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Multi-point precision binding of substrate protects LPMOs from self-destructive off-pathway processes Jennifer Sarah Maria Loose, Magnus Øverlie Arntzen, Bastien Bissaro, Roland Ludwig, Vincent G.H. Eijsink, and Gustav Vaaje-Kolstad Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b00484 • Publication Date (Web): 14 Jun 2018 Downloaded from http://pubs.acs.org on June 14, 2018

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Biochemistry

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Multi-point precision binding of substrate protects LPMOs from self-destructive off-

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pathway processes

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Jennifer S.M. Loose¹, Magnus Ø. Arntzen¹, Bastien Bissaro¹, Roland Ludwig², Vincent G.H.

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Eijsink¹ and Gustav Vaaje-Kolstad¹*

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¹Faculty of Chemistry, Biotechnology and Food Science, Norwegian University of Life Sciences

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(NMBU), Ås, Norway.

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²BOKU - University of Natural Resources and Life Sciences, Department of Food Sciences and

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Technology, Biocatalysis and Biosensing Laboratory, Vienna, Austria.

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*To whom correspondence should be addressed: Gustav Vaaje-Kolstad, Faculty of Chemistry,

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Biotechnology, and Food Science, The Norwegian University of Life Sciences (NMBU), 1432

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Ås, Norway, Tel.: +47 67232573; E-mail: [email protected]

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ABSTRACT

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Lytic polysaccharide monooxygenases (LPMOs) play a crucial role in the degradation of

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polysaccharides in biomass by catalyzing powerful oxidative chemistry using only a single

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copper ion as co-factor. Despite the natural abundance and importance of these powerful mono-

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copper enzymes, the structural determinants of their functionality have remained largely

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unknown. We have used site-directed mutagenesis to probe the roles of 13 conserved amino

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acids located on the flat substrate-binding surface of CBP21, a chitin-active family AA10 LPMO

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from Serratia marcescens, also known as SmLPMO10A. Single mutations of residues that do not

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interact with the catalytic copper site, but are involved in substrate-binding had remarkably large

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effects on overall enzyme performance. Analysis of product formation over time showed that

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these mutations primarily affected enzyme stability. Investigation of protein integrity using

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proteomics technologies showed that loss of activity was caused by oxidation of essential

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residues in the enzyme active site. For most enzyme variants, reduced enzyme stability

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correlated with reduced binding to chitin, suggesting that adhesion to the substrate prevents

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oxidative off-pathway processes that lead to enzyme inactivation. Thus, the extended and highly

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evolvable surfaces of LPMOs are tailored for precise multi-point substrate binding, which

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provides the confinement that is needed to harness and control the remarkable oxidative power of

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these enzymes. These findings are important for optimized industrial use of LPMOs as well as

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design of LPMO-inspired catalysts.

37 38 39

Keywords: Lytic polysaccharide monooxygenases, protein stability, mass spectrometry


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Biochemistry

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INTRODUCTION

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Lytic polysaccharide monooxygenases (LPMOs) are mono-copper redox enzymes that catalyze

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oxidative cleavage of glycosidic bonds in polysaccharides.1-5 Based on sequence similarity, these

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enzymes are divided into five different families of the auxiliary activities (AA) in the CAZy

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database, AA9, AA10, AA11, AA13 and AA14.6 LPMOs bind a single copper ion, which is

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coordinated by two histidine residues in a so-called histidine brace,3 bearing some similarity to

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the copper-binding site of particulate methane monooxygenases.7 The ubiquitous occurrence of

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LPMOs indicates that they are of major importance, as is also suggested by observed high

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expression levels in bacteria and fungi that degrade recalcitrant biomass.8-14 It is well established

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that LPMOs act synergistically with glycoside hydrolases and, thus, boost the enzymatic

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conversion of polysaccharides.1, 15-17

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Since their discovery in 2010, substantial efforts have been made to shed light on LPMO

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functionality and the underlying reaction mechanism.18-21 There has been consensus that catalysis

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by LPMOs requires two externally delivered electrons and involves the activation of dioxygen.1,

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22

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hydrogen peroxide is an alternative, and likely even preferred, co-substrate.23, 24 In a hydrogen

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peroxide-based catalytic mechanism electrons are only needed to activate the LPMO by reducing

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its Cu(II) to Cu(I), after which the enzyme can carry out multiple reactions using H2O2 as co-

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substrate (Fig. 1). Notably, under the conditions normally used for measuring LPMO activity,

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H2O2 will be formed from O2 at the expense of two electrons, either by reduced LPMO

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molecules that are not bound to substrate25, 26 or by reactions involving molecular oxygen and the

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reductant. LPMO reactions can be driven by a variety of reductants,27,

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molecules like ascorbate and redox enzymes like cellobiose dehydrogenase (CDH). These

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reductants show different efficiencies, which is normally ascribed to varying abilities to reduce

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the LPMO, but which could also be due to variation in the generation and stability of H2O2

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during the reaction. Clearly, assessing LPMO functionality is not straightforward, which likely

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explains in part why so little structure-function data are available for these enzymes.

Recently, this monooxygenase model of catalysis has been challenged by a study showing that


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including small

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Figure 1. Reactions involved in H2O2-driven catalysis by CBP21. In the absence of a reducing agent,

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CBP21 is present in an oxidized state, carrying a Cu(II) ion bound in the active site. In the presence of a

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reducing agent the active site copper is reduced to Cu(I) (a), activating/ priming the enzyme for catalysis.

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The primed enzyme may cleave chitin chains by binding to the surface of crystalline chitin and utilizing

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H2O2 as a co-substrate in the reaction (b). Once primed, CBP21 can perform several catalytic events

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without the need of being reduced since no external electrons are needed in this stage of the reaction (c).

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The H2O2 used by CBP21 can either come from the oxidase activity of free reduced CBP21 (d) or from

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auto-oxidation of the electron donor, an event that can be catalyzed by the presence of transition metals

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(M) in the solution (e). If a reduced CBP21 reacts with H2O2 in the absence of substrate, oxidation of the

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active site may occur, inactivating the enzyme (f). In the originally proposed LPMO mechanism, O2 is

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used directly as a co-substrate and two electrons need to be delivered to the catalytic center per catalytic

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cycle, as extensively reviewed in e.g.

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driven reactions are much slower than H2O2-driven reactions.23, 24, 29

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. This mechanism may still apply, but it is now clear that O2-

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Biochemistry

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Although some structure-function studies on LPMOs have been published,1, 16, 30, 31 information

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on the functional roles of conserved residues involved in catalysis and/or substrate binding

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remains limited. Importantly, the characterization of the few CBP21 variants published so far

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was such that possible kinetic complications related to e.g. enzyme inactivation or hydrogen

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peroxide production were overlooked. In this respect, it is worth noting that LPMOs are

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notoriously unstable under commonly used reaction conditions (e.g.

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oxidative self-destruction.23 Studying CBP21, a chitin-active LPMO from Serratia marcescens,

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also known as SmLPMO10A, Loose et al. 33 showed that stable reaction kinetics can be obtained

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if reducing power is delivered gradually, e.g. through the action of CDH. As expected, the rate of

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the CDH-driven LPMO catalysis was similar to the rate of substrate oxidation by CDH.

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The majority of substrates targeted by LPMOs are insoluble and successful oxidation of

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glycosidic bonds can only be accomplished through productive binding to an ordered assembly

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of polysaccharide chains. LPMOs have relatively flat substrate binding surfaces34, 35 that seem

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capable of interacting with multiple polysaccharide chains, likely enabling the enzymes to

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interact with the ordered surface of a crystalline substrate. Interactions with polysaccharide

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chains involve several polar interactions, whereas aromatic side chains contribute via aromatic-

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carbohydrate π-CH interactions, especially for cellulose-active LPMOs.20,

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oxidative activity was unraveled, substrate binding by CBP21 was studied by site-directed

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mutagenesis.34 This early work showed that, next to a solvent exposed tyrosine, at least four

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polar amino acids are important for substrate binding. The importance of these polar residues in

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chitin binding by CBP21 has been confirmed by NMR studies.37

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Building on these initial studies, we have carried out a site-directed mutagenesis study of the

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roles of conserved residues in the substrate binding surface of CBP21. Mutational effects were

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characterized by analyzing substrate binding and by measuring catalytic activity over time using

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a small molecule reductant or a redox enzyme (CDH) for delivery of electrons to the system. The

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results provide insight into residues that affect catalytic activity, in particular residues in the

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primary and secondary coordination sphere of the copper ion. Importantly, however, the most

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commonly observed mutational effect was decreased resistance of the enzyme against oxidative

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self-destruction, underpinning the importance of proper kinetic characterization and shedding

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new light on the role of the extended substrate-binding surface of LPMOs. Using proteomics 5 ACS Paragon Plus Environment

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), likely as a result of

21, 34, 36-38

Before its

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technologies oxidative damage to the active site of the enzyme variants was characterized in

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detail. Thus, this study sheds new light on LPMO functionality and highlights the crucial

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importance of fine-tuned enzyme-substrate interactions.

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Biochemistry

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METHODS

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Cloning, Site-directed Mutagenesis, Protein Expression and Purification. Cellobiose

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Dehydrogenase from Myriococcum thermophilum (MtCDH) with a C-terminal His6-tag was

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expressed in Pichia pastoris and purified as previously described by Zamocky et al.

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additional immobilized metal affinity step (all equipment from GE Healthcare, Little Chalfont,

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United Kingdom). The purification was carried out first by hydrophobic interaction

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chromatography (PHE-Sepharose FF resin) then by immobilized metal affinity chromatography

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(HisTrap FF resin) and as a last step by anion exchange chromatography (Q Source resin). The

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protein concentration was determined using the Bradford method (Biorad, Hercules, USA).

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CBP21 from Serratia marcescens (also known as SmLPMO10A) and all its variants were

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expressed and purified as previously reported by Vaaje-Kolstad et al.

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star (DE3) cells containing the pRSETB vector with the cbp21 gene were grown at 37°C

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overnight in two times 500 mL TB-medium containing 8.5 mM KH2PO4 and 36 mM K2HPO4,

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and 100 µg/mL ampicillin using a Harbinger LEX bioreactor (Harbinger Biotech, Toronto,

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Canada). The cells were collected by centrifugation and the protein was isolated from the

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periplasmic space using a cold osmotic shock method.40 The periplasmic extract was sterilized

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by filtration over a 0.45 µm filter. The purification of the LPMO was carried out as previously

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described by Vaaje-Kolstad et al.34. The periplasmic extract (approximately 200 mL) was

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adjusted to the binding buffer [1.0 M (NH4)2SO4, 25 mM Tris-HCl pH 8.0) and loaded onto 10

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mL chitin beads (NEB, Ipswich, USA). After the non-binding protein had passed the enzyme

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was eluted using 20 mM acetic acid. The eluted protein was immediately adjusted to 20 mM

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Tris-HCl pH 8.0. Subsequently, the enzyme solution was concentrated and the acetic acid was

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removed by ultrafiltration using an Amicon Ultra centrifugal filter with a 10 kDa cut off

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(Millipore Merck KGaA, Darmstadt, Germany).The protein concentration was determined using

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the absorbance at 280 nm and the theoretical molar absorption coefficient.

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Mutations were generated using the QuickChange II site-directed mutagenesis kit (Agilent

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Technologies, Santa Clara, USA). After DNA sequencing, the mutated expression vectors were

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transformed into chemically competent E. coli BL21 star (DE3) cells by heat shock. All protein


34

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with an

. Briefly, E. coli BL21

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variants were produced in soluble form and could be purified using standard methods described

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above. The yield for the purified CBP21 wild type was 20 mg per liter of culture.

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Table 1. Gene-specific primers used for mutagenesis PCR of CBP21 in the pRSETB expression vector. CBP21 variant Tyr54Ala Glu55Ala Ser58Ala Glu60Ala Thr111Ala Ala112Gly His114Ala Phe147Ala Ala152Arg Trp178Phe Ile180Arg Asp182Ala Thr183Ala Asn185Ala Phe187Tyr

Primer pair sense and antisense (5’ → 3’) GCGGCAGCGTGCAGGCTGAACCGCAGAGCGT ACGCTCTGCGGTTCAGCCTGCACGCTGCCGC AGCGTGCAGTACGCACCGCAGAGCG CGCTCTGCGGTGCGTACTGCACGCT GTACGAACCGCAGGCCGTCGAGGGCCTG CAGGCCCTCGACGGCCTGCGGTTCGTAC CGCAGAGCGTCGCAGGCCTGAAAGG CCTTTCAGGCCTGCGACGCTCTGCG TACCTGGAAGCTGGCCGCGCGTCACAG CTGTGACGCGCGGCCAGCTTCCAGGTA GGAAGCTGACCGGGCGTCACAGCAC GTGCTGTGACGCCCGGTCAGCTTCC GCTGACCGCCCGTGCTAGCACCACCAGCTGG CCAGCTGGTGGTGCTAGCACGGGCGGTCAGC GTTCTGCCAGGCCAACGACGGC GCCGTCGTTGGCCTGGCAGAAC CGACGGCGGCCGCATCCCTGCCGCACA TGTGCGGCAGGGATGCGGCCGCCGTCG GTGATCCTTGCCGTGTTCGACATAGCCGACACCG CGGTGTCGGCTATGTCGAACACGGCAAGGATCAC CCTTGCCGTGTGGGACAGAGCCGACAC GTGTCGGCTCTGTCCCACACGGCAAGG GGGACATAGCCGCCACCGCCAACGC GCGTTGGCGGTGGCGGCTATGTCCC GACATAGCCGACGCCGCCAACGCCTTC GAAGGCGTTGGCGGCGTCGGCTATGTC GCCGACACCGCCGCCGCCTTCTATCAGGC GCCTGATAGAAGGCGGCGGCGGTGTCGGC CACCGCTAACGCCTACTATCAGGCGATCG CGATCGCCTGATAGTAGGCGTTAGCGGTG

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Chitobiase from Serratia marcescens (SmGH20A) was expressed and purified as reported by

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Loose et al. 41. Briefly, BL21 star (DE3) cells harbouring the pET30 Xa/LIC vector with the chb

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gene were grown at 37°C to an OD600 = 0.5 in TB-medium supplemented with 100 µg/mL

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kanamycin using a Harbinger LEX bioreactor (Harbinger Biotech, Toronto, Canada). Protein

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production was induced by adding 0.3 mM IPTG (final concentration) and the culture was

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further incubated at 30°C for 5 h. The cells were harvested by centrifugation and kept at -20°C

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until use. For protein purification, the cell pellet (from approximately 300 mL culture) was

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thawed on ice and resuspended in 25 mL binding buffer (20 mM Tris-HCl pH 8.0, 5.0 mM

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imidazole) supplemented with 0.1 g/L lysozyme followed by 30 min incubation on ice. The cells 8 ACS Paragon Plus Environment

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Biochemistry

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were disrupted by sonication (Vibra cell sonicator, Sonics, Newtown, USA) at 27% amplitude in

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a repeated cycle of 5 sec on, 2 sec off, for 3 min in total. After removing cell debris by

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centrifugation, the protein extract was loaded onto 3.0 mL Ni-NTA Agarose resin (Protino,

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Machrey-Nagel, Düren, Germany). After the non-bound protein had passed the column,

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chitobiase was eluted in 20 mM Tris-HCl pH 8.0, containing 500 mM imidazole. The sample

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was concentrated and the imidazole was removed using an Amicon Ultra centrifugal filter with a

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10 kDa cut off (Millipore Merck KGaA, Darmstadt, Germany). The protein concentration was

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determined using the Bradford assay (Biorad, Hercules, USA).

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Chitinase A (ChiA, or SmChi18A)42 and Chitinase C (ChiC, or SmChi18C)43 from Serratia

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marcescens were expressed, harvested and the periplasmic extract was prepared like for CBP21.

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The periplasmic extract was sterile filtered (0.45 µm), adjusted to 50 mM Tris-HCl pH 8.0 and

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loaded onto 10 mL chitin beads (NEB, Ipswich, USA). Non-bound protein was discarded and the

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respective protein was eluted in 20 mM acetic acid. The sample was adjusted to 20 mM Tris-

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HCl, concentrated and the acetic acid was removed using an Amicon Ultra centrifugal filter with

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a 10 kDa cut off (Millipore Merck KGaA, Darmstadt, Germany). The protein concentration was

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determined using the absorbance at 280 nm and the theoretical extinction coefficient.

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Cu(II)-saturation and Desalting. All CBP21 variants were copper-saturated prior to activity

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assays. The procedure was carried out as described by Loose et al.

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solution was incubated with 3-fold molar excess of Cu(II)SO4 for 30 min at RT. The excess

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copper was removed by desalting the protein in 25 mM MES, pH 6.0 using a PD Midi-Trap G-

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25 desalting column (GE healthcare, Little Chalfont, United Kingdom).

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LPMO Activity Assay. LPMO reactions, containing 10 g/L β-chitin (France Chitine, Orange,

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France) and 1.0 µM LPMO, were buffered with 25 mM MES pH 6.0. As reductant, either 0.5

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µM MtCDH and 5.0 mM lactose or 1.0 mM gallic acid (stock solution: 100 mM gallic acid

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dissolved in 100 % EtOH) were used. The samples were incubated at 40°C with shaking at 800

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rpm in an Eppendorf Comfort Thermomixer with a temperature-controlled lid. Reactions were

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stopped by boiling for 20 min when the total amount of oxidized product was analyzed, or by

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removing the substrate by filtration using a 96-well filter plate operated by a vacuum manifold

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(Millipore) when the solubilized products only were analyzed. To be able to quantify the total 9 ACS Paragon Plus Environment

41

. In brief, the enzyme

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amount of oxidized products, the boiled samples were further degraded by incubation with a

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mixture of 3.0 µM ChiA, 3.0 µM ChiB and 4.0 µM Chb (all final concentrations) for 7 h at

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40°C, with shaking at 800 rpm. The samples obtained by filtration were degraded by incubation

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with 4.0 µM Chb for 2 h, under the same conditions. These treatments degrade the stopped

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reaction to GlcNAc and chitobionic acid.

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Production of Chitobionic Acid Standards. Chitobionic acid was produced as previously

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described by Loose et al.

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MES pH 8.0 was incubated with 0.1 g/L m-chitO44 overnight at 22°C. The complete conversion

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of chitobiose to chitobionic acid was verified by UHPLC.

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Product Analysis by UHPLC. Chitobionic acid was quantified using an Infinity 1290 UHPLC

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(Agilent Technologies, Santa Clara, USA) equipped with an Aquity BEH Amide 1.7 µm column

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(Waters, Milford, USA), and operated in HILIC (hydrophilic interaction) mode. To separate the

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oligosaccharides in the sample, a 2.1 × 150 mm column was operated at 0.4 mL/min using 15

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mM Tris HCl pH 8.0 (eluent A) and 100 % acetonitrile (eluent B) as eluents in the following

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gradient: 0 – 3.5 min, 80 % B : 20 % A; 3.5 – 12 min, gradient to 70 % B : 30 % A; 12 – 13 min,

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gradient to 55 % B : 45 % A; 13 – 14 min 55 % B : 45 % A; 14 – 15 min, gradient to 80 % B : 20

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% A, followed by reconditioning for 3 min. The elution of oligosaccharides was monitored at

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205 nm.

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The analysis of reaction supernatants, i.e. solubilized products was carried out as follows: 0 – 5

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min 74 % (B) : 26 % (A), 5 – 7 min gradient to 62 % (B) : 38 % (A), 7 – 8 min 62 % (B) : 38 %

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(A), 8 – 10 min gradient to 74 % (B) : 26 % (A), and reconditioning for 2 min. The elution of

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oligosaccharides was monitored at 205 nm.

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Substrate Binding. Binding experiments were carried out in the same conditions as the LPMO

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activity assays. Reaction mixtures containing 10 g/L β-chitin and 1.0 µM LPMO in 25 mM MES

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pH 6.0, with or without 1.0 mM gallic acid, were incubated at 40° with shaking at 800 rpm and

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samples were taken after 1, 2 and 6 h. The samples were filtered using a 96-well filter plate

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(Millipore) and the concentration of unbound protein was measured using the Bradford assay

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(BioRad, Hercules, USA). As control for determination of the total amount of protein, for each

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with minor modifications. In short, 2.0 mM chitobiose in 25 mM


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Biochemistry

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CBP21 variant, 1.0 µM LPMO was incubated in 25 mM MES pH 6.0 at 40°C and 800 rpm. The

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quantities of unbound protein in the reactions with chitin were calculated relative to the

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respective control reactions.

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Sample processing. The heat treated samples were processed according to the suspension

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trapping (STrap) protocol.45 Tryptic peptides were eluted from the STrap tips with 30 µl 80%

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acetonitrile/0.1% TFA, and dried in a speedvac. The dried peptides were subsequently

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resuspended in 10 µl loading solution (2 % acetonitrile, 0.05% TFA) and transferred to

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autosampler vials for LC-MS/MS analysis.

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NanoLC-Orbitrap MS/MS analysis of tryptic peptides. Peptides were analyzed using a

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nanoLC-MS/MS system consisting of a Dionex Ultimate 3000 UHPLC (Thermo Scientific,

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Bremen, Germany) connected to a Q-Exactive hybrid quadrupole-orbitrap mass spectrometer

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(Thermo Scientific, Bremen, Germany) equipped with a nano-electrospray ion source. Samples

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were loaded onto a trap column (Acclaim PepMap100, C18, 5 µm, 100 Å, 300 µm i.d. x 5 mm,

227

Thermo Scientific) and back flushed onto a 50-cm analytical column (Acclaim PepMap RSLC

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C18, 2 µm, 100 Å, 75 µm ID, Thermo Scientific). At the start, the columns were in 96% solution

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A [0.1% (v/v) formic acid], 4% solution B [80% (v/v) acetonitrile, 0.1% (v/v) formic acid].

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Peptides were eluted using a 40-min gradient developing from 4% to 15% (v/v) solution B in 2

231

minutes and 15% to 55% (v/v) B in 27 minutes before the wash phase at 90% B and re-

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equilibration, all at a flow rate of 300 nL/min. In order to isolate and fragment the 10 most

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intense peptide precursor ions at any given time throughout the chromatographic elution, the

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mass spectrometer was operated in data-dependent mode to switch automatically between

235

orbitrap-MS and higher-energy collisional dissociation (HCD) orbitrap-MS/MS acquisition. The

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selected precursor ions were then excluded for repeated fragmentation for 20 seconds. The

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resolution was set to R=70,000 and R=35,000 for MS and MS/MS, respectively. For optimal

238

acquisition of MS/MS spectra, automatic gain control (AGC) target values were set to 50,000

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charges and a maximum injection time of 128 milliseconds.

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Bioinformatic analysis. To get an overview of the identified peptides from CBP21 and for

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selecting correct peptides for quantification, the data were searched against a sequence database

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consisting of the 17 variants of CBP21 in a background of the complete proteome of E.coli BL21 11 ACS Paragon Plus Environment

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(downloaded from UniProt; 2871 sequences). The search engine used was Mascot46 and the

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tolerance levels for matching to the database was 7 ppm for MS and 30 mmu for MS/MS.

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Trypsin was used as digestion enzyme, and three missed cleavages were allowed. Error-tolerant

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searches were used for unbiased identification of peptides harboring potential modifications, and

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all hits with ion scores lower than 20 were omitted from the results.

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Based on these results, we selected three control peptides to be used for normalization purposes,

249

all of them distant from the active site, and from a part of the protein were observed

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modifications were at a minimum, namely STFFELDQQTPTR, TGPNSFTWK and

251

YFITKPNWDASQPLTR eluting at 20.7, 16.8 and 19.3, respectively. To quantify the

252

unmodified peptide HGYVESPASR, we used integrated peak areas in extracted ion

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chromatograms reported by the software Xcalibur (Thermo Scientific). HGYVESPASR could be

254

found eluting at 12.1 minutes, and the intensity was normalized by dividing by the sum of the

255

intensities of three control peptides. Where available, both the 2+ and 3+ ion species were used

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to generate extracted ion chromatograms. The normalized values were further normalized per

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variant by dividing them by the sum of all control peptides for all time points for the respective

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variant.

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We acknowledge that the method used for calculating oxidative damage of CBP21 variants is

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challenging and dependent on the detection quality of the selected control peptides.

261

Normalization procedures used in label-free quantification typically require hundreds of

262

identified proteins and peptides to function optimally. To get a reasonable estimate when

263

working with a single protein, we selected three control peptides, distant from the active site, and

264

from a part of the protein where observed modifications were at a minimum, and used the sum of

265

these to normalize the data.


Page 12 of 33

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Biochemistry

266

RESULTS

267

Design and initial characterization of CBP21 variants

268

Residues targeted for mutation were selected by consulting existing literature describing

269

sequence, structural and functional analysis of AA10 LPMOs.37,

270

conserved residues was performed using ConSurf.49 Residues on the substrate binding surface

271

and near the active site showing high degrees of conservation in at least one of the major AA10

272

subgroups50 were selected, resulting in the generation of 13 single amino acid variants (Fig. 2).

273

As default, residues were mutated to alanine or, if the wild-type had an alanine, glycine. There

274

are three exceptions, W178F, I180R and F187Y, all of which are mutations that reflect naturally

275

occurring variation in AA10 type LPMOs. Two additional control mutations were made

276

elsewhere on the protein surface (F147A and A152R), which were not expected to affect the

277

substrate-binding surface. All CBP21 variants showed wild type-like behavior during expression

278

and purification and the yields of purified protein where 5 – 25 mg per liter of culture. The

279

structural integrity of a subset of eight variants had previously been confirmed by circular

280

dichroism.34

47, 48

Additional analysis of

281

282 283

Figure 2. Structure of CBP21 and overview of the mutations. The side chains of residues selected for

284

mutation, as well as the side chain of the N-terminal histidine (H28) are shown as sticks and colored


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

285

purple. Panel (A) shows a side view and panel (B) shows the top view. The amino acids to which these

286

residues were mutated are indicated in the labels.

287

LPMO action generates soluble oxidized products that are often used for quantification of LPMO

288

activity. However, a fraction of the oxidized products will remain associated to the insoluble

289

material and this fraction may vary among LPMO variants. Figure 3 shows the amounts of total

290

oxidized products and soluble oxidized products for all CBP21 variants under standard assay

291

conditions. The ratio of total oxidized products versus solubilized oxidized products increased

292

with decreasing CBP21 activity. Thus, to obtain a correct comparison of WT and variant

293

activities, in the studies described below, the total amount of oxidized products was quantified at

294

each time point of the reaction.

295 296

Figure 3. Comparison of total (black) and solubilized (grey) oxidized products generated by CBP21

297

variants. The enzymes (1.0 µM) were incubated with ß-chitin (10 g/L) in the presence of MtCDH

298

(Myriococcum thermophilum CDH; (0.5 µM)) and lactose (5.0 mM) in MES buffer (pH 6.0, 25 mM), at

299

40 °C in a thermomixer (800 rpm) for 24 h. Note that the ratio between total and solubilized product

300

differs between the variants (see text for details). The error bars indicate standard deviations (n=3;

301

independent experiments).

302


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Biochemistry

303

Catalytic activity

304

Figure 4 shows product formation over time for all CBP21 variants in the presence of two

305

fundamentally different reductants: a small organic acid (gallic acid) or MtCDH (CDH from

306

Myriococcum thermophilum). The CBP21 variants differed in terms of the apparent initial rate of

307

product formation (i.e the product level at the first time point) and final product levels. The

308

outcome of the reactions depended to some extent on the reductant used, but, overall,

309

performance differences between the CBP21 variants were similar for both reductants. Two

310

notable exceptions are the A112G and F187Y variants, which gave clearly higher final product

311

levels with gallic acid compared to MtCDH. The variants E55A, E60A, H114A, I180R and

312

D182A showed particularly low product levels, whereas T111A and the two control variants

313

carrying mutations not affecting the substrate-binding surface (F147A, A152G) showed WT-like

314

behavior. All other variants had reduced final product yields.

315 316

Figure 4. Activity of CBP21 variants. (A) Structure of CBP21 showing the side chains of residues

317

selected for mutation, as well as the side chain of the N-terminal histidine (H28). Water molecules are

318

shown as red spheres. The copper ion is shown as a yellow sphere. Hydrogen-bonds are shown as black

319

dashed lines. (B&C) The graphs show total amounts of oxidized products observed after varying

320

incubation times for reactions with MtCDH/lactose (0.5 µM/5.0 mM) (B) or gallic acid (1.0 mM) (C) as

321

electron donor. Reactions were carried out in 25 mM MES buffer (pH 6.0), at 40 °C in a thermomixer

322

(800 rpm). Note that the maximum reaction times in panel A and B differ (24 h and 48 h, respectively).

323

Bars are colored red in cases where the product level is not signficantly higher than the product level


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

324

detected at the preceding time point, meaning that LPMO activity has ceased. The asterisks indicate that

325

no sample was taken at that time point. The error bars indicate standard deviations (n=3; independent

326

experiments). In the reaction conditions employed, 1 mM chitobionic acid was detected at the end of the

327

reaction, which corresponds to 4% conversion yield from chitin.

328 329

It is noteworthy that the apparent initial rates of many CBP21 variants, including Y54A, S58A,

330

T111A, F147A, A152R, W178F, T183A, and N185A, were similar to the WT (Fig. 4B and C).

331

However, over time, the catalytic activity of several of these variants ceased and clear

332

differences in product yield became apparent. Thus, enzyme inactivation took place at a rate that

333

varied between the CBP21 variants. Even for the least active variants (E60A, I180R and D182A,

334

but not E55A and H114A) small amounts of product were detected early in the reaction,

335

suggesting that also for these variants inactivation contributes to reduced performance. All

336

variants, except WT, the two control variants, and T111A, reached reaction end points, and the

337

incubation time needed to do so depended on the mutation. Clear differences between the

338

reductants were apparent: with MtCDH/lactose, initial rates were higher, but the activities of

339

many variants ceased earlier (after 2 - 4 h) compared to gallic acid (after 24 h). A112G and, even

340

more so, F187Y stand out as being substantially more active with gallic acid. With

341

MtCDH/lactose the activity of these variants had ceased at the first measuring point (2 h) and

342

final product yields were very low compared to wild-type (7% and 4% of WT for A112G and

343

F187Y, respectively). With gallic acid, product formation continued over time and the final

344

yields amounted 25% and 55% of WT levels for A112G and F187Y, respectively.

345 346

Substrate binding

347

To determine the influence of the various mutations on the binding properties of the enzyme

348

during catalysis, binding of CBP21 variants to β-chitin over time was investigated using the

349

same conditions as in the activity assay (Fig. 5). The general observation is that a stable binding

350

equilibrium was established after one hour for all CBP21 variants. Binding properties similar to

351

the WT were observed for mutation of some of the polar residues surrounding the active site


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Biochemistry

352

(T111A, T183A and N185A), the Trp buried beneath the active site (W178F), as well as for the

353

control variants (F147A and A152R). All other mutations reduced the binding to varying extents.

354

Mutation of the only solvent-exposed aromatic amino acid (Y54A) had a drastic effect, and so

355

had various mutations affecting negatively charged residues on the substrate binding surface

356

(E55A, E60A and D182A). Mutations of residues in the copper site or very close to the copper

357

site (A112G, H114A, S58A, F187Y and I180R) also reduced binding, with A112G having the

358

smallest effect.

359

Figure 5, also shows that the loss of affinity to β-chitin correlates with reduced operational

360

stability of the enzyme (Figure 4B and 4C). This correlation, which also applies to variants

361

showing the same apparent initial rate as the wild-type enzyme, suggests that precise substrate-

362

binding is a crucial factor in determining LPMO functionality. Figure 5A also shows that the

363

reduced operational stability of the LPMO affects substrate binding, since the bound fraction of

364

the protein is decreasing over time. Proteins showing a lower affinity in the beginning (below

365

60%) show a faster decrease in binding affinity compared to the strongly binding WT and

366

variants. Thus, the process that leads to enzyme inactivation also reduces substrate binding

367 368

Figure 5. Binding and activity of CBP21 variants. (A) Binding experiments were performed in

369

identical conditions as used for activity assays with gallic acid (1.0 µM enzyme 1.0 mM gallic acid, 10

370

mg/ml β-chitin in 25 mM MES pH 6.0). Reactions were incubated at 800 rpm and 40 °C, and samples

371

were collected and analyzed at 1, 2 and 6 h. The coefficient of variation was < 22 % for all time points

372

analyzed (n=3; independent experiments). Error bars are not displayed for clarity. (B) Surface

373

representations of CBP21 colored by the effect of mutations on binding after 1 h (left; data in panel A of

374

this figure) and product formation after 48 h (right; see Fig. 4C).


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

375 376

Oxidative modifications of CBP21 WT and variants

377

Bissaro et al. have shown that, in the absence of substrate and the presence of reductant,

378

activated/ primed LPMOs (i.e. containing copper in the Cu(I) state) undergo self-inactivation,

379

which is manifested by oxidative damage of residues near the catalytic center, in particular the

380

copper-coordinating histidines

381

CBP21 variants relate to oxidative inactivation, we used label-free quantitative mass

382

spectrometry for monitoring the integrity of the N-terminal tryptic peptide, containing the copper

383

coordinating His28, over time. Although we were able to observe oxidized variants of the N-

384

terminal tryptic peptide in inactivated variants (e.g. masses corresponding to oxidation [M+16]

385

or decomposition following the oxidation [M-22 or M-23]), it was found that quantification of

386

the remaining native N-terminal peptide gave better quantification of histidine oxidation (see the

387

Methods section for experimental details and the Discussion section for more discussion of this

388

topic). While this experimentally challenging approach only gives a rough estimation of

389

oxidative damage, the results show a clear trend: variants displaying reduced operational stability

390

compared to the WT also showed more evident reduction of the native N-terminal peptide over

391

time (Fig. 6). The H114A variant is an interesting exception. This inactive variant lacks one of

392

the copper ligands and will thus not be able to carry out the type of oxidative chemistry normally

393

catalyzed by LPMOs. Thus, it is not surprising that this variant does not catalyze oxidative self-

394

destruction neither. CBP21 variants with WT-like activity and WT-like substrate binding showed

395

relatively stable levels of the intact N-terminal peptide. It is worth noting that the correlation

396

between damage of the N-terminal peptide and operational stability is not absolute. In particular,

397

two variants with WT-like substrate binding ability, namely T183A and N185A, displayed

398

ceased activity between 24 and 48 h incubation time (Fig. 4), without disappearance of the native

399

N-terminal peptide (Fig. 6).

23

. To assess whether the apparent stability differences between


Page 18 of 33

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Biochemistry

400 401

Figure 6. Oxidative damage of CBP21 variants. The enzymes (1.0 µM) were incubated in 25 mM MES

402

buffer, pH 6.0, containing gallic acid (1.0 mM) and β-chitin (10 mg/ml), at 40 °C in a thermomixer (800

403

rpm). The integrity of the enzyme was analyzed at various time points. The bars show the relative amount

404

of the native N-terminal peptide of CBP21 normalized by the sum of three unmodified control peptides.

405

Orange colored small circles show the levels of oxidized products generated (same data as shown in Fig.

406

4); the data points are connected by an orange line for illustration purposes: flattening of the curve over

407

time is indicative of enzyme inactivation. * indicates that no reliable quantification of the N-terminal

408

peptide was achieved. The amount of protein bound to β-chitin after 1 h incubation under the same

409

conditions is indicated in percent beneath the name of the enzyme variant (see Fig. 5 for more details).

410

Note: The oxidative damage was not quantified at 2 h.


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

411

DISCUSSION

412

Despite the importance of LPMOs in biomass conversion processes17, 51-53 and great scientific

413

interest in the unique catalytic mechanism of these enzymes,18, 19, 21-23 site-directed mutagenesis

414

data for probing the roles of individual amino acids in catalysis are scarce.15, 16, 30 The complexity

415

of characterizing LPMO catalysis and kinetics has likely contributed to limiting this type of

416

studies. Indeed, phenomena such as non-linear kinetics, conspicuously similar turnover numbers

417

observed for unrelated enzymes acting on different substrates, and enzyme inactivation during

418

catalysis have contributed to confusion in the field. Of note, the time scales of LPMO assays

419

reported in the literature and used in the present study are usually in the multi-hour range. This

420

time range makes sense from a biological point of view, since biomass degradation is a slow

421

process, usually taking days or even months. However, such long time scales may cause issues

422

related to enzyme stability, as shown here.

423

In the current study, both an enzymatic (MtCDH) and a small molecule (gallic acid) reductant

424

were used to drive the CBP21 reaction. These reductants deliver the two electrons that are

425

needed per catalytic cycle, regardless of whether the LPMO mechanism is H2O2-based23 or

426

follows one of several proposed mechanisms with molecular oxygen as co-substrate.18, 19 Gallic

427

acid was preferred over commonly used ascorbic acid, because it is less prone to auto-oxidation27

428

and thus more likely to yield stable kinetics, as was indeed observed. Driving LPMO reactions

429

by ascorbic acid often leads to non-linear kinetics, as shown by e.g. Loose et al. 33

430

Many of the mutations made in this study had no effect on the apparent initial enzyme rate, even

431

though substrate-binding and enzyme operational stability were affected. Such a lack of effect on

432

enzyme rate could perhaps be expected for mutations far away from the catalytic center, such as

433

Y54A. However, even two mutations in the primary copper coordination sphere, F187Y in the

434

proximal axial position and A112G in the distal axial position (Fig. S1), had limited effects on

435

the apparent initial enzyme rate. The dataset contains three mutations in the secondary copper

436

coordination sphere30 (Fig. S1): W178F, affecting a buried aromatic residue that interacts with

437

Phe187 in the proximal axial copper coordination sphere; E60A, affecting a residue that is

438

structurally equivalent to a Gln residue that is conserved in AA9-type LPMOs and that is known

439

to be important for activity,16, 30 possibly due to its role in binding an oxygen species54; I180R, 20 ACS Paragon Plus Environment

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Biochemistry

440

which is a natural mutation in AA10-type LPMOs that may put an positively charged arginine

441

head group in the position of a conserved His residue that is known to be important for activity in

442

AA9-type LPMOs and30, possibly due to its role in binding an oxygen species54. Two of these

443

mutations, I180R and E60A, did reduce the apparent initial rate but it should be noted that these

444

mutations also had severe effects on substrate binding and enzyme stability, meaning that,

445

perhaps the apparent initial enzyme rates (i.e. product formation at the first measuring point) are

446

underestimated (see below for further discussion).

447

Considering the recent finding that LPMOs can use H2O2 as co-substrate,23, 24 the remarkable

448

similarity of apparent initial rates for many of the variants could be explained by assuming that

449

LPMO catalysis is rate-limited by the rate at which H2O2 is generated in the reaction mixture.

450

H2O2 will be generated by reduced LPMO molecules that are not bound to substrate25, 26 and by

451

reactions involving molecular oxygen and the reductant, and both these processes are not likely

452

to be affected by most mutations. Indeed, the observed catalytic rate (in the order of 1.3 min-1 for

453

the CDH-driven reaction) is orders of magnitude lower that the kcat of

454

determined for H2O2-driven CBP21 in a recent kinetic study.24 This leads to the very important

455

conclusion that in all LPMO engineering studies so far, including the present one, mutational

456

effects on catalytic power are likely masked by the overall much slower process of H2O2

457

generation.

458

The most important observation in the present study is the negative effect of a remarkably large

459

fraction of the mutations on CBP21 operational stability. LPMOs are known to be prone to

460

oxidative damage.38 Recently, Bissaro et al.

461

with H2O2 without being bound to substrate will be inactivated due to oxidative damage of

462

residues in and very close to the copper site. The present data show that many residues in the

463

substrate-binding surface contribute to optimizing substrate-binding and, thus, minimizing

464

enzyme inactivation. Overall, the data show a clear and remarkable correlation between reduced

465

binding to chitin (Fig. 5), a reduced operational stability during turnover (Fig. 4), and physical

466

damage to the protein (Fig. 6).

467

The mechanism resulting in the oxidative damage observed for several CBP21 variants is not

468

known, but it is likely related to uncontrolled activation of the co-substrate or to off-pathway

23

5.6 s-1 that was

showed that reduced LPMOs that are supplied


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 22 of 33

469

reactions involving a reactive intermediate. The copper-containing active site of an LPMO is

470

solvent exposed and is thus accessible to all solvent molecules unless it is shielded/confined by

471

substrate binding.55 A reduced LPMO, carrying an exposed Cu(I), that is not bound to its

472

substrate is therefore prone to react with H2O2 in solution, which will result in the formation of

473

highly reactive oxygen species (ROS) such as hydroxyl radicals. In the absence of the proper

474

substrate, such ROS will react with any organic component within a short radius. In the case of

475

LPMOs, the active site histidines are likely to be the closest to the formed ROS and are thus

476

prone to be oxidized. Indeed, an in-depth study of oxidative damage in a cellulose active LPMO

477

from S. coelicolor (ScLPMO10C) showed that such damage was limited to the direct

478

environment of the copper site, in particular the two copper-binding histidines.23 Once the

479

imidazole side chain moiety of a histidine is oxidized, the amino acid becomes unstable and will

480

decompose to an array of different products, the final stable product being asparagine or

481

aspartate,56 as indeed observed for ScLPMO10C.23 Due to the complexity of the oxidation

482

pathways, mass spectrometry-based quantification of oxidized peptides was highly challenging,

483

explaining why, here, we monitored instead the disappearance of the native peptide as a

484

quantitative proxy of oxidative damage.

485

Although most CBP21 variants analyzed in this study showed a correlation between substrate

486

binding ability and operational stability, variants T183A and N185A stand out since they bind

487

well to chitin (Fig. 5), are not especially prone to oxidation (Fig. 6), but nevertheless show

488

reduced operational stability (Fig. 4). Although there is no obvious explanation for this, it is

489

conceivable that some oxidative damage did happen, but primarily in other locations than the N-

490

terminal peptide. Another exception is the W178A variant, which did show the “expected”

491

correlation of decreased operational stability and increased oxidative damage, but showed WT-

492

like chitin binding. This mutation affects a cluster of aromatic residues located internally in the

493

protein, just below the active site (Fig. S1). Thus, Trp178 is not likely to affect substrate binding,

494

but may have a protective effect on the catalytic center.

495

While H2O2 will be continuously consumed by well-binding catalytically active LPMOs through

496

productive catalysis, weakly binding LPMOs will experience a gradually increasing H2O2

497

concentration, mediating oxidative self-inactivation. Interestingly, Eibinger et al.

498

showed that the residence time of an LPMO on cellulose is on the minute time scale, implying 22 ACS Paragon Plus Environment

57

recently

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Biochemistry

499

that exchange of LPMOs between solvent and substrate is “designed” to be slow (compared to

500

glycoside hydrolases that associate and dissociate on the second time scale). Such findings do

501

indeed make sense in the light of the results presented in the current study. On the one hand it

502

may be that the level of substrate oxidation can be regulated by substrate binding, which may be

503

beneficial in vivo. On the other hand, it cannot be excluded that LPMOs can perform multiple

504

catalytic events when bound to the substrate (i.e. when still being protected). Further, it is

505

interesting to note that LPMOs have evolved substrate binding in two ways, either by encoding

506

substrate binding properties in the catalytic LPMO domain itself, or by appending the LPMO

507

domain to a CBM. Since LPMO domains evolved with a CBM, do not bind strongly to the

508

substrate themselves,31,

509

productive manner.

510

As to the assessment of binding in this study, it is important to note that binding of course will be

511

affected by oxidative damage to the enzyme. This may explain why for several variants the

512

binding curves show a gradual loss of affinity over time, which, notably, correlates with

513

oxidation of the N-terminal peptide. Importantly, in our correlation studies we compare binding

514

conditions (1 hour, gallic acid) under which most CBP21 variants remain active (Fig. 4C).

515

Notably, also variants with strongly decreased apparent initial rates, such as E60A and D182A,

516

show signs of vary rapid enzyme inactivation, which could in part explain their apparent very

517

low binding after 1 hour. The I180R variant is special since it showed only a moderate reduction

518

in binding, but had a short operational stability accompanied by very rapid disappearance of the

519

intact N-terminal peptide. The I180R mutation occurs in nature; several cellulose-active AA10

520

LPMOs have Arg at this position. It is possible that for the I180R variant not all substrate

521

binding events provide protection against oxidative damage. Whereas productive binding allows

522

correct positioning of the reactive intermediate to carry out substrate oxidation, non-productive

523

binding may result in a similar but “idle” reactive intermediate prone to enter off-pathway

524

reactions. In this respect, it is worth noting that some cellulose-active AA10s bind strongly to

525

chitin, without cleaving it.58

526

Among the residues mutated in CBP21 variants with clearly reduced initial rates, His114 and

527

Glu55 seem absolutely essential for LPMO activity. While this is easy to understand for copper

528

coordinating His114, the role of Glu55, located far away from the copper site is intriguing. While

32

the CBM must somehow help positioning the LPMO domain in a


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

529

the solvent-exposed carboxyl group of Glu55 is as much as 7.6 Å away from the closest

530

imidazole of the copper site (Nε2 of His28), mutation of this residue had a devastating effect on

531

CBP21 functionality. Figures 5 and 6 show minimal binding and a very rapid decrease of the

532

intact N-terminal peptide, respectively. The side chain of Glu55 is highly solvent exposed and

533

seems perfectly aligned with e.g. Tyr54 to interact with a substrate chain. Its involvement in

534

substrate binding is clear from NMR studies.37 This position is not conserved amongst AA10s in

535

general, in cellulose-active AA10 this position is occupied by an Asn.59 However, the Tyr is

536

conserved in most related, chitin-active LPMOs, except in CjLPMO10A from Cellvibrio

537

japonicus where it is a Thr. Although we cannot exclude that Glu55 somehow affects the copper

538

center, it seems that the role of Glu55 is in substrate binding and, as such, the E55A mutation

539

provides a prime example of the major importance of the exquisitely evolved substrate

540

interactions that underlie LPMO functionality. Finally, it should be noted the side chain of Glu55

541

has water mediated interaction with Ser58, which is relatively close to His28 (3.8 Å; Fig. 4A)

542

and whose mutation to Ala also reduces both binding and activity negatively (Fig. 4&5). Thus, it

543

may be that mutation of Glu55 has indirect consequences for catalysis by disrupting a putative

544

interaction between Ser58 and His28 in addition to reducing substrate binding.

545

In conclusion, the present data disclose several important features of LPMOs and LPMO

546

research. Proper assessment of LPMO functionality is challenging. Clearly, single time-point

547

characterization of LPMO performance is not acceptable, since resistance against oxidative self-

548

inactivation is an intrinsic part of LPMO functionality and will be affected by variations in

549

sequence and assaying conditions. The few mutational effects that have been described in the

550

literature so far, likely represent a mixture of stability and activity effects, while the true catalytic

551

power of the LPMO in question likely was masked by H2O2 production being rate limiting.

552

Most importantly, the present data demonstrate that precise substrate-binding, involving many

553

residues in the LPMO substrate-binding surface, is crucial for LPMO functionality. Even minor

554

changes in the binding surface may have detrimental consequences. This need for multi-point

555

precision binding in order to ensure productive use of the catalytic power of LPMOs may explain

556

why some organisms have so many LPMOs. Each LPMO may be fine-tuned for one particular

557

surface, being it the various faces of a cellulose crystal or the numerous co-polymeric assemblies

558

that occur in plant cell walls. In this respect, it would be of interest to further study the 24 ACS Paragon Plus Environment

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Biochemistry

559

operational stability of LPMOs that can act on several substrates, such as various β-glucans.

560

Finally, it is intriguing to consider the role of CBMs in light of the present observations. As

561

supported by recent work by Crouch et al.

562

ascribed to CBMs may find a new meaning for LPMOs, since improved substrate binding

563

prevents auto-inactivation. While CBMs by themselves will not give “precision binding”, they

564

will obviously increase the binding efficiency thus protecting the LPMO from harmful off-

565

pathway processes.

60

and Forsberg et al. 61, the “proximity effect” often

566


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ACKNOWLEDGEMENTS

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This work was supported by the Research Council of Norway Grants 214138 (JSML and GV-K),

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249865 (JSML, GV-K and MØA), 214613 (VGHE), the European Commission (project INDOX

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FP7-KBBE-2013-7-613549) (RL), Marie-Curie FP7 COFUND People Programme (AgreenSkills

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fellowship under grant agreement n° 267196) (BB). We thank Morten Skaugen for help with

572

acquiring the proteomics data and Anne Cathrine Bunæs for assisting with protein expression

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and purification.

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ASSOCIATED CONTENT

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Supporting information. A figure displaying the active sites of various bacterial and fungal

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LPMOs is supplied as Supporting Information.


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