1 How a Lytic Polysaccharide Monooxygenase Binds Crystalline

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How a Lytic Polysaccharide Monooxygenase Binds Crystalline Chitin Bastien Bissaro, Ingvild Isaksen, Gustav Vaaje-Kolstad, Vincent G.H. Eijsink, and Åsmund K. Røhr Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b00138 • Publication Date (Web): 02 Mar 2018 Downloaded from http://pubs.acs.org on March 3, 2018

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Biochemistry

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Bastien Bissaro‡, Ingvild Isaksen‡, Gustav Vaaje-Kolstad‡, Vincent G.H. Eijsink‡ and Åsmund K. Røhr‡*

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KEYWORDS; LPMO, ENZYMES, EPR

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ABSTRACT GRAPHIC

How a Lytic Polysaccharide Monooxygenase Binds Crystalline Chitin

CHITIN,

CELLULOSE,

COMPUTATIONAL

CHEMISTRY,

COPPER

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ABSTRACT: Lytic polysaccharide monooxygenases (LPMOs) are major players in biomass

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conversion, both in Nature and in the biorefining industry. How the mono-copper LPMO

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active site is positioned relative to the crystalline substrate surface to catalyze powerful, but

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potentially self-destructive, oxidative chemistry is one of the major questions in the field. We

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have adopted a multi-disciplinary approach, combining biochemical, spectroscopic and

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molecular modeling methods to study chitin binding by the well-studied LPMO from Serratia

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marcescens SmAA10A (or CBP21). The orientation of the enzyme on a single chain substrate

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was determined by analyzing enzyme cutting patterns. Building on this analysis, molecular

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dynamics (MD) simulations were carried out to study interactions between the LPMO and

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three different surface topologies of crystalline chitin. The resulting atomistic models showed

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that most of enzyme-substrate interactions involve the polysaccharide chain that is to be

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cleaved. The models also revealed a constrained active site geometry as well as a tunnel

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connecting the bulk solvent to the copper site, through which only small molecules such as

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H2O, O2 or H2O2 can diffuse. Furthermore, MD simulations, QM/MM calculations and EPR

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spectroscopy demonstrate that rearrangement of Cu-coordinating water molecules is

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necessary when binding substrate and also provide a rationale for the experimentally observed

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C1-oxidative regiospecificity of SmAA10A. This study provides a first, experimentally

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supported, atomistic view of the interactions between an LPMO and crystalline chitin. The

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confinement of the catalytic center is likely of crucial importance for controlling the oxidative

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chemistry carried out by LPMOs and will help guiding future mechanistic studies.

1 ACS Paragon Plus Environment

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INTRODUCTION

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Enzymes known as lytic polysaccharide monooxygenases (LPMOs) act on structural,

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crystalline polysaccharides such as cellulose1–3 or chitin,4 but also various hemicelluloses5,6 or

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starch.7 This makes LPMOs key players of biomass conversion in Nature, notably during

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fungal action,8 and has rendered LPMOs crucial for the efficiency of the most recent

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commercial enzymatic cocktails employed in biorefineries.9 LPMOs may also be involved in

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microbial pathogenicity.10–12 LPMOs are mono-copper enzymes carrying out hydroxylation of

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the C1 and/or C4 carbon of the scissile glycosidic bond, leading to bond cleavage via an

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elimination reaction.13 Importantly, and in contrast to classical hydrolytic enzymes such as

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cellulases, LPMOs can cleave polysaccharide chains packed in crystalline forms4,14 thus

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disrupting the crystalline surface.15–17 Substrate hydroxylation is thought to be catalyzed by an

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activated copper-oxygen species,18–22 and requires a precise assembling of the enzyme-

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polysaccharide complex, allowing regioselective oxidative chemistry to occur and preventing

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off-pathway oxidative processes that may damage the enzyme.22

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Despite recent developments23–25 little is known about how LPMOs interact with their

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crystalline substrate. An NMR-based approach led to the identification of residues involved in

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the interaction between the chitin-active LPMO from Serratia marscecens (SmAA10A, also

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known as CBP21) and crystalline chitin,23 confirming and extending knowledge acquired

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from a previous mutagenesis study.26 Insights into the interactions between a fungal LPMO

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and (soluble) cello-oligosaccharides have been obtained from both NMR24 and

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crystallographic studies.25 The seminal crystallographic work by Frandsen et al.25 revealed

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atomistic details of the interaction between the catalytic center of the LPMO and a single

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sugar chain and also provided insights into the effect of substrate-binding on the

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configuration of the copper site. However, interactions between an LPMO and a soluble

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substrate likely differ from the interactions with a substrate embedded in a crystalline lattice.

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Currently, only computational approaches allow simultaneous visualization of an enzyme and

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a crystalline substrate at the atomic scale.

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A single study has so far addressed this matter by modeling the interaction of a fungal

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(AA9) LPMO with cellulose,27 indicating interacting residues and showing high flexibility of

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two surface loops (not present in SmAA10A). Notably, in this study, the orientation of the

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LPMO on the cellulose chain was not experimentally probed but set by analogy to previous

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molecular dynamics studies performed on carbohydrate binding modules.28 The diversity of

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loops and surface topologies displayed by LPMOs may reflect the diversity of substrate

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structures (topologies and decorations) but, so far, correlations between surface architecture

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and substrate specificity remain unknown.29,30 One interesting question is whether substrate

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interactions primarily (or only) involve the polysaccharide chain that is to be cleaved or also

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involve contacts with adjacent chains. From that point of view, chitin-LPMO systems are


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interesting to study since (i) chitin can be found as α- or β-allomorphs, displaying antiparallel

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and parallel chain arrangements, respectively, and (ii) some chitin-active LPMOs, such as

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SmAA10A, are active on both forms but with preferences.31–34

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In the present study, we aimed at answering fundamental questions pertaining to the

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geometry of the LPMO-substrate complex, the structural determinants of substrate-binding,

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and binding-induced effects on the enzyme active site and its access. Using a multi-

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disciplinary approach combining computational, biophysical and biochemical methods, we

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have produced the first, experimentally supported, fully atomistic model of an LPMO

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interacting with a crystalline polysaccharide, here chitin. Enzyme activity assays were used to

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direct model building and subsequent molecular dynamics simulations, and electron

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paramagnetic resonance (EPR) spectroscopy was used for experimental confirmation of

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computational results. All work was performed using a well-studied LPMO, SmAA10A, that

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can interact with both crystalline α- or β-chitin, allowing thorough comparisons of

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computational and experimental data. The present findings shed light on how an LPMO

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orients itself and binds to a crystalline substrate in a mode that allows regioselective catalysis

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of oxidative polysaccharide cleavage.

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MATERIALS AND METHODS

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Materials. Chemicals were purchased from Sigma-Aldrich. β-chitin extracted from squid

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pen was purchased from France Chitin (Orange, France). Ascorbic acid (100 mM) stock

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solutions were prepared in metal-free water (Trace SELECT®, Sigma-Aldrich), aliquoted,

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stored at -20 °C, and thawed in the dark for 10 min just before use.

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Production and purification of recombinant LPMOs. Recombinant LPMO from Serratia

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marcescens (SmAA10A) and mutants thereof were expressed and purified according to

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previously described protocols.26 All LPMOs used in this study, except the wild type used in

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the EPR experiments (see EPR section), were prepared in sodium phosphate buffer (50 mM,

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pH 6.0), copper-saturated with Cu(II)SO4 and desalted (PD MidiTrap G-25, GE Healthcare)

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before use.11

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SmAA10A activity test. Reactions were carried out in 2 mL Eppendorf tubes and the reaction

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volume was 200 µL (for final time point analysis) or 500 µL (for time-course monitoring).

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Typical reactions contained the LPMO (10 µM) and substrate (10 g.L-1 for β-chitin or 1 mM

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for oligosaccharides) in Tris-HCl buffer (pH 8.0, 50 mM) and were pre-incubated during 20

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min at 40 °C in a Thermomixer (1000 rpm). The reactions were initiated by adding ascorbic


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acid (to a final concentration of 1 mM). In control reactions, SmAA10A was replaced by

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Cu(II)SO4 (10 µM). For time course monitoring, 55 µL samples were taken from the reaction

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mixtures at regular intervals and soluble fractions were immediately separated from the

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insoluble substrate by filtration using a 96-well filter plate (Millipore) operated with a

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vacuum manifold. Immediately after, the filtrate was incubated at 98 °C during 15 min to

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ensure enzyme inactivation. The heat inactivated samples were then frozen (-20 °C) prior to

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further analysis.

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For qualitative product analysis, samples were analyzed by MALDI-ToF MS, as previously

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described.4 For quantitative analysis, oxidized chito-oligosaccharides were separated by high

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performance anion exchange chromatography (HPAEC) and monitored by pulsed

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amperometric detection (PAD) using a Dionex Bio-LC equipped with a CarboPac PA1

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column as previously described for cello-oligosaccharides.35 All chromatograms were

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recorded using Chromeleon 7.0 software. In all figures, DPnox refers to (GlcNAc)n-

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1GlcNAc1A,

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forms) of a chito-oligosaccharide composed of n glycosyl units. DPnox standards were

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obtained by oxidation of chito-oligosaccharides with a DP ranging from 1 to 6 by a chito-

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oligosaccharide oxidase as previously described.11

i.e. the C1-oxidized form (in equilibrium between lactone and aldonic acid

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Electron Paramagnetic Resonance (EPR) Spectroscopy. All samples contained 200 µM

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SmAA10A in 50 mM MES pH 6.0. Before use, the enzyme was saturated with Cu(II)SO4 and

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desalted (PD MidiTrap G-25, GE Healthcare), as described above. The SmAA10A-(NAG)6

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sample was prepared by dissolving 4 mg (NAG)6 (hexa-N-acetyl-chitohexaose, Megazyme)

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in 200 µL SmAA10A solution. Hydrated chitin particles were obtained by repetitive washing

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with MES buffer (50 mM, pH 6.0), after which excess buffer was removed using a 0.22 µm

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centrifugal filter (Millipore), spinning at 10,000 g for 2 minutes. Samples with α-chitin (Sea

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Garden, Norway, Avaldsnes) or β-chitin (France, Orange), both < 80 mesh-sized particles,

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were prepared by carefully packing EPR tubes with 300 µL hydrated particles before adding

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200 µL of a copper saturated 200 µM SmAA10A solution or 200 µL 50 mM MES pH 6.0

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(control reaction). After 10 minutes, excess solution was removed, resulting in a sample

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height of ~25 mm, and the samples were frozen in liquid nitrogen. EPR spectra were recorded

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using a BRUKER EleXsys 560 SuperX instrument equipped with an ER 4122 SHQE SuperX

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high sensitivity cavity and a cold finger. Spectra were recorded using 1 mW microwave

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power and 10 G modulation amplitude at 77 K. In presence of substrate the EPR spectrum

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shows a mixed population corresponding to substrate-bound and unbound fractions of the

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LPMO. The spectrum corresponding to the unbound LPMO fraction as well as the substrate-


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only background were subtracted from all substrate-containing EPR spectra. Spin

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Hamiltonian parameters were fitted using the EasySpin package.36

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Synchrotron radiation X-ray diffraction (SRXRD). Diffraction data was collected at the

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Swiss-Norwegian beamline (BM01), ESRF, France using a PILATUS2M detector and a

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wavelength of 0.7361 Å.37 Hydrated samples of the α- and β-chitin identical to what was used

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in the EPR experiments were packed in ø 0.8 mm capillaries and diffraction data was

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collected at room temperature, 60 s exposure, 1.5° s-1. Diffraction patterns were processed

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using the SNBL Tool BOX software37 and Matlab.

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Development of molecular models. The SmAA10A atomic coordinates were extracted from

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the PDB entry 2BEM:C.26 The H++ server38 predicted only His74 to have a notably altered

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pKa value, being positively charged at pH 7.0. Thus, the PDB file was updated accordingly

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(His74 -> HIP74; i.e. protonated on delta and epsilon nitrogens). The Na+ ion modeled in the

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copper-binding site in the original crystal structure was replaced by copper. All

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crystallographic water molecules were retained in the model.

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No force field parameters exist for the copper sites in LPMOs. Therefore, a minimal

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model consisting of a truncated histidine brace (N-terminal His28 atom C and His114 atom

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Cβ capped with H) with Cu(I) or Cu(II) was used to derive partial charges for both

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oxidization states. During geometry optimization of these complexes, an angle constraint (Nδ-

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Cu-Nε = 178°) was applied to retain the T-shaped Cu-3N geometry, which is a known, highly

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conserved structural feature of LPMOs, regardless of their redox state.30 The electrostatic

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potentials of the Cu(I) and Cu(II) complexes were computed using the program Gaussian 0939

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with the hybrid functional B3LYP and the 6-311G(d,p) basis set for Cu and 6-31G(d,p) for

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the remaining atoms. When computing the ESPs, the atomic radius of the Cu atom was set to

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1.8 Å. Finally, the partial charges were fitted using the RESP method implemented in

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AmberTools17.40 Bond and angle force constants describing the Cu-histidine brace

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interaction were computed from the Cu(II) containing minimal model described above to

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which two water ligands were added, resulting in a near trigonal bipyramidal starting model.

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Adding the water molecules allowed unrestrained geometry optimization (B3LYP/6-

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311G(2d,2p)) of the complex, which is a prerequisite for a correct frequency analysis and

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which resulted in a conserved T-shaped Cu-3N model with an overall distorted square

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pyramidal molecular geometry. Subsequently to frequency analysis, cartesian force constants

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were extracted from the Hessian and projected on unit vectors representing bonds and angles,

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to compute bond and angle force constants encompassing the Cu atom.41 The resulting force

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field parameters can be found in Table S1.


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The most abundant forms of crystalline chitin found in Nature are α-chitin and β-

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chitin, organized in layers of β-1,4 linked N-acetylglucosamine (NAG) chains arranged in an

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antiparallel or parallel manner, respectively. For hydrated β-chitin, only the [100] lattice plane

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exposes H1 atoms that are abstracted during LPMO catalysis (Fig. 2A). This is analogous to

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what is observed for Iβ-cellulose, the structural organization of which resembles hydrated β-

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chitin. Regarding α-chitin, the [100] lattice plane has previously been used to represent the

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polysaccharide surface42 but the [110] lattice plane should also be considered since it exhibits

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a smoother and more densely packed surface (Fig. 3B). In this study, using crystal structure

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coordinates of α-chitin43 and hydrated β-chitin44 we built models for β-chitin (β[100]) made of

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5 layers of 5 parallel (NAG)24 chains, and models for the two α-chitin types (α[100] and

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α[110]) made of 5 layers of 6 antiparallel (NAG)24 chains.

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In theory, the LPMO substrate binding patch could interact with the crystalline

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substrate surface in any orientation around an axis orthogonal to the α- and β-chitin lattice

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planes (Fig. 3A). The experimental data presented in this paper, demonstrating the orientation

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of a chitin chain bound to SmAA10A (Fig. 1), substantially reduced the degrees of freedom to

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be considered when placing the enzyme on the chitin surface. Knowing that the Cu atom must

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be close to the hydrogen atom to be abstracted during catalysis, the enzyme was placed ~5 Å

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above the chitin surface such that the Cu ion was hovering directly above the exposed H1 and

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H4 atoms, but without pre-determined selectivity. In the SmAA10A-β[100] start model the

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Cu-H1 and Cu-H4 distances were 5.7 and 5.3 Å, respectively. Corresponding distances of 5.5

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and 5.7 Å were measured for both the SmAA10A-α[100] and α[110] complexes. As a control,

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we also built start models where the enzyme was rotated 180° around the Cu atom along an

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axis orthogonal to the chitin surface. All models were solvated in a rectangular box of TIP3P

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water, with the edges at least 16 Å away from the solute, using chloride ions for

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neutralization. The solvated models contained in total ~160 000 atoms. The protein ff14SB45

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and carbohydrate GLYCAM_06-j46 force fields and our own active site parameters (see

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above) were used to generate input files for AMBER16. Input files for SmAA10A-chitin

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complexes were prepared using the force field parameters derived for Cu(I) to mimic an

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active enzyme with weak Cu-water interactions. When setting up the SmAA10A in solution

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model, the parameters derived for Cu(II) active sites, which increase the affinity for

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coordinating water molecules, were applied. The difference in water affinity for the Cu(I) and

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Cu(II) active sites originates from the dissimilar Cu partial charges (see Table S1).

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Molecular dynamics (MD) simulations. The models were subjected to 2500 steps of energy

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minimization with 5 kcal.mol-1.Å-2 positional restraints on the non-hydrogen atoms of the

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enzyme and chitin molecules to relieve abnormal molecular contacts. Then, the systems were


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heated linearly from 0 K to 300 K for 40 ps at constant volume, with restraints lowered to 1

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kcal mol-1 Å-2, using the Langevin thermostat with a collision frequency of 1 ps-1. Density

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equilibrations were run at 300 K for 0.5 ns at a constant pressure of 1 atm using the

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Berendsen barostat with a pressure relaxation time of 1 ps. The final 50 ns equilibration step

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was carried out in the NVT ensemble using the weak coupling algorithm and a time constant

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of 10 ps to regulate the temperature. In this step, 2 kcal.mol-1.Å-2 positional restraints were

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applied to the C1 atoms of the lowest layer of NAG chains of the chitin models (the “highest”

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layer being the one interacting with the LPMO). The 300 ns production runs were performed

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using the same conditions as in the final equilibration step, storing a snapshot every 20 ps,

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yielding 15,000 snapshots for each trajectory. In all simulations, we used a time step of 2 fs,

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periodic boundary conditions with a 12 Å cutoff for non-bonded interactions and PME

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treatment of long range electrostatics, while hydrogen atoms were constrained by the SHAKE

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algorithm. Simulations were carried out using the CUDA version of PEMEMD included in

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AMBER16.47 Analysis of production trajectories was performed using the cpptraj module

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included in AmberTools.48

227 228

Quantum mechanics/molecular mechanics (QM/MM) calculations. The QM/MM model of

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SmAA10A in absence of chitin (model 1) was taken from a 50 ns MD trajectory (generated

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using the same equilibration procedure as for the LPMO-chitin complexes), applying a filter

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that selects frames having an active site geometry (all non-hydrogen atoms of H28, H114,

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A112, F187, and E60) with RMSD < 1 Å relative to the crystal structure (2BEM:C).

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The QM/MM model (model 2) of SmAA10A in complex with chitin was taken from the 300

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ns SmAA10A-β[100] trajectory, applying the same filter as described above, with the addition

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of a Cu-H1 distance window of 3.8 ± 0.2 Å (the average Cu-H1 distance).

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QM/MM models 1 and 2 both included Cu(II) and the enzyme residues H28 (link atom C),

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H114, A112, F187 and E60 (link atom Cα) in the QM region. Model 1 furthermore included

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two Cu-coordinating water molecules (these two Cu-coordinating water molecules were

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observed throughout the MD trajectory). Model 2 additionally included one water molecule in

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a near equatorial position (2.9 Å away from Cu in the MD-snapshot starting structure) and

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two NAG units in the QM-region. For NAG2, the link atoms were the O1 of the reducing end

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and the C1 of the NAG unit preceding the non-reducing end.

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Finally, a plausible18,49,50 copper-oxygen reactive species ([CuO]+ core, singlet state) model,

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using the geometry optimized model 2 with oxygen replacing the Cu-coordinating water

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molecule in the equatorial position, was generated.

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The QM/MM interface in AMBER51 with ORCA352 providing the QM energy and

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gradient was used to geometry optimize the models described above. The BP86 GGA


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functional,53,54 which has previously been shown to yield conform geometries for transition-

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metal complexes,55 was selected, and dispersion was included through the Grimme’s DFT-D3

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approach with Becke-Johnson dampening.56,57 The Def2-TZVPP basis set was used for Cu

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and Def2-SVP for all other atoms,58,59 and the RI-approximation was applied to speed up the

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calculations.

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The following 3-stage geometry optimization scheme was applied to the QM/MM models; i)

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100 steps of unrestrained QM/MM optimization with the TNCG (Truncated Newton linear

255

Conjugate Gradient method with LBFGS preconditioning) algorithm, ii) 10,000 MM steps

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with the conjugate gradient method (no QM) where all QM atoms used in step i) were frozen

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using 1000 kcal.mol-1.Å-2 positional restraints, iii) TNCG QM/MM optimization until

258

convergence (drms = 10-4 kcal.mol-1.Å-1). Atoms more than 6 Å away from the QM region

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were frozen using 1000 kcal.mol-1.Å-2 positional restraints during this step.

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Computation of EPR spin Hamiltonian parameters. EPR parameters were computed from

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the QM part of the converged QM/MM geometries using ORCA352 with the hybrid functional

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PBE060 and the chain-of-spheres approximation. The (NAG)2 part of the model was omitted

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to reduce system size. This was considered safe considering that for model 2 only 2.5 % of

265

the spin density was located at (NAG)2. Scalar relativistic effects were included with ZORA.

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The basis sets Def2-TZVPP and Def2-TZVP were decontracted and used for Cu and all other

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atoms, respectively. The integration grid (Grid6) was increased for Cu (SpecialGridIntAcc 7).

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The following contributions were included when calculating the hyperfine couplings: 1) the

269

isotropic Fermi contact term, 2) the magnetic spin dipole interaction between the nucleus spin

270

and electron spin and 3) the second order contribution form spin-orbit coupling.

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RESULTS AND DISCUSSION

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Overview. The catalytic core of the LPMO, a Cu-N3 complex hosting a reactive oxygen

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species, must be carefully maneuvered close to the inert C-H bond to be functionalized

275

leading in fine to polysaccharide chain cleavage. LPMOs acting on the β(1,4) glycosidic

276

bonds of chitin or cellulose often display strict regio-selectivity, meaning that they

277

exclusively hydroxylate the C1 or C4 carbon, which are separated by 2.5 Å in the

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polysaccharide chain. Notably, there is no known inherent property of the active site itself that

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could determine this regioselectivity. LPMO action on chitin leads to oxidation at C1. The

280

LPMO active site is part of a relatively flat, solvent-exposed surface displaying conserved

281

amino acid side chains that have been shown by experiments23,26 to be crucial for enzyme

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activity. There are no available crystallographic data or models describing interactions

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between an LPMO and chitin or (soluble) chito-oligosaccharides that would allow


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rationalization of the mode of action of these enzymes. Therefore, to fill this gap and provide

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sound fundaments for future mechanistic studies, we have built atomistic models of enzyme-

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substrate complexes, the validity of which is supported by experimental data.

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Determining the relative orientation LPMO-substrate. In 2005, a site-directed mutagenesis

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study conducted on SmAA10A (then called CBP21) revealed that several surface exposed

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amino acid side chains (Tyr54, Glu55, Glu60, His114, Asp182 and Asn185) are important for

291

binding to chitin.26 Complementary to this study, and subsequent to the discovery of LPMO

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activity in 2010,4 HSQC NMR experiments demonstrated that β-chitin shields the backbone

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amides of specific residues of SmAA10A strongly (Tyr54, Glu55, Gln57, Ser58 and Thr111)

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or moderately (Gln53, Leu110, Ala112, His114 and Thr116) from deuterium exchange with

295

bulk solvent (Fig. S1).23 Most of these residues are highly conserved within the chitin-active

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AA10 subfamily. The strongly affected residues form an elongated patch on the SmAA10A

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surface (Fig. S1), stretching from Tyr54, through the Cu-site, to Asp182 (Fig. 1). Knowing

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that this patch and the Cu-site must face the surface of the crystalline substrate limits the

299

number of possible enzyme orientations. However, the enzyme can still be rotated 360º on the

300

substrate surface, leaving a substantial number of conformations to be explored.

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We hypothesized that a chitin oligomer, (N-acetyl-glucosamine)6, or (NAG)6, could

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cover this area with conserved residues and possibly be cleaved, although SmAA10A has only

303

been reported to be active on insoluble substrates.4 Using high enzyme loadings we indeed

304

detected SmAA10A activity on (NAG)6 (Figs. 1 and S2), but not on shorter oligomers (Fig.

305

S3). Importantly, the dominating oxidized product had a degree of polymerization of 4

306

(DP4ox), clearly indicating that there is one preferred binding orientation that, if we assume

307

that the substrate indeed aligns with the stretch of conserved residues, places the reducing end

308

of the hexamer near Asp182 and the non-reducing end near Tyr54 (Fig. 1 & S4A). In the case

309

of the opposite orientation, DP2ox would have been dominating (Fig. S4B), which is not the

310

case (Fig. S2A). The determination of this relative orientation allows introduction of a

311

putative subsite numbering, according to the CAZyme nomenclature,61 with subsites spanning

312

from -4 to +2, involving a stretch of exposed residues running from Tyr54 to Asp182,

313

respectively (Fig. 1). Figs. 1 and S2 also show production of DP5ox and, to a lesser extent,

314

DP3ox from NAG6, meaning that other productive binding modes are possible (Fig. S4E-H).

315

The production of considerable amount of DP5ox supports the notion that the substrate is

316

oriented with its non-reducing end pointing towards Tyr54 (Fig. S4E).


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 10 of 29

317 318 319 320 321 322 323 324 325 326 327 328 329 330 331 332 333 334 335 336 337

Figure. 1. Probing the binding orientation of (NAG)6 in SmAA10A and the role of Tyr54. Panel (A) shows HPAEC-PAD chromatograms of C1-oxidized chito-oligosaccharides released from β-chitin (10 g.L-1) or from (NAG)6 (1 mM) by SmAA10A-WT or its Y54A mutant (10 µM) after 60 min of reaction. Reactions were initiated by adding AscA (1 mM) and carried out in Tris-HCl buffer (50 mM, pH 8.0), with incubation at 40 °C in a thermomixer (1000 rpm). In control reactions SmAA10A was replaced by Cu(II)SO4 (10 µΜ). Panel (B) shows time-courses for the release of DP4ox in reactions described in panel A. Note: the enzyme is no longer active at the first time point, indicating fast inactivation, a selfdestructive reaction that was notably shown to occur in absence of substrate.22,62 Such timecourse profile with (NAG)6 is thus probably diagnostic of weak binding to this substrate. The error bars show ± s.d. (n = 3, independent experiments). Panel (C) shows a model (extracted from MD simulations performed on β-chitin; see Fig. 3) of SmAA10A interacting with (NAG)6 (see Fig. S1 for a 90°-rotated top view). The histidine brace, composed of His28 and His114, and bound Cu are shown in orange. The side chains of Tyr54 and Asp182, in subsites -4 and +2, respectively, are also shown (grey sticks). The C1 of the NAG unit located in subsite -1 is the carbon subject to hydroxylation during catalysis by SmAA10A. Abbreviations: DP, degree of polymerization; NAG, N-acetylglucosamine; NR, non-reducing end; R, reducing end.

338

To confirm the positioning of the oligosaccharide in the active site we tested the

339

effect of the Y54A mutation, in the proposed subsite -4, on the activity towards (NAG)6.

340

Expectedly, this mutation decreased the activity of SmAA10A on (NAG)6 (Fig. 1B).

341

Furthermore, the production of DP5ox was much more affected than that of DP3ox (Fig. 1A),

342

which is consistent with the notion that Tyr54 in subsite -4 is involved in the binding mode


Page 11 of 29 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

Biochemistry

343

leading to formation of DP5ox (Fig. S4E). Interestingly, the effect of the Y54A mutation on

344

the activity towards β-chitin was minimal (Fig. 1). The crystalline structure of β-chitin,

345

possibly provides additional anchoring points for the LPMO (see MD simulations below) and

346

this may dampen the effect of the Y54A mutation.

347 348

Molecular dynamics simulations of SmAA10A-chitin interactions. The experimentally

349

determined orientation of the substrate relative to the LPMO was used as input to generate

350

large scale MD start models of SmAA10A-Cu(I) interacting with α- and β-chitin. Figure 2,

351

based on analysis of transmission electron microscopy images63,64 and crystal structures43,44 of

352

α- and β-chitin, shows microfibril cross-sections and lattice planes. Any putative interaction

353

of SmAA10A with the solvent exposed surfaces [120] and [12ത0] of the microfibril would not

354

allow the enzyme active site to get close to the glycosidic bond for catalysis to occur. For the

355

[010] surfaces, the glycosidic bonds are not exposed to the solvent at all. In the case of β-

356

chitin, alike Iβ-cellulose, the surface with exposed glycosidic bonds is defined by the lattice

357

plane [100], called β[100]. In the case of α-chitin the situation is less straightforward since

358

two lattice planes could be considered, namely α[100] and α[110] (Fig. 2). The MD results

359

presented here (see below) show that SmAA10A can bind in a productive manner to all these

360

three chitin surface topologies. It is interesting to note that the mature β-chitin microfibril

361

from the diatom Thalassiosira weissflogi displays a SmAA10A accessible surface (Fig. 2A)

362

while the β-chitin microfibril from the tubeworm Lamellibrachia satsuma would need peeling

363

before a [100] surface appears (Fig. 2B). The mature α-chitin microfibril from the alga

364

Phaeocystis globose also needs trimming before a large [100] or [110] surface emerges (Fig.

365

2C). Therefore, LPMO efficiency likely depends on the source of the polysaccharide, as has

366

indeed been observed.34

367


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

368 369 370 371 372 373 374 375 376 377 378 379 380 381 382 383 384 385 386 387 388 389 390 391 392 393 394 395 396

Page 12 of 29

Figure. 2. Possible binding sites for SmAA10A on crystalline chitin fibers. The figures show scaled cross-sections of a β-chitin microfibril from T. weissflogi (A),64 a β-chitin microfibril from L. satsuma (B)64 and an α-chitin microfibril from P. globose (C)63 derived from electron micrographs and crystal structures. Each microfibril is composed of a multitude of individual chitin chains (shown as black jagged lines with an endon view, i.e. oriented perpendicular to the cross-section plane) arranged in a parallel (for A and B) or anti-parallel manner (for C). The green lines indicate the lattice plane surfaces on the crystalline chitin fibers that allow productive LPMO binding (relatively flat surfaces where H1-atoms are exposed towards the enzyme active site). The surfaces indicated by red lines are very rough and display deep grooves that are incompatible with productive LPMO binding because steric clashes between the chitin chains and the enzyme will not allow the necessary proximity between the copper center and the H1-atoms to be abstracted.


Page 13 of 29 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

Biochemistry

397

A global view of SmAA10A interacting with β[100] is shown in Fig. 3A and a movie

398

illustrating the enzyme docking on the chitin surface is available as supporting information

399

(Movie S1). End-on views (i.e. looking along the polysaccharide chains) of the β[100],

400

α[100], and α[110] crystalline chitin models reveal very different surface topologies (Fig.

401

3B). β[100] chitin is nearly planar, whereas α[100] displays deep grooves between the

402

polysaccharide chains. In the α[110] model, the polysaccharide chains overlay slightly, and

403

the grooves are shallow compared to α[100]. The MD simulations of the three SmAA10A-

404

chitin models showed that the alignments of the copper ion and the histidine brace with the

405

to-be-cleaved glycosidic bond, are similar (see Fig. S5A), with, notably, identical average

406

Cu-H1 distances (Fig. 3E). Also, the SmAA10A active site geometry is not altered

407

significantly upon substrate binding (Fig. 3C and Fig. S5B). The Cu-H1 distances are 3.8 Å

408

on average and are always more than 0.8 Å shorter than the corresponding Cu-H4 distances

409

(Fig. 3E). The closer proximity to C1 is consistent with the established C1 oxidative

410

regiospecificity of SmAA10A4 (see discussion below and Fig. 5C).

411

Both the LPMO and the chitin displayed very stable packing during all simulations

412

with average RMSDs of ~ 0.5 Å for chitin-C1 atoms and ~ 0.8 Å for SmAA10A-Cα atoms

413

(Fig. 3E). Comparison of SmAA10A-Cα atom positions relative to chitin-C1 atom positions

414

showed that the SmAA10A-chitin complexes are also relatively stable (Fig. 3E) although

415

much wider distributions are observed (see Fig. S6 for time-resolved data). SmAA10A-β[100]

416

and SmAA10A-α[110] showed considerably lower RMSD values (1.16 and 0.96 Å) compared

417

to SmAA10A-α[100] (1.49 Å) (Fig. 3E), indicating that SmAA10A can move more freely on

418

the α[100] surface, probably due to the difference in surface topology (Fig. 3B).

419

To visualize the origin of these LPMO-chitin complex RMSD distributions, a

420

principal component analysis (PCA) was applied to identify the most significant concerted

421

motions, referred to as normal modes #1 and #2, with mode #1 having the highest eigenvalue.

422

The PCA shows that the largest motion mode for SmAA10A-β[100] and SmAA10A-α[110] is

423

a rotating motion (mode #1, Fig. S7) around an axis orthogonal to the chitin surface and

424

centered close to the histidine brace. The second most significant modes show mixed

425

translational/rocking and rotational/rocking modes for SmAA10A-β[100] and SmAA10A-

426

α[110], respectively (mode #2, Fig. S7). The α[100] model, with its much more rugged

427

surface (Fig. 3B) shows an opposite arrangement of mode #1 and #2, with more

428

rocking/translational movement in mode #1 and a rotating motion for mode #2 (Fig. S7).

429

The intermolecular interactions between SmAA10A and the chitin models were

430

dissected by close contact and hydrogen bond analysis of the MD-trajectories, enabling

431

detailed examination at the residue level. Fig. 3D shows heat-maps for close contacts of

432

SmAA10A with crystalline chitin for each of the three models. The close contact analysis

433

reveals enzyme residues with atoms within a distance of 6 Å from any NAG atom in the


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 14 of 29

434

trajectory (Fig. 3F). A general observation is that the heat-maps correspond well with the

435

patch of surface-exposed, conserved amino acid residues thought to be involved in substrate

436

binding by SmAA10A (see Fig. S1). There are, however, differences between the three chitin

437

forms, likely because of their distinct surface topologies. The interactions between SmAA10A

438

and β[100] or α[110], which have relatively flat surfaces, involve a larger fraction of the

439

surface compared to α[100] (Fig. 3D). Fig. 3F shows that 71%, 70% and 66% of all measured

440

close contacts occur between the enzyme and the to-be-cleaved polysaccharide chain

441

(subsites -5 to +3) for β[100], α[100] and α[110], respectively. This is consistent with

442

biochemical data presented above but also shows that the contribution of adjacent chains is

443

not negligible in a crystalline context. The close contact analysis also reveals that the

444

aromatic ring of Tyr54 is close to the NAG unit in subsite -4 during the simulation,

445

supporting the idea that solvent-exposed aromatics in LPMOs, of which there often are only

446

few, are key factors for docking onto the polysaccharide. The hydrogen bond analysis

447

allowed us to see that residues Glu55, Thr111, His114, Gln57 and Asp182 form well defined

448

hydrogen bonds to chitin (Fig. S8B). For an in-detail view of amino acids interacting with

449

each subsite see Fig. S9.

450

Additionally, the models presented here allow to investigate the structural differences

451

that may be involved in the preference for one chitin allomorph over the other, a matter on

452

which sound biochemical data are scarce owing to substrate heterogeneity and lack of

453

systematic comparison. Experiments have shown that SmAA10A displays a binding

454

preference for β-chitin over α-chitin.23,65 Here, the close contact (Fig. 3) and, especially, the

455

hydrogen bond (Fig. S8) analyses pointed notably at Arg113 as prominent in subsite c when

456

interacting with β[100], but not interacting with α[100]. Arg113, not considered or detected

457

in previous studies, is a non-conserved residue found in the middle of the motif “TAXH”,

458

totally conserved in the chitin-active AA10 sub-family, where H is His114 of the histidine

459

brace and X = A, M, P, Q or R. Our models suggest that mapping out phenotypic sub-clades

460

(e.g. allomorphs preferences or degrees of chitin acetylation) among chitin-active AA10s may

461

require to consider non-conserved surface-exposed residues.

462

The previous mutagenesis26 and NMR23 studies aiming at mapping interactions

463

between SmAA10A and β-chitin relied on different principles and gave slightly different

464

results (Fig. S1), which now can be explained. For instance, Glu60, Asp182, and Asn185

465

were not detected in the NMR approach but found to be important in the mutagenesis study.

466

The MD simulations reveal that Glu60 and Asp182 to a large extent, and Asn185 to a lesser

467

extent, interact with β[100] through bridging water molecules (Fig. S8C). This could explain

468

why these residues are important for binding but were not detected by the NMR approach,


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Biochemistry

469

which was based on detecting protection of the backbone amides from exchange with solvent

470

by chitin binding.

471

Overall, there is a striking compliance between the previous mutagenesis and NMR

472

data and the MD-simulations presented here (Fig. S1). The present experimentally supported

473

models complete the picture by revealing details of the interaction of each amino acid with

474

crystalline chitin. As a control experiment, we also investigated the ability of the enzyme to

475

bind to the polysaccharide surface in the opposite direction by running control simulations for

476

all the model substrates where SmAA10A was rotated 180° around an axis orthogonal to the

477

chitin surface and centered on the copper atom. With α[100] or α[110]-chitins, the enzyme

478

drifted away from the substrate (results not shown). With β[100]-chitin a complex was

479

formed, which, however, did not look plausible with a O-H1 distance of 4.2 Å and a Cu-O-H1

480

angle of 86° (see Fig. 5C for comparison) and few intermolecular interactions involving

481

conserved residues were observed (not shown). Taken together, the simulations and

482

experimental data strongly indicate a substrate orientation where the non-reducing end of the

483

scissile polysaccharide chain points towards Tyr54. Surprisingly, this conclusion contrasts

484

with the crystal structure of a fungal, cellulose-active, C4-specific AA9 LPMO in complex

485

with cello-oligomers, which shows an opposite substrate/LPMO relative orientation (Fig.

486

S10).25 Despite this striking difference, in both cases, the N-terminal histidine is located in the

487

subsite hosting the sugar unit to be modified, i.e. in the +1 subsite for C4-oxidation (e.g.

488

LsAA9A) or in the -1 subsite for C1-oxidation (e.g. SmAA10A). Also, in both systems, a Tyr

489

residue is located in the negative subsites interacting with the non-reducing end of the

490

substrate, three subsites away from the subsite where catalysis occurs (Fig. S10). Altogether,

491

this suggests that the evolutionary shift between bacterial strict C1-oxidizers (belonging to

492

AA10s) and fungal strict C4-oxidizers (belonging to AA9s) may not originate from a simple

493

“small” translation of the active site over the glycosidic bond to be cleaved. Instead, an

494

appropriate and significantly different network of interactions must have been evolved so that

495

the reactive intermediate is properly poised over the target C-H bond for catalysis to occur.

496

On the contrary, it is more likely that the C1 vs C4 oxidative regioselectivity divergence

497

occurring within common phylogenetic groups (e.g. within fungal AA9s or bacterial AA10s)

498

is the result of more subtle interactions re-arrangements, as suggested by recent mutagenesis

499

studies.66,67 This inversed substrate-LPMO orientation is intriguing and its impact, on

500

substrate specificity and/or oxidative regioselectivity, deserves further investigations.

501 502 503


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

504 505 506 507 508 509 510 511 512 513 514 515 516 517 518 519 520 521 522 523 524 525 526

Page 16 of 29

Figure 3. Interactions of SmAA10A with chitin probed by MD simulations. (A) Global view of SmAA10A interacting with β-chitin (SmAA10A-β[100]); explicit solvent excluded). (B) End-on view, looking along the polysaccharide chains, of the crystalline chitin models β[100], α[100], and α[110]. The black solid lines indicate the lattice planes corresponding to the LPMO binding surfaces. (C) Averaged structures of SmAA10A in solution (brown) and bound to β-chitin (green) from 300 ns trajectories show that the copper ion coordinating amino acid side chains retain their conformation upon substrate binding (see Fig. S5 for more details). (D) Heat-mapped contact surfaces of SmAA10A when interacting with crystalline chitin. The color gradient, from blue to red, indicates the frequency of contacts from low to high, respectively. For the sake of clarity, only the NAG units responsible for most of the contacts are represented as filled rings, identified by numbers (-5 to +3) or letters (a, b, …) when belonging to the main chain or adjacent chains, respectively. (E) The left panels show the distribution of RMSD values for chitin C1 atoms (green), enzyme Cα atoms (yellow) and chitin C1 atoms versus enzyme Cα atoms (purple), whereas the right panels show the Cu-H1 (red) and Cu-H4 (blue) distances, as derived from 300 ns simulations, fitted with the kernel probability density function (the corresponding time-resolved data are shown in Fig. S7). (F) Number of close contacts between NAG units of the polysaccharide, numbered according to the binding sites indicated in panel D, and residues of SmAA10A. Close contacts with less than 20 contacts were not included in the analysis. Contacts involving sugars bound to subsites -5 to +3 dominate and their fraction of the total contacts is indicated for each chitin model (66 - 71 %). Blue bars indicate conserved residues and green bars non-conserved residues (see Fig S1 for conservation scores). See Fig. S9 for visualization of the interactions


Page 17 of 29 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

Biochemistry

527 528 529

and Fig. S8 for analysis of hydrogen bond and water bridges. Abbreviations: NAG, N-acetylglucosamine.

530 531

MD simulations reveal a gated, main access to the active site. An important aspect of LPMO

532

catalysis pertains to the accessibility of the copper site when the enzyme is bound to the

533

polysaccharide. The idea that the copper site would be confined and shielded from bulk

534

solvent, is appealing since it would allow good control of the ongoing redox chemistry, but

535

also raises questions for example because the copper site would be shielded from potential

536

reducing agents that are needed to deliver electrons. Due to the latter conundrum, the

537

existence of an intramolecular electron transfer pathway has been proposed.20,21 The existence

538

of such an electron transfer pathway has never been demonstrated and is in fact not necessary

539

when considering H2O2 as a co-substrate of LPMO catalysis.22 In a H2O2-based mechanism,

540

electron delivery to the enzyme-substrate complex is not needed, provided that the enzyme

541

was reduced prior to binding. A recent kinetic study of SmAA10A supports the involvement

542

of H2O262 and it has also been shown that reduction of the copper promotes substrate

543

binding.68

544

Our models reveal the existence of a solvent tunnel formed between SmAA10A and

545

the chitin surface upon substrate binding. The ~12 Å long tunnel, predicted using Caver,69

546

connects the bulk solvent to the copper site and consists of a narrow monolayer of water

547

molecules (Fig. 4A). The Glu60 residue, associated with water bridged substrate interactions

548

(see above), displays three different rotamers (R1, R2 and R3) (Fig. 4B), influencing access

549

to the active site (Fig. 4C). Other residues lining the tunnel, e.g. Asn185, display limited

550

fluctuations compared to Glu60. For each configuration, a priority parameter (0 ≤ Pt ≤ 1) is

551

calculated on the basis of how functional a tunnel is predicted to be with respect to molecule

552

transport and how often the tunnel is observed in the ensemble. A high Pt value indicate a

553

functional tunnel observed frequently in the ensemble. In the β-chitin model, the R1 state

554

corresponds mainly to a “closed” tunnel (with a low Pt of 0.01) (Fig. 4D). In contrast, in the

555

less populated states R2 (Pt = 0.49) and R3 (Pt = 0.59) the Glu60 side chain is rotated away

556

from the crystalline surface, increasing access to the active site. The different surface

557

topologies of α-chitin, α[100] and α[110], seem to reduce the importance of Glu60 as an

558

gatekeeper (see Table S2 for details). Importantly, for all three chitin models the tunnel

559

maximum bottleneck radius is ~1.6 Å, meaning that the tunnel is too narrow to allow large

560

molecules such as ascorbate or diphenolic compounds (which are known LPMO reductants)

561

to reach the active site. Our data are therefore in favor of a scenario where copper reduction

562

occurs prior to polysaccharide binding, as supported by a recent study showing better

563

substrate binding by the LPMO-Cu(I) versus LPMO-Cu(II).68 However, the tunnel would


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 18 of 29

564

allow molecules such as O2, O2•-, H2O2 or H2O to diffuse in and out of the active site. Given

565

the open and closed conformations observed for Glu60 we speculate that this residue is

566

involved in regulation of the access of reagents (e.g. H2O2) to the copper center and may thus

567

be crucial for LPMO activity but also for the enzyme’s operational stability. How such a

568

gatekeeping function would be controlled remains to be investigated. Of note, residues that

569

could be functionally analogous to Glu60 (mostly Glu or Gln) seems to appear in all

570

LPMOs29 and mutation of Glu60 in SmAA10A drastically reduced its boosting effect on

571

chitinase action26.

572

573 574 575 576 577 578 579 580 581 582 583 584 585 586 587

Figure 4. Accessibility of the active site in the SmAA10A-β[100] complex. “(A) An active site access tunnel (shown as beige surface), formed upon binding of SmAA10A to the polysaccharide, connects the active site copper (shown as orange sphere) with the bulk solvent. (B) MD-simulations show that the side chain of Glu60 displays three main rotamer populations indicated by R1, R2 and R3, which shape the morphology of the tunnel at proximity of the active site (see panel D). Asn185, which is also involved in the definition of the tunnel shape, displays only restricted fluctuations, not influencing the tunnel shape. (C) Snapshots representative of the different rotamer populations. (D) Approximately 500 snapshots were analyzed for each population of R1 (left panel), R2 (middle panel) and R3 (right panel) using Caver.69 The green lines indicate the resulting tunnels identified for the different populations, and the blocking effect of Glu60 is apparent for the R1 population. Parameters describing the tunnels are given in Table S2.

588 589

Insights into active-site properties upon substrate binding by QM/MM and EPR.

590

Interestingly, the MD simulations did not reveal substantial structural alterations of the active

591

site amino acids due to substrate-binding (Fig. 3C and S5B), although EPR experiments (see

592

below) clearly show that the SmAA10A copper environment is altered upon binding (Fig. 5).


Page 19 of 29 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

Biochemistry

593

Previously, it has been speculated that rearrangement of Cu-coordinating waters would cause

594

the change in the EPR spectrum when AA9s bind substrate.70 In a subsequent study, a similar

595

substrate binding-induced effect on the EPR signal was observed, but only in presence of

596

chloride, leading to the suggestion that binding of co-substrate plays a role in modulating the

597

EPR spectrum (see below).25

598 599 600 601 602 603 604 605 606 607 608 609 610 611 612 613 614 615 616 617 618 619 620 621 622 623 624

Figure 5. X-band EPR spectra of SmAA10A recorded in the absence and presence of substrate. The figures show EPR spectra (black line) of SmAA10A-Cu(II) (200 µM) in the absence of substrate (A) and in the presence of (NAG)6 (a soluble substrate) (B) or two forms of crystalline chitin, β- or αchitin (solid substrates) (C). Simulated spectra are also shown (sim, grey line); in panel C, only the simulation of the EPR spectrum recorded with β-chitin is displayed. Panel (D) shows the superhyperfine structure of the gx,yregion of the spectrum recorded for SmAA10A-Cu(II) in the presence of β-chitin. The simulation of this spectrum is based on the g-values and AzCu values determined from the experimental spectrum and Ax,yCu and AN values computed by DFT (see main text). All samples were prepared in MES buffer (50 mM, pH 6.0). All spectra were recorded at 77 K, 1 mW microwave power and 10 Gauss modulation amplitude. Simulated and computed spin Hamiltonian parameters are listed in Table 1.

625 626 627 628 629 630 631 632 633 634 635 636

To connect the computational results with spectroscopic data, two models of

637

SmAA10A-Cu(II), namely 1 (in solution, Fig. 6A) and 2 (with crystalline β-chitin, Fig. 6B),


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 20 of 29

638

were subjected to QM/MM calculations. As should be expected for an accurate Cu(II)-active

639

site model, the geometry optimized model 1 displays a water coordination mode and active

640

site geometry closely resembling the low X-ray dose structure of the chitin-active AA10A

641

from Enterococcus faecalis (EfAA10A, PDB ID 4ALC) (Fig. S11), which is homologous to

642

SmAA10A (49 % sequence identity).

643

QM/MM calculations show that model 1 displays a distorted intermediate of the

644

trigonal bipyramid and square pyramid geometries (Fig. 6A), in agreement with the

645

experimentally observed rhombic EPR spectrum (d(x2-y2) ground state) of SmAA10A (Fig.

646

5A, Table 1). Superposition of model 1 and model 2 shows that one of the Cu-coordinating

647

water molecules of model 1 would clash with the bound chitin (Fig. S12). Consequently,

648

simultaneous presence of two Cu(II)-coordinating water molecules and substrate is not

649

feasible. Thus, rearrangement of at least one of the water molecules is expected when

650

SmAA10A binds chitin. On that note, Frandsen et al. have shown that upon formation of a

651

complex between a fungal LPMO and a cello-oligomer the copper-coordinating water in the

652

axial position is displaced by the C6 hydroxymethyl group of the +1 glucose and the

653

equatorial water molecule is replaced by a chloride ion.25 Here, the QM/MM geometry

654

optimized model 2 show that the single water molecule binding to Cu(II) lies in the equatorial

655

position, resulting in a distorted square planar geometry (Fig. 6B). This change in copper site

656

geometry agrees with spectroscopic experiments, since the SmAA10A-substrate EPR

657

envelopes indicate a shift towards axial symmetry compared to the free enzyme (Fig. 5B&C,

658

Table 1).

659

The spin Hamiltonian parameters for model 1 and 2 were computed and are

660

compared to the experimentally obtained values in Table 1. Mulliken charges and spin

661

populations are listed in Table S3. The experimental spin Hamiltonian parameters were

662

acquired by simulation of the spectra (Fig. 5). The experimentally determined gx, gy, AxCu and

663

AyCu values should only be considered as estimates while gz and AzCu could be accurately

664

determined. Though the computed gz and AzCu values deviate from the experimentally

665

observed values, a well-known situation in studies on Cu-complexes by DFT71, the trends of

666

effects induced by binding to crystalline substrate were reproduced: gz values decrease and

667

|AzCu| values increase. Notably, binding of (NAG)6 had slightly different spectral features

668

compared to binding of β-chitin, where binding of (NAG)6 resulted in a weaker shift of AzCu

669

(from 346 to 560, compared to 610 MHz for β-chitin). This is not surprising, considering the

670

higher flexibility of (NAG)6 compared to crystalline β-chitin, which will lead to a larger

671

distribution of binding geometries. In the presence of (NAG)6, a weak superhyperfine

672

splitting pattern appears in the gx,y region of the spectrum (Fig. 5B). This phenomenon is

673

considerably more apparent in the presence of α-chitin or β-chitin, which seem to alter the

674

SmAA10A active site in a similar manner (Fig. 5C), despite the obvious differences in the


Page 21 of 29 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

Biochemistry

675

molecular organization and crystalline properties of the chitin materials (Fig. S13). In the

676

crystallographic study by Frandsen et al., the changes in EPR signature upon substrate

677

binding (and in presence of chloride) were proposed to originate from equatorial coordination

678

of the copper by a chloride ion and from changes in the hydrogen bonding pattern between

679

the N-terminal His and a non-Cu-coordinating water molecule.25 In contrast to this previous

680

study, where chloride spin was included when analyzing the EPR superhyperfine pattern and

681

where ligand nitrogens were described by isotropic hyperfine coupling constants,25 we here

682

applied the anisotropic values derived from DFT, without chloride being present. The use of

683

AxCu, AyCu, AN, ANδ and ANε values computed from the QM/MM model 2 enabled us to

684

reproduce the superhyperfine splitting features observed in the experimental SmAA10A-β-

685

chitin EPR spectrum with high accuracy (Fig. 5C and D). This shows that QM/MM model 2

686

(Fig. 6B) is a plausible example of the physical SmAA10A-β-chitin complex with one water

687

coordinating in equatorial position to the Cu(II) ion.

688 689 690 691 692 693 694 695 696 697 698 699 700 701 702 703 704 705 706 707

Figure 6. QM/MM geometry optimized active sites. Panel (A) shows the QM region of SmAA10A in absence of chitin with two water molecules coordinating to Cu(II) in a distorted intermediate of the trigonal bipyramid and square pyramid geometries. When bound to chitin, only one water molecule is sterically allowed to interact with Cu(II) and the copper site adopts a distorted square planar geometry (B). For both (A) and (B) the delocalized spin densities (yellow surfaces) indicate the effect of the interaction between Cu(II) and the ligands. Details of the active site geometries are shown on the right side in panel (A) and (B). The active site of SmAA10A modeled by QM/MM with an [CuO]+ core shows how the reactive oxygen species is directed to the H1 atom that is abstracted during catalysis, with a distance O-H1 of 2.08 Å and a Cu-O-H1 angle of 146° (C). The link atoms for NAG2 are indicated by “L” and shown as black stick and ball (see experimental section).

708


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

709

Finally,

we

12,18,21,22,49,50,72

geometry

optimized

a

Page 22 of 29

QM/MM

model

representing

a

+

710

plausible

711

site (Fig. 6C). In the light of the recent finding showing that H2O2 is a co-substrate for

712

LPMOs,22 and by analogy to calculations performed on cellulose-active AA9s,18,49,50 a Cu-oxo

713

or Cu-oxyl is suggested to be the species responsible for H-atom abstraction in the reaction

714

leading to hydroxylation of C1 in chitin (scheme 1). Thus, it was of interest to examine if a

715

[CuO]+ species in our SmAA10A-β[100] model would be compatible with catalysis. The Cu

716

bound oxygen atom aligns in a plane with Cu and the Cu-coordinating nitrogen atoms from

717

the histidine-brace (Fig. 6C). For this singlet species, our calculations predict a Cu-O distance

718

of 1.78 Å, while the O-H1 and O-H4 distances are 2.08 and 3.21 Å, respectively. The shorter

719

O-H1 distance, and also the Cu-O-H1 angle of 145.5°, are indeed what could be expected for

720

a pre-catalytic complex when comparing with data obtained with the analogues [FeO]2+

721

core.73 Moreover, the longer O-H4 distance indicate that H4 abstraction is energetically

722

unfavorable compared to H1 abstraction, in line with the observed C1 oxidizing activity of

723

the enzyme.

pre-catalytic complex with a [CuO] species in the SmAA10A active

724

725 726 727 728 729 730 731 732 733 734 735 736

Scheme 1. SmAA10A-catalyzed reaction mechanism of chitin oxidation. The resting state of SmAA10A (only copper-coordinating histidines are shown) is the Cu(II) state (shown as orange sphere) which undergoes a priming reduction yielding the active Cu(I) state (blue sphere). As proposed by Bissaro et al.22, following binding to chitin (represented by a chitobiose fragment), the reaction of the co-substrate H2O2 with Cu(I) leads to the release of a water molecule and formation of a copper-oxyl intermediate. As shown in the present study, the [CuO]+ core is properly poised to allow regioselective hydrogen atom abstraction (HAA) and subsequent hydroxylation at the C1 carbon of the -1 NAG unit, via a copper-oxyl, oxygen rebound mechanism.18 The resulting hydroxylated polysaccharide is then subject to a spontaneous elimination reaction leading to glycosidic bond cleavage.13

737 738

Table 1. Spin Hamiltonian parameters for experimental spectra and QM/MM modelsa Spin Hamiltonian parameters

SmAA10A

SmAA10A

SmAA10A

Model 1

+ (NAG)6

+ β-chitin

(SmAA10A)

Model 2 (SmAA10A + β-chitin)


Page 23 of 29 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

Biochemistry

g – tensor

ACu – tensor (MHz)

gx

2.018

2.025

2.023

2.034

2.048

gy gz

2.116

2.066

2.064

2.095

2.059

2.259

2.212

2.216

2.198

2.183

Ax

218

3

36

190

36

Ay

131

25

-25

-61

-25

Az

346

560

-610b

-484

-593

Ax, Ay, Az

n.a.c

n.a.

34, 34, 50

32, 32, 47

34, 34, 50

Ax, Ay, Az

n.a.

n.a.

39, 28, 29

42, 31, 32

39, 28, 29

Ax, Ay, Az

n.a.

n.a.

29, 38, 30

38, 30, 29

29, 38, 30

AN – tensor (MHz) ANδ – tensor (MHz) ANε – tensor (MHz)

739 740 741 742 743 744 745 746

a

Parameters in normal font are derived from simulation of experimental EPR spectra (assuming collinear g- and Atensors) and parameters in italics are computed by DFT (PBE0 hybrid functional and Def2-TZVPP for Cu and Def2-TZVP for other atoms, A-tensors are Euler rotated into g-tensor frame). Notably, parameters in italics found in the column “SmAA10A + β-chitin” are computed values obtained from model 2. b The negative sign suggested by DFT was kept while using the experimental determined AzCu when simulating the spectrum in Fig. 5C and D. c Not applicable.


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

747

Page 24 of 29

CONCLUDING REMARKS

748

We have demonstrated how biochemical analysis of LPMO catalytic action can be

749

used to assist the building of reliable enzyme-substrate models, by reducing the number of

750

possible starting molecular conformations. After extensive molecular dynamics simulations,

751

we were able to analyze interactions between the LPMO and its crystalline substrate,

752

revealing how evolutionary conserved amino acids secure tight binding. We could also

753

demonstrate that the conserved amino acids on the LPMO surface mainly interact with the

754

polysaccharide chain that ends up being cleaved by the enzyme. Our biochemical data and

755

models indicate that SmAA10A can cleave chitin chains at crystalline edges with very narrow

756

substrate surfaces as well as amorphous chitin substrates.

757

The LPMO-chitin complexes showed a well-defined LPMO-substrate association,

758

which was reproducible for all three models, adding confidence to the data. The positioning

759

of the LPMO on the crystalline surface brings the activated oxygen-species close to the

760

hydrogen atom on the C1 atom of the scissile bond, which is compatible with the

761

experimentally observed C1 specificity of SmAA10A. EPR spectra predicted on the basis of

762

our models and experimentally determined EPR spectra for SmAA10A interacting with

763

substrate were similar. It has been shown for several LPMOs that substrate binding alters the

764

Cu(II) EPR signal and several possible causes have been proposed. The present data

765

demonstrate that reorganization of Cu-coordinating water molecules is the most likely cause

766

of the changed EPR spectrum.

767

Importantly, the models showed that the mono-copper active site of SmAA10A is

768

connected to the bulk solvent through a narrow tunnel that would allow diffusion of small

769

molecules such as H2O, O2 and H2O2. This tunnel appears to be gated by Glu60, which is

770

highly conserved and known to be catalytically important from site-directed mutagenesis

771

studies. The accurate description of the LPMO-substrate complex presented here shows a

772

constrained molecular geometry of the catalytic center that likely is of crucial importance for

773

controlling the powerful oxidative chemistry carried out by LPMOs and that will help guiding

774

future mechanistic studies.

775 776

ASSOCIATED CONTENT

777

Supporting information. The supporting information contains supplemental figures 1 to 13,

778

tables S1-S3 and movie S1.

779 780

AUTHORS INFORMATION

781 782

Corresponding author * [email protected]


Page 25 of 29 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

Biochemistry

783 784 785 786 787 788

ORCID Bastien Bissaro: 0000-0001-8354-3892 Vincent G.H. Eijsink: 0000-0002-9220-8743 Gustav Vaaje-Kolstad: 0000-0002-3077-8003 Åsmund K. Røhr: 0000-0002-4956-4865

789

Notes

790

The authors declare no competing financial interest

791 792

Present address

793 794 795

‡

Faculty of Chemistry, Biotechnology and Food Science, Norwegian University of Life Sciences, N-1432 Ås, Norway

796 797

ACKNOWLEDGEMENTS

798 799 800 801 802 803 804 805 806 807 808 809 810

This work was supported by the Research Council of Norway grants 240967, 249865 and 262853. Computational work was performed on the Abel Cluster, owned by the University of Oslo, and the Norwegian metacenter for High Performance Computing (NOTUR) and the Extreme Science and Engineering Discovery Environment (XSEDE), which is supported by National Science Foundation grant number ACI-1548562, through allocation TGMCB090159. We would also like to acknowledge the Swiss-Norwegian Beamlines team at the European Synchrotron Radiation Facility, Grenoble, France, for assistance with powder diffraction experiments. We would also like to thank Prof. Einar Sagstuen and Efim Brondz for their assistance at the EPR laboratory, University of Oslo, Norway. Finally, we want to thank Dr. Jennifer Loose for supplying samples of the Y54A mutant of SmAA10A and of the chito-oligosaccharide oxidase.

811 812 813 814 815 816 817 818

AscA, ascorbic acid; CHOS, chito-oligosaccharides; DFT, density functional theory; DP, degree of polymerization; EfAA10A, AA10A from Enterococcus faecalis; EPR, electron paramagnetic resonance; LsAA9A, AA9A from Lentinus similis; LPMO: Lytic polysaccharide monooxygenase; MD, molecular dynamics; NAG, N-acetyl glucosamine; NR, non-reducing end; PCA, principal component analysis; QM/MM, quantum mechanics/molecular mechanics; R, reducing end; SmAA10A or CBP21: AA10A from Serratia marcescens

819 820 821 822 823 824 825 826 827 828 829 830 831 832

REFERENCES

ABBREVIATIONS

(1) Forsberg, Z., Vaaje-kolstad, G., Westereng, B., Bunsæ, A. C., Stenstrøm, Y., Mackenzie, A., Sørlie, M., Horn, S. J., and Eijsink, V. G. H. (2011) Cleavage of cellulose by a CBM33 protein. Protein Sci. 20, 1479–1483. (2) Quinlan, R. J., Sweeney, M. D., Lo Leggio, L., Otten, H., Poulsen, J.-C. N., Johansen, K. S., Krogh, K. B. R. M., Jorgensen, C. I., Tovborg, M., Anthonsen, A., Tryfona, T., Walter, C. P., Dupree, P., Xu, F., Davies, G. J., and Walton, P. H. (2011) Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components. Proc. Natl. Acad. Sci. 108, 15079–15084. (3) Phillips, C. M., Beeson, W. T., Cate, J. H., and Marletta, M. A. (2011) Cellobiose dehydrogenase and a copper-dependent polysaccharide monooxygenase potentiate cellulose degradation by Neurospora crassa. ACS Chem. Biol. 6, 1399–1406. (4) Vaaje-Kolstad, G., Westereng, B., Horn, S. J., Liu, Z., Zhai, H., Sørlie, M., and Eijsink, V. G. H. (2010) An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science 330, 219–222.


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

833 834 835 836 837 838 839 840 841 842 843 844 845 846 847 848 849 850 851 852 853 854 855 856 857 858 859 860 861 862 863 864 865 866 867 868 869 870 871 872 873 874 875 876 877 878 879 880 881 882 883 884 885 886 887 888 889 890 891 892

Page 26 of 29

(5) Agger, J. W., Isaksen, T., Varnai, A., Vidal-Melgosa, S., Willats, W. G. T., Ludwig, R., Horn, S. J., Eijsink, V. G. H., and Westereng, B. (2014) Discovery of LPMO activity on hemicelluloses shows the importance of oxidative processes in plant cell wall degradation. Proc. Natl. Acad. Sci. 111, 6287– 6292. (6) Frommhagen, M., Sforza, S., Westphal, A. H., Visser, J., Hinz, S. W. A., Koetsier, M. J., Van Berkel, W. J. H., Gruppen, H., and Kabel, M. A. (2015) Discovery of the combined oxidative cleavage of plant xylan and cellulose by a new fungal polysaccharide monooxygenase. Biotechnol. Biofuels 8, 101. (7) Vu, V. V., Beeson, W. T., Span, E. A., Farquhar, E. R., and Marletta, M. A. (2014) A family of starch-active polysaccharide monooxygenases. Proc. Natl. Acad. Sci. 111, 13822–13827. (8) Floudas, D., Binder, M., Riley, R., Barry, K., Blanchette, R. A., Henrissat, B., Martínez, A. T., Otillar, R., Spatafora, J. W., Yadav, J. S., Aerts, A., Benoit, I., Boyd, A., Carlson, A., Copeland, A., Coutinho, P. M., De Vries, R. P., Ferreira, P., Findley, K., Foster, B., Gaskell, J., Glotzer, D., Górecki, P., Heitman, J., Hesse, C., Hori, C., Igarashi, K., Jurgens, J. A., Kallen, N., Kersten, P., Kohler, A., Kües, U., Kumar, T. K. A., Kuo, A., LaButti, K., Larrondo, L. F., Lindquist, E., Ling, A., Lombard, V., Lucas, S., Lundell, T., Martin, R., McLaughlin, D. J., Morgenstern, I., Morin, E., Murat, C., Nagy, L. G., Nolan, M., Ohm, R. A., Patyshakuliyeva, A., Rokas, A., Ruiz-Dueñas, F. J., Sabat, G., Salamov, A., Samejima, M., Schmutz, J., Slot, J. C., John, F. S., Stenlid, J., Sun, H., Sun, S., Syed, K., Tsang, A., Wiebenga, A., Young, D., Pisabarro, A., Eastwood, D. C., Martin, F., Cullen, D., Grigoriev, I. V., and Hibbett, D. S. (2012) The paleozoic origin of enzymatic lignin decomposition reconstructed from 31 fungal genomes. Science 336, 1715–1719. (9) Johansen, K. S. (2016) Discovery and industrial applications of lytic polysaccharide monooxygenases. Biochem. Soc. Trans. 44, 143–149. (10) Wong, E., Vaaje-Kolstad, G., Ghosh, A., Hurtado-Guerrero, R., Konarev, P. V., Ibrahim, A. F. M., Svergun, D. I., Eijsink, V. G. H., Chatterjee, N. S., and van Aalten, D. M. F. (2012) The Vibrio cholerae colonization factor GbpA possesses a modular structure that governs binding to different host surfaces. PLoS Pathog. (Ghosh, P., Ed.) 8, e1002373. (11) Loose, J. S. M., Forsberg, Z., Fraaije, M. W., Eijsink, V. G. H., and Vaaje-Kolstad, G. (2014) A rapid quantitative activity assay shows that the Vibrio cholerae colonization factor GbpA is an active lytic polysaccharide monooxygenase. FEBS Lett. 588, 3435–3440. (12) Agostoni, M., Hangasky, J. A., and Marletta, M. A. (2017) Physiological and Molecular Understanding of Bacterial Polysaccharide Monooxygenases. Microbiol. Mol. Biol. Rev. 81, e0001517. (13) Beeson, W. T., Phillips, C. M., Cate, J. H. D., and Marletta, M. A. (2012) Oxidative cleavage of cellulose by fungal copper-dependent polysaccharide monooxygenases. J. Am. Chem. Soc. 134, 890– 892. (14) Eibinger, M., Ganner, T., Bubner, P., Rošker, S., Kracher, D., Haltrich, D., Ludwig, R., Plank, H., and Nidetzky, B. (2014) Cellulose surface degradation by a lytic polysaccharide monooxygenase and its effect on cellulase hydrolytic efficiency. J. Biol. Chem. 289, 35929–35938. (15) Vermaas, J. V., Crowley, M. F., Beckham, G. T., and Payne, C. M. (2015) Effects of Lytic Polysaccharide Monooxygenase Oxidation on Cellulose Structure and Binding of Oxidized Cellulose Oligomers to Cellulases. J. Phys. Chem. B 119, 6129–6143. (16) Villares, A., Moreau, C., Bennati-Granier, C., Garajova, S., Foucat, L., Falourd, X., Saake, B., Berrin, J. G., and Cathala, B. (2017) Lytic polysaccharide monooxygenases disrupt the cellulose fibers structure. Sci. Rep. 7, 40262. (17) Eibinger, M., Sattelkow, J., Ganner, T., Plank, H., and Nidetzky, B. (2017) Single-molecule study of oxidative enzymatic deconstruction of cellulose. Nat. Commun. 8, 894. (18) Kim, S., Stahlberg, J., Sandgren, M., Paton, R. S., and Beckham, G. T. (2014) Quantum mechanical calculations suggest that lytic polysaccharide monooxygenases use a copper-oxyl, oxygenrebound mechanism. Proc. Natl. Acad. Sci. 111, 149–154. (19) Kjaergaard, C. H., Qayyum, M. F., Wong, S. D., Xu, F., Hemsworth, G. R., Walton, D. J., Young, N. A., Davies, G. J., Walton, P. H., Johansen, K. S., Hodgson, K. O., Hedman, B., and Solomon, E. I. (2014) Spectroscopic and computational insight into the activation of O2 by the mononuclear Cu center in polysaccharide monooxygenases. Proc. Natl. Acad. Sci. 111, 8797–8802. (20) Beeson, W. T., Vu, V. V., Span, E. A., Phillips, C. M., and Marletta, M. A. (2015) Cellulose Degradation by Polysaccharide Monooxygenases. Annu. Rev. Biochem. 84, 923–946. (21) Walton, P. H., and Davies, G. J. (2016) On the catalytic mechanisms of lytic polysaccharide monooxygenases. Curr. Opin. Chem. Biol. 31, 195–207. (22) Bissaro, B., Røhr, Å. K., Müller, G., Chylenski, P., Skaugen, M., Forsberg, Z., Horn, S. J., VaajeKolstad, G., and Eijsink, V. G. H. (2017) Oxidative cleavage of polysaccharides by monocopper


Page 27 of 29 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

Biochemistry

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enzymes depends on H2O2. Nat. Chem. Biol. 13, 1123–1128. (23) Aachmann, F. L., Sorlie, M., Skjak-Braek, G., Eijsink, V. G. H., and Vaaje-Kolstad, G. (2012) NMR structure of a lytic polysaccharide monooxygenase provides insight into copper binding, protein dynamics, and substrate interactions. Proc. Natl. Acad. Sci. 109, 18779–18784. (24) Courtade, G., Wimmer, R., Røhr, Å. K., Preims, M., Felice, A. K. G., Dimarogona, M., VaajeKolstad, G., Sørlie, M., Sandgren, M., Ludwig, R., Eijsink, V. G. H., and Aachmann, F. L. (2016) Interactions of a fungal lytic polysaccharide monooxygenase with β-glucan substrates and cellobiose dehydrogenase. Proc. Natl. Acad. Sci. 113, 5922–5927. (25) Frandsen, K. E. H., Simmons, T. J., Dupree, P., Poulsen, J. C. N., Hemsworth, G. R., Ciano, L., Johnston, E. M., Tovborg, M., Johansen, K. S., Von Freiesleben, P., Marmuse, L., Fort, S., Cottaz, S., Driguez, H., Henrissat, B., Lenfant, N., Tuna, F., Baldansuren, A., Davies, G. J., Lo Leggio, L., and Walton, P. H. (2016) The molecular basis of polysaccharide cleavage by lytic polysaccharide monooxygenases. Nat. Chem. Biol. 12, 298–303. (26) Vaaje-Kolstad, G., Houston, D. R., Riemen, A. H. K., Eijsink, V. G. H., and Van Aalten, D. M. F. (2005) Crystal structure and binding properties of the Serratia marcescens chitin-binding protein CBP21. J. Biol. Chem. 280, 11313–11319. (27) Wu, M., Beckham, G. T., Larsson, A. M., Ishida, T., Kim, S., Payne, C. M., Himmel, M. E., Crowley, M. F., Horn, S. J., Westereng, B., Igarashi, K., Samejima, M., Ståhlberg, J., Eijsink, V. G. H., and Sandgren, M. (2013) Crystal structure and computational characterization of the lytic polysaccharide monooxygenase GH61D from the basidiomycota fungus Phanerochaete chrysosporium. J. Biol. Chem. 288, 12828–12839. (28) Nimlos, M. R., Beckham, G. T., Matthews, J. F., Bu, L., Himmel, M. E., and Crowley, M. F. (2012) Binding preferences, surface attachment, diffusivity, and orientation of a family 1 carbohydratebinding module on cellulose. J. Biol. Chem. 287, 20603–20612. (29) Vaaje-Kolstad, G., Forsberg, Z., Loose, J. S., Bissaro, B., and Eijsink, V. G. (2017) Structural diversity of lytic polysaccharide monooxygenases. Curr. Opin. Struct. Biol. 44, 67–76. (30) Frandsen, K. E. H., and Lo Leggio, L. (2016) Lytic polysaccharide monooxygenases: A crystallographer’s view on a new class of biomass-degrading enzymes. IUCrJ 3, 448–467. (31) Vaaje-Kolstad, G., Bøhle, L. A., Gåseidnes, S., Dalhus, B., Bjørås, M., Mathiesen, G., and Eijsink, V. G. H. (2012) Characterization of the chitinolytic machinery of enterococcus faecalis V583 and highresolution structure of its oxidative CBM33 enzyme. J. Mol. Biol. 416, 239–254. (32) Nakagawa, Y. S., Eijsink, V. G. H., Totani, K., and Vaaje-Kolstad, G. (2013) Conversion of  αchitin substrates with varying particle size and crystallinity reveals substrate preferences of the chitinases and lytic polysaccharide monooxygenase of Serratia marcescens. J. Agric. Food Chem. 61, 11061–11066. (33) Nakagawa, Y. S., Kudo, M., Loose, J. S. M., Ishikawa, T., Totani, K., Eijsink, V. G. H., and Vaaje-Kolstad, G. (2015) A small lytic polysaccharide monooxygenase from Streptomyces griseus targeting α- And β-chitin. FEBS J. 282, 1065–1079. (34) Forsberg, Z., Nelson, C. E., Dalhus, B., Mekasha, S., Loose, J. S. M., Crouch, L. I., Røhr, Å. K., Gardner, J. G., Eijsink, V. G. H., and Vaaje-Kolstad, G. (2016) Structural and functional analysis of a lytic polysaccharide monooxygenase important for efficient utilization of chitin in Cellvibrio japonicus. J. Biol. Chem. 291, 7300–7312. (35) Westereng, B., Agger, J. W., Horn, S. J., Vaaje-Kolstad, G., Aachmann, F. L., Stenstrøm, Y. H., and Eijsink, V. G. H. (2013) Efficient separation of oxidized cello-oligosaccharides generated by cellulose degrading lytic polysaccharide monooxygenases. J. Chromatogr. A 1271, 144–152. (36) Stoll, S., and Schweiger, A. (2006) EasySpin, a comprehensive software package for spectral simulation and analysis in EPR. J. Magn. Reson. 178, 42–55. (37) Dyadkin, V., Pattison, P., Dmitriev, V., and Chernyshov, D. (2016) A new multipurpose diffractometer PILATUS@SNBL. J. Synchrotron Radiat. 23, 825–829. (38) Anandakrishnan, R., Aguilar, B., and Onufriev, A. V. (2012) H++ 3.0: Automating pK prediction and the preparation of biomolecular structures for atomistic molecular modeling and simulations. Nucleic Acids Res. 40, W537–W541. (39) Frisch, M. J., Trucks, G. W., Schlegel, H. B., Scuseria, G. E., Robb, M. A., Cheeseman, J. R., Scalmani, G., Barone, V., Mennucci, B., Petersson, G. A., Nakatsuji, H., Caricato, M., Li, X., Hratchian, H. P., Izmaylov, A. F., Bloino, J., Zheng, G., Sonnenberg, J. L., Hada, M., Ehara, M., Toyota, K., Fukuda, R., Hasegawa, J., Ishida, M., Nakajima, T., Honda, Y., Kitao, O., Nakai, H., Vreven, T., Montgomery Jr., J. A., Peralta, J. E., Ogliaro, F., Bearpark, M., Heyd, J. J., Brothers, E., Kudin, K. N., Staroverov, V. N., Kobayashi, R., Normand, J., Raghavachari, K., Rendell, A., Burant, J. C., Iyengar, S. S., Tomasi, J., Cossi, M., Rega, N., Millam, J. M., Klene, M., Knox, J. E., Cross, J. B., Bakken, V., Adamo, C., Jaramillo, J., Gomperts, R., Stratmann, R. E., Yazyev, O., Austin, A. J.,


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Cammi, R., Pomelli, C., Ochterski, J. W., Martin, R. L., Morokuma, K., Zakrzewski, V. G., Voth, G. A., Salvador, P., Dannenberg, J. J., Dapprich, S., Daniels, A. D., Farkas, Ö., Foresman, J. B., Ortiz, J. V, Cioslowski, J., and Fox, D. J. (2009) Gaussian 09, Revision D.01. Gaussian Inc. Gaussian, Inc., Wallingford CT. (40) D.A. Case, D.S. Cerutti, T.E. Cheatham, III, T.A. Darden, R.E. Duke, T.J. Giese, H. Gohlke, A.W. Goetz, D. Greene, N. Homeyer, S. Izadi, A. Kovalenko, T.S. Lee, S. LeGrand, P. Li, C. Lin, J. Liu, T. Luchko, R. Luo, D. Mermelstein, K.M. Merz, G. Monard, H., D. M. Y. and P. A. K. (2017) AMBER 2017, University of California, San Francisco. (41) Seminario, J. M. (1996) Calculation of intramolecular force fields from second-derivative tensors. Int. J. Quantum Chem. 60, 1271–1277. (42) Beckham, G. T., and Crowley, M. F. (2011) Examination of the α-chitin structure and decrystallization thermodynamics at the nanoscale. J. Phys. Chem. B 115, 4516–4522. (43) Sikorski, P., Hori, R., and Wada, M. (2009) Revisit of α-chitin crystal structure using high resolution X-ray diffraction data. Biomacromolecules 10, 1100–1105. (44) Sawada, D., Nishiyama, Y., Langan, P., Forsyth, V. T., Kimura, S., and Wada, M. (2012) Water in crystalline fibers of dihydrate β-chitin results in unexpected absence of intramolecular hydrogen bonding. PLoS One (Gasset, M., Ed.) 7, e39376. (45) Maier, J. A., Martinez, C., Kasavajhala, K., Wickstrom, L., Hauser, K. E., and Simmerling, C. (2015) ff14SB: Improving the Accuracy of Protein Side Chain and Backbone Parameters from ff99SB. J. Chem. Theory Comput. 11, 3696–3713. (46) Kirschner, K. N., Yongye, A. B., Tschampel, S. M., González-Outeiriño, J., Daniels, C. R., Foley, B. L., and Woods, R. J. (2008) GLYCAM06: A generalizable biomolecular force field. carbohydrates. J. Comput. Chem. 29, 622–655. (47) Salomon-Ferrer, R., Götz, A. W., Poole, D., Le Grand, S., and Walker, R. C. (2013) Routine microsecond molecular dynamics simulations with AMBER on GPUs. 2. Explicit solvent particle mesh ewald. J. Chem. Theory Comput. 9, 3878–3888. (48) Roe, D. R., and Cheatham, T. E. (2013) PTRAJ and CPPTRAJ: Software for processing and analysis of molecular dynamics trajectory data. J. Chem. Theory Comput. 9, 3084–3095. (49) Hedegård, E. D., and Ryde, U. (2017) Targeting the reactive intermediate in polysaccharide monooxygenases. J. Biol. Inorg. Chem. 22, 1029–1037. (50) Bertini, L., Breglia, R., Lambrughi, M., Fantucci, P., De Gioia, L., Borsari, M., Sola, M., Bortolotti, C. A., and Bruschi, M. (2017) Catalytic Mechanism of Fungal Lytic Polysaccharide Monooxygenases Investigated by First-Principles Calculations. Inorg. Chem. acs.inorgchem.7b02005. (51) Götz, A. W., Clark, M. A., and Walker, R. C. (2014) An extensible interface for QM/MM molecular dynamics simulations with AMBER. J. Comput. Chem. 35, 95–108. (52) Neese, F. (2012) The ORCA program system. Wiley Interdiscip. Rev. Comput. Mol. Sci. 2, 73–78. (53) Becke, A. D. (1988) Density-functional exchange-energy approximation with correct asymptotic behavior. Phys. Rev. A 38, 3098–3100. (54) Perdew, J. P. (1986) Density-functional approximation for the correlation energy of the inhomogeneous electron gas. Phys. Rev. B 33, 8822–8824. (55) Bühl, M., and Kabrede, H. (2006) Geometries of transition-metal complexes from densityfunctional theory. J. Chem. Theory Comput. 2, 1282–1290. (56) Grimme, S., Antony, J., Ehrlich, S., and Krieg, H. (2010) A consistent and accurate ab initio parametrization of density functional dispersion correction (DFT-D) for the 94 elements H-Pu. J. Chem. Phys. 132, 154104. (57) Grimme, S., Ehrlich, S., and Goerigk, L. (2011) Effect of the damping function in dispersion corrected density functional theory. J. Comput. Chem. 32, 1456–1465. (58) Weigend, F., and Ahlrichs, R. (2005) Balanced basis sets of split valence, triple zeta valence and quadruple zeta valence quality for H to Rn: Design and assessment of accuracy. Phys. Chem. Chem. Phys. 7, 3297. (59) Weigend, F. (2006) Accurate Coulomb-fitting basis sets for H to Rn. Phys. Chem. Chem. Phys. 8, 1057. (60) Adamo, C., and Barone, V. (1999) Toward reliable density functional methods without adjustable parameters: The PBE0 model. J. Chem. Phys. 110, 6158–6170. (61) Sunna, A., Moracci, M., Rossi, M., and Antranikian, G. (1997) Glycosyl hydrolases from hyperthermophiles. Extremophiles 1, 2–13. (62) Kuusk, S., Bissaro, B., Kuusk, P., Forsberg, Z., Eijsink, V. G. H., Sørlie, M., and Väljamäe, P. (2017) Kinetics of H2O2-driven degradation of chitin by a bacterial lytic polysaccharide monooxygenase. J. Biol. Chem. jbc.M117.817593. (63) Ogawa, Y., Kimura, S., Wada, M., and Kuga, S. (2010) Crystal analysis and high-resolution


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imaging of microfibrillar α-chitin from Phaeocystis. J. Struct. Biol. 171, 111–116. (64) Ogawa, Y., Kimura, S., and Wada, M. (2011) Electron diffraction and high-resolution imaging on highly-crystalline β-chitin microfibril. J. Struct. Biol. 176, 83–90. (65) Suzuki, K., Suzuki, M., Taiyoji, M., Nikaidou, N., and Watanabe, T. (1998) Chitin Binding Protein (CBP21) in the Culture Supernatant of Serratia marcescens 2170. Biosci. Biotechnol. Biochem. 62, 128–135. (66) Danneels, B., Tanghe, M., Joosten, H. J., Gundinger, T., Spadiut, O., Stals, I., and Desmet, T. (2017) A quantitative indicator diagram for lytic polysaccharide monooxygenases reveals the role of aromatic surface residues in HjLPMO9A regioselectivity. PLoS One 12, e0178446. (67) Forsberg, Z., Bissaro, B., Gullesen, J., Dalhus, B., Vaaje-Kolstad, G., and Eijsink, V. G. H. (2018) Structural determinants of bacterial lytic polysaccharide monooxygenase functionality. J. Biol. Chem. 293, 1397–1412. (68) Kracher, D., Andlar, M., Furtmüller, P. G., and Ludwig, R. (2017) Active-site copper reduction promotes substrate binding of fungal lytic polysaccharide monooxygenase and reduces stability. J. Biol. Chem. 293, 1676-1687. (69) Chovancova, E., Pavelka, A., Benes, P., Strnad, O., Brezovsky, J., Kozlikova, B., Gora, A., Sustr, V., Klvana, M., Medek, P., Biedermannova, L., Sochor, J., and Damborsky, J. (2012) CAVER 3.0: A Tool for the Analysis of Transport Pathways in Dynamic Protein Structures. PLoS Comput. Biol. 8. (70) Borisova, A. S., Isaksen, T., Dimarogona, M., Kognole, A. A., Mathiesen, G., Várnai, A., Røhr, Å. K., Payne, C. M., Sørlie, M., Sandgren, M., and Eijsink, V. G. H. (2015) Structural and functional characterization of a lytic polysaccharide monooxygenase with broad substrate specificity. J. Biol. Chem. 290, 22955–22969. (71) de Visser, S. P., Quesne, M. G., Martin, B., Comba, P., and Ryde, U. (2014) Computational modelling of oxygenation processes in enzymes and biomimetic model complexes. Chem. Commun. 50, 262–282. (72) Meier, K. K., Jones, S. M., Kaper, T., Hansson, H., Koetsier, M. J., Karkehabadi, S., Solomon, E. I., Sandgren, M., and Kelemen, B. (2017) Oxygen Activation by Cu LPMOs in Recalcitrant Carbohydrate Polysaccharide Conversion to Monomer Sugars. Chem. Rev. acs.chemrev.7b00421. (73) Ye, S., and Neese, F. (2011) Nonheme oxo-iron ( IV ) intermediates form an oxyl radical upon approaching the C – H bond activation transition state. Proc. Natl. Acad. Sci. 108, 1228–1233.


1 How a Lytic Polysaccharide Monooxygenase Binds Crystalline

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