficiency of the peroxygenase reaction, open up new possibilities for harnessing and controlling
919
LPMO activity during degradation of lignocellulosic biomass. This potential became indeed
920
apparent from studies on the digestion of a “clean” substrate, Avicel, with LPMO-containing Cellic
921
CTec2 under anaerobic conditions (Figure 10B).31,37 Controlled pumping of low amounts of H2O2
922
into anaerobically operated bioreactors allowed tight control of LPMO activity throughout the
923
duration of the hydrolysis and revealed that LPMO activity is a direct function of the amount of
924
H2O2 added. Moreover, by controlling H2O2 supply, it was possible to develop protocols that gave
925
higher saccharification yields compared to those obtained previously in similar reactions run under
926
standard aerobic conditions.
927 928 929
3.4 Challenges in the application of LPMOs in biomass processing
930
In saccharification reactions run under aerobic conditions and with real biomass, there is
931
usually enough reducing power in the substrate to fuel the LPMO. Oxygen will either be used
932
directly, in what would be a monooxygenase reaction, or be converted to H2O2, through direct
933
reactions with reductants or catalyzed by LPMOs (Figure 11), and then used in what would be a
934
peroxygenase reaction, as discussed above. Either way, supply of oxygen is needed, which is
935
expensive in large industrial reactors. To achieve high oxygen transfer rates a high airflow must
936
be maintained and also usually a high stirrer speed, which is difficult in reactors with high dry
937
matter content.156 While oxygen supply to some extent can be controlled and monitored using an
938
oxygen electrode, controlling H2O2 concentrations is easier, since H2O2 is a liquid that is readily
939
soluble in water.
940
In industrial settings, the presence of reductants in the form of lignin and other compounds
941
naturally present in the biomass or formed during biomass pretreatment may be difficult to control.
942
Both too much and too little reducing power may be problematic. Too much reducing power may
943
lead to depletion of oxygen or H2O2 (Figure 11), whereas too little reducing power may lead to
944
impaired LPMO reduction.
945
For “clean” substrates, with no or very little lignin, the situation is manageable, and LPMO
946
activity can be controlled directly by varying the concentration of reductant, varying the O2
947
concentration or by controlled addition of H2O2, as shown in a recent study by Müller, Chylenski 41 ACS Paragon Plus Environment
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948
et al. and illustrated by Figure 10, which is derived from this study.37 Using Avicel as a substrate
949
and LPMO-containing Cellic CTec2 under anaerobic conditions, Müller, Chylenski et al. showed
950
that the degree of LPMO activity could be perfectly controlled by controlling the addition of H2O2
951
(Figure 10B) and by optimizing the conditions, they reached unprecedented high conversion
952
levels.37
953
The same study also showed that when using lignin-rich, and industrially more realistic
954
substrates, the situation became much less manageable. In these cases, including unpublished work
955
from our laboratory, running reactions under standard aerobic conditions usually is at least as
956
efficient, or even better, than running reactions with added H2O2. Likely side reactions involving
957
reductants (Figure 11, step 5) play a key role, as also suggested by recent work by Peciulyte et
958
al.155
959
One important finding in the study by Müller, Chylenski et al. is that, under most
960
conditions, the LPMOs in Cellic CTec2 became deactivated during the course of the
961
saccharification reaction.37 As discussed in detail in section 2.7, substrate depletion may be one
962
cause and it is indeed conceivable that depletion of LPMO binding sites occurs as the substrate is
963
depolymerized. As discussed in section 2.7 and illustrated in Figure 11 (step 4), there needs to be
964
balance in the system: if there are more reduced LPMOs in the system than there are productive
965
binding sites, and if there is sufficient supply of the oxygen co-substrate, non-substrate bound
966
LPMOs will suffer from auto-catalytic inactivation. Of note, in reactions with added H2O2 or with
967
conditions that lead to abundant in situ H2O2 formation, H2O2 will start accumulating, which may
968
damage the other enzymes in the cocktail, as has indeed been observed.37,151
969
A final important result from the work by Müller, Chylenski et al. is the observation that
970
even under optimal conditions for H2O2-driven degradation of Avicel (i.e. a “clean”, well-
971
controlled system), the LPMOs in Cellic CTec2 seemed to run at much lower rates than the rates
972
coming out of recent kinetic studies.33,35,37 When assuming an LPMO (TaAA9A) content of 15 %
973
in Cellic CTec2,50 the LPMO activity under optimized conditions was in the order of 1 – 5 min-1
974
(0.07 s-1), which is about 100 times less than the highest LPMO rates appearing in literature.33,35
975
This may be taken to indicate that only a small fraction of the LPMOs in Cellic Ctec2 were actually
976
engaged in substrate cleavage under these conditions, which could mean that products such as
977
Cellic Ctec2 contain more LPMOs than needed when H2O2 is directly added to the reaction. In this
978
respect, Müller, Chylenski et al. showed that further boosting of LPMO activity, by increasing the 42 ACS Paragon Plus Environment
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ACS Catalysis
979
H2O2 feed, did indeed lead to higher initial LPMO activity, but also to less efficient cellulose
980
conversion and rapid inactivation of the LPMO.37 It is possible that under these conditions, the
981
cellulases became limiting. If the cellulases are not able to uncover novel LPMO binding sites by
982
peeling off oxidized oligosaccharides from the substrate surface, LPMOs get involved in side
983
reactions and become inactivated.
984
If one accepts the idea that LPMO reactions are primarily driven by H2O2, it is conceivable
985
that the high content of LPMOs in today’s enzyme cocktails is a consequence of these cocktails
986
being designed to work with O2 as oxygen source. Under such conditions part of the LPMO pool
987
is needed to generate H2O2 from O2 in situ. When H2O2 is supplied directly, all LPMOs are
988
available for polysaccharide oxidation.
989 990 991
3.5. LPMOs and fermentative valorization of biomass-derived sugars
992
The monomeric sugars obtained from saccharification of lignocellulosic biomass may be
993
converted to a variety of products through fermentation and/or chemical processes.157
994
Fermentative production of ethanol (“second generation biofuel”) is one of the best known and
995
most explored conversions.158 Another well-known conversion concerns the production of lactic
996
acid which may be used to produce bioplastics.159,160 There are two principle ways to combine
997
enzymatic hydrolysis with fermentation: the processes can be run sequentially in a separate
998
hydrolysis and fermentation (SHF) process, or simultaneously in a so called simultaneous
999
saccharification and fermentation (SSF) process.161 Generally, if enzymes and fermenting
1000
organisms have similar pH and temperature optima, SSF processes seem to be more efficient
1001
because these are one-tank processes and because liberated sugar is immediately fermented, thus
1002
alleviating product inhibition of the enzymes.162
1003
If the efficiency of cellulase cocktails depends on LPMOs, the need of these enzymes for
1004
oxygen may create problems in SSF setups. In aerobic fermentations there will be direct
1005
competition between the needs of the microorganism and the need of the LPMO in the enzyme
1006
cocktail. In anaerobic fermentations, which are commonly used, e.g. for the production of ethanol
1007
or lactic acid, oxygen scavenging mechanisms of the microbes and process parameters (i.e.
1008
anaerobic conditions) will keep the LPMO from being active. This was first noted by Cannella and
1009
Jørgensen, who observed that production of ethanol from pretreated wheat straw using LPMO43 ACS Paragon Plus Environment
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1010
containing Cellic CTec2 and yeast resulted in a 20% higher ethanol yield in an SHF setup
1011
compared to an SSF setup.162 Cannella and Jørgensen suggested that that this difference was due
1012
to competition for oxygen between LPMOs and yeast in the SFF setup, where the yeast was
1013
believed to scavenge most of the oxygen, thus outcompeting the LPMOs. Based on current
1014
knowledge concerning the role of H2O2, another explanation would be that the yeast either
1015
inhibited in situ production of H2O2 or consumed in situ produced H2O2 faster than the LPMO. In
1016
a similar study, Müller et al. analyzed the production of lactic acid by using steam exploded birch
1017
as substrate.159 In this work, a thermophilic strain of Bacillus coagulans was used together with
1018
Cellic CTec2 to compare SSF and SHF processes carried out at 50 °C. Generally, the SHF setup
1019
yielded around 30% higher production of lactic acid, and this higher yield was associated with
1020
production of oxidized sugars, indicative of LPMO activity. So, the SHF setup was more efficient
1021
and this could be coupled to the absence of LPMO activity in the SSF setup.
1022
One exciting aspect of the recent finding that H2O2 can efficiently fuel LPMO activity is
1023
that the “oxygen battle” in SSF may be avoided. It would seem possible to run fermentations with
1024
controlled addition of H2O2 in amounts that are sufficient to drive LPMO action while being so
1025
low that they do not harm the microbe. Our own preliminary data indicate that there indeed may
1026
be considerable potential in this type of experimental approach.
1027 1028 1029
4. Conclusions and perspectives
1030
The discovery of LPMOs has revolutionized research on enzymatic biomass conversion and the
1031
industrial implementation thereof. Still, many questions remain unanswered, and several of these
1032
questions are not easy to address. One key challenge lies in the multitude of reactions that may
1033
take place in “typical” reaction mixtures, in particular when using industrial (i.e. co-polymeric &
1034
complex) substrates. Even experiments with relatively “clean” substrates and relatively well-
1035
defined reaction mixtures raise questions. For example, in the original work on chitin-active
1036
SmAA10A,1 the boosting effect of the activated LPMO on chitinase efficiency was much larger
1037
than any boosting effect ever published for a cellulose-active LPMO acting in concert with a
1038
cellulase. So far, while the contributions of cellulose-active LPMOs in cellulose cocktails are
1039
considerable and undeniable, these contributions are lower than what one could expect on the basis
1040
of the original findings for chitin conversion. One attractive potential implication is that further 44 ACS Paragon Plus Environment
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1041
improvement of enzymatic cellulose conversion may still be possible, perhaps by harnessing
1042
LPMO power in a better manner, based on the most recent insights outlined above.
1043
LPMOs are abundant and widely distributed in Nature and show a stunning sequence
1044
diversity.67,163,164 It seems likely that some LPMOs are involved in bacterial virulence and act on
1045
substrates that yet have to be discovered.165-167 Lignocellulosic biomass is a complex material not
1046
only consisting of cellulose fibrils but also containing other polymers, a variety of hemicelluloses
1047
and lignins, that may be heavily intertwined (Figure 12). It is possible that the abundance of
1048
LPMOs in lignocellulose-degrading fungi reflects the fact that various LPMOs attack various
1049
substructures in the substrate, a notion for which some evidence has recently been provided.15
1050
Considering the sequence diversity of the extended substrate-binding surfaces of LPMOs and the
1051
ability of these enzymes to act on liquid-solid interfaces, it is also conceivable that some may act
1052
on non-polysaccharide substrates such as lignin or other insoluble, recalcitrant polymers.
1053
It seems likely that the full potential of LPMOs has not yet been harnessed. Firstly, the best
1054
LPMOs, combinations of LPMOs or LPMO-cellulase combinations may not yet have been
1055
discovered. Secondly, as outlined above, there is a clear lack of understanding of how to best
1056
harness LPMO action, in laboratory experiments and industrial bioreactors alike. When it comes
1057
to industrial biorefining of lignocellulosic biomass, perhaps even pretreatment strategies may need
1058
reconsideration, since the content and nature of remaining lignin will affect the efficiency of
1059
LPMOs in subsequent saccharification reactions. The operational stability of LPMOs is a
1060
potentially major success factor that may potentially be engineered or optimized by selecting the
1061
best natural candidates.
1062
Almost seventy years after Reese et al. suggested the existence of a “substrate-disrupting”
1063
factor (C1) in what is known as the C1-Cx hypothesis for enzymatic cellulose degradation,168 and
1064
almost 50 years after Eriksson et al. demonstrated that O2 plays a role in enzymatic cellulose
1065
conversion,2 we now have access to a multitude of substrate-disrupting LPMOs. We expect these
1066
enzymes to remain in the center of attention, due to their fascinating chemistry, their abundance in
1067
Nature, and their indisputable industrial importance.
1068 1069 1070 1071
Acknowledgements 45 ACS Paragon Plus Environment
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1072
We thank current and former team members for their contributions to our LPMO research, which
1073
has been supported by the Research Council of Norway, most recently through grants 240967,
1074
243663, 243950, 257622, 256766, 268002, 262853 and 270038. Additional support was received
1075
from the Marie-Curie FP7 COFUND People Programme, through the award of an AgreenSkills
1076
fellowship (under grant agreement n° 267196), and from the French Institut National de la
1077
Recherche Agronomique (INRA).
1078 1079 1080 1081
References
1082 1083 1084 1085 1086 1087 1088 1089 1090 1091 1092 1093 1094 1095 1096 1097 1098 1099 1100 1101 1102 1103 1104 1105 1106 1107 1108 1109 1110 1111
1. Vaaje-Kolstad, G.; Westereng, B.; Horn, S. J.; Liu, Z. L.; Zhai, H.; Sorlie, M.; Eijsink, V. G. H. An Oxidative Enzyme Boosting the Enzymatic Conversion of Recalcitrant Polysaccharides. Science 2010, 330, 219-222. 2. Eriksson, K. E.; Pettersson, B.; Westermark, U. Oxidation - Important Enzyme Reaction in Fungal Degradation of Cellulose. FEBS Lett. 1974, 49, 282-285. 3. Vaaje-Kolstad, G.; Horn, S. J.; van Aalten, D. M. F.; Synstad, B.; Eijsink, V. G. H. The Non-catalytic Chitin-binding Protein CBP21 from Serratia marcescens Is Essential for Chitin Degradation. J. Biol. Chem. 2005, 280, 28492-28497. 4. Karkehabadi, S.; Hansson, H.; Kim, S.; Piens, K.; Mitchinson, C.; Sandgren, M. The First Structure of a Glycoside Hydrolase Family 61 Member, Cel61B from Hypocrea jecorina, at 1.6 Å Resolution. J. Mol. Biol. 2008, 383, 144-154. 5. Merino, S. T.; Cherry, J. Progress and Challenges in Enzyme Development for Biomass Utilization. Adv. Biochem. Eng. Biotechnol. 2007, 108, 95-120. 6. Harris, P. V.; Welner, D.; McFarland, K. C.; Re, E.; Navarro Poulsen, J. C.; Brown, K.; Salbo, R.; Ding, H.; Vlasenko, E.; Merino, S.; Xu, F.; Cherry, J.; Larsen, S.; Lo Leggio, L. Stimulation of Lignocellulosic Biomass Hydrolysis by Proteins of Glycoside Hydrolase Family 61: Structure and Function of a Large, Enigmatic Family. Biochemistry 2010, 49, 3305-3316. 7. Forsberg, Z.; Vaaje-Kolstad, G.; Westereng, B.; Bunaes, A. C.; Stenstrom, Y.; MacKenzie, A.; Sorlie, M.; Horn, S. J.; Eijsink, V. G. H. Cleavage of Cellulose by a CBM33 Protein. Protein Sci. 2011, 20, 1479-1483. 8. Phillips, C. M.; Beeson, W. T.; Cate, J. H.; Marletta, M. A. Cellobiose Dehydrogenase and a Copper-dependent Polysaccharide Monooxygenase Potentiate Cellulose Degradation by Neurospora crassa. ACS Chem. Biol. 2011, 6, 1399-1406. 9. Quinlan, R. J.; Sweeney, M. D.; Lo Leggio, L.; Otten, H.; Poulsen, J. C.; Johansen, K. S.; Krogh, K. B.; Jorgensen, C. I.; Tovborg, M.; Anthonsen, A.; Tryfona, T.; Walter, C. P.; Dupree, P.; Xu, F.; Davies, G. J.; Walton, P. H. Insights into the Oxidative Degradation of Cellulose by a Copper Metalloenzyme that Exploits Biomass Components. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 15079-15084. 10. Westereng, B.; Ishida, T.; Vaaje-Kolstad, G.; Wu, M.; Eijsink, V. G. H.; Igarashi, K.; Samejima, M.; Stahlberg, J.; Horn, S. J.; Sandgren, M. The Putative Endoglucanase PcGH61D
46 ACS Paragon Plus Environment
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from Phanerochaete chrysosporium Is a Metal-dependent Oxidative Enzyme that Cleaves Cellulose. PLoS One 2011, 6, e27807. 11. Langston, J. A.; Shaghasi, T.; Abbate, E.; Xu, F.; Vlasenko, E.; Sweeney, M. D. Oxidoreductive Cellulose Depolymerization by the Enzymes Cellobiose Dehydrogenase and Glycoside Hydrolase 61. Appl. Environ. Microbiol. 2011, 77, 7007-7015. 12. Isaksen, T.; Westereng, B.; Aachmann, F. L.; Agger, J. W.; Kracher, D.; Kittl, R.; Ludwig, R.; Haltrich, D.; Eijsink, V. G. H.; Horn, S. J. A C4-oxidizing Lytic Polysaccharide Monooxygenase Cleaving Both Cellulose and Cello-oligosaccharides. J. Biol. Chem. 2014, 289, 2632-2642. 13. Agger, J. W.; Isaksen, T.; Varnai, A.; Vidal-Melgosa, S.; Willats, W. G. T.; Ludwig, R.; Horn, S. J.; Eijsink, V. G. H.; Westereng, B. Discovery of LPMO Activity on Hemicelluloses Shows the Importance of Oxidative Processes in Plant Cell Wall Degradation. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 6287-6292. 14. Frommhagen, M.; Sforza, S.; Westphal, A. H.; Visser, J.; Hinz, S. W. A.; Koetsier, M. J.; van Berkel, W. J. H.; Gruppen, H.; Kabel, M. A. Discovery of the Combined Oxidative Cleavage of Plant Xylan and Cellulose by a New Fungal Polysaccharide Monooxygenase. Biotechnol. Biofuels 2015, 8, 101. 15. Couturier, M.; Ladeveze, S.; Sulzenbacher, G.; Ciano, L.; Fanuel, M.; Moreau, C.; Villares, A.; Cathala, B.; Chaspoul, F.; Frandsen, K. E.; Labourel, A.; Herpoel-Gimbert, I.; Grisel, S.; Haon, M.; Lenfant, N.; Rogniaux, H.; Ropartz, D.; Davies, G. J.; Rosso, M. N.; Walton, P. H.; Henrissat, B.; Berrin, J. G. Lytic Xylan Oxidases from Wood-decay Fungi Unlock Biomass Degradation. Nat. Chem. Biol. 2018, 14, 306-310. 16. Vu, V. V.; Beeson, W. T.; Span, E. A.; Farquhar, E. R.; Marletta, M. A. A Family of Starch-active Polysaccharide Monooxygenases. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 1382213827. 17. Lo Leggio, L.; Simmons, T. J.; Poulsen, J. C.; Frandsen, K. E.; Hemsworth, G. R.; Stringer, M. A.; von Freiesleben, P.; Tovborg, M.; Johansen, K. S.; De Maria, L.; Harris, P. V.; Soong, C. L.; Dupree, P.; Tryfona, T.; Lenfant, N.; Henrissat, B.; Davies, G. J.; Walton, P. H. Structure and Boosting Activity of a Starch-degrading Lytic Polysaccharide Monooxygenase. Nat. Commun. 2015, 6, 5961. 18. Levasseur, A.; Drula, E.; Lombard, V.; Coutinho, P. M.; Henrissat, B. Expansion of the Enzymatic Repertoire of the CAZy Database to Integrate Auxiliary Redox Enzymes. Biotechnol. Biofuels 2013, 6, 41. 19. Henrissat, B. A Classification of Glycosyl Hydrolases Based on Amino Acid Sequence Similarities. Biochem. J. 1991, 280, 309-316. 20. El-Gebali, S.; Mistry, J.; Bateman, A.; Eddy, S. R.; Luciani, A.; Potter, S. C.; Qureshi, M.; Richardson, L. J.; Salazar, G. A.; Smart, A.; Sonnhammer, E. L L.; Hirsh, L.; Paladin, L.; Piovesan, D.; Tosatto, S. C E.; Finn, R. D. The Pfam Protein Families Database in 2019. Nucleic Acids Res. 2019, 47, D427-D432. 21. Ciano, L.; Davies, G. J.; Tolman, W. B.; Walton, P. H. Bracing Copper for the Catalytic Oxidation of C-H bonds. Nat. Catal. 2018, 1, 571-577. 22. Lieberman, R. L.; Rosenzweig, A. C. Crystal Structure of a Membrane-bound Metalloenzyme that Catalyses the Biological Oxidation of Methane. Nature 2005, 434, 177-182. 23. Cao, L. L.; Caldararu, O.; Rosenzweig, A. C.; Ryde, U. Quantum Refinement Does Not Support Dinuclear Copper Sites in Crystal Structures of Particulate Methane Monooxygenase. Angew. Chem. Int. Ed. 2018, 57, 162-166. 47 ACS Paragon Plus Environment
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24. Aachmann, F. L.; Sorlie, M.; Skjak-Braek, G.; Eijsink, V. G. H.; Vaaje-Kolstad, G. NMR Structure of a Lytic Polysaccharide Monooxygenase Provides Insight into Copper Binding, Protein Dynamics, and Substrate Interactions. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 1877918784. 25. Beeson, W. T.; Vu, V. V.; Span, E. A.; Phillips, C. M.; Marletta, M. A. Cellulose Degradation by Polysaccharide Monooxygenases. Annu. Rev. Biochem. 2015, 84, 923-946. 26. Walton, P. H.; Davies, G. J. On the Catalytic Mechanisms of Lytic Polysaccharide Monooxygenases. Curr. Opin. Chem. Biol. 2016, 31, 195-207. 27. Frommhagen, M.; Koetsier, M. J.; Westphal, A. H.; Visser, J.; Hinz, S. W. A.; Vincken, J. P.; van Berkel, W. J. H.; Kabel, M. A.; Gruppen, H. Lytic Polysaccharide Monooxygenases from Myceliophthora thermophila C1 Differ in Substrate Preference and Reducing Agent Specificity. Biotechnol. Biofuels 2016, 9, 186. 28. Kracher, D.; Scheiblbrandner, S.; Felice, A. K. G.; Breslmayr, E.; Preims, M.; Ludwicka, K.; Haltrich, D.; Eijsink, V. G. H.; Ludwig, R. Extracellular Electron Transfer Systems Fuel Cellulose Oxidative Degradation. Science 2016, 352, 1098-1101. 29. Garajova, S.; Mathieu, Y.; Beccia, M. R.; Bennati-Granier, C.; Biaso, F.; Fanuel, M.; Ropartz, D.; Guigliarelli, B.; Record, E.; Rogniaux, H.; Henrissat, B.; Berrin, J. G. Singledomain Flavoenzymes Trigger Lytic Polysaccharide Monooxygenases for Oxidative Degradation of Cellulose. Sci. Rep. 2016, 6, 28276. 30. Bissaro, B.; Røhr, Å. K.; Skaugen, M.; Forsberg, Z.; Horn, S. J.; Vaaje-Kolstad, G.; Eijsink, V. Fenton-type Chemistry by a Copper Enzyme: Molecular Mechanism of Polysaccharide Oxidative Cleavage. bioRxiv 2016, 097022. 31. Bissaro, B.; Røhr, A. K.; Müller, G.; Chylenski, P.; Skaugen, M.; Forsberg, Z.; Horn, S. J.; Vaaje-Kolstad, G.; Eijsink, V. G. H. Oxidative Cleavage of Polysaccharides by Monocopper Enzymes Depends on H2O2. Nat. Chem. Biol. 2017, 13, 1123-1128. 32. Bissaro, B.; Várnai, A.; Røhr, A. K.; Eijsink, V. G. H. Oxidoreductases and Reactive Oxygen Species in Conversion of Lignocellulosic Biomass. Microbiol. Mol. Biol. Rev. 2018, 82, e00029-18. 33. Hangasky, J. A.; Iavarone, A. T.; Marletta, M. A. Reactivity of O2 Versus H2O2 with Polysaccharide Monooxygenases. Proc. Natl. Acad. Sci. U. S. A. 2018, 115, 4915-4920. 34. Breslmayr, E.; Hanzek, M.; Hanrahan, A.; Leitner, C.; Kittl, R.; Santek, B.; Oostenbrink, C.; Ludwig, R. A Fast and Sensitive Activity Assay for Lytic Polysaccharide Monooxygenase. Biotechnol. Biofuels 2018, 11, 79. 35. Kuusk, S.; Bissaro, B.; Kuusk, P.; Forsberg, Z.; Eijsink, V. G. H.; Sørlie, M.; Väljamäe, P. Kinetics of H2O2-driven Degradation of Chitin by a Bacterial Lytic Polysaccharide Monooxygenase. J. Biol. Chem. 2018, 293, 523-531. 36. Kuusk, S.; Kont, R.; Kuusk, P.; Heering, A.; Sørlie, M.; Bissaro, B.; Eijsink, V. G. H.; Väljamäe, P. Kinetic Insights into the Role of the Reductant in H2O2-driven Degradation of Chitin by a Bacterial Lytic Polysaccharide Monooxygenase. J. Biol. Chem. 2019, 294, 15161528. 37. Müller, G.; Chylenski, P.; Bissaro, B.; Eijsink, V. G. H.; Horn, S. J. The Impact of Hydrogen Peroxide Supply on LPMO Activity and Overall Saccharification Efficiency of a Commercial Cellulase Cocktail. Biotechnol. Biofuels 2018, 11, 209. 38. Wang, D. M.; Li, J.; Wong, A. C. Y.; Aachmann, F. L.; Hsieh, Y. S. Y. A Colorimetric Assay to Rapidly Determine the Activities of Lytic Polysaccharide Monooxygenases. Biotechnol. Biofuels 2018, 11, 215. 48 ACS Paragon Plus Environment
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39. Bissaro, B.; Isaksen, I.; Vaaje-Kolstad, G.; Eijsink, V. G. H.; Rohr, A. K. How a Lytic Polysaccharide Monooxygenase Binds Crystalline Chitin. Biochemistry 2018, 57, 1893-1906. 40. Horn, S. J.; Vaaje-Kolstad, G.; Westereng, B.; Eijsink, V. G. H. Novel Enzymes for the Degradation of Cellulose. Biotechnol. Biofuels 2012, 5, 45. 41. Eibinger, M.; Ganner, T.; Bubner, P.; Rosker, S.; Kracher, D.; Haltrich, D.; Ludwig, R.; Plank, H.; Nidetzky, B. Cellulose Surface Degradation by a Lytic Polysaccharide Monooxygenase and Its Effect on Cellulase Hydrolytic Efficiency. J. Biol. Chem. 2014, 289, 35929-35938. 42. Eibinger, M.; Sattelkow, J.; Ganner, T.; Plank, H.; Nidetzky, B. Single-molecule Study of Oxidative Enzymatic Deconstruction of Cellulose. Nat. Commun. 2017, 8, 894. 43. Vermaas, J. V.; Crowley, M. F.; Beckham, G. T.; Payne, C. M. Effects of Lytic Polysaccharide Monooxygenase Oxidation on Cellulose Structure and Binding of Oxidized Cellulose Oligomers to Cellulases. J. Phys. Chem. B 2015, 119, 6129-6143. 44. Harris, P. V.; Xu, F.; Kreel, N. E.; Kang, C.; Fukuyama, S. New Enzyme Insights Drive Advances in Commercial Ethanol Production. Curr. Opin. Chem. Biol. 2014, 19, 162-170. 45. Johansen, K. S. Discovery and Industrial Applications of Lytic Polysaccharide Monooxygenases. Biochem. Soc. Trans. 2016, 44, 143-149. 46. Cannella, D.; Hsieh, C. W. C.; Felby, C.; Jørgensen, H. Production and Effect of Aldonic Acids During Enzymatic Hydrolysis of Lignocellulose at High Dry Matter Content. Biotechnol. Biofuels 2012, 5, 26. 47. Chylenski, P.; Petrović, D. M.; Müller, G.; Dahlström, M.; Bengtsson, O.; Lersch, M.; Siika-aho, M.; Horn, S. J.; Eijsink, V. G. H. Enzymatic Degradation of Sulfite-pulped Softwoods and the Role of LPMOs. Biotechnol. Biofuels 2017, 10, 177. 48. Hu, J. G.; Arantes, V.; Pribowo, A.; Gourlay, K.; Saddler, J. N. Substrate Factors That Influence the Synergistic Interaction of AA9 and Cellulases During the Enzymatic Hydrolysis of Biomass. Energy Environ. Sci. 2014, 7, 2308-2315. 49. Hu, J. G.; Chandra, R.; Arantes, V.; Gourlay, K.; van Dyk, J. S.; Saddler, J. N. The Addition of Accessory Enzymes Enhances the Hydrolytic Performance of Cellulase Enzymes at High Solid Loadings. Bioresour. Technol. 2015, 186, 149-153. 50. Müller, G.; Várnai, A.; Johansen, K. S.; Eijsink, V. G. H.; Horn, S. J. Harnessing the Potential of LPMO-containing Cellulase Cocktails Poses New Demands on Processing Conditions. Biotechnol. Biofuels 2015, 8, 187. 51. Johansen, K. S. Lytic Polysaccharide Monooxygenases: the Microbial Power Tool for Lignocellulose Degradation. Trends Plant Sci. 2016, 21, 926-936. 52. Span, E. A.; Marietta, M. A. The Framework of Polysaccharide Monooxygenase Structure and Chemistry. Curr. Opin. Struct. Biol. 2015, 35, 93-99. 53. Frandsen, K. E. H.; Lo Leggio, L. Lytic Polysaccharide Monooxygenases: a Crystallographer's View on a New Class of Biomass-degrading Enzymes. IUCrJ 2016, 3, 448467. 54. Vaaje-Kolstad, G.; Forsberg, Z.; Loose, J. S.; Bissaro, B.; Eijsink, V. G. Structural Diversity of Lytic Polysaccharide Monooxygenases. Curr. Opin. Struct. Biol. 2017, 44, 67-76. 55. Meier, K. K.; Jones, S. M.; Kaper, T.; Hansson, H.; Koetsier, M. J.; Karkehabadi, S.; Solomon, E. I.; Sandgren, M.; Kelemen, B. Oxygen Activation by Cu LPMOs in Recalcitrant Carbohydrate Polysaccharide Conversion to Monomer Sugars. Chem. Rev. 2018, 118, 25932635.
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56. Tandrup, T.; Frandsen, K. E. H.; Johansen, K. S.; Berrin, J. G.; Lo Leggio, L. Recent Insights into Lytic Polysaccharide Monooxygenases (LPMOs). Biochem. Soc. Trans. 2018, 46, 1431-1447. 57. Frandsen, K. E. H.; Simmons, T. J.; Dupree, P.; Poulsen, J. C. N.; Hemsworth, G. R.; Ciano, L.; Johnston, E. M.; Tovborg, M.; Johansen, K. S.; von Freiesleben, P.; Marmuse, L.; Fort, S.; Cottaz, S.; Driguez, H.; Henrissat, B.; Lenfant, N.; Tuna, F.; Baldansuren, A.; Davies, G. J.; Lo Leggio, L.; Walton, P. H. The Molecular Basis of Polysaccharide Cleavage by Lytic Polysaccharide Monooxygenases. Nat. Chem. Biol. 2016, 12, 298-303. 58. Zhang, L.; Koay, M.; Mahert, M. J.; Xiao, Z.; Wedd, A. G. Intermolecular Transfer of Copper Ions from the CopC Protein of Pseudomonas syringae. Crystal Structures of Fully Loaded CuICuII Forms. J. Am. Chem. Soc. 2006, 128, 5834-5850. 59. Forsberg, Z.; Mackenzie, A. K.; Sørlie, M.; Røhr, A. K.; Helland, R.; Arvai, A. S.; VaajeKolstad, G.; Eijsink, V. G. H. Structural and Functional Characterization of a Conserved Pair of Bacterial Cellulose-oxidizing Lytic Polysaccharide Monooxygenases. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 8446-8451. 60. Petrović, D. M.; Bissaro, B.; Chylenski, P.; Skaugen, M.; Sørlie, M.; Jensen, M. S.; Aachmann, F. L.; Courtade, G.; Várnai, A.; Eijsink, V. G. H. Methylation of the N-terminal Histidine Protects a Lytic Polysaccharide Monooxygenase from Auto-oxidative Inactivation. Protein Sci. 2018, 27, 1636-1650. 61. Hemsworth, G. R.; Taylor, E. J.; Kim, R. Q.; Gregory, R. C.; Lewis, S. J.; Turkenburg, J. P.; Parkin, A.; Davies, G. J.; Walton, P. H. The Copper Active Site of CBM33 Polysaccharide Oxygenases. J. Am. Chem. Soc. 2013, 135, 6069-6077. 62. Wu, M.; Beckham, G. T.; Larsson, A. M.; Ishida, T.; Kim, S.; Payne, C. M.; Himmel, M. E.; Crowley, M. F.; Horn, S. J.; Westereng, B.; Igarashi, K.; Samejima, M.; Stahlberg, J.; Eijsink, V. G. H.; Sandgren, M. Crystal Structure and Computational Characterization of the Lytic Polysaccharide Monooxygenase GH61D from the Basidiomycota Fungus Phanerochaete chrysosporium. J. Biol. Chem. 2013, 288, 12828-12839. 63. Vu, V. V.; Beeson, W. T.; Phillips, C. M.; Cate, J. H. D.; Marletta, M. A. Determinants of Regioselective Hydroxylation in the Fungal Polysaccharide Monooxygenases. J. Am. Chem. Soc. 2014, 136, 562-565. 64. Danneels, B.; Tanghe, M.; Joosten, H. J.; Gundinger, T.; Spadiut, O.; Stals, I.; Desmet, T. A Quantitative Indicator Diagram for Lytic Polysaccharide Monooxygenases Reveals the Role of Aromatic Surface Residues in HjLPMO9A Regioselectivity. PLoS One 2017, 12, e0178446. 65. Danneels, B.; Tanghe, M.; Desmet, T. Structural Features on the Substrate-binding Surface of Fungal Lytic Polysaccharide Monooxygenases Determine Their Oxidative Regioselectivity. Biotechnol. J. 2019, 14, e1800211. 66. Forsberg, Z.; Bissaro, B.; Gullesen, J.; Dalhus, B.; Vaaje-Kolstad, G.; Eijsink, V. G. H. Structural Determinants of Bacterial Lytic Polysaccharide Monooxygenase Functionality. J. Biol. Chem. 2018, 293, 1397-1412. 67. Book, A. J.; Yennamalli, R. M.; Takasuka, T. E.; Currie, C. R.; Phillips, G. N.; Fox, B. G. Evolution of Substrate Specificity in Bacterial AA10 Lytic Polysaccharide Monooxygenases. Biotechnol. Biofuels 2014, 7, 109. 68. Li, X.; Beeson, W. T.; Phillips, C. M.; Marletta, M. A.; Cate, J. H. D. Structural Basis for Substrate Targeting and Catalysis by Fungal Polysaccharide Monooxygenases. Structure 2012, 20, 1051-1061.
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69. Beeson, W. T.; Phillips, C. M.; Cate, J. H. D.; Marletta, M. A. Oxidative Cleavage of Cellulose by Fungal Copper-dependent Polysaccharide Monooxygenases. J. Am. Chem. Soc. 2012, 134, 890-892. 70. Kim, S.; Stahlberg, J.; Sandgren, M.; Paton, R. S.; Beckham, G. T. Quantum Mechanical Calculations Suggest That Lytic Polysaccharide Monooxygenases Use a Copper-oxyl, Oxygenrebound Mechanism. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 149-154. 71. O'Dell, W. B.; Agarwal, P. K.; Meilleur, F. Oxygen Activation at the Active Site of a Fungal Lytic Polysaccharide Monooxygenase. Angew. Chem. Int. Ed. 2017, 56, 767-770. 72. Bacik, J. P.; Mekasha, S.; Forsberg, Z.; Kovalevsky, A. Y.; Vaaje-Kolstad, G.; Eijsink, V. G. H.; Nix, J. C.; Coates, L.; Cuneo, M. J.; Unkefer, C. J.; Chen, J. C. H. Neutron and Atomic Resolution X-ray Structures of a Lytic Polysaccharide Monooxygenase Reveal Copper-mediated Dioxygen Binding and Evidence for N-terminal Deprotonation. Biochemistry 2017, 56, 25292532. 73. Hedegård, E. D.; Ryde, U. Targeting the Reactive Intermediate in Polysaccharide Monooxygenases. J. Biol. Inorg. Chem. 2017, 22, 1029-1037. 74. Hedegård, E. D.; Ryde, U, Molecular Mechanism of Lytic Polysaccharide Monooxygenases. Chem. Sci. 2018, 9, 3866-3880. 75. Hemsworth, G. R.; Davies, G. J.; Walton, P. H. Recent Insights into Copper-containing Lytic Polysaccharide Mono-oxygenases. Curr. Opin. Struct. Biol. 2013, 23, 660-668. 76. Donoghue, P. J.; Tehranchi, J.; Cramer, C. J.; Sarangi, R.; Solomon, E. I.; Tolman, W. B. Rapid C-H Bond Activation by a Monocopper(III)-hydroxide Complex. J. Am. Chem. Soc. 2011, 133, 17602-17605. 77. Himes, R. A.; Karlin, K. D. Copper-dioxygen Complex Mediated C-H Bond Oxygenation: Relevance for Particulate Methane Monooxygenase (pMMO). Curr. Opin. Chem. Biol. 2009, 13, 119-131. 78. Bertini, L.; Breglia, R.; Lambrughi, M.; Fantucci, P.; De Gioia, L.; Borsari, M.; Sola, M.; Bortolotti, C. A.; Bruschi, M. Catalytic Mechanism of Fungal Lytic Polysaccharide Monooxygenases Investigated by First-principles Calculations. Inorg. Chem. 2018, 57, 86-97. 79. Wang, B. J.; Johnston, E. M.; Li, P. F.; Shaik, S.; Davies, G. J.; Walton, P. H.; Rovira, C. QM/MM Studies into the H2O2-dependent Activity of Lytic Polysaccharide Monooxygenases: Evidence for the Formation of a Caged Hydroxyl Radical Intermediate. ACS Catal. 2018, 8, 1346-1351. 80. Kjaergaard, C. H.; Qayyum, M. F.; Wong, S. D.; Xu, F.; Hemsworth, G. R.; Walton, D. J.; Young, N. A.; Davies, G. J.; Walton, P. H.; Johansen, K. S.; Hodgson, K. O.; Hedman, B.; Solomon, E. I. Spectroscopic and Computational Insight into the Activation of O2 by the Mononuclear Cu Center in Polysaccharide Monooxygenases. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 8797-8802. 81. Borisova, A. S.; Isaksen, T.; Dimarogona, M.; Kognole, A. A.; Mathiesen, G.; Várnai, A.; Røhr, A. K.; Payne, C. M.; Sørlie, M.; Sandgren, M.; Eijsink, V. G. H. Structural and Functional Characterization of a Lytic Polysaccharide Monooxygenase with Broad Substrate Specificity. J. Biol. Chem. 2015, 290, 22955-22969. 82. Hangasky, J. A.; Marletta, M. A. A Random-sequential Kinetic Mechanism for Polysaccharide Monooxygenases. Biochemistry 2018, 57, 3191-3199. 83. Cragg, S. M.; Beckham, G. T.; Bruce, N. C.; Bugg, T. D. H.; Distel, D. L.; Dupree, P.; Etxabe, A. G.; Goodell, B. S.; Jellison, J.; McGeehan, J. E.; McQueen-Mason, S. J.; Schnorr, K.;
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Walton, P. H.; Watts, J. E. M.; Zimmer, M. Lignocellulose Degradation Mechanisms Across the Tree of Life. Curr. Opin. Chem. Biol. 2015, 29, 108-119. 84. Loose, J. S. M.; Forsberg, Z.; Fraaije, M. W.; Eijsink, V. G. H.; Vaaje-Kolstad, G. A Rapid Quantitative Activity Assay Shows That the Vibrio cholerae Colonization Factor GbpA Is an Active Lytic Polysaccharide Monooxygenase. FEBS Lett. 2014, 588, 3435-3440. 85. Hofrichter, M.; Ullrich, R. Oxidations Catalyzed by Fungal Peroxygenases. Curr. Opin. Chem. Biol. 2014, 19, 116-125. 86. Matthews, S.; Belcher, J. D.; Tee, K. L.; Girvan, H. M.; McLean, K. J.; Rigby, S. E. J.; Levy, C. W.; Leys, D.; Parker, D. A.; Blankley, R. T.; Munro, A. W. Catalytic Determinants of Alkene Production by the Cytochrome P450 Peroxygenase OleTJE. J. Biol. Chem. 2017, 292, 5128-5143. 87. Shoji, O.; Watanabe, Y. Peroxygenase Reactions Catalyzed by Cytochromes P450. J Biol Inorg. Chem. 2014, 19, 529-539. 88. Hrycay, E. G.; Bandiera, S. M. Monooxygenase, Peroxidase and Peroxygenase Properties and Reaction Mechanisms of Cytochrome P450 Enzymes. In Monooxygenase, Peroxidase and Peroxygenase Properties and Mechanisms of Cytochrome P450, Hrycay, E. G.; Bandiera, S. M., Eds.; Springer International Publishing: Cham, 2015; Vol. 851, pp 1-61. 89. Krauser, J. A.; Guengerich, F. P. Cytochrome P450 3A4-catalyzed Testosterone 6βHydroxylation Stereochemistry, Kinetic Deuterium Isotope Effects, and Rate-limiting Steps. J. Biol. Chem. 2005, 280, 19496-19506. 90. Fukami, T.; Katoh, M.; Yamazaki, H.; Yokoi, T.; Nakajima, M. Human Cytochrome P450 2A13 Efficiently Metabolizes Chemicals in Air Pollutants: Naphthalene, Styrene, and Toluene. Chem. Res. Toxicol. 2008, 21, 720-725. 91. Kumar, S.; Liu, H.; Halpert, J. R. Engineering of Cytochrome P450 3A4 for Enhanced Peroxide-mediated Substrate Oxidation Using Directed Evolution and Site-directed Mutagenesis. Drug Metab. Dispos. 2006, 34, 1958-1965. 92. Joo, H.; Lin, Z. L.; Arnold, F. H. Laboratory Evolution of Peroxide-mediated Cytochrome P450 Hydroxylation. Nature 1999, 399, 670-673. 93. Girhard, M.; Kunigk, E.; Tihovsky, S.; Shumyantseva, V. V.; Urlacher, V. B. Lightdriven Biocatalysis with Cytochrome P450 Peroxygenases. Biotechnol. Appl. Biochem. 2013, 60, 111-118. 94. Shoji, O.; Fujishiro, T.; Nakajima, H.; Kim, M.; Nagano, S.; Shiro, Y.; Watanabe, Y. Hydrogen Peroxide Dependent Monooxygenations by Tricking the Substrate Recognition of Cytochrome P450BSβ. Angew. Chem. Int. Ed. 2007, 46, 3656-3659. 95. Girhard, M.; Schuster, S.; Dietrich, M.; Dürre, P.; Urlacher, V. B. Cytochrome P450 Monooxygenase from Clostridium acetobutylicum: a New α-Fatty Acid Hydroxylase. Biochem. Biophys. Res. Commun. 2007, 362, 114-119. 96. Nordblom, G. D.; White, R. E.; Coon, M. J. Studies on Hydroperoxide-dependent Substrate Hydroxylation by Purified Liver Microsomal Cytochrome P-450. Arch. Biochem. Biophys. 1976, 175, 524-533. 97. Li, Q.-S.; Ogawa, J.; Shimizu, S. Critical Role of the Residue Size at Position 87 in H2O2-dependent Substrate Hydroxylation Activity and H2O2 Inactivation of Cytochrome P450BM-3. Biochem. Biophys. Res. Commun. 2001, 280, 1258-1261. 98. Cirino, P. C.; Arnold, F. H. A Self-sufficient Peroxide-driven Hydroxylation Biocatalyst. Angew. Chem. Int .Ed. 2003, 42, 3299-3301.
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99. Fujishiro, T.; Shoji, O.; Nagano, S.; Sugimoto, H.; Shiro, Y.; Watanabe, Y. Crystal Structure of H2O2-dependent Cytochrome P450SPα with Its Bound Fatty Acid Substrate: Insight into the Regioselective Hydroxylation of Fatty Acids at the Alpha Position. J. Biol. Chem. 2011, 286, 29941-29950. 100. Girvan, H. M.; Poddar, H.; McLean, K. J.; Nelson, D. R.; Hollywood, K. A.; Levy, C. W.; Leys, D.; Munro, A. W. Structural and Catalytic Properties of the Peroxygenase P450 Enzyme CYP152K6 from Bacillus methanolicus. J. Inorg. Biochem. 2018, 188, 18-28. 101. Matsunaga, I.; Yamada, M.; Kusunose, E.; Miki, T.; Ichihara, K. Further Characterization of Hydrogen Peroxide-dependent Fatty Acid α-Hydroxylase from Sphingomonas paucimobilis. J. Biochem. 1998, 124, 105-110. 102. Lee, D.-S.; Yamada, A.; Sugimoto, H.; Matsunaga, I.; Ogura, H.; Ichihara, K.; Adachi, S.-i.; Park, S.-Y.; Shiro, Y. Substrate Recognition and Molecular Mechanism of Fatty Acid Hydroxylation by Cytochrome P450 from Bacillus subtilis: Crystallographic, Spectroscopic, and Mutational Studies. J. Biol. Chem. 2003, 278, 9761-9767. 103. Matsunaga, I.; Ueda, A.; Sumimoto, T.; Ichihara, K.; Ayata, M.; Ogura, H. Site-directed Mutagenesis of the Putative Distal Helix of Peroxygenase Cytochrome P450. Arch. Biochem. Biophys. 2001, 394, 45-53. 104. Span, E. A.; Suess, D. L. M.; Deller, M. C.; Britt, R. D.; Marletta, M. A. The Role of the Secondary Coordination Sphere in a Fungal Polysaccharide Monooxygenase. ACS Chem. Biol. 2017, 12, 1095-1103. 105. Dimarogona, M.; Topakas, E.; Olsson, L.; Christakopoulos, P. Lignin Boosts the Cellulase Performance of a GH-61 Enzyme from Sporotrichum thermophile. Bioresour. Technol. 2012, 110, 480-487. 106. Westereng, B.; Cannella, D.; Agger, J. W.; Jørgensen, H.; Andersen, M. L.; Eijsink, V. G. H.; Felby, C. Enzymatic Cellulose Oxidation is Linked to Lignin by Long-range Electron Transfer. Sci. Rep. 2015, 5, 18561. 107. Muraleedharan, M. N.; Zouraris, D.; Karantonis, A.; Topakas, E.; Sandgren, M.; Rova, U.; Christakopoulos, P.; Karnaouri, A. Effect of Lignin Fractions Isolated from Different Biomass Sources on Cellulose Oxidation by Fungal Lytic Polysaccharide Monooxygenases. Biotechnol. Biofuels 2018, 11, 296. 108. Brenelli, L.; Squina, F. M.; Felby, C.; Cannella, D. Laccase-derived Lignin Compounds Boost Cellulose Oxidative Enzymes AA9. Biotechnol. Biofuels 2018, 11, 10. 109. Várnai, A.; Umezawa, K.; Yoshida, M.; Eijsink, V. G. H. The Pyrroloquinoline-quinonedependent Pyranose Dehydrogenase from Coprinopsis cinerea Drives Lytic Polysaccharide Monooxygenase Action. Appl. Environ. Microbiol. 2018, 84, e00156-18. 110. Tan, T. C.; Kracher, D.; Gandini, R.; Sygmund, C.; Kittl, R.; Haltrich, D.; Hallberg, B. M.; Ludwig, R.; Divne, C. Structural Basis for Cellobiose Dehydrogenase Action During Oxidative Cellulose Degradation. Nat. Commun. 2015, 6, 7542. 111. Courtade, G.; Wimmer, R.; Røhr, A. K.; Preims, M.; Felice, A. K. G.; Dimarogona, M.; Vaaje-Kolstad, G.; Sørlie, M.; Sandgren, M.; Ludwig, R.; Eijsink, V. G. H.; Aachmann, F. L. Interactions of a Fungal Lytic Polysaccharide Monooxygenase with β-Glucan Substrates and Cellobiose Dehydrogenase. Proc. Natl. Acad. Sci. U. S. A. 2016, 113, 5922-5927. 112. Loose, J. S. M.; Forsberg, Z.; Kracher, D.; Scheiblbrandner, S.; Ludwig, R.; Eijsink, V. G. H.; Vaaje-Kolstad, G. Activation of Bacterial Lytic Polysaccharide Monooxygenases with Cellobiose Dehydrogenase. Protein Sci. 2016, 25, 2175-2186.
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113. Zamocky, M.; Ludwig, R.; Peterbauer, C.; Hallberg, B. M.; Divne, C.; Nicholls, P.; Haltrich, D. Cellobiose Dehydrogenase - a Flavocytochrome from Wood-degrading, Phytopathogenic and Saprotropic Fungi. Curr. Protein Pept. Sci. 2006, 7, 255-280. 114. Harada, H.; Onoda, A.; Uchihashi, T.; Watanabe, H.; Sunagawa, N.; Samejima, M.; Igarashi, K.; Hayashi, T. Interdomain Flip-flop Motion Visualized in Flavocytochrome Cellobiose Dehydrogenase Using High-speed Atomic Force Microscopy During Catalysis. Chem. Sci. 2017, 8, 6561-6565. 115. Henriksson, G.; Johansson, G.; Pettersson, G. A Critical Review of Cellobiose Dehydrogenases. J. Biotechnol. 2000, 78, 93-113. 116. Igarashi, K.; Momohara, I.; Nishino, T.; Samejima, M. Kinetics of Inter-domain Electron Transfer in Flavocytochrome Cellobiose Dehydrogenase from the White-rot Fungus Phanerochaete chrysosporium. Biochem. J. 2002, 365, 521-526. 117. Mason, M. G.; Nicholls, P.; Wilson, M. T. Rotting by Radicals - the Role of Cellobiose Oxidoreductase? Biochem. Soc. Trans. 2003, 31, 1335-1336. 118. Sygmund, C.; Santner, P.; Krondorfer, I.; Peterbauer, C. K.; Alcalde, M.; Nyanhongo, G. S.; Guebitz, G. M.; Ludwig, R. Semi-rational Engineering of Cellobiose Dehydrogenase for Improved Hydrogen Peroxide Production. Microb. Cell Fact. 2013, 12, 38. 119. Matsumura, H.; Umezawa, K.; Takeda, K.; Sugimoto, N.; Ishida, T.; Samejima, M.; Ohno, H.; Yoshida, M.; Igarashi, K.; Nakamura, N. Discovery of a Eukaryotic Pyrroloquinoline Quinone-dependent Oxidoreductase Belonging to a New Auxiliary Activity Family in the Database of Carbohydrate-active Enzymes. PLoS One 2014, 9, e104851. 120. Cannella, D.; Möllers, K. B.; Frigaard, N. U.; Jensen, P. E.; Bjerrum, M. J.; Johansen, K. S.; Felby, C. Light-driven Oxidation of Polysaccharides by Photosynthetic Pigments and a Metalloenzyme. Nat. Commun. 2016, 7, 11134. 121. Bissaro, B.; Forsberg, Z.; Ni, Y.; Hollmann, F.; Vaaje-Kolstad, G.; Eijsink, V. G. H. Fueling Biomass-degrading Oxidative Enzymes by Light-driven Water Oxidation. Green Chem. 2016, 18, 5357-5366. 122. Möllers, K. B.; Mikkelsen, H.; Simonsen, T. I.; Cannella, D.; Johansen, K. S.; Bjerrum, M. J.; Felby, C. On the Formation and Role of Reactive Oxygen Species in Light-driven LPMO Oxidation of Phosphoric Acid Swollen Cellulose. Carbohydr. Res. 2017, 448, 182-186. 123. Frommhagen, M.; Westphal, A. H.; Hilgers, R.; Koetsier, M. J.; Hinz, S. W. A.; Visser, J.; Gruppen, H.; van Berkel, W. J. H.; Kabel, M. A. Quantification of the Catalytic Performance of C1-cellulose-specific Lytic Polysaccharide Monooxygenases. Appl. Microbiol. Biotechnol. 2018, 102, 1281-1295. 124. Gusakov, A. V.; Bulakhov, A. G.; Demin, I. N.; Sinitsyn, A. P. Monitoring of Reactions Catalyzed by Lytic Polysaccharide Monooxygenases Using Highly-sensitive Fluorimetric Assay of the Oxygen Consumption Rate. Carbohydr. Res. 2017, 452, 156-161. 125. Hegnar, O. A.; Petrović, D. M.; Bissaro, B.; Alfredsen, G.; Várnai, A.; Eijsink, V. G. H. pH-Dependent Relationship between Catalytic Activity and Hydrogen Peroxide Production Shown via Characterization of a Lytic Polysaccharide Monooxygenase from Gloeophyllum trabeum. Appl. Environ. Microbiol. 2019, 85, e02612-18. 126. Simmons, T. J.; Frandsen, K. E. H.; Ciano, L.; Tryfona, T.; Lenfant, N.; Poulsen, J. C.; Wilson, L. F. L.; Tandrup, T.; Tovborg, M.; Schnorr, K.; Johansen, K. S.; Henrissat, B.; Walton, P. H.; Lo Leggio, L.; Dupree, P. Structural and Electronic Determinants of Lytic Polysaccharide Monooxygenase Reactivity on Polysaccharide Substrates. Nat. Commun. 2017, 8, 1064.
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127. Kracher, D.; Andlar, M.; Furtmuller, P. G.; Ludwig, R. Active-site Copper Reduction Promotes Substrate Binding of Fungal Lytic Polysaccharide Monooxygenase and Reduces Stability. J. Biol. Chem. 2018, 293, 1676-1687. 128. Loose, J. S. M.; Arntzen, M. O.; Bissaro, B.; Ludwig, R.; Eijsink, V. G. H.; VaajeKolstad, G. Multipoint Precision Binding of Substrate Protects Lytic Polysaccharide Monooxygenases from Self-destructive Off-pathway Processes. Biochemistry 2018, 57, 41144124. 129. Mutahir, Z.; Mekasha, S.; Loose, J. S. M.; Abbas, F.; Vaaje-Kolstad, G.; Eijsink, V. G. H.; Forsberg, Z. Characterization and Synergistic Action of a Tetra-modular Lytic Polysaccharide Monooxygenase from Bacillus cereus. FEBS Lett. 2018, 592, 2562-2571. 130. Courtade, G.; Forsberg, Z.; Heggset, E. B.; Eijsink, V. G. H.; Aachmann, F. L. The Carbohydrate-binding Module and Linker of a Modular Lytic Polysaccharide Monooxygenase Promote Localized Cellulose Oxidation. J. Biol. Chem. 2018, 293, 13006-13015. 131. Liu, P. H.; Murakami, K.; Seki, T.; He, X. M.; Yeung, S. M.; Kuzuyama, T.; Seto, H.; Liu, H. W. Protein Purification and Function Assignment of the Epoxidase Catalyzing the Formation of Fosfomycin. J. Am. Chem. Soc. 2001, 123, 4619-4620. 132. Wang, C.; Chang, W. C.; Guo, Y. S.; Huang, H.; Peck, S. C.; Pandelia, M. E.; Lin, G. M.; Liu, H. W.; Krebs, C.; Bollinger, J. M. Evidence That the Fosfomycin-producing Epoxidase, HppE, Is a Non-heme-iron Peroxidase. Science 2013, 342, 991-995. 133. Kittl, R.; Kracher, D.; Burgstaller, D.; Haltrich, D.; Ludwig, R. Production of Four Neurospora crassa Lytic Polysaccharide Monooxygenases in Pichia pastoris Monitored by a Fluorimetric Assay. Biotechnol. Biofuels 2012, 5, 79. 134. Galbe, M.; Zacchi, G. Pretreatment: the Key to Efficient Utilization of Lignocellulosic Materials. Biomass Bioenergy 2012, 46, 70-78. 135. Teeri, T. T.; Koivula, A.; Linder, M.; Wohlfahrt, G.; Divne, C.; Jones, T. A. Trichoderma reesei Cellobiohydrolases: Why So Efficient on Crystalline Cellulose? Biochem. Soc. Trans. 1998, 26, 173-178. 136. Penttila, P. A.; Várnai, A.; Pere, J.; Tammelin, T.; Salmen, L.; Siika-aho, M.; Viikari, L.; Serimaa, R. Xylan as Limiting Factor in Enzymatic Hydrolysis of Nanocellulose. Bioresour. Technol. 2013, 129, 135-141. 137. Yu, L.; Lyczakowski, J. J.; Pereira, C. S.; Kotake, T.; Yu, X. L.; Li, A.; Mogelsvang, S.; Skaf, M. S.; Dupree, P. The Patterned Structure of Galactoglucomannan Suggests It May Bind to Cellulose in Seed Mucilage. Plant Physiol. 2018, 178, 1011-1026. 138. Busse-Wicher, M.; Gomes, T. C. F.; Tryfona, T.; Nikolovski, N.; Stott, K.; Grantham, N. J.; Bolam, D. N.; Skaf, M. S.; Dupree, P. The Pattern of Xylan Acetylation Suggests Xylan May Interact with Cellulose Microfibrils as a Twofold Helical Screw in the Secondary Plant Cell Wall of Arabidopsis thaliana. Plant J. 2014, 79, 492-506. 139. Busse-Wicher, M.; Grantham, N. J.; Lyczakowski, J. J.; Nikolovski, N.; Dupree, P. Xylan Decoration Patterns and the Plant Secondary Cell Wall Molecular Architecture. Biochem. Soc. Trans. 2016, 44, 74-78. 140. Berlin, A.; Maximenko, V.; Gilkes, N.; Saddler, J. Optimization of Enzyme Complexes for Lignocellulose Hydrolysis. Biotechnol. Bioeng. 2007, 97, 287-96. 141. Selig, M. J.; Knoshaug, E. P.; Adney, W. S.; Himmel, M. E.; Decker, S. R. Synergistic Enhancement of Cellobiohydrolase Performance on Pretreated Corn Stover by Addition of Xylanase and Esterase Activities. Bioresour. Technol. 2008, 99, 4997-5005.
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142. Várnai, A.; Huikko, L.; Pere, J.; Siika-Aho, M.; Viikari, L. Synergistic Action of Xylanase and Mannanase Improves the Total Hydrolysis of Softwood. Bioresour. Technol. 2011, 102, 9096-104. 143. Hu, J. G.; Arantes, V.; Saddler, J. N. The Enhancement of Enzymatic Hydrolysis of Lignocellulosic Substrates by the Addition of Accessory Enzymes Such as Xylanase: Is It an Additive or Synergistic Effect? Biotechnol. Biofuels 2011, 4, 36. 144. Chylenski, P.; Forsberg, Z.; Stahlberg, J.; Várnai, A.; Lersch, M.; Bengtsson, O.; Saebo, S.; Horn, S. J.; Eijsink, V. G. Development of Minimal Enzyme Cocktails for Hydrolysis of Sulfite-pulped Lignocellulosic Biomass. J. Biotechnol. 2017, 246, 16-23. 145. Hu, J. G.; Arantes, V.; Pribowo, A.; Saddler, J. N. The Synergistic Action of Accessory Enzymes Enhances the Hydrolytic Potential of a "Cellulase Mixture" But Is Highly Substrate Specific. Biotechnol. Biofuels 2013, 6, 112. 146. Kojima, Y.; Várnai, A.; Ishida, T.; Sunagawa, N.; Petrovic, D. M.; Igarashi, K.; Jellison, J.; Goodell, B.; Alfredsen, G.; Westereng, B.; Eijsink, V. G. H.; Yoshida, M. A Lytic Polysaccharide Monooxygenase with Broad Xyloglucan Specificity from the Brown-rot Fungus Gloeophyllum trabeum and Its Action on Cellulose-xyloglucan Complexes. Appl. Environ. Microbiol. 2016, 82, 6557-6572. 147. Grantham, N. J.; Wurman-Rodrich, J.; Terrett, O. M.; Lyczakowski, J. J.; Stott, K.; Iuga, D.; Simmons, T. J.; Durand-Tardif, M.; Brown, S. P.; Dupree, R.; Busse-Wicher, M.; Dupree, P. An Even Pattern of Xylan Substitution Is Critical for Interaction with Cellulose in Plant Cell Walls. Nat. Plants 2017, 3, 859-865. 148. Humbird, D.; Davis, R.; Tao, L.; Kinchin, C.; Hsu, D.; Aden, A.; Schoen, P.; Lukas, J.; Olthof, B.; Worley, M.; Sexton, D.; Dudgeon, D. Process Design and Economics for Biochemical Conversion of Lignocellulosic Biomass to Ethanol. Dilute-acid Pretreatment and Enzymatic Hydrolysis of Corn Stover; Technical Report for National Renewable Energy Laboratory: Golden, CO, May 2011:NREL/TP-5100-47764. 149. Adav, S. S.; Chao, L. T.; Sze, S. K. Quantitative Secretomic Analysis of Trichoderma reesei Strains Reveals Enzymatic Composition for Lignocellulosic Biomass Degradation. Mol. Cell. Proteomics 2012, 11, M111.012419. 150. Martinez, D.; Berka, R. M.; Henrissat, B.; Saloheimo, M.; Arvas, M.; Baker, S. E.; Chapman, J.; Chertkov, O.; Coutinho, P. M.; Cullen, D.; Danchin, E. G. J.; Grigoriev, I. V.; Harris, P.; Jackson, M.; Kubicek, C. P.; Han, C. S.; Ho, I.; Larrondo, L. F.; de Leon, A. L.; Magnuson, J. K.; Merino, S.; Misra, M.; Nelson, B.; Putnam, N.; Robbertse, B.; Salamov, A. A.; Schmoll, M.; Terry, A.; Thayer, N.; Westerholm-Parvinen, A.; Schoch, C. L.; Yao, J.; Barbote, R.; Nelson, M. A.; Detter, C.; Bruce, D.; Kuske, C. R.; Xie, G.; Richardson, P.; Rokhsar, D. S.; Lucas, S. M.; Rubin, E. M.; Dunn-Coleman, N.; Ward, M.; Brettin, T. S. Genome Sequencing and Analysis of the Biomass-degrading Fungus Trichoderma reesei (syn. Hypocrea jecorina). Nat. Biotechnol. 2008, 26, 553-560. 151. Scott, B. R.; Huang, H. Z.; Frickman, J.; Halvorsen, R.; Johansen, K. S. Catalase Improves Saccharification of Lignocellulose by Reducing Lytic Polysaccharide Monooxygenaseassociated Enzyme Inactivation. Biotechnol. Lett. 2016, 38, 425-434. 152. Du, J.; Song, W. X.; Zhang, X.; Zhao, J.; Liu, G. D.; Qu, Y. B. Differential Reinforcement of Enzymatic Hydrolysis by Adding Chemicals and Accessory Proteins to High Solid Loading Substrates with Different Pretreatments. Bioprocess Biosyst. Eng. 2018, 41, 11531163.
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153. Kim, I. J.; Youn, H. J.; Kim, K. H, Synergism of an Auxiliary Activity 9 (AA9) from Chaetomium globosum with Xylanase on the Hydrolysis of Xylan and Lignocellulose. Process Biochem. 2016, 51, 1445-1451. 154. Sanhueza, C.; Carvajal, G.; Soto-Aguilar, J.; Lienqueo, M. E.; Salazar, O. The Effect of a Lytic Polysaccharide Monooxygenase and a Xylanase from Gloeophyllum trabeum on the Enzymatic Hydrolysis of Lignocellulosic Residues Using a Commercial Cellulase. Enzyme Microb. Tech. 2018, 113, 75-82. 155. Peciulyte, A.; Samuelsson, L.; Olsson, L.; McFarland, K. C.; Frickmann, J.; Østergård, L.; Halvorsen, R.; Scott, B. R.; Johansen, K. S. Redox Processes Acidify and Decarboxylate Steam-pretreated Lignocellulosic Biomass and Are Modulated by LPMO and Catalase. Biotechnol. Biofuels 2018, 11, 165. 156. Garcia-Ochoa, F.; Gomez, E.; Santos, V. E.; Merchuk, J. C. Oxygen Uptake Rate in Microbial Processes: an Overview. Biochem. Eng. J. 2010, 49, 289-307. 157. Sheldon, R. A. The Road to Biorenewables: Carbohydrates to Commodity Chemicals. ACS Sustain. Chem. Eng. 2018, 6, 4464-4480. 158. Wyman, C. E. Ethanol Production from Lignocellulosic Biomass: Overview. In Handbook on Bioethanol, Wyman, C. E., Ed.; Taylor & Francis: Washington, DC, 1996; pp 118. 159. Müller, G.; Kalyani, D. C.; Horn, S. J. LPMOs in Cellulase Mixtures Affect Fermentation Strategies for Lactic Acid Production from Lignocellulosic Biomass. Biotechnol. Bioeng. 2017, 114, 552-559. 160. de Oliveira, R. A.; Komesu, A.; Rossell, C. E. V.; Maciel, R. Challenges and Opportunities in Lactic Acid Bioprocess Design – from Economic to Production Aspects. Biochem. Eng. J. 2018, 133, 219-239. 161. Wingren, A.; Galbe, M.; Zacchi, G. Techno-economic Evaluation of Producing Ethanol from Softwood: Comparison of SSF and SHF and Identification of Bottlenecks. Biotechnol. Prog. 2003, 19, 1109-1117. 162. Cannella, D.; Jørgensen, H. Do New Cellulolytic Enzyme Preparations Affect the Industrial Strategies for High Solids Lignocellulosic Ethanol Production? Biotechnol. Bioeng. 2014, 111, 59-68. 163. Berlemont, R. Distribution and Diversity of Enzymes for Polysaccharide Degradation in Fungi. Sci. Rep. 2017, 7, 222. 164. Lenfant, N.; Hainaut, M.; Terrapon, N.; Drula, E.; Lombard, V.; Henrissat, B. A Bioinformatics Analysis of 3400 Lytic Polysaccharide Oxidases from Family AA9. Carbohydr. Res. 2017, 448, 166-174. 165. Wong, E.; Vaaje-Kolstad, G.; Ghosh, A.; Hurtado-Guerrero, R.; Konarev, P. V.; Ibrahim, A. F. M.; Svergun, D. I.; Eijsink, V. G. H.; Chatterjee, N. S.; van Aalten, D. M. F. The Vibrio cholerae Colonization Factor GbpA Possesses a Modular Structure That Governs Binding to Different Host Surfaces. PLoS Pathog. 2012, 8, e1002373. 166. Paspaliari, D. K.; Loose, J. S. M.; Larsen, M. H.; Vaaje-Kolstad, G. Listeria monocytogenes Has a Functional Chitinolytic System and an Active Lytic Polysaccharide Monooxygenase. FEBS J. 2015, 282, 921-936. 167. Agostoni, M.; Hangasky, J. A.; Marletta, M. A. Physiological and Molecular Understanding of Bacterial Polysaccharide Monooxygenases. Microbiol. Mol. Biol. Rev. 2017, 81, e00015-17.
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