Characterization of a lytic polysaccharide monooxygenase from Aspergillus fumigatus shows functional variation among family AA11 fungal LPMOs

The discovery of oxidative cleavage of recalcitrant polysaccharides by lytic polysaccharide monooxygenases (LPMOs) has affected the study and industrial application of enzymatic biomass processing. Despite being widespread in fungi, LPMOs belonging to the auxiliary activity (AA) family AA11 have been understudied. While these LPMOs are considered chitin active, some family members have little or no activity toward chitin, and the only available crystal structure of an AA11 LPMO lacks features found in bacterial chitin-active AA10 LPMOs. Here, we report structural and functional characteristics of a single-domain AA11 LPMO from Aspergillus fumigatus, AfAA11A. The crystal structure shows a substrate-binding surface with features resembling those of known chitin-active LPMOs. Indeed, despite the absence of a carbohydrate-binding module, AfAA11A has considerable affinity for α-chitin and, more so, β-chitin. AfAA11A is active toward both these chitin allomorphs and enhances chitin degradation by an endoacting chitinase, in particular for α-chitin. The catalytic activity of AfAA11A on chitin increases when supplying reactions with hydrogen peroxide, showing that, like LPMOs from other families, AfAA11A has peroxygenase activity. These results show that, in stark contrast to the previously characterized AfAA11B from the same organism, AfAA11A likely plays a role in fungal chitin turnover. Thus, members of the hitherto rather enigmatic family of AA11 LPMOs show considerable structural and functional differences and may have multiple roles in fungal physiology.

The discovery of oxidative cleavage of recalcitrant polysaccharides by lytic polysaccharide monooxygenases (LPMOs) has affected the study and industrial application of enzymatic biomass processing. Despite being widespread in fungi, LPMOs belonging to the auxiliary activity (AA) family AA11 have been understudied. While these LPMOs are considered chitin active, some family members have little or no activity toward chitin, and the only available crystal structure of an AA11 LPMO lacks features found in bacterial chitin-active AA10 LPMOs. Here, we report structural and functional characteristics of a singledomain AA11 LPMO from Aspergillus fumigatus, AfAA11A. The crystal structure shows a substrate-binding surface with features resembling those of known chitin-active LPMOs. Indeed, despite the absence of a carbohydrate-binding module, AfAA11A has considerable affinity for α-chitin and, more so, β-chitin. AfAA11A is active toward both these chitin allomorphs and enhances chitin degradation by an endoacting chitinase, in particular for α-chitin. The catalytic activity of AfAA11A on chitin increases when supplying reactions with hydrogen peroxide, showing that, like LPMOs from other families, AfAA11A has peroxygenase activity. These results show that, in stark contrast to the previously characterized AfAA11B from the same organism, AfAA11A likely plays a role in fungal chitin turnover. Thus, members of the hitherto rather enigmatic family of AA11 LPMOs show considerable structural and functional differences and may have multiple roles in fungal physiology.
Many microorganisms produce an arsenal of carbohydrateactive enzymes that concertedly operate to degrade carbohydrate biomass. These enzyme systems are fundamental to the global carbon cycle and prevent an environmental buildup of stable and abundant sugar polymers such as cellulose and chitin (1). The discovery of lytic polysaccharide monooxygenases (LPMOs) by Vaaje-Kolstad et al. (2) in 2010 changed the idea that depolymerization of cellulose and chitin is accomplished exclusively by hydrolytic enzymes or glycoside hydrolases. It is now well known that many aerobic microorganisms cosecrete LPMOs and glycoside hydrolases during growth on insoluble sugars (3)(4)(5)(6), and the contribution of LPMOs to the efficiency of chitinolytic or cellulolytic enzyme cocktails is well established (2,(7)(8)(9)(10)(11)(12)(13).
LPMOs have gained substantial interest from industries and the scientific community because of their unusual enzymatic properties. Using a single copper ion cofactor, bound in a highly conserved so-called histidine brace (14,15), LPMOs carry out oxidative cleavage of glycosidic bonds using molecular oxygen (O 2 ) (2) or hydrogen peroxide (H 2 O 2 ) (16) as a cosubstrate. The redox reaction catalyzed by LPMOs depends on reduction of the bound Cu(II) to form Cu(I) (2,17). In the case of an O 2 -driven reaction, the reaction requires two externally delivered electrons per catalytic cycle (2). In the case of H 2 O 2 -driven reaction, which tends to be much faster (16,(18)(19)(20), a reduced LPMO can carry out multiple reactions without the need for additional externally delivered electrons (16,21,22). Cellulose-active LPMOs oxidize either the C1 or the C4 carbon of the scissile glycosidic bond; some LPMOs exclusively act on C1 or C4, whereas others are less specific and produce a mixture of C1-oxidized and C4-oxidized products (23,24). This oxidation leads to spontaneous subsequent bond cleavage (25). C6-oxidized products have also been reported (26,27), but their importance and connection to the chain-cleaving function of LPMOs remain unclear. For chitin-active LPMOs, only C1-oxidizing activity has been reported. The latter activity leads to the generation of new chain ends that are lactones, which spontaneously convert to aldonic acids (2).
Despite a strictly conserved catalytic center, that is, the histidine brace, LPMOs display large sequence diversity and currently populate eight "auxiliary activity" (AA) families in the Carbohydrate-Active enZymes database (http://www.cazy. org/Auxiliary-Activities.html; as of August 2021), a sequencebased database of carbohydrate-active enzymes, namely AA families 9 to 11 and 13 to 17 (28). Fungal LPMOs appear in families 9, 11, 13, 14, and 16. Of these, the cellulose-active fungal AA9 LPMOs are the most abundant and the best studied. Remarkably, while chitin-active LPMOs are commonly found among members of the AA10 family, which lacks fungal enzymes, and while fungi are known for their ability to produce multiple chitinases and degrade chitin (29), little is known about fungal chitin-active LPMOs.
Fungal chitin-active LPMOs have been classified into the AA11 family (30). So far, only three members of this family have been subjected to some degree of functional characterization (30)(31)(32). Available structural information is limited to the incomplete crystal structure of the catalytic domain of a twodomain AA11 from Aspergillus oryzae (AoAA11; (30)). Family 11 LPMOs are the most widespread of fungal LPMOs as they occur in Dikarya and most basal fungal lineages (33). The ectomycorrhizal fungus Choiromyces venosus contains the highest number of genes (i.e., 14 genes) encoding AA11 LPMOs. AA11s are most prevalent in Ascomycota, where at least one gene encoding an AA11 LPMO has been found in 385 (99.2%) out of the 388 analyzed genomes, and where the average number of genes encoding AA11s is 3.9 ± 1.8 (33). The genome of the ascomycete Aspergillus fumigatus (34) contains three genes encoding family AA11 LPMOs, here referred to as AfAA11A (UniProt ID: Q4WF00; gene, AFUA_3G03950), a single domain enzyme, AfAA11B (UniProt ID: Q4WEH3; gene, AFUA_5G03010), a two-domain enzyme with a C-terminal domain of unknown function, and AfAA11C (UniProt ID: Q4WM72; gene, AFUA_6G10930), also a two-domain enzyme with a C-terminal domain of unknown function. Interestingly, a recent in-depth functional study of AfAA11B (32), which is very similar to AoAA11 (73% sequence identity for the catalytic domains), showed that this enzyme has low activity on chitin (i.e., much lower than chitin-active bacterial AA10 LPMOs) while being efficient when acting on soluble substrates. This raises questions as to the natural function of AA11 LPMOs.
To gain more insight into the AA11 LPMO family and oxidative chitin conversion by fungi, we have structurally and functionally characterized AfAA11A, including studies of this protein's ability to potentiate the action of chitinases. We also assessed whether this AA11 LPMO, like AA9 and AA10 LPMOs, is capable of employing H 2 O 2 as its oxygen-containing cosubstrate. Next to providing insight into the functionality of family AA11 LPMOs, our study provides the first complete crystal structure of a member of this family.

Protein production and deglycosylation
The gene encoding AfAA11A (UniProt ID: Q4WF00) was cloned in Pichia pastoris, and recombinant AfAA11A was purified to homogeneity from the culture supernatant using hydrophobic interaction chromatography and size-exclusion chromatography (Fig. 1). SDS-PAGE analysis showed that purified AfAA11A had a mass close to 30 kDa, indicating that the protein carried glycan moieties. Analysis with the NetNGlyc server (http://www.cbs.dtu.dk/services/NetNGlyc/) showed one possible N-glycosylation site, at Asn62. Indeed, treatment of the purified protein with an endo-β-N-acetylglucosaminidase from Enterococcus faecalis (EfEndo18A) (35) led to a reduction in the protein mass to a value, approximately 22 kDa, that corresponds well with the theoretical mass (21,750 Da; Fig. 1). Initial comparison of the glycosylated and deglycosylated protein batches indicated similar activities and properties, although minor, borderline significant, differences were observed in, for example, substrate binding and melting point (not shown). Here, we report the functional characterization of the glycosylated protein.

AfAA11A crystal structure
Crystallization conditions were screened for both N-glycosylated and de-N-glycosylated AfAA11A. Only one of the conditions tested resulted in a protein crystal suitable for structure analysis, for the de-N-glycosylated protein. The structure of AfAA11A was solved by X-ray crystallography, and a complete structure was obtained, which, however, lacked a bound metal ion in the histidine brace. Crystal data, as well as data collection and refinement statistics, are provided in Table 1. Of note, except for one GlcNAc attached to Asn62, the structure of this N-deglycosylated protein showed no other signs of N-glycosylation or O-glycosylation. Asn62 is not part of the flat LPMO surface containing the catalytic site and is located far away from the catalytic histidines; the Cβ-Cβ  The structure of AfAA11A (Fig. 2) shows a typical LPMO fold with a core composed of β-strands that are connected by several loops and a few short helices. The core of the protein consists of two β-sheets formed by antiparallel β-strands, which are arranged in an immunoglobulin-like β-sandwich fold. Similar to AA9s, and unlike AA10s, AfAA11A has an extended region C terminally of the β8-strand, that is, the most C-terminal strand in the β-sandwich. Pairwise alignments with all structures in the Protein Data Bank using the PDBeFold service (36) at default settings showed that AfAA11A is structurally most similar to the catalytic domain of AoAA11 (Protein Data Bank ID: 4MAH), with a Q score of 0.58 and a Cα-RMSD of 1.59 over 186 residues. These two proteins share 48% sequence identity ( Fig. 2A). AfAA11A contains a typical LPMO copper-binding site comprised of the nonmethylated N-terminal histidine and His71 (Figs. 2 and 3). The axial copper coordination positions are occupied by the hydroxyl group of Tyr133 and the methyl side chain of Ala69. This is a common arrangement in LPMOs, including chitin-active AA10s, although the amino acid residues in these axial positions may vary. In particular, bacterial chitinactive AA10 LPMOs have a phenylalanine rather than a tyrosine (Tyr133 in AfAA11A) in the proximal axial coordination position. Figure 3B shows that the copper-binding regions of AfAA11A and AoAA11 are almost identical. The variation in the orientation of His1 is likely because of the absence of a metal ion in AfAA11A, since NMR studies have shown that metal binding affects His1 in particular (15,37).
While the two AA11 enzymes have almost identical catalytic sites, they show considerable differences elsewhere in the structure. Relative to AoAA11, AfAA11A has an insertion spanning residues 16 to 26 (Figs. 2A and 3A), which contains a surface-exposed tyrosine, Tyr25, in a position similar to that of Tyr27 in CBP21 (SmAA10A), a chitin-active AA10 (38) (Fig. 3A). Previous studies have shown that this residue is important for chitin binding (38)(39)(40). AoAA11 carries two insertions relative to AfAA11A, which both have unknown locations as a major part of both of these is lacking in the AoAA11 crystal structure, as shown in Figure 3A. From the location of these insertions, it seems unlikely that they are located in the same area as the Tyr25-containing insertion in AfAA11A. Next to most of these two insertions, the AoAA11 structure lacked additional residues (151-161) that are resolved in the AfAA11A structure (residues 144-154), showing that these are located far away from the copperbinding site and the substrate-binding surface. Importantly, the insertions in AoAA11 and the additional region that is not visible in the AoAA11 structure contain a tyrosine or other aromatic residues.
Although some uncertainty exists regarding the surface of AoAA11, it is clear that the two AA11 enzymes have very different substrate-binding surfaces, as illustrated in Figure 3C, which shows that the substrate-binding surfaces of the two enzymes differ in both shape and electrostatic potential. Interestingly, both AA11 enzymes as well as CBP21 contain a small pocket close to the histidine brace, which has been previously associated with chitin-active LPMOs (41,42). While it has been proposed that this pocket accommodates an acetyl group (42), recent modeling data suggest that this pocket may give room to O 2 or H 2 O 2 in chitin-active LPMOs (43) as in accordance with suggestions by Hemsworth et al. (44).

Substrate specificity
The substrate specificity of AfAA11A was assessed by incubating the LPMO overnight with a variety of substrates in the presence of 1 mM ascorbic acid (AscA). Chromatographic analyses did not reveal any reductant-dependent product formation in reactions with phosphoric acid-swollen cellulose (PASC), Avicel, tamarind xyloglucan, birchwood xylan, beechwood xylan, acetyl glucuronoxylan from aspen, ivory nut mannan, acetylated konjac glucomannan, potato starch, heparin, hyaluronic acid, and chitosan. In contrast, soluble reaction products were observed for the reactions with chitinous substrates (α-chitin from shrimp shell and β-chitin from squid pen), and only if AscA was present (Fig. 4). Both chromatographic and MS analyses of the products generated from α-chitin and β-chitin showed that these were C1-oxidized chito-oligomers with a degree of polymerization 2 to 8 ( Fig. 4C) (45). The mass spectra showed clusters of signals that are typical for C1-oxidized oligomers, including the diagnostic peak corresponding to the Na + or K + salts of aldonic acids. For example, in the degree of polymerization 6 cluster (Fig. 4B), we observed signals corresponding to the Na + adducts of the lactone (m/z 1257.375) and aldonic acid (m/z 1275.378) forms a Values for outer shell are given in parenthesis.
d R free is the R cryst value calculated on the 5% reflections excluded for refinement.

Structural and functional variation among AA11 LPMOs
and of the sodium salt of the aldonic acid (m/z 1297.356) as well as the K + adducts of the aldonic acid (m/z 1291.343) and of the aldonic acid binding one Na + and one K + ion (m/z 1313.315).

Degradation of chitin in O 2 -driven reactions
The ability of AfAA11A to degrade α-chitin and β-chitin under conditions that are commonly used when studying LPMOs (aerobic reactions, 1 mM AscA) was initially tested at temperatures between 30 and 45 C, to find a temperature giving stable reaction kinetics that could be used in subsequent experiments. The product accumulation over time indicates not only faster LPMO rates but also increased nonlinearity, with increasing temperatures (Fig. 5, A and B). While product formation was linear for the first 6 h at all the three tested (Protein Data Bank ID: 4MAH) generated with the PyMOL alignment tool at default settings. The alignment file was further processed using the ESPript 3 server and Adobe Illustrator. The three pairs of cysteines forming disulfide bridges are labeled with green numbers. Residues missing in the crystal structure of AoAA11 are highlighted by gray shading. Secondary structure elements are indicated above (AfAA11A) and below (AoAA11) the sequence, as follows: α, alpha helix; η, 3 10 helix; β, beta-strand; T, turn. B, topology diagram of AfAA11A. C, tertiary structure of AfAA11A, with the same color coding as in panel B; secondary structure elements that are connected by a disulfide bridge (yellow; numbering as in panel A) appear in the same color (red, blue, or green). Panel C also shows the side chains of the two histidines of the copper-binding His brace (H1 and H71), a buried tyrosine beneath the histidine brace (Y133), and a surface-exposed tyrosine (Y25). temperatures (Fig. 5C), only in reactions run at 30 C, the product formation rate was stable during the full 24 h of the reaction. Hence, all subsequent reactions in this study were run at 30 C. Figure 5 also shows that AfAA11A is somewhat more active on β-chitin compared with α-chitin, both in terms of the initial rate and the product yield after 24 h. Binding studies showed that AfAA11A binds better to β-chitin compared with α-chitin (Fig. 5D), which could explain the higher activity toward the former substrate. It is worth noting that the progress curves shown in Figure 5 are similar to those typically obtained with chitin-active bacterial AA10 LPMOs when using similar reaction conditions (e.g., (8)). The low catalytic rates that can be estimated from the curves shown in Figure 5, A-C (in the range of 0.002-0.01 s −1 ) are commonly observed for LPMOs in AscA-/O 2 -driven reactions (18). Progress curves generated after storing the copper-saturated enzyme for 15 months in 50 mM bis-Tris/HCl, pH 6.5, at 4 C were similar to those shown in Figure 5, indicating that purified AfAA11A had good storage stability.

Degradation of chitin in H 2 O 2 -driven reactions
Recent studies have shown that feeding LPMOs with H 2 O 2 may lead to fast turnover of chitin or cellulose (16,20), and one study, describing a novel assay to detect LPMO-generated carboxyl groups in chitin (46), has indicated that this may also be the case for members of the AA11 family. Therefore, we analyzed if and how H 2 O 2 could be used to boost the catalytic activity of AfAA11A. For our experiments, we repeatedly added varying amounts of H 2 O 2 to reactions containing the substrate, LPMO, and AscA. Thus, we mimicked a situation where H 2 O 2 is delivered even more gradually, for example, by a H 2 O 2 -generating enzyme (47) or through continuous feeding with a pump, which has shown to be highly effective for cellulose-active LPMOs (16,48,49). When H 2 O 2 was added in Figure 3. Comparison of chitin-active LPMOs. A, the structures of AfAA11A, AoAA11, and CBP21, colored by secondary structure (helices red, strands yellow) and with additional coloring according to the sequence alignment underneath, showing side chains of residues involved in copper coordination and substrate binding (see text for details). The 11-residue insertion in AfAA11A relative to AoAA11 that includes Tyr25 appears in pink, and residues adjacent to this insertion are labeled. The 14-residue insertion near residue 100 in AoAA11 appears in blue, and residues adjacent to the insertion are labeled in both AA11 structures; note that 11 of these residues (gray shading in the sequence) could not be modeled in the AoAA11 structure. Eight of the nine residues forming an insertion near residue 165 in AoAA11 as well as 11 preceding residues could not be modeled in the AoAA11 structure; this 19-residue region is colored green, and adjacent residues are labeled. B, superposition of AfAA11A (green carbons) and AoAA11 (gray carbons); the picture shows side chains in and near the metal-binding site and a zinc atom bound to AoAA11 (purple sphere). C, a view on the putative substrate-binding surfaces of the three LPMOs shown in panel A, colored by electrostatic surface potential calculated using PyMOL. Note that the AoAA11 structure is incomplete and that the real surface thus may look differently (see main text). The arrows point at a small pocket near the catalytic center that seems to be conserved in chitin-active LPMOs, across LPMO families. LPMO, lytic polysaccharide monooxygenase.
lower concentrations (20-50 μM H 2 O 2 ; Fig. 6A), the accumulation of soluble oxidized products correlated well with the amount of H 2 O 2 added, indicating that H 2 O 2 was consumed productively between additions. After the H 2 O 2 feeding phase (Fig. 6A), the LPMO reaction proceeded at the same rate as in the control reaction with only AscA, confirming that H 2 O 2 had been consumed during the feeding phase and that the LPMO was still active. Increasing the amount of H 2 O 2 (80-200 μM per addition; Fig. 6B) gave higher initial product formation rates combined with increasingly rapid cessation of product formation. The observation that the LPMOs become inactivated when supplied with (too) high amounts of H 2 O 2 is in accordance with observations made for LPMOs in the AA9 and AA10 families (16,48).
To get an impression of the maximum rate of H 2 O 2 consumption by the LPMO, we ran reactions with reduced LPMO concentrations. The results (Fig. 6C) show that, at LPMO concentrations below 0.2 μM, the amount of LPMO became limiting for the reaction. On the other hand, dose-response effects were minimal at LPMO concentrations of 0.2 μM and higher, suggesting that in all these reactions, H 2 O 2 consumption was complete and that H 2 O 2 was limiting. The apparent initial rates that can be derived from the progress curves for the reactions with 0.05 and 0.1 μM LPMO are in the range of 0.15 to 0.20 s −1 , which is 1 to 2 orders of magnitude higher than the apparent rates observed in standard reactions with only AscA and no added H 2 O 2 . Of note, the apparent rates of AfAA11A are likely underestimated because a significant fraction of the oxidized sites, perhaps up to 50% (50), may remain in the solid residue after saccharification. It is also worth noting that the linear progress curve for the reaction with 0.2 μM AfAA11A in Figure 6C, where the level of soluble product reaches 250 μM after 4 h, implies that, in this experimental setting, one LPMO molecule can turnover at least some 1250 molecules of H 2 O 2 without becoming inactivated.

Synergy with a chitinase
The ability of AfAA11A to boost chitinase efficiency was assessed by monitoring the degradation of α-chitin and βchitin by a combination of an endochitinase, chitinase C from SmChiC (ChiC), and AfAA11A in a 1:1 M ratio (Fig. 7). The results show that AfAA11A promotes the activity of SmChiC on both α-chitin and β-chitin. When acting individually on α-chitin for 24 h (Fig. 7A), the proteins were able to solubilize 1.7 and 2.6 mM GlcNAc of the substrate, respectively, while when combined, 12.6 mM GlcNAc was produced. In the reactions with α-chitin and SmChiC only, product formation stopped after some 9 h, whereas linear product formation was achieved for 24 h in the reaction that also contained the LPMO.
Synergistic effects were also observed in reactions with βchitin (Fig. 7B). Compared with the reactions with α-chitin, these effects were smaller, which may be due to the fact that the chitinase alone was much more efficient on β-chitin (Fig. 7B) than on α-chitin (Fig. 7A). The initial rate of product formation by SmChiC was little affected by the addition of AfAA11A. On the other hand, in the reaction where SmChiC was supplemented with the LPMO, the high initial rate (as observed with SmChiC alone) was maintained for a longer time, and, consequently, higher product levels were reached earlier on in the reaction. Final saccharification yields after 24 h were also affected by the LPMO. While SmChiC and AfAA11A alone were able to produce 54.9 and 2.7 mM GlcNAc, respectively, 84.6 mM GlcNAc was produced in the reaction containing both enzymes.
To highlight the impact of the LPMO in chitin depolymerization, and to verify whether the cessation of product formation in reactions with SmChiC was caused by enzyme inactivation, denaturation, or depletion of accessible substrate, reactions were carried out in which the LPMO was added after 6 h, instead of at the start of the reaction. This experiment showed that chitin solubilization by SmChiC was enhanced upon addition of the LPMO (Fig. 7, A and B) and that, in these reactions, product formation continued for the remaining reaction period. Thus, the gradual cessation of product formation in reactions with only SmChiC is due to the enzyme not being able to access the remaining substrate and not to enzyme inactivation.

Discussion
The crystal structure of AfAA11A is only the second crystal structure of a member of the widely spread family of fungal AA11 LPMOs and the first one that shows all amino acids. The AfAA11A structure lacks a metal ion, which is remarkable considering the reported high metal affinities of these enzymes  (14, 15) but not uncommon (e.g., (38)). Nevertheless, the catalytic centers of AfAA11A and AoAA11, which has a zinc ion bound, look similar (Fig. 3B). Comparison of the structures of AfAA11A and AoAA11 beyond the catalytic centers is complicated because the latter structure lacks two regions comprising 30 residues in total. Residues 144 to 154 in the structure of AfAA11A provide structural information for residues 151 to 161 lacking in the structure of AoAA11. It is somewhat remarkable that these residues form an α-helix (α1; Fig. 2), since the fact that these residues could not be modeled in the AoAA11 structure would suggest that they are part of a flexible loop-like structure.
Even when taking into account the lacking information for AoAA11, it seems safe to conclude from the comparison of the two structures (Fig. 3) that the two enzymes have different substrate-binding surfaces. Because of the Tyr25-containing insertion that is unique for AfAA11A, the substrate-binding surface of this enzyme shows features that are characteristic for chitin-active AA10 LPMOs, such as CBP21, in particular the presence of exposed Tyr25 (Fig. 3). Experimental (38) and modeling (40) studies have shown that a tyrosine in this position is important for binding of crystalline chitin. As alluded to above, it is not possible to envisage a similar structural arrangement in AoAA11 or in previously characterized FfAA11 (31), which both lack this insertion. Interestingly, these structural differences seem to correlate with functional properties. On the one hand, we show here that the catalytic activity of AfAA11A and its ability to boost enzymatic chitin conversion are similar to what has been observed for bacterial chitin-active AA10 LPMOs. On the other hand, a recent study has shown that another AA11 LPMO from A. fumigatus, AfAA11B, which is a close homolog of AoAA11, with similar insertions and deletions relative to AfAA11A, has negligible activity toward chitin (32). Of note, AfAA11B shows high activity toward soluble chito-oligomers; the biological role of this activity remains enigmatic (32).
Our data show that AfAA11A has C1-oxidizing activity leading to production of lactone products that are in equilibrium with their aldonic acid forms. It is worth mentioning that the MALDI-TOF MS analyses suggested the presence of considerable amounts of lactones (M-2 Da) in reactions with both α-chitin and β-chitin, which could represent C4-oxidized products (since C1-oxidized lactones are expected to be hydrated to a larger extent (51)). While this observation, which is similar to observations by Wang et al. (31) for FfAA11, could indicate the presence of C4-oxidized reaction products, we did not detect unusual, that is, possibly C4-oxidized products in either hydrophilic interaction liquid chromatography (Fig. 5C) or high-performance anion exchange chromatography with pulsed amperometric detection (not shown). Compared with α-chitin, reactions with β-chitin showed both better binding at 30 C and more product formation at all three tested reaction temperatures (Fig. 5). Other chitin-active LPMOs show similar differences regarding binding to and activity on these two chitin allomorphs (45,52).
Since the discovery that LPMOs can use H 2 O 2 as a cosubstrate in 2017 (16), multiple studies have demonstrated that addition of H 2 O 2 to LPMO reactions leads to increased catalytic rates (16,20,22,46,48,53). Indeed, and in accordance with previous observations for FfAA11 (46) and AfAA11B (32), we observed increased catalytic rate upon adding H 2 O 2 to reactions with AfAA11A. The apparent catalytic rates of AfAA11A were in the range of 0.003 to 0.012 s −1 for the O 2driven reaction and in the range of 0.1 to 0.2 s −1 for the H 2 O 2driven reaction. An earlier study on FfAA11A, using the similar reaction conditions (10 g/l α-chitin, 2 μM LPMO, and 1 mM AscA) reported apparent rates of 0.5 min −1 (0.01 s −1 ) under standard aerobic conditions without H 2 O 2 and a 16-fold increase (to 0.11 s −1 ) when 100 μM H 2 O 2 was supplied (46). Of note, some claim that the so-called "O 2 -driven" reaction under standard aerobic conditions may be limited by the in situ generation of H 2 O 2 , which is needed to fuel the LPMO peroxygenase reaction (see e.g., (16,54), and see (19) for another opinion). The apparent rates for the peroxygenase reaction are much lower than a k cat value reported for CBP21 acting on chitin (6.7 s −1 ; (20)). This may reflect a true difference in enzyme efficiency, but it is important to note that the peroxygenase rate of AfAA11A is underestimated because of noncontinuous product monitoring, which may hide possibly much higher rates that are achieved right after each addition of H 2 O 2 . Additional kinetic studies are needed to determine the true maximum rate of AfAA11A.
The experiments depicted in Figure 6 show that the AfAA11A peroxygenase reaction may be limited by H 2 O 2 (Fig. 6A) or by the LPMO concentration (Fig. 6C), depending on how the reaction is set up. The H 2 O 2 -limited reactions in Figure 6A show a strong correlation between the amount of added H 2 O 2 and the amount of soluble oxidized products. In each of the three reactions with added H 2 O 2 , the level of oxidized products at the end of the H 2 O 2 -feeding period (and corrected for product levels generated in the control reaction with reductant only) correspond to some 60 to 65% of the total amount of added H 2 O 2 . It is well known from the literature that a considerable portion, in some cases up to 50%, of the oxidized sites introduced by LPMOs remain on the crystalline substrate (55), which may explain the discrepancy between the amount of added H 2 O 2 and the amount of soluble oxidized products.
As expected for LPMOs (and other enzymes with active redox centers), excessive H 2 O 2 concentrations lead to enzyme inactivation of LPMOs (16,56). Figure 6B shows that higher H 2 O 2 concentrations lead to higher initial enzyme rates and more rapid enzyme inactivation, as has been demonstrated for multiple LPMOs, including FfAA11 (46). Importantly, at lower H 2 O 2 concentrations, the enzyme seems stable, and we show that one enzyme molecule on average catalyzes at least 1250 reactions (note that this by no means is a maximum or an optimized value). This value compares well with previously published values of ≥300 (for AfAA11B acting on soluble substrate, (32)) and ≥1500 for the LPMO in a commercial cellulase cocktail (16).
Although the progress curves of Figure 5 (i.e., activity of the LPMO only) showed higher activity of AfAA11A on β-chitin compared with α-chitin, the boosting effect of the LPMO on chitinase activity was more pronounced for α-chitin (Fig. 7). αchitin is more crystalline and resilient to enzymatic degradation than β-chitin (57), and it is thus conceivable that the addition of an enzyme specialized in acting on crystalline material, that is, the LPMO, has a larger effect in reactions with α-chitin. Indeed, previous studies have shown that the extent by which an LPMO can speed up a chitinase reaction depends on substrate crystallinity (58).
In conclusion, we present the first structure of an AA11 LPMO in which all amino acids are included and show that there is considerable structural variation within the AA11 family that is correlated to functional variation. It is worth noting that the disclosed variation within the AA11 family really is remarkable. First, the structural differences in the substrate-binding surfaces seem larger than the differences seen within other LPMO families. Second, while AfAA11A degrades chitin with commonly observed reaction rates and yields, AfAA11B does not. We also show that AfAA11A can efficiently utilize H 2 O 2 as a cosubstrate and is not inactivated by acting in the "peroxygenase" mode, as long as H 2 O 2 levels are not too high. AfAA11A enhanced the solubilization of βchitin and α-chitin by SmChiC, suggesting that this LPMO may have a role in fungal chitin degradation and may be of use in the enzymatic processing of chitin-rich biomass.

Experimental procedures Reagents and substrates
All common reagents used in this study were supplied by Sigma-Aldrich and Merck Millipore, whereas Bacto Peptone and Bacto Yeast Extract were supplied by BD Biosciences, and acetonitrile was supplied by VWR. Alkaline-and acidpretreated shrimp shell α-chitin originating from Pandalus borealis was purchased from Chitinor AS and ball-milled using a Planetary Ball Mill PM100 (Retsch) equipped with a stainless-steel container and zirconium dioxide beads. Milling was performed at 450 rpm in three 5-min rounds with 2-min intervals, yielding a final particle size of 0.2 mm. β-chitin originating from squid pen (batch: 20140101) with an average particle size of 0.8 mm was purchased from France Chitine.

Protein production and purification
The AfAA11A protein was produced through heterologous expression using a P. pastoris yeast expression system (PichiaPink Strain 4; Thermo Fisher Scientific). A synthetic gene encoding AfAA11A (UniProt ID: Q4WF00; gene ID: AFUA_3G03950) including its native signal peptide was codon-optimized for P. pastoris and synthesized by GenScript and inserted behind the GAP promoter and a P. pastorisspecific Kozak sequence in the pPink-GAP vector by restriction cloning with EcoRI and Acc65I (New England BioLabs, Inc). The generated plasmid was linearized with AflII (New England BioLabs, Inc) and transformed into electrocompetent P. pastoris cells, after which LPMO-producing transformants were screened, selected, and stored as previously described (64).
For protein production, a single colony from a yeast extract-peptone-dextrose agar plate was used to inoculate 50 ml of buffered glycerol complex medium (Invitrogen) in a 250-ml Erlenmeyer flask with vent cap, which was then incubated overnight at 29 C and 200 rpm. The overnight culture was transferred to 400 ml of fresh buffered glycerol complex medium, followed by incubation overnight in a 2-l baffled shake flask with vent cap. After 24 h, the culture was supplemented with 1% (v/v) glycerol. After 48 h, the culture supernatant containing the LPMO was separated from the yeast cells by centrifugation at 10,000g for 20 min at 4 C and subsequentially filtered through a 0.45 μm polyether sulfone (PES) membrane (VWR). The presence of the LPMO was confirmed by SDS-PAGE gel analysis using Mini PROTEAN TGX Stain-Free precast gels, and fluorescence was recorded with a Gel Doc EZ Imager using the Image Lab, version 6.0.0 Standard Edition software (Bio-Rad). The culture supernatant was concentrated approximately fivefold with concomitant buffer exchange to 50 mM bis-Tris/HCl buffer, pH 6.5, using a Vivaflow 200 tangential crossflow concentrator equipped with a 10,000 molecular weight cutoff (MWCO) PES membrane (Sartorius Stedim Biotech).
For hydrophobic interaction chromatography, the concentrated culture supernatant was supplemented with ammonium sulphate to a final concentration of 2 M and centrifuged for 15 min at 10,000g and 4 C to remove precipitation. The supernatant was then applied onto a 5 ml HiTrap Phenyl FF (HS) column (GE Healthcare Lifesciences), mounted on an ÄKTA Prime Plus chromatography system and equilibrated in 50 mM bis-Tris/HCl buffer (pH 6.5) containing 2 M ammonium sulphate. Proteins were eluted by applying a 35 ml linear gradient toward 50 mM bis-Tris/HCl buffer, pH 6.5. Fractions containing AfAA11A were identified by SDS-PAGE, pooled, and concentrated using a Vivaspin 20 Centrifugal Concentrator 10,000 MWCO PES (Sartorius Stedim Biotech). The recombinant protein was further purified by size-exclusion chromatography using a HiLoad 16/600 Superdex 75 pg column (GE Healthcare) mounted on an ÄKTA purifier chromatography system (GE Healthcare), using 50 mM bis-Tris/ HCl, pH 6.5, 150 mM NaCl as eluent, and a flow rate of 1 ml⋅min −1 . Fractions containing pure LPMO were identified by SDS-PAGE and pooled, followed by concentration and buffer exchange to 50 mM bis-Tris/HCl buffer, pH 6.5, using a Vivaspin 20 Centrifugal Concentrator 10,000 MWCO PES.
To saturate the LPMO with Cu(II), the purified protein was incubated with 10-fold molar excess of CuSO 4 at 4 C for 4 h, then desalted on a Vivaspin 20 Centrifugal Concentrator 3000 MWCO PES by five cycles of concentrating and rediluting the protein solution in 50 mM bis-Tris/HCl buffer, pH 6.5 (ca. 10,000-fold dilution). The final protein solution was sterilized by filtration through a 0.22 μm Millex-GV syringe filter (Merck Millipore) and stored at 4 C.
Protein concentrations of solutions containing pure AfAA11A were determined based on absorbance at 280 nm with an Eppendorf Biophotometer D30 (Eppendorf), using the theoretical extinction coefficient of 38,765 M −1 cm −1 , calculated with the ExPASy ProtParam tool (65). The mature protein consists of 201 amino acid residues with a calculated molecular mass of 21,750 Da and a theoretical pI of 4.76 (65).

Protein crystallization
N-linked glycosylations were removed from a batch of purified and copper-saturated AfAA11A by treating the LPMO (140 μM) with an in-house-produced recombinant endo-β-Nacetylglucosaminidase (EfEndo18A, 3.4 μM; (35)) in 50 mM sodium acetate buffer (pH 6.0). The reaction (with 5 ml total volume) was incubated first at 30 C for 1 h and then at 4 C overnight. Deglycosylation was verified by SDS-PAGE, and the EfEndo18A protein was removed by hydrophobic interaction chromatography, as described previously. Fractions containing the LPMO were pooled and concentrated, and the buffer was exchanged to 50 mM bis-Tris/HCl buffer (pH 6.5) using a Vivaspin 20 Centrifugal Concentrator 10,000 MWCO PES. The resulting protein was saturated with Cu(II) and subsequentially desalted, filter sterilized, and stored according to the protocol described previously. Deglycosylated AfAA11A was successfully crystallized using conditions 1 to 15 from the commercial crystal screening kit JCSG-plus MD 1-37 Box 1 (Molecular Dimensions) in a hanging drop setup using a 24well VDX plate with sealant and 18 mm circle glass cover slides (Hampton Research). The mother liquor was made inhouse and consisted of 0.1 M bicine (pH 9.0) and 20% (w/v) PEG 6000. The best crystals were produced in droplets obtained by mixing 0.5 μl of 20 mg⋅ml −1 de-N-glycosylated AfAA11A with 1 μl mother liquor and having 200 μl mother liquor in the well. The crystallization setups were stored in the dark at room temperature, and protein crystals were observed after approximately 3.5 months.

X-ray crystallography
Protein crystals were soaked in a cryosolution containing mother-liquor with 35% glucose (w/v) before they were flash frozen in liquid nitrogen. Diffraction data were collected at the BioMAX beamline at MAX IV. Datasets were processed by XDS (66) and scaled by AIMLES (67). The CCP4i package was used to solve the structure by molecular replacement (Phaser) (68), and the structure was refined using REFMAC (69). Model manipulations were carried out using Coot (70), and molecular graphics was generated using PyMOL Molecular Graphics System, version 2.0 (Schrödinger, LLC).

Substrate specificity
The activity of AfAA11A was screened toward the following substrates: PASC, Avicel, tamarind xyloglucan, birchwood xylan, beechwood xylan, acetyl glucuronoxylan from aspen, ivory nut mannan, acetylated konjac glucomannan, potato starch, heparin, hyaluronic acid, chitosan from shrimp shell, α-chitin from shrimp shell, and β-chitin from squid pen. Reaction mixtures were set up in 100 μl total volume, contained 1 μM LPMO, 0.2 to 0.6% (w/v) substrate, and 1 mM AscA in 50 mM bis-Tris/HCl buffer, pH 6.5, and were incubated overnight at 30 C and 1000 rpm in an Eppendorf Thermomixer C (Eppendorf). In negative control reactions, AscA or the LPMO was substituted with Milli-Q water. At the end of the reaction, insoluble substrates were removed by filtration using a 0.2 μm PES 96-well filter plate (Millipore) operated in a vacuum manifold, and the soluble fraction was subsequently analyzed as outlined later.

Chitin degradation experiments
In general, reactions, with a total volume of 300 μl, were set up with 1% (w/v) substrate and 1 μM LPMO and 1 mM AscA in 50 mM bis-Tris/HCl buffer, pH 6.5, and were incubated at 30 C and 1000 rpm in thermomixers (Eppendorf Thermomixer C; Eppendorf) in standard aerobic conditions for up to 24 h. Samples (30 μl) were taken periodically, and the reaction was terminated by removing the insoluble substrate by filtration using a 0.2 μm PES 96-well filter plate (Millipore) operated in a vacuum manifold. All reactions were performed in triplicates.
When assessing synergy between chitinases and the LPMO, reactions were set up with various combinations of 1 μM LPMO (AfAA11A) and 1 μM SmChi18C, a nonprocessive endochitinase from S. marcescens (61) (30 μl) in order to maintain the reaction volume at 300 μl. Dilution in the amounts of products formed because of sampling and addition of reactant were taken into account when calculating product levels.

Substrate binding
To assess binding to chitin, 3.0 μM AfAA11A was incubated with 0.2% (w/v) α-chitin or β-chitin in 50 mM bis-Tris/HCl buffer, pH 6.5, at 30 C and 1000 rpm. At different time points, the insoluble substrate was removed by filtration using a 96well 0.2 μm PES filter plate (Millipore) installed in a vacuum manifold. The amount of LPMO remaining in solution was visualized on Mini PROTEAN TGX Stain-Free SDS-PAGE gels (Bio-Rad) with a Gel Doc EZ Imager (Bio-Rad) based on fluorescence signals and quantified with the Image Lab software, version 6.0.0. Solutions with known concentrations of AfAA11A were used as standard.

Analysis and quantification of reaction products
Product formation in reactions with cellulosic and hemicellulosic substrates was assessed by high-performance anion exchange chromatography with pulsed amperometric detection using an ICS5000 system (Dionex) equipped with Car-boPac PA1 analytical (2 × 250 mm) and guard (2 × 50 mm) columns, using a 50 min gradient protocol as previously described (71,72). Product formation in reactions with chitinous substrates was assessed by hydrophilic interaction liquid chromatography, using an UPLC (Infinity 1290; Agilent Technologies) equipped with an Acquity BEH Amide Column (130 Å, 1.7 μm, 2.1 × 150 mm) combined with a VanGuard Precolumn (Waters Corporation). The samples were adjusted to 74% acetonitrile prior to analysis, and oxidized chitooligosaccharides were separated using a 12 min gradient as reported earlier (50). For the identification of produced native and oxidized chito-oligosaccharides, standards were produced as previously described (62). To quantify conversion yields in the chitin degradation assays, soluble products (GlcNAc and GlcNAcGlcNAc1A) were analyzed using a Dionex Ultimate 3000 RSLC system (Dionex) equipped with a Rezex RFQ-Fast Acid H+ (8%) column (7.8 × 100 mm; Phenomenex) operated at 85 C (8). Data were acquired and analyzed using Chromeleon, version 7.2.9 (Thermo Fisher Scientific).
Oxidized products generated from chitin were also analyzed with MALDI-TOF MS using an Ultraflex MALDI-TOF/TOF instrument (Bruker Daltonics) equipped with a 337 nm nitrogen laser. The samples were mixed in a 1:1 ratio with a matrix solution consisting of 9 mg ml −1 2,5-dihydroxybenzoic acid and 30% acetonitrile and subsequently applied on an MTP 384TF ground steel target plate (Bruker Daltonics). Data acquisition and analysis were performed using FlexControl, version 3.4 and FlexAnalysis, version 3.4 (Bruker Daltonics), respectively.

Data availability
All data are contained within the article. The crystal structure of AfAA11A has been deposited in the Protein Data Bank under the accession code 7P3U. Conflict of interest-The authors declare that they have no conflicts of interest with the contents of this article.