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# Expanding the catalytic landscape of metalloenzymes with lytic polysaccharide monooxygenases

Alessia Munzone, Vincent G H Eijsink, Jean-Guy Berrin, Bastien Bissaro

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Alessia Munzone, Vincent G H Eijsink, Jean-Guy Berrin, Bastien Bissaro. Expanding the catalytic landscape of metalloenzymes with lytic polysaccharide monooxygenases. Nature Reviews Chemistry, 2024, 10.1038/s41570-023-00565-z. hal-04386433

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Submitted on 29 Jan 2025

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# Expanding the catalytic landscape of metalloenzymes with Lytic Polysaccharide Monooxygenases

Alessia Munzone<fnref n="1" />, Vincent G.H. Eijsink<fnref n="2" />, Jean-Guy Berrin<fnref n="1" /> and Bastien Bissaro<fnref n="1" />*

## Abstract

Lytic Polysaccharide Monooxygenases (LPMOs) play an essential role in global carbon cycle, industrial biomass processing, and microbial pathogenicity by catalysing the oxidative cleavage of recalcitrant polysaccharides. Despite initially being considered monooxygenases, experimental and theoretical studies show that LPMOs are essentially peroxygenases, employing a single copper ion and $H_2O_2$ for C-H bond oxygenation. Here, we examine LPMO catalysis, emphasizing key studies that have shaped our comprehension of their function, and address side and competing reactions that have partially obscured our understanding. Then, we compare this novel copper-peroxygenase reaction with reactions catalyzed by heme iron-enzymes, highlighting the different chemistries at play. We conclude by addressing some open questions surrounding LPMO catalysis, including the significance of peroxygenase and monooxygenase reactions in biological contexts, how LPMOs modulate copper site reactivity, and potential protective mechanisms against oxidative damage.

## Introduction

The cleavage of glycosidic bonds is at the heart of the global carbon cycle. This seemingly simple molecular event, through its impact on abundant biopolymers such as cellulose, chitin and starch, is a main driver of many biological processes. In 1953, Daniel Koshland laid the basis of our current understanding of the hydrolytic cleavage of O-glycosidic bonds by glycoside hydrolases (GHs)<fnref n="1" />, which has been enriched during the subsequent decades by various researchers<fnref n="2" />⁻⁶. Since then, the era of (glyco)genomics has revealed that this hydrolytic strategy for polysaccharide degradation has been adopted across the entire tree of life.

<footnote n="1">¹ INRAE, Aix Marseille Univ., UMR1163 Biodiversité et Biotechnologie Fongiques, 13009 Marseille, France</footnote>

<footnote n="2">² Faculty of Chemistry, Biotechnology, and Food Science, The Norwegian University of Life Sciences (NMBU), 1432 Ås, Norway</footnote>

<footnote>*Correspondence to: bastien.bissaro@inrae.fr</footnote>
For decades, the depolymerization of polysaccharides was thought to be strictly dependent on the synergistic action of hydrolytic enzymes (and, to some extent, polysaccharide lyases), until lytic polysaccharide monooxygenases (LPMOs) were discovered<sup>7,8</sup> (Fig. 1). LPMOs are equipped with a solvent-exposed mono-nuclear copper center<sup>9-11</sup> able to generate a highly reactive oxygen species that can activate a C-H bond. Such species are not accessible to Nature via the chemistry afforded by natural amino acids and can only be generated by co-factor containing enzymes, primarily heme iron enzymes such as the cytochrome P450s or the so-called unspecific peroxygenases<sup>12,13</sup>. The powerful oxidative chemistry that can be catalyzed by LPMOs is combined with a rather rare active site topology, located on a usually rather flat, solvent-exposed and extended substrate binding surface, allowing these enzymes to act on the surfaces of biomaterials such as cellulose fibrils. The disruptive action of LPMOs generates new glycosidic chain ends tractable by depolymerizing GHs, thereby boosting overall biomass turnover. The discovery of LPMO oxidative activity has profoundly modified our vision of biomass conversion, whether it is at a fundamental level or for industrial applications (such as in biorefineries)<sup>14,15</sup>.

While originally studied in the context of biomass conversion, LPMOs have now been identified in many bacterial and fungal communities and also in viruses, plants and insects, suggesting a wide range of potential biological functions<sup>16</sup>. In this context, intense research on new physiological roles has led to the discovery of LPMOs and LPMO-like proteins with unclear biological functions and for which the natural substrate remains unknown<sup>16-18</sup>. The increase in newly identified LPMOs is reflected by their distribution (as of August 2023) in eight families of the auxiliary activity (AA) class in the carbohydrate-active enzymes (CAZy) database (www.cazy.org)<sup>19</sup>, namely AA9 to AA11 and AA13 to AA17<sup>8,9,11,20-25</sup>.

To cleave glycosidic bonds in polysaccharides, unlike GHs, which mostly rely on acid/base catalysis, LPMOs hydroxylate the C-H bond of the C1 or C4 carbon (Fig. 1). This reaction involves abstraction of a hydrogen atom and formation of a carbon radical on the glucosyl unit. The bond dissociation energy (BDE) to abstract a hydrogen atom from a glycosidic C-H bonds has been calculated (on glucose models) to be between 397 kJ/mol and
423 kJ/mol (for C1-H abstraction) or 435 kJ/mol (for C4-H abstraction)<sup>26-29</sup> – for comparison, the C–H bonds of methane, CH<sub>4</sub>, have an estimated energy of 439 kJ/mol at 25°C<sup>26</sup>. However, within the crystalline lattice of a polysaccharide such as cellulose, it is plausible that these bonds are even stronger than the calculated values. This is because of the physical constraints imposed by the lattice which reduce the flexibility of the glucose ring, and thus the degree of freedom to change the interatomic angles around the carbon radical to stabilize the transition state. Such C–H bonds are thus probably one of the most inert and difficult C–H bonds to break.

By analogy with other copper-enzymes, and on the basis of experiments showing that LPMO catalysis is apparently O<sub>2</sub>-dependent, early reports concluded that LPMOs were monooxygenases, catalyzing the reaction R–H + O<sub>2</sub> + 2e<sup>-</sup> + 2H<sup>+</sup> → R–OH + H<sub>2</sub>O<sup>8,11</sup>. It was noted in multiple early reports that the ability of a mononuclear copper enzyme to catalyze such reaction raises questions as to how electrons are stored and channeled during catalysis. The initial monooxygenase paradigm was challenged by the discovery that LPMOs could catalyze a peroxygenase reaction (R–H + H<sub>2</sub>O<sub>2</sub> → R–OH + H<sub>2</sub>O) at rates that are orders of magnitude higher than those observed in apparent monooxygenase reactions<sup>30</sup>. This paradigm shift has far-reaching consequences with regard to: (i) the use of LPMOs in bioprocesses such as the degradation of lignocellulosic biomass<sup>15</sup>, (ii) the position of LPMOs in the intricate network of redox reactions at play during natural biomass conversion<sup>31</sup>, (iii) the mode of action of LPMOs with emerging novel biological functions such as pathogenesis<sup>16</sup>.

In addition, and of particular relevance for this review, the peroxygenase activity of LPMOs sheds new light on the chemistry of copper-catalyzed oxygenation reactions, hitherto thought to be restricted to O<sub>2</sub>-dependent reactions<sup>32</sup>.

Contextually, looking at the most well-studied oxidoreductases<sup>33-39</sup> using an oxygen species (O<sub>2</sub> or H<sub>2</sub>O<sub>2</sub>) as final electron acceptor, one can distinguish two main type of reactions, that is, oxygenation and oxidation, where oxygen atoms are respectively incorporated or not into the product (Fig. 2). Fig. 2 illustrates that, in terms of the reaction catalyzed, LPMOs
are in the same group as H₂O₂-dependent heme iron peroxygenases. Remarkably, LPMOs
are the first, and, so far, only example of copper-dependent peroxygenases.

In this review, we draw a picture of the mode of action of LPMOs. We provide a step-
by-step overview of their catalytic cycle together with occurring side reactions, highlighting
consensual and still debated aspects, and focusing on the unique catalytic features of these
enzymes. To evaluate the broader relevance of the chemistry involved in the different LPMO-
catalyzed on- and off-pathway reactions, we draw parallels with similar reactions catalyzed by
other (metallo)enzymes. We direct the reader towards other enlightening reviews dealing in
more detail with LPMO copper coordination chemistry⁴⁰ and *in silico* mechanistic studies⁴¹ of
LPMOs.

A step by step guide to LPMO catalysis

Like many metalloenzymes, including peroxidases or unspecific peroxygenases, it is becoming
increasingly clear that LPMOs do not catalyze just one single reaction. While their main
reaction is a hydroxylation (Fig. 3, “PO and MO paths”) leading to cleavage of glycosidic bonds
(Fig. 1C), LPMOs can also catalyze two side reactions that do not lead to oxygen incorporation
in a polysaccharide substrate: (1) an oxidase reaction (Fig. 3, “O path”) (that is, oxidation of a
reductant, generating H₂O₂ from O₂) and a peroxidase reaction (Fig. 3, “P path”) (that is,
oxidation of a (non-carbohydrate) substrate using H₂O₂ as co-substrate). In addition,
depending on the experimental conditions, LPMOs suffer from autocatalytic-inactivation (Fig.
3, “I path”). Finally, enzyme-independent reactions involving reductants that can reduce the
LPMO (Fig. 3, “X path”) affect both LPMO activity, by supplying H₂O₂ for productive reactions,
and stability, by potentially generating H₂O₂ levels that are “too” high, promoting autocatalytic
inactivation. These latter side reactions (Fig. 3, I path and X path) are almost unavoidable
owing to the presence of reducing compounds and sporadic amounts of free transition metals
that promote the oxidation of these compounds⁴²,⁴³, as discussed by Eijsink et al.⁴⁴. The
various reactions that may be catalyzed by an LPMO after its reductive activation, in the
presence or absence of the natural substrate, are shown in Fig 3. This network of interrelated
and partially substrate-dependent redox reactions may have obscured the understanding of LPMO chemistry, contributing to some uncertainty and even controversy in the existing literature.

## Main reactions of LPMOs

When productively bound to a carbohydrate substrate, an LPMO catalyzes destabilization and subsequent cleavage of the scissile glycosidic bond by what is generally agreed to be a hydroxylation reaction. Early studies of LPMOs showed that this reaction depends on the activation of molecular oxygen ($O_2$), hence the use of the traditional appellative of ‘monooxygenase’. In a monooxygenase reaction, $O_2$, two externally supplied electrons and two protons are required for each catalytic cycle ($R-H + O_2 + 2e^- + 2H^+ \rightarrow R-OH + H_2O$). The direct use of $O_2$ as co-substrate by LPMOs was first questioned when LPMOs were shown to catalyze peroxygenase reactions$^{30,45}$. Such a reaction implies that the two necessary protons, the reducing equivalents and oxygen atom are supplied simultaneously in the form of $H_2O_2$ ($R-H + H_2O_2 \rightarrow R-OH + H_2O$). While a peroxygenase reaction catalyzed by a monocopper enzyme was unprecedented at the time, this catalytic scenario solved many outstanding questions related to a monooxygenase reaction, including how protons and electrons are delivered in a timely manner to a substrate-bound LPMO (Fig. 4). In this context, it was also noted that the conditions typically used in LPMO reactions are such that $H_2O_2$ will be formed in the reaction mixture as a result of the X- and O-paths depicted in Figure 3. While controversial at first$^{46}$, it is now well established that the apparent monooxygenase activity of LPMOs in most, if not all, cases reflects a peroxygenase reaction$^{31,47-49}$.

Nevertheless, as outlined hereinafter, whether we look at the LPMOs as monooxygenases (MO path) or peroxygenases (PO path), their catalytic power derives from well-defined steps, which we detail here, that to a large extent are the same for the MO and PO paths.
Reduction step

The reduction of the resting state LPMO–Cu(II) to form the reactive Cu(I) enzyme (Fig. 3, “R path”) is the entry point required for any other enzymatic path showed in Fig 3. The one-electron reduction of an LPMO may be seen as a *priming* reaction, which implies that at the end of the catalytic cycle the enzyme is still in the Cu(I) oxidation state. This is clearly the case when LPMOs operate in “peroxygenase conditions”, which involves the use of sub-stoichiometric (relative to product formation) amounts of reducing agent (usually < 100 µM) and controlled supply of H₂O₂ to the reaction. In this respect, multiple studies have shown that a primed LPMO can catalyze between 15 and 20 peroxygenase reactions³⁰,⁵⁰,⁵¹. In contrast, when LPMOs operate in monooxygenase conditions, that is, with dissolved O₂ as source of oxygen atoms, the reducing agent needs to be added in stoichiometric amounts (relative to the reaction products), usually ≥ 1 mM, since it serves both to prime the LPMO and to drive the *in situ* generation of H₂O₂.

LPMOs usually have redox potentials (relative to a standard hydrogen electrode) in the
+200 to +300 mV range<sup>10,52,53</sup> and can be reduced by several types of electron donors: small
organic molecules (such as ascorbic acid, cysteine, glutathione, gallic acid, plant-derived and
fungal phenols<sup>52,54–56</sup>; enzymatic redox partners (such as cellobiose dehydrogenase)<sup>11,52,57–60</sup>,
and also light-harvesting molecules<sup>42,61,62</sup>.

Co-secreted redox enzymes such as cellobiose dehydrogenase (CDH) are sometimes
considered the native reducing partners of AA9 LPMOs in fungal saprotrophs. The beauty of
this enzymatic interplay lies in the necessary reducing equivalents that are gradually generated
upon oxidation of otherwise redox-inert compounds (that is, cellobiose and short cello-
oligomers), which may help ensuring optimal redox control and reaction stability. Some other
flavo-enzymes (belonging to the AA7 family in the CAZy database) were recently revealed to
be oligosaccharide dehydrogenases able to prime AA9 LPMOs and fuel the LPMO reaction
with H2O2<sup>58</sup>, quite similar to cellobiose dehydrogenase<sup>63</sup>. So far, despite an interesting
hypothesis<sup>64</sup>, no obvious natural enzymatic redox partner has been identified for bacterial
AA10 LPMOs<sup>48</sup>.
From a kinetic point of view, the reduction of Cu(II) to Cu(I) is usually not the rate-limiting step in LPMO reactions. Reduction kinetics measured by stopped-flow spectrophotometry have reported electron transfer rates ($k_{obs}$) between 0.9 and 32 s⁻¹ from CDH (at 5 µM) to AA9 and AA10 LPMOs⁵²,⁶⁵, and 420 s⁻¹ from ascorbic acid (at 1 mM) to AA10 LPMOs⁶⁶. These rates are orders of magnitude higher than those typically observed for LPMO catalysis under “monooxygenase conditions” or measured rates for reoxidation of LPMO-Cu(I) by O₂ or H₂O₂ in the absence of substrate, and in the same range as the highest rates described for LPMO peroxygenase reactions. Of note, considering these rates, one might expect that reduction could to some extent be rate-limiting in certain fast LPMO-catalyzed peroxygenase reactions carried out in the presence of low (< 1 mM) amounts of reductant, as has indeed been observed⁵¹,⁶⁷.

The dependence of LPMO activity on the type of reductant has been evident since the first studies in the field, although the cause of this dependency was initially enigmatic. What now seems clear is that the effect of the reductant on overall LPMO activity is not simply related to the reduction of the metal center, but also, and likely primarily⁴²,⁴³,⁴⁸,⁶⁷,⁶⁸, correlates with its impact on the rate of H₂O₂ production in the reaction solution (X path; **Fig. 3**).

## Substrate and co-substrate binding

The structural features that determine substrate specificity of LPMOs are still largely unknown. While it is clear that the carbohydrate-binding modules appended to some LPMOs play a role in the substrate recognition and affinity³⁰,⁶⁹,⁷⁰, the structural properties underlying variation in substrate preferences of LPMO catalytic domains are not very clear. Structural investigations of LPMO-substrate interactions using several approaches such as X-ray crystallography⁷¹,⁷², NMR¹⁰, EPR⁷³⁻⁷⁵, and modelling⁷⁶,⁷⁷, have indicated the roles of both exposed aromatic residues and exposed polar residues in substrate binding. Site-directed mutagenesis studies, including studies done before the nature of the LPMO reaction was discovered⁷,⁷⁸,⁷⁹, have confirmed that many of the surface-exposed residues are important for LPMO functionality, affecting oxidative regioselectivity⁸⁰,⁸¹, and substrate preferences⁸²,⁸³. For
more details on structural aspects of LPMOs, we direct the reader towards excellent reviews by L. Lo Leggio and collaborators<sup>71,72</sup>.

While deconstruction of the crystalline regions of insoluble polysaccharides is considered a major natural role of LPMOs, the number of known LPMOs that can also act on shorter and soluble oligosaccharides is increasing<sup>84-87</sup>. LPMOs acting on soluble substrates are more amenable to experimental characterization and some of these, such as the LPMO from *Neurospora crassa* (*NcAA9C*) and from *Lentinus similis* (*LsAA9A*), have been studied in much detail<sup>73,86</sup>. The binding of both cellulose- and chitin-active LPMOs to polymeric substrates has been studied, with binding affinities in the low micromolar range reported<sup>73,78,88</sup>. Detailed steady-state kinetic studies by Kuusk et al.<sup>89</sup> using radiolabeled chitin nanowiskers yielded a Michaelis-Menten parameter for chitin ($K_m$, chitin) of 0.58 g.L<sup>-1</sup>. Studies with oligomeric substrates tend to show low affinities with $K_d$ and $K_m$ values in the 0.1–2 mM range or even higher<sup>67,73,76,86,90</sup>. Interestingly, Frandsen and colleagues showed that cellohexaose binds very well ($K_d$ = 3.7±0.1 µM) to *LsAA9A* in the presence of 200 mM chloride ions, an observation that was taken to suggest that the presence of a partially charged oxygen species, for which chloride could be an analogue, promotes substrate binding<sup>86</sup>.

Kinetic studies have shown that the LPMO peroxygenase reaction involves formation of a ternary complex, that is, formation of a productive catalytic species only when the substrate and co-substrate are bound to the enzyme, and does not proceed according to a ping-pong mechanism (where one product is formed and released before the second substrate binds)<sup>47,89</sup>. In this respect, one question is whether the priming reduction of Cu(II) occurs before or after substrate binding. While the resting LPMO-Cu(II) form can bind to carbohydrate substrates with good affinity<sup>77</sup>, it has also been shown by Kracher et al. that this affinity is higher for the reduced LPMO-Cu(I) form<sup>88</sup>. Quantum mechanics with molecular mechanics (QM/MM) studies<sup>76</sup> suggest that the simultaneous presence of two Cu(II)-coordinating water molecules and the substrate is energetically unfavorable and that rearrangement of at least one of the water molecules would be needed for *SmAA10A* to bind chitin. Recently, Hedison et al. have shown that substrate binding increases the redox potential of the Cu site, leading to
stabilization of the reduced Cu(I) state and probably preventing side reactions from occurring⁵⁰.
We note that if Cu(II) reduction was to occur subsequently to substrate binding, an electron
transport chain through the protein would be needed as the copper site is no longer exposed
to solvent when the LPMO is bound to an insoluble substrate (Fig. 4). While some authors
have proposed the existence of such an electron delivery channel⁶¹,⁹¹ there is so far no
experimental evidence supporting its existence.

Several studies have shown that substrate binding is not affected by the presence of O₂ or H₂O₂⁸⁸,⁸⁹. However, and of crucial importance, substrate binding increases the affinity for the oxygen-bearing co-substrate, possibly for O₂ and most certainly for H₂O₂.

Considering the possible binding of oxygen, EPR studies have suggested that
substrate-binding would change the copper coordination in a way that would promote reactivity
with O₂<sup>75</sup>. Using steady-state kinetics, Hangasky et al.<sup>92</sup> found that the presence of the
substrate increases the apparent affinity for O₂ (apparent K<sub>M</sub> values in the 200 µM range).
However, in this study abiotic H₂O₂-generating reactions with the reductant (at 2 mM) (Fig.3,
X path) were overlooked. The reaction with O₂ was suggested to follow a random sequential
mechanism, which is compatible with findings from neutron crystallographic studies (done in
the absence of substrate) showing a "pre-bound state", where O₂ is bound to second sphere
residues in the active site but not to copper, which suggests that O₂ binding could precede
reduction<sup>93,94</sup>. This scenario is further supported by stopped-flow kinetics analyzed using the
Marcus theory suggesting an outer-sphere reduction of O₂, that is, a reduction of O₂ by Cu(I)
into O₂<sup>•</sup> prior to binding to Cu(II)<sup>66</sup>.

Competition experiments, single turnover reoxidation experiments and steady-state
kinetic analyses have all shown that reduced LPMOs react much faster with H₂O₂ than with
O₂, both in the absence and presence of substrate³⁰,⁴²,⁴⁷,⁴⁹,⁸⁹, although one early study has
reported relatively fast reoxidation of a fungal AA9 LPMO with O₂ consistent with an alternative
inner-sphere mechanism⁹⁵. Steady-state kinetics (with saturating substrate concentrations)
indicate K_M values for H₂O₂ in the low micromolar range (1–30 µM)⁴⁶,⁸⁹,⁹⁰,⁹⁶. Using different
LPMOs, both Jones et al.<sup>47</sup> and Bissaro et al.<sup>66</sup> found that reoxidation in solution (that is, in the absence of bound substrate) is some three orders of magnitude faster for H<sub>2</sub>O<sub>2</sub>, compared to O<sub>2</sub>. Most importantly, it was shown that the rate of reoxidation by H<sub>2</sub>O<sub>2</sub> (but not by O<sub>2</sub>) increased in the presence of substrate, by an estimated 50-fold<sup>66</sup>.

All in all, substrate and co-substrate binding in the peroxygenase reaction follow an ordered sequential mechanism as suggested by Jones et al.<sup>47</sup>. That is, all the substrates are first bound to the enzyme in a defined order before reaction occurs. Along this line, molecular dynamics (MD) simulations have shown the formation of a tunnel upon binding of SmAA10 to chitin, through which small co-substrates could diffuse to the active site cavity<sup>76</sup> (Fig. 4). Notably, the observed affinities seem tailored to prevent off-pathway reactions that may lead to enzyme damage: reduction makes the enzyme more vulnerable but promotes substrate-binding, while substrate binding promotes the chances of available H<sub>2</sub>O<sub>2</sub> being channeled to productive reactions on the substrate, rather than non-productive reactions in solution. This ordered sequential mechanism is supported by several modelling studies, including MD<sup>74</sup> and QM/MM using the dynamic<sup>95</sup> and static<sup>96</sup> approach.

**C–H bond activation: O<sub>2</sub> versus H<sub>2</sub>O<sub>2</sub> reactivity**

The LPMO-driven hydroxylation of C1–H and/or C4–H carbons in *O*-glycosidic bonds requires the formation of a copper-oxygen intermediate able to overcome the strong C–H BDE and perform the hydrogen atom abstraction (HAA) reaction.

In their seminal study on the chitin-active LPMO SmAA10, Vaaje-Kolstad et al. used labeled oxygen (<sup>18</sup>O<sub>2</sub>) to show that the oxygen atom incorporated in the final oxidized product was derived from O<sub>2</sub><sup>8</sup>. Similar results were later reported for cellulolytic LPMOs<sup>99</sup>, and were used to underpin the (apparent) monooxygenase nature of LPMOs. However, none of these studies unequivocally showed the direct use of O<sub>2</sub>, which was proposed years later to be a precursor of the more efficient, if not only, co-substrate H<sub>2</sub>O<sub>2</sub><sup>30</sup>. Indeed, it is known that in the presence of a reductant, LPMOs that are not bound to substrate behave as oxidases, catalyzing the reduction of O<sub>2</sub> to H<sub>2</sub>O<sub>2</sub><sup>100</sup> (Fig. 3, O-step; more details below). In a standard LPMO reaction under “monooxygenase conditions”, that is, containing O<sub>2</sub> and a reducing
agent, H₂O₂ will also be generated by non-enzymatic, oxidation of the reductant, in a process sometimes referred to as reductant auto-oxidation (Fig. 3, X-step), which is pH-dependent⁶⁸,¹⁰¹. Interestingly, several early studies showed a lack of apparent H₂O₂ production when LPMO was incubated with a genuine carbohydrate substrate⁸⁴,⁸⁷, and this observation was ascribed to the inability of a substrate-bound LPMO to generate H₂O₂. Still, H₂O₂ generated by reductant autooxidation and by substrate-free LPMO should have been detected. We now know that this is due to the productive consumption of H₂O₂ by substrate-bound LPMOs³⁰ in addition to the lower number of LPMOs molecules free in solution engaged in the O path.

In 2017, the H₂O₂-dependent LPMO catalysis was unveiled³⁰. The key points in favor of such mechanism are: (i) LPMO-catalysis can be achieved under anaerobic conditions if the reaction is supplied with exogenously added H₂O₂ and a reducing agent. (ii) In such reactions, the amount of reductant needed is sub-stoichiometric relative to the formation of oxidized LPMO products. (iii) The catalytic efficiency of the LPMO in H₂O₂-fueled reactions (“peroxygenase conditions”) is orders of magnitude higher than reactions driven by a combination of reductant and O₂ (“monooxygenase conditions”). (iv) When supplied with both O₂ and H₂¹⁸O₂, oxygen atoms incorporated in the reaction product originate from H₂O₂. (v) In the presence of a H₂O₂-consuming peroxidase and an excess of O₂ and reducing agent (meaning that the monooxygenase reaction is still possible), LPMO activity is inhibited. Most of these observations have also been made in subsequent studies, with different LPMOs and in different laboratories. Kinetic studies with both bacterial and fungal LPMOs have shown that oxidation of the LPMO-Cu(I) is several orders of magnitude faster with H₂O₂ than with O₂⁴⁷,⁶⁶, and that the same applies to product formation in reactions with substrate and micromolar concentrations of H₂O₂⁶⁷,⁸⁹,⁹²,¹⁰²,¹⁰³. Taken together, these observations point at LPMOs being peroxigenases.

Diving deeper into the chemistry at play, multiple computational studies on AA9 and AA10 LPMOs have focused on the possible monooxygenase and peroxigenase catalytic scenarios, as recently reviewed by Hagemann and Hedegård⁴¹. Accordingly, most of these
studies suggested that the reactive intermediate catalyzing C–H bond activation is most likely a Cu(II)–oxyl (Cu(II)–O•)²⁹,⁶⁶,⁹⁷,⁹⁸,¹⁰⁴,¹⁰⁵ (Fig. 5A), although it has not yet been experimentally isolated. Hedegård and Ryde estimated that the Cu(II)–oxyl has a C–H bond activation power of 468 kJ/mol, well above the estimated BDEs of the substrate, estimated at 434 kJ/mol for C4–H and 423 kJ/mol for C1–H²⁹. It should be noted that copper hydroxide species (Cu–OH) has also been suggested as a putative reactive intermediate by theoretical work²⁹,⁹⁸ and experimental work on inorganic model complexes¹⁰⁶.

QM/MM studies⁹⁸,¹⁰⁵ and DFT calculations⁹⁵,¹⁰⁴,¹⁰⁷ have shown that activation of O₂ by LPMO-Cu(I) leads to the formation of a Cu(II) superoxointermediate (Cu(II)-O-O•), which must be further reduced to become capable of HAA. In this pathway, two protons and an exogenous electron are added *in silico* to the distal oxygen of Cu(II)-O-O• to produce *in fine* Cu(II)-O• and H₂O (Fig. 5A). Interestingly, there is no obvious path for electrons and protons to be delivered to the copper site, which is secluded from the solvent when confined in the enzyme-substrate complex in the case of a bulky polysaccharide substrate (Fig. 4).

Alternatively, theoretical studies by Wang and collaborators on the MO route¹⁰⁵,¹⁰⁸, led to the suggestion that the formation a Cu(II)–OOH is followed by generation of H₂O₂ in the catalytic center. Here, the proximal O atom of Cu(II)–OOH would abstract a hydrogen atom from a small molecule reductant, ascorbate in this case, leading to formation of LPMO–Cu(I) and H₂O₂. The resulting situation would then allow a peroxygenase reaction and, as such, these studies by Wang et al. reconcile the MO and PO routes by showing that molecular oxygen is directly used by the LPMO to carry out a peroxygenase reaction. There is, however, a caveat in this line of thinking, which involves a physical interaction between the small molecule reductant and the Cu(II)–OOH species in the enzyme-substrate complex. While such an interaction is conceivable for LPMOs acting on a soluble linear substrate, as was the case in the Wang et al. studies, such an interaction is difficult to envisage in the case on an LPMO acting on the surface of a crystalline polysaccharide (Fig. 4).
Regarding the PO route, H₂O₂ can easily enter into the catalytic center of a substrate-bound LPMO (through a tunnel at the interface between the LPMO and the polysaccharide, so far only studied for an AA10 LPMO)⁷⁶,⁸⁹ and LPMO reactions with externally added H₂O₂ may reach rates of up to 10-100 per second⁸⁹,⁹⁰,⁹⁶,¹⁰³. Upon reaction with LPMO-Cu(I), the O-O bond in H₂O₂ has been proposed to undergo a homolytic cleavage that leads to a Cu(II)-OH species and a HO• radical, which is “caged” by a hydrogen bond network in the active site cavity, as computed first for AA9⁹⁷,⁹⁸,¹⁰⁵ and then AA10⁶⁶ LPMOs. These interactions direct the HO• radical to abstract a hydrogen atom from the Cu(II)-OH species (HAA¹, **Fig. 5A**) and form the highly reactive Cu(II)-O• that carries out the HAA reaction on the substrate (HAA², **Fig. 5A**) and subsequently hydroxylates the substrate via an oxygen-rebound mechanism. Notably, EPR spectroscopy studies have provided experimental support for the formation of a HO• radical derived from H₂O₂⁴⁷,⁶⁶. Also, recent studies have revealed the formation of transient aromatic radicals in the cores of LPMOs⁴⁷,⁵⁰,¹⁰⁹⁻¹¹¹, which suggests the presence of protective hole hopping mechanisms¹¹² that may help protecting the enzymes from radical damage.

The combination of hydroxyl radicals being formed and substrate binding having such a large effect on affinity for H₂O₂ not only support the notion that LPMOs are true peroxxygenases but also underpin the concept, proposed by Bissaro et al.³⁰,⁴⁵, that LPMOs are “enzymatic masters of Fenton-like chemistry”¹¹³.

## Glycosidic bond cleavage

In the oxygenase reaction (MO/PO paths) the Cu-Oxygen reactive species hydroxylates a C-H bond at the scissile glycosidic bond in a carbohydrate substrate. This reaction leads to a highly unstable hemiketal product that will undergo spontaneous intramolecular rearrangement leading to bond cleavage and generation of a C1- or C4-oxidized chain end (Fig. 1C)¹¹. Of note, whether this actual cleavage step is formally enzyme-dependent or not has been the object of discussion since the early studies on LPMOs¹¹,¹¹⁴, leading some researchers to discard the term “lytic” in favour of the “PMO” acronym. Nevertheless, the
adjective lytic and the LPMO acronym are widely used in the scientific community, since "lytic" underlines the occurrence of bond cleavage. This is special compared to most other monooxygenases, which tend to catalyze oxy-functionalization but not cleavage of their substrates.

Notably, oxidized C1 and C4 products display substantially different physicochemical properties which affects the qualitative and quantitative interpretation of the product profiles of LPMOs. In particular, C4-oxidized oligosaccharides are unstable under the alkaline conditions commonly used during chromatographic separation of soluble LPMO products (that is, short oxidized oligosaccharides). Confusingly, the decomposition of C4-oxidized products leads to the formation of native oligosaccharides with one less monomer compared to the original oxidized compound<sup>115</sup>. This latter phenomenon could explain why some researchers claim to have observed reductant-dependent glycoside hydrolase activity for LPMOs<sup>116</sup>. An early comparative study of multiple LPMOs from *Neurospora crassa* by Vu et al. demonstrated that standard (alkaline conditions) chromatographic analysis of product mixtures generated by C4-oxidizing LPMOs show many native products, while analysis of product mixtures generated by C1-oxidizing LPMOs do not<sup>117</sup>.
## Side reactions

LPMOs not only catalyze the hydroxylation reaction described above (that is, PO or MO paths), but have also been reported to catalyze two side reactions that do not lead to oxygen incorporation in a polysaccharide substrate, namely an oxidase reaction ("O path") and a peroxidase reaction ("P path"). LPMOs can also undergo an autocatalytic-inactivation reaction ("I path"). The understanding and consideration of all these side reactions is crucial to control LPMO catalysis.

### The oxidase reaction

Under standard aerobic conditions (that is, excess of molecular oxygen and reductant), and in the absence of a bona fide substrate, LPMOs catalyze an oxidase reaction in which the reductant is oxidized while O₂ is reduced to H₂O₂⁸⁴,⁸⁷,¹⁰⁰.

Several computational studies have explored the oxidase pathway, starting with the formation of a Cu(II)-superoxide intermediate upon the reaction between Cu(I) and O₂. Neutron protein crystallography studies by Meilleur's group allowed the experimental observation of an oxygen pre-bound state in an AA9 LPMO and the identification of Cu(II)-superoxo and hydroperoxo species⁹³,⁹⁴. In this latter work, protonation was attributed to a conserved second sphere residue (a conserved histidine in the AA9 LPMOs and a glutamic acid for most AA10 LPMOs), in line with previous crystallographic observations⁹⁵, as well as computational¹¹⁸ and mutagenesis¹¹⁹ studies.

It is important to note that the oxidase activity of LPMOs is very slow, with rates typically well below 0.1 s⁻¹, explaining the low rates of LPMO reactions under apparent "monooxygenase conditions". As such, LPMOs are outcompeted by other enzymes with regard to the efficiency of O₂ activation (Fig. 6). Equipped with co-factors suited for the two-electron reduction of O₂ to H₂O₂, many flavo-enzymes or copper-radical oxidases appear much more adequate to fulfill this function, with catalytic rates being in the range of 10⁻³ s⁻¹, compared to 10⁻⁵–10⁻¹ s⁻¹ for LPMOs³¹. While it has been suggested that the biological role of certain
LPMOs may be to generate H₂O₂ for use by other LPMOs¹²⁰ or biomass degrading peroxidases¹²¹, such a role does not seem likely in light of the relatively low oxidase activity, although this may vary with reductant and pH⁶⁸.

## The peroxidase reaction

After H₂O₂ was reported to act as a co-substrate in the LPMO-catalyzed peroxygenase reaction, Breslmayr et al. investigated whether LPMOs could harness H₂O₂ to carry out a peroxidase reaction¹²². Using a small organic molecule, 2,6-dimethoxyphenol (syringol), as chromogenic substrate and H₂O₂ as a co-substrate they demonstrated peroxidase activity for several AA9s. The rates of the LPMO peroxidase reaction appear low ($k_{cat}$ = [1-20 s⁻¹]) compared to genuine peroxidases ($k_{cat}$ = [10-10³ s⁻¹]; $k_{cat}/K_M$ = [10³-10⁷ M⁻¹·s⁻¹]) (Fig. 6). Kinetic analyses by Kuusk and Valjamae, looking at the reductant peroxidase activity (that is, LPMO-catalyzed ascorbate oxidation by H₂O₂) of a fungal LPMO, indicated similarly low peroxidase rate ($k_{cat}$ = 2.1 s⁻¹) and efficiency ($k_{cat}/K_M$ = [2.7 x 10⁴ M⁻¹·s⁻¹]). These values are 1-2 orders of magnitude lower than the peroxygenase reaction with substrate⁹⁶.

## The auto-inactivation reaction

Oxidative damage is a major threat for redox enzymes, and LPMOs are no exception. As previously discussed, reduced LPMOs are prone to reoxidation by O₂ or H₂O₂, which will generate oxygen species that, if not generated within the confinement of a bona fide enzyme-substrate complex, may oxidatively damage and eventually inactivate the LPMO³⁰,⁴³,⁴⁶,¹²³. Analysis of auto-oxidatively damaged LPMOs has shown that the copper-coordinating histidines are particularly vulnerable³⁰. Maintaining saturating amounts of a suitable substrate and keeping H₂O₂ levels low will prevent damage to the LPMO and allow the enzyme to harness available H₂O₂ in a productive manner.

Notably, there are indications that LPMOs have developed mechanisms that protect against oxidative damage, including the possible hole hopping mechanisms above. In fungal LPMOs, experimental¹²⁴ and QM/MM¹²⁵ studies suggest that the methylation of the N-terminal histidine likely also plays a protective role. A recent kinetic study estimated the probability of a
(methylated) fungal LPMO becoming irreversibly inactivated following a reductant peroxidase reaction to be in the order of 0.01⁹⁶.
## LPMOs are unique peroxygenases

$H_2O_2$ is ubiquitous in Nature. Although $H_2O_2$ can directly oxidize thiol groups in proteins and other organic molecules via two-electron oxidation¹²⁶, its strong oxidative power comes from its propensity to generate (upon one-electron reduction by reduced transition metal ions) one of the strongest known oxidants, the hydroxyl radical, $HO^•$ ($E'° = 2.31$ V at pH 7.0)¹²⁷. The eponymous reaction first described by H.J.H. Fenton¹²⁸ ($Fe^{2+} + H_2O_2 \rightarrow Fe^{3+} + HO^• + HO^•$) and later detailed by F. Haber and J. Weiss¹²⁹, entails the ferrous ion-catalyzed homolytic decomposition of $H_2O_2$ generating a hydroxide ion and a $HO^•$ radical that can indiscriminately degrade surrounding bioorganic compounds. Fenton-like reactions may also be driven by other transition metals at low oxidation state, such as $Cu(I)¹³⁰$.

Clearly, in a biological context the combination of free (reduced) transition metals and $H_2O_2$ needs to be avoided. Even so, living systems need $H_2O_2$ as a signaling molecule and as a key co-substrate in enzyme-dependent redox reactions. Enzymes that can exploit $H_2O_2$ (peroxyzymes) as reagent for selective oxidation or oxygenation reactions are widespread in biological systems and their use in the environmentally friendly production of specialty chemicals is gaining momentum¹³¹. These enzymes are mainly peroxygenases and peroxidases (including catalases) although some hydrolases have been shown to display catalytic promiscuity allowing the use of $H_2O_2$, an activity referred to as perhydrolase activity¹³². Alongside catalyzing exciting chemistry, peroxyzymes offer solutions to overcoming the so-called oxygen dilemma encountered in $O_2$-dependent reactions¹³³. This “dilemma” refers to limitations in efficiency owing to low $O_2$ solubility and limited diffusion, the need for external supply of reducing equivalents, and the risk of uncoupled production of free radicals¹³³. These factors are among the main hurdles limiting the implementation of $O_2$-using enzymes in industrial settings.
Peroxyzymes

Most H₂O₂-using enzymes display a conserved heme iron prosthetic group analogous
to that of cytochrome P450 monooxygenases (Enzyme Commission number EC 1.14.14.1)
and form, upon reaction with H₂O₂, the key reactive intermediate Compound I (Cpd I) and
H₂O¹³³,¹³⁴ (Fig. 5B). In heme peroxidases (EC 1.11.1.X), Cpd I catalyzes the oxidation of an
electron donor molecule, which is an organic substrate for most peroxidases, except in the
case of catalases (E.C. 1.11.1.6), where it is a molecule of H₂O₂. In this latter case, the final
products are O₂ and 2H₂O. In contrast, heme peroxygenases (EC 1.11.2.X), use Cpd I to
perform different oxyfunctionalizations¹³⁵, such as hydroxylation of poorly reactive C–H bonds,
and oxygenation of C=C bonds (alkene epoxidation) or heteroatoms (for example,
sulfoxidation). Thus, the outcome of the reaction catalyzed by Cpd I varies between subgroups
of peroxyzymes and depends on structural features such as active site topology, solvent
accessibility and the axial ligand of the heme iron¹³⁶⁻¹³⁸. Crystal structures of peroxidases,
catalases and peroxygenases have revealed consistent differences in the axial ligands of the
heme center: the iron atom is coordinated by a histidine or a tyrosine in canonical peroxidases
and catalases, respectively, while peroxygenases feature instead a type-1 heme-thiolate
cofactor.

Until the discovery of the primary peroxygenase activity of LPMOs³⁰, peroxygenases all featured a heme iron active site and were formally divided into 5 groups: unspecific peroxygenases (UPOs, EC 1.11.2.1), myeloperoxidases (EC 1.11.2.2), plant seed peroxygenases (EC 1.11.2.3), fatty-acid peroxygenases (EC 1.11.2.4) and 3-methyl-L-tyrosine peroxygenases (EC 1.11.2.5). Among the hitherto reported peroxygenases, only the UPOs are secreted enzymes and it is worth noting that, like LPMOs, UPOs are produced by soil-litter and wood-colonizing fungi, and may play a role in the degradation of recalcitrant biomass¹³⁹.

On a side note, canonical P450 monooxygenases are also able to perform
peroxygenation reactions, *via* a route referred to as the “peroxide shunt”, where H₂O₂, used at
high concentrations, is “forced” to act as direct surrogate for both molecular oxygen and
reducing equivalents to produce Cpd 0, the precursor of Cpd I (Fig. 5B)¹⁴⁰,¹⁴¹. While the
promiscuity of co-substrates is reminiscent of a potential similar promiscuity in LPMOs, the
difference between the two enzyme types is immense. It is experimentally evident that the
catalytic efficiencies of LPMO-catalyzed peroxygenase reactions resemble those of true
peroxygenases (~10⁶ M⁻¹s⁻¹)⁸⁹,¹⁰², whereas P450s are lousy peroxygenases¹⁴². Indeed,
employment of the peroxide shunt does not yield efficient P450 catalysis because: (i) the total
turnover number of the reaction is usually several orders of magnitude lower (often less than
10)¹⁴³,¹⁴⁴ compared to the monooxygenase reaction (and compared to the LPMO
peroxygenase reaction)³⁰,⁴⁹,⁶⁷; (ii) P450s usually have low (mM) affinities for H₂O₂ (for example
5 mM for P450 2B4)¹⁴⁵, contrasting with the low µM affinity for H₂O₂ of a substrate bound
LPMO; (iii) finally, the H₂O₂ concentrations needed to make P450s carry out peroxygenase
reactions at an appreciable rate are not physiologically relevant and cause rapid oxidative
degradation of the heme.

Comparing LPMOs with other peroxygenases

The use of a single copper atom to catalyze a peroxygenase reaction clearly separates LPMOs from heme iron-containing peroxyzymes. The use of copper may be a response to the emergence of LPMOs during evolution coinciding with the increased availability of copper that arose after the great oxygenation event¹⁴⁶. Alternatively, it may have more specific reasons, such as its simplicity and robustness (relative to heme iron) or the nature of the catalytic task that drove evolution of LPMOs.

The recalcitrance and limited solubility of common LPMO polysaccharide substrates
has undoubtedly played a role in shaping the unique molecular architecture and catalytic
center of these enzymes that are tailored to achieve efficient interfacial catalysis. Upon the
essential secretion step, a mature LPMO enzyme will feature a flat and extended substrate-
binding surface with a solvent-exposed catalytic copper site, and potentially with appended
modules (for example carbohydrate-binding modules, or CBMs)⁶⁹ enhancing the interaction
with insoluble substrates. Conversely, heme iron-containing peroxyzymes often have active
sites that are buried deep in the protein core and that are tailored to interact with small
substrates. In general, all these peroxyzymes, except UPOs, are intracellularly localized, either
in the cytosol or within the organelles. The solvent accessibility of the copper site in LPMOs
allows for electron donor promiscuity⁵², which guarantees an efficient priming reduction step.
It is noteworthy that this reductive activation step is essential for LPMOs, while the action of
peroxide-driven P450s (that is, P450s in shunt mode), UPOs and other peroxygenases is
independent from any electron supply.

The highly solvent exposed redox-active copper in LPMOs adds to the uniqueness of
these enzymes. Comparative studies of the affinity of LPMOs for Cu(I) and Cu(II)¹⁰ and the
(relatively slow) kinetics of the various off-pathway reactions discussed above show that
LPMOs have evolved to stabilize a solvent-exposed reduced copper. LPMOs bind Cu(I) much
more strongly than Cu(II) and while a reduced LPMO may engage in off-pathway reoxidation
reactions (Fig. 3), productive reactions with substrate are orders of magnitude faster, as shown
from both experimental⁹⁶ and computational studies¹²⁵.

When comparing the proposed reaction mechanism of LPMOs and heme iron-
containing peroxyzymes, it is noticeable that while a seemingly similar hydrogen-abstracting
oxygen species is being generated (Cu(II)-oxyl and Cpd I, respectively), the enzymes differ in
how they react with H₂O₂. In LPMOs, both experimental and computational data point to
homolytic cleavage of H₂O₂ and formation of a hydroxyl radical (see above, Fig. 5A). Fe-
dependent peroxidases and peroxygenases (including rare non-heme variants), follow a
pathway for H₂O₂ activation known as the Poulos-Kraut mechanism¹⁴⁷, in which H₂O₂, after
binding to Fe, is deprotonated to yield the ferric-hydroperoxide intermediate (Cpd 0; Fe(III)-O-
O-H), which then leads to Cpd I and release of H₂O upon heterolytic cleavage of the O-O bond
(Fig. 5B). This mechanism depends on acid/base catalysis facilitated by second sphere
histidine and arginine residues¹⁴⁸. Interestingly, after the discovery of the peroxygenase activity
of LPMOs, QM/MM calculations have led to suggestions that some heme iron peroxyzymes
also rely on homolytic cleavage of the O-O bond¹⁴⁹. These enzymes include certain P450s¹⁵⁰
and HppE¹⁵¹. HppE is a particularly curious case since it was shown that this non-heme mono-
iron epoxidase involved in fosfomycin production, is not an oxidase (that is, does not use O₂) as previously thought, but a peroxidase, using H₂O₂¹⁵². In an analogous manner to LPMOs, these enzymes would cope with the highly reactive hydroxyl radical species by limiting its diffusion through a H-bonding network and the presence of productively bound substrate. As shown by research on LPMOs, such stabilized HO• radicals can perform regioselective oxidations rather than trigger the autoxidation and inactivation of the enzyme.

Conclusion and outlook

The activity of LPMOs has been studied for almost two decades. The earliest report (in 2005) by Vaaje-Kolstad et al. that the protein CBP21 (then thought to be non-catalytic) had an enigmatic boost effect on chitinases⁷ was enlightened by the discovery of the underlying oxidative cleavage reaction in 2010⁸, and the role of copper played during catalysis⁹⁻¹¹. The progressive uncovering of the various facets of LPMO chemistry, including the multitude of possible redox reactions (Fig. 3) and rapid enzyme inactivation under certain conditions, studied at multiple scales – from bioreactors to atomistic theoretical studies – provides an informed picture of the mode of action of LPMOs, which act as efficient peroxygenases and, possibly, slow monooxygenases.

LPMOs have evolved to harness the catalytic potential of Cu(I) which, when not bound to the LPMO, would be hardly soluble and rapidly oxidized, to catalyze selective site-specific Fenton-like chemistry¹¹³. While LPMOs bring new chemistry and catalytic opportunities, it must be noted that they seem optimized for catalyzing interfacial reactions and that binding of the polysaccharide substrate affects the reactivity of the catalytic copper site. Additionally, the copper-binding histidine-brace motif does not provide a catalytically competent reaction center and that its reactivity and proper functionality are totally dependent on the occupancy of the additional copper coordination positions⁴⁰,¹⁵³,¹⁵⁴ and on nearby, mostly polar, “second sphere” residues⁷,⁵³,⁶⁶,⁷⁹,¹¹⁹,¹²³, which vary among LPMO families⁷¹,⁷². LPMOs also differ in terms of possible protective mechanisms, be it methylation of the N-terminal histidine (only in fungal enzymes), the presence of a redox-active tyrosine close to the copper (always in AA9s, while
it is a Phe in most AA10s), and the number and distribution of buried aromatic residues that
may be involved in hole hopping routes. We believe that this natural variability, with
implications in reactivity and protective mechanisms, must be considered when engineering
LPMOs for new, improved catalytic functions, or in the design of bio-inspired catalytic
monocopper complexes<sup>155-158</sup>.

The discovery of LPMOs peroxygenase reaction³⁰ adds a new dimension to the
peroxyzyme family, mainly populated of heme iron enzymes. Indeed, LPMOs are so far the
first and only example of peroxygenases depending on a copper active centre. Yet, the
peroxygenase nature of LPMOs has been, and to some extent still is¹⁸ controversial, ever since
its first experimental demonstration³⁰. On the one hand, the importance of the peroxygenase
reaction and the kinetic irrelevance of the monooxygenase reaction under *in vitro* conditions
seems undisputed today⁴⁵,⁴⁸,⁶⁴,⁹⁵,⁹⁶,¹⁵⁹. On the other hand, it is conceivable that functional
insights obtained in the laboratory, using model carbohydrate substrates and with potentially
artificial supplementation of electrons and co-substrates, may not give an entirely accurate
picture of what happens *in vivo*. While the studies discussed above may indicate that LPMOs
do not catalyze a monooxygenase reaction at all, the situation may be different in Nature,
where secreted enzymes will face O₂ concentrations close to 200 µM, that is, significantly
higher than expected H₂O₂ concentrations. Nevertheless, in aerobic environments, LPMOs
would also be consistently surrounded by available, water dissolved, H₂O₂ which is the reaction
product or substrate of a multitude of oxidoreductases during lignocellulose conversion³¹ and
which may also be generated through abiotic oxidation of abundantly present lignin¹⁶⁰.
Underscoring the latter, recent experiments by R. Ludwig's group have shown that, during
wood decay, stable LPMO peroxygenase reactions can occur at low steady-state H₂O₂
concentrations (µM range) and that the LPMO does not consume O₂⁴⁹. Further studies aiming
at quantifying the actual O₂ and H₂O₂ levels in relevant biological systems would be of great
interest. Staying with biology, there is increasing evidence that the role of LPMOs in Nature
goes beyond biomass degradation, as LPMOs seem involved in microbial pathogenesis in
plants, insects and humans<sup>16,18,24,161</sup>. It is conceivable that, during infection, LPMOs could be
fueled by the H<sub>2</sub>O<sub>2</sub> produced by the host's immune response (oxidative burst) or other H<sub>2</sub>O<sub>2</sub>-
generating enzymes.

Overall, the chemistry of LPMOs still holds many secrets that will hopefully be
unravelled in future studies. While LPMOs have been extensively studied and are industrially
applied in biomass processing, investigations on the role of their fascinating chemistry in
important biological processes such as in pathogenicity mechanisms or cellular development
are only just beginning<sup>16</sup>. Furthermore, we envision that the unique properties of LPMOs, such
as the solvent-exposed catalytic copper site that may engage in interfacial powerful Fenton-
type reactions, could be extended beyond the field of recalcitrant polysaccharides, for instance
for the biocatalytic recycling of synthetic polymers such as plastics.
**Acknowledgements**

We thank current and past members of our research groups that have contributed to LPMO research. We also thank our colleagues from the LPMO community. We apologize in advance to all investigators whose research cannot be appropriately discussed and cited owing to space limitations. A.M. and B.B. discloses support for the research of this work from INRAE [EvoFun project PAF_02]. V.G.H.E. discloses support for the research of this work from the European Research Council [grant number 856446].

**Author contributions**

All authors contributed substantially to discussion of the content. A.M. and B.B. wrote the first draft of the article. All authors reviewed and edited the manuscript before submission.

**Competing interests**

The authors declare no competing interests.

**ORCID**

Alessia Munzone : 0000-0002-9410-610X

Vincent Eijsink : 0000-0002-9220-8743

Jean-Guy Berrin : 0000-0001-7570-3745

Bastien Bissaro : 0000-0001-8354-3892
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**Fig. 1. Overview of the mode of action of LPMOs. (A)** Schematic view of Lytic Polysaccharide Monooxygenases (LPMOs) during cellulose depolymerization (glucose units shown as blue circles). LPMOs (shown as green triangle with yellow circle) disrupt the cellulose surface by oxidation (shown as red star) and produce new accessible chains tractable by cellulases, such as endoglucanases (EGs), and cellobiohydrolases (CBHs). EGs (shown in magenta) act by cleaving internal β-glycosidic bonds in the cellulose chain. CBHs, cleave externally, from either reducing (R) or nonreducing (NR) chain ends via CBHI (shown in light blue) or CBHII (shown in green), respectively. As a result, CBHs produce cellobiose that is further hydrolyzed to glucose by β-glucosidases (BG, shown in purple). **(B)** The active site of a fungal LPMO including a methylated N-terminal histidine residue (X=CH₃), showing the copper-oxyl (Cu(II)-O•) reactive species proposed to catalyze C–H abstraction. **(C)** LPMOs cleave glycosidic bonds by hydroxylating either the C1 or C4 carbon, yielding lactone or 4-ketoaldose products, which are spontaneously hydrolyzed or hydrated to aldonic acids or 4-gemdiols, respectively.

**Fig. 2. Schematic overview of enzymatic oxygenation and oxidation reactions.** In the reactions shown, electrons are transferred to O₂ or H₂O₂ and oxygen may (that is, oxygenation) or may not (that is, oxidation) be incorporated into the product. For each reaction, the figure shows the structure of a representative enzyme catalyzing the reaction, from left to right: homoprotocatechuate 2,3-dioxygenase (HPCD) from *Bulbophyllum fuscum* (PDB ID 2IGA)³³, cytochrome P450 (CYP) from *Homo sapiens* (PDB ID 1W0E)³⁴, unspecific peroxygenase (UPO) from *Agrocybe aegerita* (PDB ID 5OXU)³⁵, lytic polysaccharide monooxygenase (LPMO) from *Neurospora crassa* (PDB ID 4QI8)³⁶, glucose oxidase (GOX) from *Aspergillus niger* (PDB ID 1GAL)³⁷, laccase (LAC) from *Trametes versicolor* (PDB ID 1KYA)³⁸, horseradish peroxidase (HRP) from *Armoracia rusticana* (PDB ID 1W4W)³⁹. At the bottom, the co-factor employed by each representative enzyme is indicated. Co-factors provided in between brackets are co-factors that have been reported in the literature for other enzymes catalyzing the same reaction. FMN, flavin mononucleotide; FAD, flavin adenine dinucleotide.

**Fig. 3. Dissecting the chemistry of LPMO catalysis.** Overview of the Lytic Polysaccharide Monooxygenase (LPMO) reactions possibly occurring in the absence (light yellow background) and presence (light grey background) of a productively bound carbohydrate substrate. Once the LPMO-Cu(II) form is reduced to Cu(I) by a reducing agent or enzyme (R path, priming reduction), the breakdown of glycosidic bonds can efficiently occur *via* a peroxygenase (PO path) or *via* a monooxygenase (MO path) mechanism. Several pathways can lead to the *in situ* generation of H₂O₂, including the enzyme-dependent oxidase reaction (O path) or as a result of non-enzymatic oxidation of low molecular weight reductants commonly used in LPMO reactions (for example ascorbic acid and gallic acid) (X path). In the absence of a bound carbohydrate, the LPMO can also catalyze the oxidation of a non-carbohydrate substrate in what it is known as a peroxidase reaction (P path). Excessive levels of H₂O₂ resulting from the above-mentioned reactions, can lead to the autocatalytic enzyme inactivation since a reduced LPMO reacting with H₂O₂ will generate Reactive Oxygen Species (ROS) that, particularly in the absence of substrate, may damage the enzyme (I path).
**Fig. 4.** Illustration of a polysaccharide-bound LPMO. MD model of an LPMO (the AA10 from *Serratia marcescens*) bound to β-chitin<sup>76</sup>. The model was based on biochemical data (enzyme product profile) and supported by experimental EPR. The inset shows the entrance to a tunnel that connects the active site to the bulk solvent, including residues Glu60 and Asn185 (shown as yellow sticks), that limit tunnel accessibility to small molecules only (such as O<sub>2</sub>, H<sub>2</sub>O<sub>2</sub> or H<sub>2</sub>O). The copper coordinating histidines (His 28 and His 114) are shown as blue sticks and the copper atom as orange sphere. LPMO, lytic polysaccharide monooxygenase; AA10, auxiliary activity 10; MD, molecular dynamics; EPR, electron paramagnetic resonance.

**Fig. 5.** Reaction cycles in LPMOs and heme iron peroxygenases. (A) The main peroxygenase (PO) path, and monooxygenase (MO) path (in grey) proposed for lytic polysaccharide monooxygenases (LPMOs). For both paths, the LPMO enters the catalytic cycle upon priming reduction of the resting state LPMO-Cu(II) to LPMO-Cu(I). In the PO path, reaction of H<sub>2</sub>O<sub>2</sub> with the active form leads, via a Fenton-type chemistry producing a caged hydroxyl radical (highlighted in cyan), to the copper-oxyl (Cu(II)-O•) intermediate. In the MO path (that is, reaction with O<sub>2</sub>), it has also been proposed that such intermediate could be reached provided that additional electrons and protons can be supplied. The copper-oxyl intermediate is a consensus species that has been suggested to catalyze hydrogen atom abstraction (HAA) from the substrate (RH), leading via an oxygen-rebound mechanism to the hydroxylated product (R-OH) (see more details in main text). (B) Catalytic cycle of heme iron peroxygenases. After the resting state of the heme iron peroxygenase (Fe(III)) upper coordination position occupied by a water molecule) forms a complex with the substrate (RH), reaction with a molecule of H<sub>2</sub>O<sub>2</sub> has been described to follow two potential paths, either going via formation of so-called compound 0 (Cpd 0, Fe(III)-hydroperoxo complex) (path a) or, perhaps, in some cases, via formation of a hydroxyl radical<sup>149</sup> (highlighted in cyan) resulting from a Fenton-like reaction (path b) similarly to what has been reported for LPMOs. Both paths lead to the release of H<sub>2</sub>O and formation of compound I (Cpd I, Fe(IV)-oxo/porphyrin cation radical complex), which catalyzes HAA from the substrate and leads to compound II (Cpd II, Fe(IV)-hydroxo complex) and a substrate radical (R•). An oxygen-rebound mechanism leads to the hydroxylated product (R-OH) and regeneration of the resting state.

**Fig. 6.** Violin Plot of reported oxidase/peroxidase and monooxygenase/peroxygenase activities of different types of O<sub>2</sub>/H<sub>2</sub>O<sub>2</sub>-dependent enzymes. The x-axis shows various enzyme classes, with, where applicable, the CAZyme family code. The reported substrate conversion rates are Vi/[E] values, which in most cases were determined at saturating substrate concentration and thus correspond to k<sub>cat</sub>, except for the "monooxygenase" reactions of LPMOs (where reductant concentrations may have been rate-limiting) and the peroxide shunt reactions of P450s. For the latter, the reported rates are obtained under conditions employing several orders of magnitude higher concentrations of H<sub>2</sub>O<sub>2</sub> (0.6-50 mM) compared to true peroxidase and peroxygenase reactions. CAZyme, carbohydrate active enzyme; LPMO, lytic polysaccharide monooxygenase; Vi, initial velocity; k<sub>cat</sub>, catalytic constant; UPO, unspecific peroxygenase; P450, cytochrome P450; FAD, flavin adenine dinucleotide; CRO, copper radical oxidase.
**Short Summary (< 40 words)**

LPMOs are key enzymes in biomass processing and pathogenicity. They are the first known copper enzymes capable of utilizing H₂O₂ to hydroxylate C–H bonds. This review draws a portrait of the catalytic paths at play and highlights outstanding questions in their reactivity.