Full-Length Papers

Protein Science DOI 10.1002/pro.2489

Thermodynamic and Structural Investigation of the specific SDS binding of Humicola insolens cutinase David Kold1, Zbigniew Dauter2, Anne K. Laustsen3, Andrzej M. Brzozowski2, Johan P. Turkenburg2, Anders D. Nielsen4, Heidi Koldsø3, Evamaria Petersen5, Birgit Schiøtt3, Leonardo De Maria6, Keith S. Wilson2,*, Allan Svendsen6,* and Reinhard Wimmer1,* 1

Department of Biotechnology, Chemistry and Environmental Engineering, Aalborg University,

Sohngaardsholmsvej 49, DK-9000 Aalborg, Denmark 2

Structural Biology Laboratory, Department of Chemistry, University of York, Heslington, York

YO10 5DD, U.K. 3

iNANO and inSPIN centers, Dept. of Chemistry, Aarhus University, Langelandsgade 140, DK-

8000 Aarhus C, Denmark 4

Protein Characterization, Novo Nordisk A/S, DK-2760 Maaloev, Denmark

5

Department of Physics and Nanotechnology, Aalborg University, Skjernvej 4A, DK-9220 Aalborg

SØ, Denmark 6

Novozymes A/S, DK-2880 Bagsværd, Denmark

* Authors to whom correspondence should be addressed: E-mail: [email protected] (A.S.), [email protected] (K.W.), [email protected] (R.W.) Running Title: Interaction of Humicola insolens cutinase with SDS Total number of manuscript pages: 33 Electronic supplementary: 7 pages of supplementary material describing details on molecular modeling, molecular modeling results, and statistics on structure refinement available. This article has been accepted for publication and undergone full peer review but has not been through the copyediting, typesetting, pagination and proofreading process which may lead to differences between this version and the Version of Record. Please cite this article as doi: 10.1002/pro.2489 © 2014 The Protein Society Received: Mar 13, 2014; Revised: May 01, 2014; Accepted: May 01, 2014

Protein Science

Abstract The interaction of lipolytic enzymes with anionic surfactants is of great interest with respect to industrially produced detergents. Here, we report the interaction of cutinase from the thermophilic fungus Humicola insolens with the anionic surfactant SDS, and show the enzyme specifically binds a single SDS molecule under non-denaturing concentrations. Protein interaction with SDS was investigated by NMR, ITC and molecular dynamics simulations. The NMR resonances of the protein were assigned, with large stretches of the protein molecule not showing any detectable resonances. SDS is shown to specifically interact with the loops surrounding the catalytic triad with medium affinity (Ka ≈ 105 M-1). The mode of binding is closely similar to that seen previously for binding of amphiphilic molecules and substrate analogues to cutinases, and hence SDS acts as a substrate mimic. In addition, the structure of the enzyme has been solved by X-ray crystallography in its apo form and after co-crystallization with diethyl p-nitrophenyl phosphate (DNPP) leading to a complex with monoethylphosphate (MEP) esterified to the catalytically active serine. The enzyme has the same fold as reported for other cutinases but, unexpectedly, esterification of the active site serine is accompanied by the ethylation of the active site histidine which flips out from its usual position in the triad.

Keywords: cutinase, cutinase-detergent interactions, crystal structure, inhibitor complex

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Abbreviations AA: All-atom, AoC: Aspergillus oryzae cutinase, AOT: sodium bis(2-ethylhexyl) sulfosuccinate, cmc: critical micelle concentration, DEP: diethylphosphate, DNPP: diethyl p-nitrophenyl phosphate, EDTA: ethylene diamine tetraacetate, FsC: Fusarium solani cutinase, GcC: Glomerella cingulata cutinase, HiC: Humicola insolens cutinase, HSQC: Heteronuclear Single-Quantum Coherence, IPTG: Isopropyl β-D-1-thiogalactopyranoside, ITC: isothermal titration calorimetry, MD: molecular dynamics, MEP: monoethylphosphate, MWCO: molecular weight cut-off, PAGE: polyacrylamide gel electrophoresis, RMSD: root mean square deviation, RMSF: root mean square fluctuation, TES: N-[Tris(hydroxymethyl)methyl]-2-aminoethanesulfonic acid, SANS: small-angle neutron scattering, SDS: sodium dodecyl sulfate

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Introduction Cutinases (EC 3.1.1.74, Pfam family PF01083) are small (20-30 kDa) lipolytic enzymes capable of hydrolyzing a wide variety of esters, including triglycerides of varying size. To date, the threedimensional structure has been reported for three cutinases, from Fusarium solani (FsC),1 Glomerella cingulata (GcC),2,3 and Aspergillus oryzae (AoC).4 These enzymes are members of the α/β hydrolase superfamily with a central parallel β-sheet and surrounding α-helices, and active site consisting of a catalytic triad of a serine, histidine and aspartic acid. Cutinases show no interfacial activation and are therefore placed as intermediates between lipases and esterases rather than being categorized as classical lipases.5–7 This functional versatility has made them a good alternative to classical lipases for industrial use in detergent products.8 In order to function in detergent mixtures, an enzyme needs to retain its catalytic activity under the conditions used for washing, i.e. temperatures up to 60°C, and the presence of anionic and non-ionic surfactants. It was therefore of interest to characterize a cutinase (HiC) from a thermophilic organism, the fungus Humicola insolens, capable of growing at temperatures as high as 58°C.9 HiC has high sequence identities to other cutinases: 56% to FsC, 59% to GcC and 50% to AoC. Both HiC10 and FsC11 were shown to be highly sensitive towards the anionic surfactants sodium dodecyl sulfate (SDS) and dioctyl sodium sulfosuccinate (AOT). Ionic surfactants are known to denature proteins by binding to neighboring charged and hydrophobic side chains. At surfactant concentrations below the critical micelle concentration (cmc), many proteins bind monomeric detergent molecules in a specific way, while at concentrations exceeding the cmc, the protein unfolds and binds a large number of surfactant molecules. The mechanism by which this happens depends on the ionic strength of the solution, its pH, and on the absolute concentration of surfactant, since this has an influence on the shape of the

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micellar aggregates.12,13 However, even in the unfolded state, considerable residual α-helical structure can be retained.14 SDS is the most frequently used anionic surfactant. The mechanism of protein denaturation by SDS depends on the secondary structure of the protein, with all-β-sheet proteins being less prone to denaturation than α-helical or mixed α/β proteins.15,16 This is ascribed to the large number of local contacts in α-helical proteins allowing their helices to remain intact even after the overall tertiary structure has been lost, with the helical segments only starting to unfold at very high SDS concentrations. This process was recently described in detail for the interaction of SDS with myoglobin.17 The interaction of HiC with SDS has been intensively studied by a variety of methods. A combination of small-angle neutron scattering (SANS) and isothermal titration calorimetric (ITC)18 together with a combined fluorescence and ITC study19 showed that the interaction can be described by four different phases. (1). At very low concentrations, SDS binds stoichometrically to HiC, causing only minute conformational changes. (2). As the SDS concentration is increased, HiC unfolds, leading to increased exposure of hydrophobic surfaces, and to the formation of dimers and possibly larger oligomers of HiC. (3). Further increasing the SDS concentration causes the oligomers to break up leading to micelle-like aggregates of SDS bound to the unfolded HiC. (4). Further increase of the concentration leads to saturation of HiC with SDS. Unlike most proteins, HiC does not show a marked increase in molecular dimensions upon full denaturation by SDS, and SDS saturation occurs at unusually low levels (ca. 0.5 g SDS per gram of HiC). However, it is not resistant against denaturation by the detergent. Further details of its thermal stability in SDS solutions were reported and the available experimental data were combined to create a state diagram of the HiC/SDS mixture.20 One conclusion of the studies conducted by Nielsen et al.18–20 was that the protein specifically binds SDS at very low (1-2) molar ratios of SDS/HiC.

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Here we report on thermodynamic and structural studies of the specific binding of SDS to HiC. We carried out ITC measurements at very low molar ratios, determined the three-dimensional structure by X-ray crystallography, investigated the location of conformational changes upon binding of SDS by NMR spectroscopy, and probed the interaction using molecular dynamics (MD) simulations.

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Results Data deposition The X-ray data and coordinates have been deposited in the PDB, with deposition codes 4OYY (native) and 4OYL (complex). The NMR assignment data have been deposited to the BioMagRes Bank (accession no. 19770). In the PDB deposition for the MEP complex, the MEP-modifed serine has been named MIR in the protein sequence, and the ethylated histidine HIE, following PDB standard nomenclature.

ITC Titration of HiC with SDS revealed specific binding of one surfactant molecule per protein molecule (Figure 1). The binding affinity (K) of ~105 M-1 and the binding enthalpy (∆H) of -18.2 kJ mol-1 were estimated using a single set of identical sites binding model.

Denaturation of HiC measured with Fluorescence Spectroscopy The measured fluorescence signal was plotted as a function of the SDS concentration and the parameters of Equation 1 were fitted to the data points, Figure 2. At [HiC]=80µM, the fluorescence signal remains unaltered upon addition of SDS up to [SDS]≈150µM, whereafter the fluorescence signal increases.

The fold of the apo-HiC structure In both the apo-enzyme and the monoethylphosphate (MEP) complex (Figure 3), there is no electron density for the first two amino acids in the expected sequence, which may indicate truncation of the polypeptide chain. There is no well-defined density for residues beyond R193, with a clear break suggesting truncation of the C terminus as well. There is good density for the

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main chain of all residues in the range 3-193. In brief, HiC folds into a single domain, and is an αβ protein with a central 5-stranded β-sheet surrounded by helices on both sides, a typical α/β hydrolase fold.21 The central sheet is well conserved throughout the lipase-cutinase family but with significant variations in the helix and loop structures. The fold is very similar to those of the other known cutinases, FsC (PDB 1cus), GcC (PDB 3dcn) and AoC (PDB 3gbs). Superposition using the SSM algorithm22 within the CCP4mg molecular graphics program,23 Figure 3(A), gives r.m.s. deviations of 0.80, 0.81 and 1.02 Å based on 189, 185 and 184 equivalent Cα positions in the three structures respectively. The differences are mainly restricted to small variations in the surface loops and the chain termini. However, while the catalytic triad (S105, H173 and D160 in HiC) superposes closely on those of FsC and AoC, the GcG structure shows substantial changes from the other three enzymes in the loop carrying the catalytic histidine, Figure 3(B). The histidine has flipped out by about 11 Å in apo-GcC whereas it forms part of a functional triad constellation in the other three cutinases. A proposal for the significance of this histidine loop movement in relation to the mechanism has been extensively discussed.3

The HiC-MEP complex structure The overall fold is essentially the same as that of the apo-enzyme, differences being restricted to the active site region. The active S105 was modeled with a covalent link to the MEP phosphorous atom, which has three other bonds to oxygen atoms. In all three independent molecules, there is clear density for an ethyl group on only one of the phosphate oxygen atoms. Both of the “free” P-O oxygens form good H-bonding networks in the crystal, Figure 3(C). The second ethyl group appears to have been transferred to the active site histidine which is alkylated in the complex, see below. This is supported by the fact that the histidine is unmodified in the apo-enzyme.

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There are striking differences in the active site between the inhibited forms of HiC and FsC. On inhibition by DEP (PDB code 2cut), FsC shows no significant conformational change compared to the native enzyme.5 In contrast, in the HiC MEP complex, the active site is completely disrupted, with the side chains considerably displaced from one another, Figure 3(C). H173 lies about 9 Å from S105 with the space between them occupied by Y104 whose Oγ atom lies roughly half way between these two catalytic residues. H173 also lies ~8.4 Å from D160 and 6.75 Å from the Oγ atom of S105, with the space vacated by the movement of H173 being occupied by three highly ordered water molecules. There are many additional well-ordered water molecules in the active site region involved in an intensive H-bond network. Indeed the whole loop containing H173 shows substantial displacement compared to the FsC-DEP structure. This displacement also causes a smaller movement of the α-helix next to H173. The distance between the Cα atom of the histidine in these two structures is 5.2 Å and between the Cγ atoms of the side chain is 6.8 Å, reflecting the fact that H173 in HiC-MEP is completely flipped away from the catalytic serine. This disruption of the triad is reminiscent of His-loop position observed in apo-GcC. In the HiC-MEP complex, S105 and D160 retain the positions seen in the apo-enzyme and are unperturbed by the swinging out of the histidine loop. In the initial refinements of the structure, there was a significant electron density feature close to the Nε2 of H173 in all three independent molecules, Figure 3(D). This was too close to the Nε2 atom and of too low a peak height to be a metal atom, and the residue has been modeled as an ethylated histidine. Alkyl phosphates are known to be powerful and indeed toxic alkylating agents 24, and one of the ethyl groups of a diethyl-phosphate attacking Ser105 is proposed to ethylate the adjacent H173 causing it to be displaced from the triad. It is curious that such behavior is only apparent in HiC and not in FsC, but this may reflect the slightly different inhibitors used in those studies. The

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flipping out of the histidine is similar to that seen in GcC, where it has been proposed to be relevant to the mechanism.

NMR studies in solution Backbone Assignment: Backbone assignment was conducted with an array of different spectra made with a 13C/15N labelled HiC sample. The [15N-1H] HSQC spectrum showed too few signals, in fact, only 146 of the 184 expected HN/N signals were observed (84%). The amino acids that did not yield signals are: 18, 28, 29, 30, 31, 40-43, 61, 72, 73, 95, 98, 106, 110, 134-136, 145, 148 and the stretches from amino acid 156-163 and 168-176. These are mainly located in loops (Figure 4A). Weak or vanishing NMR signals are usually ascribed to conformational exchange. The HSQC spectrum of HiC featured several extremely weak signals that might arise from the undetected amino acids, however, due to the total lack of correlation peaks in the triple-resonance spectra, no assignment could be made. An HSQC spectrum with assignment is shown in Supporting Figure S1. Changing the temperature of the solution and/or addition of SDS did not change this behavior. It is noteworthy that the 168-173 loop carries the catalytic H173 while 156-163 carries the catalytic D160. Molecular Mobility: The T1/T2 of HiC was determined to be 43.5±9.3. This corresponds to a correlation time of overall tumbling τM of 21±2.5 ns.25 NMR-Titration with SDS: The overall chemical shift changes ∆δ[Hz] calculated by Equation 2 are shown in Figure 5. During the titration the migration of seven assigned HN/N peaks (amino acids 66, 68, 71, 133, 164, 165, 166 and 167) exceeded 100 Hz (calculated by Eq. 2). An additional nine amino acids (26, 35, 74, 75, 111, 130, 139, 155 and 184) shifted by 60-100 Hz. The HN/N peaks of amino acids 27, 34, 36, 37, 39, 64, 66, 67, 69, 121, 138 and 179 vanished during titration with SDS (in that case, their ∆δ was calculated from the last observable position during titration).

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Furthermore, amino acids 9, 13, 39, 40, 51, 73 and 123 featured significantly diminished HSQC signal intensities during titration with SDS. The position of the migrating and disappearing HN peaks in the 3D structure can be seen in Figure 4B-D, from which it is clear that migrating residues are mainly situated on both sides of the crevice opposite the active site. Furthermore the residues with disappearing or highly mobile HN/N peaks are mainly near the flexible loops. Chemical shift changes were converted into fraction of HiC bound to SDS, and thereby [HiC-SDS]. Titration data are shown in Figure 5 and 6. Supporting Figure S2 shows the HSQC spectra in the absence and presence of SDS, respectively, with the behavior of selected signals throughout the titration. After approximately 0.3 mM SDS, little change in the HSQC spectra could be observed, indicative of full saturation of the capacity of HiC for specific SDS-binding. Eq. 3 was used to obtain an association constant of 0.65*105 ± 0.26*105 M-1 for the interaction of HiC with SDS.

MD studies Apo-HiC MD simulations: Three AA-MD simulations were carried out to examine the stability of HiC under the conditions applied in the MD simulations. The overall stability was assessed by calculating the root mean square deviation (RMSD) of the Cα atoms and the root mean square fluctuation (RMSF) for each residue (Supporting Figure S3). It is evident that HiC reaches a stable conformation in all simulations, since the RMSD-curves reach plateaus at 1-1.5 Å. It can therefore be assumed that the enzyme is stable during the three 40 ns simulations. In the RMSF plot it is evident that the overall fluctuations are low and that the largest movements occur in the loop regions. Apo-HiC with four SDS molecules: The 12 simulations will be referred to as H4S(1.1)-H4S(1.4), H4S(2.1)-H4S(2.4), and H4S(3.1)-H4S(3.4). The first index indicates which of the three starting structures was used while the second indicates the repeat number. The stability of HiC was once

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again determined using RMSD and RMSF plots (Supporting Figure S4). It is clear that HiC reaches an equilibrium structure in H4S(1.1)-H4S(1.4), H4S(2.1)-H4S(2.4) and H4S(3.1)-H4S(3.3) since the RMSD-curves quickly reach plateaus at 1-1.5 Å and the enzyme can therefore be expected to be stable during the 40 ns simulation with four SDS molecules present. While simulation H4S(3.4) deviates from the rest and reaches a plateau around 2 Å, it is still reasonable to consider the enzyme in this computation as being stable during the simulation. In the RMSF plots the overall fluctuations are again low with the largest fluctuations restricted to the loop regions, especially in the two loops situated on either side of the active site crevice. The trajectories from each set of four simulations were combined using the CATDCD plugin for VMD1.9.126 to yield three merged trajectories. To determine where SDS comes into contact with HiC, volumetric maps were created using the built-in VolMap tool in VMD1.9.1. The maps display the volume occupied by the four SDS molecules during each of the merged trajectories, yielding three maps (Figure 7A-C). In all three, SDS is seen to occupy space close to the catalytic triad, especially in H4S(1.1)-H4S(1.4) and H4S(2.1)H4S(2.4) where the volumetric surface is mainly situated around the crevice. In H4S(3.1)-H4S(3.4) the volumetric surface is spread out on the surface of HiC, however there are additional contacts near the crevice. We defined SDS to be in contact with an amino acid if any of SDS atoms were within 4 Å of any amino acid atom, since this distance entails for hydrophobic interactions of the SDS tails as well as salt bridge interactions to the sulphate head-group.27 This was calculated for every snapshot collected for each of the 12 simulations and the results averaged. From Figure 8 it is evident that SDS is in contact with the two loops (amino acids 64-75 and 160-176) situated on either side of the catalytic triad. However, SDS also comes into contact with amino acids in a third loop (135-145) which also lies close to the catalytic triad, Figure 7D. In H4S(3.1)-H4S(3.4), SDS is seen to have the most contacts with this third lid (Figure 7C).

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Discussion Structure of HiC The overall fold of HiC is very similar to those of FsC, GcC and AoC previously deposited in the PDB as would be expected from the sequence similarity. Comparison of the active sites of the four apo-enzymes reveals that HiC, FsC and AoC are very similar, with catalytic triads correctly positioned for catalysis and small differences restricted to the loops and chain termini. These three contrast with apo-GcC where the loop carrying the catalytic histidine takes a quite different conformation causing the histidine to swing out from its expected position in the triad, reported to reflect importance of this movement in the catalytic cycle.3 In the HiC complex with MEP covalently bound to S105, the density maps in addition clearly indicate H173 to have been ethylated, presumably a side product from the reaction with DNPP. More importantly, the loop carrying the ethylated H173 has swung out to break the catalytic triad and place it on the surface of the protein in a conformation reminiscent of that seen in apo-GcC – but with a somewhat different final position. In most of the complexes formed by reactions of lipases with DNPP and similar reagents, the phosphate-serine moiety is reported to be dialkylated, for example the Humicola lanuginosa lipase DEP complex (PDB code 4tgl)28 and the FsC complex with di-isopropylphosphate (PDB code 1xzk).29 The transfer of the ethyl group of DNPP has only been previously been reported three times to our knowledge, firstly in the structure of aged tabun-inhibited murine acetylcholinesterase (PDB code 2c0p),30 secondly in the esterase EstA from the cold-adapted bacterium Pseudoalteromonas sp. 643A (PDB code 3hp4)31 which has a classic lipase fold and thirdly in a deposited but unpublished structure of FsC (PDB code 3qpa). In these structures the catalytic triad is correctly formed. The reason for the transfer of the ethyl moiety to the histidine in HiC and the flipping of this residue remains unclear.

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In the [15N-1H] HSQC, several signals of HiC which encompass the loop regions surrounding the active site were missing. Missing signals are usually explained by slow conformational exchange on a time scale comparable to the difference in resonance frequencies between the different conformations, i.e. on the millisecond time scale. A similar phenomenon was observed for the structurally similar Pseudomonas mendocina lipase that also exhibited missing signals around the active site at 303K.32 However, for that enzyme signals could be observed at 313K. The NMR spectrum of FsC did not show this feature, as all signals were visible.33 This means that the intramolecular mobility of the loops is altered in HiC as compared to FsC. The reason for this might be the origin that HiC is from a thermophilic organism, as such proteins generally possess less structural flexibility at room temperature than proteins from mesophiles.34 The B-factors from the X-ray structure were compared with calculated B-factors from the MD simulations (Supporting information Figure S3). While in both cases increased B-factors are seen close to the N-terminus, in the X-ray structure the B-factors are low in the loops around the active site whereas those in the MD simulations show raised levels in this area. Hence the MD simulations support the NMR studies, which states that movement is seen in these loop regions. The correlation time of HiC of 21 ns is only slightly larger than that found for FsC, where two independent studies reported values of 10.6 ns35 and 18.3 ns.36

Interaction with SDS ITC measurements clearly show a 1:1 binding of HiC to SDS at low molar ratios with an enthalpy of binding of 18.2 kJ mol-1 and a Ka of 105 M-1, which can be considered as strong binding. This is corroborated by the NMR titration, which yields a Ka value of 0.65*105 M-1. Fluorescence measurements show no changes at the low molar ratios / concentrations in question which indicates that the overall structure does not change as a result of this binding event. Only at higher SDS

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concentrations does the fluorescence signal of HiC change due to denaturation. HiC and FsC possess only a single tryptophan, located 22 Å away from the active site. The tryptophan fluorescence of both FsC37 and HiC19 has been shown to be highly quenched in the native protein by a nearby disulfide bond, and loss of tertiary structure therefore results in an increase of the fluorescence signal. During the SDS titration very little difference could be seen in the fluorescence signal until a SDS concentration of approximately 150 µM, where the specific HiC binding site of SDS is already saturated, as seen from ITC data, Fig. 1. The concentration of HiC was 80 µM, and the ligand binding of SDS therefore occurs without significant increase in fluorescence, i.e. loss of tertiary structure. This is further supported by the NMR data that show no sign of denaturation up to a molar ratio SDS/HiC of three (at a slightly higher HiC concentration, 110 µM). No NOEs from HiC to SDS could be detected that would allow a structure determination of the complex. However, most residues showing marked changes in chemical shift upon binding of SDS are located in the loops surrounding the active site. A smaller group of amino acid showing changes in shifts is situated below the active site residues. Chemical shifts are very sensitive to changes in the local environment, and while they do not report on the exact nature of these changes, these changes clearly indicate that binding of SDS takes place in the vicinity of the active site altering the conformation of the loops 66-75 and 164-167, as seen in the MD simulations. While NMR signals for amino acids 156-163 and 168-176 could not be observed, taking into account that all amino acids in between these unassigned stretches exhibited strong changes in chemical shifts, it is reasonable to assume that the undetectable amino acids may also be affected. It is observed from the MD simulations that amino acids 160-163 and 168-176 are in fact in contact with SDS (Figure 8). The location of the interaction site is very similar to the interaction site of FsC with detergent-like molecules, the zwitterionic dihexanoylphosphatidylcholine (C6PC)38 and non-ionic 1-(16-DOXYL-

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stearoyl)glycerol.36 For both, chemical shift changes mostly affected the loops around the active site and the bottom of the active site crevice, and in a previous NMR-study with a covalently attached inhibitor,39 these loops showed the largest chemical shift changes. An X-ray structure of cutinase with bound inhibitor O-methyl-undecylphosphonate also shows the hydrocarbon chain situated between these loops29 (an overlay is shown in Fig. 4E). It is believed that these loops are “minilids” that rearrange upon interaction with the substrate.39 In true lipases, these lids are larger and effectively block the access of the aqueous phase to the active site. Upon interaction with a hydrophobic lipid phase the lid would open generating the active form of the enzyme. Cutinases lack these lids, and are thus also active against water-soluble substrates. The interaction of the lipase from Thermomyces lanuginosus with non-ionic and zwitterionic detergents led to its activation.40–42 Although no structural data are available for this interaction, activation must imply that a true lipase also interacts with detergents in the loop/lid region by altering the conformation of the lids. Taking the available data together, cutinases appear to interact with amphiphilic molecules in the vicinity of the active site. Amphiphilic molecules can be substrates or substrate mimics like detergents, whence their mode of interaction with cutinases and lipases are similar. SDS has previously been shown to inhibit the enzymatic action of FsC,43,44 but not at concentrations (molar ratios) as low as we investigate here. It is therefore difficult to assess whether the specific binding described here inhibits the enzymatic activity of a cutinase, as the concentration of the substrate used for activity assays is much higher than the concentration of the detergent at the low molar ratios. Thus, the specific binding of the detergent is just outcompeted by an excess of substrate under these conditions. This will also be the case in industrial applications of cutinases.

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Materials and Methods Expression and purification of HiC for ITC, fluorescence and crystallography: Recombinant HiC was expressed in Aspergillus oryzae and purified to >95% determined by SDS/PAGE, as described in10, at Novozymes A/S, Bagsvaerd, Denmark. Cloning, expression and purification of HiC for NMR spectroscopy; An Escherichia coli expression plasmid was constructed by fusing the nucleotide sequences coding for a variant of E. coli alkaline phosphatase signal peptide (PhoA) from plasmid pFCEX1

45

with the cDNA coding for Humicola

insolens mutant NL-83 (Novozymes A/S). In this construct, the C-terminal residue of the signal sequence is linked to residue Q1 of HiC. Utilizing the T7/lac promoter system of the pET-11a (Novagen) vector, the HiC gene was expressed in E. coli BL21(DE3) cells, by induction with IPTG at a final concentration of 1 mM. 15N labeled HiC was produced using M9 minimal medium, with ammonium-15N chloride

15

N>98% (NH4Cl) (Cambridge Isotope Laboratories) as sole ammonium

source. Double labelled (15N/13C) HiC was produced using M9 minimal medium, with ammonium15

N chloride

15

N>98% (NH4Cl) (Cambridge Isotope Laboratories) as sole ammonium source, and

U-13C6-D-Glucose- 13C>99 (C6H12O6) (Cambridge Isotope Laboratories) as sole carbon source. The presence of the PhoA signal peptide resulted in the secretion of the protein to the periplasmic space. The cells were harvested by centrifugation at 8000g for 10 min at 277K, the cell pellet resuspended in ice cold 5 mM CaCl2 and left to incubate on ice for 5 min. The cells were re-centrifuged, the supernatant removed and the pellet resuspended and incubated for 15 minutes in ice-cold 50 mM TES buffer with 20 mM sucrose. The suspension was centrifuged and sterile filtered. Subsequently, the buffer was changed to 50 mM sodium acetate at pH 4.6 using an Amicon stirred cell with an ultrafiltration membrane with a MWCO of 10 kDa. Further purification was performed by ionexchange chromatography using an Äkta explorer with a XK16/20 column packed with SP

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Sepharose (Amersham Biosciences) with an NaCl gradient from 0-1 M. The final purification was achieved by size-exclusion chromatography using an Äkta purifier equipped with a K9/60 column packed with Superdex 75 prep grade. ITC: HiC was extensively dialyzed at 5°C against 50 mM glycine, 2 mM EDTA, pH 10.0. All ITC measurements reported here were made in this buffer, using the following chemicals: glycine, (>99.7%, Merck, Darmstadt, Germany), ethylenediaminetetraacetic acid, EDTA (>99%, Merck, Darmstadt, Germany) and sodium dodecyl sulfate, SDS (>99%, Fluka, Buchs, Switzerland) and a MCS-ITC (MicroCal Inc., Northampton, MA, USA) isothermal titration calorimeter. The reference cell was filled with water. Sample cells were loaded with a 74 µM solution of cutinase in 50 mM glycine, 2 mM EDTA, pH 10.0. The cell solution was titrated with 55 aliquots of 5 µL (1st injection only 1 µL) of 1.5 mM SDS in the same buffer solution at 38°C. The resulting heat signals were integrated using the Origin software supplied by MicroCal Inc. Binding parameters were estimated by fitting the integrated heat signals using a single set of identical sites binding model. SDS induced denaturation of HiC measured with fluorescence spectroscopy: 8.5·10-2 mM HiC was incubated in concentrations of SDS ranging from 0 mM to 1 mM for a minimum of 30 min at room temperature. Excitation was performed at λex= 295 nm and emission measured at λem= 335 nm. The [D]50% was determined by fitting the parameters from Equation 1 to the measured data.

F=

α D + β D [SDS ] + (α N + β N [SDS ]) ⋅ 10 m 1 + 10m D − N ( [SDS ]− [D ]

50%

D− N

( [ SDS ]− [ D ]50% )

)

(1)

where F is the measured fluorescence signal, αN is the signal with no SDS added and βN=dαN/d[SDS] prior to transition. αD and βD are the corresponding values after the transition region. mD-N is the sensitivity of the equilibrium constant.

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Crystallization and X-ray data collection: Extensive screening for apo-HiC gave only small, needle-like crystals. However, addition of organic solutions to the crystallization medium gave usable crystals under very narrow conditions: HiC 30 mg mL-1, 0.1 M Tris/HCl buffer pH 8.5, 50 mM lysine, 100 mg mL-1 (app. 8.6%) isopropanol, 50 mg mL-1 (app. 4.3 %) dioxane, PEG 4K 22% v/v of 50% w/v stock solutions. X-ray data to a resolution of 3.0 Å were collected on a Raxis II imaging plate scanner mounted on a Rigaku rotating anode. The space group was P212121 with cell dimensions a=125.7, b=127.3, c=134.2 Å. Initial estimates indicated there were probably eight molecules per asymmetric unit with a Matthews coefficient Vm of 3.14 and a solvent content ~61%. Native HiC was inhibited with diethyl-p-nitrophenyl-phosphate (DNPP: purchased from Sigma) with the aim of preparing a complex with diethyl-phosphate (DEP) by methods previously established for the Humicola lanuginosa28,46 and Rhizomucor miehei47 lipases. The enzyme was washed and concentrated to 30 mg mL-1 and subjected to crystallization screening. Good diffracting crystals were obtained but showed a high propensity to be twinned. The optimum crystallization conditions were protein concentration 30 mg mL-1, 0.1 M Tris/HCl pH 8.5, 50 mM lysine, PEG MME 2K 11% v/v of 50% w/v stock solutions or PEG MME 550 16% v/v of 50 % w/v stock solutions. X-ray data were collected at the EMBL beam line BW7B, DESY, Hamburg. The space group was P21 with cell dimensions a=71.63, b=66.40, c=71.98 Å, β=119°. The data were processed with DENZO48 with an Rmerge of 5.8% to a resolution of 2.0 Å. There are three molecules in the asymmetric unit, with a Vm of 2.3 Da Å-3 and a solvent content of ~47%. Structure Determination: HiC-MEP complex: All computations were performed with the CCP4

49

suite of programs unless otherwise stated . The structure was solved by molecular replacement with AMoRe50 using FsC as the search model (PDB code 1cus)51 and refined to an R factor of 0.143 and Rfree of 0.184 at 2.05 Å resolution. There are three molecules in the asymmetric unit related by a

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three-fold non-crystallographic symmetry axis. Because of the excellent electron density there was no need to impose non-crystallographic constraints and the three molecules were refined independently. The final model consists of residues 3-193 of the main chain of all three enzyme molecules, each with monoethylphosphate bound and His173 ethylated at the Nε2 position, together with a total of 337 water molecules. The side chain of R51A and the ends of R141A, N15B, R141B and R51C have no visible electron density and have been assigned zero occupancy: the last letter in each refers to the chain number. These are all arginine residues pointing into the solvent region. The contacts between the three independent molecules are mainly stabilized by hydrogen bonds and salt bridges. Structure Determination of apo-HiC: The structure was solved using the HiC-MEP complex as search model. The crystal class was P222, but the diffraction pattern displayed a very high degree of translational pseudosymmetry with three quarters of the reflections much weaker than the rest. The strong reflections fulfilled the condition h+k/2=2n with an average value of normalized structure amplitudes of 1.46, whereas for the rest of the reflections the value of was 0.86. The unit cell corresponding to the strong reflections was C-centered with its b cell dimension halved. This clearly indicated the presence of a 21 screw axis parallel to the c-direction, but due to the weakness of reflections with h,k=2n+1 it was not possible to deduce if comparable screw axes were also present along the a and b directions of the primitive lattice. The most plausible content of the C2221 asymmetric unit was three molecules of molecular weight 19.9 kDa (this corresponds to the sequence seen in the crystal, not the full sequence) with a solvent content of 45%, corresponding to 12 molecules in the primitive cell. The structure in the pseudo-cell C2221 was solved with AMoRe using the refined structure of HiCMEP as the search model, leading to clear identification of three independent molecules of HiC. The structure in the true primitive cell was solved in a somewhat non-standard way as follows. The

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transition from the C2221 cell to the primitive cell could involve four possible space groups: P2221, P21221, P22121, and P212121, since the presence or absence of screw axes along a and b was not known. In addition, the symmetry equivalent origin shift by a half-cell along the b direction in the C2221 lattice becomes non-equivalent in the primitive cell with the b-cell dimension doubled. Assemblies of 12 molecules according to the C2221 symmetry were prepared and translated in agreement with the appropriate origin definition for each of the eight possible cases. These assemblies were submitted to rigid body fitting with AMoRe without prior execution of rotation and translation searches, using all data including the subset of weak intensities. One of the eight models, corresponding to space group P212121, gave a significantly higher correlation coefficient (76% against ~65%) and lower R factor (39% against ~45%) than the rest. As for the DEP complex, the protein chain was modelled for resides 3-193. The structure was refined using REFMAC 52 to an R factor of 0.172 with an Rfree of 0.193 at 3.0 Å resolution using strict non-crystallographic symmetry averaging. Due to the limited resolution, only a very small number of water molecules were built into the model. At the end of the refinement, there was a residual feature in the active site at the 5σ level. This could not be related to any of the components in the buffers used for crystallization. While it appeared to be compatible with a moiety such as a linear thiocyanate, we have elected to model it as a set of waters in the deposited coordinates: however, these refine to B values of 2.0, the lower limit permitted by REFMAC. It is not possible to identify the moiety with confidence at this point as these data were recorded over fifteen years ago. This does not impact significantly on the rest of the structure and the conclusions drawn. A summary of the data processing and the refinement is given in Supplementary Table S1. NMR Assignment: backbone assignment was conducted using a

15

N/13C doubly-labelled HiC

sample in 95% H2O and 5% D2O (Cambridge Isotope Laboratories) with 20 mM phosphate buffer (KH2PO4/Na2HPO4) pH 6.5 and 2 mM sodium azide (NaN3). All measurements were conducted at

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298.1K using BRUKER NMR spectrometers operating at a magnetic field strength of 14.1T with either a triple resonance (H/C/N) triple-axis gradient room-temperature-probe or a TCI z-gradient cryoprobe. The following spectra were recorded: HSQC, HNCA, HNCO, HCACO, CBCA(CO)NH, CBCANH, HNHA, HBHA(CBCACO)NH, and a 3D-15N-edited NOESY with a mixing time of 50 ms as well as a 3D-15N-edited TOCSY with a mixing time of 20 ms using Bruker standard pulse programs. Data were processed using XWIN-NMR 3.6 and TopSpin 1.3 and analyzed using the CARA software package.53 The MARS54 and AutoLink55 programs for automated assignment were used. NMR Relaxation Rates: Relaxation (T1 and T2) measurements of 15N were obtained by exponential fitting of the peak intensities in 15N-HSQC spectra acquired with different relaxation delays.25,56 NMR-titration with SDS: To investigate the ligand binding of SDS by HiC a sample of 0.5 mL of 0.11 mM 15N labelled HiC in 95% H2O and 5% v/v D2O with 20 mM phosphate buffer pH 6.5 and 2 mM sodium azide, was titrated with 5 mM SDS dissolved in the same buffer. Aliquots of 3.7µl SDS solution were added and incubated for a minimum of 30 min at room temperature, whereafter [15N-1H]-HSQC spectra were recorded. Addition of one aliquot corresponds to adding 0.33 molar equivalents of SDS per HiC. All measurements were conducted with a sample temperature of 25°C. When HiC binds SDS the chemical shift of the affected HN/N pairs will change. As the exchange rate is fast on the chemical shift time scale, the recorded signal will represent the population weighted average of the chemical shifts of the free and bound form of HiC. This will result in what appears to be a migration of the HN/N peaks from one shift without SDS toward another where full saturation is achieved. The chemical shifts of HN/N cross peaks can be directly converted into a fraction of protein bound to SDS. Chemical shift changes in the 1H and combined into one ∆δ value as shown in Equation 2:

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∆δ = ∆δ H2 + ∆δ N2

(2)

where ∆δH and ∆δN are the changes in chemical shifts of the 1H and

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N nucleus, respectively,

expressed in Hz. When HiC is saturated with SDS the chemical shifts remain unaltered until denaturation of the protein begins. The denaturation results in a significant change in chemical shift for most HN pairs, and can therefore easily be detected. The association constant for the formation of a 1:1 complex is defined by Equation 3: Ka =

[HiC − SDS ] = [HiC − SDS ] [HiC ] ⋅ [SDS ] ([HiC ]0 − [HiC − SDS ]) ⋅ ([SDS ]0 − [HiC − SDS ])

(3)

Solving the quadratic equation yields Equation 4, an expression for the complex concentration as a function of the SDS concentration and the association constant:

[HiC − SDS ] = ½ [HiC ]0 + [SDS ]0 + 

2

1

 ±  [HiC ] + [SDS ] + 1  − 4[HiC ] [SDS ]  0 0 0 0 K a  K a  

(4)

[HiC-SDS] for each titration point of [SDS]0 was obtained from the chemical shift of the moving signals, Ka was obtained from fitting Equation 4 to the data. MD simulations: Two different apo-HiC setups were studied utilizing the all-atom MD (AA-MD) technique and each were run for several repeats. The first setup consists of a water box containing only apo-HiC and the second setup contained water, apo-HiC and four SDS molecules. Three runs of apo-HiC in a water box were simulated for 40 ns and the structure in the last frame of the three runs served as starting structures for the second set of simulations with SDS molecules. Each starting structure was supplied with four SDS molecules 20 Å away from the protein surface along the x- and y-axes. The SDS molecules were placed in the same location in each simulation setup, yielding only a difference within the starting structures of the enzyme. The three new setups were

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each repeated four times, resulting in 12 simulations of the second setup hereby obtaining better sampling and more reliable statistics for analysis. The total simulation time was 600 ns for both setups. See supporting information for details.

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Electronic Supplementary Materials Details on molecular modelling, molecular modelling results, figures of NMR spectra and statistics on structure refinement are available as supplementary material.

Acknowledgements We thank Miroslawa Dauter for growing the crystals of HiC, Finn L. Aachmann for recording NMR spectra, Shamkant A. Patkar, Laurent Duroux, Kristian R. Poulsen for advice on protein purification and Kim Hansen for help with fermentations. DK and RW acknowledge financial support by the Obel Foundation and the Oticon Foundation. AKL, HK and BS acknowledge financial support from the Danish Council for Independent Research | Technology and Production Sciences, the Danish National Research Foundation (DNRF59) and the Danish Centre for Scientific Computing. We thank EMBL Hamburg for providing access to beamline BW7B.

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References 1. Longhi S, Mannesse M, Verheij HM, De Haas GH, Egmond M, Knoops-Mouthuy E, Cambillau C (1997) Crystal structure of cutinase covalently inhibited by a triglyceride analogue. Protein Sci 6:275–286. 2. Nyon MP, Rice DW, Berrisford JM, Huang H, Moir AJG, Craven CJ, Nathan S, Mahadi NM, Abu Bakar FD (2008) Crystallization and preliminary X-ray analysis of recombinant Glomerella cingulata cutinase. Acta Cryst F64:504–508. 3. Nyon MP, Rice DW, Berrisford JM, Hounslow AM, Moir AJG, Huang H, Nathan S, Mahadi NM, Bakar FDA, Craven CJ (2009) Catalysis by Glomerella cingulata cutinase requires conformational cycling between the active and inactive states of its catalytic triad. J Mol Biol 385:226–235. 4. Liu Z, Gosser Y, Baker PJ, Ravee Y, Lu Z, Alemu G, Li H, Butterfoss GL, Kong X-P, Gross R, et al. (2009) Structural and functional studies of Aspergillus oryzae cutinase: enhanced thermostability and hydrolytic activity of synthetic ester and polyester degradation. J Am Chem Soc 131:15711–15716. 5. Martinez C, Nicolas A, van Tilbeurgh H, Egloff MP, Cudrey C, Verger R, Cambillau C (1994) Cutinase, a lipolytic enzyme with a preformed oxyanion hole. Biochemistry 33:83–89. 6. Schrag JD, Cygler M (1997) Lipases and alpha/beta hydrolase fold. Methods Enzymol 284:85– 107. 7. Chahinian H, Nini L, Boitard E, Dubès J-P, Comeau L-C, Sarda L (2002) Distinction between esterases and lipases: a kinetic study with vinyl esters and TAG. Lipids 37:653–662. 8. Flipsen JA., Appel AC., van der Hijden HTW., Verrips CT (1998) Mechanism of removal of immobilized triacylglycerol by liplytic enzymes in a sequential laundry wash process. Enzyme Microb Technol 23:274–280. 9. Maheshwari R, Bharadwaj G, Bhat MK (2000) Thermophilic fungi: their physiology and enzymes. Microbiol Mol Biol Rev 64:461–488. 10. Ternström T, Svendsen A, Akke M, Adlercreutz P (2005) Unfolding and inactivation of cutinases by AOT and guanidine hydrochloride. Biochim Biophys Acta 1748:74–83. 11. Creveld LD, Meijberg W, Berendsen HJ, Pepermans HA (2001) DSC studies of Fusarium solani pisi cutinase: consequences for stability in the presence of surfactants. Biophys Chem 92:65– 75. 12. Otzen DE (2002) Protein unfolding in detergents: effect of micelle structure, ionic strength, pH, and temperature. Biophys J 83:2219–2230.

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13. Otzen D (2011) Protein-surfactant interactions: a tale of many states. Biochim Biophys Acta 1814:562–591. 14. Takeda K, Sasaoka H, Sasa K, Hirai H, Hachiya K, Moriyama Y (1992) Soze and mobility of sodium dedecyl-sulfate bovine serum albumin complex as studied by dynamic light-scattering and electrophoretic light-scattering. J Colloid Interface Sci 154:385–392. 15. Manning M, Colon W (2004) Structural basis of protein kinetic stability: resistance to sodium dodecyl sulfate suggests a central role for rigidity and a bias toward beta-sheet structure. Biochemistry 43:11248–11254. 16. Nielsen MM, Andersen KK, Westh P, Otzen DE (2007) Unfolding of beta-sheet proteins in SDS. Biophys J 92:3674–3685. 17. Andersen KK, Westh P, Otzen DE (2008) Global study of myoglobin-surfactant interactions. Langmuir 24:399–407. 18. Nielsen AD, Arleth L, Westh P (2005) Interactions of Humicola insolens cutinase with an anionic surfactant studied by small-angle neutron scattering and isothermal titration calorimetry. Langmuir 21:4299–4307. 19. Nielsen AD, Arleth L, Westh P (2005) Analysis of protein-surfactant interactions–a titration calorimetric and fluorescence spectroscopic investigation of interactions between Humicola insolens cutinase and an anionic surfactant. Biochim Biophys Acta 1752:124–132. 20. Nielsen AD, Borch K, Westh P (2007) Thermal stability of Humicola insolens cutinase in aqueous SDS. J Phys Chem B 111:2941–2947. 21. Nardini M, Dijkstra BW (1999) Alpha/beta hydrolase fold enzymes: the family keeps growing. Curr Opin Struct Biol 9:732–737. 22. Krissinel E, Henrick K (2004) Secondary-structure matching (SSM), a new tool for fast protein structure alignment in three dimensions. Acta Cryst D60:2256–2268. 23. McNicholas S, Potterton E, Wilson KS, Noble MEM (2011) Presenting your structures: the CCP4mg molecular-graphics software. Acta Cryst D67:386–394. 24. Schubert C, Fiedler F (1994) Formation of pi, tau-dimethylhistidine on alkylation of trypsin with active-site-directed sulfonic acid methyl esters. J Enzym Inhib 8:173–185. 25. Kay LE, Torchia DA, Bax A (1989) Backbone dynamics of proteins as studied by 15N inverse detected heteronuclear NMR spectroscopy: application to staphylococcal nuclease. Biochemistry 28:8972–8979. 26. Humphrey W, Dalke A, Schulten K (1996) VMD: visual molecular dynamics. J Mol Graph 14:33–38, 27–28. 27. Barlow DJ, Thornton JM (1983) Ion-pairs in proteins. J Mol Biol 168:867–885.

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28. Lawson DM, Brzozowski AM, Rety S, Verma C, Dodson GG (1994) Probing the nature of substrate binding in Humicola lanuginosa lipase through X-ray crystallography and intuitive modelling. Protein Eng 7:543–550. 29. Longhi S, Nicolas A, Creveld L, Egmond M, Verrips CT, de Vlieg J, Martinez C, Cambillau C (1996) Dynamics of Fusarium solani cutinase investigated through structural comparison among different crystal forms of its variants. Proteins 26:442–458. 30. Ekström F, Akfur C, Tunemalm A-K, Lundberg S (2006) Structural changes of phenylalanine 338 and histidine 447 revealed by the crystal structures of tabun-inhibited murine acetylcholinesterase. Biochemistry 45:74–81. 31. Brzuszkiewicz A, Nowak E, Dauter Z, Dauter M, Cieśliński H, Długołecka A, Kur J (2009) Structure of EstA esterase from psychrotrophic Pseudoalteromonas sp. 643A covalently inhibited by monoethylphosphonate. Acta Cryst F65:862–865. 32. Sibille N, Favier A, Azuaga AI, Ganshaw G, Bott R, Bonvin AM, Boelens R, van Nuland NA (2006) Comparative NMR study on the impact of point mutations on protein stability of Pseudomonas mendocina lipase. Protein Sci 15:1915–1927. 33. Prompers JJ, Groenewegen A, Van Schaik RC, Pepermans HA, Hilbers CW (1997) 1H, 13C, and 15N resonance assignments of Fusarium solani pisi cutinase and preliminary features of the structure in solution. Protein Sci 6:2375–2384. 34. Likhtenshtein GI, Febbraio F, Nucci R (2000) Intramolecular dynamics and conformational transition in proteins studied by biophysical labelling methods. Common and specific features of proteins from thermophylic micro-organisms. Spectrochim Acta A Mol Biomol Spectrosc 56A:2011–2031. 35. Prompers JJ, Groenewegen A, Hilbers CW, Pepermans HA (1999) Backbone dynamics of Fusarium solani pisi cutinase probed by nuclear magnetic resonance: the lack of interfacial activation revisited. Biochemistry 38:5315–5327. 36. Poulsen KR, Sorensen TK, Duroux L, Petersen EI, Petersen SB, Wimmer R (2006) The interaction of Fusarium solani pisi cutinase with long chain spin label esters. Biochemistry 45:9163–9171. 37. Prompers JJ, Hilbers CW, Pepermans HA (1999) Tryptophan mediated photoreduction of disulfide bond causes unusual fluorescence behaviour of Fusarium solani pisi cutinase. FEBS Lett 456:409–416. 38. Sehgal P, Bang Nielsen S, Pedersen S, Wimmer R, Otzen DE (2007) Modulation of cutinase stability and structure by phospholipid detergents. Biochim Biophys Acta 1774:1544–1554. 39. Prompers JJ, van Noorloos B, Mannesse ML, Groenewegen A, Egmond MR, Verheij HM, Hilbers CW, Pepermans HA (1999) NMR studies of Fusarium solani pisi cutinase in complex with phosphonate inhibitors. Biochemistry 38:5982–5994.

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40. Mogensen JE, Sehgal P, Otzen DE (2005) Activation, inhibition, and destabilization of Thermomyces lanuginosus lipase by detergents. Biochemistry 44:1719–1730. 41. Brzozowski AM, Savage H, Verma CS, Turkenburg JP, Lawson DM, Svendsen A, Patkar S (2000) Structural origins of the interfacial activation in Thermomyces (Humicola) lanuginosa lipase. Biochemistry 39:15071–15082. 42. Yapoudjian S, Ivanova MG, Brzozowski AM, Patkar SA, Vind J, Svendsen A, Verger R (2002) Binding of Thermomyces (Humicola) lanuginosa lipase to the mixed micelles of cis-parinaric acid/NaTDC. Eur J Biochem 269:1613–1621. 43. Pocalyko DJ, Tallman M (1998) Effects of amphipaths on the activity and stability of Fusarium solani pisi cutinase. Enzyme Microb Technol 22:647–651. 44. Kolattukudy PE Cutinases from Fungi and Pollen. In: Borgstrom B, Brockman H, editors. Amsterdam: Elsevier; 1984. pp. 470–504. 45. Petersen SB, Jonson PH, Fojan P, Petersen EI, Petersen MT, Hansen S, Ishak RJ, Hough E (1998) Protein engineering the surface of enzymes. J Biotechnol 66:11–26. 46. Brzozowski AM (1993) Crystallization of a Humicola lanuginosa lipase-inhibitor complex with the use of polyethylene glycol monomethyl ether. Acta Cryst D49:352–354. 47. Derewenda U, Brzozowski AM, Lawson DM, Derewenda ZS (1992) Catalysis at the interface: the anatomy of a conformational change in a triglyceride lipase. Biochemistry 31:1532–1541. 48. Otwinowski Z, Minor W (1997) Processing of X-ray diffraction data collected in oscillation mode. Methods Enzymol 276:307–326. 49. Winn MD, Ballard CC, Cowtan KD, Dodson EJ, Emsley P, Evans PR, Keegan RM, Krissinel EB, Leslie AGW, McCoy A, et al. (2011) Overview of the CCP4 suite and current developments. Acta Cryst D67:235–242. 50. Navaza J (1994) AMoRe: an automated package for molecular replacement. Acta Cryst A50:157–163. 51. Martinez C, De Geus P, Lauwereys M, Matthyssens G, Cambillau C (1992) Fusarium solani cutinase is a lipolytic enzyme with a catalytic serine accessible to solvent. Nature 356:615–618. 52. Murshudov GN, Skubák P, Lebedev AA, Pannu NS, Steiner RA, Nicholls RA, Winn MD, Long F, Vagin AA (2011) REFMAC5 for the refinement of macromolecular crystal structures. Acta Cryst D67:355–367. 53. Keller R The Computer Aided Resonance Assignment Tutorial. 1. ed. Goldau (Switzerland): CANTINA Verlag; 2004. 54. Jung YS, Zweckstetter M (2004) Mars – robust automatic backbone assignment of proteins. J Biomol NMR 30:11–23.

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55. Masse JE, Keller R (2005) AutoLink: automated sequential resonance assignment of biopolymers from NMR data by relative-hypothesis-prioritization-based simulated logic. J Magn Reson 174:133–151. 56. Farrow NA, Muhandiram R, Singer AU, Pascal SM, Kay CM, Gish G, Shoelson SE, Pawson T, Forman-Kay JD, Kay LE (1994) Backbone dynamics of a free and phosphopeptide-complexed Src homology 2 domain studied by 15N NMR relaxation. Biochemistry 33:5984–6003.

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Figure Legends Figure 1: ITC data for the binding of SDS to HiC at 38°C in 50 mM Glycine, 2 mM EDTA, pH 10 buffer. The top panel shows the raw data (heat flow versus time) of 55 injections of 5 µl SDS aliquots. In the lower panel the peaks have been integrated to yield the enthalpy change per mole of SDS added. The solid line is a fit to the data using a single set of identical binding sites model.

Figure 2: Denaturation of HiC (80µM) by aqueous SDS measured by fluorescence spectroscopy (λexcitation= 295 nm and λemission= 335 nm). The relative fluorescence is plotted as a function of the SDS concentration, by fitting the parameters of Eq. 1 to the data points. [D]50% was determined to be 0.45mM and mD-N 2.8mM-1.

Figure 3: Views apo-cutinases, HiC (this work, grey), FsC (PDB 1cus, blue), apo-GcC (PDB 3dcn, coral) and AoC (PDB 3gbs, yellow). (a) Stereo view of the overall fold of the four enzymes shown as worms. The catalytic triad of HiC is shown as spheres. On the top left the His-loop can be seen for GcC to have moved substantially compared to the other three proteins. http://imolecules3d.wiley.com:8080/imolecules3d/review/URoXnSlgu7oWOvTt9jrThMkktSwDU5 D2FBhT6vu1EM02vFGegVk2fZ5LUvL4N0VT730/1419 (b) Stereo view showing a close up of the active site. The chain is only shown for HiC and GcC. The side chains are shown as cylinders for the four proteins, with HiC carbons colored grey, AoC and FsC in usual atom colors, and GcC coral. The triads of the first three superimpose closely, the His-loop of GcC has flipped disrupting the triad. (c) Stereo views of the active site of HiC (grey), apo-GcC (coral) and the HiC-MEP complex (pale blue), with the overall folds shown as worms.

The residues of the catalytic triads are shown as cylinders, with carbons for apo-HiC grey, HiCMEP pale blue and GcC coral. The HiC histidine has flipped away from the triad in the MEP

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complex, albeit to a different position to that of H204 in apo-GcC. (d) The electron density at the 1σ level around the MEP-Ser105 with an ethyl moiety attached to His173. The protein chain is shown in ribbon format.

Figure 4: Influence of SDS titration on NMR spectra of HiC. A: HiC-DEP with active-site (amino acids 105, 160 and 173) shown as spheres and the unassigned non-proline residues (18, 28, 29, 30, 31, 40-43, 61, 72, 73, 95, 98, 106, 110, 134-136, 145, 148, 156-163 and 168-176) in red. Residues showing the biggest chemical shift changes in HSQC spectra (Coral: ∆δ>125 Hz; residues 66, 71, 166 and 167; Yellow: 125 Hz >∆δ>100 Hz; residues 68, 164, 165 and 133), residues whose peak in HSQC spectra disappeared at some point during titration with SDS (Magenta: 27, 34, 36, 37, 39, 64, 66, 67, 69, 121, 138 and 179) and residues whose signal intensity in HSQC spectra is reduced during titration (Black: 9, 13, 39, 40, 51, 73 and 123). http://imolecules3d.wiley.com:8080/imolecules3d/review/URoXnSlgu7oWOvTt9jrThMkktSwDU5 D2FBhT6vu1EM02vFGegVk2fZ5LUvL4N0VT730/1420 B: surface of HiC. Red: residues affected by titration with SDS. Blue: unassigned non-proline residues. C: PDB structure 1XZM

29

of FsC (backbone in pale blue) with O-methyl-

undecylphosphonate esterified to active site serine, superimposed with HiC in gold, with the side chains close to the FsC ligand shown as sticks.

Figure 5: Chemical shift perturbation of HiC (calculated according to Equation 2) upon addition of 0.3 mM SDS plotted against the amino acid number.

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Figure 6: Concentration of HiC-SDS complex as a function of the total SDS concentration as calculated from the chemical shift titration data of the six strongest shifting amino acids. The solid line represents the fit for Ka=0.65*105 M-1.

Figure 7: Location of SDS molecules during MD: the secondary structure of apo-HiC is shown in light green and the catalytic triad in pink. The space, which the four SDS molecules occupy most frequently, are depicted as volumetric isosurfaces in transparent gray (isovalue 0.07). A: The volumetric surface from the combined trajectory of H4S(1.1)-H4S(1.4). B: A: The volumetric surface from the combined trajectory of H4S(2.1)-H4S(2.4). C: A: The volumetric surface from the combined trajectory of H4S(3.1)-H4S(3.4). D: The amino acids 135-145 are shown as orange van der Waals spheres.

Figure 8: The percentage of snapshots from all 12 MD simulations where a SDS molecule is in contact with the different amino acids of apo-HiC.

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95x155mm (300 x 300 DPI)

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84x72mm (300 x 300 DPI)

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140x245mm (300 x 300 DPI)

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166x218mm (300 x 300 DPI)

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160x91mm (300 x 300 DPI)

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151x106mm (300 x 300 DPI)

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127x134mm (300 x 300 DPI)

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165x112mm (300 x 300 DPI)

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Supporting information Molecular modelling Apo-HiC: The three runs of apo-HiC in a water box, each originated from the crystal structure determined for the apo-form, deposited in the PDB with code 4oyy. There are twelve independent HiC monomers in the unit cell, all closely similar in fold and chain A was chosen as the starting structure for these simulations. The model lacked the side chains of two arginine residues (Arg-51 and Arg-141), which were built on using PRIME1. The N-terminal and C-terminal ends were acetylated and Nmethylated, respectively. Protonation states were examined by PROPKA3.12. Protonation states of the two histidine residues were chosen based on surrounding residues, giving rise to His49 being modeled as the Nε-tautomer while His173, which is part of the catalytic triad, is modeled as the Nδ-tautomer. The setups were solvated by the TIP3P water model3 and two chloride ions were added to neutralize the systems. Three replicates of the system, containing 19968 atoms, were simulated for 40 ns and are referred to as HiC(1), HiC(2), and HiC(3). Apo-HiC with four SDS molecules. Three new systems containing one apo-HiC from either HiC(1), HiC(2) or HiC(3) were supplied with four SDS molecules placed 20 Å away from the protein surface along the x- and y-axes, preventing interactions between HiC and SDS at the intuition of the simulation. The parameters for SDS can be found in the CHARMM27 lipid parameter and topology file4. These three systems were run for 40 ns in four replicas, yielding 12 simulations with a total simulation time of 480 ns. These simulations will be referred to as H4S(1.1)-H4S(1.4), H4S(2.1)-H4S(2.4) and H4S(3.1)-H4S(3.4). Simulations. Each system was initially minimized by the conjugated gradient method for 15,000 steps followed by simulation using the NPT ensemble at 310 K with 1 fs

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time step. All simulations were run in NAMD2.85, utilizing the CHARMM276 force field with CMAP corrections7,8. Constant pressure at 1 atm was maintained by employing the Langevin piston method9,10 with a damping coefficient of 0.5 ps-1 and a piston period of 100 fs. PME was used to treat full electrostatics11 while van der Waals interactions were truncated at a cut-off distance of 12 Å using a switching function from 10 Å. Analysis. The RMSD and RMSF calculations as well as the volumetric maps, created from the build in VolMap tool, were calculated utilizing VMD1.9.112. VolMap divides the simulation box into a grid, where every grid point is separated by 1 Å. Each grid point is set to either 0 or 1 in each frame depending on the location of SDS. If one of the atoms in one of the SDS molecules overlays a grid point it is given the value 1 else 0. The numbers from each frame is summed together and divided by the number of frames giving a fraction of time where the grid point is occupied. This fraction is plotted in figure 9 as isosurfaces.

Supporting figure legends: Figure S1: [15N-1H]-HSQC of HiC with assignments: Peaks in red are negative (folded) peaks, presumably Arg Hε/Nε peaks (HiC contains 12 Arg residues). Peaks denoted with “*” are unassigned: they come from side chain Asn and Gln amide peaks (HiC contains 13 Asn and 7 Gln residues each potentially showing two side chain peaks) or from impurities or from some of the unassigned residues of HiC (none of them showed useful correlations in the triple resonance spectra that would allow an assignment). Figure S2: Overlay of [15N-1H]-HSQC of HiC without SDS (red) and with SDS (blue). Inserts illustrate chemical shift changes of selected residues during titration:

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SDS concentrations: black: 0 μM, blue: 37 μM, green: 74 μM, red: 111 μM, orange: 148 μM, brown: 185 μM. Figure S3: A: RMSD of HiC(1)-HiC(3). B: RMSF of apo-HiC. Figure S4: A and B: RMSD and RMSF of H4S(1.1)-H4S(1.4), respectively. C and D: RMSD and RMSF of H4S(2.1)-H4S(2.4), respectively. E and F: RMSD and RMSF of H4S(3.1)-H4S(3.4), respectively.

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Table S1. Data and refinement statistics Data set PDB Code Wavelength (Å) Space group Cell parameters (Å,º)

Native 4oyy 1.54 P212121 a = 125.55 b = 127.0 c = 134.51

MEP complex 4oyl 0.89 P21 a = 71.63 b = 66.40 c = 71.98 β = 119.3 99,112 36,132 20-2.05 (2.09–2.05) 0.058 (0.115) 97.4 (89.3) 2.8 (1.9) 10.5 (2.6) 2.33 3 0.143 0.184 4599 337 19.9 19.2

Total reflections 195,605 Unique reflections 43,331 Resolution (Å) 20-3.0 (3.16-3.0) Rmerge 0.118 (0.324) Completeness (%) 99.5 (97.5) Redundancy 4.5 (3.6) 5.9 (2.3) VM (Å3/Da) 3.14 Mol. per asymmetric unit 12 Rcryst 0.172 Rfree 0.193 No. of non-hydrogen atoms 16996 No. of water molecules 106 Mean B value (Å2) 28.1 Mean B value protein (Å2) 28.1 RMS deviation from ideality Bonds (Å) 0.011 0.017 Angles (º) 1.5 1.7 Values in parenthesis correspond to the high resolution shell.

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References

1. Schrödinger (2010) New York. 2. Li H, Robertson AD, Jensen JH (2005) Very fast empirical prediction and rationalization of protein pKa values. Proteins 61:704–21. 3. Jorgensen WL, Chandrasekhar J, Madura JD, Impey RW, Klein ML (1983) Comparison of simple potential functions for simulating liquid water. J Chem Phys 79:926. 4. Feller SE, MacKerell AD (2000) An Improved Empirical Potential Energy Function for Molecular Simulations of Phospholipids. J Phys Chem B 104:7510– 7515. 5. Phillips JC, Braun R, Wang W, Gumbart J, Tajkhorshid E, Villa E, Chipot C, Skeel RD, Kalé L, Schulten K (2005) Scalable molecular dynamics with NAMD. J Comput Chem 26:1781–802. 6. MacKerell, AD, Bashford D, Dunbrack, RL, Evanseck JD, Field MJ, Fischer S, Gao J, Guo H, Ha S, Joseph-McCarthy D, et al. (1998) All-Atom Empirical Potential for Molecular Modeling and Dynamics Studies of Proteins. J Phys Chem B 102:3586–3616. 7. MacKerell AD, Feig M, Brooks CL (2004) Improved treatment of the protein backbone in empirical force fields. J Am Chem Soc 126:698–9. 8. Mackerell AD, Feig M, Brooks CL (2004) Extending the treatment of backbone energetics in protein force fields: limitations of gas-phase quantum mechanics in reproducing protein conformational distributions in molecular dynamics simulations. J Comput Chem 25:1400–15. 9. Martyna GJ, Tobias DJ, Klein ML (1994) Constant pressure molecular dynamics algorithms. J Chem Phys 101:4177. 10. Feller SE, Zhang Y, Pastor RW, Brooks BR (1995) Constant pressure molecular dynamics simulation: The Langevin piston method. J Chem Phys 103:4613. 11. Darden T, York D, Pedersen L (1993) Particle mesh Ewald: An Nlog(N) method for Ewald sums in large systems. J Chem Phys 98:10089. 12. Humphrey W, Dalke A, Schulten K (1996) VMD: Visual molecular dynamics. J Mol Graph 14:33–38.

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Figure S1: [15N-1H]-HSQC of HiC with assignments: Peaks in red are negative (folded) peaks, presumably Arg Hε/Nε peaks (HiC contains 12 Arg residues). Peaks denoted with “*” are unassigned: they come from side chain Asn and Gln amide peaks (HiC contains 13 Asn and 7 Gln residues each potentially showing two side chain peaks) or from impurities or from some of the unassigned residues of HiC (none of them showed useful correlations in the triple resonance spectra that would allow an assignment). 78x56mm (300 x 300 DPI)

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Figure S2: Overlay of [15N-1H]-HSQC of HiC without SDS (red) and with SDS (blue). Inserts illustrate chemical shift changes of selected residues during titration: SDS concentrations: black: 0 µM, blue: 37 µM, green: 74 µM, red: 111 µM, orange: 148 µM, brown: 185 µM. 76x55mm (300 x 300 DPI)

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Figure S3: A: RMSD of HiC(1)-HiC(3). B: RMSF of apo-HiC. 259x103mm (300 x 300 DPI)

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Figure S4: A and B: RMSD and RMSF of H4S(1.1)-H4S(1.4), respectively. C and D: RMSD and RMSF of H4S(2.1)-H4S(2.4), respectively. E and F: RMSD and RMSF of H4S(3.1)-H4S(3.4), respectively. 163x194mm (300 x 300 DPI)

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