Study of the Structural and Dynamic Effects in the FimH Adhesin upon

Article pubs.acs.org/jmc

Study of the Structural and Dynamic Effects in the FimH Adhesin upon α‑D‑Heptyl Mannose Binding Sophie Vanwetswinkel,†,‡ Alexander N. Volkov,*,†,‡ Yann G. J. Sterckx,†,‡ Abel Garcia-Pino,†,‡ Lieven Buts,†,‡ Wim F. Vranken,†,‡ Julie Bouckaert,‡,¶ René Roy,§ Lode Wyns,†,‡ and Nico A. J. van Nuland*,†,‡ †

Jean Jeener NMR Centre, Structural Biology Brussels, Vrije Universiteit Brussel, Pleinlaan 2, 1050 Brussels, Belgium Molecular Recognition Unit, Structural Biology Research Center, VIB, Pleinlaan 2, 1050 Brussels, Belgium § Department of Chemistry, Université du Québec à Montréal, P.O. Box 8888, Succ. Centre-Ville, Montréal (QC), Canada H3C 3P8 ‡

S Supporting Information *

ABSTRACT: Uropathogenic Escherichia coli cause urinary tract infections by adhering to mannosylated receptors on the human urothelium via the carbohydrate-binding domain of the FimH adhesin (FimHL). Numerous α-D-mannopyranosides, including α-D-heptyl mannose (HM), inhibit this process by interacting with FimHL. To establish the molecular basis of the high-affinity HM binding, we solved the solution structure of the apo form and the crystal structure of the FimHL−HM complex. NMR relaxation analysis revealed that protein dynamics were not affected by the sugar binding, yet HM addition promoted protein dimerization, which was further confirmed by small-angle X-ray scattering. Finally, to address the role of Y48, part of the “tyrosine gate” believed to govern the affinity and specificity of mannoside binding, we characterized the FimHL Y48A mutant, whose conformational, dynamical, and HM binding properties were found to be very similar to those of the wild-type protein.

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INTRODUCTION Located at the tips of the type-1 pili of uropathogenic Escherichia coli (UPEC) strains, the FimH adhesin is the most prevalent causative agent of human urinary tract infections (UTIs). Bacterial invasion is effectively mediated by the adhesion of FimH to mannosylated receptors on the urinary epithelium through its amino-terminal, carbohydratebinding domain, referred to as lectin domain (FimHL). The carboxy-terminal pilin domain of FimH (FimHP) anchors the whole protein to the pili rod on the bacterial surface. In the native context of the fimbriae, both domains are interdependent, and the sugar affinity is allosterically regulated through a molecular mechanism established in an elegant study by LeTrong and colleagues.1 Briefly, FimHP makes internal contacts with FimHL causing the loosening of the mannosebinding pocket and thereby maintaining FimH in a low-affinity state. Upon separation of the domains through sugar interaction and/or tensile forces, the FimHL conformation rearranges toward a high-affinity state featuring a tighter binding site. Because of their frequency of occurrence, UTIs represent an important public health issue. Women are particularly affected, as almost half of them will experience at least one infection at some point in their lives. Moreover, despite the current availability of antibiotic treatments, UTIs recur two or more © 2014 American Chemical Society

times within months of a primary infection in 20−30% of the patients and evolve into a chronic form in about 5% of the cases.2 Combined with the threat of increasing antibiotic resistance among bacterial strains, this context has been fueling the research toward the discovery of efficient FimH antagonists with therapeutical value. Most of the reported studies aiming at designing and optimizing new drugs have been performed on the isolated FimHL domain displaying the high-affinity conformation.2−6 Indeed, the latter is small and easy to produce as opposed to the full-length protein, which is prone to aggregation due to the incomplete Ig-fold of FimHP lacking a βstrand donated by the FimG adaptor subunit in the mature fimbriae.1 Since the discovery that aryl α-D-mannosides were potent inhibitors of FimH-mediated bacterial adhesion,7 a lot of effort has been directed toward the development of new synthetic derivatives with improved antiadhesive properties. An important step in this direction was made in a study reporting the crystal structure of FimHL bound to butyl mannose and demonstrating that alkyl mannosides act as high-affinity (nanomolar) FimH ligands,8 which established the structural basis of sugar binding. The subsequent crystal structure of Received: October 28, 2013 Published: January 29, 2014 1416

dx.doi.org/10.1021/jm401666c | J. Med. Chem. 2014, 57, 1416−1427

Journal of Medicinal Chemistry

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The ensemble of the 25 water-refined lowest-energy FimHL structures (Figure 1A) is in excellent agreement with the

FimHL in complex with oligomannose-3, a physiological ligand mimic, confirmed that the binding was mostly driven by a single sugar ring embedded in a deep, polar monosaccharide binding pocket making the most of the protein contacts.9 This work thereby validated the earlier idea that monomannosides were sufficient for sugar recognition and tight binding and could act as potent competitive inhibitors of the natural ligand.10,11 Since then, synthetic monomannosides carrying various aglycone chains have been reported and their binding to FimHL has been characterized. Several crystal structures of FimHL−monomannoside complexes are available,2,4 illustrating that, as in the case of butyl mannose, the aglycone interacts via van der Waals contacts and/or aromatic stacking with a hydrophobic collar surrounding the binding pocket. More specifically, the interaction is mediated by the residues Tyr48, Tyr137, and Ile52, forming the so-called “tyrosine gate”. In a work combining structural and thermodynamic studies, Wellens et al. suggested that changes in the orientation and dynamics of these residues extending from the binding pocket, in particular the Tyr48, could govern the binding affinity and specificity of synthetic antagonists.2 To address the role of the tyrosine gate in more detail, we have studied FimHL by NMR spectroscopy and obtained the solution structure of the apo form. We investigated the structural and dynamics effects of heptyl mannose (HM) binding in solution by isothermal titration calorimetry (ITC), nuclear magnetic resonance (NMR) spectroscopy, and smallangle X-ray scattering (SAXS). Moreover, we solved the crystal structure of the FimHL−HM complex, for which there were no atomic details available so far. In parallel, the Y48A FimHL mutant was prepared and its binding properties and dynamics were compared with that of the wild-type (wt) protein.

Figure 1. Solution structure of the apo wt FimHL. (A) Ensemble of the 25 lowest energy structures represented as ribbons. (B) Overlay of the cartoon representation of the model closest to the mean (in blue) with the crystal structure of the FimHL−oligomannose-3 complex (PDB 2VCO; the protein is in gray and oligommanose-3 in orange). The tyrosine 48 (in open conformation) is shown in sticks. The inset shows the sugar binding site residues Y48, I52, and Y137 forming the “tyrosine gate”. Molecular graphics were generated with PyMOL15 and Chimera.16

experimental data showing no significant distance violations. The satisfactory red-orange-green (ROG) CING validation score14 and Ramachandran statistics (Table S1, Supporting Information) further illustrate the good quality of the solution structure. The atomic coordinates were deposited to the Protein Data Bank under the accession number 3zpd. The solution structure was obtained in the absence of any ligand, whereas, to the best of our knowledge, all crystal structures reported so far for the isolated FimHL domain contain either mannose, synthetic mannosides, or ethylene glycol, mimicking the O2, O3, and O5 hydroxyls of α-Dmannopyranose,2 at the sugar binding site. Interestingly, in the solution structure of the apo FimHL, the tyrosine 48, which has been proposed to govern the sugar binding affinity and specificity,2 adopts an “open” conformation similar to the one observed in the X-ray structure of the complex with oligomannose-3,9 a natural substrate analogue (Figure 1B). Overall, the NMR structure of the apo FimHL and crystal structures of the FimHL−ligand complexes are very similar, with an average backbone root-mean-square deviation (RMSD) of 1.44 Å (Table S2, Supporting Information). Yet, while the secondary structure elements superimpose well, several loops, especially those located near the binding site and appearing well-defined in the crystal structures (Figure S2, Supporting Information), display different conformations. It is noteworthy that the NMR ensemble shows higher variability for these regions compared to the rest of the protein (Figure 1A), most likely due to smaller number of NOE distance restraints obtained for these residues (Figure S3, Supporting Information). In principle, such structural heterogeneity could arise from local loop motions; however, as discussed below, NMR relaxation analysis reveals no significant protein dynamics at

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RESULTS AND DISCUSSION Structure and Dynamics of the Apo FimHL. The good peak dispersion in the [1H,15N]-heteronuclear single-quantum coherence (HSQC) spectrum (Figure S1, Supporting Information) and high quality of the triple-resonance data enabled nearly complete assignment (backbone 96.5%, sidechain 1H 97.6%, side-chain non-1H 84.3%, BMRB entry 19066) of the apo wild-type (wt) FimHL 1H, 13C, and 15N nuclei, including all non-proline backbone amides. The NMR structure of the wt FimHL in its apo form was solved using NOE-derived distance restraints and a combination of dihedral angle restraints. All unambiguous distance restraints were generated with CYANA12,13 based on the automatic assignment of the 3D NOE cross-peaks. About 80% of the NOEs were unambiguously assigned, leading to an average of 29.2 NOE restraints per residue; the remaining incompleteness is most likely due to the few missing chemical shift assignments and a semiautomated peak-picking procedure applied to crowded spectra. In addition to the unambiguous restraints, nine distance restraints were added manually to account for the ambiguity of the Phe1 aromatic proton assignments. Indeed, these protons were tentatively assigned based on the 2D NOESY and 3D 13C-NOESY-HSQC spectra, where the ambiguity could not be resolved. In addition, the Phe1 Hδ and Hε resonances could not be identified in the 2D (HB)CB(CGCD)HD and (HB)CB(CGCDCE)HE experiments. Omitting these additional restraints during the structure calculation resulted in an unrealistic Phe1 conformation fully exposed to the solvent. 1417



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Figure 2. SAXS analysis for the apo and HM-bound FimHL. Buffer-subtracted scattering curves of (A) apo and (C) holo FimHL measured at four different concentrations: 2.0, 4.0, 6.0, and 7.5 mg/mL. The curves are shown in different shades of blue and green for apo and holo FimHL, respectively. In both cases, the curves are displaced vertically relative to each other and are plotted on an arbitrary intensity scale for clarity. The gray traces depict the experimental error. The insets show the Guinier regions of the respective scattering curves. (B) The merged scattering curve for apo FimHL (black dots, gray traces depict the experimental error) is very well described by an apo FimH monomer. The red and blue lines represent the CRYSOL fit for the apo FimHL NMR structure and the fit for the best ab initio model, respectively. An overlay of the apo FimHL NMR structure (blue ribbon) and the ab initio model (gray beads) is shown on the right. (D) The percentages of holo FimHL monomer and dimer as a function of concentration obtained by a linear combination of scattering from the dimer and monomer by OLIGOMER (χ2 in black). Given for comparison, the χ2 in red correspond to the fits with 100% of monomer. The fits of the different OLIGOMER runs are shown as red traces in C.

backbone dynamics on the ps−ns and μs−s time scales. Although the present experiments did not address the motions on the intermediate, ns−μs time scale, a time regime not easily accessible to the solution NMR spectroscopy,17 given the protein backbone rigidity on either side of this time-window, large-scale ns−μs protein dynamics appear to be rather unlikely. Being sensitive to the shape of the protein, the R2/R1 ratio can be used to determine the hydrodynamic properties of the molecule.18 The analysis of the R2/R1 data shows that the diffusion tensor of the apo wt FimHL is accurately described by an axially symmetric particle with Dpar/Dper = 1.46 ± 0.02 and a rotational correlation time tc = 11.2 ns. Though the absolute value should be taken with caution due to the self-association propensity of FimHL (see below), the latter value is in a good agreement with τc = 10.1 ns obtained from the structure-based hydrodynamic calculations (see Experimental Section). The behavior of the apo FimHL in solution was also investigated by SAXS under the conditions similar to those of the NMR experiments (Figure 2A). The final scattering curve is fitted well by the apo FimHL solution structure, and ab initio shape reconstruction adequately produces a model that matches the protein monomer (Figure 2B). Thus, the SAXS analysis confirms the validity of the apo FimHL NMR structure. Structural and Thermodynamic Characterization of the wt FimHL−HM Interaction. HM is routinely used as a

the backbone level and shows that FimHL is essentially rigid on the ps−ns and μs−ms time scales. Given that HM binding to FimHL in solution affects only the sugar binding site residues (see below), we believe that the differences observed for the apo and holo FimHL structures reflect the experimental discrepancies between the solution NMR spectroscopy and X-ray crystallography, rather than conformational changes induced by the sugar binding. Indeed, in FimHL-ligand X-ray structures, the loops surrounding the binding site are often engaged in intermolecular contacts dictated by the crystal lattice, which might explain the difference in conformations compared to the NMR structure determined in this work. Except for several isolated outliers and slightly increased R2 rates for the residues 135−141, the apo wt FimHL exhibits flat R1 and R 2 relaxation profiles (Figure S4, Supporting Information), indicating the absence of significant backbone dynamics on the ps−ns time scale. Furthermore, backbone amides of the apo wt FimHL show flat R2 relaxation dispersion profiles, implying that either the protein undergoes no conformational exchange on the μs−ms time scale or the difference in chemical shifts between the exchanging species is small.17 Finally, a single set of well-resolved peaks in the HSQC spectra signifies the presence of a single protein species on the NMR chemical shift time scale (ms−s). Taken together, the NMR relaxation data demonstrate the absence of protein 1418



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As many of the resonances in the HSQC spectra of the apo and the HM-bound wt FimHL were virtually identical, their NMR assignments could be easily transferred from the former spectrum to the latter. These were verified, and the assignment of the HSQC spectrum of the wt FimHL−HM was completed by triple-resonance NMR experiments (see Experimental Section). Significant line broadening was observed upon sugar addition, and the consequent loss of resolution resulted in a limited assignment of the theoretically observable side-chain atom resonances in the FimHL−HM complex (backbone 96.3%, side-chain 1H 85.4%, side-chain non-1H 71.8%, BMRB entry 19256). Despite this line broadening, which effectively prevented the calculation of the NMR structure of the HM-bound FimHL form, the analysis of the backbone amide chemical shift perturbations indicated that, also in solution, HM binding only affects the sugar binding site residues (Figure 4). Moreover, the

reference in drug discovery programs because it is easily accessible, is water-soluble, and remains one of the strongest monovalent FimHL binders described so far.2−6 However, in the absence of structural data, the reasons for this high affinity remained subject to speculation.2 Here, we obtained the crystal structure of the wt FimHL−HM complex at 1.6 Å resolution (Figure 3A; Table S3, Supporting Information). Clear electron

Figure 3. Cartoon representation of the crystal structure of the FimHL−HM complex. (A) Overlay of the crystal structures of FimHL bound either to HM (this work, in green) or to butyl mannose (PDB 1UWF, in gray). HM and butyl mannose are shown in black and gray, respectively; the tyrosine 48 (in closed conformation) is shown in sticks. (B) Head-to-tail dimer as observed in the crystal.

density was observed for HM, including its aglycone tail, which appeared to be well ordered. With a 0.11 Å backbone rmsd, the structure features virtually identical Cα traces compared to the butyl mannose-bound FimHL.8 As in the latter, the Tyr48 is found to be in the closed conformation. The ligands in both structures are also highly superimposable, with their alkyl moieties extending out of the mannose-binding pocket toward the hydrophobic patch formed by Tyr48, Tyr137, and Ile52. However, while this patch is only partially covered in the case of butyl mannose, the HM aglycone tail (longer by three carbon atoms) allows additional van der Waals contacts, which could account for the ∼20 fold higher binding affinity. Moreover, contrary to what was observed for the apo FimHL solution structure, the loops surrounding the sugar binding site of the HM-bound protein adopt conformations similar to the one seen in all previously crystallized FimHL complexes, which confirms that they are well-defined in X-ray structures (Figure S2, Supporting Information). In the case of our FimHL−HM complex, these loops are definitely involved in crystal contacts. Indeed, the crystal packing appears to involve the formation of head-to-tail dimers, where the sugar binding site of one molecule contacts the carboxy-terminal part of a second one (Figure 3B). Favorable protein−protein interactions are further reinforced by the HM ligand, contributing sugar−protein contacts.

Figure 4. Combined backbone amide chemical shift perturbations (Δδavg) induced upon HM binding to the wt FimHL (A) plotted for each observed backbone amide, and (B) mapped onto a surface representation of the FimHL−HM crystal structure (PDB 4lov, this work). Residues are colored by the Δδavg as defined in the ramp. Proline and unassigned residues are shown in light gray, and HM in dark gray sticks.

protein−sugar interaction was found to be in slow exchange regime on the NMR chemical shift time scale as expected from the nanomolar affinity of HM for the wt FimHL (see ITC below and ref 2). NMR exchange regime is defined by the exchange constant (kex) and the difference in the chemical shifts between the free and bound forms (Δδ) as fast (kex ≫ Δδ), intermediate (kex ≈ Δδ), or slow (kex ≪ Δδ). The latter is manifested by the presence of two sets of peaks, corresponding to free and bound forms, at nonsaturating ligand concentrations. 1419



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Figure 5. ITC binding curves. (A) Direct titration of HM into the wt FimHL and (B) reverse titration of the wt FimHL into HM. The top and bottom panels show, respectively, the raw data after the baseline correction and the integrated data corrected for the heat of dilution of the ligand. The solid line in the bottom panel is the best fit of the data to an n identical and independent site-binding model. See Table 1 for the measured thermodynamic parameters.

Table 1. ITC Thermodynamic Parameters for the Binding of HM to the FimHL WT and Y48A Mutant wt (direct) wt (reverse) Y48A (direct)

n

Kd (nM)

ΔH (kcal/mol)

-TΔS (kcal/mol)

ΔG (kcal/mol)

0.95 ± 0.01 0.96 ± 0.01 1.04 ± 0.01

17.1 ± 2.7 19.2 ± 4.6 94.3 ± 12.9

−11.59 ± 0.08 −11.65 ± 0.09 −8.99 ± 0.08

1.00 ± 0.12 1.12 ± 0.17 −0.64 ± 0.11

−10.59 ± 0.14 −10.53 ± 0.19 −9.63 ± 0.14

As expected, HM displayed a strong affinity for the FimHL (KD ∼ 17−19 nM), in good agreement with the previously reported value of ∼7 nM.2 Although the thermodynamic signature is consistent with the binding being driven by a relatively large enthalpic contribution (ΔH ∼ 11.6 kcal/mol), here we observed only a slight entropic loss (−TΔS ∼ 1 kcal/ mol vs 2.65 kcal/mol in ref 2) (Table 1). This discrepancy likely reflects differences in the (de)solvation energies of the binding partners due to the different experimental conditions used in this and earlier works. Analysis of the HM-Induced FimHL Monomer−Dimer Equilibrium in Solution. To investigate whether a monomer−dimer equilibrium exists in solution upon addition of HM to FimHL, SAXS studies were carried out. Analysis of the ln[I(q)] vs q2 plots (Figure 2C, inset) using Guinier approximation ln[I(q)] = ln[I(0)] − q2Rg2/3 allows extracting I(0) and the Rg, which are, respectively, the forward scattering intensity (related to the effective molecular weight of the scattering particle) and the apparent radius of gyration. As can be seen from the rise of I(0) and Rg with the holo FimHL concentration (the latter increases from 17.90 Å to 21.24 Å), the addition of HM to the FimHL causes an increase in particle size. A similar trend is observed in the plot of the distance distribution function, p(r) (Figure 6A), which features increased occurrences of the larger particle diameters with the rising protein concentration. Thus, unlike for the apo FimHL, the scattering curves for the holo FimHL show significant concentration dependence, suggesting possible sugar-induced protein dimerization. Indeed, in contrast to the Guinier regions

Addition of HM to the wt FimHL does not change the pattern of the R1 and R2 relaxation profiles but leads to a uniform decrease in the R1 and concomitant increase in the R2 rates for all FimHL backbone amides (Figure S4, Supporting Information). This behavior indicates an increase in the overall rotational correlation time (τc) of the system, most likely arising from partial, sugar-induced FimHL dimerization (see below). In a similar fashion, small systematic shifts of the relaxation profiles are observed at a higher concentration of the apo protein (Figure S5, Supporting Information), which suggests that FimHL shows signs of self-association also in the absence of the sugar, confirming the observations of a recent report.19 Like the apo protein, FimHL−HM shows flat R2 relaxation dispersion profiles. Overall, these findings suggest that the HM binding has virtually no effect on the protein backbone dynamics. In contrast to the apo FimHL, the R2/R1 data for the HM-bound protein could not be reliably analyzed with a simple model, most likely due to the presence of several species populated at the monomer−dimer equilibrium (see below). The binding of HM to the wt FimHL was measured by ITC in the same conditions as used for the NMR experiments. As it is practically difficult to accurately determine the concentration of the sugar stock solution, and thereby the binding stoichiometry, both direct and reverse titrations were performed (Figure 5). In both cases, the observed number of binding sites (n) was close to 1, confirming the 1:1 stoichiometry of the FimHL−HM interaction. 1420



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Figure 6. SAXS curve fitting using different shape dimers for the holo FimHL. (A) p(r) functions of apo and holo FimHL support the occurrence of a concentration-dependent dimerization in the presence of HM. (B) Different AUTODOCK solutions for the holo FimHL dimer are shown in purple, yellow, and red, respectively. The crystallographic dimer is shown in green, and the HM is depicted in a sphere representation. (C) Fits of OLIGOMER runs for the different dimers shown in B to the experimental data of the highest holo FimHL concentration (black dots; gray traces represent the experimental error). (D) Same as C for head-to-tail, tail-to-tail, and head-to-head dimers.

AUTODOCK run, different solutions with distinct shapes were selected and used for fitting the scattering curve at the highest holo FimHL concentration. As can be seen from Figure 6C, only the elongated crystallographic dimer produces a good fit compared to the other, more compact, dimeric forms. However, because of the inherently low resolution of the SAXS experiment, we cannot unambiguously define the head/tail arrangement of the protein units as the head-to-tail, head-tohead, and tail-to-tail dimers provide equally good fits of the SAXS curves (Figure 6D). Although analyzed in the context of

of the apo FimHL curves, the ones measured for the holo FimHL are not parallel (compare Figures 2A, C). If HM-induced holo FimHL dimerization is at play, it should be possible to fit the different concentration curves by the linear combinations of the scattering of a holo FimHL monomer and dimer using OLIGOMER.20 Given that no prior information on the shape of the putative holo FimHL dimer is available, and that the crystallographic head-to-tail dimer could be a crystallization artifact, different holo FimHL dimers were generated using AUTODOCK (Figure 6B).21 From the 1421



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Figure 7. NMR relaxation analysis of the wt FimHL−HM monomer−dimer equilibrium. (A) R2/R1 profiles calculated for the apo monomeric FimHL (filled circles), head-to-tail crystallographic dimer of the HM-bound FimHL (filled triangles), and the linear combination of the predicted values for the 83% monomer and 17% dimer mixture (black open circles) are compared to the experimental data (red open circles). The experiments were conducted on a 1.08 mM wt FimHL−HM sample in 20 mM sodium phosphate/100 mM NaCl pH 6.0 at 25 °C and 600 MHz. (B) Q factor as a function of the dimer population (p). See text for details.

indicate that the translational diffusion coefficient, D, of the crystallographic, HM-bound FimHL dimer is expected to be 1.41 times smaller than that of the monomeric protein. With the D = 1.27 ± 0.01 × 10−10 m2 s−1 measured by NMR diffusion-ordered spectroscopy (DOSY) for the apo FimHL, a D = 0.90 × 10−10 m2 s−1 is expected for the FimHL−HM protein dimer. Assuming an equilibrium mixture of 17% dimer and 83% monomer as established by the NMR relaxation analysis (see above), an overall D = 1.19 × 10−10 m2 s−1 is expected for the FimHL−HM system, which is indeed very close to the value (D = 1.20 ± 0.01 × 10−10 m2 s−1) obtained by DOSY on the HM-bound FimHL sample. Thus, the NMR diffusion measurements agree with the monomer−dimer equilibrium in FimHL−HM solutions as established by SAXS and NMR relaxation analyses. Study of the FimHL Y48A Mutant. In an effort to further investigate the role of the tyrosine 48, we prepared a FimHL Y48A mutant. The backbone amide resonances for both the apo (BMRB entry 19255) and HM-bound (BMRB entry 19254) proteins were assigned by comparing the [1H,15N]HSQC and 15N-NOESY-HSQC spectra with those of the wt protein. Note that, in the case of the bound form, the Asp47 and Asn96 backbone amide chemical shifts were not identified. Chemical shift perturbation analysis confirmed that the mutation did not affect the overall structure of the protein because its effects were restricted to the direct neighbors of residue 48 (Figure S8A, Supporting Information). Moreover, just as observed for the wt FimHL, the HM binding shifts were limited to the sugar recognition site (Figure S8B, Supporting Information). The affinity of HM for the Y48A mutant was assessed by ITC (Figure S9, Supporting Information). Compared to the wt protein, we observed ∼5-fold decrease in Kd, accompanied by a slight decrease of enthalpy, partly compensated by an entropy gain (Table 1). These relatively small changes in the binding constant and thermodynamic parameters do not support an active role of the residue 48 in HM recognition. Rather, they can be explained by differences in solvent reorganization upon binding, because an entropy gain is generally due to water molecules being expelled from the complex interface.22 Indeed, the replacement of the hydrophobic solvent-exposed tyrosine 48 by an alanine potentially could create an additional site occupied by ordered water molecules.

the crystallographic dimer, the NMR relaxation data do not help to resolve this ambiguity as the overall shape of the assembly, which largely determines the relaxation properties of the individual atoms, remains the same irrespective of the head/ tail orientation. Nevertheless, it is clear that the holo FimHL dimer is elongated (Figures 6C,D). Analysis of the different concentration curves using the holo FimHL monomer and the elongated (head-to-tail) dimer demonstrates that the population of the latter increases with the protein concentration. The lowest and highest holo FimHL concentrations correspond, respectively, to the monomeric protein and the equilibrium mixture of the 80% monomer and 20% dimer (Figure 2D). This HM-induced dimerization seems to be an intrinsic property of the system, as performing the experiments under different buffer conditions yields very similar results (Figures S6 and S7, Supporting Information). To complement the SAXS experiments and characterize the sugar-promoted FimHL dimerization under the conditions of the NMR experiments, we have performed the following analysis. First, the R2/R1 profiles for the backbone amides of the FimH L monomer and dimer were obtained from the corresponding atomic coordinates (filled symbols in Figure 7A). Then the sum of the population-weighted R2/R1 values of the individual species, (R2/R1)calcd, was calculated at varying dimer populations (p), and the agreement between the experimental R2/R1 and the (R2/R1)calcd was assessed at each p value by computing a Q factor (see Experimental Section). The lower the Q factor, the better the agreement between (R2/ R1)calcd and the experimental data; thus, the global minimum of the Q = f(p) function corresponds to the best solution for the p value (Figure 7B). Here, the smallest Q = 0.12 is obtained at p = 0.17, suggesting that the NMR relaxation data can be well explained by the presence of 17% protein dimer and 83% monomer in the FimHL−HM solution (Figure 7A). The p value derived from the NMR relaxation measurements (p = 0.17) is in good agreement with that obtained from the SAXS experiments conducted at the highest protein concentration (p = 0.2; see above). Thus, the present Q = f(p) analysis of the R2/ R1 data complements the SAXS findings and provides an independent means to assess the sugar-induced FimH L monomer−dimer equilibrium in solution. These results are confirmed further by NMR diffusion measurements. Structure-based hydrodynamic calculations 1422



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The periplasmic extract was loaded onto a 10 mL Source 30S column (GE Healthcare) pre-equilibrated in 20 mM sodium acetate pH 4.0. After washing with 20−30 mL of the equilibration buffer, the protein was eluted with a 100−150 mL gradient of 0−500 mM NaCl in 20 mM sodium acetate pH 4.0. The protein-containing fractions were directly neutralized with 1 M Tris-HCl pH 8.0, pooled, exchanged into the required buffer, and then concentrated to 5−20 mg/mL in a Hydrosart centrifugal device (5000 cutoff, Vivaspin). The Y48A mutation was introduced into the pET24a(+)-derived FimHL expression plasmid via whole plasmid synthesis polymerase chain reaction.25 The Y48A FimHL variant was produced and purified as described for the wt protein. As they were prepared from cultures in M9 minimal medium, all the protein samples (labeled and not) used in this study were in the apo form. The samples were found to be very stable (upon at least 6 month storage at 4 °C) at pH 6.0 and under relatively low ionic strength (20 mM sodium phosphate containing 100 mM NaCl), buffer conditions ideal for biomolecular NMR spectroscopy. Proteins were routinely stored in a cold-room or kept at −80 °C for longer-term storage. NMR Chemical Shift Assignments. NMR experiments were performed at 298 K (unless mentioned otherwise) either in-house on Varian NMR Direct-Drive Systems 600 and 800 MHz spectrometers, the latter equipped with a salt tolerant 13C-enhanced PFG-Z cold probe, or at the University of Utrecht (BIO-NMR partner, The Netherlands) on Bruker Avance II 900 MHz and Bruker Avance DRX 600 MHz spectrometers, both equipped with 5 mm TCI cryo HCN ZGRD cryogenic probes. All NMR data were processed in NMRPipe26 and analyzed in CCPN.27 A set of 2D [1H,15N]-HSQC, [1H,13C]-HSQC, 3D NOESYs [15NNOESY-HSQC, and 13C-NOESY-HSQC for aliphatics and aromatics, mixing time 100 ms] and assignment spectra [CBCA(CO)NH, HNCACB, HNCO, HBHA(CO)NH, C(CO)NH, and HCCHTOCSY28 as well as (HB)CB-(CGCD)HD and (HB)CB(CGCDCE)HE for aromatics29] was first recorded for the apo form on a sample containing 1 mM [13C, 15N] double-labeled FimHL in 20 mM sodium phosphate/100 mM NaCl pH 6.0 and 6% D2O for the lock. Then HM was added to the sample till saturation, and the same suite of experiments was acquired for the sugar-bound form. For completion, 2D NOESY spectra were also recorded for both forms using unlabeled FimHL (0.5 mM) resuspended in D2O after lyophilization of a protein solution in 20 mM sodium phosphate/100 mM NaCl pH 6.0. Samples containing 1 mM of apo and HM-bound [15N] labeled Y48A in 20 mM sodium phosphate/100 mM NaCl pH 6.0/6% D2O were used for the acquisition of [1H,15N]-HSQC and 15N-NOESY-HSQC spectra . Nearly complete H, N, and C nuclei assignment was obtained for the apo FimHL, and above 80% of these resonances were determined for the HM-bound form following a standard procedure.30 Sequential backbone assignments were obtained by connecting 13Ca and 13Cb frequencies from the HNCACB and CBCA(CO)NH spectra at the 1 H, 15N frequencies of every peak in the [1H,15N]-HSQC spectrum. Subsequently, 1Hα, 1Hβ, and side-chain 13C chemical shifts were obtained from the HBHA(CO)NH and C(CO)NH spectra, respectively. The defined 1Hα−13Cα and 1Hβ−13Cβ resonances were then used for the assignment of the 1H and missing 13C frequencies of aliphatic side chains from the HCCH-TOCSY spectrum, and the 13CO resonances were deduced from the HNCO spectrum. Aromatic 1H chemical shifts were obtained from the combined use of the (HB)CB(CGCD)HD and (HB)CB(CGCDCE)HE spectra, based on the correlation with the assigned 13Cβ resonances; the corresponding aromatic 13C frequencies were taken from the [1H,13C]-HSQC spectrum. Finally, the heavy-atom resonances of the asparagine and glutamine residues were assigned from the CBCA(CO)NH spectrum by connecting the 1H,15N sidechain amide resonances to the corresponding Cα/Cβ or Cβ/Cγ chemical shifts while the side-chain NH2 were similarly obtained from the HBHA(CO)NH spectrum and checked using the 3D 15NNOESY-HSQC spectra. For the apo and HM-bound Y48A mutant, backbone amides were assigned based on overlays and comparison of the [1H,15N]-HSQC

For both apo and HM-bound Y48A FimHL, the R1 and R2 profiles (Figure S10, Supporting Information) and the flat R2 relaxation dispersion curves are virtually identical to those of the wt protein, indicating that the introduced mutation neither alters the protein backbone dynamics nor perturbs the sugarinduced FimHL monomer−dimer equilibrium.

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CONCLUDING REMARKS We have established the molecular determinants of the highaffinity HM binding to FimHL. Overall, our data do not support an active role of Y48 in HM recognition and demonstrate its limited influence on protein backbone dynamics. From the methodological perspective, this work illustrates the importance of the combined use of several biophysical techniques (i.e., SAXS, NMR relaxation, and diffusion measurements) for comprehensive analysis of protein monomer−dimer equilibrium in solution. Our results, in particular the findings of the sugar-induced dimerization of the isolated FimHL domain, unlikely to occur in the context of the full-length protein, could benefit future studies of ligand binding to FimH and are expected to inform the ongoing drug discovery efforts.

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EXPERIMENTAL SECTION

Heptyl α-D-Mannopyranoside (HM). The sugar was synthesized at the Department of Chemistry, the University of Quebec (Montréal, Canada), through Fischer glycosidation by coupling D-mannose and 1heptanol catalyzed with camphorsulfonic acid.23 Expression and Purification of FimHL. Recombinant wt FimHL wt and its Y48A mutant were produced under the control of a T7 promoter using a construct where the endogenous FimH signal peptide was replaced with the PelB one to increase the yield of periplasmic export. Therefore, the gene encoding the residues 1 to 158 of the FimH adhesin from the uropathogenic E. coli (UPEC) strain J96 was amplified from the plasmid pMMB91,9 appended with the PelB sequence and subcloned into the pET24a(+) vector (Novagen). The protein was expressed from the resulting construct transformed into E. coli C43 (DE3) cells24 grown in M9 minimal medium (6.8 g/L Na2HPO4, 3 g/L KH2PO4, 1 g/L NaCl) containing 25 mg/L kanamycin and supplemented with 2 mM MgSO4, 0.2 mM CaCl2, trace elements (60 mg/L FeSO4·7H2O, 12 mg/L MnCl2·4H2O, 8 mg/ L CoCl2·6H2O, 7 mg/L ZnSO4·7H2O, 3 mg/L CuCl2·2H2O, 0.2 mg/ L H3BO3, and 50 mg/L EDTA), BME vitamin mix (Sigma), and 1 g/L NH4Cl plus 4 g/L D-glucose. Bacteria were grown at 37 °C till OD600 nm reached 0.6−0.8. At this point, the protein expression was induced with 1 mM IPTG, and the culture temperature was lowered to 30 °C. After overnight incubation, the cells were harvested by centrifugation, and the bacterial pellets were resuspended in 30 mM Tris-HCl pH 8.0 buffer containing 20% sucrose and stored at −80 °C. The same procedure was applied to the expression of the uniformly labeled [15N] and [13C, 15N] proteins, except that 1 g/L 15NH4Cl (CortecNet) and 2 g/L 13C6-glucose (Cambridge Isotope Laboratories) were used as the sole nitrogen and carbon sources. To purify the FimHL domain, the frozen cells were thawed, 1 mM EDTA and protease inhibitors were added (0.1 mg/mL AEBSF-HCl and 1 mg/mL leupeptin, Roche), and periplasmic extracts were prepared. For this, the thawed bacterial suspensions were cleared by centrifugation (12 000 rpm for 15 min), the supernatant (sucrose fraction) kept on ice, and the pellet resuspended in ice-cold 5 mM MgSO4. After incubation of the latter at room temperature and slight agitation for 10−15 min, the solution was spun down (20 000 rpm for 20 min), and the supernatant was pooled with the sucrose fraction to obtain the total periplasmic extract, which was dialyzed against 20 mM sodium acetate pH 4.0 at 4 °C for 3 to 4 h. The sample was harvested and cleared by centrifugation, and the pH and conductivity were checked and, if necessary, adjusted by dilution with water to pH 4.0 and ≤1 mS/cm, respectively. 1423



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spectra from the corresponding forms of the wt protein. When possible, ambiguities were resolved by examining the 3D 15N-NOESYHSQC spectra. Chemical shift perturbations of the 15N and 1H nuclei were analyzed by overlaying the [1H,15N]-HSQC spectra of the apo and holo proteins and of the wt and Y48A mutant. The combined chemical shifts perturbations (Δδavg) were derived from eq 1: Δδavg = [(Δδ N/6.51)2 + (Δδ H)2 ]0.5

with a diffusion delay of 125 ms, and gradient amplitudes varied in the range of 6−60 G/cm (21 values arrayed). Signal attenuation was fitted to the Stejskal−Tanner equation;35 the obtained average diffusion constants (D) were D = 1.27 ± 0.01 × 10−10 m2/s and D = 1.20 ± 0.01 × 10−10 m2/s for the apo and HM-bound protein, respectively. Solution Structure Calculation of the WT Apo FimHL. The apo FimHL 3D 15N- and 13C-NOESY-HSQC spectra were peak picked with the assistance of the ATNOS software36 from the UNIO’10 package (version 2.0.2). The extracted NOE cross-peaks were then checked manually prior to being used as an input for automated assignment and structure calculations in CYANA version 2.113 together with a combination of dihedral restraints predicted from TALOS+37 and DANGLE.38 Final structure refinement in explicit solvent was performed using the RECOORD protocol.39 The 25 lowest-energy structures were retained for the final analysis and structure validation in CING.14 Crystal Structure Determination of the WT FimHL−HM. Crystals of the HM-bound FimHL were observed after six months of storage at 277 K of a 0.5 mM sample (∼8.5 mg/mL protein) in D2O containing 20 mM sodium phosphate/100 mM NaCl at pH 6.0. A series of these crystals were cryoprotected by transferring them to drops consisting of 1 M HM in 20 mM sodium phosphate/100 mM NaCl pH 6.0/25% glycerol. The crystals were subsequently vitrified in liquid nitrogen. X-ray data were collected at the PROXIMA-1 beamline of the SOLEIL synchrotron (Gif-Sur-Yvette, France). The structure was determined by molecular replacement using the structure of apo-FimH (Protein Data Bank (PDB) number 4AUU) as a search model. The coordinates from heptyl α-D-mannose were docked into the map from the initial molecular replacement model resulting from PHASER, and the resulting model was subsequently refined. The final model was obtained after alternating cycles of refinement with phenix.ref ine40 and manual build using Coot41 and has an Rfree of 15.5% and Rwork of 19.2% with excellent statistics (see Table S3, Supporting Information). SAXS Studies of the WT Apo and HM-Bound FimHL. Small angle X-ray scattering (SAXS) studies were conducted at the PETRA 12 beamline (DESY, Hamburg, Germany). Prior to analysis, the sample was dialyzed into a buffer matching the NMR conditions (20 mM Mes, 100 mM NaCl pH 6.0) and concentrated (Vivaspin Hydrosart 5000 cutoff) to 7.5 mg/mL (0.44 mM). The dialysis buffer was filtered (0.22 μm) and kept as a buffer blank for the experiments. For the apo FimHL, aliquots of the sample were prepared at different concentrations (2.0, 4.0, 6.0, and 7.5 mg/mL) by diluting the protein stock with the filtered dialysis buffer. The holo FimHL stock was prepared by adding a 5-fold excess of HM to the apo protein, and different concentrations of the holo FimHL were prepared as for the apo FimHL. For both forms, scattering curves were collected at the different protein concentrations and the buffer was measured before and after each sample measurement. The samples were exposed for 0.05 s to the beam with flow of 0.2 mL/min. The collected SAXS data were processed and analyzed with the ATSAS package.20 After buffer subtraction, the scattering curves were compared. For the apo FimHL, this analysis revealed a slight concentrationdependency. The scattering curves were analyzed and merged accordingly to yield a final scattering curve,42 which was used for ab initio modeling. Briefly, 20 ab initio models were generated by DAMMIF43 and averaged using DAMAVER.44 A starting bead model was extracted from the averaged dummy-atom model for the calculation of a final model in DAMMIN.45 The scattering curve of one of the structures of the NMR ensemble (model 21) was compared to the experimental SAXS data set using CRYSOL.46 For the holo FimHL, the comparison of buffer-subtracted scattering curves revealed a significant concentration effect believed to be the effect of HM-induced FimHL dimerization. The scattering curves measured at different concentrations were not merged and were thus treated separately. Each curve was analyzed using OLIGOMER20 to obtain the fractions of monomeric and dimeric holo FimHL in solution. The same SAXS measurements and data analysis were performed on apo and holo FimHL at pH 7.4. Therefore, samples at 2.0, 4.0, 6.0,

(1)

where ΔδN and ΔδH are the chemical shift perturbations of the amide nitrogen and proton, respectively. NMR Backbone Dynamics Experiments. The backbone 15N R1, R2, and Carr−Purcell−Meiboom−Gill (CPMG) R2 relaxation dispersion experiments were typically recorded at 600 MHz on ∼1.1 mM [15N] single-labeled apo and HM-bound protein samples in 20 mM sodium phosphate/100 mM NaCl pH 6.0/6% D2O. For the R1 and R2 measurements, additional data were acquired at different protein concentrations and/or different buffers (see main text for details). Relaxation values were obtained from series of 2D experiments with coherence selection achieved by pulse field gradients using the experiments described previously.31 The 15N R1 and R2 relaxation rates were measured from spectra with different relaxation delays: 100 (in duplicate), 200, 300, 400, 500, 700, 900, 1200, and 1500 (in duplicate) ms for R1, and 10 (in duplicate), 30 (in duplicate), 50, 70, 90, 110, 150, and 190 ms for R2. The CPMG relaxation dispersion experiments32 were recorded with 0, 25, 50 (in duplicate), 75, 125, 175, 225, 350, 550, 750 (in duplicate), and 1000 Hz pulse repetition rates. All data were processed in NMRPipe,26 and the relaxation parameters and their corresponding errors were extracted with CCPN.27 NMR Relaxation Analysis. Starting from the experimental R2/R1 values obtained for the 15N backbone atoms, the diffusion tensor of the apo FimH was obtained with R2R1_diffusion or Quadric_diffusion software packages from A. Palmer’s lab. The residues with the measured 15N R2 higher than one standard deviation from the average ⟨R2⟩ (obtained for the entire set of the FimH backbone amides) likely exhibit significant internal motions18 and, thus, were excluded from the analysis. The model selection was performed using the F statistics as described elsewhere.33 The R2/R1 and the hydrodynamic parameters for the FimH monomer and dimer were calculated with HYDRONMR34 from the atomic coordinates of the apo FimH (lowest-energy NMR structure, this work) and the head-to-tail FimH−HM dimer (X-ray structure, this work), respectively. For the latter, the relaxation parameters were averaged over the values calculated for each of the two individual FimH molecules constituting the dimer. As explained above, the residues with the measured 15N R2 higher than one standard deviation from ⟨R2⟩ were excluded from further analysis. For each backbone amide in the final data set, the population-weighted average of the R2/ R1 ratios, (R2/R1)calcd,i, was obtained at varying populations of the dimer, p, eq 2:

(R 2/R1)calcd,i = p(R 2/R1)dimer,i + (1 − p)(R 2/R1)monomer,i

(2)

At each p value, the agreement between the experimentally determined (R2/R1)exp and the (R2/R1)calcd for the entire protein was assessed by calculating the Q factor (eq 3):

Q=

∑i [(R 2/R1)calcd, i − (R 2/R1)exp , i ]2 2 ∑i (R 2/R1)exp ,i

(3)

The lower the Q factor, the better the agreement between the calculated parameters and the experimental data, with the best solution for the p value found at the global minimum of the Q = f(p) function (Figure 7B). NMR Diffusion Measurements. Samples containing ∼1.1 mM [15N] single-labeled apo or HM-bound FimHL in 20 mM sodium phosphate/100 mM NaCl pH 6.0/6% D2O were measured on the Varian NMR Direct-Drive Systems 800 MHz instrument. Dbppsteghsqcse (with ni = 1 and phase = 1) experiments were run at 298 K 1424



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and 8.2 mg/mL were prepared in the same way as above, except that 20 mM HEPES, 150 mM NaCl pH 7.4 was used as dialysis and dilution buffer. Isothermal Titration Calorimetry. Experiments were performed at 25 °C in an ITC200 calorimeter (GE Healthcare, USA). All the protein and sugar solutions were prepared in 20 mM sodium phosphate/100 mM NaCl pH 6.0 and degassed before use. For the direct titrations, the HM at 200 μM was titrated into a 10 μM solution of either the FimHL wt or its Y48A mutant. For the reverse titration, the FimHL wt at 400 μM was titrated into HM at 12 μM. Titrations were performed with 26 injections of 1.5 μL, using a delay between injections of 140 s. In each case, the first data point was discarded, and the integrated data were fitted to a binding model assuming n identical and independent sites using MicroCal Origin 7.0.

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described in this work. We are grateful to the BIO-NMR Transnational Access program for providing the measurement time at the Utrecht NMR facility (TA project BIO-NMR00095). The Petra III (DESY, Hamburg, Germany) and PROXIMA1 (SOLEIL synchrotron Gif-sur-Yvette, France) beam lines are acknowledged for collection of the SAXS and crystallographic data, respectively. S.V. acknowledges financial support of the framework for bilateral cooperation between the Fonds voor Wetenschappelijk Onderzoek Vlaanderen (FWO) and the Ministère du développement économique, de l’innovation et de l’exportation (MDEIE), Québec (research project G.A060.10N). The Hercules Foundation financed the equipment and infrastructure used in this work. A.N.V. and A.G.P. are FWO postdoctoral researchers, W.V. is a Brains Back to Brussels fellow, and Y.S. is a FWO predoctoral researcher.

ASSOCIATED CONTENT

S Supporting Information *

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CING analysis of the apo FimHL NMR structure, RMSD between the apo FimHL NMR structure and deposited crystal structures of FimHL complexes, details of the X-ray data collection and refinement of the FimHL−HM complex, [1H,15N]-HSQC spectrum of the apo wt FimHL, local RMSDs of the FimHL crystal structures, NOE coverage of the apo FimHL solution structure, backbone-amide NMR relaxation profiles of the apo and HM-bound wt and Y48A FimHL, SAXS scattering curves and data analysis of apo and HM-bound FimHL in HEPES buffer, chemical shift perturbation analysis of Y48A FimHL, and the ITC titration of HM into Y48A FimHL. This material is available free of charge via the Internet at http://pubs.acs.org.

ABBREVIATIONS USED AEBSF, 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride; DOSY, diffusion ordered spectroscopy; FimHL, FimH carbohydrate-binding domain, also referred to as FimH lectin domain; HEPES, 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid sodium salt; HM, α-D-heptyl mannose; heptyl α-Dmannopyranoside; IPTG, isopropyl β-D-1-thiogalactopyranoside; ITC, isothermal titration calorimetry; Mes, 2-(Nmorpholino)ethanesulfonic acid; UPEC, uropathogenic E. coli; upl, upper limit distances; UTI, urinary tract infection; SAXS, small angle X-ray scattering

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Accession Codes

The FimHL solution structure and FimHL−HM crystal structure have been deposited at the PDB data bank (entries 3zpd and 4lov, respectively). All chemical shifts and T1 and T2 relaxation data have been deposited at the BMRB (ID19066 for the wt FimHL, ID19256 for the wt HM-bound FimHL, ID19255 for the apo Y48A FimHL mutant, and ID19254 for the HM-bound Y48A FimHL mutant).

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REFERENCES

(1) Le Trong, I.; Aprikian, P.; Kidd, B. A.; Forero-Shelton, M.; Tchesnokova, V.; Rajagopal, P.; Rodriguez, V.; Interlandi, G.; Klevit, R.; Vogel, V.; Stenkamp, R. E.; Sokurenko, E. V.; Thomas, W. E. Structural basis for mechanical force regulation of the adhesin FimH via finger trap-like beta sheet twisting. Cell 2010, 141, 645−655. (2) Wellens, A.; Lahmann, M.; Touaibia, M.; Vaucher, J.; Oscarson, S.; Roy, R.; Remaut, H.; Bouckaert, J. The tyrosine gate as a potential entropic lever in the receptor-binding site of the bacterial adhesin FimH. Biochemistry 2012, 51, 4790−4799. (3) Klein, T.; Abgottspon, D.; Wittwer, M.; Rabbani, S.; Herold, J.; Jiang, X.; Kleeb, S.; Lüthi, C.; Scharenberg, M.; Bezençon, J.; Gubler, E.; Pang, L.; Smiesko, M.; Cutting, B.; Schwardt, O.; Ernst, B. FimH antagonists for the oral treatment of urinary tract infections: From design and synthesis to in vitro and in vivo evaluation. J. Med. Chem. 2010, 53, 8627−8641. (4) Han, Z.; Pinkner, J. S.; Ford, B.; Obermann, R.; Nolan, W.; Wildman, S. A.; Hobbs, D.; Ellenberger, T.; Cusumano, C. K.; Hultgren, S. J.; Janetka, J. W. Structure-based drug design and optimization of mannoside bacterial FimH antagonists. J. Med. Chem. 2010, 53, 4779−4792. (5) Schwardt, O.; Rabbani, S.; Hartmann, M.; Abgottspon, D.; Wittwer, M.; Kleeb, S.; Zalewski, A.; Smiesko, M.; Cutting, B.; Ernst, B. Design, synthesis and biological evaluation of mannosyl triazoles as FimH antagonists. Bioorg. Med. Chem. 2011, 19, 6454−6473. (6) Pang, L.; Kleeb, S.; Lemme, K.; Rabbani, S.; Scharenberg, M.; Zalewski, A.; Schädler, F.; Schwardt, O.; Ernst, B. FimH antagonists: structure-activity and structure-property relationships for biphenyl αD-mannopyranosides. ChemMedChem 2012, 7, 1404−1422. (7) Firon, N.; Ashkenazi, S.; Mirelman, D.; Ofek, I.; Sharon, N. Aromatic alpha-glycosides of mannose are powerful inhibitors of the adherence of type 1 fimbriated Escherichia coli to yeast and intestinal epithelial cells. Infect. Immun. 1987, 55, 472−476. (8) Bouckaert, J.; Berglund, J.; Schembri, M.; De Genst, E.; Cools, L.; Wuhrer, M.; Hung, C. S.; Pinkner, J.; Slättegård, R.; Zavialov, A.; Choudhury, D.; Langermann, S.; Hultgren, S. J.; Wyns, L.; Klemm, P.; Oscarson, S.; Knight, S. D.; De Greve, H. Receptor binding studies

AUTHOR INFORMATION

Corresponding Authors

*Phone: (+32) 2 629 1025; fax: (+32) 2 629 1963; e-mail: [email protected]. *Phone: (+32) 2 629 3553; fax: (+32) 2 629 1963; e-mail: [email protected]. Present Address ¶

Unité de Glycobiologie Structurale et Fonctionnelle, UMR du CMRS 8576, Bâtiment C9, Avenue Mendeleiev, Université des Sciences et Technologies de Lille 1, 59 655 Villeneuve d’Ascq, France. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Klaartje Houben (Bijvoet Center for Biomolecular Research, Utrecht University, The Netherlands), Evgeny Tishchenko and Peter Sandor (Agilent, Santa Clara, CA) for the acquisition of NMR data and useful discussions on their interpretation, and Tze Chieh Shiao (Department of Chemistry, Université du Québec à Montréal, Canada) for the α-D-heptyl mannose synthesis. Prof. De Greve (VUB, Brussels) is acknowledged for providing the template DNA, subsequently used in the PCR cloning of the expression vector 1425



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disclose a novel class of high-affinity inhibitors of the Escherichia coli FimH adhesin. Mol. Microbiol. 2005, 55, 441−455. (9) Wellens, A.; Garofalo, C.; Nguyen, H.; Van Gerven, N.; Slättegård, R.; Hernalsteens, J. P.; Wyns, L.; Oscarson, S.; De Greve, H.; Hultgren, S.; Bouckaert, J. Intervening with urinary tract infections using anti-adhesives based on the crystal structure of the FimHoligomannose-3 complex. PLoS One 2008, e2040. (10) Abgottspon, D.; Ernst, B. In vivo evaluation of FimH antagonists - a novel class of antimicrobials for the treatment of urinary tract infections. Chimia 2012, 66, 166−169. (11) Cusumano, C. K.; Pinkner, J. S.; Han, Z.; Greene, S. E.; Ford, B. A.; Crowley, J. R.; Henderson, J. P.; Janetka, J. W.; Hultgren, S. J. Treatment and prevention of urinary tract infection with orally active FimH inhibitors. Sci. Transl. Med. 2011, 3, 109ra115. (12) Güntert, P.; Mumenthaler, C.; Wüthrich, K. Torsion angle dynamics for NMR structure calculation with the new program DYANA. J. Mol. Biol. 1997, 273, 283−298. (13) Herrmann, T.; Güntert, P.; Wüthrich, K. Protein NMR structure determination with automated NOE assignment using the new software CANDID and the torsion angle dynamics algorithm DYANA. J. Mol. Biol. 2002, 319, 209−227. (14) Doreleijers, J. F.; Sousa da Silva, A. W.; Krieger, E.; Nabuurs, S. B.; Spronk, C. A. E. M.; Stevens, T. J.; Vranken, W. F.; Vriend, G.; Vuister, G. W. CING: an integrated residue-based structure validation program suite. J. Biomol. NMR 2012, 54, 267−283. (15) DeLano, W. L. The PyMOL Molecular Graphics System; DeLano Scientific: Palo Alto, 2002. (16) Pettersen, E. F.; Goddard, T. D.; Huang, C. C.; Couch, G. S.; Greenblatt, D. M.; Meng, E. C.; Ferrin, T. E. UCSF Chimera - a visualization system for exploratory research and analysis. J. Comput. Chem. 2004, 25, 1605−1612. (17) Palmer, A. G., III. NMR characterization of the dynamics of biomacromolecules. Chem. Rev. 2004, 104, 3623−3640. (18) Rule, G. S.; Hitchens, T. K. Fundamentals of Protein NMR Spectroscopy; Springer: Dordrecht, 2006; pp 431−474. (19) Bouckaert, J.; Li, Z.; Xavier, C.; Almant, M.; Caveliers, V.; Lahoutte, T.; Weeks, S. D.; Kovensky, J.; Gouin, S. G. Heptyl α-Dmannosides grafted on a β-cyclodextrin core to interfere with Escherichia coli adhesion: An in vivo multivalent effect. Chemistry 2013, 19, 7847−7855. (20) Petoukhov, M. V.; Franke, D.; Shkumatov, A. V.; Tria, G.; Kikhney, A. G.; Gajda, M.; Gorba, C.; Mertens, H. D. T.; Konarev, P. V.; Svergun, D. I. New developments in the ATSAS program package for small-angle scattering data analysis. J. Appl. Crystallogr. 2012, 45, 342−350. (21) Morris, G. M.; Huey, R.; Lindstrom, W.; Sanner, M. F.; Belew, R. K.; Goodsell, D. S.; Olson, A. J. Autodock4 and AutoDockTools4: Automated docking with selective receptor flexiblity. J. Comput. Chem. 2009, 16, 2785−2791. (22) Jelesarov, I.; Bosshard, H. R. Isothermal titration calorimetry and differential scanning calorimetry as complementary tools to investigate the energetics of biomolecular recognition. J. Mol. Recognit. 1999, 12, 3−18. (23) André, S.; Cañada, F. J.; Shiao, T. C.; Largartera, L.; Diercks, T.; Bergeron-Briek, M.; el Biari, K.; Papadopoulos, A.; Ribeiro, J. P.; Touaibia, M.; Solís, D.; Menéndez, M.; Jiménez-Barbero, J.; Roy, R.; Gabius, H. J. Fluorinated carbohydrates as lectin ligands: Biorelevant sensors with capacity to monitor anomer affinity in 19F-NMR-based inhibitor screening. Eur. J. Org. Chem. 2012, 4364. (24) Miroux, B.; Walker, J. E. Over-production of proteins in Escherichia coli: mutant hosts that allow synthesis of some membrane proteins and globular proteins at high levels. J. Mol. Biol. 1996, 260, 289−298. (25) Weiner, M. P.; Costa, G. L.; Schoettlin, W.; Cline, J.; Mathur, E.; Bauer, J. C. Site-directed mutagenesis of double-stranded DNA by the polymerase chain reaction. Gene 1994, 151, 119−123. (26) Delaglio, F.; Grzesiek, S.; Vuister, G. W.; Zhu, G.; Pfeifer, J.; Bax, A. NMRPipe: A multidimensional spectral processing system based on UNIX pipes. J. Biomol. NMR 1995, 6, 277−293.

(27) Vranken, W. F.; Boucher, W.; Stevens, T. J.; Fogh, R. H.; Pajon, A.; Llinas, M.; Ulrich, E. L.; Markley, J. L.; Ionides, J.; Laue, E. D. The CCPN data model for NMR spectroscopy: Development of a software pipeline. Proteins 2005, 59, 687−696. (28) Sattler, M.; Schleucher, J.; Griesinger, C. Heteronuclear multidimensional NMR experiments for the structure determination of proteins in solution employing pulsed field gradients. Prog. Nucl. Magn. Reson. Spectrosc. 1999, 34, 93−158. (29) Yamazaki, T.; Forman-Kay, J. D.; Kay, L. E. Two-dimensional NMR experiments for correlating 13Cb and 1Hd/e chemical shifts of aromatic residues in 13C-labeled proteins via scalar couplings. J. Am. Chem. Soc. 1993, 115, 11054−11055. (30) Higman, V. A. Protein NMR: A practical guide. http://www. protein-nmr.org.uk (accessed January 29, 2014). (31) Farrow, N. A.; Muhandiram, R.; Singer, A. U.; Pascal, S. M.; Kay, C. M.; Gish, G.; Shoelson, S. E.; Pawson, T.; Forman-Kay, J. D.; Kay, L. E. Backbone dynamics of a free and phosphopeptidecomplexed Src homology 2 domain studied by 15N NMR relaxation. Biochemistry 1994, 33, 5984−6003. (32) Palmer, A. G., III; Kroenke, C. D.; Loria, J. P. Nuclear magnetic resonance methods for quantifying microsecond-to-millisecond motions in biological macromolecules. Methods Enzymol. 2001, 339, 204−238. (33) Mandel, A. M.; Akke, M.; Palmer, A. G., III. Backbone dynamics of Escherichia coli ribonuclease H1: Correlations with structure and function in an active enzyme. J. Mol. Biol. 1995, 246, 144−163. (34) García de la Torre, J.; Huertas, M. L.; Carrasco, B. HYDRONMR: Prediction of NMR relaxation of globular proteins from atomic-level structures and hydrodynamic calculations. J. Magn. Reson. 2000, 147, 138−146. (35) Stejskal, E. O.; Tanner, J. E. Spin diffusion measurements: Spin echoes in the presence of a time-dependent field gradient. J. Chem. Phys. 1965, 42, 288−292. (36) Herrmann, T.; Güntert, P.; Wüthrich, K. Protein NMR structure determination with automated NOE-identification in the NOESY spectra using the new software ATNOS. J. Biomol. NMR 2002, 24, 171−189. (37) Shen, Y.; Delaglio, F.; Cornilescu, G.; Bax, A. TALOS+: A hybrid method for predicting protein backbone torsion angles from NMR chemical shifts. J. Biomol. NMR 2009, 44, 213−223. (38) Cheung, M. S.; Maguire, M. L.; Stevens, T. J.; Broadhurst, R. W. DANGLE: A Bayesian inferential method for predicting protein backbone dihedral angles and secondary structure. J. Magn. Reson. 2010, 202, 223−233. (39) Nederveen, A. J.; Doreleijers, J. F.; Vranken, W. F.; Miller, Z.; Spronk, C. A. E. M.; Nabuurs, S. B.; Güntert, P.; Livny, M.; Markley, J. L.; Nilges, M.; Ulrich, E. L.; Kaptein, R.; Bonvin, A. M. J. J. RECOORD: A recalculated coordinate database of 500+ proteins from the PDB using restraints from the BioMagResBank. Proteins 2005, 59, 662−672. (40) Afonine, P. V.; Grosse-Kunstleve, R. W.; Adams, P. D. The Phenix refinement framework. CCP4 Newsl. 2005, 42, 1−7. (41) Emsley, P.; Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr., Sect. D: Biol. Crystallogr. 2004, 60, 2126−2132. (42) Mangenot, S.; Leforestier, A.; Vachette, P.; Durand, D.; Livolant, F. Salt-induced conformational and interaction changes of nucleosome core particles. Biophys. J. 2002, 82, 345−356. (43) Franke, D.; Svergun, D. I. DAMMIF, a program for rapid ab initio shape determination in small-angle scattering. J. Appl. Crystallogr. 2009, 42, 342−346. (44) Volkov, V. V.; Svergun, D. I. Uniqueness of ab initio shape determination in small-angle scattering. J. Appl. Crystallogr. 1993, 36, 860−864. (45) Svergun, D. I. Restoring low resolution structure of biological macromolecules from solution scattering using simulated annealing. Biophys. J. 1999, 76, 2879−2886. (46) Svergun, D. I.; Barberato, C.; Koch, M. H. J. CRYSOL: A program to evaluate X-ray solution scattering of biological macro1426



Article

molecules from atomic coordinates. J. Appl. Crystallogr. 1995, 28, 768− 773.

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